Laboratory Sample Submission Guidelines

When shipping animals/samples, include a completed Diagnostic & Research Histology Laboratory submission form with the shipment. Place the submission form in a Ziploc-type bag to prevent it from getting soiled or contaminated. Please note we will not be able to process animals/samples submitted without completed submission form.

Live Animal
Pack each animal separately in approved shipping container following guidelines of live animal transport. Ensure each animal and shipping container are clearly identified. Packing animals separately prevents transfer of ectoparasites from animal to animal.

Carcass
Pack each animal carcass in separate Ziploc-type bag. Ensure each bag is clearly identified and sealed. Immediately after euthanasia, place carcass in shipping box lined with ice pack refrigerants to keep the carcass cold and deliver to the lab promptly. Do not freeze the carcass.

Sample collection
All samples for bacteriology must be collected aseptically to prevent microbial contamination and ideally before any antimicrobial treatment is initiated.

For specimen collection, only commercially available culture swab systems consisting of swab and tube with transport medium are used. Dry swabs are not suitable. Tissues, fluids (e.g. urine, aspirates, exudates) and feces must be collected in sterile containers or vials. We do not accept samples in syringes with needles attached. Label each culture swab or container with animal ID, collection site, and date & time of collection. Specimens that cannot be delivered to the lab the same day must be refrigerated and delivered next day. Contact us if you need to obtain culture swabs and/or sterile containers.

Live Animal
Pack each animal separately in approved shipping container following guidelines of live animal transport. Ensure each animal and shipping container are clearly identified.

Carcass
Pack each animal carcass in separate Ziploc-type bag. Ensure each bag is clearly identified and sealed. Immediately after euthanasia, place carcass in shipping box lined with ice pack refrigerants to keep the carcass cold, and deliver to the lab promptly. Do not freeze the carcass.

Live Animal
Pack each animal separately in approved shipping container following guidelines of live animal transport. Ensure each animal and shipping container are clearly identified. Packing animals separately prevents transfer of ectoparasites from animal to animal.

Carcass
Pack each animal carcass in separate Ziploc-type bag. Ensure each bag is clearly identified and sealed. Immediately after euthanasia, place carcass in shipping box lined with ice pack refrigerants to keep the carcass cold, and deliver to the lab promptly. Do not freeze the carcass.

Fecal Samples
Collect fecal sample with clean disinfected forceps in sterile container. Sample size: at least 10 fresh fecal pellets/mouse, 3-5 fresh fecal pellets/rat, 3-5 g fresh feces/medium or large animal. Deliver the sample(s) on ice pack refrigerants the same day, If same-day delivery is not possible, keep samples refrigerated (do not freeze) and deliver next day. Fecal samples for examination for motile protozoan trophozoites must be examined within 30 min after collection.

Live Animal
Pack each animal separately in approved shipping container following guidelines of live animal transport. Ensure each animal and shipping container are clearly identified.

Carcass (fresh)
Pack each animal carcass in separate Ziploc-type bag. Ensure each bag is clearly identified and sealed. Immediately after euthanasia, place carcass in shipping box lined with ice pack refrigerants to keep the carcass cold, and deliver to the lab promptly. Do not freeze the carcass.

Specimen in Fixative
Tissues must be placed in fixative (preferably 10% neutral buffered formalin) immediately after collection to avoid autolysis and desiccation. Ratio of tissue to fixative must be at least 1:10 (e.g. 30 g mouse carcass is placed in at least 300 mL of formalin). If submitting mouse or rat carcass, the body cavity must be cut open from chin to anus to expose all internal organs to formalin and to ensure proper tissue fixation (the diaphragm must be also cut to expose heart and lungs). Inadequate tissue fixation causes autolytic changes that may prevent proper tissue evaluation. Label container with animal/sample ID, date of fixation and type of fixative used (e.g. formalin). Containers must not leak.

Suitable specimens for necropsy and/or histopathology are those in good post-mortem state that are not showing sings of advanced autolysis.

Each primary receptacle must clearly identify its contents. Required information on the primary receptacle (choose all that apply): animal/ sample ID, date and time of animal euthanasia and/or sample collection, site of collection, type of sample, type of fixative used. If biohazard material is suspected, a biohazard label must be attached to the outer packaging. All other labeling and marking information must be attached to the outside of the package as required by DOT regulations.

Please include a copy of the completed submission form with your samples and an empty slide box with capacity for the work to be performed if you do not wish to be charged for one. If the form is missing or incomplete, your work order will not be initiated.

Samples will be processed on a first come, first served basis. The turnaround time depends on both the size and complexity of your project and the lab’s current work load. If you have any questions about completing the submission form, tissue preparation, procedures, services, or if you would like help planning the integration of histology into your experiments, please contact ingrid.barta@ubc.ca or call the histology lab at 604-822-7091.

The submission of a high-quality sample is a crucial component for a successful evaluation. Please see below for important information about specimen fixation.

How should I handle unfixed tissue?
To avoid crushing artifact, be gentle and try to avoid pinching or grasping the tissue too tightly. Try handling tissue by grasping areas adjacent to the tissue you desire to collect (e.g. manipulate intestine by holding onto the mesenteric fat instead of the intestine itself). If tissue is particularly bloody, rinse it with cold normal saline or PBS before fixation.

What is the purpose of fixation?
To prevent autolysis and putrefaction, tissues must be fixed as soon as possible after cessation of blood flow. This is especially important for intestinal and CNS tissue. There are a myriad of fixative choices, each with its own advantages and disadvantages. 10% Neutral Buffered Formalin (NBF) has been shown to be the most useful and forgiving fixative and is therefore most commonly used.

If tissue not sufficiently fixed (fixative expired, concentration too low, volume too low or not enough time), the lab cannot guarantee the quality tissue morphology or staining. H&E staining is fairly forgiving but proper fixation is very important for achieving accurate special staining.

How can fixation be achieved?
Fixation can be done by cardiac perfusion of fixative or by immersion fixation. Perfusion fixation followed by completion of fixation by immersion is best but is not always feasible. For immersion fixation, you must use at least 10x the volume of fixative relative to the size of the specimen. The fixative needs to penetrate the tissue from all sides. Therefore, if tissue floats in the fixative, cover it with a piece of paper towel or gauze to ensure it is fully submerged at all times; if a large portion of the tissue rests on the bottom of the container, place some loosely arranged gauze under it. Always use fixative that is within its expiry date or prepared fresh. We strongly advise purchasing a commercial fixative.

How long should fixation take?
Time of fixation depends on a variety of factors including the fixative used, the size of the specimen and temperature. For 10% neutral buffered formalin, the most commonly used and forgiving fixative, the time required to penetrate the tissue is approximately the square of the depth of the tissue (at room temperature). Thus, for a piece of tissue 5mm thick, the time to complete penetration is 5² = 25 hours.

Some tissue types (e.g. skin, gut) curl when fixed, especially if fixed in a thin tube. Tissue cannot be straightened after fixation without damaging it so if it is important that the tissue remains flat for orientation purposes, please secure the tissue before fixing. Please contact the lab if you require instructions for this.

Tissues should be placed in 70% ethanol and kept at 4°C if they are to be stored long term (>3 months).

What happens next?
Chemical fixation continues after fixative fully penetrates the tissue, further stabilizing it. This is desirable for preserving morphology but it can come at the expense of altering or blocking antigens you may wish to detect with IHC or IF. In this case, it is advisable to transfer your samples to 70% ethanol after fixation. Tissues should be placed in 70% ethanol and kept at 4°C if it is to be stored long term (>3 months).

If using a fixative other than 10% NBF or 4% PFA, please contact the lab if you are unsure of the appropriate storage medium.

Fixed tissue submitted in 10% NBF or 4% PFA, 70% Ethanol or other appropriate storage medium.

CLEARLY label all containers, tubes, etc. with a CONCISE ID that match sample IDs you provide in the submission form. Cassette and slide labelling software does not support long sample IDs. Refer to the submission form for acceptable fields (note 6 characters max for each field).

Pre-trimmed tissue already placed in cassettes and submitted in storage medium as above.

Ensure storage medium covers all cassettes sufficiently. Label cassettes clearly in PENCIL. Do not use narrow-necked containers (easy to get the cassettes in but difficult to remove). Do not over-stuff the cassette or squeeze tissue into the cassette. The tissue should be small enough that it comfortably fits into the cassette without touching the sides. Additionally, the tissue must be NO MORE than 5mm in thickness in at least one dimension (usually this is relative to the plane of desired sectioning); ideal thickness is 3-4mm. Use of biopsy pads: contact the lab to determine whether appropriate for your samples.

If you are submitting tissue in ethanol, ensure that your container labels are written in solvent-resistant ink (no permanent markers). When possible, please avoid submitting tissues in glass containers to reduce the risk of breakage and injury to yourself and others. Please identify the storage medium on all containers, tubes, etc.

If known, please indicate fixative used. Blocks in a Ziploc-type bag or box is sufficient provided the blocks are protected from damage during transit.

If sectioned somewhere other than at the ACS Diagnostic & Research Histology Laboratory or if sections are frozen, please indicate fixative used and section thickness on the submission form. Frozen section slides must be labeled with pencil as permanent markers will wash out in the solvents used during staining.

Submit clean slides: remove excess dried resinous (permanent) mounting medium from the slides and wipe clean any fingerprints. Unfortunately, we cannot accept slides coverslipped with aqueous mounting medium. Please include a labeled USB flash drive (or other external drive) with your submission for saving the image files. Please indicate the number and IDs of slides to be scanned for each magnification. Available magnifications: 2x, 10x, 20x and 40x with slide label capture option. Please inquire for scanning at higher magnifications.

Output image format is .vsi so images are viewable using:

• Olympus OlyVIA (PC only)
• OlympusViewer Plugin (ImageJ)
• QuPath (Windows/Mac/Linux)

Please refer to UBC Animal Care Committee, Policy 003: Transportation of Rodents from a UBC Facility to Another Institution.

Fresh, unfixed carcasses must be placed on ice pack refrigerants immediately after euthanasia (or kept in refrigerator until shipped) and delivered to our lab promptly on the same day to minimize post-mortem changes. Do not freeze carcasses. Samples such as blood, serum, urine, feces, culture swabs, and other biological materials need to be sent on ice pack refrigerants. It is important to keep carcasses and samples cold (at refrigerator temperature) at all times until they are delivered to the lab. Samples that are already frozen (e.g. serum, plasma) must remain frozen during transport and cannot thaw. Contact our lab for more information regarding sample storage/transport.

Each sample must be placed in a leakproof primary receptacle (e.g. tube, vial, container). The primary receptacle (containing the sample itself) is then placed in a secondary leakproof packaging (e.g. Ziploc-type bag or container). If the sample is liquid, an absorbent material is placed between the primary receptacle/s and the secondary packaging to absorb any spills. For sensitive and fragile samples, we recommend using triple packaging (see Transport off UBC Campus).

Triple packaging is required: leakproof primary receptacle is placed in a leakproof secondary packaging (containing absorbent material if the sample is liquid). The secondary packaging is then placed in a rigid outer packaging that is strong enough to hold the samples (e.g. fibreboard or Styrofoam box). In the box, air pouches, bubble wrap or similar material is placed to secure the samples and keep them in upright position.