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1. Introduction and Definitions:

Use the following table to determine the most appropriate site for blood collection based on the volume required.

The following are “good practice” guidelines recommended for blood collection volumes, sites and needle gauges. As a general principle, sample volumes and number of samples should be kept to a minimum. As a general guide, up to 7.5% of the total blood volume can be taken on a single occasion from a normal, healthy animal on an adequate plane of nutrition with minimal adverse effects; 10% once every two weeks and 15% once every four weeks. For repeat bleeds at shorter intervals, a maximum of 1.0% of an animal's total blood volume can be removed every 24 hours. The acceptable quantity and frequency of blood sampling is dependent on the circulating blood volume of the animal and the red blood cell (RBC) turnover rate (RBC life span of the mouse: 38-47 days / RBC life span of the rat: 42-65 days). Always taken into consideration must be:

• The species to be sampled
• The size of the animal to be sampled
• The age and health of the animal to be sampled
• The effects of handling stress
• The collection site
• The frequency of sampling necessary
• The training and experience of the personnel performing the collection
• The suitability of sedation and/or anesthesia
• The minimum volume required for analysis. The maximum permitted blood volume includes blood lost during collection. As a general rule, 20 drops = 1 mL (i.e. 5 drops = 250 uL)

When collecting blood it is very important that the handler is able to recognize signs of shock and anemia. The combined effect of sample volume and sample frequency without appropriate fluid replacement can cause an animal to go into hypovolaemic shock or become anemic. Packed cell volume, haemoglobin level, red blood cell and reticulocyte counts should be monitored throughout a series of bleeds using the results from the first sample from each animal as the baseline for the animal.

• Signs of hypovolemic shock include a fast and thready pulse, pale dry mucous membranes, cold skin and extremities, restlessness, hyperventilation, and a sub-normal body temperature.
• Signs of anemia include pale mucous membranes of the conjunctiva or inside the mouth, pale tongue, gums, ears or footpads (non-pigmented animals), intolerance to exercise and with severe anemia, increased respiratory rate when at rest.

If>10% blood volume is required, it is recommended to replace collected blood volume by 3–4 times the volume of blood collected with isotonic fluids (i.e. fluids with same tonicity as blood, such as 0.9% saline, 5% dextrose or Lactated Ringer’s solution).

The Circulating Blood Volume (CBV) of an adult mouse is ~72 ml/kg (0.072ml/g).

• 1% (maximum) of the CBV can be collected every 24 hours.
• 7.5% (maximum) of the CBV can be collected in a single collection, once per week.
• 10% (maximum) of the CBV can be collected in a single collection, once per every 2 weeks.
• 15% (maximum) of the CBV can be collected in a single collection, once per every 4 weeks.

To calculate blood collection volumes:
Body weight x Circulating Blood Volume = Total Blood Volume (TBV)

• TBV x % (based on desired frequency of collection) = allowable volume to be collected.
• i.e. For a single collection once per week: 20 g x 0.072 ml/g = 1.44 ml/g then 1.44 x 0.075 (7.5% for once per week sample) = 0.1 ml is the maximum allowable volume.

This schedule allows for recovery time for the animals as illustrated in the following table:

2. Materials:

• Heat lamp
• Restrainers as required
• Sterile syringes
• Sterile needles (multiple sizes ranging from 23-30g)
• Sterile scalpel blades
• Lancets
  • <20g = 4.0 mm
  • 20g – 40g = 5.0 mm
  • >40g = 5.5 mm
• Gauze
• Alcohol swabs
• Petroleum jelly
• Collection tubes
• Isotonic fluids such as Lactated Ringers or 0.9% NaCl

3. Procedures:

• Only University Animal Care Committee (UACC) approved blood collection techniques can be performed.
• The minimal volume required should be collected at all times.
• All collections should be performed by trained and competent individuals.
• The smallest needle size for the collection location (avoiding hemolysis) should be used.
• Each and every animal requires a new sterile syringe and a new sterile needle/lancet. Prepare all equipment in advance.
• Only three attempts per site should be practiced. If unsuccessful, allow another trained person to collect the sample.
• Apply pressure with gauze until hemostasis occurs.

Tail Vein (intravenous)

• Each and every animal requires a new sterile needle/lancet.
• Place mouse in restraining tube.
• Swab the tail with alcohol.
• Using a heat lamp, direct the heat only to the tail for 5-10 seconds for vasodilation.
• Hold tail and locate lateral tail veins, refer to Figure 5.
• Insert 25 gauge needle at a 10o angle to puncture the vein.
• For serial collections, a small nick in the tip of the tail using a sharp, sterile scalpel blade is permitted. This technique works well for clot dislodgment and repeat sampling. Hemostasis must be ensured.
• Withdraw needle and collect blood drops in capillary tube.
• Apply pressure with gauze until hemostasis occurs.