LAS Pro July-August 2021 by aalasoffice

July/August 2021

Focusing on the Possibilities: Exploring LAS Trends Diving In: Water Quality and Pain Management in Fish Forecasting the Future, 3D Printing, and LAS Innovations Veiled Chameleon Husbandry

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July/August 2021 Vol. 9 Issue 4

INSIDE THIS ISSUE...

Zebrafish Water Quality Challenges

Pain Management in Fish

A Future Forward Focus

Exploring NHP Shortage Solutions

Wellbeing and Safe Workplaces

3D Printing Innovations

Zebrafish husbandry hinges on water quality, a complicated chemical process.

Fish have become increasingly popular animal models leading to discussions regarding how potential pain in fish is approached and managed.

AALAS leadership discusses thoughts on the future of LAS, our association, and member services.

Health, safety, and global competitiveness depends on appropriate animal model availability.

Incorporate nontraditional employee measurements into staff hiring and retention strategies.

Discover how using 3D printing solutions can meet the needs of LAS professionals and enhance facility operations.

July/August 2021

Focusing on the Possibilities: Exploring LAS Trends Diving In: Water Quality and Pain Management in Fish Forecasting the Future, 3D Printing, and LAS Innovations Veiled Chameleon Husbandry

2 Laboratory Animal Science Professional July 2021

On the cover: Pictured with Mina the chameleon is Alex Muensch, BS, MS, LATG, a Research Technician II at the Stowers Institute for Medical Research in Kansas City, MO. Photo provided by Philippe Noguera, Stowers Institute for Medical Research.

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DEPARTMENTS 5 Publisher’s Note

52 Tech Tips

6 People & Places

64 Across the Pond

Embracing the future

New hires, promotions, awards, memorials

8 PROfiles

Meet Frankie Howell

Insights on techniques and tactics

Meetings in the wake of COVID-19

66 AALAS Foundation News

Support the AALAS Foundation through their 2021 fundraising events

36 Inside the IACUC

67 Crossword

40 Career & Training

68 Ad Index

Communication channels

Connection opportunities, teamwork, and pilot program launches

48 DIY

Watermelon training chips

4 Laboratory Animal Science Professional July 2021

Zebrafish

PUBLISHER’S NOTE Staff Publisher Ann Turner Associate Publisher Chris Lyons

Learning from the Past, Embracing the Future

Managing Editor John Farrar Associate Editor Liz Rozanski Ad Sales John Farrar

As I was paging through this issue of LAS Pro, I had two vivid memories pop into my mind. I distinctly remember caring for a colony of guppies when I was ten years old. And I also remember time spent in front of a duplication machine preparing worksheet packets for my students when I taught school. The guppy memory was spurred by the two feature articles we have on fish and how they have developed into a vital animal model in biomedical research. The duplication machine is an ancient ancestor to the 3D printers LAS pros are now using to meet needs in their facilities. Beyond aquatics and 3D printing, this issue has several articles focused on the future. When we look at the past and evaluate where we are now, multiple lessons come to mind, and AALAS’ importance to LAS’ future is prominent. When the Animal Care Panel met in Chicago over 75 years ago, the first discussions were about food and diet for animal models. The second discussion focused on caging. Moving from table scraps to a formulated diet was a big step for research. Fast forward multiple decades, and we all know the integral part proper nutrition plays in our facilities and institutions. Caging is a dynamic and critical component in research. The users of wooden cages from years ago would be stunned to review the varieties of cages now available for a host of animal models. Comparing the past to the present and projecting into the future can be challenging. However, one aspect I know will remain constant is the value AALAS has had and will continue to have to LAS pros worldwide. Our members and supporters continue to be engaged and working on discoveries and developing innovations and therapies we may not even know we need yet. As we continue to grow, learn, evolve, and innovate, I encourage you to be proud of our history and look forward to our industry’s prospects. I hope to see all of you in Kansas City, Missouri, for our 72nd AALAS National Meeting in the not-so-distant future. But in the meantime, best wishes for a happy and healthy summer!

Ann Turner

Publisher Executive Director American Association for Laboratory Animal Science

Design/Production Zara Garza

Editorial Advisory Board Leslie Birke Louisiana State Univ Andrew Burich Benaroya Research Institute Bob Dauchy Tulane Univ School of Medicine David DeOrnellis Champions Oncology Penny Devlin Pennsylvania State Univ College of Med Sonia Doss Duke Univ Medical Center Kelly Ethun Emory University Glenn Jackson Cornell University Richard Marble Alpha Genesis Inc Elizabeth Nunamaker Univ of Florida Sara Oglesby Abbvie Karuna Patil Seattle Children's Research Institute Amy Pierce Tulane Univ School of Medicine Stacy Pritt UT Southwestern Medical Center Robin Tucker Georgetown Univ

Mission Statement Laboratory Animal Science Professional (LAS Pro) is the official magazine for American Association for Laboratory Animal Science members. LAS Pro provides a wide range of useful resources and knowledge to the association’s 14,000 laboratory animal science professionals who are involved in advancing responsible laboratory animal care and use to benefit people and animals. All signed articles, including, committee reports, news, and commentary, reflect the individual views of the authors and are not official views of AALAS. Authorization to photocopy portions for personal or internal use is granted by the American Association for Laboratory Animal Science. Photocopying for purposes of resale or outside distribution is prohibited unless written approval is obtained from the AALAS Director of Communications. Copyright 2021 by the American Association for Laboratory Animal Science. Laboratory Animal Science Professional (USPS 010-730) is published bimonthly by the American Association for Laboratory Animal Science, 9190 Crestwyn Hills Drive, Memphis, TN 38125. Periodicals Postage paid at Memphis, TN 38101 and additional mailing offices. POSTMASTER: Send address changes to AALAS, 9190 Crestwyn Hills Drive, Memphis, TN 38125-8538.

American Association for Laboratory Animal Science 9190 Crestwyn Hills Drive Memphis, TN 38125-8538 Phone: 901-754-8620 Fax: 901-753-0046 E-mail: info@aalas.org Web: www.aalas.org

July 2021 Laboratory Animal Science Professional 5

PEOPLE & PLACES

New hires, promotions, meeting updates, and memorials.

Congratulations! AALAS Recognizes Certification Achievements

ALAS certifications are a recognized path to climbing the laboratory animal science career ladder. Gregory Lawson, DVM, PhD, DACLAM, recently contacted AALAS to celebrate Kayla Lynch earning her Certified Manager of Animal Resources (CMAR) certificate. Coincidentally, the day Kayla received her notification, she also received an employee recognition award! Dr. Lawson shared Kayla’s good news with his colleagues and noted that the CMAR certification is a 1–2-year study commitment and is just one of the AALAS certifications. “Completion, as with the ALAT, LAT, and LATG, takes a commitment to studying during off time, even when tired and mentally exhausted. Kayla is a great example of committing to a task and not giving up until it is completed,” Lawson said. Seventeen individuals earned the CMAR in 2020, and so far, 4 have earned the certificate in 2021. AALAS recognizes these achievements. Congratulations!

2020 CMAR • • • • • • • • •

Tasha Decker Heather McKay Jannelle Monnas Heather Bonanno Ashley Dean Carley Batson Carrie List Chad Rancourt Brandon Baker

• • • • • • • •

Amy Parrillo Syed Zaidi Sarah Laraway Wendy Coyne Claudia Diaz Matthew Keller Rachael Koch Shannon Thomas

2021 CMAR • • • • • •

Amy Davis Jessica Hendricks Rachel Rodriguez Kayla Lynch Tiffanie Whitty Nicole Collette

ALAT, LAT, and LATG Certifications

For 2020, 748 technicians achieved the ALAT certification, 401 earned the LAT, and 192 completed the LATG. For the first quarter of 2021, a total of 293 technicians have earned a certification: ALAT, 172; LAT, 86; and LATG, 35. Congratulations!

Standing Committee for the Care and Use of Animals in Research Announced

he National Academies of Sciences, Engineering, and Medicine has announced a standing committee tasked with engaging with stakeholders regarding new processes, formats, and topics for future updates or additions to the Guide for the Care and Use of Laboratory Animals (the Guide). Now in its 8th edition, the Guide serves as the basis for accreditation of institutions worldwide. The standing committee is not an advisory or oversight committee that will make recommendations for changes to the Guide; rather it will provide a venue for the exchange of ideas and knowledge sharing among those involved in scientific research and animal care and use in the academic, 6 Laboratory Animal Science Professional July 2021

government, private, and non-profit sectors. To learn more, visit: https://www.nationalacademies.org/our-work/standingcommittee-for-the-care-and-use-of-animals-in-research Members are: Chair: James G. Fox Members: William W. Bowerman, IV; Douglas L. Brining; Kelly R. Decker; Jeffrey Everitt; Michael E. Goldberg; Susan B. Harper; Richard Nakamura; Nathaniel Powell, Jr.; Robert S. Sikes; Jerrold Tannenbaum; Katherine Thibault; Sue VandeWoude

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minutes with...

Frankie Howell, CMAR, RLATG

PRO-files in LAS Facility/Employer: UT Southwestern Medical Center’s Animal Resource Center Job Title: Manager of Animal Care How did you get in this field? I worked at a veterinary hospital to put

myself through college. One of the veterinarians I worked with asked me to run anesthesia at Howard University for a cardiac research procedure on a cat. After the procedure, she took me on a tour through the animal facility and talked to me about opportunities the field had to offer. From that moment on, I knew I wanted to shift my career to laboratory animal research management and started working toward that goal.

Who were your mentors? I’ve had several mentors. Jack Dyer, my high

school Ag teacher; Dr. Kathryn Nepote, DMV, the Attending Veterinarian at the University of Maryland; Dr. Tannis Johnson, DVM, APAW Veterinary Hospital in New Carrollton, Maryland; Sheryl Richeson at Johns Hopkins University; and Julie M. Wood, CMAR.

What are your current interests in animal science? Everything. I come

from a farming and ranching background and have always been an animal lover to my core. I am interested in everything from improvements in food production to innovations in companion animal medicine and techniques. I’m also interested in pure research and animal welfare in academia.

Where do you see yourself in 5 years? I hope to continue leading and

mentoring a team of leaders, working with scientists in a premier animal research institution, and looking for the next generation of laboratory animal professionals to develop.

Getting Personal

What is your favorite part of your job? I like seeing someone I hired and mentored be successful and accomplish goals they never dreamed they could achieve.

What companion animals do you have?

What advice do you have for others just beginning their animal science career? Learn as much as you can every single day. Challenge yourself. Talk to

I have 3 dogs: an American Pitbull, Miss Daphne Delilah of York; a Jack Russell and Dachshund mix, Cleopatra Gingersnap Schnitzlehooven; and a Labrador and Aussie Shepherd mix, Gangsta Orphan Annie of Balko.

Best binge-watching TV series? Yellowstone

What is the last book you read?

A Team of Leaders – Empowering Every Member to Take Ownership, Demonstrate Initiative, and Deliver Results by Paul Gustavson and Stewart Liff.

Where is your favorite vacation spot?

I love the Hill Country in Texas, but if you name a river where I can float and be in the sun and nature, that place might be my favorite, too.

What is your favorite dessert? Angel Food Cake

your investigative staff and learn about the research they are performing with the animals you care for. Get your certifications. Network with others in the field. Get involved in your AALAS branch. Make this a career because the field has far more to offer than “just a job.”

What is the most rewarding aspect of your career? Direct rewards are

helping people accomplish their career goals. Indirect rewards are seeing the scientists I work with publish articles on the discoveries they made through using the animal models housed in our animal facilities. It is rewarding to know we are helping to change the world.

What impacts have the pandemic had on your work experience?

Early on, it was a little rough. We were writing the playbook as contact tracings, and potential mass exposure situations occurred. We were thinking about risk management and how to prepare for the worst scenarios while still making sure the staff, animals, and scientists had what they needed to keep the research going. Learning how to mitigate fear as a leader became a daily topic when dealing with employees and educating them on how to stay safe. Together, we got through the crisis without sacrificing quality in our work or welfare to the animals. Through strong leadership and education, the animal care staff let their purpose drive them through the fear of potential exposures while they were at work and they became the definition of "essential" by coming to work every day while others stayed home. While the pandemic is not yet over, we have found our rhythm. Work is ramping back up here at UT Southwestern and we are starting to be able to work more closely together again. I miss the days when we could get the team together in a large group for team building and celebratory events. It is difficult to believe it has been over a year since this all began, and who knows when it will end.

8 Laboratory Animal Science Professional July 2021

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Zebrafish are freshwater fish, which are reasonably tolerant of a range of water parameters compared to other common species. 10 Laboratory Animal Science Professional July 2021

FEATURE

Aquatics

Zebrafish Water Quality for Laboratory Animal Technicians By Christine Archer RVT, RLAT

ebrafish are an increasingly popular model organism in many aspects of animal research; however, some aspects of their husbandry needs are quite disparate from that which most lab animal technicians are trained. Possibly the most important aspect of zebrafish husbandry is water quality, which has impacts on all areas of the life of a laboratory zebrafish. Water quality is a very complicated chemical concept, with the different parameters often intertwined in ways that seem daunting for many technicians. However, with a solid grasp of the different parameters of fresh water and how they interact with each other and have implications for fish health, lab animal technicians can establish a solid background to manage a recirculating water system housing zebrafish. Zebrafish are freshwater fish, which are reasonably tolerant of a range of water parameters compared to other common species. However, wide shifts in water parameters are stressful on fish, so it is important that facilities work to define a standard normal parameter set to maintain via regular testing and system maintenance. Many zebrafish facilities can share or even publish their water parameter ranges, which may be a good start for establishing what normal values to strive for in a zebrafish facility. Since water parameters can impact on fish behavior and research outcomes, having the system parameters recorded and available to researchers is also valuable when it comes time to publish. The water parameters most important for freshwater fish like zebrafish are water temperature, pH, conductivity, alkalinity, hardness, dissolved gas, and nitrogenous waste. Each must be monitored regularly and logged to monitor for changes or abnormalities over time. A simple spreadsheet can be used or printed and filled in by hand as part of a maintenance and health check binder. In the case of multiple systems requiring monitoring, a water quality binder kept in the area where most water tests are conducted can be helpful. Water tests come in various forms, from simple dipsticks common in the pet trade to state-of-the-art photometers with digital outputs and data tracking. Regardless of the methods used, what matters is that the tests utilized are reliable and repeatable. Wherever possible, have backup testing methods (i.e., a dipstick test to back up a photometer tester) to double-check and triple-check any unusual test results. What follows is a summary of the most important water parameters, implications for fish health, frequency of monitoring, and how parameters may impact each other.

Water Temperature

Zebrafish are poikilotherms. This means their body temperature varies with the ambient temperature of their environment. While zebrafish can tolerate a wide temperature range from 20°C-40°C, most housing for juvenile and adult zebrafish in research is maintained at 24°C-28°C. Due to commonly utilized developmental staging publications, zebrafish embryos are often incubated at 28.5°C. Water temperature can impact almost all other water parameters, so it is very important that it remains as stable as possible. Temperature shifts of more than a degree Celsius over 24 hours can be stressful for zebrafish. Water has a fairly high heating capacity compared to room air, so heating or cooling water can take some time, which can be beneficial when trying to maintain consistent water temperatures. Water temperature should be monitored constantly in a system, using at least one temperature probe, preferably hooked up to an alarm system that will notify users if the temperature shifts outside an acceptable range. If possible, set the range within less than 2 degrees Celsius on either side of the desired temperature setpoint. Water temperature should also be checked manually daily during system checks, and daily temperatures should be recorded. If daily temperatures are changing more than a degree day to day, operations of the system and ability to heat water or room air should be assessed. If it is not possible to maintain steady warmer temperatures, it is better to run a zebrafish system at a cooler temperature that can be kept stable over a long period.

pH, also known as the power of hydrogen, is a complex topic. However, at its root in the context of zebrafish water quality, pH is a measurement of how acidic or basic the water is. pH is measured on a logarithmic scale from 0 to 14, with 0 being most acidic and 14 being most basic. 7 is neutral pH, in between acidic and basic. On this type of scale, a pH change of 1 unit represents a significant change in the pH of the water, on a factor of 10. Therefore, even a change of one pH unit in a short time can cause significant stress to a fish, as pH changes will impact most other water parameters. There is no standardized pH value known for zebrafish, and they’ve been observed in a range of slightly acidic (pH ~6) to slightly basic (pH ~8) environments. However, in the closed recirculating systems laboratory zebrafish live in, the organJuly 2021 Laboratory Animal Science Professional 11

FEATURE

Aquatics be achieved by adding a buffer to the water system, such as sodium bicarbonate. Low alkalinity, below 50mg/L, could put a fish system at risk of pH crash. Alkalinity should be monitored weekly and adjusted as needed by the gradual addition of sodium bicarbonate until a range of 50-150mg/L is achieved.

Hardness Zebrafish (Danio rerio).

isms that help reduce and remove nitrogenous wastes thrive at a pH close to 7, or ‘neutral.’ Most zebrafish systems will be maintained within a one half unit of this range, and pH should be monitored daily at a minimum. Most commercial zebrafish systems can have a pH probe installed, which will provide constant monitoring and alarm, should the pH move out of the desired range, usually if pH changes more than a half unit on either side of the setpoint. Double-check the pH probe values with a second pH tester, using either a handheld meter or test strips.

Conductivity

Coarsely, conductivity is a measure of the salinity of water, measured in microsiemens (µS). While zebrafish are freshwater fish, they can benefit from a small amount of dissolved salts in their water. Freshwater fish are constantly under some level of osmotic stress, as their body systems work to counter the loss of ions and water infiltration from their surroundings. The fish body is saltier than the water they live in, and osmosis is working to balance the water and the internal fish environment. Have slightly saltier water can reduce the systemic demands on zebrafish from this process. Zebrafish are often kept at below 2000µS. In context, seawater is approximately 50,000µS, and tap water ranges from 200-1000µS depending on region. There is no standard conductivity level for zebrafish, but setpoints in most fish facilities will be in the range of 500-2000µS, and small amounts of sea salt are added to maintain the desired values. It is important to keep the conductivity reasonably stable, not permitting it to change more than 50-100µS per day. Conductivity should be measured daily, and much like pH and temperature, in-dwelling probes exist, which can be installed in a zebrafish system for constant monitoring and alarming purposes. Conductivity can be double-checked with a handheld digital meter.

Alkalinity

Some basic ions, like bicarbonates in solution, will impact many aspects of water parameters. In the context of a zebrafish housing system, alkalinity is the ability of the water itself to resist acidification. Alkalinity should be at a level that will help to prevent large pH swings due to increasing carbon dioxide gas levels or after all the fish have been fed. This can 12 Laboratory Animal Science Professional July 2021

Certain mineral ions in water, such as calcium or magnesium, will increase the ‘hardness’ of the water, which is measured in mg/L like alkalinity. The water hardness in a zebrafish system is often correlated with both the alkalinity and the conductivity. The main source of the ions that contribute to hardness comes from the sea salt used to control conductivity, and in some cases the buffer solution used to maintain an acceptable alkalinity level. There is no known best water hardness value for zebrafish, but it should remain stable and be monitored weekly. Measured hardness values and how they change relative to those measured for conductivity and alkalinity should be noted, as they may help to indicate issues with stabilizing the other parameters.

All common atmospheric gases are found in solution in freshwater, but oxygen (O2), carbon dioxide (CO2), and Nitrogen (N2) are the most important gases to be aware of in freshwater systems. They can be more challenging to monitor than other water parameters. Knowledge of how their relative saturation can impact water systems is important when considering a healthy water system for zebrafish. Oxygen is necessary for zebrafish to live. They absorb oxygen dissolved in water primarily across their gills and partially via other body surfaces in contact with water. Dissolved oxygen levels can deplete quickly when water is stagnant, especially after feeding fish or in crowded tank environments. A heavily loaded zebrafish system may experience dangerously low dissolved oxygen (DO) levels after a feeding has been completed, or if an issue arises with circulation of clean water to the housing tanks. Meters and test kits, and continuous in-system monitoring probes are valuable to ensure DO doesn’t drop below 4mg/L in the system. Test at least weekly or when fish appear distressed and are gasping for air near the surface of the water. Carbon dioxide is a by-product of respiration in the zebrafish, as well as from other organisms living in biofilms and system detritus. It is generally off gassed to the atmospheric air in the room by water agitation as it flows from the tank and

into the water treatment areas. Zebrafish are fairly tolerant of elevated CO2; however, increasing levels can also increase the acidity of the water. Test kits can be used to ensure CO2 levels stay below about 15-20mg/L. Nitrogen is the most abundant gas in the air we breathe, and it is also prevalent in solution in freshwater. However, when situations exist that may cause pressurized atmospheric air to enter the water system, the N2 concentration can reach levels which are harmful or even fatal to fish, causing injuries to blood vessels and multiple organs, and leading to symptoms and eventual death from gas bubble disease, which resembles decompression sickness or “the bends” known to divers. Pressurized air can end up in a water system due to a leaky connection that may suck in air, or when a large bubble or slug of air is left in pipes or filter canisters after maintenance when they are closed up to be restarted. Another source of nitrogen supersaturation is very cold municipal water in cooler climates. Tap water in the winter is often supersaturated with gas. Care must be taken when using this water in topping up systems without first treating or warming the affected water. Supersaturation can be prevented with good aeration of water in systems, which can be achieved with degassing columns or simply by agitating water open to the air during the filtration process. Generally, monitoring for supersaturation events can be accomplished with a meter that measures total gas pressure. It should stay around 100% (pressure relative to barometric pressure), and values at or around 110% or greater are cause for alarm.

Nitrogenous Waste

There are three nitrogenous waste products a zebrafish system is faced with: ammonia, nitrite, and nitrate. These are all related and have varying impacts on other water parameters and fish health overall. The process of nitrification and oxidation of nitrogenous waste products is complex, but a general grasp of what the products are and how they are related to one another is helpful to develop an understanding of their impact on zebrafish health and water system operation. Zebrafish produce ammonia as a waste product. It is excreted primarily via their gills and via urine. Zebrafish are generally fed a high protein diet, which contributes to their ammonia excretion and increased ammonia from the breakdown of uneaten food and fecal waste products. High levels of ammonia can be stressful for zebrafish and may cause injury or death when levels continue to increase. An established zebrafish housing system, usually at least 2-4 months old, will have beneficial bacteria inhabiting a biological filtration system that converts ammonia to nitrite. If this biological filter is not established, ammonia must be removed via regular large water changes or via ammonia adsorbing filter media. Ammonia is present in two forms in a freshwater system, ionized and unionized, the latter of which is the toxic form for fish. When measuring ammonia using most test kits, the reading is generally a sum of the two forms of ammonia, also known as total ammonia nitrogen or TAN. The balance of the two forms of ammonia in a system varies based on water temperature, pH, and conductivity. For the sake of simplicity when over-

Zebrafish produce ammonia as a waste product.

seeing a system as a new technician, strive for a level of zero total ammonia, measured at least weekly or more frequently if ammonia is found to be present. Beneficial bacteria produce nitrite in the water system, which converts ammonia. Nitrite is also toxic to fish and can cause symptoms similar to carbon monoxide poisoning. Along with testing ammonia levels, kits should be used to test nitrite levels at least weekly, and the value should be close to 0. Nitrate is the final product of bacterial biofiltration and is produced when nitrite is converted. Nitrate is far less toxic to fish but should not be permitted to accumulate beyond 100200mg/L, tested at least weekly along with the other nitrogenous waste products. Nitrate is generally removed from the water system via regular water changes, and increasing levels can lead to excess algae growth, even below levels dangerous to fish.

Conclusion

Water chemistries in freshwater systems like those required to keep laboratory zebrafish healthy are somewhat challenging in their complexity, and achieving certain parameters may be difficult depending on the housing situation and specific type of housing system. While zebrafish users and caretakers should aspire to a more standardized set of desired water parameters for laboratory zebrafish across most facilities, the most important thing to consider about water parameters is that they should remain as stable as possible, as long as they are within a safe range for zebrafish health and welfare. In many cases, unstable water parameters can have more detrimental effects than levels that are slightly approaching the range of what is considered “acceptable” for housing zebrafish. Work to keep the system stable, and then work to see if it is possible to gradually change system parameters to the values you desire while still maintaining stability. For more detailed information on water chemistry for zebrafish and the interplay of different parameters, read Chapter 29: Water Quality for Zebrafish of The Zebrafish in Biomedical Research (2019, Academic Press). Christine Archer, RVT, RLAT, is the Zebrafish Facility Coordinator at the University of Colorado’s Anschutz Medical Campus and President-Elect for the Zebrafish Husbandry Association. July 2021 Laboratory Animal Science Professional 13

FEATURE

Aquatics

Fish have become increasingly popular animal models.

Fish Analgesia: Can We Do Betta? By Frederic Chatigny, DVM, MSc and Julie Balko, VMD, DACVAA

espite an ever-growing body of evidence to support the argument that fish are capable of pain perception, there continues to be ongoing controversy as to whether or not fish truly feel pain.3, 13, 5 This contention in combination with the paucity of pharmacologic analgesic data in fish and their status as a lower organism on the sentience scale has led to stark contrast in pain management strategies between fish and mammals. Fish have become increasingly popular animal models in many fields of research: developmental biology, cancer, toxicology, drug discovery, molecular genetics and investigation of various diseases, including Alzheimer’s. 14, 11 As live animal models, they are frequently exposed to noxious stimuli associated with a myriad of diagnostic and therapeutic procedures, including fin biopsies and telemetric device implantation, among others. While such procedures in mammals would unequivocally be coupled with routine pain management, in contrast, the standard use of analgesics in fish is far more limited.

Nociception or Pain…Does It Matter?

The controversy surrounding pain perception in fish is mainly centered upon the generally accepted definitions of nociception and pain. According to the International Association for the Study of Pain (IASP) nociception is “the neural process of encoding noxious stimuli” and pain is “an unpleasant sensory and emotional experience associated with, or resembling that associated with, actual or potential tissue damage.”12 Put in simpler terms, nociception is the messaging system and pain is how one feels about the message. Both sides of the controversy accept the fact that fish experience nociception; they possess nociceptors* and respond to noxious stimuli with a nocifensive response. However, the debate focuses on the line between nociception and pain and whether fish can truly generate an emotional response. This discussion is incredibly complex and calls into question complicated concepts such as consciousness and emotions. While this debate is unlikely to end anytime soon, it should not change how we approach and manage potential pain in fish.

*Elasmobranch, cartilaginous fish of a group that comprises the sharks, rays, and skates, are notable exceptions as nociceptors have not been documented in those species as of yet.

14 Laboratory Animal Science Professional July 2021

The aforementioned IASP definition of pain also includes the caveat that “…inability to communicate does not negate the possibility that a… nonhuman animal experiences pain.” Furthermore, the American College of Veterinary Anesthesia and Analgesia (ACVAA) adds that “…regardless of the clinical signs demonstrated, if there is any doubt that an animal may be experiencing pain, then a trial treatment with analgesics is indicated.”1 Both of these statements support the precautionary principle, which argues that in light of the lack of scientific consensus, it is more appropriate to assume that fish feel pain.9 The authors support this recommendation and argue that this rationale has benefits for both fish and scientist. As mentioned above, individuals on both sides of the fish pain debate agree that fish display nocifensive behaviors and these behaviors may negatively impact research results. Additionally, it is well recognized that unalleviated surgical pain can have a myriad of negative consequences including effects on immune function, food consumption, social behavior, and metabolism, among others.7,4 Thus, while untreated pain can have obvious and significant individual animal welfare implications, from a purely scientific standpoint, it can also have a tremendous impact on study data. Therefore, until proven otherwise, fish should be assumed to experience pain and analgesic therapy provided where appropriate.

Advances in Welfare Assessment

The general principle of welfare assessment in any veterinary species is to detect deviations from normal in both physiologic (e.g., appetite, opercular rate, heart rate, cortisol level) and behavioral (e.g., activity, location in the tank, social dynamics, fear of novel object or space) variables. Welfare assessment in fish is particularly challenging for a host of reasons. First, knowledge of normal (versus abnormal) behavior and physiology may be unknown or may vary wildly between different species and life stages. Second, these variables may also be significantly impacted by the presence of a human observer, in both fish response and human bias. Third, there is currently no optimal or “gold standard” test to assess fish welfare; grimace scales, commonly employed in mammalian species, are not • The use of analgesics in fish is currently applicable of utmost importance, both for in fish and most of optimal animal welfare and sound the generally accepted scientific data. mammalian indicators • The debate surrounding pain of pain are difficult or perception in fish continues, but simply inappropriate this should not impede the provito use in fishes.2 Lastly, sion of appropriate and timely pain there is also signifimanagement. cant bias in that fish • Welfare assessment in fish is challenging, but new tools are pavare blatantly alien to ing the way for more objective and humans as our expericlinically applicable techniques. ences and environment • Immersion offers a refined and differ significantly promising route for analgesic from theirs. This potherapy in fish, but more research tentially muddies the is warranted. water of pain recog-

Key Points

The use of analgesics in fish is of utmost importance, both for optimal animal welfare and sound scientific data.

nition and subsequent observer assessment of analgesic need. This attitude has been called the ‘evidence-threshold’ approach meaning that there is insufficient evidence, as determined by an observer, to conclude that a specific animal is in pain.15 Fish welfare assessment is still in its infancy, but recent advances have attempted to mitigate some of the aforementioned problems. An automated system to document and objectively characterize fish activity has recently been developed.8 Essentially, it uses remote cameras and tracking software to generate large-scale, real-time three-dimensional (3D) assessments of fish activity. That system was then used in the creation of a Fish Behavior Index (FBI) for zebrafish, a sort of compendium that characterizes specific key behaviors exhibited in response to noxious laboratory procedures. This not only aids in determination of normal or anormal behaviors in this species but allows assessment of the efficacy of analgesic drugs. A similar compendium, called the Zebrafish Behavior Catalog (ZBC), has also been proposed.10 All of these automated tools offer non-invasive, objective, and efficient methods of assessing behavior in fish in order to reduce human bias, support scientific inquiry, and improve welfare. While these systems have only been applied in zebrafish and extrapolation to other, especially larger, species should be practiced with caution, the advent of this technology paves the road for similar advances in other fish species.

Immersion Analgesia on the Horizon?

Administration of analgesics via immersion represents an innovative and potentially hugely beneficial pain management strategy for fish. Drug administration to fish, just like for other animals, should minimize physiologic and psychologic stress as much as possible. Injections can be particularly stressful for fish as they commonly require manual restraint (often out of water) which can damage their beneficial biofilm, there is a possibility of pain or tissue damage at the injection site, and serial handling is usually indicated. Additionally, injection of small species or large numbers of fish is often time- and resource-prohibitive. Administration of drugs via immersion offers a refined means to mitigate some of these challenges as this technique is noninvasive and can theoretically be used continuously and/or on multiple animals at once. July 2021 Laboratory Animal Science Professional 15

Welfare assessment in fish is challenging, but new tools are paving the way for more objective and clinically applicable techniques..

entists may be understandably reticent to use analgesics due to concerns for unpredictable or unwanted effects on study results that might outweigh any benefits to animal welfare. While this is a concern worth consideration, published studies generally show a lack of side effects from tested drugs and as previously mentioned, unrelieved pain could have significant effects on study data. Thus, given their positive risk: benefit ratio, scientists should be encouraged to use analgesics in fish whenever pain is suspected. While these studies do demonstrate promise with the use of immersion analgesics in fish, caution should be practiced with interpretation, application, or extrapolation of this data. Attention needs to be given to the particular species, noxious stimulus, and potential author bias.6 There is also the need to consider proper disposal of tank water following use, especially in the case of controlled substances. Considerable interspecies variation has been reported and unforeseen effects could affect experimental results. Whenever possible, pilot studies are recommended to evaluate the most appropriate analgesic strategy for a particular species and research circumstance. As this data would then be of potential use to the larger veterinary community, the authors encourage sharing of the results of any pilot analgesic studies to continue to build on and improve fish welfare. Immersion offers a refined and promising route for analgesic therapy in fish, but more research is warranted.

Objective data regarding the use of immersion analgesics in fish remains scarce. While there is presently no overwhelming evidence of efficacy for most analgesics in fish, especially when administered via immersion, beneficial effects on behavior and physiologic parameters have been reported.6 This is especially true for opioids, particularly morphine, as well the local anesthetic drug lidocaine. Beneficial effects have also been observed with some nonsteroidal anti-inflammatory drugs (NSAIDs), but data on their use via immersion is limited. Sci16 Laboratory Animal Science Professional July 2021

Frederic Chatigny, DVM, MSc, is the Clinical Veterinarian at Université de Sherbrooke, QC, CA. Julie Balko, VMD, DACVAA, is an Assistant Professor of Anesthesiology at North Carolina State University, College of Veterinary Medicine in Raleigh, NC. REFERENCES 1. American College of Veterinary Anesthesiologists (ACVAA). [Internet]. 2017. Position statement: the treatment of pain in animals. [Cited 2017 Jul 5]. Available at: http://www.acvaa.org/ docs/Pain_Treatment.

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2. Anil SS, Anil L, Deen J. [Internet]. 2002. Challenges of pain assessment in domestic animals. J Am Vet Med Assoc. 220(3):313–9. [Cited 2016 Jun 28]. Available at: http://www. ncbi.nlm.nih.gov/pubmed/11829261. 3. Braithwaite V. 2010. Do fish feel pain? Oxford (UK): Oxford University Press. 4. Carbone L, Austin J, Becker J, Bannon A, Calvo M, Caspani O. [Internet]. 2016. Pain and laboratory animals: publication practices for better data reproducibility and better animal welfare. Gao C-Q, editor. PLoS One. 11(5):e0155001. doi:10.1371/journal.pone.0155001. [Cited 2017 Jul 5]. Available at: http://dx.plos.org/10.1371/journal.pone.0155001. 5. Chatigny F. 2019. The Controversy on fish pain: a veterinarian’s perspective. J Appl Anim Welf Sci. 22(4). doi:10.1080/10 888705.2018.1530596. 6. Chatigny F, Creighton CM, Stevens ED. [Internet]. 2018. Updated review of fish analgesia. J Am Assoc Lab Anim Sci. 57(1): 5–12. [Cited 2018 Feb 13]. http://www.ncbi.nlm.nih. gov/pubmed/29402345. 7. Committee on Recognition and Alleviation of Pain in Laboratory Animals NRC. [Internet]. 2009. Recognition and Alleviation of Pain in Laboratory Animals. National Academies Press. [Cited 2017 Jul 5]. Available at: http://www.ncbi.nlm.nih.gov/ pubmed/20662126. 8. Deakin AG, Buckley J, AlZu’bi HS, Cossins AR, Spencer JW, Al’Nuaimy W, Young IS, Thomson JS, Sneddon LU. {internet]. 2019. Automated monitoring of behaviour in zebrafish after invasive procedures. Sci Rep. 9(1):9042. doi:10.1038/s41598019-45464-w. [Cited 2019 Jul 3]. Available at: http://www. ncbi.nlm.nih.gov/pubmed/31227751. 9. Jones RC. [Internet]. 2016. Fish sentience and the precautionary principle. Anim Sentience Jones Comment Key Fish

Pain. 016. [Cited 2016 Jul 19]. Available at: http://myweb. csuchico.edu/~rcjones/. 10. Kalueff A V, Gebhardt M, Stewart AM, Cachat JM, Brimmer M, Chawla JS, Craddock C, Kyzar EJ, Roth A, Landsman S, et al. [Internet]. 2013. Towards a comprehensive catalog of zebrafish behavior 1.0 and beyond. Zebrafish. 10(1):70–86. doi:10.1089/zeb.2012.0861. [Cited 2016 Dec 2]. Available at: http://www.ncbi.nlm.nih.gov/pubmed/23590400. 11. Rahman Khan F, Sulaiman Alhewairini S. [Internet]. 2019. Zebrafish (Danio rerio) as a model organism. In: Current Trends in Cancer Management. IntechOpen. [Cited 2021 Feb 16]. Available at: www.intechopen.com. 12. Raja SN, Carr DB, Cohen M, Finnerup NB, Flor H, Gibson S, Keefe FJ, Mogil JS, Ringkamp M, Sluka KA, et al. [Internet]. 2020. The revised International Association for the Study of Pain definition of pain: concepts, challenges, and compromises. Pain. 161(9): 1976–1982. doi:10.1097/j. pain.0000000000001939. [Cited 2021 Feb 16]. Available at: https://journals.lww.com/10.1097/j.pain.0000000000001939. 13. Rose JD, Arlinghaus R, Cooke SJ, Diggles BK, Sawynok W, Stevens ED, Wynne CDL. [Internet]. 2014. Can fish really feel pain? Fish Fish. 15(1): 97–133. doi:10.1111/faf.12010. [Cited 2016 Jun 30]. Available at: http://doi.wiley.com/10.1111/ faf.12010. 14. Santana S, Rico EP, Burgos JS. [Internet]. 2012. Can zebrafish be used as animal model to study Alzheimer’s disease? Am J Neurodegener Dis. 1(1): 32–48. [Cited 2016 Jul 11]. Available at: http://www.ncbi.nlm.nih.gov/pubmed/23383380. 15. Schofield Bvsc JC, Aclam D, Williams Bvsc VM. [Internet]. 2002. Analgesic best practice for the use of animals in research and teaching - an interpretative international literature review. [Cited 2017 Jul 5]. Available at: http://www.fao.org/ fileadmin/user_upload/animalwelfare/analgesic-practice.pdf.

Aquatics Featured in May JAALAS The May issue of JAALAS featured two articles on aquatics.

• Behavioral and Reproductive Effects of Environmental Enrichment and Pseudoloma neurophilia infection on Adult Zebrafish (Danio rerio) By Jenny M Estes, Michelle L Altemara, Marcus J Crim, Craig A Fletcher, and Julia W Whitaker

• Comparison of Juvenile Feed Protocols on Growth and Spawning in Zebrafish By Stephen C Frederickson, Mark D Steinmiller, Tiffany Rae Blaylock, Mike E Wisnieski II, James D Malley, Lauren M Pandolfo, and Daniel Castranova The May issue of JAALAS is available to AALAS Silver and Gold members on the website. Please log in (https://www.aalas.org/) and visit our Publications tab to view the online content. While COVID-19 may have impacted the delivery of physical copies of the AALAS journals and magazine, members may always access digital editions through the AALAS website. Members must be logged in to the website to access materials. Journal articles will become public 6 months after publication. AALAS membership is required to view all publications.

18 Laboratory Animal Science Professional July 2021

72nd AALAS NATIONAL MEETING KANSAS CITY, MO OCTOBER 17 - 21, 2021

AALAS Heads to Kansas City! Each fall since 1950, the American Association for Laboratory Animal Science has held its annual National Meeting. Attendees come together to enjoy the workshops, lectures, poster sessions, and exhibits. Exhibitors have an opportunity to interact with AALAS members from the academic community, research institutions, government organizations, and commercial companies. The AALAS National Meeting is the largest gathering in the world of professionals concerned with the production, care, and use of laboratory animals. Visit aalas.org/national-meeting for details!

July 2021 Laboratory Animal Science Professional 19

FEATURE

Future Focus

Looking into an AALAS Crystal Ball

AALAS leadership share thoughts on the future of our organization, lab animal science, and industry trends. By Liz Rozanski, BA

OVID-19 delivered stunning blows to personal and professional plans. Illness, canceled trips, work-fromhome routines, virtual schools, and supply shortages resulted in plans and expectations for 2020 being tossed aside. Innovation with a hefty dose of resiliency became the status quo. Now, over 18 months later, we are moving forward. AALAS is planning a live meeting in Kansas City, MO, with a virtual component for attendees unable to travel. Beyond Kansas City, our leadership prepares for the association’s and LAS’ future. AALAS Executive Committee members, and association Executive Director, Ann Turner shared thoughts on what the future might hold. They were asked to dust off their crystal balls, to look and imagine where we are headed.

Future Forecasting

If you had to guess what laboratory animal science would look like in 10 or 20 years, what would your answer be? Executive committee members identified two immediate impacts to the future of LAS: technology and generational change. AALAS President, Doug Taylor, noted that generational blends in the “office” would deliver changes to not just what we do but how we do it. “We're in an era where people spanning 4-5 generations are working together. This is unprecedented in history, and I would expect it to continue. This means that the very young have an opportunity to influence the thoughts and directions taken by senior leadership, which could be impactful, whether simply in changing outdated practices or embracing new, untested technology,” Taylor explained. Emphasizing the possible technological advances, AALAS Doug Taylor, AALAS President Vice President, Marc Hulin, noted changes driven by significant tech investments. “I envision a significant investment in technologies allowing us to monitor animal activity remotely without having to engage the animal by entering 20 Laboratory Animal Science Professional July 2021

the room to assess care and welfare. Looking 20 plus years out, I see the true adoption of complex in vitro models to replace the use of animals in certain arenas for biomedical research,” Hulin said. Tim Mandrell, AALAS’ Secretary/Treasurer, noted a move towards fewer animals, increased automation, and an emphasis on certification. Tim Mandrell, AALAS Secretary/Treasurer “Technology will allow for more powerful data from each animal, and fewer animals will be used. Also, advancements in caging will make all cages "smart cages," and AALAS certified technicians will know how to set up, maintain the equipment and download data, and perform daily care. There will be continued improvement in technology such that automation in cage processing will become more prevalent, even in smaller programs,” Mandrell said. He noted increased technology and training requirements would necessitate that most research technicians will be required to have AALAS certifications. Vice President-Elect, Pam Straeter, agreed that animal numbers would decrease. “Technology will be further refined and advanced such that the numbers of animals used in research will decline significantly,” Straeter said. She added that virtual education and conferences would continue to expand. She said virtual meetings have become so commonplace that in-office check-ins or gatherings are now held online at her facility. AALAS Executive Director, Ann Turner, agreed with Straeter’s assessment. “AALAS will explore the expansion of our training and educational program to meet the needs of the emerging generation of laboratory animal science professionals,” Turner said.

COVID Spurs Planning Changes

Over the past 18 months, we have adjusted to COVID and the impacts on our professional and personal lives. When it comes to long-range planning, how the pandemic influenced and will continue to influence operations is still being examined and considered.

“Hopefully, we are coming out of the COVID pandemic, one of the most treacherous times in modern history. While many of us want to get back to normal, we must realize that ‘normal’ has changed rapidly and will continue to do so,” Turner explained. Straeter agreed that normal has changed and anticipates long-term effects. “We do long-range planning to a certain extent, covering animal enrichment, equipment, and facility advancements that enhance animal welfare and staff ergonomics. Our two to fiveyear plans were put temporarily on hold, and although we are not fully out of the pandemic, we are beginning to pick up on those paused plans,” Straeter said. Mandrell shared that typically he has planned for one-year and five-year goals. He notes, however, that in academia, it is more challenging to predict changes in academic missions. “COVID has been a wake-up call for preparedness and continuity of business plans. We think about disaster plans, which continue to be important, but pandemic/workforce issues are also important and require some different approaches that many of us learned on the fly this year,” he explained. From a staffing perspective, COVID response has created a more fluid workforce and has often seen employees working less than full time, Mandrell noted. “That is the downside. The upside, if there is one, is the necessity/opportunity to have more employees cross-trained. On a given day, there may be staffing shortages in animal husbandry, cage processing, or technical services, and a welltrained/cross-trained workforce can meet those needs,” he said. In addition to cross-trained workers, Taylor noted the broader acceptance of remote work. “We are otherwise back to business as usual as most of my workforce has been vaccinated. We are pretty well versed in disaster planning and in this field, and my observations are that we have weathered the COVID storm adeptly, all things considered,” Taylor said.

A Magic Wand

In addition to asking AALAS leadership to look into a crystal ball, we gifted them with magic wands to make systemic changes in laboratory animal science. Marc Hulin noted he would use this gift to foster diversity and inclusion in laboratory animal science now and forever. "Although widely recognized in the workplace that diversity of thought and innovation are instrumental to success, unconscious preferences for people who are like us continues to severely challenge our ability to create

Marc Hulin, AALAS Vice President

diversity and inclusion. An area we need to educate ourselves on is the impact of unconscious bias in the workplace. We have a tremendous opportunity to begin to "move the needle" on changing social injustice in the US by educating ourselves and individuals in our workplace on unconscious bias/implicit bias," Hulin explained. Doug Taylor focused on transparency. Noting the relative lack of transparency in the field, he would create a way that all research programs maintaining animals could fully disclose in a meaningful and public way precisely what they do and how they do it. Being president, Taylor was granted a second wand wave to bring about international standards for animal colony health monitoring, and in the case of rodents, without the use of sentinel animals. “Big picture stuff,” Pam Straeter said. “I’d help LAS staff to recognize the importance of external involvement through volunteerism and to embrace the enhanced availability of technology. I’d also encourage people to see beyond the day-to-day work.” Tim Mandrell wants to see smart caging for rodents, industry standards for enrichment, and more integration of AALAS certified technicians into research laboratories. “It should be a requirement Pam Straeter, AALAS Vice President-Elect that anyone who works with laboratory animals is certified. Advanced certification would be required for technicians to perform more technical procedures,” Mandrell added.

A Final Glance to the Future

As we live through the pandemic and analyze its current and projected impacts on laboratory animal science, measuring how AALAS, as an association, will experience this pivot to a new normal is not easy. Relevancy, resiliency, and flexibility weave through the Executive Committee’s answers. “We've had some discussions about remaining relevant in the face of generational changes, and this issue will continue to impact AALAS. Membership is dependent upon the value a potential member assigns to the association,” Taylor said. He added that post-COVID, the nature of national meetings and conferences, in general, are probably changed forever. Turner explained that conferences and conventions have been central to associations since the first associations in ancient Greece; however, the nature of those gatherings have evolved over time. “The pandemic put the changing nature of mass gatherings in warp speed. People love and need to connect. Many prefer that connection to be in person and some prefer virtual connection,” she said. July 2021 Laboratory Animal Science Professional 21

She noted that regardless of preference, associations will continue to provide opportunities for people to connect, exchange information, and plan for the advancement of society through meetings. “While the AALAS National Meeting in 2021 and beyond may look different from the one in Denver in 2019 and the virtual one in 2020, the AALAS family will gather and celebrate who we are and what we do!” Turner added. Hulin and Straeter focused on remaining flexible and innovative. “The last year has forced our industry to continue to be flexible and adaptable to any circumstance,” Hulin noted. “I believe this pandemic has taught us innovative ways to accomplish the critical work needed in research, and anywhere really. I hope we recall the great benefit and professional growth gained from being a volunteer, gathering in person, and sharing experiences,” Straeter said. Professional growth was also on Tim Mandrell’s mind. He hopes that AALAS’ role in certifying technicians, providing high-quality training materials, and promoting responsible

care will be elevated. “As we move forward, I hope major research organizations and institutions will not only recognize the importance of AALAS certification but endorse AALAS certification and training as the best option for all researchers and research assistants/technicians,” Mandrell said.

Last Word

With half of 2021 in the rear-view mirror and the National Meeting only months away, Ann Turner sees reasons for hope for laboratory animal science and AALAS’ continued growth. “Biomedical research, science, and vaccines have taken center stage during the entire pandemic. Science does matter, and now more than ever, that message resonates with large portions of our global society. AALAS is well-positioned to be the leader in educating our front-line workers: laboratory animal science professionals,” Turner said. Liz Rozanski, BA, is Communications Manager at AALAS in Memphis, TN.

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FEATURE

Future Focus

Nonhuman Primate Shortage: Exploring Solutions By Matthew R. Bailey

he United States faces a potential shortage of nonhuman primates (NHPs) for biomedical research.2,6 A rise in demand and reduction in availability have exacerbated the situation from the onset of the coronavirus pandemic. Public/ private synergies to expand domestic colonies could provide a possible solution to the shortage. U.S. breeding and conservation assistance programs with other countries may also provide new avenues for U.S. medical research to obtain internationally sourced nonhuman primates. The race to develop coronavirus vaccines and the resumption of research activities not related to COVID-19 together have led to an increased demand for NHPs. Approximately 68,000 monkeys were engaged in research programs in 2019.9 According to studies conducted by the NIH, future demand could exceed this number. Yet currently the U.S. is experiencing a reduction in the availability of NHPs for research.

though this partnership.4 By establishing similar collaborations with Southeast Asian countries that breed NHPs, the U.S. could establish new avenues to import Old World monkeys like rhesus macaques to the U.S. The U.S. research community should also explore the possibility of public/private research sector partnerships to establish a strategic resource for domestically bred monkeys. A new strategic resource could potentially complement the critical work being conducted at the federally funded National Primate Research Centers. Public and private resources are desirable to cover the costs of establishing, maintaining, and expanding a strategic primate resource.

Conclusion

Most of the world is emerging from the coronavirus pandemic with new vaccines developed thanks in large part to NHPs

China Bans NHP Exports

At the onset of the pandemic, the Chinese government issued a temporary ban on the sale and transport of wild animals, then made the ban permanent.8 China was a major supplier of NHPs for the rest of the world. Roughly 60% of the estimated 30,000 NHPs imported into the United States came from China until the country implemented its embargo on exportation of NHPs.7 It is unclear when or if the Chinese government will resume exportation of NHPs. Vietnam, Cambodia, and Mauritius continue to export research monkeys to the U.S.7 They supply approximately one third of the NHPs the U.S. obtained from foreign sources.7 However, it appears unlikely these countries can make up for the shortage stemming from China’s embargo. India ceased exportation of NHPs to the U.S. in 1977 after the Indian press reported they were being used in military research.5 Several other Southeast Asian countries including the Philippines stopped exporting nonhuman primates to the U.S. when some monkeys were found to be contaminated with a strain of Ebola.3,1 It is therefore incumbent upon the U.S. research community to find new sources of research nonhuman primates to avoid the short- and long- term effects of a shortage on America’s biomedical research sector.

New Partnerships Emerge

The partnership between the U.S. and the Pan American Health Organization offers one example the U.S. could replicate with other countries. The National Institutes of Health (NIH) and the Peruvian Primatology Project collaborate on efforts to conserve Peru’s wild monkey population. They also work together to improve Peruvian monkey breeding facilities. The NIH has imported New World monkeys from Peru 24 Laboratory Animal Science Professional July 2021

The race to develop coronavirus vaccines and the resumption of research activities not related to COVID-19 together have led to an increased demand for NHPs.

Researchers are already building on the coronavirus vaccine technologies to develop new and promising vaccines and therapies for diseases that range from cancer to HIV/AIDS, and research with NHPs will continue to be an essential part of the research and development process.

and other animal models. Researchers are already building on the coronavirus vaccine technologies to develop new and promising vaccines and therapies for diseases that range from cancer to HIV/AIDS, and research with NHPs will continue to be an essential part of the research and development process. COVID-19 was not the first pandemic our nation has successfully managed, and it certainly will not be the last. For that reason, the U.S. must proactively ensure it has adequate resources for medical research, including appropriate laboratory animals. Our future health, safety and global competitiveness depend on it. Matthew R. Bailey is the President of the National Association for Biomedical Research in Washington, DC. REFERENCES 1. Al Jazerra. [Internet]. 2015. Philippines bans monkey exports over Ebola deaths. [Cited 18 May 2021]. Available at: https://www.aljazeera.com/news/2015/9/10/philippines-bans-monkey-exports-over-ebola-deaths 2. Biodiversity Heritage Library. [Internet]. 1978. Interagency primate steering committee national primate plan. [Cited 18 May 2021]. Available at: https://www.biodiversitylibrary.org/ item/247593#page/3/mode/1up 3. Centers for Disease Control and Prevention. [Internet]. 2015. Surveillance for ebola virus in wildlife, thailand, Emerging infection diseases journal Volume 21, Number 12—December 2015. [Cited 18 May 2021]. Available at: https://wwwnc. cdc.gov/eid/article/21/12/15-0860_article

4. Held JR. 1981. Breeding and use of nonhuman primates in the USA. Int J Study Anim Prob 2(1): 27-37. 5. LeCornu A, Rowan AN. [Internet]. 1978.Trends in the use of nonhuman primates in biomedical programs. Lab Anim 12: 235-242. [Cited 18 May 2021]. Available at: https://journals. sagepub.com/doi/pdf/10.1258/002367778781088576 6. National Institutes of Health. [Internet]. 2018. National institutes of health nonhuman primate evaluation and analysis parts 1 and 2, 2018, [Cited 18 May 2021]. Available at: https://orip.nih.gov/sites/default/files/508%20NHP%20Evaluation%20and%20Analysis%20Final%20Report%20-%20 Part%201%20Update%2030Oct2018_508.pdf and https:// orip.nih.gov/sites/default/files/NHP%20Evaluation%20and%20 Analysis%20Final%20%20Report%20-%20Part%202%20 Final%20508%2021Dec2018_002.pdf 7. National Research Council (US) Institute for Laboratory Animal Research. [Internet]. 2011. Animal research in a global environment: meeting the challenges: proceedings of the november 2008 international workshop. Washington (DC): National Academies Press. [Cited 18 May 2021]. Available at: https://www.ncbi.nlm.nih.gov/books/NBK91512/ 8. Research Professional News. [Internet]. 2021. Animal research advocates seek urgent action on china export ban. [Cited 18 May 2021]. Available at: https://www.researchprofessionalnews.com/rr-news-europe-regulation-2021-2-animal-researchadvocates-seek-urgent-action-on-china-export-ban/ 9. United States Department of Agriculture. [Internet]. 2019. 2019 Annual report animal usage by fiscal year. [Cited 18 May 2021]. Available at: https://www.aphis.usda.gov/animal_welfare/ annual-reports/2019/fy19-summary-report-column-F.pdf

July 2021 Laboratory Animal Science Professional 25

FEATURE

Future Focus

The Psychologically Safe Workplace By Jarrod Nichol, MBA, PMP, CLSSBB, Prosci

he title of this article may perplex you, but the concept is gaining traction in our industry and you are going to hear about in the years ahead. Ponder this, when you interview a Veterinary Technician, what are you looking for in the potential new hire? Are items like punctuality, work ethic, attention to detail, and previous experience key factors to measure a successful employee? Do you focus the bulk of your time on those points? If so, you may be missing a new approach in the biomedical research industry. Instead of assessing traditional factors, look at an employee’s ability to adapt, their drive to learn new techniques and approaches, and above all, to speak up when they have something to say. Robots in Research

Years ago, cage washes all over the world were retrofitted with robotics. Do you remember why we did this? The most common answer was efficiency and ergonomics. As advances in robotics continue, one of the upcoming trends will be to slowly replace staff with robots or equipment that will alert the technicians which cages need to be changed or which animals need to be examined. There are already fully automated research facilities in Asia, where the cages are changed entirely by robots. Veterinary staff are present to oversee animals that millions of sensors monitor continuously. This is becoming more commonplace and if our workforce does not adapt, it will be overtaken by robotic replacements. After visiting research facilities across North America, several in Europe, and even a handful in Asia, I can say robotic replacement is happening sooner than we realize. Most organizations continue to focus on traditional qualities such as a staff that is punctual, follows instruction, and does not cause disruptions. Instead, we should be looking for employees who are challenging the status quo, willing to report the root causes of errors, and look beyond the confines of their job description. Employee characteristics such as these will draw a distinct difference between automation and human abilities. The global COVID-19 pandemic may have radically changed most industries, but an essential service industry like laboratory animal sciences has not seen the same level of evolution. Our staff must come in, regardless of the weather, the political climate, or in this case, a worldwide pandemic. Why change if we don’t have to?

A Lean Lesson in Change

In 2004 Dr. Donna Matthews Jarrell, Director of the Center for Comparative Medicine at Massachusetts General Hospital, asked that same question.1 Why change? She realized that 26 Laboratory Animal Science Professional July 2021

without change, the problems the center was facing would continue in perpetuity. What was her solution? She introduced the Toyota Production System (TPS) to her workforce and never looked back. The Toyota Production System is a process improvement and management system that organizes manufacturing, logistics, as well as a client-focused approach to vastly improve manufacturing quality and output. The methodology was developed by Toyota. While the system focuses on operational efficiency, at its heart, it is about people and empowering them to impact their day-to-day work. A decade ago, this new trend would set out to reshape our industry, and now it is being fully implemented. The Vivarium Operational Excellent Network (www.voenetwork.com) is a testament to Dr. Jarrell’s vision. The network features worldwide organizations that have come together to share best practices in the field of continuous improvement in the biomedical research industry. Dr. Jarrell was right, challenging the status quo and changing was the right thing to do. As we continue to tighten operational efficiency with robotics, another question is popping up, what is next? If we have already seen an evolution regarding operations and empowering of staff, how do we move forward to save workforces from a robotic revolution?

Psychological Safety

The answer lies in a trend gaining traction in many other industries, the concept of psychological safety. I first heard of psychological safety from Harvard University behavioral scientist, Amy Edmondson, who wrote The Fearless Organization: Creating Psychological Safety in the Workplace for Learning, Innovation, and Growth, in 2019. She describes psychological safety as a belief that employees should not be afraid to speak their mind and are encouraged to do so without the fear of repercussion. This concept may sound simple but developing

a workplace culture conducive to such behavior is exceedingly difficult. TPS or another Continuous improvement program are a great first step. They open the door to empowerment, giving people a voice, and a chance to directly impact their work processes. If you already have such a program, then it is time

Psychological safety is a belief that employees should not be afraid to speak their mind and are encouraged to do so without the fear of repercussion. This concept may sound simple but developing a workplace culture conducive to such behavior is exceedingly difficult. to take the next step. Start by rewarding people for bringing issues to light and for challenging actions they believe to be incorrect. How you might ask, by simply recognizing these people at town halls, in your corporate newsletter, or just having the Director come forward and personally thank the individual or team. This is not about being one of those employees. It is about feeling safe enough to ask, “Why are we doing this?” and “Did you forget to…,” etc. By creating psychologically safe environments, problems are addressed far quicker than ever before because people do not stay silent, so you are made aware of issues often before they become huge problems that end up costing a lot of money, or issues to the research being done. Additionally, researchers will be impressed by the dramatic increase in customer service and

the quality of animal care and welfare will be directly affected by the new work culture. Under a psychologically safe culture, people will reveal their full potential and readily become an asset to their organization beyond their defined role. When people feel safe to respectfully challenge work processes, ideas, or even the evolution of a unit, they also look for ways to be more involved beyond their positions. When you mix psychological safety with the advances in robotics, you suddenly start to develop wholly unique and original positions. An animal health technician can easily become an animal welfare and robotics expert or a human interaction specialist. The options are as endless and the reward to the company is great. What could have become an industry of automatons could instead be filled with dramatically unique roles and a bright future for everyone involved in the industry. However, only by making that first step into changing the way we select and manage people will we be able to establish the next big trend in the industry, the psychologically safe workplace. Jarrod Nichol, MBA, PMP, CLSSBB, Prosci, is the Associate Director, Operations and Administration in the Comparative Medicine and Animal Resources Centre at McGill University in Montreal, Quebec, Canada. REFERENCES 1. Jarrell DM. [Internet]. 2017. Donna Matthews Jarrell, DVM, DACLAM Bio. [Cited 11 May 2021]. Available at: https://www.eventscribe.com/2017/primr-iacuc/ajaxcalls/PresenterInfo.asp?efp=TkVJQkJLTlozOTYy&PresenterID=267630&rnd=0.8298016 July 2021 Laboratory Animal Science Professional 27

FEATURE

Future Focus

Innovation with 3D Printing

By Kim Froeschl, MLAS, CMAR, CPTD; Michaela Workman, BS, RALAT; and Melissa Dragon, MBA, CMAR

s laboratory animal science professionals, we are often presented with unique challenges in obtaining the precise equipment needed to support animal studies due to the limited availability of commercially manufactured equipment such as restrainers, blood tube holders, and anesthesia equipment. Recently, the use of innovative technology, specifically 3D printers, has become an efficient, economical, and practical approach to meeting these needs. Each of the four Pfizer Comparative Medicine (CM) sites (Groton, CT; Pearl River, NY; Cambridge, MA; La Jolla, CA) has unique study needs, and we are often challenged to find solutions that will improve animal welfare, ergonomics, data collection, and/ or safety. When a specific need cannot be met with a product from our commercial partners, 3D printer technology has been helping us to innovate and create those needed solutions. We’re fortunate to have a scientist with a 3D printer (Form2, Makerbot Replicator: https://www.makerbot.com/3d-printers/ replicator/) in his lab and CAD software ((Autodesk Fusion 360 https://www.autodesk.com/) to create prototypes. He helped us develop several products that fit our specific needs, but his time was limited. We enlisted the help of a contract CAD Designer for a few projects, but this was time-consuming and expensive. Therefore, we focused on expanding our

Figure 1. Form3 Printer; Courtesy of Form Labs.

internal resources. As project ideas and solutions grew, it was evident that having a printer and CAD software in our department would be beneficial. After researching options, we decided on a resin-based 3D printer (Form3 by Form Labs; https://formlabs.com/3d-printers/form-3/), (Figure 1) due to ease of use, reliability, and affordability. Like other 3D printer technologies, resin-based printers print objects layer-by-layer but use a pool of resin stored in a cartridge below the print base rather than an extrusion filament that deposits layers to the top of the printed object. Its highly advanced light processing unit (LPU) ensures precise, reliable prints each time. With various resins available, it allows the printing of extremely realistic objects; for example, some resins are tough and rigid, while others are flexible and elastic. This allows creating a wide variety of prints, from autoclavable objects to flexible, realistic animal models. The printer does have a size limitation of 5.7×5.7×7.3 inches, but this has not yet been a limitation in our designs. One of our team members also taught herself the CAD program (Autodesk Fusion 360) to create the designs proposed. The printing process is a relatively straightforward 3 step process of design, printing, and washing/curing the product. Print time varies depending on the size and density of the object, but it is approximately a 7-hour process which can run overnight.

3D Print Projects Figure 2. Caliper measuring printed tumor.

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Mouse Tumor Model Training new staff to perform tumor measurements accurately and consistently on mice is challenging due to the variation

Figure 3. CAD drawing of a tumor.

Figure 4. Mice with tumor prototype.

in tumor shape, firmness, and caliper pressure applied. By creating soft, flexible 3D printed tumor models (Figure 2-3), we can improve consistency in measurement and eliminate the use of live animals for this training. The printed tumor models can be inserted into inanimate mice to better replicate the actual measuring of tumors. Additionally, a prototype mouse with a tumor was printed from existing full-body 3D micro-computed tomography (mCT) scans of an anesthetized mouse with a subcutaneous tumor. Using additional software, we converted the mCT files to a printable file (Figure 4). The next step will be to print the model with a flexible or elastic resin to make the mouse look and feel as real as possible.

Rodent Anesthesia Nose Cone Clamp

Anesthesia Nose Cone for Rat CSF Collection Standard rat nose cones for inhaled anesthesia did not allow for correct positioning of the head to collect cerebrospinal fluid (CSF) from the cisterna magna of rats. By 3D printing nose cone inserts onsite, we were able to quickly create a few prototypes (Figure 5) to trial and determine the optimum design and size needed. The new nose cone can be positioned flat against the rat restraint platform. This prevents the rat’s head from popping out of the nose cone during manipulations, ensuring continuity of anesthesia and improved animal welfare (Figure 6). Single Tube Holder for Submandibular Blood Collection Commercially available tube racks are designed to hold dozens of vials but don’t hold tubes well for rodent facial/submandibular vein blood collection. Commercial racks hold tubes low on the benchtop and have deep wells, which prohibit good visualization of the blood volume during collection. We now create multiple tube holder types to meet user-specific needs and preferences (Figure 7). Mouse Restrainer for Intravenous (IV) Dosing Custom-made cone-shaped mouse restrainers for IV dosing were expensive ($400 each) and did not hold up to cage washing. By designing our restrainer, we were able to create the exact dimensions needed, chose a material (nylon) that would hold up to cage washing, and print new restrainers as needed (Figure 8).

Figure 5. Standard nose cone, Prototype 1, final design.

Figure 6. Rat anesthetized using printed nose cone.

Technicians performing rodent procedures using gas anesthesia were challenged with keeping the nose cone in place for the duration of the technique. A clamp was designed that would connect to the work surface and hold the nose cone in place. The nose cones are now held securely in place regardless of the duration of the procedure (Figure 9). July 2021 Laboratory Animal Science Professional 29

Figure 7. Three styles of blood tube holders.

Figure 8. IV mouse restrainer; custom-made (clear) and 3D printed (white).

Figure 9. Clamp for rodent anesthesia nose cone.

Conclusion

All procedures involving animals were in accordance with regulations and established guidelines and were reviewed and approved by Pfizer’s Institutional Animal Care and Use Committee.

Drawing on the innovative ideas of the team and available internal resources, we were able to resolve several unmet needs with innovative brainstorming and 3D print technology. By bringing together several individuals with different backgrounds, we quickly put these projects in motion, improved animal welfare, efficiency, and effectiveness for our CM staff and scientists, and met these challenges extremely economically. Acknowledgments: Pfizer colleagues: Tim Winton, Mary Kate Montgomery, and Corey Dasilva.

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Kim Froeschl, MLAS, CMAR, CPTD, is a Comparative Medicine Global Trainer at Pfizer Inc in Pearl River, NY. Michaela Workman, BS, RALAT, is a Comparative Medicine Research Support Coordinator at Pfizer Inc in La Jolla, CA. Melissa Dragon, MBA, CMAR, is a Comparative Medicine Global Trainer at Pfizer Inc in Groton, CT.

FEATURE

Future Focus

A 3-D Printed Apparatus for Imaging Multiple Rats Simultaneously

By Nicholas J. Harrison, DVM, MS; Kate L. Shumway, DVM, DACVR; Sarah A. Hansen, DVM, MS, DACLAM; Charles A. Maitz, DVM, PhD, DACVR; Lori A. Thombs, MS, PhD; and Brian K. Flesner, DVM, MS, DACVIM

reclinical studies routinely require the use of advanced diagnostic imaging techniques to evaluate animal models of human disease.2 Rodent models are commonly used to gather data prior to human clinical trials, but computerized tomography (CT) in rodents can be difficult due to their small size and constant movement necessitating general anesthesia. Improving the convenience and cost of imaging rats has the potential to advance preclinical study outcomes by facilitating longitudinal translational studies. Although microCT (computerized tomography) has been developed and used for high anatomic resolution of small animals, the equipment may be unavailable for use or difficult to access.1 In addition, the high resolution (typically 50 to 100 um3 voxel spacing) afforded by this equipment may not be necessary for a given experiment. Limitations for either micro-CT or clinical CT include anesthesia to limit image distortion due to motion, biosecurity, and cost. To address several of these constraints, we created a 3D printed apparatus, allowing simultaneous imaging of up to nine rats under gas anesthesia. Rats were anesthetized in series and stacked in a 3×3 fashion. A full article outlining the apparatus in use was published in the April 2021 issue of Comparative Medicine.3

Materials and Methods Creating the Apparatus

An 8-mo-old, intact male rat was measured for the development of the imaging bed. The tip of the nose to the tail base measured 21 cm, the tail itself was 16 cm long, and the height of the rat when prone was 3.75 cm. For repeat cylindrical printing, a commercially available water

Figure 1 A-F. Components of the apparatus. (A) Plastic bottle restraint device. (B) Extension sets with 4-way stopcocks and extension sets in series. (C) Completion of anesthetic circuit. (D) Three rats anesthetized on the first row of the apparatus. (E) Completion of imaging of top row of anesthetized rats, with one rat anesthetized in the top middle position. (F) Nine rats under inhalant anesthesia and imaged on a clinical CT machine.

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bottle (diameter, 6.5 cm; length, 24.5 cm) made of PVC plastic was determined to be the appropriate size and material. The label and adhesive material were removed with a commercially available product containing terpene hydrocarbons and C9–11 ethoxylated alcohols (Goo Gone, Weiman Products, Gurnee, IL). The base of the water bottle was sectioned Read more about this apparatus in and the edges sanded the April 2021 issue of Comparative to prevent injury to Medicine (Vol 71, No 2, Pages 116122) animals, to a final length of 24 cm. A 20-mL syringe (Terumo, Tokyo, Japan) was transected at the 5-mL mark and inserted into the top of the bottle. To stabilize its attachment to the bottle, the syringe was secured in place by using a bead of glue (Gorilla Glue, Sharonville, OH) at the junction of the syringe and the lip of the bottle, thus completing the restraint device (Figure 1 A). We made 9 individual restraint devices. To allow for flow-by anesthesia, a series of 24-in fluid extension sets (Baxter Healthcare, Deerfield, IL) were connected by using 4-way stopcocks and luer connections (Cole Parmer, Vernon Hills, IL), allowing simultaneous delivery of isoflurane to all 9 restraint devices. The nine 4-way stop-cocks were connected linearly, with a fluid extension set connected to each stopcock. This arrangement allowed attachment to each restraint device. An extension set was connected at each end of the line of stopcocks to allow connection to the anesthetic machine (Figure 1 B). A 5-mm uncuffed endotracheal tube was attached to a 4-way stopcock. The plastic connector of the endotracheal tube was removed, and approximately 1 cm of the tip of the tube was transected to make the end flat. The fluid extension set from each end of the line of stopcocks then was connected to the individual stopcock, completing the circuit, and allowing for flow from both ends of the chain of stopcocks (Figure 1 C). The endotracheal tube was attached to the circuit of the anesthetic machine, allowing flow-by delivery of isoflurane to each rat during the imaging procedure. A scaffold was created to hold modified water bottles in place during imaging. Modified water bottles were recreated by using an Autodesk Fusion 360 (San Rafael, CA). A 3D representation of the bottle was made, and 3 scaffolds were 3D-printed additively by using polylactic acid (0.3 mm layers, 10% infill, and 3 shells) on a Makerbot Z18 (MakerBot Industries, New York, NY). These devices were stacked, which allowed the anesthesia and scanning of as many as 9 animals simultaneously (Figure 1 D through F).

Measuring Attenuation in Each Chamber

To determine effect on attenuation between individual chambers and rows/columns in the device, a standardized phantom plug (0.99 Hounsfield units [HU]) was imaged in each chamber. Under our IACUC-approved protocol nine rats were then anesthetized and placed in the apparatus and four organs were evaluated: liver, kidney, femur, and brain.

Results

All animals were maintained at an appropriate anesthetic plane in the apparatus with isoflurane anesthesia (1-5%) for the duration of experiment. A statistically significant, but clinically negligible, effect on attenuation was noted between rows, but not columns in the kidneys (p=0.04) and the phantom (p<0.0001). We attribute this finding to the absence of a top layer of the apparatus, which thus created asymmetric attenuation and beam hardening through the device. This apparatus allowed us to successfully and simultaneously image nine rats in a clinical CT machine, with clinically negligible effects on attenuation. Future iterations of this apparatus include versions with complete enclosure for biosecure imaging. Nicholas J. Harrison, DVM, MS, is a Clinical Veterinarian at Charles River Laboratories in Ashland, OH.

Kate L. Shumway, DVM, DACVR, is a Radiologist at VetCT in Columbia, MO. Sarah A. Hansen, DVM, MS, DACLAM, is with IDEXX BioAnalytics in Columbia, MO. Charles A. Maitz, DVM, PhD, DACVR (Radiation Oncology) is Assistant Professor in the Department of Veterinary Medicine and Surgery, College of Veterinary Medicine at the University of Missouri in Columbia, MO.

Lori A. Thombs, MS, PhD, is an Associate Professor in the Department of Statistics, College of Arts and Science at the University of Missouri, Columbia, MO. Brian K. Flesner, DVM, MS, DACVIM (Oncology), is an Assistant Professor in the Department of Veterinary Medicine and Surgery, College of Veterinary Medicine at the University of Missouri in Columbia, MO. REFERENCES 1. Paulus MJ, Gleason SS, Kennel SJ, Hunsicker PR, Johnson DK. 2000. High resolution X-ray computed tomography: an emerging tool for small animal cancer research. Neoplasia 2:62-70. 2. Puaux AL, Ong LC, Jin Y, Teh I, Hong M, Chow PK, Golay X, Abastado JP. 2011. A comparison of imaging techniques to monitor tumor growth and cancer progression in living animals. Int J Mol Imaging 2011:321538. 3. Harrison NJ, Shumway KL, Hansen SA, Maitz CA, Thombs LA, Flesner BK. 2021. A 3D-Printed Apparatus for Imaging Multiple Rats Simultaneously. Comparative Medicine 71(2):116-22.

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FEATURE

Future Focus

Don’t WAG Your Finger at 3D Printing: Proper F/air Canister Position with a 3D Printed Holder By Jennifer L Booth, DVM, MS, DACLAM

aste anesthetic gas (WAG) is defined as any anesthetic gas that leaks into the surrounding room. Exposure to WAG is presumed to be an occupational health hazard for personnel who operate or work near anesthesia units. Research in humans and animals has shown that exposure to these gases may result in many deleterious effects. Short-term, acute exposures can lead to complaints of nausea, dizziness, headaches, and fatigue, while more long-term, chronic exposures potentially result in the development of cancer, reduced fertility, birth defects, spontaneous abortions, and liver and kidney disease.1 To help mitigate the risk of personnel exposure, it is imperative to understand the potential ways that anesthesia gas can leak into the environment so that best practices can be implemented to prevent or minimize this. (Figure 1) When considering common sources for leaks from the system, the proper use of the scavenger can sometimes get overlooked. But it really shouldn’t because they serve as a crucial component in reducing WAG exposure. As oxygen

flows through the anesthesia unit, anesthetic gas in the tubing will pass through the scavenger, containing activated charcoal, which adsorbs the anesthetic from the air that is being discharged into the environment. Circumstances that can diminish the scavenger’s ability to adsorb anesthetic gas include situations in which the charcoal has reached the maximum adsorbency, oxygen flow rates are too high, or the scavenger is positioned poorly. Scavengers have an adsorption limit that can be tracked by weight. Many of these scavengers come with directions printed on the canister, instructing the user to discard it once the recommended weight has been reached. Operating the anesthesia unit at high flow rates will not only cause your charcoal to reach the adsorbency limit quicker, but the high flow could allow some of the gas to bypass the charcoal, leading to WAG production. The position of the canister is important because the overall design of the canister and the location of the aeration holes will determine the optimal position for adsorption efficiency. The proper position of these canisters is not always intuitive,

Figure 1. Potential sources of waste anesthetic gas. Anesthetic gas may be released into the environment from processes involving an anesthetized patient, handling of the anesthesia in liquid form, and directly from the anesthesia system.

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to elevate the canister in the upright position. Commercial holders for these canisters are readily available for purchase, typically averaging about $15 per holder. Instead of purchasing a commercial holder, I believed that I could design a holder that could be 3D printed. By utilizing the resources available at Penn State College of Medicine, I developed a 3D printed holder, at a fraction of the cost, only $0.15 per holder! (Figure 2 C-D)

Materials and Methods

To develop the 3D design, measurements of the outer diameter of the F/air canister were taken to ensure that the holder was an appropriate size. Tinkercad.com was then used to design a holder that would elevate the canister in the upright position, allowing the gas to enter the canister and discharge through the aeration holes without obstruction. The design file is free to the public under Creative Commons licensing and accessible at https://www.tinkercad.com/things/dfjyl4v6Nwa. Once the design was completed, the holder was then printed with a 3mm polylactic acid plastic filament, commonly referred to as PLA, on a LulzBot Taz Mini printer. This plastic is durable enough to withstand hand-cleaning with detergents and may further be sanitized by spraying with a disinfectant.

Conclusion

Figure 2 A-D. Use of a 3D printed holder to elevate the F/air canister in the upright position. A) The F/air canister without the holder sits flat against the table top when in the upright position. B) The aeration holes of the F/air canister are on the bottom of the canister. C) The F/air canister has been placed into a 3D printed holder which elevates it off of the table. D) The holder prevents occlusion of the aeration holes on the bottom of the canister.

as they often do not come with directions that offer guidance about positioning. A commonly used scavenger at our institution is the F/air canister (AM Bickford, Wales Center, New York). On a recent inspection of our animal facilities and laboratory spaces, it was noticed that the F/air canisters were either standing upright directly on the tabletop or lying sideways. Since the aeration holes are located on the bottom of the canister, sitting it directly on a flat surface will occlude the airflow (Figure 2 A-B). Laying the canister sideways is also not best because the canisters were not designed to operate in this position. To solve this problem, it was recommended that a holder be purchased

Exposure of personnel to WAG in the workplace is a concern due to the potential of harmful acute and chronic effects. Elimination or minimization of WAG exposure can be achieved when there is an understanding of the potential sources of WAG, and then best practices are developed to mitigate exposures. The scavenger is an important component of many anesthetic machines because they adsorb anesthetic gas discharged from the breathing circuit before being expelled into the air. However, to optimize gas adsorption, the scavenger must be used appropriately, including assurance that the position of the canister allows proper aeration. F/air canisters with aeration holes on the bottom have a design that could cause users to inadvertently occlude the holes. To avoid this, some people may lay the canister sideways, which is also not ideal. By utilizing 3D printing technology, I was able to design a cost-efficient holder for the F/air canister, helping to mistake-proof the positioning of the canister. 3D printing is a technology that is still relatively new and may not be widely available. However, if your institution has 3D printing capabilities, you may want to think outside the box and try your hand at designing a solution for your facility needs! Jennifer Booth, DVM, MS, DACLAM, is an Assistant Professor of Comparative Medicine at Penn State College of Medicine in Hershey, PA. REFERENCE 1. Smith JC. 2008. Waste anesthetic gas safety. p. 183–193. Anesthesia and analgesia in laboratory animals, 2nd ed. London (United Kingdom): Academic Press

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Inside the IACUC

You have heard it before, and we are saying it again, communication is the cornerstone of good IACUC operations. But how exactly does an IACUC maintain good communications with its biggest stakeholder group – the institution’s research community? During the COVID-19 pandemic, IACUCs also had the opportunity to re-examine existing communication channels and develop new ones. This article, conceived and developed by Katy Hall, takes a deeper dive into IACUC communications and how communication channels can be best leveraged depending on the message to be conveyed. Katy and I also go into how developing an understanding of what methods and channels are preferred based on the desired outcome, is key to successful communications. I hope this review of IACUC communication channels augments your current understanding of what works, and what does not, when it comes to connecting with research communities served by IACUCs. Even minor changes in IACUC communications can lead to big results! Stacy Pritt, DVM, MS, MBA, CPIA, CHRC, DACAW

Examining an IACUC Cornerstone: Communication Channels By Kathryn Hall, MA, RLATG and Stacy Pritt, DVM, MS, MBA, CPIA, CHRC, DACAW

ffective communication is an essential component of any Institutional Animal Care and Use Committee’s (IACUC’s) operations. With the movement to remote work due to the SARS-CoV-2 pandemic, and phased reopening plans under discussion and consideration, now is a good time to reassess communication channels and their use. A broad definition of communications is “the exchange of information.” For effective communications to occur, you need an information source, or sender, and an audience, or recipient, of that same information. Sources of communication use some type of media, or platform, to facilitate sharing information (a communications channel). When looking at communications emanating from the IACUC, the first step is to determine the communication’s intent. Examples of the different reasons or goals for communication include: • Purely informational – Recipients who receive the information may have learned something new, such as when the next AAALAC site visit will occur. But they are not expected to respond nor change any behaviors or outcomes based on the communication. • Response requested or required – Recipients who receive this communication are expected to respond in some manner as the communication represents a response request. • Behavioral or outcome change desired – Recipients who receive this type of communication are expected to change their operation’s behaviors or outcomes. If change is not achieved, there could be negative consequences. An IACUC can utilize several different platforms to effectively communicate with the animal research community at their institution. During the SARSCoV-2 pandemic, many IACUCs have embraced technology to bridge the gap formed by social distancing guidelines and some communication platforms changed from in-person to virtual. This article will describe common approaches for IACUC communication to stakeholder groups (e.g., researchers) and the determining factors for which communication channel to utilize to support effective communication. These communication channels will be divided into one-way, two-way, and combination methods, and are relevant as we move into post-pandemic operations.

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One-way Communication

First, let’s discuss one-way communication, also known as linear communication. Imagine a teacher giving a lecture or an author who writes a book. The teacher and the author are the senders of information. The students in the classroom and the readers of the book are the recipients or the audience. One-way communication works well with clear and succinct information when no response or change of behavior is required of the recipient. There are generally few opportunities for clarification during one-way communication, so it is important information shared using one way communication is easily understandable. The following are examples of one-way communication: • Websites • Newsletters/Periodic Updates • Posted Signage

Websites

Many IACUCs maintain a website for internal users to access important information and current materials. A website can be a fantastic platform to share forms, policies, and FAQs at the institutional level and can serve as an up-to-date resource for all users involved in the animal care and use program. IACUC websites should link to other important departmental landing pages such as Occupational Health and Lab Animal Resources (Veterinary Resources). Other departmental websites such as Grants or Research Administration should link to the IACUC’s webpage. When designing and maintaining the page, a comprehensive review of all potential external and internal links should be undertaken. The website should be easy to navigate and provide up-to-date information for the intended audience. If visitors to the page find the information is inaccessible (e.g., broken links) or out-of-date, they will be less likely to use it and refer others to the site.

Newsletters/Periodic Updates

An IACUC may choose to periodically send news-

letters or email updates to the research community highlighting events or updated information. Many types of newsletters exist. A newsletter can be in a hard copy format, such as a printed bulletin that is distributed to targeted groups within the institution’s facilities, or in a digital format such as through email. Newsletters or periodic updates are a great way to bring attention to pertinent information or invite the community to visit an updated resource, such as a website. When using newsletters or periodic updates as a one-way communication channel, material included must be concise and formatted to draw attention to important information and improve readability. Care must be taken with newsletters, or other periodic updates, to ensure consistent formatting and regularity. For example, will they be sent monthly, quarterly, semi-annually, or as needed? Will special editions be needed for important activities such as AAALAC site visits? What type of information will always be contained in the update, such as policy updates, personnel contact information, and regulatory updates?

Posted Signage

Using posted signage is a traditional mainstay method of one-way communication. Since the start of the SARS-CoV-2 pandemic, you have probably seen posted signage outlining safety measures for entering establishments and physical distancing symbols on floors. An IACUC can use posted signage to share information, such as how to report animal welfare concerns and share contact information. When using posted signs, ensure the information is legible and the sign is appropriately mounted. Programs also need to be aware of the potential of “sign fatigue” when sign usage becomes overdone and if individuals find it impossible to review all posted signs. Only the most current and relevant signs should be displayed at any given time.

Two-way Communication

Now let’s discuss two-way communication, also known as bidirectional communication. This style of communication aims to provide information from the sender to the recipient and encourage/request some sort of response, feedback, action, change, and/ or outcome from the recipient. Remember the famous cellular network provider commercial asking, “Can you hear me now?” while talking on the phone in different locations. There was a pause after the question, then a response, “Good.” The person’s response on the other end of the call provided feedback to help the caller identify where coverage was. Two-way communication is a potent form of communication for IACUCs to utilize. Providing information and receiving feedback or requesting action from different animal research community areas can strengthen the entire animal care and use program. When asking for feedback or requesting action, attention must be given to asking the right questions and facilitating participation. The following are examples of two-way communication: • Town Halls • Targeted Meetings

Town Halls

A town hall meeting is also known as an “all-hands” meeting which, as the name states, includes all hands, in other words,

everybody. This type of communication can include a variety of individuals involved in the animal research program. An all-hands meeting for an institution’s animal research program provides a platform for IACUCs to share important information or direct members of the institution’s animal research community to additional information sources. This type of platform lends visibility to IACUC leadership and recent IACUC advancements. Holding an all-hands meeting before an AAALAC site visit is an excellent opportunity for everyone to receive important updates and begin their work towards preparing for the site visit.

Targeted Meetings

IACUCs can boost communications with a targeted approach, either in-person or virtual. An IACUC can tailor its message within meetings to large groups (e.g., departments) or one-onone meetings. Platforms available to facilitate targeted meetings include, but are not limited to, phone, face-to-face, virtual meetings, and open office hours (in-person or virtual). Utilizing a targeted approach, incorporating both large- and small-scale opportunities, can bolster two-way dialogs and encourage stakeholder participation.

Combination Methods

Combination methods, as the name implies, employ both oneway and two-way communication strategies. For example, we all know that sending an informational type of email update rarely gets individuals to change behaviors. Therefore, an IACUC could consider holding a town hall or targeted meeting before or after the announcement of a major policy change via email.

Summary

Our society’s new normal may not resemble life before the pandemic. As we begin to emerge from our current conditions and enter the next phase of this historical event, it is a great time to reflect. While reflecting, take time to review the IACUC communication styles and platforms in use at your institution. Asking questions such as: “Are we using methods to share information effectively? Have any new methods been adopted over the past year? Are any methods no longer serving the IACUC or research community?” may provide some insight. No matter what method or combination of methods your institution uses to communicate, make sure the IACUC communicates often and routinely with all animal research community members and encourages them to communicate with the IACUC. Having an open and collegial communication structure will benefit both the IACUC and the animal care and use program at your institution. The opinions and assertions expressed herein are those of the author(s) and do not necessarily reflect the official policy or position of the Uniformed Services University or the Department of Defense. Kathryn Hall, MA, RLATG, is a Henry M. Jackson Foundation for the Advancement of Military Medicine, Inc. employee supporting the Office of Research Compliance at the Uniformed Services University of the Health Sciences in Bethesda, MD. Stacy Pritt, DVM, MS, MBA, CPIA, CHRC, DACAW is Assistant Vice President at UT Southwestern Medical Center in Dallas, TX. July 2021 Laboratory Animal Science Professional 37

Opioid Analgesic For subcutaneous use in mice and rats only. CAUTION: Federal law restricts this drug to use by or on the order of a licensed veterinarian. LEGAL STATUS--In order to be legally marketed, a new animal drug intended for a minor species must be Approved, Conditionally Approved, or Indexed by the Food and Drug Administration. THIS PRODUCT IS INDEXED--MIF # 900-014. Extra-label use is prohibited. This product is not to be used in animals intended for use as food for humans or food-producing animals. WARNING: ABUSE POTENTIAL, LIFE-THREATENING RESPIRATORY DEPRESSION, and ACCIDENTAL EXPOSURE Abuse Potential Ethiqa XR contains buprenorphine, a high concentration (1.3 mg/mL) opioid agonist and Schedule III controlled substance with an abuse potential similar to other Schedule III opioids. The high concentration of Ethiqa XR may be a particular target for human abuse. Buprenorphine has opioid properties that in humans may lead to dependence of the morphine type. Abuse of buprenorphine may lead to low or moderate physical dependence or high psychological dependence. The risk of abuse by humans should be considered when storing, administering, and disposing of Ethiqa XR. Persons at increased risk for opioid abuse include those with a personal or family history of substance abuse (including drug or alcohol abuse or addiction) or mental illness (suicidal depression). Because of human safety risks, this drug should be used only with veterinary supervision. Do not dispense Ethiqa XR. Life-Threatening Respiratory Depression The concentration of buprenorphine in Ethiqa XR is 1.3 mg/mL. Respiratory depression, including fatal cases, may occur with abuse of Ethiqa XR. Ethiqa XR has additive CNS depressant effects when used with alcohol, other opioids, or illicit drugs that cause central nervous system depression. Because of the potential for adverse reactions associated with accidental injection, Ethiqa XR should only be administered by a veterinarian or laboratory staff trained in the handling of potent opioids. DESCRIPTION Ethiqa XR is an injectable suspension of extended-release buprenorphine. Buprenorphine hydrochloride, an opioid analgesic, is the active ingredient in Ethiqa XR. Lipid-bound buprenorphine hydrochloride is suspended in medium chain fatty acid triglyceride (MCT) oil. Lipids encapsulate the buprenorphine limiting diffusion which provides for larger doses and prolonged action.1,2 Ethiqa XR has a slightly yellow to white opaque appearance. Each mL contains approximately 1.3 mg buprenorphine hydrochloride. The sterile product contains cholesterol, glyceryl tristearate, and buprenorphine hydrochloride suspended in MCT oil. Buprenorphine Formula C29H41NO4

INDICATIONS Ethiqa XR is indicated for the control of post-procedural pain in mice and rats. MOUSE DOSAGE AND ADMINISTRATION Wear protective clothing when administering Ethiqa XR (see Human Safety Warnings). Shake the vial briefly before each use to ensure uniform suspension. If stored refrigerated, bring to room temperature before use. Use aseptic techniques to withdraw the dose into a disposable 0.5 or 1 mL syringe. A 20 to 23 gauge needle should be used for injections due to the viscosity of the drug suspension. The dosage of Ethiqa XR is a single subcutaneous injection of 0.05 mL per 20 gram mouse (3.25 mg/kg body weight). Therapeutic drug concentrations are maintained for 72 hours after the initial dose. If needed, a single repeat dose may be administered 72 hours after the initial dose. Secure the mouse in a scruff-of-the-neck hold. Insert the needle into the dorsal subcutaneous space created by the scruff hold. Inject the entire dose into the dorsal subcutaneous space. An oily sheen may be observed in the dorsal fur of the mouse after injection due to leakage of the oil-based drug suspension from the injection site. The oily sheen may last for 4 to 5 days postinjection. Leakage from the injection site can be minimized by slowly injecting Ethiqa XR into the subcutaneous space. The mouse can be returned to its cage immediately after receiving Ethiqa XR. Do not return any unused drug suspension from the syringe back into the vial. Once the vial is broached, Ethiqa XR can be stored at 15° to 25°C (59° – 77°F) or refrigerated for 28 days. DO NOT FREEZE. RAT DOSAGE AND ADMINISTRATION Wear protective clothing when administering Ethiqa XR (see Human Safety Warnings). Shake the vial briefly before each use to ensure uniform suspension. If stored refrigerated, bring to room temperature before use. Use aseptic techniques to withdraw the dose into a disposable 0.5 or 1 mL syringe. A 20 to 23 gauge needle should be used for injections due to the viscosity of the drug suspension. The dosage of Ethiqa XR is a single subcutaneous injection of 0.1 mL per 200 gram rat (0.65 mg/kg body weight). Therapeutic drug concentrations are maintained for 72 hours after the initial dose. If needed, a single repeat dose may be administered 72 hours after the initial dose. Secure the rat in a passive restraint tube or by holding with a heavy glove with one person to secure the rat and a second person to administer the drug. Insert the needle in the dorsal subcutaneous space. Inject the entire dose into the dorsal subcutaneous space. An oily sheen may be observed in the dorsal fur after injection due to leakage of the oil-based drug suspension from the injection site. The oily sheen may last for 4 to 5 days post-injection. Leakage from the injection site can be minimized by slowly injecting Ethiqa XR into the subcutaneous space. The rat can be returned to its cage immediately after receiving Ethiqa XR. See CONTRAINDICATIONS and Rat PRECAUTIONS for additional information on bedding. Do not return any unused drug suspension from the syringe back into the vial. Once the vial is broached, Ethiqa XR can be stored at 15° to 25°C (59° – 77°F) or refrigerated for 28 days. DO NOT FREEZE. CONTRAINDICATIONS Only administer Ethiqa XR by subcutaneous injection. Ethiqa XR is not intended for intravenous, intra-arterial, intrathecal, intramuscular, or intra-peritoneal injection. Do not use on mice or rats with pre-existing respiratory deficiencies. Do not keep rats on wood chip-type bedding after administration of Ethiqa XR. HUMAN SAFETY WARNINGS Not for use in humans. Keep out of the reach of children. Human User Safety while handling Ethiqa XR: Two trained staff for administration: Ethiqa XR should only be handled and administered by a veterinarian, veterinary technician, or laboratory staff trained in the handling of potent opioids. To prevent human adverse reactions or abuse, at least 2 trained administrators should be present during injection of Ethiqa XR. Protective covering: To prevent direct contact of Ethiqa XR with human skin or mucous membranes when handling the suspension, protective clothing is recommended. Mucous membrane or eye contact during administration: Direct contact of Ethiqa XR with the eyes, oral or other mucous membranes of humans could result in absorption of buprenorphine and the potential for adverse reactions. If accidental eye, oral or other mucous membrane contact is made during administration, flush the area with water and contact a physician. Skin contact during administration: If human skin is accidentally exposed to Ethiqa XR, wash the exposed area with soap and water and contact a physician. Accidental exposure could result in absorption of buprenorphine and the potential for adverse reactions. Drug Abuse, Addiction, and Diversion of Opioids: Controlled Substance: Ethiqa XR contains buprenorphine, a mu opioid partial agonist and Schedule III controlled substance with an abuse potential similar to other Schedule III opioids. Ethiqa XR can be abused and is subject to misuse, abuse, addiction,

and criminal diversion. Ethiqa XR should be handled appropriately to minimize the risk of diversion, including restriction of access, the use of accounting procedures, and proper disposal methods, as appropriate to the laboratory setting and as required by law. Abuse: Abuse of Ethiqa XR poses a hazard of overdose and death. This risk is increased with concurrent abuse of alcohol and other substances including other opioids and benzodiazepines. Buprenorphine has been diverted for non-medical use into illicit channels of distribution. All people handling opioids require careful monitoring for signs of abuse. Drug abuse is the intentional non-therapeutic use of a prescription drug for its rewarding psychological or physiological effects. Abuse of opioids can occur in the absence of true addiction. Storage and Discard: Ethiqa XR is a Class III opioid. Store in a locked, substantially constructed cabinet according to DEA and local controlled substance guidelines. Discard broached vials after 28 days. Any unused or expired vials must be destroyed by a DEA registered reverse distributor; for further information, call 1-833-384-4729. Physician information: Ethiqa XR injectable suspension is a mu-opioid partial agonist (1.3 mg buprenorphine/mL). In the case of an emergency, provide the physician with the package insert. Naloxone may not be effective in reversing respiratory depression produced by buprenorphine. The onset of naloxone effect may be delayed by 30 minutes or more. Doxapram hydrochloride has also been used as a respiratory stimulant. PRECAUTIONS Mice The safety of Ethiqa XR has not been evaluated in pregnant, lactating, neonatal, or immune-compromised mice. As with other opioids, buprenorphine may cause sedation, decreased blood pressure, decreased heart rate, decreased gastrointestinal mobility, and respiratory depression. Use caution with concomitant administration of Ethiqa XR with drugs that cause respiratory depression. The use of paper or soft bedding for up to 3 days following administration of Ethiqa XR should be considered. Normal mice may exhibit an obtunded response to stimuli up to 4 hours after receiving Ethiqa XR. Buprenorphine is excreted in the feces (see Clinical Pharmacology section below). Coprophagy may lead to ingestion of buprenorphine or its metabolites by mice treated with Ethiqa XR and untreated cage mates. Rats The safety of Ethiqa XR has not been evaluated in pregnant, lactating, neonatal, or immune-compromised rats. As with other opioids, buprenorphine may cause sedation, decreased blood pressure, decreased heart rate, decreased gastrointestinal mobility, and respiratory depression. Use caution with concomitant administration of Ethiqa XR with drugs that cause respiratory depression. Rats may exhibit signs of nausea including pica up to 3 days post-treatment. Rats should be maintained on paper or soft bedding to avoid ingestion of wood chip-type bedding after administration of Ethiqa XR. Pica involving wood chip-type bedding can be lethal in rats. Buprenorphine is excreted in the feces (see Clinical Pharmacology section below). Coprophagy may lead to ingestion of buprenorphine or its metabolites by rats treated with Ethiqa XR and untreated cage mates. ADVERSE REACTIONS Mice No adverse reactions were observed in 20 to 25 gram young adult male and female mice after a single subcutaneous injection of Ethiqa XR at a dose 5 times the indicated dose. Laboratory parameters evaluated in the study included hematology and clinical chemistry; histopathology was also performed. In a second study, adult male and female mice received Ethiqa XR subcutaneously at 5 times the indicated dose for three doses at four day intervals. A surgical procedure was performed on the study mice prior to receiving each of the three doses of Ethiqa XR. Mortality was seen in two male mice after the third surgical procedure and dose of Ethiqa XR (total dose of 49 mg buprenorphine/ kg body weight in 8 days). Weight loss has been observed in mice treated post-procedurally with Ethiqa XR. Rats Adverse reactions were evaluated in 180 to 200 gram young adult male and female rats after a single injection of Ethiqa XR. A surgical procedure was performed on the rats prior to administration of a single dose at the intended dose of 0.65 mg/kg or a single dose of 2, 6 or 10-fold excess dose. Adverse reactions also were evaluated in male and female rats administered 2, 6 and 10 times the intended dose for three doses at four day intervals. A surgical procedure was performed on the rats prior to administration of the first of three doses. Laboratory parameters evaluated in the study included hematology, clinical chemistry, urinalysis, histopathology, and bodyweight. Signs of nausea were observed at all dose levels within 24 hours of the dose. Signs included self-licking, self-gnawing and efforts to eat wood-chip bedding. Mortality was seen in 1 of 36 rats exposed to wood chip bedding. Necropsy revealed the stomach and esophagus were compacted with bedding, the bladder was abnormally distended and the urine contained blood. Mortality was seen in 3 of 222 rats treated with Ethiqa XR due to technical complications with serial bleeding of the jugular vein. For technical assistance, or to report an adverse drug reaction, please call Fidelis Pharmaceuticals LLC at 1-833-384-4729. For additional information about adverse drug experience reporting for animal drugs, contact FDA at 1-888-FDA-VETS or http:// www.fda.gov/AnimalVeterinary/SafetyHealth. CLINICAL PHARMACOLOGY3 Buprenorphine can act as an agonist and antagonist at different classes of opioid receptors. Agonism at the mu opioid receptor and, in some cases, antagonism at the kappa or delta opioid receptors are possible underlying mechanisms for the ceiling effect and bell-shaped dose-response curve of buprenorphine. Studies with knockout mice have shown that the antinociceptive effect of buprenorphine, which is mediated primarily by the mu opioid receptor, is attenuated by the ability of the drug to activate the opioid receptor like (ORL-1) receptor. The drug can be described as a ‘full’ and a ‘partial’ agonist at the same receptor depending on the specific assay. There appears to be no ceiling effect for analgesia, but there is a ceiling effect for respiratory depression. Pharmacokinetic studies with bolus injections of buprenorphine in mice and rats provide similar models. After bolus intravenous administration, plasma levels decline tri-exponentially. The drug is n-deakylated in the liver to norbuprenorphine (NBN), an active metabolite. Studies have shown that glucuronide metabolites of buprenorphine and NBN are also metabolically active, and can approximate or exceed the concentration of the parent drug. Un-metabolized drug excreted in the urine and feces one week after injection was 1.9 and 22.4% of the dose, respectively, and 92% of the dose was accounted for in one week.3 Mice Pharmacokinetic parameters of Ethiqa XR were studied in 6-8 week old male and female Balb/c mice following a single subcutaneous injection of 3.25 mg/kg bodyweight. Clinically significant blood levels were observed up to 72 hours after subcutaneous injection. Rats Pharmacokinetic parameters of Ethiqa XR were studied in 8 week old male and female Fischer rats following a single subcutaneous injection of 0.65 mg/kg bodyweight. Clinically significant blood levels were observed up to 72 hours after subcutaneous injection. HOW SUPPLIED Ethiqa XR is supplied in a multi-use glass vial containing 3.0 mL of injectable drug suspension. Ethiqa XR

3 mL vial

NDC 86084-100-30

U.S. Patent No. 8,461,173 STORAGE INFORMATION Store between 15° and 25°C (59° – 77°F) or refrigerated. DO NOT FREEZE. If stored refrigerated, bring to room temperature before use. Once broached, the multi-dose vial should be discarded after 28 days. REFERENCES 1. Mishra et al., Drug Delivery and Transl. Res, 2:238-253; 2012. 2. Bethune et al., The role of drug-lipid interactions on the disposition of liposome-formulated opioid analgesics in vitro and in vivo. Anesth Analg. 93(4):928-33; 2001. 3. Guarnieri et al., Lab Animal, 41(11): 337-343; 2012. Manufactured for: Fidelis Pharmaceuticals LLC CCIT Incubator 675 US Highway One, Suite B113 North Brunswick, NJ 08902 833-384-4729 www.EthiqaXR.com Fidelis Pharmaceuticals® and EthiqaXR® are registered trademarks of Fidelis Pharmaceuticals LLC, a Delaware Corporation. February 2020

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WARNING: ABUSE POTENTIAL, LIFE-THREATENING RESPIRATORY DEPRESSION, and ACCIDENTAL EXPOSURE Abuse Potential This formulation contains buprenorphine, a high-concentration (1.3 mg/mL) opioid agonist and Schedule III controlled substance with an abuse potential similar to other Schedule III opioids. The high concentration may be a particular target for human abuse. Buprenorphine has opioid properties that in humans may lead to dependence of the morphine type. Abuse of buprenorphine may lead to low or moderate physical dependence or high psychological dependence. The risk of abuse by humans should be considered when storing, administering, and disposing of Ethiqa XR. Persons at increased risk for opioid abuse include those with a personal or family history of substance abuse (including drug or alcohol abuse or addiction) or mental illness (suicidal depression). Because of human safety risks, this drug should be used only with veterinary supervision. Do not dispense Ethiqa XR. Life-Threatening Respiratory Depression The concentration of buprenorphine in Ethiqa XR is 1.3 mg/mL. Respiratory depression, including fatal cases, may occur with abuse of Ethiqa XR. Ethiqa XR has additive CNS depressant effects when used with alcohol, other opioids, or illicit drugs that cause central nervous system depression. Because of the potential for adverse reactions associated with accidental injection, Ethiqa XR should only be administered by a veterinarian or laboratory staff trained in the handling of potent opioids. Important Safety Information for Rats and Mice For Rats and Mice: Only administer Ethiqa XR by subcutaneous injection. Ethiqa XR is not intended for intravenous, intra-arterial, intrathecal, intramuscular, or intra-peritoneal injection. Do not use on mice or rats with pre-existing respiratory deficiencies. Do not keep rats on wood chip-type bedding after administration of Ethiqa XR. Use caution with concomitant administration of Ethiqa XR with drugs that cause respiratory depression. For Humans: Ethiqa XR should only be administered by a veterinarian or laboratory staff trained in the handling of potent opioids. Protective clothing is recommended to avoid direct contact with human skin or mucus membranes which could result in absorption of buprenorphine and adverse reactions. Not for use in humans. For more information, consult the Prescribing Information including the boxed warning located on the next page. ethiqaxr.com 833-EthiqaXR (833-384-4729) © 2021 Fidelis Pharmaceuticals, LLC February 2021 FID-ETH-024

Management/Career & Training

ACE Provides AALAS Members Unique Connection Opportunities Enhance your platform knowledge regarding security and new functions. By Mary Kathryn Billings, MBA, MAc, CAE

he AALAS Community Exchange (ACE) has hosted over 6,100 discussions since its inception. The ability to share with and learn from peers in real-time continues to strengthen the knowledge, professionalism, and camaraderie of AALAS members. The AALAS Open Forum is comprised of the entire AALAS membership (13,600+). We all know the importance of virtual connection and learning, particularly during the pandemic. The Open Forum community has proven to be an excellent tool for member engagement. AALAS is proud to provide a way for members to reach their peers. Here are some tips and tricks to keep in mind while using this networking tool.

Tips: Security

As this collaboration platform continues to grow, keep security in mind. The idea of community is open collaboration and networking. However, particular areas and topics are sensitive in nature. Use the anonymous posting feature to safeguard personal and institutional safety. Since ACE is open to all AALAS members, it is important to recognize that members’ security depends upon the security at users' institutions. AALAS requires a member sponsorship for anyone to join, but institutions’ hiring and screening processes vary. When posting on a sensitive topic, post anonymously to protect your identity and your institution. By posting anonymously, identities and facilities are hidden. Questions will still be distributed in the daily digest. To post anonymously, click add under the latest discussion posts section on the community homepage. You should see the option to post your message anonymously. All you need to do is check the box. You will not automatically receive replies to your message in real-time since your identity and email are hidden. Be sure to click the follow star after your message is posted to receive real-time email notifications of all replies.

40 Laboratory Animal Science Professional July 2021

Tips: Calendar

A new calendar system is available on the ACE. On April 1, the AALAS calendar moved from the AALAS website to the ACE platform. The move enabled the calendar to function more robustly, and information will continue to be accessible to nonmembers. Users can now download events to personal email calendars! First, go to the ACE Events Calendar. Select the event you would like to attend. View the event description and included links on the event listing. To download an event from the ACE calendar to your email calendar, click the Download to Your Calendar button. Additional details on submitting information to the calendar for publication are available on our website: https://community.aalas.org/calendar

Tips: Advanced Member Search

The advanced search function in the ACE member directory allows you to search for other AALAS members based on various factors, including location, AALAS branch, credentials, AALAS certification, and AALAS specialized training. The advanced search tool creates instant connections with your peers. Want some study tips? Search for members based on AALAS certification credentials. Thinking about attending ILAM and need a first-hand account of a prior attendee’s experience? Search for members based on ILAM in the specialized training section. The possibilities and connections are endless. Don’t miss out on this great opportunity to make new connections!

Make Connections with ACE

ACE is full of opportunities to make connections with your peers. It continues to be an excellent resource and a great way to reach others in the LAS field. Thank you for being a part of our community! Mary Kathryn Billings, MBA, MAc, CAE, is the Strategic Initiatives Manager at AALAS in Memphis, TN.

Management/Career & Training

Go Team: Research Services within a Biopharmaceutical Company Launching an initiative to provide specialty research services on a broader scale. By Donna Strasburg, LATG and Paige Ebert, MBA, CMAR, RLATG

bbVie is a global biopharmaceutical company headquartered in North Chicago, IL. AbbVie’s Comparative Medicine department is responsible for the care and use of the research animals. It provides operational support, including husbandry, cagewash, facility upkeep, behavior management, veterinary support, including preventative medicine,

emergency care, and surgical services, animal procurement, and research services. The veterinary technicians (VTs) and animal care technicians (ACTs) have historically supported the research teams on weekends to cover dosing requests for rodents. They have been the primary group responsible for nonhuman primate study work. In 2010, the department launched an initiative to provide specialty research services on a broader scale for the research staff. That was the inception of the Research Services Group (RSG). The group was formed by transitioning one veterinary technician to the group and a facility manager with a technical background. The initiative started on a small scale and provided over 1,800 hours of support in the first year. This was primarily through support of one therapeutic group Monday-Friday and the occasional weekend/holiday request. In 2011 a full-time veterinary technician was added to the team and one additional technician later in the year. In 2011 more than 3,500 hours of support were provided. The RSG team was quickly becoming an integral part of the AbbVie research program. The initiative began because of the sporadic cycle of research work. As the pipeline ebbs and flows, the number of in vivo studies and resources required to support them generally follows suit. The RSG group was committed to filling the temporary headcount needs to augment the workload of the research teams with a flexible headcount concept that any group could utilize.

Training Program Refresh

Donna Strasburg is a Research Technologist at AbbVie and is a member of the RTS team.

In 2017 the training program in Comparative Medicine was ready for a refresh. The decision was made to combine RSG and the training program, and the group became Research and Training Services (RTS). With that merger, the team grew to 4 technologists and a manager to oversee the group and support the technical work as needed. The merger July 2021 Laboratory Animal Science Professional 41

it became clear that if we wanted to provide high quality, consistent support across the different therapeutic areas we needed to invest more time and resources.

Embracing an Expanded Role

The RTS team provides data collection such as body weights, tumor measurements, nest scoring, and food and water consumption. Pictured are Paige Ebert and Deb Weisbecker.

allowed for training work to be shared when the research requests were at a minimum. Technical work, creating and revising training documents, and hands-on training became part of all members’ daily job tasks allowing for increased diversity, expansion of skill sets and more team members to support the startup of studies and weekend/holiday coverage. Currently, the RTS team has 4 members whose primary role is technical support and one member whose primary role is coordinating training activities. ACTs and veterinary technicians supplement the full-time members of the team. This is extremely helpful not only for being able to support more research requests but to cover the daily overtime and weekend/ holiday work. This paradigm allows for maximum flexibility. Having ACTs work with the RTS team allows them to expand their skills, add diversity to the day and provide another growth opportunity. The last 3 hires in the group were all from the ACT team. In the past few years, the volume of technical work requests has grown, as has the level of both complexity and scientific engagement. Comparative Medicine provided primarily basic support consisting of dosing and tissue collection in rodents and monkeys before the RTS group was formed. Most of this work was conducted by our veterinary technicians and a few ACTs. This research support was done in addition to their other full-time duties. As the requests became more numerous and complex,

42 Laboratory Animal Science Professional July 2021

RTS has expanded the research support offered to include highly complex model development, study planning, and surgical services. The more routine services provided are dosing and bleeding of mice, rats, dogs, nonhuman primates (NHPs), and rabbits. We also provide data collection such as body weights, tumor measurements, nest scoring, and food and water consumption. Animal identification is an additional service frequently requested that includes tattoos and RFID chips in all species. Rodent surgical services involve several standard models such as mini-pump placement, cannulations, and perfusions. The team supports the maintenance of large animal surgical models, including a colony of vascular and/or gastro-intestinally ported dogs and NHPs. Automated blood sampling (ABS) is conducted in rats, dogs, and NHPs on a routine basis, and several models have been developed and refined by the RTS team. The team works with the 3Rs/Animal Welfare group to complete studies that target assessing and implementing refinements and new technologies. Investigative staff use the RTS team as subject matter experts and frequently consult with members to improve models by creating and learning new techniques. RTS has protocols that allow the team to fulfill study requests by investigators who do not hold protocols. Training is a core service that RTS provides, including handling, restraint, and technical training for all animal users, and the administrative side for creating and updating training materials, assignment and tracking of training, and expertise in software systems. Requests for study support are generally made by submitting a request form through a SharePoint workflow. Once a request is submitted, it is routed to the scheduler. The request is then reviewed, and any follow-up needed for clarification is done with the submitter. Once the request is approved, the scheduler adds it to the RTS shared calendar and assigns the work to the appropriate technician(s). Assigned work is based on skill set and availability. When scheduling ACTs and VTs for support, an email or invitation is sent with the pertinent information. It is policy to submit requests at least 3 days in advance; however, the team is “all for one” and works to fulfill all requests even if they are last minute. By using a shared calendar, all RTS members are kept up to date since transparency is key to encourage team members to back one another up when available. RTS often juggles several different studies, making strong communication and agility key attributes for team members. The RTS team is responsible for their daily work but can depend on their manager for support and assistance when necessary, with technical work. The group frequently collaborates with the surgical veterinarian and other

new skills and having a different type of direct support of the research. While RTS often provides after-hours and weekend support, the work is balanced between all team members. The group has set work hours, but flexibility in the department allows for scheduling adjustments to achieve a good work-life balance.

Conclusion

The RTS team continues to thrive 11 years after its original formation. The core team and support members provide increasingly diverse services.

veterinarians to ensure that the highest quality work is performed and the highest animal welfare standards are achieved. The AbbVie RTS team provides supplemental headcounts for customers who may have restrictions due to technical know-how or personnel numbers. With RTS support, more studies can be executed with high-quality results. Due to the ever-changing types of requests received, the team is regularly learning and developing new skills, which helps keep the job interesting and provides personal development, which is a strong motivator. Also, the veterinary technicians and ACTs also have growth opportunities and increased job satisfaction by learning

The RTS team continues to thrive 11 years after its original formation. The core team and support members provide increasingly diverse services, and the “if we build it, they will come” persona has come to fruition. For the past 11 years, the hours of service provided have increased. In 2019 more than 7,000 hours of support were provided. The research staff collaborates with the team and continues to broaden the horizons of what is possible for this highly functioning service group. Disclosure: All authors are employees of AbbVie and may own AbbVie stock. AbbVie sponsored and funded the study; contributed to the design; participated in the collection, analysis, and approval of the final publication. Donna Strasburg is a Research Technologist in the Department of Comparative Medicine at AbbVie in North Chicago, IL

Paige Ebert, MBA, CMAR, RLATG, is an Operations Manager and Manager of RTS in the Department of Comparative Medicine at AbbVie in North Chicago, IL.

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Management/Career & Training

Launching a Pilot Socialization and Enrichment Plan By Britannia Robinson, BS, RLATG and Susan Rosati, MSEd, CVT, LATG

nvironmental enrichment and socialization aim to positively effect animals’ physical and psychological wellbeing and achieve high-quality research outcomes. A welltrained and engaged staff is key, and combining departmental resources can help attain these goals. As husbandry technicians learn new job tasks and grow in their roles, many are eager to learn new techniques and seek out training opportunities. The University of Pennsylvania’s Laboratory Animal Resources (ULAR) department created a program with two goals in mind. We wanted to ensure the highest levels of animal care and wellbeing through socialization and enrichment and utilize internal resources to educate husbandry technicians and provide career growth opportunities.

For animal handling, the ULAR Scientist Training Team, along with veterinary consultation, set S/E minimum duration times for each species (swine and guinea pigs, 5-10 minutes and rabbits, 10-15 minutes). ULAR’s Research Training Team was approved to use all S/E toys and food items in this program by facility and veterinary staff. After an assessment at week 4, technicians were approved to work alone once a week (See Appendix 1).

Pilot Program

In 2018, ULAR established the Animal Behavior, Socialization and Enrichment Curriculum for husbandry technicians. With departmental leadership approval, the ULAR Scientist Training Division announced the program to ULAR staff. Technicians would contact their managers if interested. Three husbandry department technicians showed interest and volunteered to pilot the program. The program provides in-depth didactic training and hands-on learning. Training topics include a review of species-specific behavior, the effects of socialization and enrichment (S/E), a review of Institutional Animal Care and Use (IACUC) Guidelines, Departmental Standard Operating Procedures (SOPs), and regulations covering S/E. Also, topics such as appropriate enrichment devices and the importance of personal interactions are discussed. Speakers include training staff, as well as clinical veterinary staff. Hands-on training occurred with guinea pigs, rabbits, and swine that were used for species-specific training courses for scientists. We hypothesized that consistent handling and interaction with the animals would make the animals calmer and easier to handle during the training sessions. Managers received a S/E training syllabus to facilitate staff scheduling (Appendix 1). Training would take 1-2 hours up to twice a week, and managers worked with the technicians’ schedules to ensure S/E was rotated into their husbandry schedule while maintaining their daily routines. Meetings were held as needed to discuss the technician’s schedule and to answer any questions. 44 Laboratory Animal Science Professional July 2021

Hands-on training occurred with guinea pigs, rabbits, and swine that were used for species-specific training courses for scientists.

Week 1: Overview of species behavior of guinea pigs, rabbit, and swine. Understanding the importance of socialization and enrichment in laboratory animals. Lecture A June 4th 10:15am, OVQ Conference Room Guinea Pigs and Rabbits

Lecture B June 7th 2pm, OVQ Conference Room Swine

Week 2 and 3: Shadowing ULAR Research Trainers on the enrichment and socialization of guinea pigs, rabbits, and swine. Goals • Visiting Stemmler and Translational Research Building (TRC-7), donning PPE, traffic pattern • Shadowing and practicing socialization/ enrichment with trainers • Observation and review of clinical records and documentation procedures (with documentation practice by end of week 3)

Expectations • Trainees will understand the traffic pattern between species and their housing building • Learn basic S/E training, handling, and behavior of the species • Understand the importance of documentation in laboratory clinical records

Week 4: ULAR Research Trainer observation of trainees. Goals • Observation by ULAR Training Team - Stemmler and Translational Research Building (TRC-7) • Perform all socialization/enrichment for species • Observation of documentation procedures

Assessment • Trainers must be comfortable and demonstrate: • Handling guinea pig, rabbit, and pig species • Performing all socialization and enrichment • Documentation in the clinical records and S/E training log

The first cohort showed that providing S/E to the animals created a less stressful environment for both animals and humans.

Periodic reviews were completed for each technician, including clinical record documentation. If there were any questions regarding documentation, technicians were contacted to discuss the forms. Managers were copied on all reviews. After one year of commitment to the pilot program, technicians were recognized for their contribution to the program at a departmental-wide meeting. Certificates of completion were awarded.

hiding in the back, show excitement when coming out of the cage for socialization time, move from pen to pen for cleaning seamlessly, enjoy being held or touched for longer periods, sit on laps, follow commands such as sitting, spinning, jumping up, and would approach technicians for treats. Animals handled and socialized weekly provided the ULAR Scientist Training Team with smoother training sessions for researchers, easing stress for researchers new to working with a species or are nervous. The program also encouraged improving workplace skills. Workplace fundamentals such as time management, communication, and problem-solving are all skills needed for success. Technicians worked with their managers to ensure their daily schedules, and their S/E schedule, were completed. They learned to contact the S/E coordinator and their cohort when absences, vacation days, or other issues occurred that could prevent successful completion of the S/E. Due to the success of this initial pilot program, we continued with a second cohort of technicians. However, as experienced S/E technicians, the first cohort has been asked to assist with S/E for the animals even after the pilot program completion. They have helped the ULAR Scientist Training Team when other technicians were unavailable to provide S/E. Managers have been supportive, and the technicians have been eager and willing to continue helping with socialization and enrichment.

Pilot Study Findings

Challenges

Weeks 5-8: Socialization and Enrichment of Species. Based on assessment approval from Week 4: All S/E Trainees will be completed in Knowledge Link for the Socialization and Enrichment Training Curriculum. S/E Trainees will provide S/E to guinea pigs, rabbits, and pigs without supervision 1x a week based on their manager’s schedule. Periodically, clinical records will be checked for completion and compliance of S/E during this time. Any concerns, discrepancies or incompletions will be brought up to the attention of the S/E trainees and facility manager(s). Weeks 9 – 11 Socialization and enrichment of species will continue 1x a week. Weeks 12 and up Socialization and enrichment of species will be set to 2x a week*. ACTs will work with their manager’s schedule. *Note: If S/E cannot be done 2x a week for any reason, the manager must contact the ULAR Training Coordinator/Developer.

Appendix 1

The first cohort showed that providing S/E to the animals created a less stressful environment for both animals and humans. Animals would easily come to the front of the cage versus

Throughout the training program’s development and implementation, we expected challenges. One challenge was related to the timing of the socialization and enrichment. Knowing July 2021 Laboratory Animal Science Professional 45

When developing a program like this, communication methods should be discussed before program implementation.

Next Steps and Conclusion

As the program grows, additional enrichment options will be explored. Having an environment that mimics the species’ natural habitat is ideal, but for many facilities may not be possible. By investing time to improve and incorporate new enrichment practices and training processes, the program can become stronger. Next steps will include: • the development of a train the trainer session where previous cohorts can assist in training new S/E program cohorts, • increasing participant numbers to 5 or 6, which will provide more rotation throughout campus and give more technicians an opportunity to take part in the program, • monthly training development and enrichment lectures, • alternative enrichment toys and housing, and • introduction of new species to the program.

By investing time to improve and incorporate new enrichment practices and training processes, the program can become stronger.

when husbandry staff clean the runs or change the cages helped the technicians schedule S/E sessions. Otherwise, technicians might have to wait until the husbandry staff is finished. The ULAR Scientist Training Team also informed the technicians not to provide S/E on certain days if a handson lab session was scheduled. Another challenge involved the animals’ housing location. The university has over 14 animal facilities on campus. Some of the S/E technicians did not work in the building where the S/E training animals were housed. Managers and technicians should be aware of the locations as some areas might need to be entered first or last depending upon the technician’s primary facility. This can make scheduling more challenging, but it ensures they followed specific traffic patterns and presented a teachable moment on traffic patterns and related policies. We also found that different people have a favorite communication style for receiving their S/E schedule and any updates. Some liked email, whereas others preferred text messages. 46 Laboratory Animal Science Professional July 2021

With the positive feedback from our technicians about their experience, the improved behaviors of the animals involved, and departmental support, the Scientist Training Team will continue to expand the Socialization and Enrichment Program. Working with the departmental husbandry and veterinary staff, we will ensure enhancement of animal behavior and enrichment education through this program. In addition, the curriculum will continue to focus on career development skills. We are excited that this program was added to our Animal Care Technician career ladder, giving participants an opportunity for job reclassification. In summary, human and animal participants have benefited. We are eager to continue to offer this opportunity to technicians interested in developing professional skills and laboratory animal science knowledge. Acknowledgments: We would like to thank our first cohort of the program. Denise Arttaway, Dana Major, and Steve Gooseby. They played a crucial role in implementing this program. Thanks to their commitment to the program, we had a successful launch of our pilot, and we can expand and grow the program. We are grateful to Denise, Dana, and Steve’s devotion and care of the training animals and their continuous motivation and willingness to help even after the pilot ended. Their dedicated efforts have set up a strong foundation for this program and provided insight for enhancing our socialization and enrichment methods.

Britannia Robinson, BS, RLATG, is the Manager of Training Development at the University of Pennsylvania (University Laboratory Animal Resources) in Philadelphia, PA. Susan Rosati, MSEd, CVT, LATG, is the Assistant Director of Training Operations at the University of Pennsylvania (University Laboratory Animal Resources) in Philadelphia, PA.

Keeping your facility clean can be as easy as 1 2 3

The first step on TechTrak removes the larger particles stuck to the bottom of shoes.

The second step removes smaller particles and whatever was missed on the first step.

The third step removes any remaining particles, trapping all debris to the mat until cleaned.

July 2021 Laboratory Animal Science Professional 47

Better Products. Better Science. 516.781.0755 ancare.com

ENRICHMENT ITEM

TOYS, TREATS, AND TIPS

DIY: Dehydrated Watermelon Training Chips By Kelsey Lambert, BS, LATG

his simple DIY training treat is great for NHP, swine, rabbits, rodents, and dogs. All dietary items should be approved by your veterinarian for the intended species and individual before use. Note that dehydrated watermelon may resemble muscle tissue!

Ingredients:

• Fresh Watermelon (Figure 1) • We have found NHPs prefer dehydrated apple over the use of other processed or sugary items for training. • We have also found that animals requiring low calorie items, love having dehydrated cucumbers as a training reward.

Supplies:

• Dehydrator • Parchment paper • Sealable plastic bags with sticker labels (or permanent marker) • Food cutting knife • Cutting gloves

Directions:

1. Cut thin slices no more than ¼ in thick (thicker slices will take longer to dehydrate). Watermelon slices should not contain rind.

Figure 1

Figure 2

48 Laboratory Animal Science Professional July 2021

2. Line dehydrator trays with parchment paper. Please follow guidance for your particular dehydrator. (Figure 2). 3. Set dehydrator temperature to 135° (this is the fruit setting for our dehydrator) for at least 18 hours. a. Dehydration times can take between 18-24 hours depending on humidity levels in the room and thickness of slices. 4. Ideal consistency of dehydrated watermelon should be chip-like and easy to break. If it is more taffy-like or flexible, it will need to remain in the dehydrator longer. It is still edible in this consistency; however, it is not as ideal for training purposes – as you cannot create smaller pieces easily and it becomes a little sticky (Figure 3). 5. Once at an ideal consistency, allow the dehydrated watermelon to cool before placing in a storage container/bag. Label your item with the day it was made and an expiration date (in our experience, we have not had to store this longer than 14 days, so we are unsure how long it will stay good for). 6. Can be stored at room temperature or in the refrigerator.

Figure 3

Responsible Conduct of Research

Courses

Need Responsible Conduct of Research Training?

Course content covers the following NIH-recommended subject areas for RCR: 1. Conflict of interest – personal, professional, and financial

2. Policies regarding human subjects, live vertebrate animal subjects in research, and safe laboratory practices 3. Mentor/mentee responsibilities and relationships 4. Collaborative research 5. Peer review

6. Data acquisition, laboratory tools; management, sharing and ownership

Course Titles:

1. Responsible Conduct of Research Introduction 2. Responsible Conduct of Research Part 1: Shared Values 3. Responsible Conduct of Research Part 2: Planning Research 4. Responsible Conduct of Research Part 3: Conducting Research

5. Responsible Conduct of Research Part 4: Reporting and Reviewing Research 6. Avoiding Financial Conflict of Interest in Federal Research

7. Research misconduct and policies for handling misconduct

Additional Bioethics Courses Include:

9. Ethical issues in biomedical research, and the environmental and societal impacts of scientific research

2. Ethical Case 1: Mouse in a Parisitology Experiment

8. Responsible authorship and publication

1. Ethical Decision-Making in Animal Research

3. Ethical Case 2: Rat with Partial Paralysis

Find Them in the New Research Ethics Library

• 9190 Crestwyn Hills Drive, Memphis, TN 38125 July 2021 Laboratory Animal Science Professional 49 email: info@aalas.org www.aalas.org

American Association for Laboratory Animal Science Phone: 901.754.8620

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TECNIPLAST NEWS

The following is sponsored content

Clean and Dirty Side Material Transportation: Additional Savings in Space and Operating Costs

uildings are being designed with more modular construction allowing spaces to be easily converted between holding rooms and procedure space. Other existing spaces face similar challenges in converting areas between rodent, primate, or aquatic housing to keep up with changing research needs. Spaces designated for bedding/food storage falls also into such category of areas the project must consider the modularity and flexibility. Vacuum conveying technology is a proven method of connecting spaces while adapting to modern day flexible designs. It surrounds items around us as part of everyday life. Industrial plants producing building materials and other plastic products to food and beverage plants which stock our grocery store shelves. The same technology made its way into vivarium facilities over 20 years ago and continues to evolve in modern facilities with operational flexibility at the forefront of equipment design criteria. Processing equipment can be equipped to handle a wide range of bedding types including lightweight materials and the redesigned mechanical area reduces the system footprint by over 50% compared to prior generations while granting a proven concept reliability blended with energy efficient concepts to reduce as much as possible the operating costs. Companies such as Tecniplast continue to meet new challenges in facilities with the introduction of the Eolus lines of material handling equipment. The Eolus products cover both clean material delivery to dispensing units and soiled material removal from the facility 50 Laboratory Animal Science Professional July 2021

where it is staged for removal from the site. Its compact and modular construction allow from placement is small mechanical areas and, in some cases, can have the technical areas combined (clean material and dirty material transportation made by a combo-compact technical area) for additional savings in space and operating costs. Systems are completely scalable, and solutions range in size from a wall pass through to offerings designed to traverse through mechanical spaces and levels to the most extreme case of transporting material underground to a dedicated offloading structure. The backwards flow design advances considerations solutions beyond an either positive pressure or negative pressure waste container. Conversations about what might be added or combined provided one facility the opportunity to eliminate the need for third party waste removal and manage their wase removal with its own resources. Another located in Europe has able to utilize fire hose type connections to its waste container and have material removed by a vacuum truck with similar connection hoses. Whatever the specific solution, linking remote equipment areas introduces a layer of complexity beyond standard equipment installation. Space and routing of transportation piping must be routed through existing utility spaces or within a coordinated BIM model. Tecniplast is equipped to manage with its team of experienced project managers and 3D aided design department. They have been able to provide design assistance in early project planning stages and maintain involvement is project implementation. Andrew Arvanites, Business Manager Washing and Automation, Tecniplast. aarvanites@tecniplastusa.com

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UNITY EXCH EXCHANG ANGE AA NITY E AALA E AALAS C LAS COM TY EX T M I I U Y M N N E T MUNI M I U U O G N M AA MU CO HAN AS C OMM XCH OMM OM NITY ANG M E L S E C C C U H A O A G X Y S L S E M C A N T C I A A A X A M Y L S IT O UN GE S C CH AAL E A TY E AALA LAS C AS COMM MMUN AALA TY EX CHAN EXCHANG CHANGE MUNI ANGE ANGE AA MUNI AAL NGE ITY EX AL AS CO ALAS COM X M A Y L N E A E T H O I A U Y G C N C A E T M H N I X MMU E A CHA MUN COM ITY E ALAS HANG CHANGE Y EXC O X G N ANG A M S E C C N U H A X O Y A E L S C E M T C ITY XCH ANG Y EX E AA COM TY EX NITY AALA E AALAS MUNI I G S N M E N A U O G L A MMUN OMMUNIT MMUNITY E NITY EXCH U MM AA S C XCH CHAN EXCHANG C AALA AALAS CO LAS COMM COM NGE MMU S CO NITY E TY EX ALAS ITY S XCHA CHANGE A E N E A AALA AALAS CO S COMMU A MUNI L U Y G M A E T E M I N A O X G E C UN HA NG A E OM HAN AALA OMM ALAS UNITY XCHA Y EXC ANGE GE A LAS C HANG

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TECH TIPS

Insights in husbandry, enrichment, and new techniques and tactics

Figure 1. Female chameleon climbing on an artificial vine, demonstrating this species' adaptations to arboreal habitats.

Husbandry Techniques for a Research Colony of Veiled Chameleons (Chamaeleo calyptratus) By David Jewell, LATG; Alex Muensch BS, MS, LATG; Richard Kupronis, BS, LATG; and Diana P. Baumann, BSc (Hons), RLATG, CMAR

eiled chameleons (Chamaeleo calyptratus) are a large flamboyant species native to Western Yemen and the southern tip of Saudi Arabia. They inhabit valleys and ephemeral stream beds in the desert known as wadis, which collect water during seasonal heavy rains allowing vegetation to grow. As a solitary arboreal species, only one male is found per tree, which they will aggressively defend. This species is adaptable to a wide range of niches within their habitat, being found on acacia trees and various shrubs, as well as in gardens and farmland. Though primarily insectivores, they have been known to occasionally consume small vertebrates such as birds and smaller lizards, as well as vegetation. Veiled chameleons are a non-traditional lab animal species offering huge potential as a research model organism, particularly in evolutionary and developmental biology. The early stage of embryonic growth at the time of oviposition allows researchers to study a continuous range of anatomical development. These animals possess several unique adaptations to their environment, from their turret-mounted eyes that allow for stereoscopic vision, to their zygodactylous feet and 52 Laboratory Animal Science Professional July 2021

prehensile tails that aid their arboreal lifestyle (Figure 1). The unique morphology that allows them to project their tongue also makes them superb ambush predators (Figure 2). Lastly, their amazing ability to change color and pattern is primarily used in social signaling within the species, such as conveying territoriality and breeding receptiveness. Although most chameleon species can be challenging captives, C. calyptratus have been successfully kept and bred in captivity for over thirty years. Despite their relative familiarity, our challenge is to replicate their preferred habitats in a practical way while also maintaining the rigorous welfare standards of laboratory animal husbandry. We incorporated the general principles offered in the Guide for the Care and Use of Laboratory Animals when designing our program of management and care, along with current literature on captive care specifically relating to this species. Our Attending Veterinarian actively participates in all husbandry and welfare decisions and regularly performs a walk-through of the reptile rooms. We are an AAALAC International accredited facility, and all studies are approved by the IACUC. The following describes our cur-

rent solutions in achieving this goal but is not meant to be an all-inclusive description of basic husbandry practices.

Primary Enclosure

Figure 2. Female about to use her tongue to capture prey. A PVC coupling holds a deli cup of hornworms, while a strip of yellow squash is held by a clothespin.

Our animals are housed in Dragon Strand® XL cages (Figure 3) which feature solid back and side panels that provide visual barriers between animals in adjacent cages and help to retain humidity. They also feature a large mesh screen door in front, along with a separate smaller door underneath, allowing cage access at different heights. Cable ties are used to fasten perches onto the built-in ledges, from which artificial vines and foliage are supported. Plastic gutter guard is attached along the cage sides for additional climbing structure. All adult cages are elevated on plastic dunnage racks for easier cleaning of the cage interior as well as the floor below. Cage interiors are spot cleaned daily, with cages and furnishings being disinfected every 4 mo. The Guide does not provide recommendations for non-traditional species and cleaning schedules are based upon the need to provide a beneficial, non-stressful environment balanced with minimal disruption of the animals. When breeding chameleons, it is important to keep a container filled with a suitable digging medium within the cage for adult females (Figure 4). A 2:1:1 ratio of organic sphagnum peat moss, topsoil, and playground sand yields the best results for our chameleons to construct a stable tunnel. During daily checks for eggs, the substrate is stirred and tested for adequate moisture levels by inserting fingers to try to form a tunnel. If the mixture is too dry, water is added until this consistency is achieved. The lay tubs are elevated using PVC stands for increased ergonomic cage maintenance.

Lighting and Temperature

Figure 3. Row of cages for adult chameleons, each measuring 23.75 in W × 23 in D × 48 in H. The leftmost is for a male while the 2 on the right house females, who are provided lay tubs set on PVC stands.

Figure 4. Female starting to dig in lay tub before laying eggs. Rubbermaid® tubs, 17 in L × 11 in W × 8 in H, provide adequate substrate depth for our chameleons to dig a tunnel before laying, while still being easy to remove.

Providing the correct lighting and temperatures is essential to the health and wellbeing of captive chameleons. As they are poikilothermic animals, their core temperature is determined by that of their environment. An ambient daytime temperature of 78° F is maintained with a nighttime drop to 74° F. Overhead room LED lighting is on a 12:12 on/off light cycle and cage lighting is on a 9:15 cycle. The chameleon room does not have windows and lighting cycles are kept constant year-round. Zoo Med™ Reptisun® T5 10.0 fluorescent bulbs are used to provide adequate levels of UVA and UVB wavelengths, and Zoo Med™ 160W Powersun® mercury vapor bulbs to provide basking temperatures between 95 to 115° F, visible light, and additional UV. This setup gives our animals a range of options within their environment where they can thermoregulate and photoregulate. All bulbs are tested prior to use with handheld UVB model 6.2 and UVA+B model 6.5 (Solarmeter® by Solartech, Inc.) meters, and a spectrometer ( Jaz by Ocean Optics, Inc.) to ensure no UVC wavelengths are present, which have been known to cause burn injuries in captive animals.

Hydration

Captive chameleons rarely drink from water bowls, so misting is crucial to their survival. It has been suggested that the head casque of veiled chameleons serves to collect and channel waJuly 2021 Laboratory Animal Science Professional 53

Figure 5. Ultrasonic greenhouse humidifier. PVC piping helps to distribute water vapor evenly through the room.

Figure 6. Hornworms, Dubia roaches, and superworms used to feed chameleons on days alternating with crickets.

The misters are fed by a holding tank connected to facility plumbing. This reservoir automatically refills as it is used, ensuring a constant source of water without the manual labor of refilling. To achieve ambient humidity levels above 50%, an ultrasonic greenhouse humidifier (Ocean Mist® model #MHS10) (Figure 5) operates at timed intervals throughout the day. Filters are used in the water lines feeding both the misters and humidifier to decrease sediment infiltration. The timing of operation for these systems can be fine-tuned throughout the year to account for seasonal fluctuations in environmental humidity.

Nutrition

Figure 7. Hatchling cages, 16 in W × 16 in D × 20 in H, set in drainage trays. The wire shelving unit, 72 in W × 24 in D × 75 in H, also supports lighting and misting fixtures.

ter to their mouths. An automated misting system controlled by a digital timer is utilized, providing hydration to each cage through individual nozzles that can be adjusted directionally.

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54 Laboratory Animal Science Professional July 2021

As with any animal, providing proper nutrition is vital. A rotating feeding schedule of ¾ in crickets, superworms, hornworms, and Dubia roaches is provided (Figure 6). Supplementation alternates between Rep-Cal® calcium, Rep-Cal® Herptivite® multivitamin, and Sticky Tongue Farms© Miner-All (a mixture of calcium and other minerals). Crickets are also gut loaded with Mazuri® Better Bug® diet along with nutrient dense vegetables such as kale, sweet potato, and carrots. Worms are offered to chameleons in plastic deli cups placed in 4 in PVC couplings. These holders are attached by cable ties to the built-in ledges in the cage, where misting water can-

rows on a wire shelving unit (Figure 7). We have incorporated the same misting source used by our adults. To reduce excess water in the cages, we developed a novel drainage system consisting of plastic trays which support each cage, from which we connect hoses to collect wastewater into a common receptacle for disposal. For a heat source, 60W incandescent lamps sit directly on each cage top, and 4-ft twin-bulb light fixtures with T5 10.0 (Zoo Med™ ReptiSun®) bulbs hang above each row of cages to provide UV. A hatchling can reach adulthood in 6 months and a considerable amount of food is consumed during this growth period. Hatchling chameleons are initially fed only 1/8 in crickets using the same supplements, but as they grow are offered increasingly larger size crickets and mealworms along with vegetables, until reaching maturity where they switch to the adult diet schedule.

Conclusion

For a species that is widely kept and bred in captivity, we have incorporated a variety of husbandry adjustments to suit the rigorous practices of a lab animal facility. Using these methods, we have consistently met research needs throughout the past decade, while promoting the welfare of our animals. The intent of this article was to share with the reader these practices, which may also be suitable for other arboreal reptilian species. Acknowledgments: The authors would like to thank the following technicians for their many years of dedicated and caring work with our chameleon colony: Nikki Inlow, Christina Piraquive, and Kristy Winter.

A hatchling chameleon

not accumulate and drown the insects. Vegetables are offered every other day along with worms, and we quickly learned our chameleons did not like leafy greens, preferring squash, bell peppers, carrots, and sweet potatoes. Strips of these vegetables are fastened to the cage foliage using plastic clothespins.

Hatchling and Juvenile Husbandry

In the case of housing our young chameleons, many of the methods already discussed are used but with some modifications. Zoo Med™ ReptiBreeze® screen cages are placed in

David Jewell, LATG, is a Research Technician III at Stowers Institute for Medical Research in Kansas City, MO.

Alex Muensch, BS, MS, LATG, is a Research Technician II at Stowers Institute for Medical Research in Kansas City, MO.

Richard Kupronis, BS, LATG, is Reptile Manager at Stowers Institute for Medical Research in Kansas City, MO.

Diana P. Baumann, BSc (Hons), RLATG, CMAR, is Head, Reptile & Aquatics at Stowers Institute for Medical Research in Kansas City, MO.

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July 2021 Laboratory Animal Science Professional 55

TECH TIPS

Insights in husbandry, enrichment, and new techniques and tactics

Solutions to Minimize Urine Spraying in Single Housed Male New Zealand White Rabbits (Oryctolagus cuniculus) By Alexis Morgan, BS, LAT; Sarah Thurston, BS, LAT, CLABP; Lisa Burlingame, BS, LVT, LATG; and Tara Martin, DVM, MS

n the wild, rabbits are strongly territorial and often react aggressively to strangers invading their home range.1,2 Scent marking is one way rabbits mark territory. Rabbits leave urine, feces, or skin secretions at the borders of their territory to repel intruders and to communicate with neighbors.2-4 Scent may also be used to reinforce social hierarchy, even in animals that share a territory.3 Within our colony of New Zealand White rabbits (Oryctolagus cuniculus), we observed urine spraying in singly housed males. Urine spraying can be associated with health conditions such as fur matting, dermatitis, and urine scalding, but little information is available on treatment or prevention of this behavior. We hypothesized that rabbits engaged in urine spraying as a territorial marking response related to the presence of dominant males within the sprayers’ zone of defense. Further, we hypothesized that alternating neighboring rabbits would change the social hierarchy by increasing the sprayers’ distance from dominant males, thus reducing urine spraying.

Figure 1. Urine Spraying Intervention Process.

56 Laboratory Animal Science Professional July 2021

Materials and Methods

All animals were housed in an AAALAC accredited facility under IACUC approved protocols. Two separate rooms were evaluated, each containing both male and female New Zealand White rabbits with a total population of approximately 95 animals per day. Both rooms were stable and had not recently undergone major changes (new technician, major surgical procedures, building construction, etc.) which would account for behavioral changes. Animals were housed in standard non-ventilated rabbit racks (Allentown, Allentown, PA) measuring 756 mm × 127 mm × 750 mm. Rabbits had individual access to water ad-libitum via a lixit on an automatic watering system and were fed a daily diet consisting of 0.75 cups of chow (Laboratory Rabbit Diet HF 5326, LabDiet, St Louis, MO) and a handful of timothy hay (Timothy Meadow Hay, Oxbow Animal Health, Omaha, NE). All rabbits are provided with one low-value enrichment item per rabbit as a baseline enrichment that is rotated every rack cycle (2

Figure 2. Urine Spraying Scale Visual Representation.

Urine Spraying Scale 0 1 2 3

No Spraying Minimal Moderate Excessive

Table 1. Urine Spraying Scale.

wk). Daily observations were conducted by trained husbandry personnel to monitor for signs of distress or health conditions, including urine spraying. If urine stains were determined to be severe, an Animal Treatment Report (ATR) was generated to request evaluation from a veterinary technician. We identified

20 single-housed intact male rabbits with a history of varying degrees of urine spraying due to non-clinical issues and used them to develop a Urine Spraying Scale (Table 1.) Interventions occurred in three phases – see Figure 1 for a visual representation of this process. Phase 1: On initial observation, urine spraying rabbits were placed on an increased enrichment regimen where new enrichment was provided, and old enrichment removed every Monday, Wednesday, and Friday. This was continued for 1 rack cycle (2 wk), at which point enrichment frequency was increased to daily if urine spraying continued. This is standard for all behavioral issues in rabbits at the University of Michigan.5 Enrichment was provided from varying categories to increase novelty, and whenever possible, destructible enrichment was provided (Table 2) because it is especially valued by rabbits.

Table 2. Categories of Enrichment.

July 2021 Laboratory Animal Science Professional 57

Figure 3. Urine Spraying Results.

Phase 2: If spraying continued despite providing daily enrichment for at least two weeks, then at the next rack change, the rabbit was relocated to a new cage, directly next to a rabbit without a history of urine spraying. A non-urine sprayer was chosen by cleanliness of fur, which was assigned ‘0’ based on the Urine Spraying Scale (Table 1). Two weeks following the first relocation, before the next cage change, observations were noted using the Urine Spraying Scale (Figure 2). Phase 3: If urine spraying had been eliminated, the rabbit remained housed next to the same non-urine spraying neighbor. If urine spraying was still observed, the rabbit was moved to a single freestanding rack, which eliminated any direct neighbor. Visual and olfactory contact with other rabbits in the room was maintained, and alternative enrichment opportunities were further explored.

Metric

Stopped Continued Spraying n=13 to Spray n=4 1.6 1.7

Average Age (Years) Average Spraying 1.3 Score (Table 1.) Average Body 3.1 Condition Score (BCS) Progression to 50 Matting (%)

1.9 3.3 50

Table 3. Demographic Information.

Results

Phase One (Increased Enrichment) Following placement on daily enrichment for a minimum of 2 wk, all 20 rabbits continued urine spraying. All rabbits were moved to phase 2 of intervention. 58 Laboratory Animal Science Professional July 2021

Phase Two (New Neighbor) Of the 20 initially identified rabbits, 3 were euthanized for study-related reasons, giving a final sample size of 17 rabbits. Of these 17 rabbits, urine spraying was eliminated in 10 (59%) following relocation next to a non-urine spraying neighbor. 7 of 17 (41%) rabbits required additional intervention in phase 3. (Figure 3) Phase Three (Single Rack) Seven rabbits were placed in single racks with no immediate neighbors to either side. Of these, urine spraying was eliminated in 3 rabbits and reduced in 2, meaning 5 rabbits (71%) showed improvement. (Figure 3) Overall Of 17 affected rabbits, 13 (76%) showed elimination, and 2 showed reduction of spraying following movement away from initial neighboring rabbits. Demographics and Clinical Information The average age of rabbits that did and did not respond to intervention was similar at 18–19 mo old. Rabbits that continued to spray had a slightly higher average initial spraying score and body condition score. In both groups, 50% of rabbits with urine spraying progressed to matting (Table 3). Mats were removed via combing or shaving by veterinary technicians. Body condition scores were assessed using a rabbit Body Condition Score Chart6 which ranges from 1 (very thin) to 5 (obese).

Conclusion

This data supports our hypothesis that urine spraying in male rabbits is a territorial marking behavior, perhaps meant to reinforce dominance hierarchies. When rabbits were placed next to a new neighbor, the majority stopped urine spraying. We recommend leaving rabbits with their new neighbors for one cage-change cycle (2 wk) to assess effectiveness. If animals continue to spray, consider rotation to a different neighbor or, if possible, placement in a cage where they have no adjacent neighbors.

Because rabbits are a social species, placement in cages without neighbors must be weighed carefully against the effects of increased isolation. Urine spraying is a normal behavior in rabbits and, absent other signs of stress and a low urine spraying score, rabbits may be able to be left with a neighbor if careful monitoring is employed. Rabbits with urine staining should be routinely checked for mats or skin lesions such as redness or scabbing. If other neighbors are trialled, animals should be carefully monitored to ensure that this repeated change to the social environment does not lead to additional signs of stress. Separation via a divider limits the social behaviors rabbits can engage in to reinforce dominance hierarchies.7 This could make olfactory communication, like urine spraying, more important. Future work should examine the impact of social housing on urine spraying prevalence in rabbits. In addition, additional methods of reducing urine-spraying should be examined for those animals that continue to spray despite changing their social condition. Alexis Morgan, BS, LAT, Unit for Laboratory Animal Medicine, University of Michigan, in Ann Arbor, MI.

Sarah Thurston, BS, LAT, CLABP, Unit for Laboratory Animal Medicine Refinement and Enrichment Advancements Laboratory, University of Michigan, in Ann Arbor, MI. Lisa Burlingame, BS, LVT, LATG, Unit for Laboratory Animal Medicine Refinement and Enrichment Advancements Laboratory, University of Michigan, in Ann Arbor, MI.

Tara Martin, DVM, MS, Unit for Laboratory Animal Medicine Refinement and Enrichment Advancements Laboratory, University of Michigan, in Ann Arbor, MI. REFERENCES 1. Mykytowycz R. 1968. Territorial marking by rabbits. Scientific American 218(5):116–129. 2. Mykytowycz R. 1973. Reproduction of mammals in relation to environmental odours. J Reprod Fertil Suppl 19:433–446. 3. Sneddon I. 1991. Latrine use by the European rabbit (Oryctolagus cuniculus). J Mamm 72(4):769 – 775. 4. Arteaga L, Bautista A, Martines-Gomez M, Nicolas L, Hudson R. 2008. Scent marking, dominance and serum testosterone levels in male domestic rabbits. Physiology & Behavior 94:510 – 515. 5. University of Michigan. [Internet] 2020. Rabbit Social Housing. Animal Care and Use Program. [Cited March 3, 2021] Available at https://animalcare.umich.edu/animal-use/rabbit-social-housing 6. Ontario Rabbit. [Internet] 2014. Body Condition Score Chart. Ontario Farm Innovation Program. [Cited March 3, 2021] Available at http://ontariorabbit.ca/?page_id=359 7. Thurston S, Burlingame L, Lester PA, Lofgren J. 2018. Methods of pairing and pair maintenance of New Zealand White rabbits (Oryctolagus cuniculus) via behavioral ethogram, monitoring, and interventions. J Vis Exp 133:57267. doi: 10.3791/57267

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July 2021 Laboratory Animal Science Professional 59

TECH TIPS

Insights in husbandry, enrichment, and new techniques and tactics

Exploring Isolator-raised Heterozygous Nude Mice as Soiled Bedding Sentinels: Balancing the 3Rs with Protecting Colony Health By Hannah M. Atkins, BS, DVM, PhD, DACVP; Jennifer L Booth, DVM, MS, DACLAM; Sarah E. Clark, DVM, DACLAM; Lori A. Davis, BAS, CVT; and Tiffany L. Whitcomb, DVM, DACLAM

ealth monitoring programs are a cornerstone of laboratory animal care and an important component of minimizing biomedical research variables. Programs are unique to each institution and vary based on the type of research, the animals' immune status, the resources available, and the risk to the animal and the institution if a disease goes undetected. Historically, health monitoring for colonies of mice has involved exposing sentinel animals with a known health status to soiled bedding from the colony animals, with the goal of sentinel mice becoming infected and producing detectable levels of antibodies, parasites, or viral antigens. Exposing sentinel mice to soiled bedding allows for monitoring colony health without impacting research by directly testing and/or euthanizing colony mice. Recent advances in technology have emerged that can replace the soiled bedding sentinel (SBS) mouse with environmental testing methods. These new testing methods involve performing PCR from filters located in the exhaust of rodent housing systems.5,6,9,11 Drawbacks to these testing methodologies, like false-positive results and false-negative results, are being refined to the point of preventing the need for SBS for many institutions.5,6,9 However, SBS remain the best option for health monitoring in facilities with housing systems that were not designed for filtration of exhaust air, as is our facility's case. Here we discuss the advantages and disadvantages of using high health status heterozygous nude mice (Het-Nude) as an SBS that preserves the 3Rs and provides significant financial savings.

Het-nude mice were suggested as a potential alternative stock by the vendor. These mice are isolator raised and of a high health status as part of the production of athymic nude mice (Foxn1nu/nu). Even though we were aware that Het-nude mice are used at other institutions as SBS, there was some concern about whether heterozygous nude mice were immunologically normal. Several references discussed that Het-nude mice have lower thymic weights than wild type Foxn1+/+ from a decreased lymphoid population2,8,10 and fewer bone marrow stem cells.4 One reference described Het-nudes as able to produce antibodies,7 which increases their promise for use as SBS. To determine whether Het-nude mice would provide appropriate health monitoring for our colonies, we compared seroconversion rate and titer levels between CD1-elite and Het-nude SBS. After we received approval for the study from the IACUC, mice of each stock were pair housed in rooms that had historically been positive for MNV, MHV, or both and systematically exposed to soiled bedding every 1-2 wk (depending on cage type) for a minimum of 12 wk (shown in both conventional and ventilated caging in Figures 1 and 2). A blood sample was collected for serology at 3 wk, 9 wk, and final sampling. At necropsy, thymic and splenic weights were gathered, and histology of the thymus was performed. Due to the lack of significant differences in antibody titers and seroconversion rate between the two, we elected to use Het-nude mice as SBS in our barrier rooms.3

Het-nudes as SBS

There are many advantages to using Het-nude mice as SBS in a health monitoring program (Figure 3). One benefit to using Het-nude mice is that it helps to uphold the principles of the 3R’s (reduction, replacement, and refinement). Het-nude mice are primarily used to mate females with male Foxn1nu/nu, due to insufficient lactation of female Foxn1nu/nu mice. This mating scheme results in 50% of the litter being Foxn1nu/nu (commonly used in oncology research) and an excess of Het-nude mice,

Our facility monitored SBS in the barrier facilities for opportunistic agents due to housing many immunocompromised strains. To protect the existing colonies and prevent confounding sentinel results, they needed to be negative for opportunistic agents at the vendor and upon arrival to our facility. Isolator-raised CD1 mice with a high health status made this possible. However, there were occasionally availability issues. 60 Laboratory Animal Science Professional July 2021

Advantages and Disadvantages of Het-Nudes as SBS

Figure 1. An individually ventilated cage from a barrier facility.

which don’t have a common research need. Providing a new purpose for excess Het-Nude mice could essentially reduce the numbers of isolator-raised CD1 mice generated to support sentinel programs. An additional benefit to using Het-nude mice as SBS is that they seroconvert to MNV and MHV at similar rates as isolator-raised CD1 mice used as SBS.3 Further benefits to using Het-nude mice as SBS are their pleasant disposition, reduced cost (one-third of the cost of isolator-raised CD1 mice), and availability. Although there were many advantages to using Het-nude mice as sentinels, we encountered some disadvantages to using this strain in a health monitoring program. First, our study was limited to testing for MHV and MNV. Empirically, the Hetnude SBS have detected the same bacterial and viral pathogens as the isolator-reared CD1 mice. Still, controlled testing needs to be conducted to confirm susceptibility to other infectious and opportunistic pathogens. Another disadvantage of HetNudes as SBS is that Foxn1nu/nu mice are highly susceptible to Corynebacterium bovis, putting Het-nudes at risk of being exposed in the event of an outbreak. Corynebacterium bovis is a small, gram-positive rod bacterium that can cause chronic infections and clinical signs that often manifest as a scaly skin disease in immunocompromised mice, such as the Foxn1nu/nu, however immunocompetent mice, such as the Het-nude, are asymptomatic.1 Based on individual institutional concerns, further screening of Het-nudes for C. bovis may be necessary prior to use as SBS.

Conclusions

Figure 2. One conventional (open top) cage from a conventional facility.

Figure 3. Advantages and Disadvantages of Using Isolator-reared HetNude Mice as Sentinels.

Het-nudes are a readily available, economical choice for SBS that support the 3Rs. Het-nudes are also isolator-reared, making their immune system naïve to most common mouse pathogens and opportunistic microorganisms. The study described here showed no significant difference in detectable MNV and MHV antibody levels after exposure to soiled bedding from an exposed colony. However, there are several disadvantages to using Het-nudes as SBSs, including limited empirical evidence regarding post-exposure opportunistic organism antibody production and that Het-nude mice may serve as fomites for C. bovis in the event of an outbreak. Therefore, additional controlled studies with different organisms and multiple serum antibody testing points are needed. It may be essential to screen mice at delivery when using Het-nudes as sentinels in mouse populations susceptible to C. bovis. In conclusion, Het-nudes are a cost-effective, resource-savvy, institution- and colony-specific SBS option for surveying common pathogens in mice. Ethics statement: All studies described in this article were approved by the Penn State College of Medicine Institutional Animal Care and Use Committee. The work described here was funded by the Penn State College of Medicine, Department of Comparative Medicine with some of the Het-nude mice donated by Charles River and some of the antibody testing donated by IDEXX Laboratories.

July 2021 Laboratory Animal Science Professional 61

Tiffany L. Whitcomb, DVM, DACLAM, is an Associate Professor of Comparative Medicine and Associate Director of Animal Resources at Penn State College of Medicine in Hershey, PA.

Read more about this original research published in the January 2021 issue of JAALAS (JAALAS 60:152-159: Antibody Titers and Seroconversion Kinetics of Outbred Swiss and Heterozygous Nude Soiled-bedding Sentinels for Murine Norovirus and Mouse Hepatitis Virus).

Hannah M. Atkins, BS, DVM, PhD, DACVP, is an assistant professor and veterinary pathologist at the Penn State College of Medicine in Hershey PA.

Jennifer Booth, DVM, MS, DACLAM, is an Assistant Professor of Comparative Medicine at Penn State College of Medicine in Hershey, PA.

Sarah E. Clark, DVM, DACLAM, is an Attending Veterinarian with Covance by Labcorp Animal Welfare and Veterinary Services in Ann Arbor, MI.

Lori A. Davis, BAS, CVT, is a Veterinary Support Associate-Quality Assurance Technician in the Department of Comparative Medicine at Penn State College of Medicine in Hershey, PA.

62 Laboratory Animal Science Professional July 2021

REFERENCES 1. Burr HN, Lipman NS, White JR, Zheng J, Wolf FR. 2011. Strategies to prevent, treat, and provoke Corynebacterium-associated hyperkeratosis in athymic nude mice. J Am Assoc Lab Anim Sci 50(3):378-88. 2. Chen L, Xiao S, Manley NR. 2009. Foxn1 is required to maintain the postnatal thymic environment in a dosage-sensitive manner. Blood 113:567-574. 3. Clark SE, Davis LA, Booth JL, Atkins HM, Whitcomb TL. 2021. Antibody Titers and Seroconversion Kinetics of Outbred Swiss and Heterozygous Nude Soiled-bedding Sentinels for Murine Norovirus and Mouse Hepatitis Virus. J Am Assoc Lab Anim Sci 60:152-159. 4. Holub M, Necas E, Jirásková Z. 1998. Essential defects of athymic nude mice affect also the nu/+ heterozygotes. Folia Microbiol (Praha) 43:491–492. 5. Mailhiot D, Ostdiek AM, Luchins KR, Bowers CJ, Theriault BR, Langan GP. 2020. Comparing Mouse Health Monitoring Between Soiled-bedding Sentinel and Exhaust Air Dust Surveillance Programs. J Am Assoc Lab Anim Sci 59:58–66. 6. Miller M, Brielmeier M. 2018. Environmental samples make soiled bedding sentinels dispensable for hygienic monitoring of IVC-reared mouse colonies. Lab Anim 52:233–239. 7. Mink JG, Radl J, van den Berg P, Haaijman JJ, van Zwieten MJ, Benner R. 1980. Serum immunoglobulins in nude mice and their heterozygous littermates during aging. Immunology 40:539–545. 8. Pantelouris EM. 1968. Absence of thymus in a mouse mutant. Nature 217:370–371. 9. Pettan-Brewer C, Trost RJ, Maggio-Price L, Seamons A, Dowling SC. 2020. Adoption of Exhaust Air Dust Testing in SPF Rodent Facilities. J Am Assoc Lab Anim Sci 59(2):156–162. 10. Scheiff JM, Cordier AC, Haumont S. 1978. The thymus of nu/+ mice. Anat Embryol (Berl) 153:115–122. 11. Zorn J, Ritter B, Miller M, Kraus M, Northrup E, Brielmeier M. 2017. Murine norovirus detection in the exhaust air of IVCs is more sensitive than serological analysis of soiled bedding sentinels. Lab Anim 51:301–310.

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Across the Pond: Animal Welfare Ethical Review Body (AWERB) Meetings in the Wake of COVID-19 By Nicky Windows and Alanah Mudie

nder the Animals (Scientific Procedures) Act 1986, it is a legal requirement in the UK to have an Animal Welfare Ethical Review Body (AWERB) within every facility that uses, breeds, or supplies lab animals. With animal welfare at the front of research decisions, AWERB meetings are an opportunity to raise 3Rs awareness and applicability. Staff handling animals are to be provided significant advice on animal welfare issues, including acquisition, accommodation, care, and use.

This process does not end with license approvals; the AWERB follows up to monitor the level of care and welfare provided for animals housed or used in the establishment. Project development is monitored based on the effect on the animals and whether the project is running as planned. Regular assessments to advise on better 3Rs use take place. Discussions between AWERB members and the applicants are vital to ensure all staff involved are advised and aware of welfare and ethical issue responsibilities. Training opportunities are identified to ensure all research is done at the highest level. Ultimately, an AWERB is responsible for helping to promote a “Culture of Care” within the establishment and the relevant wider community.

Sharing Experiences

In February, members from different AWERB committees met to discuss their experiences with moving online. Usually, an AWERB will meet regularly to fulfill all tasks, including reviews of project license applications and surrounding issues. After an unexpected and unprecedented year, these meetings have continued almost entirely online. Understandably, this changes the meeting dynamic, but it’s important committees continue to fulfill their functions while research continues. It was vital to learn about different AWERB members’ experiences to gauge how they feel and check that standards were still being maintained.

Going Virtual

After several surveys, delegated members discussed committee opinions surrounding the pros and cons of meeting online. This included feedback from Penny Hawkins (RSPCA), Matthew Allen (University of Cambridge), Kally Booth (Institute of Animal Technology; University of Dundee), Bentley Crudgington (AstraZeneca), Ngaire Dennison (Laboratory Animal Vet Association; University of Dundee) & Cathy Abbott (University of Edinburgh). 64 Laboratory Animal Science Professional July 2021

While responses offered a wide range of feedback, surveys from all speakers provided several repeating themes, and the consensus was similar across most AWERBs involved. Time-saving opportunities were noted; in a fast-paced and busy industry, time is valuable! People were happy to conduct meetings with fewer tangents and more focus. Online meetings provided structure and efficiency. Ultimately, technology enhanced organizing meetings and sticking to agendas. Moving to online meetings has reduced travel, saved money, and reduced carbon footprints. With computer access, anyone wanting to be involved in an AWERB meeting can do so. Accessibility has potentially boosted the opportunity to get colleagues more involved with their AWERB. Survey results indicated an improvement in meeting attendance. With platforms such as Zoom, Teams, Google Meet, etc., becoming the normal way of communication, attendees have utilized the technology for their benefit. Options like chat boxes for comments or questions have reduced public comment fears. Online meeting accessibility has not gone unnoticed.

Potential Concerns

It was noted that virtual situations make it harder to read the room’s mood on certain topics. While this may sound like a negative, it also means that the mood cannot influence others’ decisions. However, when discussing particularly sensitive topics, the ability to read the mood can help ensure attendees are comfortable and happy. Other issues come with meeting online. Even with a year’s experience, technical issues happen. Connection issues, bandwidth problems, cameras and microphones not working, the list goes on. How many times have we all said, “You’re on mute!” by now? Privacy has been a concern for some. The material discussed in an AWERB meeting can be sensitive. Uninvited attendees being a part of the meeting is problematic, so having cameras

on is important. Recording meetings presents personal security and privacy worries. While recordings are ideal for keeping minutes and reviewing meetings, the recordings need to be stored securely. Unsurprisingly, the hardest part has been the lack of faceto-face, personal networking. AWERB meetings are ideal opportunities to spend time with colleagues. Kally Booth from the University of Dundee agreed. “From the presentation, I think AWERB are possibly running more efficient now they are online. I think going forward, meetings will remain heavily online with one or two face-to-face because people clearly miss that interaction,” Booth said.

Maintaining Standards

Maintaining the longstanding “Culture of Care” that continues to improve welfare standards across the country has been essential, even during these extraordinary times. Welfare should continue to remain a priority no matter the circumstances. AWERBs have done an excellent job ensuring this and should be proud of the work they have put into their everyday roles, and committees. “I feel it is really important to maintain the same standard of the AWERB. The running of the AWERB not only contributes to good science, it ensures animal welfare is kept at the highest possible standard. These standards cannot be allowed to slip. The AWERB meetings will also be offering support to NACWOs, scientists, vets, and lay people as this is also really important during this time as we are all finding things a little difficult,” Booth said to presenters. Ngaire Dennison, also from the University of Dundee, shared that while in general, the online platforms had a positive impact on AWERB meetings, the survey of Named Veterinary Surgeons the Laboratory Animal Veterinary Association had undertaken indicated vets were concerned some members,

specifically NACWOS and lay people, may be less likely to provide input during meetings. With a focus on keeping things running and getting used to the technology, Dennison thinks that going forward, AWERBs should review issues of participation and interaction to make sure everybody still feels welcome and involved. “The chair is key, and if that person isn’t confident in the role, they should seek support/training,” Dennison emphasized. Bentley Crudgington is an independent member on the AWERB at AstraZeneca. He also felt there is an opportunity to remove participation barriers to ensure all voices are heard at the AWERB. “We often report that our AWERBs are ‘well attended’ and ‘productive,’ but these metrics do not consider how fully attendees feel they can participate or how it feels when they do or can’t. For example, the AWERB cannot discuss the things that are never said. Online platforms allow for committees and contributions to be made anonymously. This may be something we continue to make use of as we move back towards hybrid and face-to-face meetings,” Crudgington commented.

Conclusion

While AWERBs are working hard and keep things moving and keep research environments safe for animals, there is also a level of care required to keep colleagues engaged and ensure everybody is working together to produce better research for a better future ultimately. Nicky Windows is a Global Commercial Manager with Datesand in the United Kingdom. Alanah Mudie is a Social Media Executive with Datesand in the United Kingdom. July 2021 Laboratory Animal Science Professional 65

Support the AALAS Foundation! News from the AALAS Foundation, including the “Swing into Science” Contest

into

The monkey has played an important role in COVID-19 research – that’s why the AALAS Foundation is celebrating the monkey in several of this year’s activities. The “Swing into Science” virtual contest challenges contestants to creatively paint/decorate a 10-inch wooden monkey. The contest includes four categories: Individual, Corporate, Branch, and Institution/Organization. A winner will be selected from each category. Additionally, a Best of Show winner will be chosen by AALAS leadership members, and a Fan Favorite will be awarded to the entry whose photo receives the most likes in the “Swing into Science” contest photo gallery on the AALAS Foundation’s Facebook page. All winners will be announced, and prizes awarded at the AALAS Foundation’s Appreciation Reception held October 20, 2021, at the AALAS National Meeting in Kansas City, MO. Visit https://tinyurl.com/AF-MONKEY for more information and to register for the contest. The deadline to enter the contest is September 1, 2021.

“Celebrate the Monkey” COVID-19 Awareness Lapel Pin

Speaking of celebrating the monkey – the AALAS Foundation is also excited to announce that it has developed a “Celebrate the Monkey” lapel pin as part of its Celebrate Animal Research & Education (CARE) public awareness program. Anyone making a $5.00, or more, in-person donation at the AALAS Foundation booth during the National Meeting in Kansas City, MO, will receive a “Celebrate the Monkey” COVID-19 awareness lapel pin. Anyone making a $10 or more online donation to the AALAS Foundation between October 15 and December 31, 2021, will also receive a “Celebrate the Monkey” lapel pin – while supplies last.

Virtual Silent Auction

The AALAS Foundation is hoping its second annual virtual Silent Auction will be a huge success, and, for that, it needs your help. Please consider donating an auction item. Not sure what to donate? Items that always prove popular are animal-related gifts, event tickets, jewelry, gift baskets, gift certificates, hotel getaways, travel packages, sporting goods and memorabilia, unique items from your area, and of course, and the latest must-have electronics. All auction item donors are, once again, asked to hold their auction item donations and agree to ship the item(s) directly to the winning bidders after the virtual Silent Auction. If you would like to donate, you can do so now. Go to https://tinyurl.com/AF-2021-Auction to pledge your auction item donation. Auction item pledges must be received by September 30, 2021. 66 Laboratory Animal Science Professional July 2021

THE MONK EY BRATE CELE

The AALAS Foundation also plans to conduct its first hybrid live auction – virtual and in-person - which will take place following its Appreciation Reception on Wednesday, October 20, from 6:30-8:30 pm at the AALAS National Meeting in Kansas City, MO. The Appreciation Reception is always a fun and lively event. This year’s Appreciation Reception theme is a sports tailgate party, so plan on wearing your favorite sports team attire and join us for an incredible evening. The monies raised through the annual contests and auctions help support the AALAS Foundation and its mission. The Foundation’s mission is to inform the public about the critical role of animals in research and educate the public about all the compassionate professionals working in the laboratory animal science field (psst..that’s all of YOU!).

AALAS Crossword By Kari Buchanan, RLATG

Zebrafish

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4 What unit is used for measurement of conductivity? 6 How do zebrafish distinguish between kin and non-kin? 10 The zebrafish is a member of the ______family of fish. 11 What vaccination is required to work with aquatic species? 12 Zebrafish are typically housed at how many fish per liter? 13 What is the most common route of administration of an anesthetic agent in zebrafish? 14 What sense organ runs along the side of the zebrafish body that allows them to detect movements and vibrations in the water? 18 What is the scientific name for zebrafish? 20 What part of the fish helps in locomotion? 21 What fish organ extracts oxygen from the water? 23 Baby fish after they hatch are called: 24 Fish should not be used for egg collection more than once per_________.

1 What is the name of the geneticist that initially identified zebrafish as a research model? 2 The release of eggs and sperm is known as: 3 What is one of the most common species of fish used in laboratory animal facilities? 5 What scientific word describes the shell surrounding and protecting the zebrafish embryo? 7 Although they don’t have lungs, zebrafish have proved particularly useful in the study of what disease manifested in the respiratory system? 8 The ability to regrow a missing part of the body is called: 9 What organ of the fish helps maintain buoyancy? 15 The zebrafish genome is organized into______________chromosome pairs. 16 What is the small plastic container use to hold embryos and larvae in called? 17 What term is used when fish are gulping air at the surface of the tank? 19 Which structure covers the gills? 22 What disease is caused by the oval-shaped, dinoflagellate algae parasite called Oodinium pilularis?

July 2021 Laboratory Animal Science Professional 67

AALAS Answers Zebrafish 1

G S E P 6 7 8 9 O L F A C T O R Y S E N S R W U E W G N B G I 12 13 E F I V E E I M S N R N B 14 T G C E L R U R A E L A D 18 I O T D 20 S F I N S I E I I O R N S N 21 22 G I L L S V E E 23 R L A R V V 24 W E E K T

Z M I C R O S I E M E N S H B N N OW R 11 R T E T A N U S N I F O I L I N E S H 17 P R I O P I N G

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A T E R A L W 16 E P 19 A N I O R E T P T Y E R F R I I C D V U I A E L S U H M

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