LAT Chapter 11 Procedures PDF

LAT Chapter 11 Common Technical Procedures Technical procedures are a significant part of the role of a LAT. Proficiency at these techniques is therefore essential for a technician and acquiring the skills and practice to perform them should be a training priority. Conducting technical procedures sk...

LAT Chapter 11 Common Technical Procedures Technical procedures are a significant part of the role of a LAT. Proficiency at these techniques is therefore essential for a technician and acquiring the skills and practice to perform them should be a training priority. Conducting technical procedures skillfully is essential for animal welfare, as animals that are handled and treated gently by a proficient technician tend to exhibit less stress and better outcomes. Technical Procedures in Terrestrial Species Substance Administration • Injection routes: (in order of fastest absorption to slowest) • • • • • intravenous (IV) intraperitoneal (IP) intramuscular (IM) subcutaneous (SC or SQ) intradermal (ID) • The rate of absorption of the injected substance into the body depends upon the route of injection • Injections can also be administered into: • Cerebrum of the brain (intracerebral or IC), the • Retroorbital sinus, and the • Hollow cavity of a bone (intraosseus) [pic]. • Substances may also be administered by gastric gavage, which is an injection into the stomach lumen. The techniques for each route vary by species and ALL require a general understanding of local anatomy at the injection site. Intravenous Injections For an intravenous (IV) injection, the needle is inserted into a blood vessel. This injection route is commonly used for fast absorption and distribution of a compound in the body. • Some medicines or test substances may only be administered IV because they may irritate tissues when given outside the vessels. • Various blood vessels can be used for IV injections, depending on the species. • Prep: Prior to an injection, the injection site may be shaved or plucked and prepped with an antiseptic to disinfect the skin and make the vessel easier to see. • Occlude: Light pressure from a tourniquet or an assistant’s hand is applied proximal to the injection site, causing the vein to distend with blood and become more visible. Be careful not to apply excessive pressure in squeezing a limb or tail; too much pressure would hurt the animal unnecessarily. Veins are easily collapsible, so only light pressure is needed to occlude the vein. • Needle Insertion: The vein should be punctured at an angle of about 30 degrees to the skin surface, with the bevel of the needle facing away from the skin. Verify that the needle is placed correctly by aspirating gently, that is, by pulling slightly on the syringe's plunger. Blood aspirated into the syringe or needle hub (known as a flash) indicates proper placement of the needle inside the vein. • Release the pressure: On the vein prior to starting the injection so that the blood circulation will carry off the solution as it is injected. Failure to release the pressure holds the solution in the vessel at the injection site, causing it to leak out of the vein at the puncture site into the surrounding tissues. • Hemostasis: When the injection is completed, pressure should be applied over the injection site simultaneously with the withdrawal of the needle. This pressure should be maintained for a short time to allow the puncture site to form a clot (hemostasis), which prevents bleeding into the surrounding tissues (a hematoma) from forming and keeps the injected material from leaking out of the vein. IV – not from a US-based facility • https://www.youtube.com/watch?v=hanSVyDFwCo Intravenous Injections - Mice & Rats • • • • Site: The lateral tail vein is commonly used for injections of small volumes in mice and rats. Restraint: Secure unanesthetized animals in a restrainer; anesthetized animals do not require restraint. Locate: Locate the lateral tail vein (swabbing the area with alcohol helps make the vein visible); Occlude: Pressure can be applied proximally if it helps visualize and access the vein. A tourniquet may be devised by threading a loop of suture through a 3 mL syringe; moving the plunger controls the size of the suture loop to either constrict or release the tourniquet. • Needle Insertion: The vein should be punctured at an angle of about 30 degrees to the skin surface, with the bevel of the needle facing away from the skin. Verify that the needle is placed correctly by aspirating gently, that is, by pulling slightly on the syringe's plunger. Blood aspirated into the syringe or needle hub (known as a flash) indicates proper placement of the needle inside the vein. • Release the pressure: On the vein prior to starting the injection so that the blood circulation will carry off the solution as it is injected. Failure to release the pressure holds the solution in the vessel at the injection site, causing it to leak out of the vein at the puncture site into the surrounding tissues. • Hemostasis: When the injection is completed, pressure should be applied over the injection site simultaneously with the withdrawal of the needle. This pressure should be maintained for a short time to allow the puncture site to form a clot (hemostasis), which prevents bleeding into the surrounding tissues (a hematoma) from forming and keeps the injected material from leaking out of the vein. https://www.youtube.com/watch?v=UsUwT2e8W48 Intravenous Injections - Mice & Rats Intraperitoneal Injections • Intraperitoneal (IP) injections are made into the abdominal cavity of an animal. The term “peritoneum” refers to the tissue lining this cavity. • Danger: Care must be taken to avoid puncturing structures such as the bladder and cecum, so IP injections are commonly administered into the lower right quadrant of the abdomen (Figure 11.2). • Tilt down: Rodents should be restrained with their heads tilted downward to allow abdominal structures to fall out of the way. • Needle position: Insert the needle bevel up, at a 30-degree angle into the lower right quadrant of the abdomen. This places the injection away from the abdominal midline so as not to inject into the urinary bladder. • Aspirate: Always aspirate the syringe prior to an IP injection to confirm proper needle placement. A yellowish aspirate may indicate the bladder has been punctured. Likewise, a greenish-brown fluid suggests the needle has penetrated the cecum or intestines, and blood would indicate a vessel has been punctured. • Replace: If any aspirate is seen, the needle should be withdrawn, discarded and the procedure started again. Reusing a needle and syringe contaminated by an aspirate could cause injury or infection at the injection site. Intraperitoneal Injections https://www.youtube.com/watch?v=GbF6kK8bIXI Intramuscular Injections Intramuscular (IM) injections are generally given into a large muscle mass. Usually such injections are administered into the hind limb (Figure 11.3) in either: • The quadriceps muscle group, which is cranial (anterior) to the femur • The biceps muscle, which is lateral to the femur. Injecting these muscles avoids the sciatic nerve and the femoral vein, artery, and nerve that are located deep within the tissues caudal and medial to the femur; these structures can be injured by the needle or irritated by the material injected. Alternative locations: • Caudal thigh muscles: May be used in large species. Direct the needle caudally to avoid the femoral vessels and sciatic nerve. • Occasionally, IM injections are given in the muscles of the back or shoulders instead of in the hind limbs. • To inject IM in many species, grasp and stabilize the muscle mass with one hand and perform the injection with the other hand. • Aspirate: Prior to injecting the material, gently pull back the plunger on the syringe to ensure that no blood is aspirated into the syringe, which verifies that the needle tip is not inside a vessel. • Replace: If blood is aspirated, remove the needle, and apply pressure to the area to stop bleeding, then use another site for the IM injection. • Volume: Be careful not to inject too much material into a muscle; too large a volume would damage the tissue. In a mouse, the maximum IM injection volume is 50 microliters (0.05 mL). Subcutaneous Injections “Subcutaneous” means “below the skin” • Subcutaneous (SC or SQ) injections are placed between the skin and the underlying muscle (in the hypodermis). • Using one hand, gently pinch and pull up the skin to form a tent which allows for a large subcutaneous space into which the injection is made. • With your other hand, orient the hypodermic needle bevel up and parallel to the body at the base of the tent; then insert it through the skin into the subcutaneous space. • Avoid misdirecting the needle and entering the underlying muscle (changing the route of injection to IM) or passing through the other side of the tent (losing the injected material on the skin surface). • Danger: Direct the needle away from your fingers to prevent needlestick injury and inadvertently injecting yourself with the compound. • Volume: In many species, large volumes of non-irritating fluids may be administered SC because the skin is only loosely adhered to underlying tissues. Intradermal Injections • Intradermal (ID) injections are given into the dermal layers of the skin (Figure 11.4B). • Although the procedure is not painful, anesthesia is commonly used to immobilize the animal for intradermal injections because it is difficult to accurately place the compound in a conscious animal. A routine ID injection is for tuberculosis testing of nonhuman primates. Test material is injected into the eyelid intradermally. This location allows for easy reading of the test results. • Needle size: A 25 gauge or smaller hypodermic needle is often used with a 1 mL syringe. The skin is shaved and prepped, and the needle is inserted at a shallow angle (10–20 degrees) between the layers of the skin. • Placement: The needle is advanced only until its bevel is inserted within the skin. • Volume: Only a small volume (0.1 to 0.2 mL) may be injected at a single site. Multiple injections are necessary for a larger quantity. • QC: Correct placement of an intradermal injection will result in a bleb, which is a bulge observed following ID injection of fluid. Intracranial Injections https://youtu.be/41P3hJEa96w • Many compounds in the circulating blood are blocked from entering the brain by the blood-brain-barrier. The blood-brain barrier is a tissue lining in the blood vessels of the brain. This tissue lining separates the circulating blood from the brain, thus protecting the brain tissue from many chemicals. • To bypass the blood-brain barrier and infuse compounds such as viral vectors and fluorescent compounds directly into the brain, • Injections may be made through the skull of an anesthetized animal and into the cerebrospinal fluid (CSF) within the ventricles of the brain. It is vital to identify the anatomical structures of the skull, particularly the sagittal suture and the points at its cranial and caudal ends, which are known as the bregma and lambda, respectively • The injecting 30-gauge needle must be positioned correctly to access the ventricles. • This may take place in a stereotaxic apparatus (discussed in more detail in Chapter 10) that holds the head steady. • Alternatively, this technique may be done freehand, particularly if the injections are being performed in neonatal animals that have thinner bones in the skull. Intracranial Injections https://youtu.be/41P3hJEa96w • Content advisory: This video shows the full procedure, which includes exposure of the skull after a surgical incision on the scalp of a mouse. There is another method that uses a template instead of the stereotax. Retroorbital Injections • While intravenous injections in a rodent’s tail vein are an excellent way of administering substances into the bloodstream, the technique is difficult to master. If the needle is not correctly inserted into the vein, any substance injected can be lost in the surrounding tissues. • Injection in the retroorbital sinus is another method that may be used in rodents, although this route is generally discouraged because of the potential for an eye injury. • The retroorbital sinus is a complex network of blood vessels behind the eyeball (Figure 11.6). • To inject retroorbitally (RO): • • • • Anesthetize the animal and place it in lateral recumbency. Grasp the skin of the face and pull it taut, so the eyeball protrudes slightly. Apply a topical ophthalmic anesthetic to numb the area and prevent drying of the cornea. Insert a 25-gauge needle, bevel side down, into the “corner” of the eye, where the eyelids meet (the medial canthus) at a 30-degree angle toward the midline. • Inject the substance and gently remove the needle. • Release the facial skin, so the eyeball returns to the socket and keeps the liquid from leaking out. • Volume: Adult Mice - no greater than 150 microliters. Neonates no more than 10 microliters. Gastric Gavage https://researchanimaltraining.com/articles/oral-gavage-in-the-mouse/ • Gastric gavage: The administration of a solution directly into the stomach by intubation or injection down the esophagus. • This method is especially effective for rodents and rabbits because these species are physiologically unable to regurgitate stomach content. • A special gavage needle with a round, bulbous tip is used to prevent injuring the animal’s esophageal lining (Figure 11.7). The needle used may be rigid or flexible, curved, or straight. The choice is according to operator preference. • The choice of needle length and gauge depends on the species, size of animal, and viscosity of the solution to be injected. • The needle is attached to a syringe containing the material to be gavaged. • To administer the solution, the animal should be restrained in a vertical position with one hand. • Mouse Example: • • • • • • • Restrain by scruffing With the other hand, check the needle length by measuring the needle from the corner of the mouth to the xiphoid process at the caudal end of the sternum to ensure it is the correct length and will not puncture the stomach. Gently introduce the bulb of the needle into the animal’s mouth, and using the syringe and needle as a lever, tilt the animal's head to a vertical position in alignment with its body (Figure 11.8). Gently advance the needle. No force should be used; gravity and the weight of the needle itself should be sufficient to advance the needle into the stomach. If any resistance is felt, immediately stop, remove the needle, and start the procedure over. Once the needle tip is in the stomach of the animal, slowly administer the gavage solution into the stomach. Keep the needle stationary until all material is expelled from the syringe; this prevents aspiration of the material into the lungs. Once the administration is complete, gently, and slowly withdraw the needle. With proper training and practice, this procedure can be performed easily and efficiently with minimal stress to the animal. Gastric Gavage - mouse https://researchanimaltraining.com/articles/oral-gavage-in-the-mouse/ Gastric Gavage – Large Animals • The stomach tube should be measured from the corner of the mouth to the xiphoid process of the sternum to prevent injury to the animal. A mark may be made on the tube with tape or an indelible pen to show how far to advance it. • Place a bite bar in the animal’s mouth and secure it in place with one hand. • The gavage tube is then inserted into the animal’s mouth through a hole in the bite bar and advanced until it reaches the pharynx. • Slight resistance will be encountered at the pharynx, usually causing the animal to swallow, which permits the advancement of the tube into the esophagus and then the stomach. • Ensure that the tube does not enter the trachea, as delivering the gavage solution into the lungs would most likely cause death. Inadvertent entrance into the larynx will cause paroxysmal coughing and struggling; if this happens, quickly remove the tube, and allow the animal to recover before proceeding. • Following administration of the solution, the tube is folded or kinked to prevent aspiration of any remaining liquid during the withdrawal of the tube. • Virtually any species can be gavaged if an appropriate bite bar and stomach tube can be found. Bite bars are easy to make. For example, a syringe casing can be cut, and the rough edges smoothed to make a bite bar suitable for animals ranging from rabbits to sheep. • Two species that are notable for unique anatomical features of the pharynx affecting intubation are the guinea pig and the chinchilla. The soft palate of these species is attached to the base of the tongue, forming a membrane between the mouth and pharynx. A small hole in the membrane called the palatal ostium allows food, water, and air to pass through. Great care must be taken when gavaging these animals to prevent damage to this membrane, which is rich in blood vessels and can hemorrhage if damaged. Gastric Gavage – large animal https://www.youtube.com/watch?v=P7PUU1eLZIQ Blood Collection Blood Collection • Collecting blood from laboratory animals is a common procedure used to aid in the diagnosis of disease or to support data generation in experimental studies. • The total blood volume in an animal’s body equals approximately 6% of its total body weight. This figure varies somewhat, depending on such factors as species, age, and sex, but 6% is a good average. This means that a 3 kg rabbit will have a total blood volume of about 180mL. (Note that this calculation can only be done in the metric system because of the convertibility of weight and volume units of measure.) • When blood samples are collected at frequent intervals, institutional guidelines on blood collection limits ensure that the health of the donor animal is not adversely affected. • A common guideline is that a volume equal to 1% of the animal's total body weight can be safely taken every 2 weeks without causing side effects associated with blood loss, such as anemia. For a 3 kg rabbit, this means that 30 mL of blood can be safely taken every 2 weeks. • The vessel and the amount of blood to be collected are determined by the species and type of tests to be performed. • Venous blood collection is similar to the technique for intravenous injections. The skin site is prepped with an antiseptic both for skin disinfection and vein visualization. Unlike intravenous injection, however, the proximal pressure on the vein is maintained throughout the blood collection, keeping the vein distended to pool blood to be drawn. Once the required volume of blood has been collected, pressure on the vein is released. As the needle is withdrawn, pressure is applied to the puncture site until hemostasis is achieved. Saphenous Vein • The saphenous veins on the lateral side of the lower rear leg may be used for blood collection in both large and small laboratory animals. • Anesthesia is not necessary, but appropriate restraint is important. • Prepare: Shave the site and prep with an antiseptic. • Large animals, the vein is cannulated with a needle and blood is collected into a syringe or a vial (such as a Vacutainer®). • Small animals such as mice and rats, a hypodermic needle or lancet is used to puncture the vein, not to cannulate it (like testing blood sugar). Blood is collected as it wells onto the skin surface, either wicked up by a capillary tube or dripped into a small vial. When collecting blood from the skin surface, silicone grease may be applied onto the skin, after disinfection and prior to vein puncture, to help the blood well up and to minimize clotting. https://vimeo.com/170066703 Saphenous Vein Hold off with gauze once done collecting https://vimeo.com/170066703 Facial or Maxillary Vein • Facial or maxillary vein blood collection is an easy and safe way to collect a blood sample in mice and rats (Figure 11.10), and it is preferred as an alternative to retroorbital bleeding (see description of this procedure later in the chapter). • Anesthesia is not required; the animal can be manually restrained by scruffing and pulling the skin taut over the face. If it gasps (repeatedly opens its mouth), lessen the tension on the nape so the animal can breathe easily. • To collect blood, a needle or lancet is used to puncture the vessel behind the mandible and in front of the ear canal. A lancet is often preferred because a needle can be inserted too deeply into the tissues. • Align the lancet caudal to the angle of the mandible, extending toward the ear. Locate the approximate midpoint of the line represented by the lancet; this is the puncture site on the cheek. This midpoint coincides in mice with a small indentation in the skin that can be used as a landmark. In white rats, a helpful landmark is a gray spot on the jawline, directly below the lateral canthus (corner) of the eye; the puncture site is approximately 0.5 cm behind the gray spot. • The puncture is made with the lancet held at a 30-degree angle. • Blood should flow immediately and can be collected into an appropriate container. • Following blood collection, maintain gentle pressure at the puncture site until hemostasis occurs. https://www.youtube.com/watch?v=RNyWrhHHwqc Facial or Maxillary Vein Hold off with gauze once done collecting https://www.youtube.com/watch?v=RNyWrhHHwqc Tail Snip (Transection) {Arterial vessel} • A tail snip can be used to collect several small drops of blood from mice or rats. • The end of the tail is prepped with a disinfectant, then 1 to 2 mm of the tip is snipped off using a sterile scalpel blade or scissors (Figure 11.11). • Blood dripping from the tail is collected in a capillary tube. • A gauze sponge should be applied with pressure at the site to control any bleeding following the procedure. A drop of tissue glue may also be used to control bleeding. • Using a tail snip in animals older than weaning age is controversial, as the cartilage of the tail is replaced by bone. In these animals, anesthesia is strongly recommended. Retroorbital Sinus or Plexus {Arterial Vessel} • Retroorbital blood collection is a method of collecting blood from species with a large venous sinus, such as mice, hamsters, gerbils, ferrets and guinea pigs, or a plexus posterior to the eye like the rat. Pigs may be bled from a venous sinus ventral to the eye. • Retroorbital blood collection procedures are performed with the animal under anesthesia, although it can be done on conscious animals using a topical anesthetic. • The retroorbital route is generally discouraged because of the potential for an eye injury. Other routes that are shown to be effective for injections and blood collection are preferred. The retroorbital technique is described here, however, because it continues to be used in some settings. • To access the site in a rodent: • Secure the head between the thumb and forefinger and insert a hematocrit (capillary) tube, or alternatively a needle, at the medial canthus of the orbit of the eyeball. • Direct the tube toward the back of the eye socket (Figure 11.12). The tube is inserted at a 45degree angle at the medial canthus of the eye, avoiding the globe. • While gently advancing the tube, carefully rotate it to puncture the sinus and collect blood. • At the end of the procedure, the eyelid should be held closed for a few seconds to allow the punctured sinus to clot. It is also common practice to apply a small amount of ophthalmic ointment onto the eye following the procedure. • Repeated sampling should be minimized; institutional policies may limit repetition intervals and require an alternation of eye used. Cardiac puncture • Cardiac puncture (puncturing the heart) is used when a large volume of blood is needed from small animals (Figure 11.13). It is performed as a terminal procedure; the animal is euthanized at the end of the blood collection. • How to: • Place the animal under general anesthesia. • Method 1: Place animal on its side and access the heart the intercostal spaces (between the ribs). • Method 2: Place the animal in dorsal recumbency and direct the needle cranially in the angle between the xiphoid process of the sternum and the ribcage. In this approach, the needle penetrates the diaphragm to reach the heart. • Method 3: Introduce the needle from the top of the rib cage, inserting the needle into the right ventricle. • Withdraw blood slowly to prevent the heart from collapsing. Maintain either a mild steady vacuum or a pulsing vacuum by action of the syringe plunger allows the chambers of the heart to refill with blood during the procedure. • It is possible for even an experienced technician to unintentionally lacerate the lung, the pericardial sac, or the heart wall, thus killing the animal. Therefore, technicians performing this procedure should be highly trained and proficient. https://www.youtube.com/watch?v=p4xJo7dqrgU Cardiac puncture Which method are they using? https://www.youtube.com/watch?v=p4xJo7dqrgU Jugular vein • The jugular vein is used for collecting larger volumes of blood from research animals. Rats, dogs, pigs, and ruminants such as sheep and goats are routinely bled from the jugular vein as it is easily accessed, and a relatively large volume of blood can be taken. • In large animals, vacutainers may be used for sampling. In this method, the sample collection tube is evacuated, so blood flows into it without needing to be under pressure. • Insert the needle into the vein and once it is positioned correctly, attach the sample collection and collect the sample. • One benefit of using vacutainers is that the blood samples can be collected directly into a tube containing anticoagulant, or if the cells are not required, a tube containing a gel that will separate the cells from the serum after centrifugation. https://www.youtube.com/watch?v=SOE8-0rZGdE Jugular vein https://www.youtube.com/watch?v=SOE8-0rZGdE Arterial Blood • Central (auricular) Ear Artery: Collection site for rabbits and pigs via the central ear (auricular) artery. This artery is easily seen along the center of the ear lobe or pinna (Figure 11.14). Larger quantities of blood can be drawn from this artery than from the smaller marginal ear veins where blood flow is slow (under venous pressure). • Femoral Artery: Used for dogs & NHPs. • The technique for arterial blood sampling is similar to venous blood collection, except that proximal pressure is not applied to the artery during blood collection because the artery is always filled with sufficient blood for collection. • Arteries take much longer to clot than veins, be sure to apply pressure to the artery for several minutes following withdrawal of the needle to prevent hemorrhage and hematoma formation. • Do not return the animal to its cage or pen until you are certain the bleeding has stopped. • Hemorrhage or tissue irritation will impair further use of the vessel, so take extra precautions when collecting serial blood samples. Figure 11.11. Other Procedures Urinary Catheterization • A urinary catheter is placed into the urethra of the animal to allow urine to flow out of the body for collection of urine, or to facilitate urination (most often male cats with a blocked urethra). In some species of animals, such as male cats, a catheter may be used to unblock the urethra. • What is a catheter? A catheter is usually a flexible tube with a tapered end. Catheters are sized in the French scale, abbreviated as Fr. For example, an 8 Fr catheter measures 2.7 mm around the outside diameter. • A complicating factor in urinary catheterization is the bacterial flora of the urethra. Although the bladder and proximal urethra are normally sterile, the distal urethra usually contains bacteria, which makes it impossible to avoid carrying some bacteria into the bladder. Thus, even with flawless aseptic technique, the possibility of introducing bacteria into the bladder exists. Every effort must be made to minimize the introduction of pathogens into the urinary tract. • Another complicating factor in urinary catheterization is the internal location of the urethral orifice in females of most species. This location makes urinary catheterization more difficult than in males because seeing the urethral orifice is generally necessary for catheter insertion. • To insert: Apply sterile lubricating jelly to the tip, which is then advanced gently through the urethra into the bladder. • To catheterize females, a vaginal speculum (a device that is inserted to hold open a body passageway) and good illumination are required. A common type of catheter used in large animals is a Foley catheter, which has a balloon in the tip and a syringe port at the other end (Figure 11.15). Once the catheter (with the balloon uninflated) is passed through the urethra and its tip reaches the bladder, a syringe is connected to the port to inject fluid or air, which inflates the balloon. The inflated balloon holds the catheter tip in the bladder, preventing it from slipping out. Urinary Cystocentesis • If a sterile urine sample is desired, then the bladder may be tapped, a procedure which is called cystocentesis. The animal should be tranquilized, and the skin site for needle penetration may be numbed using a local anesthetic. Cystocentesis is then collected by passing a hypodermic needle through the surgically prepared abdomen and into the bladder. • https://www.youtube.com/watch?v=aftdk02aJi0 Urinary Cystocentesis Endotracheal Intubation • Often when extended anesthesia is required, as for surgery, an endotracheal tube is inserted into the trachea to provide an airway in which gas anesthetics or oxygen can be administered. The endotracheal tube is valuable for controlling the airway when an animal has to be resuscitated, such as when breathing has ceased due to the anesthesia or the surgical procedure. • The animal is first anesthetized with a short-acting injectable anesthetic, which permits the tube to be passed into its trachea (Figure 11.16). Anesthesia is necessary because endotracheal intubation is impossible in a conscious animal. • The endotracheal tube is connected to a gas line from an anesthesia machine, which maintains the anesthesia through administration of an anesthetic gas. • To place an endotracheal tube in large animals, the tube is often guided by a laryngoscope that illuminates and exposes the larynx (the opening to the trachea; Figure 11.17). The laryngoscope consists of a handle, a long blade, and a light at the blade tip. The blade has smooth edges, so it does not traumatize tissues. The animal is anesthetized and placed in a recumbent position appropriate for the surgery. The technician opens the animal’s mouth and inserts the blade of the laryngoscope into the oral cavity. The laryngoscope is advanced until the larynx is visible. Endotracheal Intubation • A topical anesthetic, often lidocaine, may be sprayed onto the larynx to prevent tissue swelling due to any trauma caused by inserting the tube through the larynx. After allowing the topical anesthetic to take effect, the technician introduces the endotracheal tube into the oral cavity, passes it through the larynx, and advances it into the trachea. • Proper placement of the tube in the trachea must be verified before connecting the tube to a gas anesthesia machine or initiating another procedure. There are multiple ways for verifying endotracheal tube placement: • Listen to the end of the tube for breath sounds, hold a very light fiber or animal hair to the tube opening and observe its movement as the animal breathes. • Hold a stainless-steel surface up to the end of the tube and look for condensation indicating expiration. • If the tube is clear, watch for alternating condensation and clearing within the tube itself. • Once proper placement is confirmed, a syringe filled with air is used to inflate a cuff that will hold the endotracheal tube in place. • This cuff is located on the outside of the tube, near its tip within the trachea. • The cuff expands with air to seal the airway around the tube so that ventilation via the tube can be controlled. The inflated cuff also prevents the passage of fluids, like secretions from the mouth and throat, into the lungs. • Once the cuff is inflated, the tube should not be moved, rotationally or up or down the trachea. That is because motion of the tube would cause damage to the tracheal lining from the inflated balloon. • Once the balloon cuff is inflated, the endotracheal tube can be attached to any of several devices. • If the endotracheal tube must be repositioned inside the trachea, always deflate the balloon first. Then once the tube has been repositioned, it is safe to reinflate the balloon. Endotracheal Intubation • The endotracheal tube can be attached to any of several devices, such as an: • Anesthetic machine. As the animal breathes, it inhales anesthetic. • Automatic ventilator that drives the animal’s breathing. The ventilator may be a component of an anesthetic machine. • AmbuR bag for manual ventilation. • While anesthetized, the animal’s respiration is just one of the parameters that should be monitored. Endotracheal Extubation • After the procedure, at an early stage of anesthetic recovery, the animal will start to cough and gag. • At that point, the endotracheal tube is removed, allowing the animal to breathe on its own. • Before removing the tube, the cuff must be deflated by aspirating air through the injection port. When the pilot balloon has collapsed, it is safe to withdraw the endotracheal tube. • NOTE: In small animals, like mice and rats, endotracheal tubes are often placed using an otoscope to illuminate and magnify the view of the larynx. An endotracheal tube for these species lacks a cuff; the tapered tube seats itself in the larynx to seal the airway around the tube. Technical Procedures in Aquatic Species The primary focus of this section is on procedures for the two most common aquatic laboratory species, the zebrafish (Danio rerio) and the African clawed frog (Xenopus laevis). The procedures described are also generally applicable to other aquatic species used in the laboratory. Substance Administration Handling and restraint of aquatic species are more difficult than with mammalian species, particularly for complex or time-consuming technical procedures. Some procedures that are commonly performed in mammalian research animals, such as intravenous substance administration, are not possible or practical in aquatic species. There are two routes of administration that are unique to aquatic species: immersion and intracoelomic injections. Immersion • Most fish and many amphibians perform their gas exchange through either gills or the skin. Both these structures contain rich capillary beds that allow gas exchange to occur, but also makes fish and amphibians sensitive to chemicals in the water. This means that these animals may be immersed for administration of test agents, medication, or anesthesia. • In this technique, aquatic animals are placed in water containing a drug, such as the anesthetic MS-222, which is absorbed either through the gills of fish and some amphibians (branchial absorption), or through the skin in larval fish and all amphibians (cutaneous absorption). • Due to their lack of easily accessible veins and the impossibility of using inhalant anesthetic, immersion is the preferred method of anesthetizing fish. • When administering compounds by immersion, the chemical activity of the compound should be determined, as the rate of compound uptake can be influenced by water chemistry, temperature, and light. Intracoelomic Injections • Unlike mammals, fish and amphibians do not possess a peritoneal cavity. Instead, the internal body cavity that houses the abdominal and thoracic organs is called the coelom. • Like the peritoneal cavity, the coelom may be used as a site for substance administration, referred to as intracoelomic or ICo. • Amphibians: • Hold in dorsal recumbency, head pointed downwards so the viscera fall away from the injection site. • Insert the needle into the lower right quadrant. • Aspirate gently to ensure no blood is seen, otherwise the needle may be in the spleen. • Discard: If blood is seen, remove and discard the needle. Use a fresh needle for administering the treatment or test substance. • Fish: Take care to not perforate the gut. • Fast for 24 hours to avoid the abdominal viscera, which are sensitive to many compounds. • Administer along the midline, cranial to the anus (urogenital pore) and caudal to the pectoral fins. The urogenital pore must be avoided when administering compounds. Percutaneous Injections • Zebrafish are seldom given percutaneous injections due to their small body size and the small volume that can be given. • One exception to this is when fish are injected with visible implant elastomer (VIE) tags for identification. • In this process, a small amount of nontoxic silicone dye is injected just under the skin in multiple anatomical locations, such as by the tail, under the eye, or by the pectoral fin. • Amphibians may be injected just under the skin of the back, cranial to the cloacal papilla. This is where the dorsal lymph sacs are located, proximal to the posterior lymph hearts. • VOLUME: Using a small (22-28 ga) needle, a volume less than 0.5 mL may be injected, just under the skin and medial to the lateral line. https://www.youtube.com/watch?v=79fSt_e7YtU Intramuscular Injections • Fish IM: Use the dorsal epaxial or abdominal muscles.Care should be taken to avoid the lateral line and the ventral blood vessels. • Go Slow: Administration should be slow to prevent leakage. • Volume: less than 0.05 mL/50g fish due to the small size of Zebrafish • Amphibians IM: Use the quadriceps muscle. • Volume: The maximum volume is usually less than 0.5 mL, depending on the size of the amphibian. Gastric Gavage Gastric gavage may be performed on amphibians, but it should be noted that they absorb drugs poorly through the stomach. Xenopus may regurgitate when stressed, or if noxious substances are administered by gastric gavage. • To gavage a Xenopus, • Anesthetize the animal until it ceases swimming motions and loses its righting reflex when placed on its back. • A 1.5-inch gavage needle should be introduced at the corner of the mouth and guided gently into the stomach. Either a soft plastic or steel gavage needle may be used, but in both cases, care should be taken not to use a needle that is too long, as the stomach may be perforated. • Fish: Administer by mouth either by mixing the compound into food or gelatin, by bioencapsulating compounds into brine shrimp larvae, or by esophageal gavage. • One drawback of using medicated feed is the difficulty in accurately quantifying intake. • If esophageal gavage is performed, fish should be fasted prior to gavage so the solution flows into the esophagus freely. A flexible rubber tube is inserted into the mouth and into the esophagus past the gills (approximately 1 cm in zebrafish). • Volume: In general, the dose rate for fish should not exceed 1% of body weight. Blood Collection - Amphibians • For many species of frogs and salamanders, the mid-ventral abdominal vein can be used for blood collection. • Xenopus lack both a tongue and a tail and thus, do not have a lingual or tail vein, which are two anatomical sites that are commonly used for blood sampling in other amphibian species. • The easiest method for blood sampling is to anesthetize the animal and take blood by cardiocentesis. Cardiocentesis (also known as a cardiac puncture) is usually a terminal procedure that must only be performed on animals that are anesthetized and will be euthanized at the end of the procedure. • Blood samples should be handled gently, as for any species. • The choice of anticoagulant used for blood collection is important, as using EDTA may cause hemolysis. An alternative to EDTA is heparin or citrate, although it should be noted that some clotting may occur with either of these two anticoagulants. Blood Collection - Zebrafish • Zebrafish are challenging to take blood samples from due to their size, small blood volume, and lack of accessible peripheral vasculature. • Caudal vein: commonly used to take blood, but it has a very small diameter. • To draw blood from this site, the vein must be cut with a scalpel at the base of the caudal fin. However, this means any sample acquired using this technique can be contaminated with tissue fluid, skewing the clinical chemistry results. • To extract an uncontaminated sample, cardiocentesis has been a preferred method, although a terminal procedure. • Doral aorta: An alternative method which uses a very fine diameter glass pipette to extract blood from the dorsal aorta. • This procedure is minimally invasive so that serial samples may be collected. • If serial samples are taken from zebrafish, samples should not exceed 3 microliters because of their very small blood volume. • Tail Puncture: Used for larger fish for repeated blood collections. Gamete Collection • Oocytes and sperm are used for studies in molecular biology, embryology, and biochemistry. • Egg and sperm collection are necessary for in vitro fertilization, and large numbers of synchronously developing embryos can be obtained from fish or amphibians. • The terms “stripping” or “milking” are commonly used to refer to collecting either ova from females or sperm from males without harming them. • Oocytes and sperm may be cryopreserved. • In fish, semen is known as milt. Sperm Collection • Zebrafish – Stripping Technique: • • • • Anesthetize the fish following protocol-approved methods. Dry the ventral side near the urogenital pore completely as water will activate the sperm. Transfer the male, ventral side up, to a sponge animal holder and position on a stereomicroscope to view the urogenital opening. Expel milt by positioning smooth forceps on both sides of the fish and applying pressure, while moving the forceps in a cranial to caudal direction. • • Milt is collected in the calibrated end of a 10 μL microcapillary tube. Transfer the fish to system water and allowed to recover from anesthesia. • Zebrafish – Testes dissection technique: • • • • Euthanize the male fish and dry by rolling on paper towels Transfer the fish transferred to a Petri dish and position under a stereomicroscope Using dissection scissors, open the abdomen Retract the intestines and swim bladder cranially to expose the testes. The testes can be identified as bilateral white tissue just ventral to the swim bladder. • Xenopus: • • Pretreat the frogs with an injection of synthetic human gonadotropin releasing hormone (GnRH) 1–5 hours post injection, hold the male frog over a Petri dish and milt is released. • Xenopus – lavage: Sperm can be also be harvested by gentle mechanical stimulation of the abdomen or by cloacal lavage • Gently insert a sterile Pasteur pipette into the cloaca and introduce a small volume of sterile saline; this is often sufficient to start the release of milt • Xenopus – Terminal Testes harvest: • • • • Deeply anesthetize the frog before placing in dorsal recumbency. Make a horizontal incision in the lower abdomen to expose the organs of the coelom, and then make a vertical incision, so the flaps of skin and musculature can be retracted Push aside the organs and fat bodies to exposed and remove the testes. Euthanize the frog once the collection is complete Oocyte Collection https://www.youtube.com/watch?v=3ldTyIpcM_4 • Zebrafish females: • • • Anesthetize and then dry by rolling the fish on a paper towel (as with the males, careful attention must be given to drying the ventral surface of the fish near the urogenital pore because water activates the gametes). Hold the female between the thumb and index finger, gently squeeze while applying light pressure on the ventral abdomen in the direction of the urogenital opening while rolling fingers slightly. This maneuver easily expresses the ova. Transfer the female to system water to recover from anesthesia. • Xenopus females: Prepare for egg harvest by either a one- or two-step induction process. • Hormone Two-step process: • • • Inject the frog with pregnant mare serum (PMS) into the dorsal lymph sac 3–7 days before use to induce oocyte maturation. To stimulate egg laying, an injection of human chorionic gonadotropin (hCG) is administered into the dorsal lymph sac, and the frog is placed in a tub of either tank water or buffered egg laying solution to lay her eggs. The precise onset and duration of egg laying varies from frog to frog and from injection to injection. • Hormone One-step process: administer a single dose of hCG (without the PMS pre-treatment) and the frog will lay her eggs the next day. • Manual expression: • • Hold the female and apply gentle pressure, while stroking downward on the ventrum. Care should be taken to stroke the female gently, as bruising and rupture of internal organs can occur if too much pressure is applied. This procedure mimics the natural mating behavior called amplexus, where the male climbs on the back of the female and grasps her tightly with his forearms. After 1–2 minutes of stroking, the female will start laying eggs. • Surgical Laparotomy: • • • • • • Decisions about surgical site prep, anesthesia, and post-op analgesia should be under the direction of the veterinary staff. Sterile gloves and instruments are required. After the skin and muscle layer incisions are made, a portion of the ovary can be excised to reveal sufficient oocyte mass for collection. After removal of the desired number of oocytes, the remaining ovarian mass is carefully placed back in the coelomic cavity. Closing the skin and muscle in 2 layers with absorbable suture avoids the need to remove sutures at a later time. A limit of 6 surgeries over the life of the animal is the recommended maximum, with 3 to 6 months rest time between surgeries. Oocyte Collection https://www.youtube.com/watch?v=3ldTyIpcM_4 Tissue Sampling • It is often necessary to obtain a small tissue sample from aquatic species for genotyping. With fish, this is usually accomplished by taking a small clip of the caudal fin from an anesthetized animal. Recovery is fast and the fin regenerates. • Tissue samples may also be taken from Xenopus, but only when they are in the larval stage. The tadpoles must first be lightly anesthetized, and then 3 – 5 millimeters of the tip of the tail cleanly severed with a scalpel. Do not remove too much tissue, because cuts made too far cranially to the tail tip are likely to sever the major blood vessels running along the tail, leading to excessive bleeding and death. Common Laboratory Tests Laboratory animal technicians may be assigned to perform routine laboratory tests, such as blood tests, fecal examination, and urinalysis, on the animals under their care. These tests may be requested for veterinary exams or to collect study data. Blood Tests • Tests are run on mammal and aquatic blood samples to yield information on the cell types and chemical constituents that are present. The values of these parameters are interpreted by a veterinarian to determine the animal’s status of health or disease. • For each type of blood test ordered, specific anticoagulants are used. An anticoagulant is a chemical that blocks the clotting of blood (coagulation). • Tube types for various tests differ chiefly by the presence or absence of an anticoagulant. There are multiple anticoagulant agents, each of which acts by a different mechanism. Specific anticoagulants are used for each type of blood test ordered. Blood tubes with an invisible interior coating of anticoagulant are commercially available. The type of anticoagulant, or lack of one, in a blood tube is signified by the color of the rubber stopper or cap on the tube. This color scheme is consistent among manufacturers. • • Tubes intended for serum have an interior coating of silicone, which activates and hastens clotting. Some tube types contain a gel polymer which, during centrifugation, separates the plasma from the cells, or the serum from the clot. This polymer has a density intermediate between the plasma/serum and the other fraction of the sample, so it separates out these layers through centrifugation. • If blood is collected in a syringe, it must be rapidly transferred to the correct type of blood tube for the test to be run. • Upon stoppering a blood tube, it is important to gently invert the tube and mix the additive with the sample. Do not shake the tube, as that can cause hemolysis and contaminate the plasma or serum with cell fragments, affecting test results. Gentle handling and refrigeration are necessary to protect and preserve the samples until testing. • Separated serum and plasma may be stored frozen but freezing will destroy the cells in whole blood. • Be sure to label each tube appropriately and completely. Blood Tests • Microhematocrit tube, aka capillary tube: A common blood collection container used to determine anemia, (a deficiency in the number or size of red blood cells). • A small sample of blood is collected into a microhematocrit tube and spun down in a centrifuge. • The tube is then held against a chart and the reading is taken. • Microhematocrit tubes may be heparinized (interior coated with heparin) or plain (no anticoagulant). • Be sure to use the type of tube specified by the veterinarian for these tests. Urinalysis • Urinalysis can be used to test for abnormalities in the urinary tract, including the kidneys and the bladder, and some metabolic functions. • Urinalysis consists of visual inspection of the urine sample for color and turbidity (cloudiness), measurement of the specific gravity, chemical analysis, and centrifugation to examine the heavier particles, or sediment in the urine. The specific gravity test is used to determine how well the kidneys are able to concentrate the urine. • Specific Gravity Test: A drop of urine is placed onto an instrument called a refractometer. The laboratory animal technician looks through the refractometer lens and takes a specific gravity reading from the instrument. • Chemical analysis : Is performed using a dipstick. The dipstick has pads that contain different chemical reagents to test for a specific substance in the urine. The dipstick is dipped into the urine and the colors of the pads on the dipsticks are read against a chart to determine the values for chemicals in the urine. Some common factors measured using dipsticks are pH, protein level, glucose, and the presence of blood in the urine. • Centrifugation: Used to detect the presence of cells or debris that may indicate a health problem. A urine sample is placed into a centrifuge tube and spun down, forcing the sediment in the urine into the bottom of the tube. The urine is poured off, and a sample of the sediment is placed onto a slide. Commonly, a drop of stain is added to the sediment on the slide to aid its examination under a microscope. • White blood cells (WBCs) and bacteria are signs of kidney or bladder infections. • Mineral sediment may indicate bladder stones. • Protein and cellular casts of nephron tubules provide information about kidney malfunction. Parasite Tests • Fecal examination is commonly used to test for internal parasites, also called endoparasites, or to identify other abnormalities such as blood in the stool. An infected animal must be treated to rid it of any parasites and the animal may need to be removed from an experimental group or placed in quarantine until infestation has been resolved. • Fecal floatation: A sample collected from the animal is placed into a prepared solution, often from a kit, to break it up. A drop of the fecal mixture is then placed onto a slide and examined under a microscope. Parasites may be present in the feces as either eggs or mature organisms, depending on the parasite species. • Cellophane tape test: Used to detect pinworms of the Syphacia genus, an endoparasite found in the intestines of rodents. • Because these pinworms lay their eggs on the exterior of the animal, around the anus, these eggs can be picked up using the sticky side of a piece of clear tape pressed against the anal area (Figure 11.19). • The tape is placed on a microscope slide, sticky side down, to view the pinworm eggs (Figure 11.20). • PCR Testing: Parasites may also be detected using PCR. The benefits of PCR are numerous. The sensitivity of PCR means only a tiny amount of material is required for detection, and unlike serology, detection is not dependent on the immunocompetence of the subject. PCR can be used to test different types of samples; feces, fur, or anal swabs, and both endo- and ectoparasites can be detected using this method. Vaginal Cytology - Lavage • Assessing the reproductive status of rodents is a necessary step in performing both timed matings and the study of reproductive dysfunction in toxicological studies. • In rodents, the ratio of different cells types in the vagina correspond to the stage of estrus and thus, the levels of reproductive hormones present. • Proestrus stage, the cells in the vagina are predominantly nucleated epithelial cells • Estrus: The majority of cells will be clumps of anuclear, cornified squamous epithelial cells. • Invasive techniques such as taking swabs of the vaginal canal may affect reproductive status and induce an inflammatory response that confounds cytological analysis. For these reasons, vaginal lavage may be used instead. • Vaginal lavage allows the cells of the vaginal canal to be sampled and visualized with crystal violet stain. Staining the cells makes them easy to identify, so determining the current stage of the estrus cycle is straightforward.