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NC3Rs: National Centre for the Replacement Refinement & Reduction of Animals in Research
Guidance

Blood sampling: Rat

Approaches for sampling blood in the rat, covering non-surgical, surgical and terminal techniques.

General principles

You should read the general principles of blood sampling page before attempting any blood sampling procedure.

Decision tree

The volume of blood removed and the frequency of sampling should be based on the purpose of the scientific procedure and the total blood volume of the animal. As a general principle, sample volumes and number of samples should be kept to a minimum. Further advice is given in the general principles.

How much blood does a rat have?

On average, rats have around 64 ml of blood per kg of bodyweight.

A rat weighing 400 g would therefore have a total blood volume (TBV) of approximately 64 ml/kg x 0.4 kg = 25.6 ml.

How to decide on the most appropriate blood sampling technique for rats?

The two tables below are designed to assist in determining the amount of blood to sample from the animal, and depending on that volume, the most appropriate techniques to use.

1. Do you require more than one blood sample from the same rat?

YES

NO

Maximum <10% TBV (= 2.56 ml) on any single occasion AND <15% TBV (= 3.84 ml) in 28 days

Maximum <10% TBV (= 2.56 ml)

For repeat bleeds at short intervals, suggested limit <1% TBV (= 0.25 ml) in 24 hours AND consider catheterisation

OR terminal sample under general anaesthesia (volume unrestricted)

2. How much blood do you require?

 

Total of <3.0 ml

Total of <3.0 ml

Total of >3.0 ml

General anaesthesia required

General anaesthesia not used

General anaesthesia required; non recovery

Blood vessel catheterisation

 

 

 

Saphenous vein

Tail vein / Temporary cannula

Jugular vein

Sublingual vein*

 

Cardiac puncture

Abdominal / thoracic blood vessel

Retro-orbital**

Decapitation**

* Not widely used [1]

** Blood may be mixed with tissue fluid

Microsampling

Advances in bioanalytical techniques have opened up the potential to use smaller sample volumes (microsamples of ≤50µl) to assess drug and chemical exposure in blood, plasma and/or serum.

Information on microsampling (e.g. study designs, sampling protocols, videos) can be found in our dedicated microsampling resource. The technique for sampling is as described for tail vein below.

 

Resources and references 

  1. Zeller W et al. (1998). Refinement of blood sampling from the sublingual vein of rats. Laboratory Animals 32(4): 369-76doi: 10.1258/002367798780599910

  2. Diehl KH et al. (2001). A good practice guide to the administration of substances and removal of blood, including routes and volumes. Journal of Applied Toxicology 21(1): 15-23. doi: 10.1002/jat.727

  3. Parasuraman S et al. (2010). Blood sample collection in small laboratory animals. Journal of Pharmacology and Pharmacotherapeutics 1(2): 87-93. doi: 10.4103/0976-500X.72350

  4. Powles-Glove N et al. (2014). Assessment of toxicological effects of blood microsampling in the vehicle dosed adult rat. Regulatory Toxicology and Pharmacology 68(3): 325-31. doi: 10.1016/j.yrtph.2014.01.001

Blood vessel catheterisation (surgical)

Technique

Blood vessel catheterisation should be considered when repeated samples are required, as it avoids multiple needle entries at any one site. It is suitable for use in all strains of rat and can be used to take blood from the femoral artery and vein, carotid artery, jugular vein, vena cava and dorsal aorta. Surgery is required, using aseptic technique to prevent post-operative infection, and appropriate anaesthesia and analgesia to minimise pain. Rats should be allowed to regain their pre-operative body weight before blood samples are taken.

The catheter is surgically implanted and connected to a transcutaneous skin button that is secured at the dorsal midline scapula region. Use of a transcutaneous skin button is preferable to a jacket or harness as it has been shown to have a lower incidence of adverse effects. Transcutaneous skin buttons with septums are closed systems, eliminating the need to keep animals tethered during periods when animals are not being sampled from.

Animals that have been implanted with a catheter and transcutaneous skin button can be group-housed immediately after surgery (with use of a protective cap). Additionally, skin buttons have no impact on the animal's movement. Because the transcutaneous skin button is a closed system, caging, bedding and environmental enrichment do not need to be modified due to concern over contamination. Skin buttons should always be accessed utilising appropriate aseptic technique to prevent contamination of the catheter.

In comparison, jacket and tether systems can restrict free movement and rats may need to be housed singly after surgery, which will further negatively affect their welfare. The caging, bedding and environmental enrichment need to be appropriate to prevent the tether becoming entangled and the wound contaminated. In addition, the bedding needs to be sand free.

Catheter selection is dependent upon the vessel to be catheterised, and expected duration of patency. For example, for longer-term patency, a rounded tip polyurethane catheter would be an ideal option. For a short-term study, a square tip polyurethane catheter would be acceptable. Consideration should be given to catheter material and potential for evaporation.

Routine flushing and catheter maintenance should be performed (no more than once per week) for animals implanted with a catheter and transcutaneous button. Solutions infused into the catheters should always be sterile and pharmaceutical grade when possible.

Small catheters will increase the risk of blood clotting (large catheters can abrade the blood vessel wall). To prevent this, the catheter requires regular maintenance (e.g. regular flushing with an appropriate flush solution. See our preventing thrombosis page for more information).

Blood should be collected aseptically. Usually, 0.1 - 0.2 ml can be taken per sample. Depending on the sample volume and scientific purpose, up to six samples over a two hour period or up to 20 samples over a 24-hour period may be taken. Sterile saline with anticoagulant should be flushed into the catheter after blood sampling to prevent the blood from clotting. Aseptic technique is more easily followed with the use of a transcutaneous skin button with septum; maintaining sterility with an open system and pin requires careful effort with a heightened risk of contamination. 

A pharmaceutical grade sterile locking solution should be used to lock the catheter after a series of samples have been taken, allowing flushing to be avoided for a number of days.

The following should be checked daily for all animals implanted with a subcutaneous skin button and catheter:

  • Animal demeanor (bright, alert, responsive).
  • Hydration status.
  • Food consumption. 
  • Normal urine/fecal output.
  • Incision site(s) appears to be healing well, without evidence of swelling, inflammation, infection, or skin abrasions. 

*Daily patency checks are not needed for transcutaneous skin buttons. 

**Animal weight can be monitored per facility protocol, but weight loss is not a standard observation for this procedure.

Changes in any of the above may require veterinary advice or treatment, or may indicate that a humane endpoint has been reached and appropriate action should be taken.

Summary

Consideration Recommendation
Number of samples It is recommended, up to six samples may be taken in a two hour period or up to 20 samples over a 24 hour period, depending on sample volume.
Sample volume 0.1 - 0.2 ml
Equipment
  • Catheter, 1-3 French gauge (dependent upon vessel to be catheterised)
  • Transcutaneous skin button
  • Surgical supplies
  • Syringe
  • Mating injector to access skin button 
  • Sterile flush and lock solutions
Staff resource One person is required to take the blood sample. However, further staff resource is required for surgery, post-operative care for as long as necessary for the individual animal, and daily animal observations post-surgery.
Adverse effects
  • Infection: 1-5%, usually due to breach in aseptic technique during surgery or when accessing skin button. 
  • Obstructed catheter: 1-5%. 
  • Skin abrasions from scratching post-operatively: 1-5%. 
  • Seroma formation: 1-5%, should not impact catheter patency.

Be sure to use our advice on vascular catheters to reduce the incidence of adverse effects.

Resources and references

  1. Feng J et al. (2015). Catheterization of the carotid artery and jugular vein to perform hemodynamic measures, infusions and blood sampling in a conscious rat model. Journal of visualized experiments: JoVE (95): 51881. doi: 10.3791/51881

  2. Gunaratna PC et al. (2004). An automated blood sampler for simultaneous sampling of systemic blood and brain microdialysates for drug absorption, distribution, metabolism and elimination studies. Journal of Pharmacological and Toxicological Methods 49(1): 57-64. doi: 10.1016/S1056-8719(03)00058-3

  3. Bardelmeijer HA et al. (2003). Cannulation of the jugular vein in mice: a method for serial withdrawal of blood samples. Laboratory Animals 37(3): 181-7. doi: 10.1258/002367703766453010

  4. Ling S and Jamali F (2003). Effect of cannulation surgery and restraint stress on the plasma corticosterone concentration in the rat: application of an improved corticosterone HPLC assay. Journal of Pharmacy and Pharmeutical Sciences 6(2): 246-51. PMID: 12935437

  5. Nolan TE and Klein HJ (2002). Methods in vascular infusion biotechnology in research with rodents. Institute for Laboratory Animal Research journal 43(3): 175-82. doi: 10.1093/ilar.43.3.175

  6. Bellinger D (2015). Harness versus button device with automated blood sampler. Poster presentation, 66th AALAS National Meeting, Phoenix, AZ, 1-5 November.

  7. Hagstedt T (2019). Improved chronic vascular catheterization in unrestrained conscious rats. Poster presentation, AstraZeneca Global 3Rs Award event.

Tail vein (non-surgical)

Technique

Tail vein sampling is suitable for all strains of rat. With proper training, the procedure is quick and simple to perform. However, this technique requires the rats to be warmed in order to dilate the blood vessel prior to taking the sample. This can be stressful and can cause dehydration due to salivation, in addition to increasing metabolic rate, which may affect the experimental data. As a result, other routes such as saphenous vein sampling should be considered where possible. View the tail vein sampling technique below.

The lateral tail vein is usually used and 0.1 - 2 ml of blood can be obtained per sample depending on the size of the rat, sampling frequency and scientific justification. The tail may need to be washed with diluted Hibiscrub (1%) in order to see the blood vessel. Finger pressure 5 cm from the tail tip can enhance the visibility of the tail vessels.
 

To avoid bruising and damage to the tail, normally no more than eight blood samples should be taken per session and in any one 24-hour period. Where it is necessary and justifiable to take more, the use of temporary cannulation or surgical catheterisation should be considered. The number of attempts to take a blood sample should be minimised (no more than three needle sticks in any one attempt) and sufficient time should be given for the tail to recover between blood sampling sessions. Alternate sides of the tail should be used and needle punctures moved towards the tail base.

If it is necessary to warm the animal, a warming cabinet should be used (38oC for up to 10 minutes). Rats should be carefully monitored, including checking for signs of hyperthermia and dehydration. The time the rat is in the warming cabinet should be recorded and the cabinet should be calibrated regularly to avoid hyperthermia; digital displays should not be relied upon. It is important to ensure the temperature in the cabinet is uniform and that there are no 'hot spots'. Alternatively, a warm bath at a maximum of 38oC can be used to warm just the tail of the rat. The temperature of the bath should be monitored to prevent the tail being scalded.

The lateral tail vein is usually accessed approximately one-third along the length of the tail from the tail tip, moving towards the base of the tail for multiple samples. Blood samples should only be taken from the base of the tail if no vein is visible elsewhere. Taking the first sample/s from the proximal end of the tail can result in a perivascular clot and inflammation that significantly reduces blood flow to the distal portion of the vessel.

Proper aseptic technique should be used throughout the procedure. Untrained rats will need to be restrained which can cause stress; therefore, the duration of restraint should be minimised. Habituated rats are much less stressed by manual restraint than by restraint in a tube.
 

A rat that has been trained to undergo tail vein blood collection. Following training and acclimatisation, the rat appears calm and cooperative and dosen’t require restraint during the procedure.

Blood flow should be stopped by applying finger pressure to the soft tissue. A finger should be placed at the blood sampling site for approximately 30 seconds before the animal is returned to its cage.

Summary

Consideration Recommendation
Number of samples No more than eight blood samples should be taken per session and in any 24-hour period, depending on sample volume.
Sample volume 0.1 - 2 ml
Equipment 21G - 23G needle or butterfly needle or lance
Staff resource One person is required to take the blood sample if a tube restrainer is used. Two people are required if the rat is held for sampling.
Adverse effects
  • Infection <1%
  • Haemorrhage <1%
Other Rats may be warmed to dilate the blood vessel. Care should be taken to avoid hyperthermia and dehydration.

Resources and references

  1.  Lee G and Goosens KA (2015). Sampling blood from the lateral tail vein of the rat. Journal of visualized expepriments: JoVE 99: e52766. doi: 10.3791/52766

  2. Diehl KH et al. (2001). A good practice guide to the administration of substances and removal of blood, including routes and volumes. Journal of Applied Toxicology 21(1): 15-23. doi: 10.1002/jat.727

  3. Morton DB et al. (1993). Removal of blood from laboratory mammals and birds. Lab Animal 27 (1): 1-22. doi: 10.1258/00236779378108241

  4. Fluttert M et al. (2000)A refined method for sequential blood sampling by tail incision in rats. Laboratory Animals 34(4):372-8. doi: 10.1258/002367700780387714

  5. Lee G and Goosens KA (2015). Sampling Blood from the Lateral Tail Vein of the Rat. Journal of Visualized Experiments: JoVE  (99): 52766. doi: 10.3791/52766

  6. Zou W et al. (2017). Repeated blood collection from tail vein of non-anesthetized rats with a vacuum blood collection system. Journal of Visualized Experiments: JoVE (130): 55852. doi: 10.3791/55852

Jugular vein (non-surgical)

Technique

Sampling from the jugular vein can be used with all strains but requires a high degree of competence to avoid harm to the rat. Its use should be limited, for instance, to studies where blood collection is required immediately after dosing (e.g. inhalation and infusion studies) or where a sampling site distal to the dosing site is required (e.g. intravenous studies). Warming of the rat is not required.

One person is required to take the blood sample and another to restrain and monitor the rat. If necessary the person taking the blood sample can restrain the head by the use of a head cap. Blood is taken from a small triangular patch of skin just under the scapula. The head is tilted at an angle in the head cap, which makes the site of sampling prominent. If required visualisation of the sampling area can be improved by trimming the animal's fur. Single person jugular sampling is also possible, though this does require a very high level of skill.

Sampling should be carried out aseptically. 0.1 - 2 ml (normally 0.1 - 0.3 ml) of blood can be collected per sample and, depending on the sample volume and scientific justification, up to eight samples in a 24-hour period. The number of needle sticks at each attempt should be a limited to three. If more samples are needed, then surgical catheterisation or temporary cannulation of a different blood vessel should be considered. 

Blood flow should be stopped before the rat is returned to its cage by applying gentle pressure to the blood sampling site for thirty seconds.

Summary

Consideration Recommendation
Number of samples No more than eight blood samples should be taken in a 24-hour period.
Sample volume 0.1 - 2 ml (normally 0.1 - 0.3 ml)
Equipment 23G ( 1" long) needle
Staff resource Two people: one to take the blood sample and another to restrain and monitor the rat.
Adverse effects
  • Bruising
  • Infection <1%
  • Haemorrhage <1%
Other A high degree of competence is required to perform this technique.

Resources and references

  1. Laboratory animal welfare training exchange (2007). Jugular Blood Collection in a Conscious Rat.

  2. Lucas RL et al. (2004). Collection and preparation of blood products. Clinical techniques in small animal practice 19(2): 55-62. doi: 10.1053/j.ctsap.2004.01.007

  3. Thrivikraman KV et al. (2002). Jugular vein catheterization for repeated blood sampling in the unrestrained conscious rat. Brain research protocols 10(2): 84-94. doi: 10.1016/s1385-299x(02)00185-x

  4. Goldkuhl R et al. (2010). Plasma concentrations of corticosterone and buprenorphine in rats subjected to jugular vein catheterization. Laboratory animals 44(4): 337-43. doi: 10.1258/la.2010.009115

Saphenous vein (non-surgical)

Technique

Sampling from the lateral saphenous vein is a relatively quick method of obtaining blood samples from all strains of rat. It does not require the rats to be warmed.

Slides and videos of restraint, shaving and sampling for this technique with mice are available on the websites of the Norwegian Reference Centre for Laboratory Animal Science and Alternatives.

Blood is collected from the lateral saphenous vein which runs dorsally and then laterally over the tarsal joint. To improve visualisation of the vein the sample site can be shaved with electric clippers. Shaving need only be repeated as the hair grows again.

The rat should be restrained either manually or using a restraint tube. This can cause stress and therefore the duration of restraint should be minimised. Where a restraint tube is used, it should be appropriate to the size of the rat. All forms of restraining equipment should be frequently washed to prevent pheromonally-induced stress or cross-infection.

To collect blood, the hind leg should be immobilised in the extended position by applying gentle downward pressure immediately above the knee joint. This stretches the skin over the ankle, making it easier to clip and immobilise the saphenous vein. Please note that hair removal by shaving with a scalpel blade is no longer recommended as it removes the epidermal layers of the skin. Aseptic technique should be used. Anaesthesia is not necessary but may be used on welfare grounds for animals that are difficult to hold. Where sedatives contain peripheral vasodilators, doses should be low to avoid prolonged bleeding from the puncture site. The number of attempts to take a blood sample should be minimised (no more than three needle sticks in any one attempt). Blood is collected by capillary action into a haematocrit tube or passively into a tube.

Blood flow can be stopped by gentle finger pressure over the puncture site, or simple relaxation of operator's grip on the animal's leg. Animals should not be returned to their cage before the blood flow has stopped. 

No more than four blood samples should be taken in any 24-hour period. If more samples are needed, then surgical catheterisation or temporary cannulation should be considered. For multiple samples, the scab or the blood clot should be removed.

Rats can show temporary favouring of the limb following sampling from the saphenous vein.

Summary

Consideration Recommendation
Number of samples No more than four blood samples should be taken within any 24-hour period
Sample volume Up to 0.2 ml may be taken for a single sample, which can usually be repeated at 2-week intervals without disturbances to haematological status. Alternatively, multiple smaller samples (e.g. 0.02 ml daily) may be drawn, taking into account limits on sample volume.
Equipment 23G needle or lance
Staff resource One person if a restraint tube is used. Where manual restraint is used, two people are required: one for handling the rat and one for taking the blood sample.
Adverse effects
  • Bruising
  • Haemorrhage
  • Infection
  • Temporary favouring of the limb

Resources and references

  1. Diehl KH et al. (2001). A good practice guide to the administration of substances and removal of blood, including routes and volumes. Journal of applied Toxicology 21(1): 15-23. doi: 10.1002/jat.727

  2. Van Herck H et al. (2001). Blood sampling from the retro-orbital plexus, the saphenous vein and the tail vein in rats: comparative effects on selected behavioural and blood variables. Laboratory Animals 35(2): 131-9. doi: 10.1258/0023677011911499

  3. Luzzi M et al. (2005). Collecting blood from rodents: a discussion by the Laboratory Animal Refinement and Enrichment ForumAnimal Technology and Welfare 4(2): 99-102.   

  4. Hem A et al. (1998). Saphenous vein puncture for blood sampling of the mouse, rat, hamster, gerbil, guinea pig, ferret and mink. Laboratory animals 32(4): 364-8. doi: 10.1258/002367798780599866

Temporary cannula (non-surgical)

Technique

Temporary cannulation of the lateral tail vein should be considered when repeated blood samples are required over a short period of time (e.g. a few hours) as it avoids multiple needle entries and the associated damage to the tail vein. The technique may be combined with normal tail vein bleeding to accommodate all the blood samples required by a given protocol but reduce the number of needle entries and reduce the time the rat must spend in a warming cabinet (since warming may not be necessary for taking blood via the temporary cannula). It is suitable for use in all strains of rat, and the animals can be group-housed during the study period. 

Tail bleeding normally requires the rats to be warmed in order to dilate the blood vessel prior to taking the sample. This may be stressful and can cause dehydration due to salivation, in addition to increasing metabolic rate, which may affect the experimental data. Restraining the rat for long periods of time in a restraint tube may also cause it to become hot, which will be stressful to the rat and may affect the blood parameters. View this technique below.
 

No surgery is required. An intravenous catheter is inserted into the vein by puncture of the skin and is taped in situ. A heparin flush is used (0.1 ml) after placement and between samples to prevent clotting. An access port is inserted into the exteriorised end of the cannula, which stops the blood from flowing, and the catheter is taped into place.

An appropriate aseptic technique should be used during the procedure. The tail may need to be washed in order to visualise the blood vessel.

0.1 - 2.0 ml (normally 0.1 - 0.3 ml) can be taken per sample, and depending on the sample volume and consideration of the effects of repeated warming and restraint, no more than six samples in a two hour period or eight samples in a 24-hour period.

The lateral tail vein is usually accessed approximately one-third along the length of the tail from the tail tip. Cannulation should not be attempted at the base of the tail, as this could result in a perivascular clot and inflammation that significantly reduces blood flow to the distal portion of the vessel. If cannulation is unsuccessful, direct venepuncture may be used as the alternative. The number of attempts to take any blood sample should be minimised (no more than three needle sticks in any one attempt).

warming cabinet is used prior to the cannulation (39oC for up to 10 minutes). Subsequent warming prior to sampling may not be required so long as blood is 'free flowing'. If necessary, the rat can be warmed for a short period (up to five minutes) as required. The rat should be carefully monitored, including checking for signs of hyperthermia and dehydration. The time the rat is in the warming cabinet should be recorded and the cabinet should be calibrated regularly to avoid hyperthermia; digital displays should not be relied upon. It is important to ensure the temperature in the cabinet is uniform and that there are no 'hot spots'. Alternatively, a warm bath at a maximum of 39oC can be used to warm just the tail of the rat. The temperature of the bath should be monitored otherwise the tail can be scalded.

Rats need to be restrained which can cause stress and therefore the duration of restraint should be minimised. Restraint can either be manual or using a restraint tube. Anecdotal evidence suggests that holding the rat is less stressful than using a restraint tube. Where a restraint tube is used, it should be appropriate to the size of the rat in order to avoid damage to the tail, testes, limbs and back. All forms of restraining equipment should be frequently washed to prevent pheromonally-induced stress or cross-infection.

Summary

Consideration Recommendation
Number of samples Ideally, no more than six samples should be taken in a two hour period, or eight samples in a 24-hour period, depending on sample volume and consideration of the effects of repeated warming and restraint. 
Sample volume 0.1 - 2.0 ml (normally 0.1 - 0.3 ml)
Equipment 22G 0.90 mm i.v. catheter
Staff resource One person is required to take the blood sample if a tube restrainer is used. Two people would be required if one were to restrain the rat manually. For large batches of animals, two people are required: one person to take the blood sample and one to operate the warming cabinet.
Adverse effects
  • Bruising
  • Infection <1%
  • Haemorrhage <1%
Other Rats are warmed to dilate the blood vessel and care should be taken to avoid hyperthermia and dehydration. The time the animal is exposed to warming and restraint should be kept to a minimum.

Resources and references

  1. Lee G and Goosens KA (2015). Sampling blood from the lateral tail vein of the rat. Journal of visualized experiments: JoVE (99): 52766. doi: 10.3791/52766

  2. Ling S and Jamali F (2003). Effect of cannulation surgery and restraint stress on the plasma corticosterone concentration in the rat: application of an improved corticosterone HPLC assay. Journal of Pharmacy and Pharmaceutical Sciences 6(2): 246-51.  PMID: 12935437

  3. Nolan TE and Klein HJ (2002). Methods in vascular infusion biotechnology in research with rodents. Institute for Laboratory Animal Research journal 43(3): 175-82. doi: 10.1093/ilar.43.3.175

Retro-orbital (terminal)

Technique

Retro-orbital bleeding should only be performed under terminal anaesthesia because of the severity of adverse effects that can occur with this technique, even in skilled hands (summarised below).

Also referred to as peri-orbital, posterior-orbital and orbital venous sinus bleeding. 

Blood is collected from the venous sinus. The rat is restrained, the neck gently scruffed and the eye made to bulge. A capillary tube/pipette is inserted medially, laterally or dorsally. Blood is allowed to flow by capillary action into the capillary tube/pipette. The sample obtained is a mixture of venous blood and tissue fluid, and is not representative of venous blood.

Summary

Consideration Recommendation

Number of samples

One 

Sample volume

>3ml when performed as a terminal procedure

Equipment

A glass capillary tube or Pasteur pipette.

Staff resource

One person is required to take the blood sample.

Other

Procedure should be carried out under terminal anaesthesia.

Adverse effects
  • Retro-orbital haemorrhage resulting in haematoma and excessive pressure on the eye
  • Corneal ulceration, keratitis, pannus formation, rupture of the globe and micro-ophthalmia caused by proptosis of the globe
  • Damage to the optic nerve and other intra-orbital structures which can lead to deficits in vision and blindness
  • Fracture of the fragile bones of the orbit and neural damage by the micro-pipette
  • Penetration of the eye globe itself with a loss of vitreous humour

Resources and references

  1. Jo EJ et al. (2021). Comparison of murine retroorbital plexus and facial vein blood collection to mitigate animal ethics issues. Laboratory Animal Research 37(1): 12. doi: 10.1186/s42826-021-00090-4 

  2. Harikrishnan VS et al. (2018). A comparison of various methods of blood sampling in mice and rats: Effects on animal welfare. Laboratory Animals 52(3): 253-64. doi: 10.1177/0023677217741332 

  3. Sharma A et al. (2014). Safety and blood sample volume and quality of a refined retro-orbital bleeding technique in rats using a lateral approach. Lab Animal 43(2): 63-6. doi: 10.1038/laban.432

  4. Luzzi M et al. (2005). Collecting blood from rodents: a discussion by the Laboratory Animal Refinement and Enrichment ForumAnimal Technology and Welfare 4(2): 99-102. 

  5. Van Herck H et al. (1992). Histological changes in the orbital region of rats after orbital puncture. Lab Animal 26(1): 53-8. doi: 10.1258/002367792780809048

  6. Van Herck H et al. (1998). Orbital sinus blood sampling in rats as performed by different animal technicians: the influence of technique and expertise. Lab Animal 32(4): 377-86. doi: 10.1258/002367798780599794

Abdominal/thoracic blood vessel (terminal)

Technique

Appropriate for all strains of rat, this is a suitable technique to obtain a single, large, good-quality blood sample from a euthanised rat or a rat under terminal anaesthesia. A sample size of 5-10 ml can be collected from the hepatic portal vein, or 10-15 ml from other abdominal/thoracic vessels, depending on the size of the rat. As the heart is not punctured, this technique can be used where it is necessary to avoid cardiac damage.

Blood is collected either from the abdominal aorta, caudal or dorsal aorta, vena cava or hepatic portal vein which are accessed via a laparotomy or thoracotomy. Removal of connective tissue and application of finger pressure is necessary to dilate the vessel. Blood should be withdrawn slowly to prevent the vessel collapsing. Deep surgical anaesthesia is necessary. 

Summary

Consideration Recommendation
Number of samples One
Sample volume Up to 10 ml from the hepatic portal vein, or 15 ml from other abdominal/thoracic vessels, depending on the size of the rat.
Equipment 19 - 21G needle
Staff resource One person is required to take the sample.

Resources and references

  1. Parasuraman S et al. (2010). Blood sample collection in small laboratory animals. Journal of Pharmacology and Pharmacotherapeutics 1(2): 87-93. doi: 10.4103/0976-500X.72350

  2. Morton DB et al. (2001). Refining procedures for the administration of substances. Laboratory animals 35(1): 1-41. doi: 10.1258/0023677011911345

Cardiac puncture (terminal)

Technique

Cardiac puncture should not be used if the peritoneum needs to be lavaged to harvest cells, as this technique can cause blood to escape into the peritoneal cavity.

Cardiac puncture is a suitable technique to obtain a single, large, good quality sample from a euthanised rat or a rat under deep terminal anaesthesia if coagulation parameters, a separate arterial or venous sample or cardiac histology are not required. It is appropriate for all strains of rat.

A sample of 10 - 15 ml of blood can be obtained depending on the size of the rat and whether the heart is beating. Blood samples are taken from the heart, preferably the ventricle, which can be accessed either via the left side of the chest, through the diaphragm, from the top of the sternum or by performing a thoracotomy. Blood should be withdrawn slowly to prevent the heart collapsing.

Summary

Consideration Recommendation
Number of samples One
Sample volume Up to 15 ml
Equipment 19G - 21G needle
Staff resource One person is required to take the blood sample.

Resources and references

  1. Beeton C et al. (2007). Drawing blood from rats through the saphenous vein and by cardiac puncture. Journal of visualized experiments: JoVE (7): 266. doi: 10.3791/266

  2. Parasuraman S et al. (2010). Blood sample collection in small laboratory animals. Journal of Pharmacology and Pharmacotherapeutics 1(2): 87-93. doi: 10.4103/0976-500X.72350  

  3. Morton DB et al. (2001). Refining procedures for the administration of substances. Laboratory animals 35(1): 1-41. doi: 10.1258/0023677011911345

Schedule 1 stunning and decapitation (terminal)

Technique

Although suitable for all rat strains, this technique should only be used in rare circumstances and where there is exceptional scientific justification.

Further, the ASPA limits its use to rodents of body weight 1kg or less. The primary reason for using this technique is to obtain a large volume of blood that has not been affected by anaesthetic drugs or carbon dioxide. A large volume of blood can be collected from the trunk if necessary, but it should be noted there is a risk of contamination from other body fluids and tissues.

In order to be deemed a Schedule 1 method of euthanasia, rats which have been stunned must be determined as dead before decapitation can take place (e.g. via confirmation of cessation of circulation or exsanguination - see Section 1(4) of the amended ASPA). This method should only be carried out by people competent in this method for the species and size of the animal. Training for stunning and decapitation should be undertaken on dead animals.

Summary

Consideration Recommendation
Number of samples One
Sample volume Up to 10 ml
Equipment Suitable sharp instrument to decapitate, (e.g., guillotine or sharp scissors).
Staff resource One person is required to take the blood sample.
Other A high level of expertise is required for this technique.

Decapitation (terminal)

Technique

This technique should only be used in exceptional circumstances. In the UK this technique is not a Schedule 1 method of euthanasia and therefore personal and project licence authority is required.

Trunk blood is collected from the site where the animal is decapitated, under deep terminal anaesthesia. It should be noted there is a risk of contamination from other body fluids and tissues. Training for decapitation should be undertaken on dead animals.

Summary

Number of samples One
Sample volume Up to 10 ml
Equipment Suitable sharp instrument to decapitate (e.g. guillotine or sharp scissors).
Staff resource One person is required to take the blood sample.
Other A high level of expertise is required for this technique

You should read the general principles of blood sampling page before attempting any blood sampling procedure.

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