Before you begin...
To access the Online Protocol you will be required to submit a user name/id and password.  See the Frequently Asked Questions (FAQ) for specific instructions for gaining access.

What to expect...
You can use this new web-based system to create new protocols, submit amendments, renew protocols, submit Personnel Qualifications forms, and update your personal information. You can save your work at any time as a draft and return later to edit. When you submit your protocol application by choosing the SUBMIT button at the end of the form, the protocol is locked and is sent for IACUC Review. There are a few changes to the way information is collected because the data you enter goes directly into a database.  The protocol application will let you know before you submit if there is critical information missing and you may correct before you submit.

A building process...
As you are guided through the application, your protocol will be custom-built for you. The “Procedures to be performed” section asks you to choose the procedures you will use and creates a protocol with only the questions that are applicable to your studies.Over time, you will create Amendments, Rewrites, and Renewals for this protocol. When you create one of these applications, all of the data from the previous application will be "pulled forward" and easy to amend.

Previously submitted protocols...
Protocols submitted prior to the 1/24/05 implementation of the Online Protocol have already been imported into the new system.  The old system did not capture data from all sections of an application, so the data we have imported is limited.  Also, since the old system was not in the best shape, there may be discrepancies between what we have and what you know is correct.  Please do not be alarmed by this.  Feel free to correct these errors when you come across them.

Finding a protocol...

It’s easy to get a list of your current protocols.  From the menu choose "List the existing protocols for PI ID #", leave the field empty and hit the SUBMIT button. You will receive a list of your current protocols, the status of the protocol and buttons to edit or print those protocols.  If you have permission granted to access another PI’s protocols, you may enter the ID# in the field and hit the SUBMIT button.

People...
We require your help to keep the contact information for all of your personnel up to date.  From the menu choose “Edit your personal information”.  Don’t forget that new personnel must fill out a Personnel Qualification online form AND you must add the new person to each protocol by filling out an Amendment.

General Tips and Recommendations… 

Use only one species per protocol. If you do the same experiments on another species, simply choose the "clone" menu choice to produce a copy and then edit for the second species. 

 

The IACUC recommends that you combine all experiments for each particular species into one protocol application.

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Frequently Asked Questions… 

 How do I get a password? What if I forgot my password?

Click here to request a login and you will receive an email response within 24 business hours. Only UCSD employees and students may obtain access to this system.

 Where do I get help for computer problems?

The IACUC web site is best viewed with IE 5.0/higher or Netscape 6.0/higher, resolution 800x600 and COOKIES TURNED ON. If cookies are not turned on, you will not be able to submit an online protocol. Do not use your browser BACK button when in the online form. For technical assistance, please call your departmental computer support first, then call 858-534-1086.

 

Questions about Forms

 Where do I get help with online forms?

First, click on the red question mark or the hyperlinks that you see on the form for further explanation about each question on the form. If you still need help, call 858-534-6069. 

Who must apply to the UCSD IACUC?

You must have a Protocol for Animal Use approved by the UCSD Institutional Animal Care and Use Committee (IACUC) before you can conduct any project involving research, teaching, or testing using live, vertebrate animals at UCSD. The protocol must be approved by the IACUC before you can begin the proposed activity.  Only UCSD faculty and individuals otherwise eligible to apply for research grants through UCSD may be the Principal Investigator on an Animal Use Protocol. The use of animals in research, teaching and testing is a privilege of faculty at UCSD, not a right, and it may be withheld or revoked by the IACUC. 

Can I save my work and come back later?

Yes.  Scroll to the bottom of the form and hit the button “Save as a Draft”. You may come back at anytime and continue to fill out the form. 

Can I put more than one species on my protocol?

No.  Each protocol should include all of the experiments and procedures that you will perform on one species. 

What should I do if I am working on more than one species?

Use the menu choice “Clone a Protocol” to create a copy of any protocol; the clone will appear with a new Protocol Number.  You may then edit any aspect of the protocol and submit. 

Can I let someone else fill out my Animal Use Protocols?

The UCSD IACUC encourages Principal Investigators to write and submit each Animal Use Protocol.  The online form makes it easy to give complete and correct answers.  However, a staff member may fill out an online protocol with the permission of the Principal Investigator.

 Can I let someone else fill out my Personnel Qualifications Form?

No. The IACUC has determined that each person who uses animals must fill out his/her own Personnel Qualifications (PQ) Form.  You may not fill out this form for another person. 

How long does it take to review my Personal Qualifications Form before I receive approval?

Your Personal Qualifications Form will be reviewed within one week and you will be notified of any classes required or other requirements. 

 Can I attend training classes prior to filling out and submitting a Personnel Qualifications Form?

Yes, although it is recommended that you fill out the Personnel Qualifications Form as early as possible to avoid any possible delays.

 Will training taken at other universities count in lieu of the UCSD training program? 

Anyone performing animal research at UCSD must take, at a minimum ,Orientation to Research at UCSD. Additional training may be required depending on background experience, procedures, and/or species.

 Do I need to fill out all sections of the Animal Use Protocol?

Yes.  You need to answer each question in order to submit the Animal Use Protocol. If you do not answer every question you not be allowed to submit the Protocol.  If an answer is incomplete, your Animal Use Protocol may be delayed and you will receive an email from the IACUC asking for more information.

 Why is there data missing in my Animal Use Protocol?

The UCSD IACUC has recently launched a new Online Protocol System. There are many features which will make it much easier to complete your forms, keep track of your approvals, “clone” old forms to create new applications, view your current forms and change information online.  The old system did not capture data from all sections of an application, so the data we have imported is limited.  Also, since the old system was not in the best shape, there may be discrepancies between what we have and what you know is correct.  Please do not be alarmed by this.  Feel free to correct these errors when you come across them. 

What if I can’t answer a question on the form?

First, check out the hyperlink help for the question. If you still need help, please call the IACUC office at 858-534-6069.  An IACUC administrator may help you or refer you to an analyst for more help.   

Can I see a list of my Animal Use Protocols?

Yes. After you log in you will have a menu of options.  Choose List the existing protocols for PI ID #.  If you leave the field blank it will default to a list of the protocols from the logged-in user.  You may also enter another ID number in this field. 

How can I decide which Pain and Distress Category my experiments are in?

The USDA regulations require UCSD to report each year on the number of animals used and the pain category of each animal.  We must therefore ask you to report both prospectively and retrospectively on your animal use and category. A good explanation is available in Pain and Distress Categories C, D & E.   

Must I complete the Animal Protocol application if I use only snails, shrimp, bees, worms, coral, or microorganisms in my research to teaching/instruction activities?

No. You only need to complete the animal use application if you are working on a vertebrate animal. PHS Policy defines an animal as “any live, vertebrate animal used or intended for use in research, research training, experimentation, or biological testing, or for related purposes”.  The Animal Welfare Act Regulations (AWAR) define an animal as “any live or dead dog, cat, nonhuman primate, guinea pig, hamster, rabbit, or any other warm-blooded animal, which is being used or is intended for use for research, teaching, testing, experimentation, or exhibition purposes, or as a pet.  This term excludes birds, and rats and mice bred for use in research.”  The IACUC ensures the campus is in compliance with all regulations by applying the AWAR and PHS regulations to all vertebrate animal use on campus for teaching and research.   

Who should be listed on a protocol?

Everyone with direct animal contact should be listed. New personnel should be added to a protocol prior to their use of laboratory animals. Each person must also complete and submit a Personnel Qualifications Form. 

Can I get help with writing my animal use protocol?

Yes. For questions on federal assurances, funding or experimental design, email iacuc@ucsd.edu or call 858-534-6069. For consultation on animal procedures or veterinary care issues email kjenne@ucsd.edu or call 858-822-5687.  For consultation on anesthesia, analgesia or surgical issues email jfujimoto@ucsd.edu or call 858-534-8612. 

What if all I need is some blood or tissue, or I want to buy some antibodies?

You must have a Tissue Transfer Approval before you can import animal tissue (fixed or unfixed) from another investigator within UCSD or from an outside source. Generally, the tissue source must have an NIH Animal Welfare Assurance. 

 

How do I?

 How do I get animal facility access?

1.       Log in from the IACUC Home Page

2.       Fill out a Personnel Qualifications (PQ) form and submit

3.       Ask your P.I. to add you to each protocol that you will work on or to send an email to iacuc@ucsd.edu.

4.       You will receive an email confirmation of your PQ with a list of the mandatory classes you must attend.

5.       Attend Orientation to Animal Research at UCSD after you register online.

6.   Go to the Animal Care Program office to get your photograph taken and hand scanned for access. This service is available each Tuesday and Thursday 1:30 to 4:00 p.m.

Note: your ID badge and facility access will not be issued until you have completed the above steps.

Who do I call if I have a question about animal anesthetics, analgesia, surgery, monitoring, or training in procedures?

The Animal Care Program has many services, equipment and training available to you.  For a consultation on anesthetics or analgesia call 858-534-8612.  For other services see the Animal Care Program Index of Services web page. 

Who can I call with a question that is not on these FAQs?

Please call the Animal Welfare Program at 534-6069.  We will be glad to answer your question or direct you to the help you need. 

Who should I contact if my project involves hazardous biological agents or radioactive material?

UCSD Environmental Health and Safety has a website with detailed information about obtaining a BUA or RUA and also contains valuable information on the employee occupational health program, ergonomics, chemical safety,  and hazardous waste. 

Do I need to participate in the Occupational Health Program and medical surveillance?

You will need to take the risk assessment screening questionnaire located online at the EH&S web site to determine if you are required to enroll in the program or if it is recommended that you enroll in the program.

  

When?

 How long does it take to review my Animal Use Protocol before I get approval?

You must allow 6 – 8 weeks for the approval process after you submit your application.  Your Animal Use Protocol application goes through a very thorough review process where it is reviewed by veterinary staff, Environmental Health and Safety, administrative professional staff, and the entire Institutional Animal Care and Use Committee at a monthly meeting. 

What happens to the protocol after it is submitted?

All protocols receive veterinary review, administrative review by Animal Welfare Program staff to assure that all animal care and use conforms to PHS, USDA, and UCSD requirements, and (as necessary) Environmental Health and Safety (EH&S) review and Conflict of Interest (COI) review. Depending on these reviews, the Principal Investigator may be asked for additional information or clarifications by telephone, email, or letter. These responses may then receive additional veterinary and administrative review. When complete, the protocol is reviewed by the UCSD IACUC. One member may be appointed "Primary Reviewer" for a particular protocol. If the Primary Reviewer has questions not previously addressed in veterinary or administrative review, he or she may contact the PI directly. At the monthly meeting of the IACUC, the Primary Reviewer presents the protocol, and his/her recommendations, for discussion. The IACUC will vote to either:

• Approve
• Approve with Conditions
• Require Modifications Before Approval
• Defer
• Withhold Approval  

Will I be notified of the expiration and renewal dates of my Animal Use Protocol?

It the responsibility of each Principal Investigator to track the expiration dates of each protocol.  You will receive a courtesy notice from the IACUC (Animal Welfare Program) allowing you enough time to complete your renewal.  If you do not respond by the deadline for renewal, your protocol will expire and you cannot purchase or use animals on this campus.

 What are the meeting dates and submission deadlines for the IACUC?

The IACUC meets monthly on the third Wednesday of each month.  The deadline for each meeting is the third Friday of the previous month.  A yearly calendar of these dates is published.

How long is a protocol approval good for?

      Three years. You should write the animal use application to include all work and all animal requested for three years. You will be required to rewrite the protocol application every third year.  However, federal regulations also require you to update the information on your protocol annually.

How do I renew my protocol?

      From the IACUC website choose log in.  After you log in you will have a menu of options.  Choose Annual Renewal.

   When should I report changes or modifications to my protocol?

     Every change or modification to your protocol must be approved by the IACUC before you begin. From the IACUC website, choose log in. After you log in you will have a menu of options. Choose Amend Protocol.

 If I received word that my protocol was approved, when can I start my work?

     If your approval letter states full approval you may begin work immediately.  If your approval letter lists conditions then you must fulfill the conditions before beginning the work. If your approval letter states “Modifications Required…” you must answer the questions asked of you and wait to receive approval to begin your work.

Can I order animals before I have an approved Animal Use Protocol?

No.

  

 What is the UCSD Animal Care and Use Program?

Federal Regulations require that each institution have an Animal Care and Use Program to assure that all Animal Welfare laws, regulations and policies are implemented and consistently followed at the institution. At UCSD the Animal Care and Use Program consists of three components which work together – the IACUC, the IACUC Office and the Animal Care Program 

What is the IACUC?

The Institutional Animal Care and Use Committee oversees the University's animal care and use programs, facilities and procedures and ensures the appropriate care, use and humane treatment of animals being used for research, testing and education. The IACUC is also responsible for reviewing all animal use protocols, ensuring compliance with federal regulations, inspecting animal facilities and laboratories and overseeing training and educational programs. The IACUC serves as a resource to faculty, investigators, technicians, students, and staff, providing guidance in planning and conducting all animal use procedures in accordance with the highest scientific, humane, and ethical principles. The faculty members of this Committee are appointed by the Institutional Official (Vice Chancellor for Research) from each school and functional unit of the University. 

What is the IACUC Office?

The IACUC Office is the administrative, regulatory and compliance arm of the Animal Care and Use Program.  You should contact the IACUC Office to get an Animal Use Protocol approved, renewed or amended. You should contact the IACUC Office to report animal concerns.

 What is the Animal Care Program?

The Animal Care Program is the veterinary care, housing, and husbandry arm of the Animal Care and Use Program.  You should contact the Animal Care and Use program to arrange for purchasing animals, housing animals or to report animal health concerns. You should contact the ACP with facility or caging concerns. 

What is the meaning of this term/abbreviation/acronym?    Please check our glossary. 

What is our PHS Assurance Number?

University of California San Diego and SIO	A3033-01
VASDHS and VMRF	A3659-01

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If you wish to propose a study that uses a species not on this pull down list, please email vetservices@acp.ucsd.edu.  ACP veterinary staff will respond to you within 24 hours to discuss the species, housing requirements, and location.  If approved, you will be notified that the new species has been added to the pull down list.

Goals of the research must be clearly written to explain the purpose, scope, and significance of the research being proposed. It should be written in language that can be understood by a college graduate with no medical background. Scientific jargon should be held at a minimum. Abstracts that are submitted to scientific meetings and abstracts which are parts of a grant proposal generally have too much scientific jargon and assume considerable knowledge of the subject being discussed. Not all scientific disciplines are represented on the Committee and the non-faculty members of the Committee should be able to clearly understand the intent and significance of the study.

Explain your rationale for using animals in your studies. The rationale should include reasons why non-animal models cannot be used. Consider in vitro testing, computer modeling, cell lines, organisms lower on the phylogenetic scale, bacteria, fungi, plants, drosophila, non-vertebrates. Check these sites for help:

http://www.vetmed.ucdavis.edu/Animal_Alternatives/main.htm

http://www.nal.usda.gov/awic/

http://altweb.jhsph.edu/

United States Department of Agriculture (USDA) defines animal as any live or dead dog, cat, nonhuman primate, guinea pig, hamster, rabbit, or any other warm-blooded animal, which is being used, or is intended for use for research, teaching, testing, experimentation, or exhibition purposes, or as a pet. This term excludes: birds, rats of the genus Rattus and mice of the genus Mus bred for use in research, and horses not used for research purposes and other farm animals, such as, but not limited to livestock or poultry, used or intended for use as food or fiber, or livestock or poultry used or intended for use for improving animal nutrition, breeding, management, or production efficiency, or for improving the quality of food or fiber. With respect to a dog, the term means all dogs, including those used for hunting, security, or breeding purposes.

The United States Animal Welfare Act (AWA) regulations require that principal investigators consider alternatives to procedures that may cause more than momentary or slight pain or distress to animals, and to provide a written narrative description of methods and sources used to determine that alternatives were not available. These requirements are explained in USDA policy 11 and policy 12.  The PHS policy requires compliance with the AWA and the Guide for the Care and Use of Laboratory Animals.

Procedures classified as category D or category E

Category D: Potential pain or distress relieved with anesthetics, analgesics and/or tranquilizer drugs or other methods for relieving pain or distress.
Category E: Pain or distress or potential pain or distress that is not relieved with anesthetics, analgesics and/or tranquilizer drugs or other methods for relieving pain or distress.

 

Animals to be used in Category D or E procedures should be of an appropriate species and quality and the minimum number required to obtain valid results. Methods such as mathematical models, computer simulations, and in vitro biological systems should be considered whenever possible. Proper use of animals including avoidance or minimization of discomfort, distress and pain consistent with sound scientific practices is imperative.  Faculty, students, and staff should consult Essentials for Animal Research, a useful resource for investigators.

How to satisfy regulatory requirements for alternatives

Principal Investigators often ask about the requirement to search for alternative procedures to those involving alleviated or unalleviated pain or distress.  The AWA requires the Principal Investigator to consider alternatives to procedures that may cause pain or distress to animals and to provide a written narrative in their protocol of the methods used and sources consulted to determine the availability of alternatives. Alternatives should be considered during planning of the animal use proposal and should form part of the routine scientific literature search and not be separate from it. If a bona fide alternative method is identified, the written narrative should justify why this alternative was not used.

Alternatives should aim at avoiding or minimizing discomfort, distress or pain without compromising research goals using the so-called “3 Rs”. The 3 Rs include: replacement with no-animal systems when possible, reduction in the number of animals required to obtain scientifically valid data through better experimental design and refinement techniques that decrease or eliminate pain or distress. All three approaches should be considered.

The federal requirements can be met by any of the following:

• Database search (consult more than one major database): include names of databases searched, date the search was performed, period covered by the search, keywords and/or the search strategy used.
• Attendance of colloquia or conferences.
• Consultation with subject experts besides the PI, include consultant's name, qualifications and date and content of the consult.

Animal Welfare Information Center (AWIC), an information service of the National Agricultural Library, was created specifically by Congress to provide information on ways to minimize pain and distress. AWIC can formulate search strategy, select key words and databases, access unique databases, train in conducting effective alternatives searches, and perform no-cost or low-cost electronic database searches. AWIC has developed an anesthesia and analgesia database that is now available on ALTWEB.

Other sources for alternatives searches are:

• University of California Center for Animal Alternatives
• Antibody Resource Page
• USDA Web Site: USDA Web Site: http://www.aphis.usda.gov/ac/9CFR99.html
• Animal Care Factsheet:  http://www.aphis.usda.gov/oa/pubs/awact.pdf ).
• Animal Welfare Information Center http://www.nal.usda.gov/awic/
• USDA web site:   http://www.aphis.usda.gov/ac/polmanpdf.html
• The GUIDE:  http://www.nap.edu/readingroom/books/labrats/
• ALTWEB web site at Johns Hopkins:  http://altweb.jhsph.edu/

The IACUC recognizes that, for certain situations, a computerized literature search may not be an appropriate means of identifying alternatives.  In such cases, the investigator should record the source of expert information, qualifications and the date of consultation. The PI must also supply a narrative that includes all findings from the search for alternative methods.

All strains, breeds or genotypes used in these studies must be listed in this section. Although you only see one line on the form, more lines will be created each time you hit the SAVE STRAIN button. It is essential that the IACUC reviews potential health issues due to phenotypes expressed in genetically altered animals. You may describe these phenotypes briefly in this section and later in the Pain and Distress section you will indicate how you will minimize and recognize pain and distress in these animals.

The Lab ID section is for your optional use if you have a shortened strain designation ("nickname") that you would like to enter.

This is the ANIMAL NUMBERS SECTION.  Here you request the total number of animals you will need for studies during the THREE YEAR course of this protocol. Although in years past you were asked to request numbers for one year, this is no longer true.  You are asked to plan and justify the animal numbers for THREE YEARS.

Step One: Plan out your experimental groups.  The IACUC has determined that each experimental group must be listed and described.

Step Two: Determine the pain/distress classification for each experimental group of animals by referring to the IACUC Policy on Pain and Distress. Breeding colony groups are always Pain/Distress category C.

Step Three: Fill out the table; to add new groups simply press the SAVE GROUP button. If you have indicated that you will run your own breeding colony (i.e. you checked the box for breed animals in the Procedures to be Performed Section) then two groups will automatically appear in this table - Colony Maintenance and Transferred. You cannot delete these groups but you can place a 0 for number of animals if you do not transfer animals out of the protocol.

EXAMPLE:

12. Experimental Groups for 3 Years

	Number of Animals in each Category of Pain/Distress
Experiment (or Group)	C	D	E
	14556	180	0	Grand Total= 14736

Step Four: Press the REFRESH button to automatically total for you the numbers of animals requested.

Step Five: For each group/experiment that you have listed a text field will be created.  Enter the brief description of the studies that will be performed on the group.  This should be an overview that enables the IACUC reviewer to understand your experimental design and be able to understand what procedures (including treatments, data collection, endpoints) will be done on each animal through the course of the study. Provide a brief explanation of numbers requested.

EXAMPLE:

Transferred to Another Protocol - Describe experimental design. Provide calculations to justify the number of animals requested. Do not include procedural details here.

We work closely in collaboration with Dr John White. In order to consolidate breeding efforts some of the strains that are on our mutual program project are bred under this protocol and then they are transferred as needed to Dr. White's experimental protocol. Dr. White’s protocol number is S09240. We transfer an estimated 1109 mice per year of BLR2, BLR9, IL-1R, IL6, MjD88 and LSS2 to Dr White's protocol. We also transfer C57BL/6 and BALB/c mice when needed as controls from our colony. Over 3 years 3345 mice will be transferred.

Breeding Colony Maintenance - List all animals used as sires, dams, offspring with unwanted genotypes or phenotypes or any animals euthanized (or died) without any experiment. Provide calculations that justify the number requested.

We generally maintain 2-3 breeder cages of each line listed in Question 11. jlr7 now being backcrossed onto the B6 background TyD88: this strain is difficult to breed and we maintain homozygous and het breeding NoD mice are purchased and bred to MRN T cell transgenic mice to generate the M/XN strain GFP are het breeding for lethality reasons TNFa transgenic are het breeding - about 1/3 have the correct genotype TOGe are being backcrossed to the C57Bl/6 background 20 individual strains with 6 addition strains generated by intercross or backcross 2-6 breeding pairs with exchanges 4x/year=600x3years=1800 Some animals do not have the appropriate genotypes in strains that are maintained heterozygously for lethality reasons. Mice that are sacrificed due to unwanted genotype include: GDP wt, TNF YT, MJD88 het and wt. We are also backcrossing mice onto the B6 background and the wt offspring are not used.Estimated 330/yr x 3years=990

Immune Response - Describe experimental design. Provide calculations to justify the number of animals requested. Do not include procedural details here.

To examine a role for Slc7a2 in modulating the immune response, we will carry out standard experiments to assess T-dependent and T-independent immune cell function. Mice are immunized with T-dependent (DNP-KLH) or T-independent (DNP-Ficoll) antigens by ip injection. Blood samples are collected every 5 days and the mice are boosted with antigen at 20 days. After 35 days the mice are euthanized and tissues collected. 10 mice/group x 3 groups/genotypes x 1 repeat = 60 mice/year

Tumor transplantation - Describe experimental design. Provide calculations to justify the number of animals requested. Do not include procedural details here.

The interaction between mammary tumor cells and the surrounding stromal cells has been shown to affect tumor growth and progression. To study the importance of genetic differences in iNOS and Slc7a2 in the mammary epithelium versus the mammary stroma, we will transplant 1 mm pieces of tumor tissue of one genotype (wild type, iNOS-/- or Slc7a2-/-) into the fat pad of mice of another genotype (wild type, iNOS-/- or Slc7a2-/-) and assess the growth rate and histologic phenotype of the transplanted tissues. Mammary fat pads are cleared and tumor tissue removed from donor females is implanted into the cleared fat pad. Tumors of different genotypes are transplanted into the fat pads of mice of different genotypes (all congenic in the C57Bl/6 strain) to assess epithelial/stromal interactions. Mice will be euthanized when tumors reach 1 cm in diameter. These studies are carried out in the C57Bl/6 strain. 10 mice/group x 3 genotypes x 2 experiments = 60 mice/year

More help is available in Animal Experimental Design

Question 13. Justification of the number of animals requested.  How were the sample size, number of groups, and number of repetitions determined?

A key principle in the ethical use of animals in research, testing and teaching is that the number of animals used in each project is the minimum necessary to obtain valid and meaningful results. By law the IACUC must review the number of animals requested in each protocol and agree that the number is appropriately justified in terms of the stated goals of the project.

 

The number of animals should be based on the experiments designed to test the hypothesis or otherwise achieve the goals of the project. That is, the number of animals must be justified by the scientific objectives. A brief description or outline of the proposed experiments should be provided to the IACUC that includes:

·          The purpose of each experiment or set of related experiments,

·          The number and species/strain of animals per group/subgroup,

·          The number of groups/subgroups, and

·          The number of control and experimental animals

 

If the project involves euthanizing animals for tissue, the protocol should explain how the number of animals requested relates to the amount of tissue/cells needed for particular in vitro experiments. Animal numbers cannot be justified on the basis of how many experiments the lab personnel can perform in a week, month, etc; nor can the cost of the animal be the primary justification.

 

Research and testing studies should be designed to test statistical significance of the result(s) with a minimum number of animals.The number of animals requested can be based on previous experience or published information using similar procedures, and whenever possible, justified statistically. Statistical techniques should be used to maximize the analysis of the data generated from each animal, thus reducing the total number of animals necessary for a particular set of data.The number of animals must be sufficient to obtain meaningful data. Replication within experiments can lead to a seemingly large number of animals being used; however this may be well-justified and necessary. Biological variation in the test system can obscure the effects of a given intervention if the sample size is insufficient to obtain a statistically significant result (e.g., n=1 is generally not acceptable). It is useful to consult with a statistician or utilize a statistical software package during the design.

Pilot Studies

Depending on the nature of the pilot study, the number of animals can be justified in terms of:

·          Determining the feasibility of a procedure or technique,

·          Assessing the effect of a procedure on the animal, or

·          Generating sufficient data to allow the investigator to project the approximate number of animals that will be needed to reach statistical significance (such as power analysis).

 

All animals involved in the project must be included in the protocol and justified. This includes not only experimental animals, but also donor animals, breeding pairs, pregnant mothers, and offspring that cannot be utilized because of genotype, sex, etc. Any need for additional animals due to anticipated losses due to morbidity, mortality or other known difficulties with the experimental procedures should be explained and justified.

Repetition of Experiments

Repetition of experiments is important in certain cases. For example, changes in technology that greatly enhance the data resolution may justify a series of experiments being repeated. Also, comparisons between species are important if certain specific phenomena are to be shown to be more generally applicable. Repetition that is a direct repeat of a previous study is less readily justified and investigators are discouraged from such studies. The Animal Welfare Act requires that investigators state that a proposed activity is not "unnecessarily duplicative" of previous studies.

 

More help is available in Animal Experimental Design and Online Statistics Textbook

Section 16. In Vivo Blood/Tissue Collection

If you agree to follow the guidelines as stated in the IACUC Policy, you can simply check the box.  If you are asking for a variation from the policy you must describe your procedure in the text field and provide justification.Too much blood collected at any one time may cause hypovolemic shock, physiological stress and even death. If smaller volumes are collected too frequently, anemia may result.

 

As a general rule, 20% of the total blood volume can be collected at one time every 2-4 weeks, or 1% at more frequent intervals of 24 hours or more. The total blood volume can be calculated as approximately 6% of body weight. The estimated volume at exsanguination is approximately half of the total blood volume.

 

Mice

 

To improve vasodilation effects in rodents, it is helpful to warm the entire patient. This can be accomplished in 10-15 min at 40^o C with a special commercially available warming chamber. Care should be taken to prevent overheating.

The choice of anesthetics is an important consideration when collecting blood from rodents due to physiologic effects of the anesthetic. Consult with an ACP veterinarian.

If you are not experienced in blood collection techniques, training is recommended. If you have questions or comments about any of the above techniques, contact an ACP veterinary technician at vetservices@acp.ucsd.edu.

 

Total blood volume = 6% of lean body weight.Maximum blood collection = 10% of total blood volume every two (2) weeks.

Example: 30 gm mouse = 0.03 kg x 0.06 = 1.8 ml x 0.10 = 0.18 ml

Collection Site	Advantages	Disadvantages
Lateral tail vein	·    Anesthesia not required ·    Vein is easily accessed	·         Must be securely restrained ·         Yields only small quantities ·         Some specialized equipment needed
Orbital Sinus or Plexus	·    Large volumes of blood can be collected	·         Anesthesia is required
Submandibular Lancet	·    Anesthesia not required ·    Excellent technique for serial blood sampling ·    Moderate volume of blood can be collected	·         Requires some specialized training ·         Some specialized equipment required.
Lateral Saphenous Vein	·    Anesthesia not required ·    Excellent technique for serial blood sampling ·    Moderate volume of blood can be collected	·         Requires some specialized training ·         Some specialized equipment required.
Cardiac Puncture	·    Maximum volume of blood can be collected	·         Requires deep anesthesia. ·         Non-survival procedure only

 

Rabbits

Total blood volume = 6% of lean body weightMaximum blood collection = 20% of total blood volume every 2 - 4  weeks 

Examples:
             4 lb rabbit    =   1.80 kg x 0.06   =          108 ml x 0.20   =           21.6 ml 
             6 lb rabbit    =   2.72 kg x 0.06   =          163 ml x 0.20   =           32.6 ml 
             8 lb rabbit    =   3.60 kg x 0.06   =          216 ml x 0.20   =           43.2 ml

            10 lb rabbit    =  4.50 kg x 0.06   =          270 ml x 0.20   =           54.0 ml 

 

Rabbits may follow the above bleeding schedule as long as packed cell volume (PCV) and total plasma proteins (TPP) are monitored.  The occurrence of anemia, hypoproteinemia, or unthriftiness require appropriate supplementation and a rest from further bleeds.  The duration of this rest will be determined by the attending veterinarian.  Animals should be weighed weekly if on the above bleeding schedule and their weights recorded appropriately.

 

Collection Site	Advantages	Disadvantages
Marginal Ear Vein	·         Anesthesia not required ·         Vein is easily accessed ·         Yields small - moderate quantities of blood	·         Must be securely restrained ·         Some specialized equipment is needed ·         Topical anesthetic is recommended
Central Ear Artery	·         Large quantities of blood can be collected	·         Topical anesthesia is strongly recommended (due to the possibility of arterial spasm) ·         Training recommended
Lateral Saphenous Vein	·         Anesthesia not required ·         Collection of small quantities of blood	·         Training recommended ·         Some specialized equipment needed
Cephalic Vein	·         Anesthesia not required ·         Collection of small quantities of blood	·         Training recommended ·         Some specialized equipment needed
Jugular Vein	·         Large quantities of blood can be collected	·         Anesthesia is recommended ·         Requires specialized training
Anterior Vena Cava	·         Maximum quantity of blood can be collected	·         Requires anesthesia ·         Requires skill ·         Risk of cardiac tamponade
Cardiac Puncture	·         Maximum quantity of blood can be collected	·         Requires deep anesthesia ·         Non survival procedure

  

 

Rats

 

To improve vasodilation effects in rodents, it is helpful to warm the entire patient. This can be accomplished in 10-15 min at 40^o C with a special commercially available warming chamber. Care should be taken to prevent overheating.

The choice of anesthetics is an important consideration when collecting blood from rodents due to physiologic effects of the anesthetic. Consult with an ACP veterinarian.

If you are not experienced in blood collection techniques, training is recommended. If you have questions or comments about any of the above techniques, contact an ACP veterinary technician at vetservices@acp.ucsd.edu.

 

Total blood volume = 6% of lean body weight.Maximum blood collection = 10% of total blood volume every 2 weeks. Example: 250 gm rat = 0.25 kg x 0.06 = 15 ml x 0.10 = 1.5 ml

 

 

Collection Site	Advantages	Disadvantages
Lateral tail vein	·         Vein is easily accessed ·         Anesthesia not required ·         Yields moderate quantities	·         Specialized equipment is needed ·         Animal must be securely restrained
Ventral Tail Artery	·         Large quantities of blood can be collected	·         Anesthesia is required ·         Requires training
Orbital Sinus or Plexus	·         Large volume of blood can be collected	·         Anesthesia is required ·         Requires training
Lateral Saphenous Vein	·         Large quantities of blood can be collected ·         Anesthesia not required ·         Excellent technique for serial sampling	·         Requires training
Anterior Vena Cava	·         Large quantities can be collected	·         Requires anesthesia ·         Requires specialized training
Cardiac Puncture	·         Maximum volume of blood can be collected	·         Requires deep anesthesia ·         Non survival procedure

 

 

Terminal Blood Withdrawal

 

Terminal bleeds are only allowed on animals under general anesthesia, and the animal's death must be verified at the end of the bleed. An alternative euthanasia method is recommended after the blood withdrawal.

A general rule: An animal's blood volume is 10% of its body weight, and only about half of that can be recovered when the animal is bled out. Therefore, as a terminal bleed, 5-6 percent of an animal's body weight is a reasonable amount of blood (ml) that may be collected at exsanguination.

Section 17. Administration of Anesthetic, Analgesic, Therapeutic and Experimental Agents

 

This section allows you to list all administered substances by adding lines as need. The IACUC is interested in:

 

·          The possible adverse effects to animal health and welfare from administered test agents.

·          The anesthetics and analgesics that will be used to minimize pain and distress

 

Standard of Care: Anesthesia and Analgesia

 

Animal anesthesia, analgesia and pain management are crucial components of the animal use protocol.

 

The standard of care at UCSD is to prevent animal pain whenever possible and to treat animal pain whenever diagnosed. Exceptions to these principles are permitted only in the small number of protocols approved by the Institutional Animal Care and Use Committee as USDA Category E.

 

Multi-modal anesthetic and analgesic regimens combine drugs from a variety of classes. They are designed to maximize the desired effects while minimizing those unhealthy side effects that occur with over-reliance on a single agent. 

 

The ideal anesthetic/analgesic regimen

 

The ideal anesthetic/analgesic regimen must balance several goals:

1. It should provide pre-emptive analgesia so that animal pain is already being treated as the general anesthetic is wearing off, to prevent sensitization (“ramp-up”) of pain sensory mechanisms, and to lower the overall amount of general anesthetic required for the procedure.
2. It should be precisely titratable to assure that animals receive adequate anesthesia to block pain sensation, to produce unconsciousness, and to produce immobility without experiencing hemodynamic instability or life-threatening anesthetic overdoses.
3. It should not interfere with the study that the animals are on.
4. It should not result in unhealthy post-operative side-effects.
5. It should not cause pain or distress on induction or recovery
6. It should be compatible with available equipment and available medication

To meet the first goal, ACP & the IACUC advocate pre-emptive analgesia, using some combination of four drug classes 30 minutes or more prior to surgery. The four classes of injectable drugs are:

• opioid analgesics (such as buprenorphine)
• non-steroidal anti-inflammatory drugs (such as carprofen, meloxicam, ibuprofen)dissociative anesthetic/sedatives (ketamine, tiletamine)
• local/regional anesthetics (lidocaine, bupivicaine, proparacaine)

 

 

Volatile anesthetics (isoflurane, halothane, sevoflurane) delivered via a precision vaporizer best meet the second goal. Adjusting the inhaled percentage of anesthetic gas to deepen anesthesia is far safer than repeated redosing of injected drugs. Volatile anesthetics are easier to decrease as well, even compared to drugs for which there is an injectable antagonist or reversal agent. A major shortcoming of the inhalant anesthetic agents is the lack of residual analgesia once the vaporizer has been turned off.  

 

The very best anesthetic plans are only as good as the skill and care with which they are applied. Training is available from the Animal Care Program training and veterinary staff. Veterinary consultation (jfujimoto@ucsd.edu) is encouraged when planning any potentially painful study (and is required by law for USDA-covered species).  

 

 

Drug dosages and frequencies of administration

 

Drugs should be listed in the protocol with approximate dose ranges. These are starting points which must be titrated up or down for the individual animal, or for the particular application (procedures conducted, animal age and strain differences). When laboratory experience finds that recommended dose ranges are consistently too high or too low for the particular application, the veterinarian should be informed, and a protocol modification submitted to the Institutional Animal Care and Use Committee.

Anesthetics are always titrated to effect. It is not acceptable to conduct surgical procedures unless the animal is fully anesthetized.

Analgesic doses and frequencies are more difficult to gauge. Caution is required for overnight pain management. Most analgesics administered at 5 pm will not still be effective at 8 am the next morning. Newer, longer-lasting non-steroidal anti-inflammatory analgesics may have longer durations of action than available opioids; they are frequently co-administered with an opioid to combine potency of effect with duration of action.  

 

 

Safe and effective animal anesthesia

 

General guidelines on use of anesthetics and analgesics are posted. Plans for intra- and post-operative monitoring must be included in the IACUC protocol application, and then practiced as written.

Animals should be acclimated to their surroundings for several days prior to major procedures.

 

Supplemental administration of warmed fluids (lactated ringers solution or isotonic saline) and maintenance of body temperature will improve anesthetic safety for the animals.

 

 

Commonly used anesthetics and analgesic agents 

 

Inhalant agents

 

Isoflurane and Halothane

 

The standard inhalant anesthetics for laboratory animal use are either isoflurane or halothane, delivered to effect in concentrations of 1-3% in oxygen (up to 5% for initial induction), using a precision vaporizer. Advantages: Advantages of inhalant agents include rapid induction and recovery, with the ability to precisely titrate the level of anesthesia. Disadvantages: Disadvantages include the cost and logistics of using precision vaporizers, occupational exposure concerns, the risk of fatal overdosage if an open system is used instead of a precision vaporizer, and depressed respiratory rate and decreased blood pressure. In addition, once animals awaken from gas anesthesia, there is no residual analgesic activity. Concurrent use of ketamine combinations and/or opioid and/or non-steroidal anti-inflammatory analgesics is strongly encouraged if the procedure is likely to result in any residual pain.Several individual laboratories have their own isoflurane vaporizers, and the Animal Care Program maintains several vaporizers for laboratory use. Occupational safety is a serious concern. Inhalants must be directly vented out of the room, or (less reliable), adsorbed in a charcoal canister filter. Filters must be weighed and replaced before they reach target weight (usually an increase of 50 gm).  

 

Nitrous Oxide (N₂O)

 

May be used 50:50 or 60:40 with oxygen as carrier gas for inhalant anesthetics such as Isoflurane. Nitrous oxide is not acceptable as sole anesthetic agent for surgery, but it may lower the required dose of inhalant.

 

Other inhalant agents

 

Other agents and techniques may be used for inhalant anesthesia, only when specifically approved by the IACUC in the animal use protocol.

Methoxyflurane is useful for open-system use in rodents (inside an appropriate fume hood). It is not currently readily available in the United States.

"Open-drop" inhalant anesthesia is acceptable with rodents only for some very short procedures. Diluting halothane in oil may make this option safer for the animals.

Ether is an irritant and a fire hazard, and its use is generally not allowed unless approved by EH&S and IACUC.

Carbon dioxide is a potent anesthetic, but concentrations are difficult to control, making the margin of safety unacceptably low

Nitrous oxide is a less potent anesthetic/analgesic gas in most animals than it is in people. It can be used up to 50% in oxygen as a carrier gas for inhalant agents such as isoflurane and halothane, and may thereby reduce the required concentration of the other agent required. Occupational exposure is potentially dangerous so direct venting is required (charcoal filters do not absorb nitrous oxide).

 

Dissociative anesthetics

 

Ketamine & Tiletamine

Ketamine is a widely used anesthetic in a variety of species. In low doses, ketamine provides chemical restraint with some analgesia. In higher doses, it may provide short-term surgical anesthesia in some species. In most instances, ketamine is used in combination with other injectable agents. Tiletamine is similar to ketamine; it is primarily used in combination with zolazepam as the drug Telazol. Advantages of Ketamine: Advantages of ketamine are its wide margin of safety in most species and its analgesic action. In combination with other drugs, it can provide surgical plane of anesthesia for about one half hour. Disadvantages of Ketamine: Disadvantages of ketamine include some irritancy due to low pH, and insufficient anesthesia in some species and strains (especially mice) for some procedures. Ketamine is a Class III controlled substance. Advantages of Telazol: A low volume of injection is required. Like ketamine combinations, it can occasionally produce short-term anesthesia, though rarely of sufficient depth for surgery. It is more useful as an induction agent prior to general inhalant anesthesia, or for chemical restraint for short non-surgical procedures. Disadvantages of Telazol: Telazol must be stored under refrigeration once reconstituted. It is not safe for use in rabbits (kidney disease). Telazol is a Class III controlled substance.

Ketamine combinations  Ketamine-a2-agonists (Xylazine or Medetomidine)

Ketamine may be combined with the a2-agonists Xylazine or Medetomidin the same syringe to produce a deep level of sedation. In some situations in some species and strains an adequate depth of anesthesia for surgery may be attained. In other cases, this sedation may require an inhalant agent to achieve surgical anesthesia. It is generally safer to titrate to effect with inhalant anesthetic from a precision vaporizer than with supplemental injections of ketamine. Advantages: Advantages of ketamine-a2-agonist combinations are that they may be combined in one syringe, that they may produce short-term surgical anesthesia with good analgesia, and that recovery can be hastened by reversing the a2-agonist with Atipamezole or Yohimbine. Disadvantages: Disadvantages of ketamine-a2-agonist combinations are that they will not reliably reach surgical anesthesia in all cases, and that they can cause profound cardiac depression. Xylazine may cause vomiting, especially in cats. Ketamine is a Class III controlled substance. Caution for use: if a ketamine a2-agonist combination is used for surgery longer than 20 minutes, animals will likely require additional anesthetic. Redosing with ketamine rather than the combination is usually safer, as the cardiovascular depression of a2-agonists is often longer-lasting than the sedation or analgesia produced. Adding acepromazine to the ketamine-a2-agonist combination may result in deeper and/or longer plane of anesthesia in small rodents, especially rats, and possibly some strains of mouse as well.

Ketamine-benzodiazepines (Midazolam or Diazepam)

Ketamine may be combined with the benzodiazepines Midazolam or Diazepam in the same syringe to produce a deep level of sedation. In most cases, this sedation will require an inhalant agent or other anesthetic to achieve surgical anesthesia. In most applications, Midazolam is preferred, as it can be injected intramuscularly; intramuscular injection of propylene glycol (the carrier in injectable diazepam) can cause painful, sterile abscesses and is discouraged. Advantages: Advantages of ketamine-benzodiazepine combinations are that they may be combined in one syringe and will produce deep sedation with moderate analgesia as well as amnesia. Recovery from ketamine-midazolam is often smoother than recovery from ketamine alone. Disadvantages: Disadvantages of ketamine- benzodiazepine combinations are that they will not reliably reach surgical anesthesia in most cases. Diazepam should be restricted to intravenous or intraperitoneal use. Ketamine is a Class III controlled substance while the benzodiazepines are in Class IV. Pharmacologically, Telazol is a dissociate-benzodiazepine combination.

Barbiturates

 

Though superseded in most applications by newer anesthetic, barbiturates still have their place in the animal laboratory. They are most frequently used in terminal or acute studies, as recovery can be prolonged and unpleasant, especially in larger animals. Barbiturates are often the anesthetic of choice when neurophysiological recordings are being conducted, such as visual or auditory evoked responses. Concurrent use of an analgesic (opioid or non-steroidal anti-inflammatory drug) is encouraged as it may improve pain relief with barbiturate use, and lower the required dose of barbiturate.

Sodium pentobarbital (Nembutal) and sodium thiopental (Pentothal) are currently the two most commonly used barbiturates. The duration of action of pentobarbital is considerably longer than that of thiopental. Advantages: Barbiturates do not depress cortical evoked responses to the extent that other anesthetics might. Animals do not feel pain when they are at a surgical plane of anesthesia. Once stable anesthesia has been achieved, it may be longer lasting than with most other injectable agents. Barbiturates are the most common of the injected euthanasia solutions, as they reliably produce unconsciousness before respiratory depression and death. Disadvantages: Disadvantages of barbiturates include a narrow margin of safety, primarily associated with respiratory depression. Pain sensation is only decreased at surgical planes of unconsciousness, and may even be heightened (hyperalgesia) at subanesthetic doses. Larger animals may experience a distressful anesthetic recovery. Outside of the vein (perivascular, or intraperitoneal) barbiturates can be irritating; barbiturates for IP injection should be diluted to a strength of 6 mg/kg. Barbiturates are Class II controlled substances, except for some Class III euthanasia solutions

 

 a2-agonists (Xylazine or Medetomidine)

The a2-agonists (Xylazine or Medetomidine) are hypnotic analgesics with significant pain relief. Used as sole agents, they do not produce sufficient depth of anesthesia for even minor surgical procedures. Combined with ketamine, and possibly supplemented with inhalants or local or topical analgesics, they may be useful during surgery. In some species, medetomidine appears to lead to greater anesthetic depth than does xylazine, and it is more reliably antagonized by atipamezole. Advantages: a2-agonists are that they produce profound analgesia of short duration, can be combined with ketamine (and in rodents, acepromazine) to produce deeper anesthesia, they are not controlled substances, and they are reversible with IP or subcutaneous atipamezole (yohimbine is sometimes used for xylazine reversal). They are not irritant when injected via intramuscular or intraperitoneal routes. Disadvantages: Disadvantages in most species include cardiovascular depression (decreased heart rate, decreased cardiac output, and hypotension), which is somewhat controlled by use of atropine or glycopyrrolate. a2-agonists cause a transient hyperglycemia which may have research implications. Xylazine often causes transient nausea and vomiting, especially in cats. Rapid IV administration of reversal agent has produced seizures in some species. Caution for use: If a ketamine a2-agonist combination is used for surgery longer than 20 minutes, animals will likely require additional anesthetic. Redosing with ketamine rather than the combination is usually safer, as the cardiovascular depression of a2-agonists is often longer-lasting than the sedation or analgesia produced.

Propofol

 

Propofol can produce general anesthesia in animals, as a sole agent with continuous infusion for surgery, or as a pre-anesthetic for endotracheal intubation. It is valued for its fast recovery time, even after prolonged administration. Advantages: Animals recover from propofol in minutes, even after prolonged administration. Disadvantages: Propofol has minimal analgesia at sub-anesthetic doses. It can be a profound respiratory depression, and may also cause hypotension. Because of its rapid elimination, it must be administered IV, and so is of limited use in small rodents. Unused propofol from an opened ampule should be discarded after use and not stored for future use.

 

Opioids

 

Opioid drugs are important components of many surgical anesthesia regimens, and are the most potent available post-procedural analgesics. Drugs in this group vary in their potency as well as their duration of action. Fentanyl, oxymorphone, buprenorphine and butorphanol are the most commonly used opioids in laboratory animal care, though others may be used on occasion. Fentanyl is the most potent of the three, but also the shortest acting. Buprenorphine is longer-acting and is good for most post-operative applications. Butorphanol may be more efficacious than buprenorphine for birds and for cats. Buprenorphine and butorphanol are mixed agonist/antagonists at different opioid receptors; they produce a less profound respiratory depression than full agonists, but also have a “ceiling effect” in the degree of analgesia produced with increasing doses. Opioids are most often administered by injection. Oral use is effective, but requires much higher doses because of "first-pass" liver metabolism when absorbed from the gut. Pre-emptive analgesic use is strongly recommended - buprenorphine may be administered when the general anesthetic is administered, or at any time during surgery. Respiratory depression is minimal, though sleep time may be lengthened. Pre-emptive use enhances pain management during the immediate post-surgical period. Though it increases animal handling (a stressor), administration of the analgesic 30 minutes prior to the initial surgical incision maximizes the analgesic efficacy in most situations. Advantages: Opioids are potent analgesics. Concurrent use with inhalant or barbiturate general anesthesia will lower the required dose f the anesthetic. Disadvantages: Opioids can suppress respiration (more marked effect in fentanyl than in buprenorphine). Opioids may increase locomotor activity, and may cause pica (abnormal ingestion of non-food items such as bedding) in rats. Alternatively, they may sometimes cause sleepiness and slower recovery from general anesthesia. Fentanyl has a very short duration of action in most animal species. Opioids are controlled substances. Cautions for use: Buprenorphine has found favor as the longest-acting opioid analgesic. However, this duration of action is closer to 6 hours in most situations than it is to 12 hours. 12 hours is the absolute maximum dosing interval for use of buprenorphine for post-procedural pain.

Non-steroidal anti-inflammatory drugs (NSAIDs)

The advent of newer, more potent, more specific anti-inflammatory agents has increased their usefulness in laboratory animal use. Most reduce fever, reduce inflammation, and provide varying degrees of analgesia (acetaminophen does not significantly reduce inflammation). Advantages: Carprofen, ketoprofen, ketorolac, and meloxicam may have duration of analgesic action up to 24 hours. They may be used concurrently with anesthetics, with opioid analgesics, and with local anesthetic/analgesics. Injectable NSAIDs are useful for accurate dosage and administration to small rodents. Oral flavored analgesics are useful for mild pain in nonhuman primates. They are not controlled substances (some are by veterinary prescription only, and must be obtained through Laboratory Animal Resource Center Disadvantages: NSAIDs may decrease clotting ability, of possible concern following surgery. Gastric upset and even ulceration may occur, especially with prolonged use. Prolonged use carries the risk of kidney or liver disease. Cautions for use: Cats are particularly susceptible to toxic effects of NSAIDs. Acetaminophen is never administered to cats; other NSAIDs should be used only at the dose and frequency recommended. Undesired side effects are more likely with increasing length of usage - for most situations, limit use of NSAIDs to 3-4 days per animal, except under veterinary supervision. Do not use in dehydrated animals, or in animals with kidney or liver dysfunction.

 

Local anesthetic/analgesic drugs (lidocaine and bupivicaine)

 

Local anesthetic/analgesic drugs (lidocaine and bupivicaine) may be useful both during surgery, and post-operatively. They block nerve conduction when applied locally at sufficient concentration. Lidocaine has a fast onset of action, and provides a couple of hours of analgesia. Bupivicaine has a slower onset of action (up to 30 minutes) but provides up to 12 hours of residual analgesia. Both are infiltrated subcutaneously at the surgical site, or (especially in larger animals) may be used regionally (epidural, intrathecal, intercostal). Lidocaine cream (EMLA or ELAMax) is used topically on shaved, intact skin prior to venipuncture, though it requires 30-60 minutes or more of contact with skin to reach full effect. Tricaine methanesulfonate (MS-222) is a related compound used as a general anesthetic for fish and frogs. Advantages: Intra-operative use can augment the pain relief of general anesthetics, and reduce the need for frequent redosing. Bupivicaine can augment the post-operative analgesic action of opioids and/or NSAIDs. They are not controlled substances. At appropriate doses, they have minimal cardiovascular effect. Disadvantages: Intramuscular and intravenous injection should both be avoided. Systemic toxicity (including seizures and death) can result from overdosage (more likely to occur with smaller subjects) and with accidental intravenous injection. Lidocaine may sting when first injected.

 

 Miscellaneous agents

Urethane, choral hydrate, equithesin, sodium thiamylal, a-chloralose have some specialized use in laboratory animal anesthesia. Their use should be discussed with an ACP veterinarian as you are writing your protocol.

Section 18. Paralyzing Agents

When neuromuscular blocking (NMB) drugs are used on animals in experimental research, it is critical that the animals be adequately anesthetized.  NMB drugs paralyze muscles, thereby making it possible for animals to experience pain or discomfort without being able to move or withdraw from a painful stimulus.  The following guidelines for the use of NMB drugs are designed to paralyze muscles for the shortest time possible, once adequate dose or depth of anesthesia has been assured.

 

INDICATIONS

NMB drugs may be used for specific indications, such as preventing animal motion:

i)      during surgery which is technically delicate

ii)    during physiological recordings where muscle activity would seriously impair the precision of the measurements.

 

NMB drugs are not indicated for routine endotracheal intubation, positive pressure ventilation, or abdominal or thoracic surgery.

 

When it is unclear if a NMB is necessary for a project, the investigator should consult with the UCSD veterinarians at the Animal Care Program.  In some instances a pilot procedure may be required following committee approval to determine whether these drugs are necessary for the study and what an adequate depth of anesthesia would be.  An alternative to the pilot procedure is a modification in the experimental design; e.g., where surgery/instrumentation is performed without NMB drugs, and data collecting post-instrumentation is performed under an adequate level of anesthesia with NMB drugs.

 

DEPTH OF ANESTHESIA

The most important determination is whether the depth or dose of anesthesia is adequate for a given type of surgery or investigative procedure.  Since animals will not be able to move in response to pain, their blood pressure (BP) and heart rate (HR) must be carefully monitored and the anesthetic dose adjusted appropriately.  Both the upper and lower range of acceptable BP and HR values need to be specified in the proposal.  The parameters of any other signs (e.g., pupil size) which might be used for depth assessment also need to be specified.  These values should be determined before NMB drug administration; e.g., during preparatory surgery, pilot studies, or from previous experimental procedures not using NMB drugs.

 

A detailed anesthesia record must be completed for each investigative or surgical procedure utilizing NMB drugs.  The anesthesia record should document:  i) timing and dose of anesthetic and NMB drug administration, ii) vital signs and depth of anesthesia signs obtained during the procedure, and iii) neuromuscular response to the peripheral nerve stimulator (see Paragraph 5 below).  A copy of each anesthetic record must be maintained by the Principal Investigator, and should be readily available for review or inspection at all times.

 

ADMINISTRATION

The kind and quantity of dose of NMB drugs should be chosen to provide only brief periods (e.g., 20 minutes) of neuromuscular blockade, so that adequacy of anesthesia (lack of movement in response to a noxious stimulus) can be assured on a frequent and regular basis.

 

MONITORING NEUROMUSCULAR BLOCKADE

When NMB drugs are used for experimental procedures in which NMB drugs are used for more than a single short event, peripheral nerve stimulation (PNS) tests are recommended to document that neuromuscular function has recovered during the non-paralyzed periods of anesthesia-depth assessment.  Note:  If animal Welfare are to recover from the experimental procedures, clinical signs of neuromuscular recovery should complement the above nerve stimulation tests (e.g., normal breathing pattern, sustained head lift, or ability to stand). SPECIAL EQUIPMENT

 

The proper use of the following equipment is required:

1.      mechanical ventilator  

2.      reliable means of quantifying heart rate and blood pressure

 

PROTOCOL APPROVAL

Approval of an Animal Use Protocol with the use of NMB drugs depends on whether the above provisions have been satisfied, and necessitates a detailed valid scientific justification.

Section 19. Surgery

All surgical procedures in animals must be pre-approved by the IACUC. The campus veterinarian is required to provide guidance or oversight to surgery programs and post-surgical care. Both the IACUC and Campus Veterinarian are required by federal law to suspend animal activities not in compliance with federal regulations.

Definitions

 

Survival surgery: a surgical procedure from which an animal is expected to regain consciousness.

 

Non-Survival Surgery: a surgical procedure from which an animal is euthanized before regaining consciousness.

 

Major Survival surgery: a surgical procedure that penetrates and exposes a body cavity or produces substantial impairment of physical or physiologic functions. Examples include laparotomy, thoracotomy, craniotomy, orthopedic procedures, limb amputation and enucleation.

Minor Survival Surgery: a surgical procedure that does not expose a body cavity and causes little or no physical impairment. Examples include wound suturing, peripheral vessel cannulation, and placement of subcutaneous implants.

 

 

General Responsibilities of Principal Investigators  

 

1. Assuring adequately trained surgical staff:

 

Each person involved with a study needs appropriate training to adequately perform the duties required. Training can be provided by the laboratory or through classes offered by ACP or individual Principal Investigators.

 

Adequate surgical training must be provided to ensure that good surgical technique is practiced including asepsis, gentle tissue handling, minimal dissection of tissue, appropriate use of instruments, effective hemostasis and suturing techniques. Individuals trained in human surgery may need additional training in other species’ variations in anatomy, physiology, the effects of analgesic and anesthetic drugs, or in postoperative procedures. The Guide states that, "The PHS Policy and the AWRs place responsibility with the IACUC for determining that personnel performing surgical procedures are appropriately qualified and trained in the procedures to be performed."  This is accomplished by Committee assessment of qualifications of personnel submitted through Personnel Qualifications forms.  As such, only people listed on an approved protocol and specified as participants in surgical procedures may participate in those procedures. The protocol application process includes a place to list personnel who will perform surgical procedures as well as space for describing each person’s qualifications and training. Adding additional surgical personnel must be approved by amendment to an approved protocol before participation in surgery.

 

 2. Performing procedures as approved by the IACUC:

 

All surgical procedures to be performed on animals at UCSD must be described in an animal protocol and approved by the IACUC before being performed on animals. The description of these procedures is part of the protocol application process. Post-surgical care must be addressed in the protocol, including where animal recovery will take place, how often the animals will be monitored, and the criteria for cessation of monitoring. Changes to an approved surgical procedure must also be approved by the IACUC before implementation.

 

 Policies and Guidelines for Rodent Surgery

Specific Responsibilities of Principal Investigators:

 

1. Assuring appropriate presurgical planning, presurgical evaluation and preparation

 

Presurgical planning is critical to the success of surgical procedures in animals. Presurgical planning should include input from all members of the surgical team. The surgical plan should identify personnel, their roles and training needs; equipment and supplies required for the procedures planned; the location and nature of the facilities in which the procedures will be conducted; and preoperative animal health assessment and postoperative care.

Appropriate pre-surgical evaluations are required to assure that the prospective patient is in good health. The withholding of food is not necessary in rodents unless specifically required by the protocol or surgical procedure. Water should not be withheld unless required by the protocol. Withholding of food for more than six hours should be discussed with a veterinarian.
 

2. Performing all surgical procedures in facilities approved by the IACUC

 Survival Surgery can be performed in a small, dedicated space that allows for the minimization of contamination from other activities in the room during surgery. In general, rodent survival surgery should be performed in the vivarium in order to decrease the risk of catastrophic contamination of animals colonies associated with transport of animals out of and back into the vivarium. While survival surgical procedures can performed outside the vivarium, this practice depends on policies associated with specific vivaria. Facilities for rodent survival surgery must be specified and approved by the IACUC in the animal use protocol.

 

Non-Survival Surgery can be performed outside of approved surgical suites but must be specified and approved by the IACUC in the animal use protocol.
 

3. Performing surgical procedures with appropriate surgical technique

Survival Surgery

Modified Aseptic Technique includes:

1. Shaving or hair removal in the operative field unless it can be documented that this is not indicated.
2. Preparation of the surgical site with an antiseptic solution such as povidone iodine. It is recommended that instruments be maintained sterile throughout all procedures. At a minimum, instruments must be sterile at the start of the first surgery of a daily series of surgeries. Accepted methods of instrument sterilization.
3. The use of exam or surgical gloves. Gloves should be cleaned (disinfected) between each animal.
4. Sterilization of implanted devices (catheters, osmotic pumps, etc.).
5. All rodent surgery must be performed in an area dedicated for that purpose.

Non-Survival Surgery

For studies in which local bacterial contamination of tissues, or sepsis, could influence study outcomes, standard aseptic technique is recommended. Such studies include long term (>8 hours) terminal surgeries and collection of samples for tissue or microbial culture.

 

At a minimum, it is recommended that lab personnel wear dedicated clothing (e.g. lab jacket), gloves, and other appropriate personal protective equipment when euthanizing animals for tissue collection. In addition, the area used for tissue collection should not be used for other purposes during the tissue collection and the area should be cleaned and disinfected after each use.
 

4. Providing proper anesthesia and analgesia

Animals must be properly anesthetized for all surgical procedures as required by federal regulations. Consultation or assistance is available through the ACP (Caution: exposure to waste anesthetic gas can result in occupational disease for people who are overexposed to these chemicals. Contact EH&S for assistance).

 

The extent of monitoring necessary during surgery depends upon the anesthetic regimen, the animals' clinical condition, and the invasiveness of the procedure and may need to include body temperature, heart, and respiratory rates.

 

Body temperature should be maintained (for non-survival surgery lasting at least 45 minutes, body temperature should also be maintained).

 

External heat sources, such as recirculating water blankets or "Delta Phase Pads" (Braintree Scientific) are recommended. Heat lamps and commercial household-type heating pads are not permitted.

5. Providing appropriate post-surgical care and analgesia

 

It is the responsibility of the principal investigator to assure that their animal receives adequate post-surgical care. This care begins with recovery from anesthesia and may extend for days or weeks depending on post-surgical outcome and study design. The post-surgical care of animals encompasses weekends and holidays, and should be performed by trained individuals.

 

Continuous and careful observation of animals immediately postoperatively requires systematic monitoring of thermoregulation, cardiovascular and respiratory function, and postoperative pain and discomfort.

 

During recovery from anesthesia, the administration of supportive fluids, analgesics, and other drugs may be indicated. Animals should not be left unattended until they are physiologically stable, conscious, and able to maintain sternal recumbency without assistance.

After anesthetic recovery, monitoring is often less intense but should include attention to basic biologic functions of intake and elimination and behavioral signs of postoperative pain, monitoring for post-surgical infections, monitoring of the surgical incision, bandaging as appropriate, and timely removal of skin sutures.

Appropriate medical records must be maintained and any complications or concerns should be reported to the veterinary staff of the Animal Care Program.

6. Maintaining appropriate documentation of activities associated with surgery

P.I. must assure documented observations daily for five days. If problems occur at a later date in the study, the animal must be observed daily until resolved. If permanently instrumented, after the initial 5 days, observations will be made weekly until removal of the instrument of final disposition of the animal.

 

Policies and Guidelines for all other Mammalian Surgery

 

 

Specific Responsibilities of Principal Investigators:

 

1. Assuring appropriate presurgical planning, presurgical evaluation and preparation

 

Presurgical planning is critical to the success of surgical procedures in animals. Presurgical planning should include input from all members of the surgical team, including the surgeon, anesthetist, veterinarian, surgical technicians, animal care staff, and investigator. The surgical plan should identify personnel, their roles and training needs; equipment and supplies required for the procedures planned; the location and nature of the facilities in which the procedures will be conducted; and preoperative animal health assessment and postoperative care.

Appropriate pre-surgical evaluations are required to assure that the prospective patient is in good health. All animals scheduled for any survival surgical procedure must receive a preoperative physical evaluation by a qualified person. This may range from brief observations to identifying specific laboratory baseline values and must be documented.

 

It is advantageous in many studies to introduce the animals to your laboratory setting for training or for habituation to a restraint device before beginning the project. Investigators must assure that animals are appropriately fasted before administration of general anesthetics and elective surgical procedures.

 

Non-Human Primates: Food should generally be withheld 8 -12 hours before anesthesia in adult non-human primates.

Ruminants: Food should generally be withheld 24 - 48 hours prior to anesthesia and water 12 - 24 hours prior to anesthesia in adult ruminants.

Dogs: Food should generally be withheld 8 -12 hours before anesthesia in adult dogs.Pigs: Food should generally be withheld 8 -12 hours before anesthesia in adult pigs. 

 

2. Performing all surgical procedures in facilities approved by the IACUC

 

Survival Surgery must be conducted only in facilities intended for that purpose as defined by the NIH Guide and approved by the IACUC. The following sites are currently approved Survival Surgery Areas at UCSD: BSB B104, CTF A050 and C310, SRL 106 and 114, EFS CVL, and the SWC surgery suite. Investigators with special circumstances may request a variance from the IACUC.

 

Non-Survival Surgery can be performed outside of approved surgical suites but must be specified and approved by the IACUC in the animal use protocol. 

 

3. Performing surgical procedures with appropriate/adequate surgical technique

 

Survival Surgery

 

All survival surgery must be performed using aseptic technique. Aseptic technique encompasses a number of practices and procedures to reduce microbial contamination of surgical sites to the lowest possible practical level. In general, survival surgical procedures should be performed using the same aseptic practices common in veterinary and human medicine. The guidelines below are to be followed when performing survival surgery:

 

Surgeon:

Preparation and attire of the surgeon includes the following:

Surgical scrub/hand wash
Standard surgical attire: mask, shoe covers, cap, sterile gloves and gown. 

 

Animal Preparation:

The incision site and an adequate surrounding area should be prepared by removing all hair. If the hair is excessively dirty, the animal may require bathing before the surgical procedure. Usually after anesthesia has been induced, the hair is removed by clipping with electric clippers. A razor or depilatory cream may also be used.

The skin should then be cleaned and disinfected. A chlorhexidine or iodine-based detergent (e.g. Povidone, Betadine) and a sterile gauze sponge can be used to scrub the area. The surgical site should always be cleaned by scrubbing along the proposed incision line and then proceeding outward. The sponge should never be brought from the contaminated edge of the surgical area back into the clean center.

 

An ophthalmic lubricating ointment may be applied to the eyes to prevent drying.

 

The use of sterile surgical drapes is required.

 

Instruments: 

All instruments that come into direct contact with the surgical area must be sterile.  Sterilization of instruments can be achieved in a number of ways:
a) Steam (autoclave)
b) Chemical sterilants (Note, alcohol is not a sterilant.)
c) Ethylene oxide (arranged through the UCSD School of Medicine. Ethylene Oxide is a regulated chemical carcinogen; any use at UCSD must be approved by Environment, Health, and Safety)

 

If surgeries are to be performed on consecutive animals, surgical instruments must be sterilized between animals. This can be achieved by using multiple surgical packs, chemical sterilants, or use of a hot bead sterilizer.

 

Non-Survival Surgery

 

For studies in which local bacterial contamination of tissues, or sepsis, could influence study outcomes, standard aseptic technique is recommended. Such studies include long term (>8 hours) terminal surgeries and collection of samples for tissue or microbial culture.

At a minimum, it is recommended that lab personnel wear dedicated clothing (e.g. lab jacket), gloves, and other appropriate personal protective equipment when euthanatizing animals for tissue collection. In addition, the area used for tissue collection should not be used for other purposes during the tissue collection and the area should be cleaned and disinfected after each use.

 

4. Providing proper anesthesia and analgesia

 

Animals must be properly anesthetized for all surgical procedures as required by federal regulations. Consultation or assistance is available through the ACP (Caution: Exposure to waste anesthetic gas can result in occupational disease for people who are overexposed to these chemicals. Contact EH&S for assistance).

 

The extent of monitoring necessary during surgery depends upon the anesthetic regimen, the animals' clinical condition, and the invasiveness of the procedure and may need to include body temperature, heart, and respiratory rates. The frequency of monitoring must be stated in the protocol and documented during the procedure. Such monitoring may need to be continuous at 5-15 min intervals. Other necessary variables may include blood pressure, pulse quality, urine output, hematocrit, ECG, blood gases and pH.

 

Therapeutic drugs administered during the procedure must be recorded.

 

Intravenous parenteral fluids must be administered when required to maintain normal hydration in invasive surgical or extended anesthesia cases.

 

Body temperature should be maintained (for non-survival surgery lasting at least 2 hours, body temperatures should also be maintained). External heat sources, such as recirculating water blankets or "Delta Phase Pads" (Braintree Scientific) are recommended. Heat lamps and commercial household-type heating pads are not permitted. 

 

5. Providing appropriate post-surgical care and analgesia

 

It is the responsibility of the principal investigator to assure that their animal receives adequate post-surgical care. This care begins with recovery from anesthesia and may extend for days or weeks depending on post-surgical outcome and study design. The post-surgical care of animals encompasses weekends and holidays, and should be performed by trained individuals.

Continuous and careful observation of animals immediately postoperatively requires systematic monitoring of thermoregulation, cardiovascular and respiratory function, and postoperative pain and discomfort.

 

Post-anesthetic monitoring guidelines:

• During recovery from anesthesia, the administration of supportive fluids, analgesics, and other drugs may be indicated.
• Animals should not be left unattended until they are physiologically stable, conscious, and able to maintain sternal recumbency without assistance.
• After anesthetic recovery, monitoring is often less intense but should include attention to basic biologic functions of intake and elimination and behavioral signs of postoperative pain, monitoring for post-surgical infections, monitoring of the surgical incision, bandaging as appropriate, and timely removal of skin sutures.
• Appropriate medical records must be maintained and any complications or concerns should be reported to the veterinary staff of the Animal Care Program.

 

6. Specific Requirements for Survival Surgical Procedures

 

All animals undergoing surgical procedures must have documented observations daily until suture removal and adequate healing of the surgical site (generally 7-10 days). After adequate healing has occurred, documented observations will be made according to the approved animal use protocol, with a minimum of weekly documented observations.

The Principal Investigator or a designated staff member must observe animals that exhibit pain, distress, discomfort, handicap, or other health complications daily during the period of time that the problem exists.

Any procedure that involves exposed instrumentation, catheterization, bandage, etc. must be observed daily until removal of the apparatus and adequate healing of associated sites has occurred. Externalized catheters or instrumentation must be covered or contained to prevent damage that might result in exsanguinations. Such dressings must be kept clean.

Animals with health complications or at significant risk of postoperative infection should have body temperature monitored and documented during these periods. Animals with externalized instrumentation should have body temperature monitored and documented according to the frequency described in the protocol.

Analgesia must be provided as described in the approved IACUC protocol. If an animal exhibits unalleviated pain or distress not approved by the IACUC, veterinary staff consultation is required.

 

 7. Maintaining appropriate documentation of activities associated with surgery

Post-Surgical Records

The post-surgical record is composed of three specific forms: Post-Anesthetic Recovery Record, Daily Progress Notes, and Daily Observation Sheets. All three forms must be used with all non-rodent mammalian species with the exception of rabbits, which do not require a Daily Observation Sheet. The investigator is responsible for maintaining only two of the forms (Post-Anesthetic Recovery and Daily Progress Notes). Animal husbandry staff completes the Daily Observation Sheets. The IACUC or veterinary staff must approve any modifications to these forms. Standard forms are available at the office of each animal facility.

Post-Anesthetic Recovery Record

Used on day of procedure only. Frequency of observations and physiologic parameters monitored and recorded are at the discretion of the investigator but should concur with the post-surgical care described in the Animal Use Protocol as well as with the UCSD Post-surgical Monitoring Guidelines Particular attention should be given to the periodic recording of thermoregulation, cardiovascular and respiratory function, as well as postsurgical pain or discomfort.

Daily Progress Notes

Two types:

    
• Daily Progress Notes - Page One
  Initiated on the day following the procedure
  Top of form must be filled out completely
• Daily Progress Notes - Continuation Page
  Initiated when Page One is complete

Healthy animals are observed a minimum of once a day. Frequency of observations of sick animals (those with medical problems or complications) depends upon the severity of the condition and the treatment regimen. After adequate healing has occurred, documented observations will be made according to the approved animal use protocol, with a minimum of weekly documented observations. It is recommended that body temperature of chronically instrumented animals be monitored on a regular basis. All observations of animals must be entered in the medical record.

Daily Observation Sheet

Top portion must be filled out completely by investigator staff. Sheet is maintained by the animal facility staff and is kept on a clipboard on the door of the animal's kennel or on the wall outside the room and be returned to the animal facility staff at the time of final disposition of the animal. If your protocol stipulates that animals are to be trained or habituated to a restraint device before surgery, these sessions should be documented.

 

Section 20. Food or Water Restriction

 

Food and/or fluid restriction may be required in order to achieve a variety of research and/or husbandry objectives. It has been shown that some methods of food and/or fluid restriction are physiologically and/or psychologically stressful and, if restriction is allowed to exceed acceptable levels, can be physically harmful to an animal.

 

1. Food and/or fluid restriction that deviates from the normal husbandry procedures should be described in the IACUC Protocol Application.

 

2. Withholding of food and/or water for a period of greater than 18 hours requires scientific justification and a literature search for alternatives.

 

3. The investigator is responsible for assuring that specially formulated diets are nutritionally adequate and palatable.

 

4. The investigator should plan to monitor parameters such as body weight, hydration status, body condition, and food consumption.

 

5. Restriction endpoints should be specified in advance. Examples of specific endpoints include:

a. Failure of growing animals to gain weight.
b. Loss of greater than 20% of the body weight of a mature animal.
c. Body condition score i.e. animal is underconditioned: (a) segmentation of vertebral column is evident; (b) dorsal pelvic bones are readily palpable. 

 

Conditioned Response Research Protocols:

a. The amount of food/fluid used should be the minimum level that will achieve the objective.
b. If food/fluid restriction is to be used in an experimental protocol, the method of restriction and scientific justification for its use should be clearly explained in the IACUC Protocol Application submitted for review and approval by the IACUC.
c. Restriction must be based on a measurable parameter such as percentage of ad libitum intake, percentage of body weight compared to a control animal (paired; ad libitum intake), or length of time access of food/fluid is allowed per 24 hours.
d. Consideration must be given to alternative methods and/or modifications to food and/or fluid restriction.
e. In order to make a knowledgeable determination of an appropriate level of food/fluid restriction, it is necessary to know what normal quantities of food or fluid are required for maintenance of the species. Life stage (growth, pregnancy, lactation, geriatric) and state of health must also be taken into consideration in determining maintenance requirements.
f. Unless scientifically justified, food and/or water restriction should be introduced incrementally to allow for physiological and psychological adaptation. 

 

Nutrition Studies:

a. The principal investigator is responsible for assuring the proper formulation and nutritional adequacy of these diets.
b. Specific arrangements for feeding and diet storage should be provided in the IACUC Protocol Application and arranged with the Animal Care Program.
c. These rations frequently vary in form and in palatability. The animals should be closely monitored to insure that an adequate diet is consumed. 

 

Pre-anesthetic Fasting:

a. For non-rodent and non-rabbit species, food may be withheld for up to 18 hours prior to an anesthetic procedure (i.e. overnight fasting). Water should be available during the overnight fast but may be removed in the morning on the day of surgery.
b. In most cases, pre-anesthetic fasting is not required for rodents and rabbits. However, specific surgical procedures may require an overnight fast and should be explained in the IACUC Protocol Application.

Section 21. Restraint

Restraint is the use of manual, mechanical, or chemical means to limit some or all of an animal's normal movement for such purposes as examination, collection of samples, and drug administration. Typically, animals are restrained for brief periods, usually minutes, in most research applications. Often animals can be trained, through use of positive reinforcement, to present limbs or remain immobile for brief procedures.

If restraint devices are required, they should be suitable in size, design, and operation to minimize discomfort or injury to the animal. If restraint devices are required, they need to be reviewed by the Institutional Animal Care and Use Committee (IACUC) and identified in the protocol with a description of the device, and the duration of restraint.

Prolonged restraint should be avoided unless it is essential for achieving research objectives. Approval by the IACUC requires scientific justification, a description of the restraint device, the duration of restraint, monitoring procedures and methods to minimize animal distress (e.g. acclimation to the device).

 

Important guidelines for restraint of any duration:

• When restraint devices are used, they should be specifically designed to accomplish research goals that are impossible or impractical to accomplish by other means or to prevent injury to animals or personnel.
• The period of restraint should be the minimum required to accomplish the research objectives.
• If possible, animals placed in restraint devices should be given training to adapt to the equipment and personnel.
• Provision should be made for observation of the animal at appropriate intervals.
• Veterinary care should be provided if lesions or illnesses associated with restraint are observed. The presence of lesions, illnesses, or severe behavioral change often necessitates temporary or permanent removal of the animal from restraint.

Section 22. Tumors/Neoplasia

Investigators producing tumors in rodents should use this document as a reference in preparing their Animal Care and Use Protocols.  These guidelines may be reviewed and followed as written, or exceptions may be requested.

Legal, regulatory, and moral guidelines require that animal pain, distress, and suffering be minimized in any experiment. In order to minimize pain and distress in studies on experimental neoplasia in rodents, investigators must address issues concerning tumor burden, the status of the tumor (e.g. ulceration), and criteria for euthanasia. The following guidelines are intended to provide investigators with an indication of the standards and criteria for euthanasia that have been considered acceptable by the IACUC.  The IACUC will evaluate an investigator’s request for an exception to these policies. The scientific rationale for the exception must be clearly stated.  

In general, tumor burdens should not be so large as to interfere with ambulation, eating, drinking, defecating, and urination.  Subcutaneous tumor burdens in rodents should not exceed 20% of the animal's body weight.  In the case of a 25 gram mouse, this would represent a single subcutaneous nodule of approximately 2.5 cm. diameter. Without a specific exception justified in the animal protocol, animals should be euthanized before tumors reach this size.

Ulceration is often an inevitable complication of subcutaneous tumors.  Animals with ulcerations or necrotic tumors must be sacrificed unless an exception has been approved.

Animals with skin ulcerations, particularly if immunocompromised, may have subclinical infections which can influence the tumor biology and confound experiments.  The investigator must monitor animals with ulcerations at least daily.  Animals must be sacrificed if the ulcerated tumor is painful (animal reacts to probing of the tumor) or if there is evidence of significant infection. 

In addition to the daily health checks performed by the animal care staff, documented daily health checks should be initiated by the investigator before the tumor begins to interfere with the physiological function of the animals. 

It is a violation of federal guidelines if, on daily health checks by the animal care staff or the PI,  the condition of the animal is found not to be in accordance with the approved protocol.

Investigators must administer euthanasia in moribund animals. The IACUC requires that an investigator judge when euthanasia is appropriate for moribund rodents based on objective signs.

Examples of Clinical Signs for Judging Moribundity:

1. inability to remain upright.
2. impaired ambulation (unable to reach food or water)
3. breathing rate very slow, shallow, and labored
4. any obvious prolonged illness including such signs as lethargy (drowsiness, aversion to activity, lack of physical or mental alertness), prolonged inappetence, bleeding, difficulty breathing, central nervous system disturbances, or chronic diarrhea or constipation
5. evidence of muscle atrophy or other signs of emaciation (body weight is not always proportionate)

Death as an endpoint studies are generally not approved by the Institutional Animal Care and Use Committee. Investigators are expected to justify the use of death as an endpoint in the animal use protocol application and the written justification will be reviewed by the IACUC. If approved, investigators are expected to monitor animals at least daily in these studies including weekends and holidays.

The scientific justification for use of death as an endpoint must include:

1. what alternatives were considered and why morbundity can’t be used in place of death what additional information is gained in the interval between moribund condition and death
2. the number of animals in survival duration protocols should be clearly stated, as well as the statistical techniques used to estimate the numbers in the study groups.

Section 23. Death as an Endpoint

Death as an endpoint studies are generally not approved by the Institutional Animal Care and Use Committee. Investigators are expected to justify the use of death as an endpoint in the Animal Use Protocol application and the written justification will be reviewed by the IACUC. If approved, investigators are expected to monitor animals at least daily including weekends and holidays. The scientific justification for use of death as an endpoint must include:

1. what alternatives were considered and why morbidity can’t be used in place of death what additional information is gained in the interval between moribund condition and death
2. the number of animals in survival duration protocols should be clearly stated, as well as the statistical techniques used to estimate the numbers in the study groups.

The IACUC recognizes that unexpected results may occur and the Principal Investigator may need to apply for exceptions to these guidelines at a critical time period in an approved protocol. When an immediate change is scientifically justified, the PI needs to submit a protocol amendment.

Section 24. Arthritis

Footpad injections with Complete Freund's Adjuvant (CFA) are not acceptable unless scientifically justified in your protocol and approved by the IACUC.

The use of intradermal injections must be scientifically justified over other routes of administration and must be approved by the IACUC.

Routes, volumes, dosages, dosing schedules, and contents of injections must be written into the protocol.

All injection dates will be written clearly on the cage card to facilitate care of animals with ACP husbandry staff.

 

The site of injection will be monitored daily post-injection for evidence of severe inflammation, ulceration, or for very large granuloma formation. The animals will be monitored daily for evidence of pain, distress, or infection resulting from the injection. The protocol will state what will be done to relieve the pain and distress for animals showing these signs. A schedule for monitoring animals and injection sites must be included in the protocol. If early signs of infection at the injection site are detected, the ACP veterinary staff will be consulted regarding treatment versus euthanasia. Animals with ulcerated injection sites will be monitored a minimum of three times per week by the PI or his/her staff. Animals must be treated and observed daily or euthanized if there is evidence of significant infection.

 

Arthritis measurements will be determined regularly after animals show difficulty ambulating or pain on manipulation of affected joints. The frequency of these measurements is stated in the protocol. A description of how measurements are performed will be stated in the protocol.

When severity of arthritis prevents easy access to food and water, the ACP staff will be notified that these animals are to receive water bottles with long sipper tubes and food in the bottom of the cage.

 

Animals in painful studies such as adjuvant arthritis must be administered effective analgesics unless the lack of analgesics is scientifically justified in your protocol.

Adjuvant arthritis experiments will be terminated by day 30 unless described in your protocol and approved by the IACUC. Animals will also be euthanized if:

·          Weight loss exceeds 20%

·          Animals are unable to eat or drink

·          Excessive skin breakdown occurs in arthritic joints

·          General health of the animal is at a level requiring euthanasia for humane reason

 

It is a violation of federal guidelines if the condition of the animal is found not to be in accordance with the approved protocol on daily health checks by the animal care staff or the PI. The IACUC recognizes that unexpected results may occur and the PI may need to submit an amendment to apply for exceptions to these guidelines at a critical time period in an approved protocol.

Section 25. Polyclonal Antibody Production

The Principal Investigator must provide a specific rationale for selection of species, adjuvant, route, sites and handling of antigens when completing the Animal Use Protocol. 

 

When an adjuvant is necessary to accomplish experimental goals, the investigator should select the adjuvant that causes the least amount of associated pain and discomfort.  Therefore, the use of other adjuvants causing less inflammation than complete Freund's adjuvant (CFA) is desirable.

 

Repeated inoculations of CFA, OR footpad, lymph node or intradermal inoculations with CFA are not acceptable unless scientifically justified and approved in advance by the IACUC.  The P.I. must provide data showing that usual methods do not give the result necessary for the experiment(s) proposed and that the requested method would give the desired result.  Note that quantity of antibody is not sufficient justification.

 

 Freund’s Complete Adjuvant (CFA) can cause severe inflammation and ulceration at the site of injection if used improperly. CFA should be used only for the initial immunization, with Freund’s Incomplete Adjuvant (IFA) used for subsequent booster injections. Other adjuvants should be considered before CFA and IFA. CFA should only be used if no appropriate alternatives are available.  

 

Freund’s adjuvant

FIA consists of 85% mineral oil or paraffin oil and 15% mannide monooleate (Arlacel A) as emulsifier. With the addition of heat-killed mycobacteria (M. butyricum or M. tuberculosis) the mixture is termed CFA. CFA is known to commonly produce undesirable side effects including granuloma formation, tissue necrosis and sloughing, abscesses, and fever. Other deleterious systemic effects, such as polyarthritis, have been reported. CFA is considered a human biohazard, such that accidental self inoculation, or splash in the eye have been shown to cause painful sequelae not readily amenable to treatment.

 

Other Adjuvants

Less problematic alternatives to Freund’s adjuvant are available and should be considered. RIBI Adjuvant System®, Specol®, TiterMax®, Montanide IAS50, and Montanide ISA70 are commonly used as appropriate alternatives. Noninflammatory adsorptive adjuvants such as alum and aluminum hydroxide gel may also be considered.

 

Consideration and justification must be given in the animal use protocol for selection of the laboratory animal species, adjuvant, volume per injection site, site of administration, number of sites, and response required. Particularly with the use of Freund’s adjuvant, it is important to note that the severity of potentially painful inflammatory reactions may be minimized by injection of a small volume of inoculum per site and the use of multiple, sufficiently separated, injection sites when appropriate.

 

Routes of Administration

 

Injections should be subcutaneous or, in rodents, intraperitoneal. Choice of other routes, such as intradermal are discouraged and must be scientifically justified by the investigator.  

 

For multiple subcutaneous sites, not more than 0.25 ml per SC site should be used for rabbits, 0.5 ml SC for sheep and goats, and 0.1 ml SC or 0.2 ml IP for mice. It is recommended that no more than five sites are used.   

 

If intradermal injections are scientifically justified by the P.I. and approved by the IACUC, no more than 0.05 ml may be injected at a site. Sites should be well separated to prevent consolidation of inflammatory responses.  

 

Subcutaneous inoculations should not be done in areas over bony protruberances such as the spine.

 

No injections should be done in the foot or footpad.

 

Frequency of Boosters

 

The frequency of boosters must be addressed in the animal use protocol. Two to three weeks is generally considered the minimum time period between the initial and subsequent immunizations. Booster immunizations cannot use CFA.

 

Monitoring

 

It is the Principal Investigator’s responsibility to ensure the animals are regularly checked. This is in addition to the daily checking done by animal technicians.

 

Investigators should observe the animals for evidence of pain or distress, and for evidence of lesions such as swelling, abscess or fistula formation, and infection or ulceration at the immunization sites and the ACP veterinary staff should be notified if these clinical problems are found.

 

The animal weight should regularly be compared to initial animal weights and this should be indicated in the protocol and documented.

 

Sites of inoculation must be examined daily or an alternate schedule must be described in the Animal Use Protocol and approved by the IACUC.

 

Records

 

The date of each injection and any adjuvant used must be recorded on the cage card, in a chart in the animal room, or on procedure stickers.

Section 26. Monoclonal Antibody Production

The Institutional Animal Care and Use Committee will not approve the use of the ascites method to produce monoclonal antibodies unless the investigator uses the following criteria to justify that the use of animals is necessary when completing the Animal Protocol Application:

 

1.       The proposed use of animals is scientifically justified;

2.       Methods that avoid or minimize discomfort, distress and pain (including in vitro methods) have been considered; and

3.       The latter have been found unsuitable.

 

In order to be in compliance with current NIH policy, it is the responsibility of the Institutional Animal Care and Use Committee (IACUC) to critically evaluate these criteria and appropriately document that the three criteria listed above have been fulfilled.

 

The 1997 workshop on alternatives in Monoclonal Antibody Production concluded that in vitro methods for MAb production should be the accepted method because the ascites method causes pain and suffering Thus use of the ascites method should be the exception and should require rigorous and well-documented justification. Justification should not be based solely on cost or convenience. If adequate justification is not provided for an exception, then in vitro  methods are required.

 

 If it is determined by the IACUC that the ascites method to product MAbs is justified, the investigator should do everything possible to reduce the number of animals used and to minimize pain and distress. Mice used for ascites production may experience problems such as respiratory distress, circulatory shock, difficulty walking, drinking and eating (with resulting dehydration). Furthermore, mice may develop solid tumors during the process.  In order to minimize pain and suffering, mice should be observed by the investigator daily including weekends and holidays to monitor the degree of abdominal distention and signs of illness.

 

Ascites fluid should be removed by peritoneal tap before abdominal distention is great enough to cause discomfort or interfere with normal activity. The maximum frequency of taps is every other day. 

 

Mice should be euthanized when they show signs of illness such as huddling, ruffled coat, reluctance to move when touched and cold ears. 

 

To avoid dehydration, supplements such as moistened chow can be provided.  Subcutaneous fluid therapy may be indicated.   

 

References: 

¹ OLAW Reports, National Institutes of Health, 11/17/97

 ² "Conclusions & Recommendations, Alternatives in Monoclonal Antibody  Production" - Workshop of Johns Hopkins Center for Alternatives  OLAW

Section 27. Wire bottom cages

In accordance with The Guide, the IACUC does not approve rodent studies using wire bottom cages except in certain cases when the investigator provides adequate scientific justification for this in the Animal Use Protocol.  It is recommended that investigators approved for studies with wire bottom cages provide resting boards for the rodents in the caging and it is also recommended that the caging be flattened bars rather than wire. Rodent foot abnormalities tend to appear in heavier and older rodents.

Section 28.  Imaging or Irradiation

An investigator who proposes to use a shared imaging or irradiation facility must address transportation issues, public health and animal health considerations and must have an approved SOP in place for the facility and the species proposed.  

 

For the fMRI center, this is a summary of the steps that a P.I. must follow to obtain approval:  

1) Review the Facility SOP (Standard Operating Procedure)

2) Submit a Proposal to the fMRI Center Biomedical Applications Committee (fCBAC)

3) Submit an Animal Protocol and required supporting documents to the UCSD Institutional Animal Care and Use Committee (IACUC)

4) Submit Personnel Qualifications (PQ) to IACUC 

 

See the IACUC Policy for more information.

Section 29.  Other Procedures Not Listed

In your answer, list and describe all procedures that are not described elsewhere on the prototocol. Include:

1. Non-Surgical Physical Manipulations	2. Special Equipment Monitoring/Measurements	3. Environmental manipulations/Non Standard Housing	4. Non Standard Husbandry Food/Water Manipulations
Procedures w/wo anesthesia not described in Q19 that require physical manipulation of the animal subject	Procedures w/wo anesthesia not described in Q19 or Q28 that require the use of specialized equipment and extended manipulation of the animal subject outside its home environment	Specialized Equipment and Procedures whose purpose is to house animals in an experimental environment the animal would not receive in its home cage	The addition of any compound to standarized chow or drinking water for any experimental purpose or conditioning
IV infusions	Echocardiography	Hypoxic protocols/hypobaric chambers	Antibiotic prophylaxis
Transcardial perfusions	Non-invasive or telemetric measurements like ERG, EEG, ECG	Temperature manipulations	Agents administered for genetic manipulation; disease model induction, or cellular identification
Limb casting	Fluorescent Microscopy	Refrigeration boxes	Special formulated diets
Endoscopy	Nebulizers	Light cycle manipulations
Non-standard injections--intraocular; non-catheterized intrathecal, intra-hepatic, retroorbital, intranasal, intra- vaginal, footpad, lymph node	Tethering	Light/Dark Boxes; Reverse Light Cycle Equipment
Warming Devices		Metabolism Caging
Ventilation Studies		Acoustic Chambers
Burn Models
Manual Egg Collection

Questions 30 – 33.  Potential Animal Pain and Distress/Euthanasia

The UCSD policy is adopted from the U.S. Government Principles for the Utilization and Care of Vertebrate Animals Used in Testing, Research, and Training:

 

·          Proper use of animals, including the avoidance or minimization of discomfort, distress, and pain when consistent with sound scientific practices, is imperative. Unless the contrary is established, investigators should consider that procedures that cause pain or distress in human beings may cause pain or distress in other animals.

 

·          Procedures with animals that may cause more than momentary or slight pain or distress should be performed with appropriate sedation, analgesia, or anesthesia. Surgical or other painful procedures should not be performed on unanesthetized animals paralyzed by chemical agents.

 

·          Animals that would otherwise suffer severe or chronic pain or distress that cannot be relieved should be painlessly killed at the end of the procedure or, if appropriate, during the procedure.

 

·          The living conditions of animals should be appropriate for their species and contribute to their health and comfort. Normally, the housing, feeding, and care of all animals used for biomedical purposes must be directed by a veterinarian or other scientist trained and experienced in the proper care, handling, and use of the species being maintained or studied. In any case, veterinary care shall be provided as indicated.

 

·          Investigators and other personnel shall be appropriately qualified and experienced for conducting procedures on living animals. Adequate arrangements shall be made for their in-service training, including the proper and humane care and use of laboratory animals.

 

·          Where exceptions are required in relation to the provisions of these Principles, the decisions should not rest with the investigators directly concerned but should be made by an institutional animal care and use committee. Such exceptions should not be made solely for the purposes of teaching or demonstration.

 

UCSD employs three Pain and Distress Categories C, D, and E (corresponding to the USDA reportable pain categories) as follows: 

 

Category C includes only procedures that are considered to produce minimal, transient, or no pain or distress in animals when performed by a competent individual. The definition of USDA category C also emphasizes that protocols involve no more than momentary or slight pain or distress and no use of pain-relieving drugs.

 

Category D includes procedures that have the potential to produce pain or distress in animals, but which are performed using appropriate and adequate anesthetics, analgesics, or tranquilizers to alleviate the pain or distress. 

 

Category E includes potentially painful or distressing procedures that are performed without appropriate and adequate anesthesia, analgesia, or tranquilizers; or are not followed with appropriate measures to alleviate pain or distress; or are not amenable to relief by therapeutic measures. 

 

General approach to pain/distress categorization

 

Investigators and reviewers should employ the following two sets of questions and criteria: 

 

1. Comparison with humans. What would be an equivalent or comparable procedure or state in humans, and would it cause more than minimal or transient pain or distress? If pain might be expected, would it be necessary to treat it and how? What would the consequences be of not treating the pain? Animal procedures for which the human equivalent causes very transient, minor, or no pain or distress, or treatment is not necessary for the majority of individuals, should generally be classified as Category C procedures, unless animals show objective signs of pain or distress.
2. Objective signs of pain and distress in animals. Do animals develop objective signs of pain and distress following the procedure?
   Such signs include, but are not limited to:
   
   • Changes in activity level: reduced spontaneous motor activity, recumbent position, delayed response to handling, or observation of pacing, restlessness or lameness.
   • Changes in appearance: hunched position, ruffled fur, decreased grooming, discharge around nose and eyes.
   • Changes in temperament: increased aggression, guarding, reluctance to interact.
   • Vocalizations: teeth-grinding, chattering, whining, whimpering.
   • Changes in feeding behavior: decreased food and water consumption, reduction in body weight, urine, or stool output.
   • Physiologic changes: heart rate, respiratory rate, blood pressure, body temperature, skin color.
   • Appearance of surgical site: erythema, swelling or discharge, excessive licking or chewing.
   
   Procedures that cause animals to exhibit objective signs of pain or distress spontaneously and to a significant extent should generally be categorized as Category E (or Category D if treated appropriately with analgesics or anesthetics) unless the investigator can justify variance. Importantly, the constellation of several parameters of pain and distress is given greater weight in evaluation than any one of these signs when observed in isolation.

Examples

 

The following examples illustrate the approach to assigning experimental procedures to categories: (NOTE: These lists are examples and are not complete lists of every procedure; investigators may contact the IACUC for further guidance) 

 

Examples for Category C (procedures that are minimal, transient, or involve no pain or distress)

• Handling, weighing, behavioral observation
• Injections and non-surgical fluid collection of modest volumes (i.p., i.v., s.c.)
• Identification (ear punching, tattooing, ear notching)
• Tail clipping of mice less than 4 weeks old; less than 5 mm
• Food and water restriction without clinical health problems in closely monitored animals
• Mild to moderate ascites formation without systemic effects
• Infections without objective signs of pain or distress
• Inflammatory conditions without systemic effects (e.g. localized to skin)
• Cancer studies, limited to subcutaneous tumors of small - moderate size, which adhere to Policy
• Gavage
• Euthanasia followed by tissue collection
• Pain models in which animals can compensate for any potential pain by behavioral modification

Examples for Category D (procedures with the potential to produce pain or distress in animals, but which are performed using appropriate and adequate anesthetics, analgesics, or tranquilizers to alleviate the pain or distress) 

• Surgical procedures with appropriate and effective anesthesia and analgesia
• Blood or tissue sampling with appropriate anesthesia or tranquilizers
• Physical trauma under anesthesia

Examples for Category E (potentially painful or distressing procedures that are performed without appropriate and adequate anesthesia, analgesia, or tranquilizers because they would adversely affect the results or interpretation of data)

• Studies with significant mortality or death as an endpoint
• Procedures leading to clinically significant (>10%) weight loss
• Noxious stimuli from which there is no escape

Pain and distress in breeding colonies

Breeding of genetically altered animals raises special concerns as spontaneous phenotypes can occur in the absence of specific interventions. For example, an animal might be born healthy but then develop symptoms that would necessitate inclusion into Category E. In addition, the phenotype of a new mutation is often not known or predictable. Unexpected phenotypes which fall into Category E will require the Principal Investigator to submit an amendment to the applicable Animal Use Protocol.

 

Analgesia

 

All animals that undergo a painful procedure such as surgery and are likely to experience post procedural pain, should receive preemptive analgesic medication. The Principal Investigator in consultation with the ACP Veterinarian should use their professional judgment to determine the need for analgesic medication. Procedures for analgesic administration and a plan for postoperative care should be prepared in advance. Exceptions to this policy must be justified scientifically in writing and approved by the IACUC. Where doubt exists regarding the potential for interference with an experiment, consideration should be given to performing a pilot study designed to investigate the possibility of interference.  

 

Note: Pain is more easily controlled by administration of analgesics before pain starts than after the animal is experiencing pain. Therefore, analgesics should be administered preemptively before the animal recovers from anesthesia. 

 

Recognizing Pain

 

Anthropomorphic assessment of the degree of pain or discomfort experienced by an animal can be difficult and inaccurate. While the physiologic mechanisms of pain perception are similar in all mammalian species the ability to tolerate and cope with pain may be vastly different in different species. For example, prey species such as rodents have adapted to hide overt signs of pain to avoid signaling to a predator that they are ill and would be an easy meal. Therefore, a rodent that is experiencing mild to moderate pain may display no clinical signs associated with its discomfort. Moderate to severe pain in rodents frequently leads to changes in normal physiology or behavior. Accurate recognition of these changes requires that research personnel have some knowledge of normal behavior and physiology for the species they are using.

 

General clinical signs associated with pain in most species:

• Changes in activity level: reduced spontaneous motor activity, recumbent position, delayed response to handling, or observation of pacing, restlessness or lameness.
• Changes in appearance: hunched position, ruffled fur, decreased grooming, discharge around nose and eyes.
• Changes in temperament: increased aggression, guarding, reluctance to interact.
• Vocalizations: teeth-grinding, chattering, whining, whimpering.Changes in feeding behavior: decreased food and water consumption, reduction in body weight, urine, or stool output.
• Physiologic changes: heart rate, respiratory rate, blood pressure, body temperature, skin color.
• Appearance of surgical site: erythema, swelling or discharge, excessive licking or chewing.

Analgesic Drugs and Delivery Methods

There are many drugs and routes of administration available for alleviation of pain in all species. Some drugs, such as the opiates (example: buprenorphine or morphine), provide pain relief through their action on the central nervous system. Corticosteriods and non-steroidal anti-inflammatory drugs (NSAIDS), relieve pain by decreasing inflammation. Long acting local anesthetics such as bupivacaine, provide postoperative analgesia by their direct action blocking nerve conduction.  Common analgesics for rodents and rabbits are shown in the chart below. Check the ACP web site for the Recommended Dosage Chart for analgesics in many other species. For a consultation with the veterinary anesthesiologist about analgesia in rodents or other species please contact vetservices@acp.ucsd.edu.  

Recommended Dosage Rates for Analgesic Drugs in Rodents and Rabbits
Drug Name	Mouse	Rat	Rabbit
Bupivicaine local anesthetic; injected or dripped onto incision site	Maximum total dose of 4 mg/kg given once at the end of surgery	Maximum total dose of 4 mg/kg given once at the end of surgery
Buprenorphine mild pain; longer duration of action	0.05-0.1 mg/kg SC every 8-12 hours	0.01-0.05 mg/kg SC every 8-12 hours	0.01-0.05 mg/kg IM, SC or IV every 6-12 hours
Butorphanol moderate pain; short duration of action	1.0-2.0 mg/kg SC every 4 hours	1.0 - 2.0 mg/kg SC every 2- 4 hours	0.1-0.5 mg/kg SC, IM,IV every 4 hours
Carprofen (Rimadyl)	5 mg/kg SC every 24 hours	5 mg/kg SC every 24 hours	4 mg/kg SC every 24 hours
Flunixin (Banamine)	2.5 mg/kg SC every 12-24 hours	2.5 mg/kg SC every 12-24 hours	1.0 mg/kg SC every 12  - 24 hours
Ketoprofen (Toradol)	5 mg/kg SC every 24 hours	5 mg/kg SC every 24 hours	3 mg/kg SC every 24 hours
Morphine moderate to severe pain; short duration	2 - 5 mg/kg SC every 4 hours	2 - 5 mg/kg SC every 4 hours	2 - 5 mg/kg SC, every 4 hours
Analgesic effectiveness must be evaluated in each animal due to variations in response between individuals and strains.

Question 34. Method of Euthanasia

The Institutional Animal Care and Use Committee requires investigators to follow the recommendations of the Panel on Euthanasia of the American Veterinary Medical Association unless scientific justification is provided. For chemical agents, please describe agent, route, and dose. For physical methods without anesthetic, provide detailed scientific justification. Describe how confirmation of death will be assured.

Physical methods such as cervical dislocation and decapitation in the absence of anesthesia are not considered an acceptable means of euthanasia unless required for the scientific goals of the project and justification is provided.

Confirmation of Death: Euthanasia and death must be verified (e.g. cardiac and respiratory arrest, circulatory collapse, brain death). After euthanasia by inhalant anesthesia or carbon dioxide, personnel should ensure that death has occurred by carefully monitoring animals for the absence of heart beat and/or respirations, or by performing cervical dislocation, decapitation, exsanguination, removal of an organ, perfusion, or cutting the chest open to induce pneumothorax.

The Principal Investigator has the responsibility to assure that all personnel understand the nature of the hazardous agents to be used in the protocol and understand the precautions, equipment, personal protection equipment needed.

 

The P.I. has the responsibility to ensure that all personnel have completed the Survey and enrolled in the Medical Monitoring Program as necessary.

 

The P.I. must assure on this protocol that all necessary BUA, RUA and IBC approvals are obtained from Environmental Health and Safety before any work is started on the studies involving hazardous agents.

 

If the hazardous agent you propose to use is not on the pull-down list in the protocol, please call EH&S (Steve Kowalewsky) at 534-6715 to discuss the agent with the Biosafety Officer.  He can add it to the list after the discussion.

If research will be funded by an external sponsor (grant, contract, or gift), the P.I. must assure that all necessary approvals have been obtained from Office of Contract and Grant Administration and Conflict of Interest Office.