How can I take blood from a mouse? - briefly
Induce brief anesthesia, warm the mouse, and make a small incision at the distal tail with a sterile scalpel to collect blood into a heparin‑treated capillary tube. For larger volumes, perform a retro‑orbital puncture using a fine glass capillary under the same anesthetic conditions, maintaining aseptic technique.
How can I take blood from a mouse? - in detail
Collecting murine blood requires preparation, proper anesthesia, and strict adherence to volume limits and aseptic technique.
Begin with equipment: sterile syringes (27‑30 G), fine needles, heparin‑ or EDTA‑coated microtubes, gauze, disinfectant (70 % ethanol), warming pad, and an appropriate anesthetic agent (isoflurane vaporizer or injectable mix). Verify that the animal’s weight is recorded to calculate permissible draw (no more than 10 % of total blood volume per 24 h, approximately 0.2 ml per 20 g mouse).
Anesthesia: induce and maintain a surgical plane using isoflurane (2–3 % induction, 1–2 % maintenance) or a ketamine/xylazine cocktail (dose adjusted to body weight). Confirm loss of reflexes before proceeding.
Select a sampling site:
- Tail vein – suitable for repeated small volumes (≤0.1 ml). Warm tail for 2–3 min, locate lateral vein, puncture with a 27‑G needle at a shallow angle, collect blood into a pre‑coated tube, apply gentle pressure to stop bleeding.
- Submandibular (facial) vein – yields 0.1–0.2 ml in a single draw. Restrain mouse, make a brief incision at the mandibular angle, insert a 30‑G needle, aspirate, then apply pressure with gauze.
- Retro‑orbital sinus – provides 0.1–0.2 ml quickly, reserved for terminal procedures or when other sites are unsuitable. Position mouse laterally, insert a 27‑G heparinized needle at the orbital rim, draw blood, then apply pressure to the eye for several seconds.
- Cardiac puncture – terminal method, obtains up to 1 ml. Perform after deep anesthesia, open thoracic cavity, insert a 23‑G needle into the left ventricle, aspirate, then euthanize according to protocol.
After collection, transfer blood to chilled microtubes, mix gently to prevent clotting, and process immediately or store at appropriate temperature. Monitor the mouse during recovery; keep on a warming pad until ambulatory. Record date, time, volume, and site for each sample.
Maintain documentation of institutional animal care and use committee (IACUC) approval, ensure personnel are trained in microsurgical techniques, and dispose of sharps in compliance with biosafety regulations.