Compound Administration IV

Lab Animal Research

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Overview

Source: Kay Stewart, RVT, RLATG, CMAR; Valerie A. Schroeder, RVT, RLATG. University of Notre Dame, IN

There are many commonly used routes for compound administration in laboratory mice and rats. Protocols may, however, require the use of the less commonly used routes: intracardiac, footpad, and retro-orbital injections. Specialized training is essential for these procedures to be performed successfully. Justification for these routes may need to be provided to gain Institutional Animal Care and Use Committee (IACUC) approval.

Cite this Video

JoVE Science Education Database. Lab Animal Research. Compound Administration IV. JoVE, Cambridge, MA, (2017).

Principles

Intracardiac administration has been used in a variety of applications, including the development of an animal model of bone cancer metastasis, as well as an examination of the effects of direct intracardiac delivery on the outcome of myocardial infarction. This procedure is often done through the use of an ultrasound to guide the needle into the correct location in the heart.2 However, when performed correctly utilizing the proper landmarks, this procedure can be performed without the use of ultrasound visualization.

Due to the invasive nature of the procedure, the use of intracardiac injection must be scientifically justified in an IACUC protocol. Only one survival injection should be permitted. This procedure requires the use of a general anesthetic, either inhalant or injectable, as per the guidelines established within an organization. Needle selection should be the smallest size possible that will allow for the viscosity of the material injected; generally, a 27-30 gauge needle is used. Injection volumes range from 100 µL to a maximum of 300 µL.

Intravenous injections in the tail of mice are both challenging and often unsuccessful. An alternate route of intravenous administration is through the retro-orbital plexus. While this technique necessitates training and skill to perform, studies have shown that there is a higher success rate with the retro-orbital injection than with lateral tail vein injection.3, 4, 5 Anesthesia is required to prevent the mouse from moving during the procedure. General inhalant anesthesia delivered either via a bell jar or an induction chamber attached to a precision vaporizer is effective. However, if inhalant will be used, be aware that the animal will begin to recover quickly once it is removed from the chamber, so one must be ready to perform the injection. A topical ophthalmic anesthetic (tetracaine or proparacaine) is recommended when multiple injections are to be performed.

The orbital venous structure of the mouse and rat are different. The mouse has a sinus or convergence of several vessels, including the supraorbital vein, dorsal nasal vein, the inferior palpebral vein, and the superficial temporal veins that fill the space in the orbit around the eye. In the rat orbital area, there is a network or plexus of vessels. As with all injections, the needle selected should be the smallest size possible; generally a 27-30 gauge needle. Although there have been reports of larger volumes, the maximum volume is 150 µL per eye.3, 4, 5 One injection per eye, per day, is recommended, with a total of two injections per eye for survival procedures. Also, there should be at least a one-day interval between injections. For a nonsurvival procedure, volumes up to 500 µL can be administered.

Despite the controversy, the use of the foot pad as an injection site is still required for some studies. It has been demonstrated that when injected via the foot pad, the antibody response in some mouse strains was significantly stronger than when injected into the hock.6 All animals must be closely monitored for signs of pain, level of food consumption, and for normal ambulation. Self-mutilation of the foot can occur to the extent of the foot being destroyed. This is a sign of chronic pain. Any animal demonstrating self-mutilation should be called to the attention of the veterinary staff immediately.

Footpad measurements should be done daily as soon as obvious swelling has occurred. Endpoints must be in place according to IACUC guidelines. Generally, the animal must be euthanized when the lesion or tumor interferes with the animal's ability to ambulate or reach food and water. The maximum volume that can be injected into a footpad is 50 µL. A 29-30 gauge needle is recommended for the injection.

Procedure

1. Intracardiac injection

  1. Landmarks and positioning: Position the mouse or rat either in right lateral recumbency (with the left side facing upward) or in dorsal recumbency, and identify the landmarks.
    1. Position the heart approximately level with the point of the elbow and just to the left of the sternum.
    2. Insert the needle between the ribs at the point of the elbow.
    3. In an animal in dorsal recumbency, insert the needle into the chest parallel to the table.
    4. In an animal in lateral recumbency, insert the needle into the chest perpendicular to the table.
  2. Injection of the article
    1. Ensure there is minimal movement of the needle once it is inserted into the heart to prevent the muscle from being torn and causing bleeding into the pericardial sac, as this compromises cardiac function.
    2. Aspirate the syringe to determine proper placement. There is often a flash of blood into the hub of the syringe.
    3. Hold the syringe so that the hand is not repositioned on the syringe for injection once the needle has been positioned in the heart.
    4. Inject the article in a slow and steady manner.
    5. Withdraw the needle slowly to minimize bleeding.
    6. Mice that experience respiratory distress, seizure, cyanosis, prolonged ataxia, or other sequelae related to injection trauma must be evaluated by the veterinarian staff and euthanized according to their recommendation.

Figure 1
Figure 1. Intracardiac injection in mice.

2. Intravenous injection utilizing the retro-orbital plexus

  1. Injection landmarks and positioning
    1. Place the animal on a flat surface in lateral recumbency.
    2. Insert the needle into the medial canthus of the eye at a 45° angle to the nose.
    3. The needle depth must be sufficient to penetrate the conjunctival tissues and advance behind the ocular orbit and into the ocular plexus. When placing the needle, it should not encounter the bone at the back of the orbit.
    4. Place the index finger on the top of the head with the thumb on the jaw.
    5. Gently pull back and downward to tighten the skin and protrude the eyeball.
    6. Take care not to apply pressure on the trachea and restrict air flow.
  2. Injection
    1. Place the needle behind the eye and not intra-ocularly.
    2. Ensure that the needle has minimal movement once inserted into the retro-orbital plexus, or the vessels will rupture causing bleeding and loss of the agent into the tissues behind the eye. Hold the syringe so that the hand is not repositioned on the syringe for injection once the needle has been positioned in the retro-orbital plexus.
    3. Do not aspirate with the syringe, as you will collapse the vessels.
    4. Inject the article in a slow and steady manner.
    5. Withdraw the needle and apply light pressure to the eye to control bleeding and to provide hemostasis.
    6. Mice that experience swelling, conjunctivitis, or other sequelae related to injection trauma must be evaluated by the veterinary staff and treated or euthanized according to their recommendation

Figure 2
Figure 2. Retro orbital injection in mice.

3. Footpad Injection

  1. For restraint, place the animal in a restraint tube with one hind foot isolated, and extend by grasping the skin above the stifle.
  2. Wipe the foot with water or alcohol to remove debris prior to injecting.
  3. Inject subcutaneously into the center of the hind foot forming a small bleb at the injection site. To avoid the blood vessel that runs the length of the foot, inject just off the midline parallel to the vessel.

Figure 3
Figure 3. Footpad injection in mice and rats.

Intracardiac, retro orbital and footpad are some of the specialized injection methods that biomedical researchers use for experiments necessitating delivery of compounds via these atypical routes.

An intracardiac injection delivers the compound into the left ventricle allowing the substance to directly enter the arterial circulation. The retro orbital route is an alternative to tail vein injection and is used to deliver the compound into the venous circulation. And a footpad injection involves subcutaneous administration of the article into the animal's hind foot. This video will illustrate the considerations, procedures and applications of these special injection techniques.

Let's begin with some background information and things one should consider before starting these administration procedures.

Intracardiac administration is often done through the use of an ultrasound to guide the needle into the correct location in the heart. However, if performed correctly utilizing the proper landmarks, the administration can be performed without the use of ultrasound visualization. Note that the procedure requires the use of a general anesthetic, and only one injection per animal is permitted for survival procedures. Generally a 27-30 gauge needle is used for this injection and maximum volume of administration is 100 and 300 microliters for mice and rats, respectively.

For intravenous injection via the retro orbital route, one should have a fair understanding of the orbital venous structure. A mouse has a sinus where several veins-namely the supraorbital, dorsal nasal, inferior palpebral, and superficial temporal-converge. Whereas in rats, there is a network or plexus of several veins. The injection is performed into the sinus or the plexus directly. Like intracardiac, this procedure also requires use of general anesthesia, and only one injection per eye per day is recommended with a total of two injections per eye for survival procedures. As with all injections, the smallest size needle should be selected-generally 27-30 gauge-and the recommended maximum volume is 150 μL per eye.

Despite the controversy, the use of footpad injection is still required for some studies, typically related to inflammation and tumor growth. Note that the injections can only be performed on one foot, never bi-laterally. And the footpad measurements should be done daily as soon as obvious swelling has occurred. A 29-30 gauge needle is recommended for the injection and the maximum volume recommended is 50 μL. Following any injection, all animals must be closely monitored for signs of pain, level of food consumption, and for normal ambulation. Generally the animal must be euthanized when the lesion or tumor interferes with the animal's ability to ambulate or reach the food and water.

Now let's learn the injection procedures, starting with the intracardiac injection. We will demonstrate the procedure in a mouse, but the landmarks and the protocol for a rat are similar.

The first step is to prepare the syringe. Recall a 29 gauge needle and 1 cc syringe is appropriate for mice. And the maximum volume for intracardiac injection is 100 microliters. When drawing the solution, leave a small amount of air between the plunger and the injection material. This is to allow for blood to enter the syringe as it is placed into the heart.

To start, anesthetize the animal using inhalant or injectable anesthetics. Review the considerations for maintaining general anesthesia in another video of this collection. Next, position the animal in dorsal recumbency position on an insulated platform. Then, tape the forelimbs to the platform and place a piece of tape horizontally across the abdomen above the hips. This is to further steady the animal and avoid any movement once the needle has been inserted. Next, using a swab, wet the animal's chest with 70% alcohol.

To pinpoint the injection site, first locate the xiphoid and the manubrium sternum. Then, find the midpoint between the two landmarks. 1-2 mm left of this point, is the needle insertion landmark. Using a cotton-tipped applicator, apply povidone iodine to mark the needle insertion site.

To inject, direct the needle perpendicular to the table, and insert it to the depth of about 2 mm. Then, apply a very slight backpressure to the plunger. A bright red oxygenated blood should enter the syringe hub, which confirms proper placement. Hold the syringe in same spot and inject the material slowly and steadily over the course of 30 to 60 seconds. Rapid injection can result in clumping of the cells and clogging of the arteries, a shock to the system due to the temperature of the substance being significantly lower than the body temperature, or an expansion of the ventricle and disruption of the heart rhythm.

Once the material has cleared the syringe, slowly and carefully remove the needle without any lateral movement as that can damage the heart muscles. Then immediately release the tape from the forelegs and abdomen and place the animal in prone position in a clean cage with sufficiently deep bedding to act as an insulating layer. Note that one half of this recovery cage is on a heating source and the anesthetized animal is situated on the heated side of the cage. This prevents hypothermia, and as the animal recovers from the anesthesia it will be able to move off the heated side as desired.

Next, let's learn the method for intravenous injection utilizing the retro-orbital plexus in rats. Again, we will demonstrate the procedure on a mouse, but the landmarks and the protocol for rats are similar.

Attach the appropriate needle to the selected syringe and fill in the injection material. Remember, generally one would use a 27-30 gauge needle with the smallest syringe possible and a maximum volume of 150 microliters.

To start the procedure, first anesthetize the animal. Then, place it on a flat surface in lateral recumbency position. Now place your index finger on the top of the head and the thumb on the jaw, and gently pull back and down. This is to tighten the skin and protrude the eyeball. Take care not to apply pressure on the trachea and restrict airflow. If planning multiple injections, apply topical ophthalmic anesthetic, like tetracaine or proparacaine.

Insert the needle into the medial canthus of the eye at a 45° angle to the nose. The depth must be sufficient to penetrate the conjunctival tissues and advance into the ocular orbit and into the sinus. It should not encounter the bone at the back of the orbit. To avoid rupturing of the blood vessels, ensure that the needle has minimal movement once inserted. Do not aspirate, as that will collapse the vessels. Inject the article in a slow and steady manner. Then, withdraw the needle gently and apply light pressure to the eye to control bleeding and to provide hemostasis.

Lastly, let's review footpad injection method in mice and rats. To start, attach the appropriate needle and fill the syringe with the correct volume. This procedure can be done in conscious animals.

Place the animal in a restraint tube with one hind foot isolated and extended by grasping the skin above the stifle. Wipe the foot with water or alcohol to remove debris prior to injecting. To avoid the blood vessel that runs the length of the foot, the injection landmark is at the center, but just off of the midline, closer to the toes.

Place the needle bevel up at the injection site directing it towards the heel. Inject the article slowly and steadily to avoid rapid distention of the foot tissues. This will cause the footpad to swell as the injection material fills that subcutaneous space. On a small animal's foot the swelling from the injection can extend to the heel, whereas in a larger animal it will be more localized.

After the injection, observe the animals daily and if persistent swelling is present or if there are lesions or tumors as a result of the experimental protocol, then, using a caliper, perform the footpad measurement. This instrument measures the foot thickness in millimeters and helps in quantitation of swelling.

Now let's discuss a few example experiments utilizing intracardiac, retro orbital and footpad injections.

One of the many applications of intracardiac administration is development of an animal model of cancer metastasis. Here, researchers used this route to inject tumor cells that possess propensity for bone colonization. In the following days, they studied tumor growth in bones using X-ray and fluorescence imaging techniques. In another study, the retro orbital route was used to inject specific antibodies that label neutrophils. Then, with help of intravital imaging, the scientists were able to track the migration pattern of the labeled cells.

Lastly, investigators often use footpad injection to analyze inflammatory response. In this experiment, researchers isolated peripheral blood mononuclear cells from human blood samples, mixed them with different test antigens and injected the solutions into the animal's footpad. Finally, they performed foot measurements to quantify the swelling response due to different antigens.

You've just watched JoVE's final installment on the usual and specialized compound administration techniques.

Just to recap, in the first part we reviewed the most common parenteral route. In the second chapter, we discussed the enteral and topical procedures. The third installment dealt with the first set of atypical procedures like intradermal, intranasal, and intracranial in neonates. Lastly, here we discussed three additional routes that biomedical researchers use in labs for specific purposes.

After watching this series you should have a much better understanding of different administration techniques and you should also know the general and specific considerations related to these protocols of compound administration As always, thanks for watching!

Summary

The administration of compounds into animals can have a significant effect on both the wellbeing of the animal and the outcome of the experimental data and scientific value. The proper method of delivery is essential to the success of the experiment. Many factors must be considered to determine the best route, including the scientific aim of the study, the pH of the substance, the required dosage volume, the viscosity of the substance, and the wellbeing of the animals. Technical expertise is also a requirement for all injection methods.

References

  1. Morton, D.A., Jennings, M., Buckwell, A., Ewbank, R., Godfrey, C., Holgate, B., Inglis, I., James, R., Page, C., Sharman, I., Verschoyle, R., Westall, L., and Wilson, A.B. 2001. Refining procedures for the administration of substances Report of the BVAAWF/FRAME/RSPCA/UFAW Joint Working Group on Refinement. Members of the Joint Working Group on Refinement. Laboratory Animals. 35. 1-41
  2. Prendiville , T.W., Qing, M., Lin, Z., Zhou, P., He, A., and Pu, W.T. 2014. Ultrasound-guided Transthoracic Intramyocardial Injection in Mice. Journal of Visualized Experiments. 90 | e51566.
  3. Yardeni, T., Eckhaus, M., Morris, H.D., Huizing, M., and Hoogstraten-Miller, S. 2001. Retro-orbital injection in mice. Lab Animal. 40:5. 155-171.
  4. Steel, C., Stephens, A., Hahto, S., Singletary, S., Ciavarra, R. 2008. Comparison of the lateral tail vein and the retro-orbital sinus as routes of intravenous drug delivery in a transgenic mouse model. Lab Anim. 37. 26-31.
  5. Timm, K.I. 1989. Orbital venous anatomy of the Mongolian gerbil, with comparison to the mouse, hamster, and rat. Laboratory Animal Science. 39:3. 262-265.
  6. Kamala, T. 2007. Hock immunization: a humane alternative to mouse footpad injections. Journal of Immunological Methods. 328. 204-214.

1. Intracardiac injection

  1. Landmarks and positioning: Position the mouse or rat either in right lateral recumbency (with the left side facing upward) or in dorsal recumbency, and identify the landmarks.
    1. Position the heart approximately level with the point of the elbow and just to the left of the sternum.
    2. Insert the needle between the ribs at the point of the elbow.
    3. In an animal in dorsal recumbency, insert the needle into the chest parallel to the table.
    4. In an animal in lateral recumbency, insert the needle into the chest perpendicular to the table.
  2. Injection of the article
    1. Ensure there is minimal movement of the needle once it is inserted into the heart to prevent the muscle from being torn and causing bleeding into the pericardial sac, as this compromises cardiac function.
    2. Aspirate the syringe to determine proper placement. There is often a flash of blood into the hub of the syringe.
    3. Hold the syringe so that the hand is not repositioned on the syringe for injection once the needle has been positioned in the heart.
    4. Inject the article in a slow and steady manner.
    5. Withdraw the needle slowly to minimize bleeding.
    6. Mice that experience respiratory distress, seizure, cyanosis, prolonged ataxia, or other sequelae related to injection trauma must be evaluated by the veterinarian staff and euthanized according to their recommendation.

Figure 1
Figure 1. Intracardiac injection in mice.

2. Intravenous injection utilizing the retro-orbital plexus

  1. Injection landmarks and positioning
    1. Place the animal on a flat surface in lateral recumbency.
    2. Insert the needle into the medial canthus of the eye at a 45° angle to the nose.
    3. The needle depth must be sufficient to penetrate the conjunctival tissues and advance behind the ocular orbit and into the ocular plexus. When placing the needle, it should not encounter the bone at the back of the orbit.
    4. Place the index finger on the top of the head with the thumb on the jaw.
    5. Gently pull back and downward to tighten the skin and protrude the eyeball.
    6. Take care not to apply pressure on the trachea and restrict air flow.
  2. Injection
    1. Place the needle behind the eye and not intra-ocularly.
    2. Ensure that the needle has minimal movement once inserted into the retro-orbital plexus, or the vessels will rupture causing bleeding and loss of the agent into the tissues behind the eye. Hold the syringe so that the hand is not repositioned on the syringe for injection once the needle has been positioned in the retro-orbital plexus.
    3. Do not aspirate with the syringe, as you will collapse the vessels.
    4. Inject the article in a slow and steady manner.
    5. Withdraw the needle and apply light pressure to the eye to control bleeding and to provide hemostasis.
    6. Mice that experience swelling, conjunctivitis, or other sequelae related to injection trauma must be evaluated by the veterinary staff and treated or euthanized according to their recommendation

Figure 2
Figure 2. Retro orbital injection in mice.

3. Footpad Injection

  1. For restraint, place the animal in a restraint tube with one hind foot isolated, and extend by grasping the skin above the stifle.
  2. Wipe the foot with water or alcohol to remove debris prior to injecting.
  3. Inject subcutaneously into the center of the hind foot forming a small bleb at the injection site. To avoid the blood vessel that runs the length of the foot, inject just off the midline parallel to the vessel.

Figure 3
Figure 3. Footpad injection in mice and rats.

Intracardiac, retro orbital and footpad are some of the specialized injection methods that biomedical researchers use for experiments necessitating delivery of compounds via these atypical routes.

An intracardiac injection delivers the compound into the left ventricle allowing the substance to directly enter the arterial circulation. The retro orbital route is an alternative to tail vein injection and is used to deliver the compound into the venous circulation. And a footpad injection involves subcutaneous administration of the article into the animal's hind foot. This video will illustrate the considerations, procedures and applications of these special injection techniques.

Let's begin with some background information and things one should consider before starting these administration procedures.

Intracardiac administration is often done through the use of an ultrasound to guide the needle into the correct location in the heart. However, if performed correctly utilizing the proper landmarks, the administration can be performed without the use of ultrasound visualization. Note that the procedure requires the use of a general anesthetic, and only one injection per animal is permitted for survival procedures. Generally a 27-30 gauge needle is used for this injection and maximum volume of administration is 100 and 300 microliters for mice and rats, respectively.

For intravenous injection via the retro orbital route, one should have a fair understanding of the orbital venous structure. A mouse has a sinus where several veins-namely the supraorbital, dorsal nasal, inferior palpebral, and superficial temporal-converge. Whereas in rats, there is a network or plexus of several veins. The injection is performed into the sinus or the plexus directly. Like intracardiac, this procedure also requires use of general anesthesia, and only one injection per eye per day is recommended with a total of two injections per eye for survival procedures. As with all injections, the smallest size needle should be selected-generally 27-30 gauge-and the recommended maximum volume is 150 μL per eye.

Despite the controversy, the use of footpad injection is still required for some studies, typically related to inflammation and tumor growth. Note that the injections can only be performed on one foot, never bi-laterally. And the footpad measurements should be done daily as soon as obvious swelling has occurred. A 29-30 gauge needle is recommended for the injection and the maximum volume recommended is 50 μL. Following any injection, all animals must be closely monitored for signs of pain, level of food consumption, and for normal ambulation. Generally the animal must be euthanized when the lesion or tumor interferes with the animal's ability to ambulate or reach the food and water.

Now let's learn the injection procedures, starting with the intracardiac injection. We will demonstrate the procedure in a mouse, but the landmarks and the protocol for a rat are similar.

The first step is to prepare the syringe. Recall a 29 gauge needle and 1 cc syringe is appropriate for mice. And the maximum volume for intracardiac injection is 100 microliters. When drawing the solution, leave a small amount of air between the plunger and the injection material. This is to allow for blood to enter the syringe as it is placed into the heart.

To start, anesthetize the animal using inhalant or injectable anesthetics. Review the considerations for maintaining general anesthesia in another video of this collection. Next, position the animal in dorsal recumbency position on an insulated platform. Then, tape the forelimbs to the platform and place a piece of tape horizontally across the abdomen above the hips. This is to further steady the animal and avoid any movement once the needle has been inserted. Next, using a swab, wet the animal's chest with 70% alcohol.

To pinpoint the injection site, first locate the xiphoid and the manubrium sternum. Then, find the midpoint between the two landmarks. 1-2 mm left of this point, is the needle insertion landmark. Using a cotton-tipped applicator, apply povidone iodine to mark the needle insertion site.

To inject, direct the needle perpendicular to the table, and insert it to the depth of about 2 mm. Then, apply a very slight backpressure to the plunger. A bright red oxygenated blood should enter the syringe hub, which confirms proper placement. Hold the syringe in same spot and inject the material slowly and steadily over the course of 30 to 60 seconds. Rapid injection can result in clumping of the cells and clogging of the arteries, a shock to the system due to the temperature of the substance being significantly lower than the body temperature, or an expansion of the ventricle and disruption of the heart rhythm.

Once the material has cleared the syringe, slowly and carefully remove the needle without any lateral movement as that can damage the heart muscles. Then immediately release the tape from the forelegs and abdomen and place the animal in prone position in a clean cage with sufficiently deep bedding to act as an insulating layer. Note that one half of this recovery cage is on a heating source and the anesthetized animal is situated on the heated side of the cage. This prevents hypothermia, and as the animal recovers from the anesthesia it will be able to move off the heated side as desired.

Next, let's learn the method for intravenous injection utilizing the retro-orbital plexus in rats. Again, we will demonstrate the procedure on a mouse, but the landmarks and the protocol for rats are similar.

Attach the appropriate needle to the selected syringe and fill in the injection material. Remember, generally one would use a 27-30 gauge needle with the smallest syringe possible and a maximum volume of 150 microliters.

To start the procedure, first anesthetize the animal. Then, place it on a flat surface in lateral recumbency position. Now place your index finger on the top of the head and the thumb on the jaw, and gently pull back and down. This is to tighten the skin and protrude the eyeball. Take care not to apply pressure on the trachea and restrict airflow. If planning multiple injections, apply topical ophthalmic anesthetic, like tetracaine or proparacaine.

Insert the needle into the medial canthus of the eye at a 45° angle to the nose. The depth must be sufficient to penetrate the conjunctival tissues and advance into the ocular orbit and into the sinus. It should not encounter the bone at the back of the orbit. To avoid rupturing of the blood vessels, ensure that the needle has minimal movement once inserted. Do not aspirate, as that will collapse the vessels. Inject the article in a slow and steady manner. Then, withdraw the needle gently and apply light pressure to the eye to control bleeding and to provide hemostasis.

Lastly, let's review footpad injection method in mice and rats. To start, attach the appropriate needle and fill the syringe with the correct volume. This procedure can be done in conscious animals.

Place the animal in a restraint tube with one hind foot isolated and extended by grasping the skin above the stifle. Wipe the foot with water or alcohol to remove debris prior to injecting. To avoid the blood vessel that runs the length of the foot, the injection landmark is at the center, but just off of the midline, closer to the toes.

Place the needle bevel up at the injection site directing it towards the heel. Inject the article slowly and steadily to avoid rapid distention of the foot tissues. This will cause the footpad to swell as the injection material fills that subcutaneous space. On a small animal's foot the swelling from the injection can extend to the heel, whereas in a larger animal it will be more localized.

After the injection, observe the animals daily and if persistent swelling is present or if there are lesions or tumors as a result of the experimental protocol, then, using a caliper, perform the footpad measurement. This instrument measures the foot thickness in millimeters and helps in quantitation of swelling.

Now let's discuss a few example experiments utilizing intracardiac, retro orbital and footpad injections.

One of the many applications of intracardiac administration is development of an animal model of cancer metastasis. Here, researchers used this route to inject tumor cells that possess propensity for bone colonization. In the following days, they studied tumor growth in bones using X-ray and fluorescence imaging techniques. In another study, the retro orbital route was used to inject specific antibodies that label neutrophils. Then, with help of intravital imaging, the scientists were able to track the migration pattern of the labeled cells.

Lastly, investigators often use footpad injection to analyze inflammatory response. In this experiment, researchers isolated peripheral blood mononuclear cells from human blood samples, mixed them with different test antigens and injected the solutions into the animal's footpad. Finally, they performed foot measurements to quantify the swelling response due to different antigens.

You've just watched JoVE's final installment on the usual and specialized compound administration techniques.

Just to recap, in the first part we reviewed the most common parenteral route. In the second chapter, we discussed the enteral and topical procedures. The third installment dealt with the first set of atypical procedures like intradermal, intranasal, and intracranial in neonates. Lastly, here we discussed three additional routes that biomedical researchers use in labs for specific purposes.

After watching this series you should have a much better understanding of different administration techniques and you should also know the general and specific considerations related to these protocols of compound administration As always, thanks for watching!

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