Retro Orbital Pain

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Retro Orbital Pain
Pain behind the eyes is a common complaint for those patients experiencing tempormandibular joint dysfunction (TMD). The retro-orbital bony complex contains the greater and lesser wings of the sphenoid. Since the lateral and medial pterygoid muscles insert into the medial and pterygoid plates, chronic contracture of these muscles could result in the torquing action of the sphenoid. Pressure behind the eyes, dagger feelings and or ice pick feels along with blurred vision can also be associated with TMD. Patient with limited mouth opening often experience these kind of symptoms. The sphenomandibularis muscle (an internal muscle behind the eyes), runs from the mandible (internal lower jaw) and inserts into the sphenoid bone just behind the eyes.

Educating Yourself About TMJ Anatomy of the Temporomandibular Joint Over Closed Bites – TMD Class II Division 2 Type Problems Degenerative Joint Disease: Clinical Considerations Limited Mouth Opening Problems Ear Congestion Feelings Tinnitus (Ringing in the Ears)

Tongue Posture and Abnormal Swallowing Patterns Contributing to Hyper Muscle Activity and TMD Airway Restrictions

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Where is retro-orbital pain?

Clinical evaluation has associated this muscle with retro-orbital headache. The pain may be localized at the temple or it may also be represented with painful symptoms at the muscle insertion on the internal oblique line of the mandible, intra-orally.

What causes orbital pain?

What Causes Eye Pain? – Causes of eye pain fall into two broad categories: ocular pain and orbital pain.

  • Conjunctivitis is one of the most common eye problems, Conjunctivitis can be an allergic, chemical, bacterial, or viral inflammation of the conjunctiva (the delicate membrane lining the eyelid and covering the eyeball). Pinkeye is a nonmedical term usually referring to conjunctivitis caused by a respiratory virus, because the conjunctiva gets inflamed and turns a pinkish color. Conjunctivitis is usually not associated with the symptom of pain. Itching, redness, and drainage are typical symptoms and signs associated with conjunctivitis.
  • Corneal abrasions and corneal ulcerations are common conditions that cause eye pain. The cornea is the transparent surface of the eye, and it contains many pain-sensitive nerve endings. Abrasions occur from scratches to the surface of the cornea, such as from trauma, a foreign body in the eye, or overuse of contact lenses, Ulcerations occur from primary infections of the cornea or infected abrasions.
  • Keratopathies are a variety of conditions of the cornea and can be a cause of ocular pain.
  • Foreign bodies, usually located on the cornea or in the conjunctiva, are objects or materials that give the sensation that something is in eye. Foreign bodies produce eye pain similar to that of corneal abrasions.
  • Chemical burns and flash burns can be significant causes of eye pain. Chemical burns come from eye exposure to acid or alkaline substances, such as household cleaners or bleach. Flash burns occur from intense light sources when improper or no eye protection is worn while performing arc welding or being exposed to the ultraviolet rays of tanning booths. Even an intense sunny day can cause a corneal flash burn from reflected ultraviolet light.
  • Blepharitis is a condition that causes mild eye discomfort when plugged oil glands at the eyelid edges cause inflammation of the eyelid.
  • A sty or a chalazion causes eye pain because of local irritation. Either of these conditions cause a lump you can see or feel within the eyelid. The lump is a result of a blocked oil gland within the eyelid. This lump causes irritation to the eye, can be very painful to the touch, and may occur in both children and adults.
  • Acute angle closure glaucoma can cause severe ocular or orbital pain. However, most cases of glaucoma are of the open-angle variety and are painless. An increase in intraocular pressure, or internal eye pressure, causes glaucoma. This can ultimately lead to defects in vision and even blindness if left untreated. Intraocular pressure can increase because of a blockage of outflow or increased production of aqueous humor (the fluid that bathes the inner eye). Glaucoma typically occurs in older adults.
  • Iritis is an inflammation of the iris, or colored part of the eye, that causes one to feel deep eye or orbital pain, usually accompanied by blurred vision and light sensitivity.
  • Scleritis is a rare cause of severe eye pain and is often associated with systemic illness.

Orbital pain is described as a deep, dull ache behind or in the eye. This pain is often caused by diseases of the eye.

  • Optic neuritis is an inflammation of the optic nerve, The optic nerve connects to the back of the eye. The cause of this inflammation can be from multiple sclerosis, viral infections, or bacterial infections and can cause symptoms such as pressure behind the eye together with changes in vision and eye pain, especially on movement of the affected eye.
  • Sinusitis, which is a bacterial or viral infection or allergic reaction in the sinuses, can cause a sensation of orbital or eye socket pain. Pain coming from the sinus cavities can be interpreted as eye pain.
  • Migraines and cluster headaches are a very common cause of orbital eye pain.
  • Painful ophthalmoplegia is the combination of orbital pain and eye muscle weakness. In addition to pain, there is double vision when both eyes are open. Causes include various inflammatory conditions of the orbit.
  • Tooth pain resulting from problems with the upper teeth may present as pain in the orbit or below the eye.
  • Traumatic events, such as a penetrating injury to the eye, a blow to the eye with a foreign object, and motor vehicle collisions, are causes of significant eye pain and injury. Scratches to the cornea typically associated with traumatic events are very painful. These are common eye problems that lead people to seek medical attention.

What is a one sided headache and retro-orbital pain?

Cluster Headache – Cluster headache is a common cause of referred ocular and periocular pain. It is presumed that interplay between the sphenopalatine ganglion and the trigeminal ganglion is responsible for the patient’s perception of eye pain when suffering from cluster headache.

  1. Cluster headache derives its name from the pattern of its occurrence: namely, the headaches occur in clusters followed by headache-free remission periods.28 Unlike other common headache disorders that affect primarily females, cluster headache occurs much more often in males by a ratio of 5:1.
  2. Much less common than tension-type headache or migraine headache, cluster headache is thought to affect approximately 0.5% of the male population.

The onset of cluster headache occurs in the late third or early fourth decade, in contradistinction to migraine, which almost always manifests itself by the early second decade. Unlike migraine, cluster headache does not appear to run in families and cluster headache sufferers do not experience aura.

  • Attacks of cluster headache will generally occur approximately 90 minutes after the patient falls asleep.
  • This association with sleep is reportedly maintained when a shift worker changes to and from nighttime to daytime hours of sleep.
  • Cluster headache also appears to follow a distinct chronobiologic pattern that coincides with the seasonal change in the length of daylight.

This results in an increased frequency of cluster headaches in the spring and fall. During a cluster headache period, attacks occur two to three times a day and last for 45 minutes to an hour. Cluster headache periods usually last for 8 to 12 weeks, interrupted by remission periods of less than 2 years.

  • In rare patients, the remission periods become shorter and shorter and the frequency may increase up to 10-fold.
  • This situation is termed chronic cluster headache and differs from the more common episodic cluster headache described previously.
  • Cluster headache is characterized as a unilateral headache that is ocular, retro-orbital, and temporal.

The pain has a deep burning or boring quality. Physical findings during an attack of cluster headache may include Horner syndrome, consisting of ptosis, abnormal pupil constriction, facial flushing, and conjunctival injection ( Fig.49-15 ). Additionally, profuse lacrimation and rhinorrhea is often present.

  1. The ocular changes may become permanent with repeated attacks.
  2. Peau d’orange skin over the malar region, deeply furrowed and glabellar folds, and telangiectasia may be observed.
  3. Attacks of cluster headache may be provoked by small amounts of alcohol, nitrates, histamines, and other vasoactive substances and occasionally by high altitude.

When the attack is in progress, the patient may not be able to lie still and may pace or rock back and forth in a chair. This behavior contrasts to that in other headache syndromes, during which patients seeking relief will lie down in a dark, quiet, room.

  1. The pain of cluster headache is said to be among the worst pain that mankind suffers from.
  2. Because of the severity of pain associated with cluster headaches, the clinician must watch closely for medication overuse or misuse.
  3. Suicides have been associated with prolonged, unrelieved attacks of cluster headaches.

There is no specific test for cluster headache. Testing is aimed primarily at identifying occult pathology or other diseases that may mimic cluster headache (see “Differential Diagnosis”). All patients with a recent onset of headache thought to be cluster headache should undergo MRI testing of the brain.

If neurologic dysfunction accompanies the patient’s headache symptomatology, the MRI should be performed with and without gadolinium contrast medium and MR angiography should also be considered. MRI testing should also be performed in those patients with previously stable cluster headache who are experiencing an inexplicable change in headache symptoms.

Screening laboratory testing including a erythrocyte sedimentation rate, complete blood cell count, and automated blood chemistry should be performed if the diagnosis of cluster headache is in question. Ophthalmologic evaluation including measurement of intraocular pressures is indicated in those patients suffering with headache who experience significant ocular symptom.

In contradistinction to migraine headache, where most patients experience improvement with the implementation of therapy with β-adrenergic blockers, patients suffering from cluster headache will usually require more individualized therapy. A reasonable starting place in the treatment of cluster headache is to begin treatment with prednisone combined with daily sphenopalatine ganglion blocks with local anesthetic.29 A reasonable starting dose of prednisone would be 80 mg given in divided doses tapered by 10 mg per dose per day.

If headaches are not rapidly brought under control, the inhalation of 100% oxygen is added via a close-fitting mask. If headaches persist and the diagnosis of cluster headache is not in question, a trial of lithium carbonate may be considered. It should be noted that the therapeutic window of lithium carbonate is small and thus this drug should be used with caution.

A starting dose of 300 mg at bedtime may be increased after 48 hours to 300 mg twice a day. If no side effects are noted, after 48 hours the dose may again be increased to 300 mg three times a day. The patient should be continued at this dosage level for a total of 10 days, and the drug should then be tapered downward over a 1-week period.

Other medications that can be considered if the just- mentioned treatments are ineffective include methysergide and sumatriptan and sumatriptan-like drugs. Read full chapter URL: https://www.sciencedirect.com/science/article/pii/B9780721603346500534

What is retro-orbital area?

: situated or occurring behind the orbit of the eye.

What does retro orbital pain feel like?

Some headaches may feel like they are behind the eyes and can feel like a deep pressure or even throbbing pain. These headaches are known as Retro-orbital Headaches.

Is orbital eye pain serious?

Orbital eye pain and pain around the eye sockets – The eye is an extremely complex organ, leading to many potential causes of orbital eye pain. The following are the most common conditions and diseases that may cause pain around the eye sockets.

Glaucoma: When people ask “What does pressure behind the eyes mean?” they are typically referencing glaucoma, a disease caused by increased intraocular pressure. While the most common type of glaucoma, open-angle glaucoma, is typically painless, a rarer, fast-acting and dangerous type of glaucoma called angle-closure glaucoma can cause redness, severe pain, and vision loss. (For more information, read our article on the,) Iritis: Iritis is a rare condition in which the iris (the colored part of the eye) becomes inflamed. Side effects include deep orbital pain, reduced vision, redness, and light sensitivity. Migraines, tension headaches, cluster headaches: All three of these types of headaches can create the sensation of pain originating from behind the eye. Note that ocular migraines are different than having eye pain from a migraine; ocular migraines typically last for thirty minutes to an hour and can result in either temporary vision loss or blindness in one eye. Optic neuritis: Optic neuritis is the inflammation and/or infection of the nerve that connects your eye to your brain. Pain caused by optic neuritis often increases with eye movement. Patients may also experience temporary vision loss and headaches. Orbital cellulitis: Orbital cellulitis is an infection of the inside of your eye socket. It can occur after eye trauma, eye surgery, or as the result of infections spreading from other parts of your body (especially the teeth and sinuses). Orbital cellulitis creates redness, pain, and swelling, discharge, and fever, and can lead to permanent vision loss without immediate treatment. Sinusitis/Sinus infection: Yes, your sinuses can also cause pain around your eye sockets—or at least the sensation of eye pain. The congestion and inflammation associated with sinus infections can lead to increased pressure in the sinuses, which can then radiate to your eyes. Toothache: A toothache can cause both headaches and eye pain by pain referred via the nerves that run throughout your facial structure (especially the trigeminal nerve).

Why do I have a pain behind my eye?

Pain behind the eye can result from eyestrain, migraine, dental problems, glaucoma, giant cell arteritis, and other causes. Treatment will depend on the cause, but applying cool or warm compresses may help. Pain behind the eye is a nonspecific symptom as it can be associated with many different health conditions.

Common types of pain behind the eye vary from dull aches to sharp and intense pains. Some people experience a sharp pain behind while others have a deeper pain inside the head, Symptoms can also include tearing, sensitivity to light, redness, vision changes, or pain during eye movement. This article examines the possible causes of pain behind the eye, treatments, alternative therapies, and when to consult a doctor if the pain persists.

Reviewing the possible causes for pain behind the eye may provide people with a better sense of the signs of discomfort and when to seek medical help. There are as many as 300 types of headaches, including those that may cause pain behind the eye. The specific causes are known for only about 10% of headaches.

  • Where a person feels pain does not necessarily correspond to what is causing it.
  • Many different health issues can cause pain behind the eye, including the following: Straining the eyes can leave them feeling dry, tired, and blurry.
  • Research has shown that if a person stares at something for an extended time, they tend to blink less, so eyes become less moist.

People should keep screens at a comfortable distance and take breaks from digital devices to reduce eyestrain. The following may put people at risk of eyestrain:

spending long hours staring at a screen being exposed to glare straining eyes in poor lighting driving long distancesstruggling to get by without glasses or an updated prescription when neededother underlying vision problems

What does an orbital migraine feel like?

Migraine aura affecting your vision – Migraine aura is a wave of activity in the brain traveling through the brain. The location of the wave of activity in the brain determines the type of aura. The most common type of aura is a visual aura. About 90% of people who have migraine with aura have this type.

It’s thought that auras are usually visual because such a large portion of the brain processes visual information. If the wave of activity goes through other areas of the brain such as the sensory or language centers, then the person would have sensory (for example, tingling in the tongue, face or arm) or language auras.

The auras usually last for about five minutes to an hour. Aura can sometimes occur without a headache. A migraine aura that affects your vision is common. Visual symptoms don’t last long. A migraine aura involving your vision will affect both eyes, and you may see:

  • Flashes of light
  • Zigzagging patterns
  • Blind spots
  • Shimmering spots or stars

These symptoms can temporarily get in the way of certain activities, such as reading or driving. But migraine with aura isn’t usually considered serious.

What kind of headache is like pain behind one eye?

Cluster headaches are excruciating attacks of pain in one side of the head, often felt around the eye. Cluster headaches are rare. Anyone can get them, but they’re more common in men and tend to start when a person is in their 30s or 40s.

What is retro-orbital sinus?

Retro Orbital, Tail and Intra-cardiac Blood Collection in Rodents | Lab Animal Research | JoVE Trial ends in Source: Kay Stewart, RVT, RLATG, CMAR; Valerie A. Schroeder, RVT, RLATG. University of Notre Dame, IN Blood collection is a common requirement for research studies that involve mice and rats.

The method of blood withdrawal in mice and rats is dependent upon the volume of blood needed, the frequency of the sampling, the health status of the animal to be bled, and the skill level of the technician.1 All methods discussed-retro-orbital sinus bleeds, initial tail snip bleeds, and intracardiac bleeds-require the use of a general anesthesia.

Prior to the bleeding procedure, the type of sample required must be determined. Experimental procedures could require whole blood, plasma, or serum. For whole blood, an anticoagulant must be added to the sample. Plasma, which contains fibrinogen and other clotting factors when separated from the red blood cells, can be extracted from an anticoagulated sample.

Serum is obtained through blood collection without an anticoagulant. The serum will result from centrifugation of the sample once a clot has formed. As the sample has clotted, the serum will not contain fibrinogen or other clotting factors. Both plasma and serum are obtained through the use of a centrifuge run at 2200-2500 RPM for a minimum of 15 minutes.

For a sample that must yield whole blood or plasma, an appropriate anticoagulant must be used. Commonly used anticoagulants for laboratory animals are heparin, sodium citrate, and ethylenediamine tetraacetic acid (EDTA); selection of which is based on research needs.

Sequester-a liquid form of EDTA, heparin, and sodium citrate-can be loaded directly into the syringe to coat the surfaces. This allows contact of the anticoagulant directly as the blood is drawn, aiding in the prevention of clotting. As rat blood clots faster than most mammalian blood, it is essential that the correct ratio of anticoagulant to blood be used for blood collection.

Needle selection is based on the size of the animal and the site of the venipuncture. In general, the larger the bore of the needle, the more rapidly the sample can be collected. Less damage to the blood cells is another benefit to larger needles. However, the main disadvantage to large-bore needles is the potential damage to the vessel.

  • On mice and rats, the choices of size range from 20-29 gauge needles that are 0.5-1.5 inches in length.
  • If a needle is too long, not only is it awkward to use, but having the extra space in the needle could result in clotting.
  • The appropriate needle size is listed for each method in the procedures section.

The size of the required sample must also be predetermined. Due to the small size of the mouse or rat, the maximum amount of blood collection must be calculated for a survival bleed. An average mouse weighing 25 grams has a total blood volume of 1.8 ml; the average rat weighing 250 grams has a total blood volume of 16 ml.

  1. For a single blood sample on a mouse or rat without fluid replacement, the maximum blood volume that can be safely removed is 10% of the total blood volume, or 7.7-8 µl/g.
  2. Thus for an average mouse, 10% of its blood volume is 193-200 µl.
  3. For an average rat of 250 grams, this is equivalent to 1.9-2.0 ml.

Studies have shown that removing more than 15% of the blood volume can cause hypovolemic shock.1,2 However, with fluid replacement, up to 15% of the total blood volume-or 12 µl/g-can be removed. For a 25 gram mouse, this is equivalent to 300 µl; for a 250 gram rat, it is equivalent to 3 ml.

  • For fluid replacement, the fluids should be warmed and given subcutaneously.
  • If it is necessary to take multiple samples, the blood volume drawn is reduced.
  • The maximum blood volume that may be drawn per week is no more than 7.5% of the total blood volume, or 6 µl/g.
  • For a 25 gram mouse, this is equivalent to 145-150 µl per week.
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For a 250 gram rat, this is equivalent to 1.45-1.50 ml per week. If sampling will occur every 2 weeks, up to 10% of the total blood volume (8 µl/g) may be drawn. This is equivalent to 200 µl every 2 weeks for an average mouse, and up to 2.00 ml every 2 weeks for a 250 gram rat.

One study, performed on rats with the average weight of 250 grams, revealed that when blood volumes of 15-20% were removed, it took more than 29 days for blood levels to normalize.1,2 For repeated blood collection, fluid replacement does not allow for a larger blood volume or more frequent blood collection, as it only replaces volume.

The animal will need time to replenish blood cells. The use of the retro-orbital plexus has been a common practice in the past. However, many concerns about the humaneness of this procedure have arisen. During the procedure, excessive movement of the hematocrit tube once placed in the medial canthus of the eye can cause damage to the surrounding tissues, resulting in swelling of the eyelids and/or conjunctival membranes.

The swollen tissues can cause the eyeball to protrude far enough so that closure of the eyelid is impeded, potentially resulting in corneal drying and damage. Pain from swelling can trigger scratching and self-mutilation that results in enucleation of the eye. Improper placement of the hematocrit tube during a retro-orbital bleed can sever the optic nerve, resulting in blindness.

If the hematocrit tube is advanced at an improper angle, the eye can be forced out of the orbit, allowing the eyelids to fall behind the eyeball. If this occurs, it is very difficult to correctly replace the eye into the socket. Other issues that can arise include fracturing of the fragile orbit bones, penetration of the eye globe that results in the loss of vitreous humour, or the formation of a hematoma behind the eye that can result in extreme pain due to the pressure on the eye and surrounding structures.

  1. Despite all of these concerns, if a skilled technician performs the procedure and the animal is fully anesthetized with a general anesthetic, such as isoflurane inhalant anesthesia, retro-orbital bleeding has been shown to be an effective method of blood collection in rodents.
  2. The anatomical structure of the orbital area is different between the mouse and rat.

The mouse has the retro-orbital sinus-a collection of vessels that create a sinus in the orbital area. In the orbit of the rat eye, there is a plexus of vessels that flow behind that eye; however, they do not form a sinus, as in the mouse. Consequently, it is easier to perform this procedure on mice.

For repeated sampling collection via the retro-orbital plexus, a minimum of 10 days between bleeds is required to allow the tissues in the area to heal. Although general anesthesia is recommended, the procedure can be performed in mice without general anesthesia if a topical ophthalmic anesthetic, such as proparacaine or tetracaine, is applied prior to the procedure.

As rats do not have the retro-orbital sinus, and because their membranes around the orbit are much stronger, it is mandatory to anesthetize them for this procedure. Serial samples of a small volume can be obtained by using a tail clip method. The initial amputation of the tail must be limited to a tail tip, approximately 0.5-1.0 mm in length in mice and 2.0 mm in rats.1 The tail snip procedure for blood collection allows for serial collections by disrupting the scab or clot of the original cut at the end of the tail.

Generally, additional amputation of the tail tip is not necessary. Volumes of blood collected range from 20-100 µL for mice and 75-150 µL for rats. The amount collected is variable between animals and can be influenced by age, health status, and weight. The sample collected from a tail snip can contain both arterial and venous blood, along with tissue product contamination.

The sample quality decreases if the tail is stroked or “milked” to obtain more blood. To increase blood flow, the tail can be heated with warm compresses, a heat lamp, or submersion in warm water. Pressure should be applied to the tail tip for hemostasis, and animals should be checked every 5-10 minutes to ensure hemostasis has been achieved.

  1. Hemostasis is often delayed with repeated sampling.
  2. A styptic powder may be used for hemostasis.
  3. For the initial amputation, anesthesia (general or local) is recommended.
  4. Subsequent bleeding should not require anesthesia, especially as the animals become habituated to the procedure.
  5. Anesthesia will cause a drop in blood pressure, making blood collection with this technique difficult.

An alternative to a tail snip is the tail vessel nick. This procedure is easily performed on both mice and rats. However, as with the tail snip, the samples may be contaminated with tissue products, especially in the mouse. For rats, a hypodermic needle is inserted into the vessel, and the blood is collected from the hub of the needle.

One study demonstrated the use of a tourniquet placed above the needle puncture site to aid in blood collection.3 A syringe is not used to draw the blood out of the vessel, as the pressure created from the syringe will collapse the vessel. This method can also be used for serial sampling, as a clot can be removed to cause the site to bleed again.

As with tail snips, it is imperative to ensure hemostasis by applying pressure to the site and rechecking the animal every 5-10 minutes. Often, studies require a nonsurvival, large blood sample that is collected through exsanguination via an intracardiac bleed or the caudal vena cava.4 Approximately half of the total blood volume can be collected from a mouse or rat by cardiac puncture.

This is equivalent to 40 µl/g or approximately 1 ml for an average 25 gram mouse. A 250 gram rat would yield approximately 10 ml of blood. The animal must be anesthetized for exsanguination. Inhalant anesthesia or CO 2 narcosis can be used by a proficient technician; injectable anesthesia can also be used.

However, there may be a decrease in blood pressure and circulation, which could decrease the amount of blood collected. The caudal vena cava method requires that the animal be deeply anesthetized to surgically expose the vessel. CO 2 narcosis is not sufficient, as the heart must be beating and the animal breathing during blood withdrawal.

  • During the procedure, too rapid of blood withdrawal can cause the vessel to collapse onto the bevel of the syringe, occluding the opening and preventing blood collection.
  • Also, the vessel walls are thin, and thus movement of the hand and needle must be avoided to prevent rupture or leaking of blood from the needle entry site.

As the needle is not passing through the skin, this method results in the collection of a sterile sample. Adjunctive euthanasia methods must be employed to ensure that the animal does not recover from anesthesia. This method is often followed by cardiac or aortic perfusion.

  1. The intracardiac method can be performed either with the animal restrained manually once it is anesthetized (closed method), or the heart can be surgically exposed as per the protocol for caudal vena cava blood collection method (open method).
  2. For the closed method, the landmarks for needle placement are the groove formed by the rib cage at the xiphoid process, on the animal’s left side.1.

Retro-orbital bleed

Equipment

Prepare a bell jar, or anesthetic induction chamber, to administer an anesthetic gas such as isoflurane. When using a bell jar, it is imperative that the liquid anesthetic does not come into contact with the animal, to avoid absorption through the skin. A platform with small holes can be used. Microhematocrit tubes that hold 50-75 microliters are preferred. Mylar wrapped tubes are less likely to break between the fingers of the operator and should be considered as a safety measure. Several paper towel thicknesses, or other insulating materials, are placed on the work surface to maintain the animal’s body heat during the procedure.

Preparation and positioning of the animal

The animal is anesthetized with an inhalation anesthetic, such as isoflurane, in a bell jar or gas anesthesia induction chamber, to effect. Once the animal is fully anesthetized, it is removed and placed in lateral recumbency. The eye is protruded by placing a finger on the top of the head and along the jawline, and pulling the skin back and down. Avoid applying pressure to the trachea, as that may collapse or occlude the airway causing death by asphyxia.

Blood withdrawal

The microhematocrit is placed in the medial canthus of the eye and directed caudally at a 30-45° angle from the plane of the nose. Apply pressure while gently rotating the hematocrit tube. This will cut through the conjunctival membranes and rupture the ocular plexus. The blood will flow into the hematocrit tube by capillary action. Avoid pushing so deep that you hit the bone at the back of the ocular cavity. Once blood begins to flow, maintain pressure to keep the eye protruded. To collect multiple tubes of blood, it is not necessary to place the next tube into the ocular plexus, as the blood will continue to flow and can be collected as it comes from the medial canthus. To stop bleeding, release the skin and allow the eye to return to the normal position. Apply pressure to the orbit to ensure hemostasis.

Figure 1. Retro orbital blood withdrawal in mice.2. Tail bleed procedures: tail snip and tail nick

Equipment

A sterile scalpel blade, preferably a number 11 blade or a single-sided razor blade, is used to make the initial amputation for the tail snip method. Scissors should not be used because the cut made by scissors is crushing, thus promoting clotting and reducing blood flow. For the tail nick procedure, a number 11 or 15 scalpel blade is used to make the cut. A restraint tube that allows access to the tail of the mouse is prepared. Absorbent paper towels or gauze are used as the substrate for performing the tail snip. Collection tubes or hematocrit tubes are also required. Styptic powder should be available to aid in hemostasis.

Restraint

The animal is placed into the tube such that the tail is accessible. For Broome type restrainers, the animal is pulled rump first into the tube. For other tubes, the animal is placed head first. Animals are secured into the tube such that they cannot turn around or withdraw the tail. Some mice will allow the tail snip and blood collection with minimal manual restraint if they are allowed to grab a rough surface. Some rats will require inhalation anesthesia for this method of blood collection.

Blood withdrawal

The tail is wiped with warm water to remove debris and cause slight vasodilation. DO NOT use hot water. For the tail snip, the tail is extended, and the very end of the tail (0.5-1 mm for mice and up to 2 mm for rats) is cut with the scalpel blade. For the tail nick, the tail is extended, and a cut is made with the scalpel blade approximately 2/3 the distance from the rump, directly over the lateral tail vein. The tail can be stroked from rump to tip to encourage blood flow; however, this will decrease the quality of the sample. The blood is collected from the tip or nick using hematocrit tubes or allowed to drip into a collection vial.

3. Cardiac blood collection

Equipment

For a mouse, a 3 cc syringe with a 22-25 gauge x 1″ needle is preferred. Smaller syringes do not have the same back pressure and can make blood withdrawal more difficult. Needles smaller than 25 gauge restrict the flow of blood, leading to increased clotting and damage to the blood cells. Needles shorter than 1″ may not reach the level of the heart when approaching from the diaphragm. For a rat, a 10-12 cc syringe with an 18 gauge x 1.5″ needle is preferred. Depending on the size of the rat, a smaller syringe may not hold the entire blood volume to be collected, and thus the syringe would have to be changed during the procedure. Needles smaller than 20 gauge restrict the flow of the blood, leading to increased clotting. Needles shorter than 1.5″ may not reach the level of the heart when approaching from the diaphragm. A blood collection tube of sufficient size is used to hold the blood collected.

Restraint

Proper restraint is essential to the success of this method. The animal is held by the scruff with the body hanging vertically. It is important that the body be straight to prevent deflection of the heart or a twisting of the chest. An alternative position is dorsal recumbency when placing the needle between the ribs on the animal’s left side. This is especially useful for very large rats or when multiple animals are to be bled.

Figure 2. Cardiac blood withdrawal with mouse held vertically.

Blood withdrawal

The approach from the posterior aspect, puncturing the diaphragm is more easily accomplished when the mouse or rat is held vertically by the scruff.

The needle is advanced in the notch just to the left of the animal’s xiphoid. The needle should be parallel to the spine and placed just under the ribs. The heart is located approximately at the level of the elbow. Place the needle, bevel up, into the chest, and puncture the heart. Apply slight back pressure with the syringe. If the needle is in the heart, blood will flow into the syringe. Wait until blood has filled the syringe before adding additional back pressure on the syringe.

The lateral approach from the animal’s left side requires positioning the animal in dorsal recumbency.

The point of entry is measured against the point of the elbow on the chest wall. The heart is located approximately at the level of the elbow. The needle is inserted perpendicular to the plane of the table at a point midway on the chest wall as measured dorsoventrally. Place the needle, bevel up, into the chest, and puncture the heart. Apply slight back pressure with the syringe. If the needle is in the heart, blood will flow into the syringe. Wait until blood has filled the syringe before adding additional backpressure on the syringe.

Figure 3. Cardiac blood withdrawal with mouse in dorsal recumbency position.

Technical tips

The normal heart is situated with the apex pointing to the left. In rare instances, the heart may be reversed, resulting in difficulty in puncturing the heart. Excessive back pressure on the syringe may collapse the heart, occluding the needle bevel and stopping blood flow into the syringe. Applying back pressure and releasing it repeatedly will initiate clotting in the syringe. Gently applying pressure to the liver can force additional blood volume into the circulatory system, making it available for withdrawal.

4. Posterior vena cava blood withdrawal

Equipment

A TB syringe with a 25-29 gauge needle is used for blood collection in the mouse. For rats, a 10-12 cc syringe with a 22-25 gauge x 1″ needle is required. A surgical platform, dissection tray, or other surface to secure the animal is needed, along with ties, tape, or pins to affix the limbs in position. Injectable anesthesia or inhalation anesthesia is necessary. If using inhalation anesthesia, it is desirable that the anesthetic be delivered via a precision vaporizer with a nose cone. The procedure length is such that using an induction chamber without additional anesthetic gas delivery will not provide sufficient time to complete the blood withdrawal before the animal revives. Iris scissors for the mouse, or operating room sharp-blunt scissors for the rat, are required, along with small atraumatic thumb forceps, and a 2″x 2″ gauze sponge.

Restraint

When the animal is fully anesthetized, as determined by toe pinch or tail pinch, the animal is placed in dorsal recumbency. The limbs are secured to the platform with tape or pins. The limbs should be extended away from the body.

Withdrawal

The skin is lifted and a small transverse cut is made through the skin just above the pelvis in females, or just above the prepuce in males. The point of the scissors is placed into the cut, and a midline incision is made through the skin from the pelvis/prepuce to the xiphoid. The skin is reflected laterally to each side. Blunt dissection may be necessary to loosen it from the underlying muscle. The muscle is lifted, and a small transverse cut is made through the muscle just above the skin cut. The point of the scissors is placed into the abdomen and a midline incision is made through the muscle to the xiphoid. Be sure to angle the scissors’ point upward to avoid cutting any organs. Cut transversely along the curve of the ribs on each side. Exercise extra caution not to puncture the liver. Gently move the intestines to the animal’s left to expose the posterior vena cava. Place a gauze pad on the liver, and rest the index and middle finger on the liver. With the other hand, insert the needle, bevel upward, into the vena cava midway between the junction of the renal vessels and the iliac bifurcation. Slowly withdraw the blood while applying pressure on the liver.

Figure 4. Blood withdrawal from posterior vena cava. Blood collection is a common requirement for several research studies that involve mice and rats. The choice of method for blood withdrawal in these animals is dependent upon many factors like, the volume of blood needed, frequency of the sampling, health status of the animal to be bled, and the skill level of the technician.

  1. Here, we will review these considerations and outline blood collection procedures including the retro-orbital eye bleed, tail snips and nicks, as well as intra-cardiac blood collection.
  2. For other methods, see the second video in this series.
  3. Before delving into the blood withdrawal protocols, let’s first review some general considerations including sample type, needle selection, and the maximum blood volume that can be collected.

Prior to collecting blood from a mouse or a rat, the type of blood sample required must be determined. Experimental procedures could require whole blood, plasma, or serum. If collecting whole blood, an anticoagulant must be added to the sample to prevent clotting.

  • Commonly used anticoagulants include heparin, sodium citrate, and ethylenediamine tetraacetic acid, abbreviated as EDTA.
  • Anticoagulants can be loaded directly into the syringe to coat the surfaces.
  • This allows contact of the anticoagulant directly as the blood is drawn aiding in the prevention of clotting.

Because rodent blood clots rapidly, it is essential that the correct ratio of anticoagulant to blood be used. Plasma collection requires centrifuging the whole blood WITH anticoagulant. Following the spin, the translucent liquid above the WBC and platelet layer is plasma.

It contains fibrinogen and other clotting factors. On the other hand, serum is collected from whole blood sample WITHOUT anticoagulants. And because the sample has clotted, the serum, which is the top player, does not contain fibrinogen or other clotting factors. Needle selection is based on the size of the animal and the site of the venipuncture.

In general, large bore needles cause less damage to blood cells and enable more rapid blood collection; but are more likely to cause vessel damage. Needle length should also be considered. If a needle is too long, it could be awkward to use, or blood could begin to clot while still inside the needle.

The choices of size ranges from 18 to 29 gauge and 0.5 to 1.5 inches in length. The appropriate needle size for each method will be discussed in the procedures section. Lastly, because of the small size of rodents, there is a maximum amount of blood that can be collected from a single blood draw, which will not cause serious harm to the organism.

Blood withdrawal could be without or with fluid replacement – usually done using 0.9% physiological saline. The upper limit in each case is listed in the text protocol below. Furthermore, some experiments require multiple sample collection and in such cases along with fluid replacement animal will need time in between to replenish blood cells as well.

  1. Again, there is a maximum amount that can be collected during serial collection, and the upper limits are listed in the protocol below.
  2. After reviewing some general considerations, let’s jump into the specific blood withdrawal techniques, starting with retro-orbital bleeding – a technique used by scientists to collect small volumes from the vessels near the eye.

Note that the anatomical structure of the orbital area is different between the mouse and rat. The rats have a plexus of vessels that flow behind the eye, whereas the mouse has a collection of vessels that create a retro orbital sinus, which makes it is easier to perform this procedure in mice.

  • Begin by grabbing a tube for blood collection.
  • Micro hematocrit tubes that hold 50-75 microliters are preferred.
  • Lay down several paper towels or other insulating materials on the work surface.
  • This is to maintain the animal’s body heat during the procedure.
  • Now anesthetize the animal using an inhalation anesthetic such as isoflurane.

Once the animal is fully anesthetized, remove it from the chamber and place it down on its side that is in in lateral recumbency position. Next, place a finger on the top of the head and along the jaw line and pull the skin back and down to induce eye protrusion.

  • Avoid applying pressure to the trachea as that may cause death by asphyxia.
  • Subsequently, place the micro-hematocrit tube in the medial canthus of the eye and direct it caudally at a 30 to 45 degree angle from the plane of the nose.
  • Apply pressure while gently rotating the tube.
  • This will cut through the conjunctival membranes and rupture the ocular plexus or sinus.
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The blood will flow into the hematocrit tube by capillary action. Avoid pushing the tube so deep that you hit the bone at the back of the ocular cavity. Once blood begins to flow, maintain pressure to keep the eye protruded. To stop bleeding, release the skin and allow the eye to return to the normal position.

  • Apply pressure to promote hemostasis.
  • For repeated sample collection, allow a minimum of 10 days between the bleeds.
  • This provides tissues some time to heal.
  • Although retro-orbital bleeding is a common procedure, there are many concerns about its humaneness.
  • These include swelling due to excessive movement of the hematocrit tube.

This in turn can cause the eyeball protrusion and impede closure of the eyelid resulting in corneal drying, damage, and pain, which can trigger scratching and self-mutilation. Improper placement of the hematocrit tube can sever the optic nerve resulting in blindness.

  • Another possible complication is that the eye can be forced out of the orbit, allowing the eyelids to fall behind the eyeball.
  • Furthermore, issues can arise from the fracturing of the fragile orbit bones, penetration of the eye globe resulting in the loss of vitreous humor, or the formation of a hematoma behind the eye that can result in extreme pain.

Despite all of these concerns, if a skilled technician performs the procedure and the animal is fully anesthetized, retro-orbital bleeding is an effective method of blood collection in rodents. Now let’s review the considerations and procedures for tail bleeding, which allows collection of a serial samples of small volumes.

The equipment needed for this procedure include a sterile number 11 scalpel. Scissors should not be used because the cut made by scissors is crushing, which can promote clotting and reduce blood flow. Other instruments are a restraint tube that allows access to the animal’s tail; absorbent paper towels; collection or hematocrit tubes and styptic powder – to aid in hemostasis.

Start by securing the animal into the restraint tube. Then, wipe the tail with warm water to remove debris and to cause slight vasodilation. DO NOT use hot water.Extend the tail and with the scalpel blade snip the very end of the tail to collect the blood using hematocrit or collection tubes.

  • The tail can be stroked or “milked” from rump to tip to encourage blood flow.
  • This will, however, decrease the quality of the sample.
  • To stop bleeding, apply pressure to the tail tip with a gauze pad.
  • The styptic powder may be used to achieve hemostasis.
  • Check the animals every 5 to 10 minutes to ensure hemostasis has been achieved, which might be delayed after repeated sampling.

The sample collected from a tail snip can contain both arterial and venous blood, along with tissue product contamination. However, this procedure for blood collection allows for serial collections by disrupting the scab or clot of the original cut at the end of the tail.

An alternative blood collection method to a tail snip is the tail vessel nick, which is relatively less invasive. For this, using the same scalpel blade, make a small cut directly over the lateral tail vein, approximately two-third the distance from the rump. As with tail snips, blood can be collected in collection or hematocrit tubes.

And it is imperative to assure hemostasis by applying pressure to the site and rechecking the animal every 5-10 minutes. However, as with the tail snip, the samples may be contaminated with tissue products. Often studies that require a non-survival large blood sample, which is accomplished through exsanguination via an intra-cardiac bleed or the caudal vena cava.

For intra-cardiac method in mice, you need a 3 cc syringe with a 22 -25 gauge 1 inch needle. And for rats, a 10-12 cc syringe with an 18 gauge 1.5 inches needle is preferred. See the protocol below to understand the why these needs and syringes are ideal. Start by euthanizing the animal using carbon dioxide.

Following euthanasia, hold the rodent by the scruff with the body hanging vertically. This restrain is critical as the body should be straight to prevent deflection of the heart or a twisting of the chest. Note that the heart is located approximately at the level of the elbow.

  • The insertion side is in the notch just to the left of the xiphoid, parallel to the spine and under the ribs.
  • Insert the needle, bevel up, into the chest and puncture the heart.
  • Apply slight backpressure with the syringe.
  • If the needle is in the heart, blood will flow into the syringe.
  • Wait until the blood has filled the barrel before adding additional backpressure.

Approximately half of the total blood volume can be collected from a mouse or rat by cardiac puncture. This is equivalent to approximately 1 mL of blood from an average mouse and approximately 10 mL of blood from an average rat An alternative position is dorsal recumbency when using the lateral approach.

In this case, place the needle between the ribs on the animal’s left side. The point of entry is measured against the point of the elbow on the chest wall. Insert the needle, bevel up, perpendicular to the plane of the table at a point midway on the chest wall. Apply slight back pressure with the syringe.

If the needle is in the heart blood will flow into the syringe. Again, wait until the blood has filled the barrel before adding additional backpressure. Note that in either position, excessive backpressure may collapse the heart occluding the needle bevel and stopping blood flow into the syringe.

  • Another method to collect cardiac blood is through the caudal vena cava.
  • The equipment needed for this procedure are an appropriate syringe with a correct size needle attached; scissors for opening the abdominal cavity, small atraumatic thumb forceps and gauze sponge.
  • This technique requires that the animal be deeply anesthetized and maintained under anesthesia throughout the procedure.

CO2 narcosis is not an option, as the animal heart must be beating for this procedure. Place the animal in dorsal recumbency position, and secure the limbs to the platform. The limbs should be extended away from the body. Now lift the skin with forceps and use scissors to make a small transverse cut through the skin just above the pelvis in females or prepuce in males.

  1. Next, place the point of the scissors into the cut and make a midline incision through the skin from the pelvis or prepuce to the xiphoid.
  2. With the skin laterally reflected, lift the muscle and make a small transverse cut through the muscle, just above the skin cut.
  3. Place the point of the scissor into the abdomen and make a midline incision through the muscle to the xiphoid.

Be sure to angle the scissor point upward to prevent cutting any organs. Cut transversely along the curve of the ribs on each side. Be careful not to puncture the liver. Gently move the intestines to the animal’s left to expose the posterior vena cava. Place a gauze pad on the liver and rest your index and middle fingers on it.

  1. With your other hand, insert the needle, bevel up into the vena cava, midway between the junction of the renal vessels and iliac bifurcation.
  2. Slowly withdraw the blood while applying pressure on the liver.
  3. Avoid hand movement, as that might cause the vessel rupture.
  4. Also, too rapid blood withdrawal can cause the vessel to collapse onto the bevel occluding the opening and preventing blood collection.

The main advantage of this technique is the ability to collect a sterile sample because the needle does not pass through the skin. Lastly, let’s look at some applications of these blood withdrawal techniques. Immuno-oncology is an emerging field, and researchers in this area often perform blood collection to study the immune cells at different stages of cancer development.

For example, here researchers collected cardiac blood from cancer-bearing mice to isolate and quantify neutrophils at ten, twenty and thirty days following tumor engraftment. On the other hand, blood composition is also frequently studied by physiologists. Like in this study, researchers were interested in evaluating kidney function in diabetic animals.

In order to do that, these scientists first injected a dye into a diabetes animal model. Next, they then used tail snip method to collect blood at several time-points to evaluate dye concentration in blood, which was ultimately used to calculate glomerular filtration rate that highlighted the difference in kidney function following diabetes induction.

  • Lastly, stem cells researchers use blood samples to evaluate the success of incorporation of donor cells into the recipient’s system.
  • Here, the investigators first transplanted bone marrow cells from a male mouse into a wild type and genetically modified female animal via the tail vein injection.
  • Next, they collected blood from the retro orbital sinus of the recipient mouse to study the genomic DNA of blood cells using polymerase chain reaction.

This provided the percentage of donor cells engraftment in the two types of animals. You’ve just watched JoVE’s first installment on blood withdrawal techniques. Please see the next video in series to review how to perform other commonly employed techniques of blood collection in lab animals.

As always, thanks for watching! Blood collection for mice and rats can be accomplished with a variety of techniques. Although many factors, such as sample size, frequency of sampling, and the size and age of the animal influence this, the most essential component is the skill level of the technician performing the sample collection.

For the methods described here, the proper use of anesthetics is also crucial for quality samples and the wellbeing of the animals.

Guidelines for the survival bleeding of mice and rats.2010. Diehl, K.H., Hull, R., Morton, D., Pfister, R., Rabemampianina, Y., Smith, D., Vidal, J.M., and van de Vorstenbosch, C.2001. A good practical guide to the administration of substances and removal of blood, including routes and volumes. Journal of Applied Toxicology,21.15-23. Omaye, S.T., Skala, J.H., Gretz, M.D., Schaus, E.E., and Wade, C.E.1987. Simple method for bleeding the unanaesthetized rat by tail venipuncture. Laboratory Animals,21.261-264. Adeghe, A.J-H. and Cohen, J.1986. A better method for terminal bleeding of mice. Laboratory Animals,20.70-72.

Blood collection is a common requirement for several research studies that involve mice and rats. The choice of method for blood withdrawal in these animals is dependent upon many factors like, the volume of blood needed, frequency of the sampling, health status of the animal to be bled, and the skill level of the technician.

Here, we will review these considerations and outline blood collection procedures including the retro-orbital eye bleed, tail snips and nicks, as well as intra-cardiac blood collection. For other methods, see the second video in this series. Before delving into the blood withdrawal protocols, let’s first review some general considerations including sample type, needle selection, and the maximum blood volume that can be collected.

Prior to collecting blood from a mouse or a rat, the type of blood sample required must be determined. Experimental procedures could require whole blood, plasma, or serum. If collecting whole blood, an anticoagulant must be added to the sample to prevent clotting.

  1. Commonly used anticoagulants include heparin, sodium citrate, and ethylenediamine tetraacetic acid, abbreviated as EDTA.
  2. Anticoagulants can be loaded directly into the syringe to coat the surfaces.
  3. This allows contact of the anticoagulant directly as the blood is drawn aiding in the prevention of clotting.

Because rodent blood clots rapidly, it is essential that the correct ratio of anticoagulant to blood be used. Plasma collection requires centrifuging the whole blood WITH anticoagulant. Following the spin, the translucent liquid above the WBC and platelet layer is plasma.

It contains fibrinogen and other clotting factors. On the other hand, serum is collected from whole blood sample WITHOUT anticoagulants. And because the sample has clotted, the serum, which is the top player, does not contain fibrinogen or other clotting factors. Needle selection is based on the size of the animal and the site of the venipuncture.

In general, large bore needles cause less damage to blood cells and enable more rapid blood collection; but are more likely to cause vessel damage. Needle length should also be considered. If a needle is too long, it could be awkward to use, or blood could begin to clot while still inside the needle.

  • The choices of size ranges from 18 to 29 gauge and 0.5 to 1.5 inches in length.
  • The appropriate needle size for each method will be discussed in the procedures section.
  • Lastly, because of the small size of rodents, there is a maximum amount of blood that can be collected from a single blood draw, which will not cause serious harm to the organism.

Blood withdrawal could be without or with fluid replacement – usually done using 0.9% physiological saline. The upper limit in each case is listed in the text protocol below. Furthermore, some experiments require multiple sample collection and in such cases along with fluid replacement animal will need time in between to replenish blood cells as well.

Again, there is a maximum amount that can be collected during serial collection, and the upper limits are listed in the protocol below. After reviewing some general considerations, let’s jump into the specific blood withdrawal techniques, starting with retro-orbital bleeding – a technique used by scientists to collect small volumes from the vessels near the eye.

Note that the anatomical structure of the orbital area is different between the mouse and rat. The rats have a plexus of vessels that flow behind the eye, whereas the mouse has a collection of vessels that create a retro orbital sinus, which makes it is easier to perform this procedure in mice.

  1. Begin by grabbing a tube for blood collection.
  2. Micro hematocrit tubes that hold 50-75 microliters are preferred.
  3. Lay down several paper towels or other insulating materials on the work surface.
  4. This is to maintain the animal’s body heat during the procedure.
  5. Now anesthetize the animal using an inhalation anesthetic such as isoflurane.

Once the animal is fully anesthetized, remove it from the chamber and place it down on its side that is in in lateral recumbency position. Next, place a finger on the top of the head and along the jaw line and pull the skin back and down to induce eye protrusion.

  1. Avoid applying pressure to the trachea as that may cause death by asphyxia.
  2. Subsequently, place the micro-hematocrit tube in the medial canthus of the eye and direct it caudally at a 30 to 45 degree angle from the plane of the nose.
  3. Apply pressure while gently rotating the tube.
  4. This will cut through the conjunctival membranes and rupture the ocular plexus or sinus.

The blood will flow into the hematocrit tube by capillary action. Avoid pushing the tube so deep that you hit the bone at the back of the ocular cavity. Once blood begins to flow, maintain pressure to keep the eye protruded. To stop bleeding, release the skin and allow the eye to return to the normal position.

Apply pressure to promote hemostasis. For repeated sample collection, allow a minimum of 10 days between the bleeds. This provides tissues some time to heal. Although retro-orbital bleeding is a common procedure, there are many concerns about its humaneness. These include swelling due to excessive movement of the hematocrit tube.

This in turn can cause the eyeball protrusion and impede closure of the eyelid resulting in corneal drying, damage, and pain, which can trigger scratching and self-mutilation. Improper placement of the hematocrit tube can sever the optic nerve resulting in blindness.

Another possible complication is that the eye can be forced out of the orbit, allowing the eyelids to fall behind the eyeball. Furthermore, issues can arise from the fracturing of the fragile orbit bones, penetration of the eye globe resulting in the loss of vitreous humor, or the formation of a hematoma behind the eye that can result in extreme pain.

Despite all of these concerns, if a skilled technician performs the procedure and the animal is fully anesthetized, retro-orbital bleeding is an effective method of blood collection in rodents. Now let’s review the considerations and procedures for tail bleeding, which allows collection of a serial samples of small volumes.

The equipment needed for this procedure include a sterile number 11 scalpel. Scissors should not be used because the cut made by scissors is crushing, which can promote clotting and reduce blood flow. Other instruments are a restraint tube that allows access to the animal’s tail; absorbent paper towels; collection or hematocrit tubes and styptic powder – to aid in hemostasis.

Start by securing the animal into the restraint tube. Then, wipe the tail with warm water to remove debris and to cause slight vasodilation. DO NOT use hot water.Extend the tail and with the scalpel blade snip the very end of the tail to collect the blood using hematocrit or collection tubes.

  1. The tail can be stroked or “milked” from rump to tip to encourage blood flow.
  2. This will, however, decrease the quality of the sample.
  3. To stop bleeding, apply pressure to the tail tip with a gauze pad.
  4. The styptic powder may be used to achieve hemostasis.
  5. Check the animals every 5 to 10 minutes to ensure hemostasis has been achieved, which might be delayed after repeated sampling.

The sample collected from a tail snip can contain both arterial and venous blood, along with tissue product contamination. However, this procedure for blood collection allows for serial collections by disrupting the scab or clot of the original cut at the end of the tail.

  1. An alternative blood collection method to a tail snip is the tail vessel nick, which is relatively less invasive.
  2. For this, using the same scalpel blade, make a small cut directly over the lateral tail vein, approximately two-third the distance from the rump.
  3. As with tail snips, blood can be collected in collection or hematocrit tubes.

And it is imperative to assure hemostasis by applying pressure to the site and rechecking the animal every 5-10 minutes. However, as with the tail snip, the samples may be contaminated with tissue products. Often studies that require a non-survival large blood sample, which is accomplished through exsanguination via an intra-cardiac bleed or the caudal vena cava.

For intra-cardiac method in mice, you need a 3 cc syringe with a 22 -25 gauge 1 inch needle. And for rats, a 10-12 cc syringe with an 18 gauge 1.5 inches needle is preferred. See the protocol below to understand the why these needs and syringes are ideal. Start by euthanizing the animal using carbon dioxide.

Following euthanasia, hold the rodent by the scruff with the body hanging vertically. This restrain is critical as the body should be straight to prevent deflection of the heart or a twisting of the chest. Note that the heart is located approximately at the level of the elbow.

The insertion side is in the notch just to the left of the xiphoid, parallel to the spine and under the ribs. Insert the needle, bevel up, into the chest and puncture the heart. Apply slight backpressure with the syringe. If the needle is in the heart, blood will flow into the syringe. Wait until the blood has filled the barrel before adding additional backpressure.

Approximately half of the total blood volume can be collected from a mouse or rat by cardiac puncture. This is equivalent to approximately 1 mL of blood from an average mouse and approximately 10 mL of blood from an average rat An alternative position is dorsal recumbency when using the lateral approach.

  • In this case, place the needle between the ribs on the animal’s left side.
  • The point of entry is measured against the point of the elbow on the chest wall.
  • Insert the needle, bevel up, perpendicular to the plane of the table at a point midway on the chest wall.
  • Apply slight back pressure with the syringe.

If the needle is in the heart blood will flow into the syringe. Again, wait until the blood has filled the barrel before adding additional backpressure. Note that in either position, excessive backpressure may collapse the heart occluding the needle bevel and stopping blood flow into the syringe.

Another method to collect cardiac blood is through the caudal vena cava. The equipment needed for this procedure are an appropriate syringe with a correct size needle attached; scissors for opening the abdominal cavity, small atraumatic thumb forceps and gauze sponge. This technique requires that the animal be deeply anesthetized and maintained under anesthesia throughout the procedure.

CO2 narcosis is not an option, as the animal heart must be beating for this procedure. Place the animal in dorsal recumbency position, and secure the limbs to the platform. The limbs should be extended away from the body. Now lift the skin with forceps and use scissors to make a small transverse cut through the skin just above the pelvis in females or prepuce in males.

Next, place the point of the scissors into the cut and make a midline incision through the skin from the pelvis or prepuce to the xiphoid. With the skin laterally reflected, lift the muscle and make a small transverse cut through the muscle, just above the skin cut. Place the point of the scissor into the abdomen and make a midline incision through the muscle to the xiphoid.

Be sure to angle the scissor point upward to prevent cutting any organs. Cut transversely along the curve of the ribs on each side. Be careful not to puncture the liver. Gently move the intestines to the animal’s left to expose the posterior vena cava. Place a gauze pad on the liver and rest your index and middle fingers on it.

With your other hand, insert the needle, bevel up into the vena cava, midway between the junction of the renal vessels and iliac bifurcation. Slowly withdraw the blood while applying pressure on the liver. Avoid hand movement, as that might cause the vessel rupture. Also, too rapid blood withdrawal can cause the vessel to collapse onto the bevel occluding the opening and preventing blood collection.

The main advantage of this technique is the ability to collect a sterile sample because the needle does not pass through the skin. Lastly, let’s look at some applications of these blood withdrawal techniques. Immuno-oncology is an emerging field, and researchers in this area often perform blood collection to study the immune cells at different stages of cancer development.

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For example, here researchers collected cardiac blood from cancer-bearing mice to isolate and quantify neutrophils at ten, twenty and thirty days following tumor engraftment. On the other hand, blood composition is also frequently studied by physiologists. Like in this study, researchers were interested in evaluating kidney function in diabetic animals.

In order to do that, these scientists first injected a dye into a diabetes animal model. Next, they then used tail snip method to collect blood at several time-points to evaluate dye concentration in blood, which was ultimately used to calculate glomerular filtration rate that highlighted the difference in kidney function following diabetes induction.

Lastly, stem cells researchers use blood samples to evaluate the success of incorporation of donor cells into the recipient’s system. Here, the investigators first transplanted bone marrow cells from a male mouse into a wild type and genetically modified female animal via the tail vein injection. Next, they collected blood from the retro orbital sinus of the recipient mouse to study the genomic DNA of blood cells using polymerase chain reaction.

This provided the percentage of donor cells engraftment in the two types of animals. You’ve just watched JoVE’s first installment on blood withdrawal techniques. Please see the next video in series to review how to perform other commonly employed techniques of blood collection in lab animals.

How often does retro-orbital bleeding occur?

  1. Anesthesia is required for retro-orbital bleeding.
  2. The following anesthetic agents are recommended for this procedure: Mice – Ketamine (90 mg/kg) and Xylazine (10 mg/kg), IP Pentobarbital (50mg/kg), IP Proparacaine (Ophthetic®) – 1 drop per eye Isoflurane (Drop Method) – contact vet staff Rats – Ketamine (75 mg/kg) and Xylazine 10 mg/kg), IP Pentobarbital (50 mg/kg), IP
  3. Topical anesthetic, alone, may only be used in mice. Rats, who require more restraint due to their size, must be bled under general anesthesia. Note that there is the potential for contamination of the blood sample if topical anesthetic is used.
  4. Use of any other anesthetic agents must be identified in the IACUC application.

Prior to and during the procedure the following parameters should be monitored at a minimum of 5 minute intervals:

  • Respiratory rate
  • Response to noxious stimulus
  • Spontaneous movement
  1. During recovery from anesthesia, the following clinical parameters must be monitored at a minimum of 5 minute intervals until the animal is ambulatory. · Respiratory rate · Movement · Ability to maintain sternal recumbancy
  2. To protect the animal from hypothermia they should be placed on a water recirculating heating blanket, or covered well, to conserve body temperature. Animal should never be placed on metal surfaces.
  3. It is estimated that animals will recover within 30-60 minutes postoperatively.
  1. Standard heparinized or non-heparinized micro-hematocrit capillary tubes can be used for blood collection.
  2. The animal is held by the back of the neck and the loose skin of the head is tightened with the thumb and middle finger.
  3. The tip of the capillary tube is placed at the medial canthus of the eye under the nictitating membrane.
  4. A short thrust past the eyeball will enter the slightly resistant membrane of the sinus. The eyeball itself remains uninjured.
  5. As soon as the sinus is punctured, blood enters the tubing by capillary action. It may be helpful to retract the tube to facilitate blood flow.
  6. When the allowable amount of blood is collected, the tube is withdrawn and slight pressure with a piece of gauze on the eyeball is used to prevent further bleeding.

Analgesia is not required for this procedure but the use of topical anesthetic (e.g., Proparacaine – 1 drop per eye) decreases pain post-procedurally.

  1. The maximum amount of blood that may be withdrawn at one time from this location is 1% of the animal’s body weight (e.g., 0.2 ml from a 20 gm adult mouse and 2.0 ml’s from a 200 gm adult rat).
  2. Blood can only be collected once per week from one eye. Subsequent bleeds should use alternate eyes.
  3. The maximum number of bleeds for each animal is two bleeds per eye.
  4. If a project requires a greater volume of blood withdrawal, more frequent bleeds or an increase in the total number of bleeds, any change in the parameters listed must be scientifically justified in the IACUC application.
  5. Alternate bleeding sites, such as the saphenous or tail veins, should be considered.

Potential adverse effects from this procedure include:

  • anesthetic related respiratory distress
  • eye infection
  • peri-orbital swelling, redness and/or hematoma formation
  • blindness
  1. Animals should be monitored at least twice weekly after each retro-orbital bleed.
  2. If adverse effects are seen, the investigator should consult immediately with the veterinary staff regarding treatment options (410-955-3713). If animals have acute adverse reactions to the anesthetic agents (respiratory distress and/or lack of recovery), they must be euthanized immediately.

Animals should be euthanized if the eyeball is acutely damaged, if treatment of an injured/infected eye is unsuccessful and/or if bilateral blindness occurs.

What is the retro-orbital bleeding method?

Retro‐orbital bleeding is a method of blood collection whereby the retro‐orbital sinus in mice or the retro‐orbital plexus in rats is penetrated with a capillary tube. Retro‐orbital sampling has a greater potential than other blood collection routes to result in complications.

What does MS eye pain feel like?

A common visual symptom of MS is optic neuritis — inflammation of the optic (vision) nerve. Optic neuritis usually occurs in one eye and may cause aching pain with eye movement, blurred vision, dim vision or loss of color vision. For example, the color red may appear washed out or gray.

How do you know if your retina is inflamed?

Retinal inflammatory disease (or uveitis ) is an eye condition that causes dysfunction of the retina and, in the most severe cases, substantial vision loss. It is most common in people between the ages of 20 and 60. Retinal inflammatory disease may be caused by an autoimmune disorder that affects multiple systems within the body, or by an infection or trauma to the eye.

Does TMJ cause pain behind eye?

Guiding you in the fight against facial and neck pain. – Oftentimes, a misaligned jaw causes these symptoms. Dr. Stafford is our designated TMJ dentist who has dedicated her continuing education to obtaining specialized training for treating jaw joint, facial pain, and sleep disorders.

Headaches or Migraines One of the top symptoms patients will complain about experiencing are frequent headaches and migraines. Oftentimes, they don’t even know that TMJ is the cause and instead spend their time trying endless medications or treatments that don’t work. TMJ causes headaches because the position of the lower jaw is off. By correcting this misalignment, patients can say goodbye to headaches for good. Pain Behind the Eyes When someone has TMJ dysfunction, it can cause them to overuse their chewing muscles. These muscles are attached to the jaw from behind the eye socket. As a result of hyperactivity of this muscle, patients can experience pain behind their eyes or a headache. Vertigo & Ear Pain Due to the temporomandibular joint’s close proximity to the ears, it’s common for ear symptoms to arise with TMJ, such as ear pain, congestion, and ringing. When there are issues in the inner ear, it can cause a postural imbalance which can lead to feeling dizzy or experiencing vertigo. Neckaches & Tingling in the Fingers When the chewing muscles experience hyperactivity, this also usually means there’s hyperactivity in the head and neck muscles. When head and neck muscles experience hyperactivity, they might constrict or put pressure on radial nerves, which leave the nervous system at the neck. This may cause patients to experience tingling sensations in their fingers.

Clicking/Popping in the Jaw Joint When you open and close your mouth, if you hear any clicking or popping, this is the sound of the disc that rests between the two bones in your jaw slipping out. When it slips out, the jaw bone jumps to put the disc back in place, which can lead to popping or clicking noises. Clenching/Grinding the Teeth When the jaw is at a resting state, the teeth should rest slightly apart, only to touch when swallowing. If the upper and lower jaw experience disharmony in their position, it can lead to clenching and grinding, which can wear down teeth and lead to receding gums. Insomnia The area of the brain that controls TMJ and chewing muscles is also responsible for alertness and wakefulness. If there is hyperactivity in the chewing muscles and TMJ, it can cause patients to suffer insomnia. Sore Jaw Muscles & Facial Pain A misalignment of the TMJ will often force the body to compensate by holding chewing muscles in a position that is non-resting. This will then cause hyperactivity of chewing muscles which can cause facial pain, fatigue from chewing, facial swelling, inability to hold the mouth open for long periods or open all the way.

What are the signs of orbital inflammation?

Symptoms and Signs of Inflammatory Orbital Disease – Symptoms and signs of inflammatory orbital pseudotumor typically include a sudden onset of pain along with swelling and erythema of the eyelids. Proptosis, diplopia, and vision loss are also possible.

In cases of reactive lymphoid hyperplasia or IgG4-related orbital disease, there are typically few symptoms other than proptosis or swelling. Ophthalmopathy in TED may occur before the onset of hyperthyroidism or as late as 20 years afterward, and frequently worsens or abates independently of the clinical course of hyperthyroidism.

Of patients with TED, 5% may have hypothyroidism, and some patients show typical ophthalmopathy in the presence of normal thyroid function (“euthyroid Graves disease”). Symptoms and signs of TED include those that are specific to the condition (ie, eyelid retraction) as well as the nonspecific symptoms seen in almost all orbital inflammation (ie, proptosis, diplopia, periorbital edema, retrobulbar pain).

Corticosteroids, radiation therapy, and/or immunomodulating drugs Some surgery

Treatment for inflammatory orbital pseudotumor depends on the type of inflammatory response and may include oral corticosteroids, radiation therapy, and one of several immunomodulating drugs. In difficult cases of inflammatory orbital pseudotumor, particularly those with granulomatous inflammation, some initial success has occurred with monoclonal antibodies against tumor necrosis factor (TNF)-alpha or with lymphocyte depletion using rituximab, if the inflammation is primarily vasculitis.

  1. Treatment of ophthalmopathy in patients with Graves disease may require selenium, corticosteroids, orbital radiation, and sometimes surgery.
  2. Teprotumumab, an insulin -like growth factor 1 (IGF-1) receptor inhibitor, is effective therapy for moderate-to-severe ophthalmopathy ( 1 Treatment reference Inflammatory orbital disease is a benign space-occupying inflammation involving orbital tissues.

Inflammatory orbital disease, also called orbital pseudotumor, is inflammation that can affect. read more ). Treatment of concomitant hyperthyroidism includes thionamides, radioiodine, or surgery, However, radioiodine therapy may accelerate progression of ophthalmopathy and is therefore contraindicated in the active phase, which is typically determined by clinical signs and symptoms as indicated by the clinical activity score.

Ronquillo Y, Patel BC : Nonspecific orbital inflammation (NSOI). StatPearls Publishing, Treasure Island, FL, 2020.

NOTE: This is the Professional Version. CONSUMERS: View Consumer Version Copyright © 2023 Merck & Co., Inc., Rahway, NJ, USA and its affiliates. All rights reserved.

Is orbital inflammation painful?

Any or all of the structures within the eye socket (orbit) may become inflamed because of a bodywide inflammatory disorder or an inflammatory disorder that affects only the orbit. People of all ages can be affected. Inflammation can be brief or long lasting, may or may not be caused by an infection, and can recur.

Symptoms vary depending on which structures are actually inflamed. In general, symptoms start rather suddenly, typically over a few days. Pain and redness of the eyeball or eyelid usually occur. Pain can be severe and incapacitating at times. Abnormal bulging of the eyes ( proptosis Eyes, Bulging Bulging or protruding of one or both eyes is called proptosis or exophthalmos.

Exophthalmos is usually used when describing bulging eyes caused by Graves disease, a disorder causing overactivity. read more ), double vision, and vision loss are also possible. Symptoms related to IgG4-related orbital inflammation, on the other hand, are usually minimal. Rarely is there any discomfort, but rather proptosis and eyelid swelling Eyelid Swelling A person may experience swelling in one or both eyelids.

Computed tomography or magnetic resonance imaging Biopsy Other tests to determine the cause

Drugs to treat inflammation (corticosteroids) Radiation therapy or drugs to change immune response and treat the underlying cause

Many disorders causing inflammation of the orbit are treated with a corticosteroid drug, which can be given by mouth. Corticosteroids can be given by vein (intravenously) if the inflammation is severe. Radiation therapy or drugs and treatments that change the body’s immune responses may sometimes be used.

Generic Name Select Brand Names
rituximab RIABNI, Rituxan, RUXIENCE, truxima

NOTE: This is the Consumer Version. DOCTORS: VIEW PROFESSIONAL VERSION VIEW PROFESSIONAL VERSION Copyright © 2023 Merck & Co., Inc., Rahway, NJ, USA and its affiliates. All rights reserved.

How long should I be worried if my eye hurts?

Summary – Eye pain can range from mild to severe and can be caused by various conditions. If the pain persists or is accompanied by other symptoms, seek medical attention immediately. Treatment options depend on the cause of the pain, so getting a proper diagnosis is important.

Where is eye strain pain located?

Unlike other types of headaches, eye strain headaches are rarely associated with vomiting or nausea. Pain behind your eyes. The pain is usually located behind or around your eyes. The area might feel sore or tired.

Where is optic nerve pain located?

Optic nerve – The optic nerve is a bundle of nerve fibers that serves as the communication cable between your eyes and your brain. The nerve fibers have a special coating called myelin. Signs and symptoms of optic neuritis can be the first indication of multiple sclerosis (MS), or they can occur later in the course of MS,

  • MS is a disease that causes inflammation and damage to nerves in your brain as well as the optic nerve.
  • Besides MS, optic nerve inflammation can occur with other conditions, including infections or immune diseases, such as lupus.
  • Rarely, another disease called neuromyelitis optica causes inflammation of the optic nerve and spinal cord.

Most people who have a single episode of optic neuritis eventually recover their vision without treatment. Sometimes steroid medications may speed the recovery of vision after optic neuritis. Optic neuritis usually affects one eye. Symptoms might include:

Pain. Most people who develop optic neuritis have eye pain that’s worsened by eye movement. Sometimes the pain feels like a dull ache behind the eye. Vision loss in one eye. Most people have at least some temporary reduction in vision, but the extent of loss varies. Noticeable vision loss usually develops over hours or days and improves over several weeks to months. Vision loss is permanent in some people. Visual field loss. Side vision loss can occur in any pattern, such as central vision loss or peripheral vision loss. Loss of color vision. Optic neuritis often affects color perception. You might notice that colors appear less vivid than normal. Flashing lights. Some people with optic neuritis report seeing flashing or flickering lights with eye movements.

Eye conditions can be serious. Some can lead to permanent vision loss, and some are associated with other serious medical problems. Contact your doctor if:

You develop new symptoms, such as eye pain or a change in your vision. Your symptoms worsen or don’t improve with treatment. You have unusual symptoms, including vision loss in both eyes, double vision, and numbness or weakness in one or more limbs, which can indicate a neurological disorder.

What is orbital pain behind eye?

WHAT IS ORBITAL EYE PAIN? – Eye pain can be experienced as discomfort in the in the area behind and around it (called orbital eye pain) or in the eye itself. Most people with orbital eye pain feel it the behind the eyes. While pain in the eyeball is mainly caused by medical conditions, orbital eye pain can be related to dental and bite problems, including tooth infections in the upper jaw and temporomandibular (TMJ) disorder, characterized by dysfunction in the jaw joint.

Migraines and sinus problems can also cause orbital eye pain; these conditions can be aggravated by bite and jaw joint problems. JAWBONES ARE CONNECTED TO THE EYES. The connection between the jaws and the eye area comes from the nerves. The three nerve branches that run through the upper jaw, lower jaw and eye area originate from the same nerve in the brain, called the trigeminal nerve.

These nerves can transmit pain from one area of the other.

What is the difference between ocular and orbital pain?

Eye Pain: Ocular and Orbital You may have experienced eye pain at some point in your life. Some of them are caused due to minor problems, and they go away without needing any medication, while some problems need the attention of an eye specialist. Eye pain can be a symptom of a wide range of problems; a sign of something as plain as refractive error or a sight-threatening condition like glaucoma.

Therefore, you should not ignore eye pain even if it is mild. The pain could be categorized into two types: ocular, and orbital pain. Ocular pain is one that you experience on the eye’s surface whereas orbital pain occurs within the eye. Depending upon the accompanying symptoms, your doctor will determine the cause of your pain and discomfort, and recommend a suitable treatment plan.

Ocular pain Ocular pain refers to the pain that you experience on the surface of the eye. The pain is often accompanied by sensations of itching and burning. You may experience ocular pain because of irritation caused due to a foreign particle, infection, or trauma. Causes of ocular pain

Foreign object: One of the most common causes of ocular pain is when a foreign object like a piece of dirt, a tiny twig or use of eye cosmetics causes infection around the eyes. Conjunctivitis: It is one of the most common eye infections that affects the front of the eye and the underside of the eyelid. It causes mild pain but it is quite uncomfortable. The eye pain due to conjunctivitis is often accompanied by a burning sensation. Sty : It is an eye infection that causes a bump on the eyelid. It is quite painful and often the area around the bump becomes swollen. Eye irritation : There are various factors that can cause eye irritation. And one of them is wearing dirty contact lenses. Also,exposure to irritants such as bleach and pollen can cause eyes to pain.

Orbital pain Eye pain that is experienced within the eye is called orbital pain. If you are suffering from orbital pain, you will feel excruciating pain. Orbital pain should not be ignored as it can be a sign of a serious eye disease and it requires proper eye treatment.

Glaucoma : The condition can cause eye pain, along with headache and nausea. In the case of an acute-angle-closure glaucoma eye pressure inside the eye increases leading to pain. Glaucoma, when left untreated for long, can lead to loss of vision. Optic neuritis: When the nerve connecting the back of the eyeball to the brain, known as the optic nerve, becomes inflamed, you may experience eye pain along with changes in vision.

There are many other eye conditions that could cause you an eye pain, in which case you must consult an eye specialist. Seeing an eye doctor will help you with the diagnosis and they may recommend a treatment accordingly. In the case of eye pain along with loss of vision or blurry vision or sudden change in vision, treat it as is a sign of emergency and seek immediate medical attention.

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: Eye Pain: Ocular and Orbital