Presentation on theme: "Handling Lab mice. Environment Caging Feed Bedding Water Sanitation Provisions for off-hour care Environmental monitoring and maintenance."— Presentation transcript:
Handling Lab mice
Environment Caging Feed Bedding Water Sanitation Provisions for off-hour care Environmental monitoring and maintenance
Table Minimum Space Requirements for Housing Laboratory Animals in Cages a WeightFloor area/animalCage height b Animal(g)(in 2 )(cm 2 )(in)(cm) Mouse< > Rat< > a Guidelines are derived from Guide for the Care and Use of Laboratory Animals (ILAR, 1985).b From the resting floor to the cage top From Current Protocols in Immunology Online
High-quality cages INDIVIDUALLY VENTILATED CAGES laminar air rack From TENCIPLAST
Ventilation efficiency with low velocity Air Speed Visibility Easy access
Metabolic and Diuresis Cages From TENCIPLAST
Diuresis cage From TENCIPLAST
Metabolic cage From TENCIPLAST
Refrigerated Collection Rack
Diets Chemically defined diets Custom-mix diets Natural diets Purified diets Semi-purified diets Other diets Animal Technologies Ltd.Animal Technologies Ltd. Bio-Serv Dyets Inc. Harlan Netherlands Harlan Teklad ICN Biomedicals, Inc. Lactamin AB Lillico Biotechnology NLS Animal Health Nutritional Research Assoc., Inc.Nutritional Research Assoc., Inc. P.J. Noyes Company, Inc. Purina LabDiet Research Diets, Inc. Scanbur BK (Denmark) SE Lab Group Special Diets Services Zeigler Bros. Inc.
Water Fresh uncontaminated and sterile water Water purification systems deionization reverse osmosis ultrafiltration Hyperchlorination/acidific ation
Table Recommended Relative Humidity and Dry Bulb Temperature for Animals Housed in Cages a Dry bulb temperature b Animal Relative humidity (%) CC FF Mouse Rat Hamster Rabbit a Guidelines from ILAR, b ILAR, 1965, From Current Protocols in Immunology
Ventilation Oxygen Thermal loads Air contaminants 10 to 15 room air changes per hour w/o recirculation
Illumination Regular diurnal light cycle Light levels: 323 lux (30-foot candles) 1 meter above the floor
Mouse Mouse Handling and Manual Restraint Mouse handling and manual restraint. Apply slight, rearward traction on the tail (A). Grasp skin behind ears with thumb and index finger (B). Transfer the tail from the preferred hand to beneath the little finger of the hand holding the scruff of the neck (C). From Current Protocols in Immunology
Rodent restrainers From Current Protocols in Immunology Rodent restrainers. With animal under control as described for handling and manual restraint, place the head at opening of the box while maintaining tension on the tail. Allow animal to crawl in and place the securing block appropriately. Size Length of time Injection and sampling
Ear Notch or Punch for Mouse, Rat and Hamster From Current Protocols in Immunology
mouse Two people may be necessary for intramuscular injections in rats, one to restrain the rat and another to retract and immobilize the rear leg and to inject into heavy musculature of the upper thigh. For the mouse From Current Protocols in Immunology
Tented intracapsular area
From Current Protocols in Immunology 1. Fill syringe with injectate and remove air bubbles. Removal of air bubbles is critical to avoid air embolism. 2. Place mouse in a restrainer. 3. Warm the tail with a heat lamp or by immersing in warm water to dilate vessels. 4. Swab the tail with 70% ethanol on a gauze sponge or swab. 5. Immobilize the tail with gentle traction. 6. Visualize the lateral tail vein and insert needle parallel to the vein 2 to 4 mm into the lumen. Keep the bevel of the needle facing up. 7. Inject slowly. No bleb should form if needle was properly located. If a bleb appears, indicating failure to cannulate the vein, additional attempts may be made proximally. Thus it is helpful to make the first attempt at injection as close to the tip of the tail as possible. 8. Withdraw the needle and apply digital pressure if necessary to achieve hemostasis. INTRAVENOUS INJECTION OF MOUSE
Intraperitoneal injection of the mouse For the mouse, rat, or hamster, expose the animal's abdomen, tilting the head downward. Insert needle into the lower left or right quadrant of the abdomen, avoiding the abdominal midline. From Current Protocols in Immunology
FOOTPAD INJECTION OF THE MOUSE From Current Protocols in Immunology Spread the toes of one hind foot and introduce the needle into the soft tissue pad of the plantar surface. Alternatively, the injection can be made subcutaneously between the second and third digits. <50 μl
BLOOD COLLECTION FROM TAIL VEIN OF MOUSE AND RAT USING MICROHEMATOCRIT TUBE From Current Protocols in Immunology Visualize a sampling site of the lateral tail vein at approximately midpoint on the length of the tail. Collect blood from the hub of the needle with the microhematocrit tube.
BLOOD COLLECTION FROM ORBITAL SINUS OR PLEXUS OF MOUSE AND RAT Slowly, and with axial rotation, advance the tip of the microhematocrit tube gently towards the rear of the socket until blood flows into the tube. Remove the microhematocrit tube from orbit and dab excess blood from the site with a gauze sponge or swab moistened in saline or PBS. Introduce the end of the microhematocr it tube at the medial canthus of the orbit. From Current Protocols in Immunology
Carbon Dioxide Asphyxiation 60% to 100% CO2 (tank or house carbon dioxide) or dry ice Impervious container with raised floor and lid or CO2 chamber with lid Precharge the impervious container or the CO2 chamber with CO2 prior to introducing the animals. If dry ice is used as the CO2 source, a raised floor is necessary to prevent animal contact with the dry ice. Placing warm water on the dry ice facilitates vaporization and filling of the container. Unconsciousness will occur within 30 sec, but animals should be left in the container for several minutes to ensure death. 4. Verify death by lack of cardiac pulse and fixed and dilated pupils prior to carcass disposal. Cause perivascular edema in the lungs or alveolar hemorrhage. Neonates are resistant to the effects of high levels of CO2
Cervical Dislocation of Mouse Institutional Animal Care and Use Committee policy. Remove the mouse from its cage by grasping the base of the tail. Place it on a smooth, hard surface without releasing the tail. Place the pencil or metal rod firmly behind the ears and across the neck. Pull the tail sharply to the rear while pressing down on the neck (to quickly dislocate the cervical vertebrae).
Removal of Lymphoid Organs Cytotoxicity or proliferation assays Analysis of cytokine production or response Isolation of B, T, and adherent cell subpopulations for culture or reconstitution of other animals.
From Current Protocols in Immunology Wet the fur with 70% ethanol to sterilize the area and reduce the possibility of contamination. Make a midline incision with iris scissors. Retract the skin above the head and below the thighs by pulling it with gloved fingers.
Remove the thymus Make an incision in the chest, beginning at the xiphoid and extending to the neck with surgical scissors. Retract the ribs with curved forceps. It may be necessary to crack the ribs for effective retraction. The thymus is a yellowish-white bi-lobed organ found just under the ribs, attached above the heart in the midline. Grasp each lobe of the thymus with curved forceps and gently pull the thymus away. Place the thymus tissue in several milliliters of balanced salt solution or tissue culture medium in a tissue culture plate. Thymic size varies with age, being maximal at 3 to 5 weeks in the mouse and shrinking to a minimum at ~6 months.
The lymph nodes and spleen of the mouse. From Current Protocols in Immunology Remove the lymph nodes The major lymph nodes are found in the axillary, cervical, inguinal, and mesenteric regions. Typically, 1- 5x10 7 lymph node cells can be isolated from a single mouse. Grasp the lymph nodes with curved forceps and pull them free of attached tissue. Occasionally, it is necessary to dissect adjacent fat. Place the lymph nodes in several milliliters of HBSS or tissue culture medium in a tissue culture plate
Remove the spleen Make a 1-in. incision at the left of the peritoneal wall with surgical scissors. The spleen is attached to the greater curvature of the stomach by connective tissue. Grasp as much of the spleen as possible with curved iris forceps. Gently pull the spleen free of the peritoneum, tearing the connective tissue behind the spleen. Alternatively, the top and bottom portions of the spleen can be grasped and torn from the connective tissue one part at a time. It is usually more efficient to remove the spleen all at once, particularly if large numbers of animals will be used. Place the spleen in several milliliters of HBSS or tissue culture medium in a tissue culture plate.
HBSS (Hanks' balanced salt solution) 0.40 g KCl (5.4 mM final) 0.09 g Na2HPO4·7H2O (0.3 mM final) 0.06 g KH2PO4 (0.4 mM final) 0.35 g NaHCO3 (4.2 mM final) 0.14 g CaCl2 (1.3 mM final) 0.10 g MgCl2·6H2O (0.5 mM final) 0.10 g MgSO4·7H2O (0.6 mM final) 8.0 g NaCl (137 mM final) 1.0 g D-glucose (5.6 mM final) 0.2 g phenol red (0.02%; optional) Add H2O to l liter and adjust pH to 7.4 with 1 M HCl or 1 M NaOH Filter sterilize and store up to 1 month at 4°C HBBS can be purchased from Biofluids or Bio-Whittaker. HBSS may be made or purchased without Ca2+ and Mg2+ (CMF-HBSS). These components are optional and usually have no effect on an experiment; in a few cases, however, their presence may be detrimental. Consult individual protocols to see if the presence or absence of these components is recommended. Bottles should be kept tightly closed to prevent CO2 loss and subsequent alkalinization.1 M HCl1 M NaOH