Cranial Vena Cava: The cranial vena cava is located in the thoracic inlet between the first pair of ribs. The right side of the animal should be used to prevent damaging the phrenic nerve, which is anatomically more protected on the right side of the animal than the left. Hitting the phrenic nerve may alter the function of the diaphragm and can result in life-threatening cardiac or respiratory problems. For piglets, a 20-gauge × 1.5-inch needle is used. An 18- to 20-gauge × 1- to 1.5-inch needle is appropriate for small pigs (up to approximately 25 kg). For pigs weighing more than 25 kg an 18- to 20-gauge × 1.5- to 3.5-inch needle is used. Large adults require a 16- to 18-gauge × 4- to 4.5-inch needle.

To collect blood from a small pig, the animal is placed on its back (dorsal recumbency) on a 45-degree incline with the head lower than the hips. The head is extended and front legs pulled caudally. The jugular furrow is visualized, and the needle with a syringe attached is inserted in the furrow lateral to the manubrium of the sternum. The needle is pointed toward the caudal aspect of the top of the opposite shoulder blade (Figure 20-55). As the needle is inserted, the syringe plunger should be pulled slightly to maintain negative pressure. When blood enters the syringe, the needle is held in place while the desired amount of blood is aspirated.

Larger hogs are restrained with the use of a hog snare in a standing position with the head slightly elevated. The person with the syringe crouches in front of the right side of the pig facing the body of the pig or can crouch to the side of the right shoulder facing the neck. The needle is inserted into the right jugular furrow lateral to the manubrium (Figure 20-56), directed toward the shoulder. The cranial vena cava in large pigs is deep (4 inches may be required to reach the lumen).

Jugular Vein: The jugular vein can be used for a blood sample collection in pigs of any age. It is located in the jugular furrow. It is not as deep as the vena cava, so fewer potential complications are associated with its use. The jugular vein has a smaller diameter than the vena cava and is more difficult to access, especially in large or heavy pigs. The needle size selected depends on the size of the animal. A jugular venipuncture in piglets can be done with a 20-gauge × 1.5-inch needle. Large pigs may require 16-gauge × 3- to 3.5-inch needles. To prevent the puncture of the phrenic nerve, the right side of the animal should be used when possible. The needle should be inserted cranially to the manubrium where the jugular furrow appears deepest. The animal is restrained as for a vena cava venipuncture. An imaginary horizontal line that passes through the shoulders and manubrium sterni is visualized. A second line is visualized that extends from the manubrium sterni to the scapula at an angle of 45 degrees with the first line. The needle is inserted perpendicular to the skin at the intersection of the second line with the deepest part of the right jugular fossa. The needle is directed caudodorsally and should not be angled toward either scapula. Because the vein is superficial, the syringe plunger should be retracted slightly as soon as the needle penetrates the skin. The needle is advanced until blood is aspirated into the syringe. Once a sufficient volume of blood has been collected, the needle is removed.

The technician should be aware that fewer complications and a lower risk for incidental injury to the animal are associated with a venipuncture of the more distal veins listed below.

Auricular Vein: The auricular vein is located near the lateral border of the pinna of the ear. It is easily visualized on the dorsal side of the ear and can be seen even more clearly by placing digital pressure at the base of the lateral surface of the ear. The ear is held and digital pressure (or rubber-band tourniquet) applied at the base of the ear to distend the vein. A needle with a syringe attached can be inserted into the vein while gently pulling back on the plunger. To prevent the collapse of the vein with aspiration, some people prefer to insert the needle and allow the blood to drip from the needle hub directly into the uncapped collection tube. For most pigs, a 20-gauge × 1-inch needle is appropriate. For large adult pigs, an 18- to 19-gauge × 1-inch needle may be used. Vacutainer collection needles and tubes may also apply too much suction, and as in the case of excess pressure on the syringe plunger, these tend to cause the vein to collapse. When an adequate blood sample has been collected, pressure is released from the base of the ear, the needle removed, and pressure applied to the insertion site. For repeated sampling, the placement of an IV catheter should be considered.

Peripheral Leg Veins: The cephalic vein on the front leg, saphenous vein on the hind leg, and branches of veins located on the lower limbs are accessible in small pigs and can be used for venous sampling. The veins can be visualized and are readily accessible in anesthetized pigs. To prevent the collapse of the vein, the needle is inserted, and blood is allowed to drip from the hub of the needle into the open collection tube (Figure 20-57). The collection of blood from the standing and restrained pig can be done by placing hand pressure or a tourniquet above the collection site to distend the vein. A 20-gauge × 1- to 1.5-inch needle is inserted at approximately a 45-degree angle to the skin in the direction of the body.

Coccygeal Vein: Though this site is not commonly used, it can be used for a venipuncture in adult pigs with intact (not docked) tails. A 20-gauge × 1-inch needle is inserted in the ventral midline of the tail perpendicular to the skin. Small blood samples can be obtained from this site.

Orbital Sinus (Medial Canthus of Eye): The orbital sinus is adjacent to the medial canthus of the eye and can be used for the collection of small volumes of blood. A 20- to 22-gauge × 1 inch-needle can be used for piglets. Sampling on larger pigs may require a 16- to 18-gauge × 1.5-inch needle. The needle is inserted into the medial canthus deep into the third eyelid (nictitating membrane) and advanced at a 45-degree angle toward the opposite jaw until bone is felt. The needle is then rotated between the fingers until blood enters the hub. The syringe is attached and blood collected with gentle aspiration. When the collection is complete, the needle is removed and digital pressure applied over the medial canthus with the head elevated. A microcapillary tube with the end broken to form a rough point can be used in lieu of a needle (Figure 20-58).

Free-catch Method

Urination may be encouraged by placing the horse in a freshly bedded stall. Sometimes standing the horse on a grassy area will encourage urination. Other suggestions include running water on cement and tickling the prepuce with a piece of straw. Some race and performance horses have been conditioned to urinate when whistled to. Recumbent neonates frequently will urinate when they are assisted to stand. The technician should be prepared by having a urine collection container within reach when helping a foal to rise or supporting it in a standing position.

Cleansing of the external genital area is not necessary when collecting urine for drug testing.

Urinary Catheterization

Supplies needed:

Free-Catch Method

Llamas and alpacas urinate and defecate on communal dung piles, so, if possible, the animal should be lead to a dung pile. Attaching a collection cup to the end of a broom or dowel facilitates collection without requiring personnel to be close enough to distract the animal. Both males and females urinate in a caudal direction while in a squatting position. Complete urination usually takes 30 to 60 seconds.

Urinary Catheterization of Llamas and Alpacas

In addition to the sigmoid shape of the penis, which makes passage of urinary catheters extremely difficult, male llamas and alpacas have a membranous flap at the ischial arch that prevents passage of urinary catheters into the bladder, so this procedure cannot be done in males.

Free Catch

Place the animal in a chute or stanchion and allow the animal to relax. Urination in cows may be encouraged by lightly stroking the vulva tip and the skin beneath the vulva with fingers or straw. The technician may also try repeated parting of the lips of the vulva to elicit urination. Steers and bulls may oblige with urination if the prepuce is massaged and splashed with warm water. Urine is collected into a container or, depending on the required analysis, a urine dipstick (litmus paper test strip) can be held directly in the stream of urine. This technique is often used to monitor urine pH and ketones.

Urethral to bladder catheterization in male cattle is virtually impossible because of the anatomy and therefore is not performed by the technician.

Cows can be catheterized with a small-diameter (0.5 cm) catheter using similar techniques as used for mares. With sterile gloved and lubricated hand, a finger is inserted into the suburethral diverticulum, and the catheter tip is guided over the diverticulum and into the external urethral orifice.

Free Catch

Both ewes and does tend to urinate immediately after rising following a period of recumbency. The technician should be prepared to collect a urine sample when the animal rises. This is the easiest approach and causes the least stress to the animal.

Urination may be induced in ewes by occluding the nostrils for up to 45 seconds while the animal is standing. This causes stress to the animal. The animal will indicate discomfort by struggling, and as the nostrils are released and the animal allowed to breathe again, she will urinate. Two people are required for this procedure to go smoothly. One occludes the nostrils and restrains the ewe while the other catches the urine.

Holding the nostrils of does seldom results in urination. Sometimes urination may be encouraged by placing the animal in a new stall or pen.

Male ruminants may respond to manual stimulation of the prepuce, but the technician should be prepared with a collection container and make every effort to collect a sample when the animal obliges.

Urinary Catheterization

Samples for California Mastitis Test

The California mastitis test (CMT) is commonly used to identify the presence of mastitis in cows (or does and ewes). The test involves the use of a white plastic test paddle with four cups labeled A to D and a reagent fluid (Figure 20-67).

The teats are cleaned and dried. It is not necessary to clean the entire udder. If washed completely, there is an increased risk of introducing contaminants from the udder into the teat orifice. Strip a small amount from one teat into one well of the paddle. Do the same for the remaining teats and note which teat was milked into each well (Figure 20-68). An equal volume of CMT reagent solution (one part milk to one part reagent solution) is added to each well, the paddle is gently moved to swirl the milk, and the resultant solution is graded based on gel formation.

Stomach tube—For adult cattle: medium to large diameter with internal diameter no less that 1.5 cm because smaller size is more likely to become obstructed with ingesta. For calves, sheep and goats, small and medium foal stomach tubes can be used

Water-based lubricant

Frick speculum (cattle)

PVC pipe “speculum,” block of wood with hole cut in center, roll of tape (sheep, goats)

Dose syringe

Sample collection container

Rumenocentesis

This method is seldom done because of the ease and safety of the stomach tube method described earlier.

The ventral abdomen caudal to the xiphoid process and left of the ventral midline is clipped and surgically prepared. The veterinarian inserts a needle with a syringe attached (14-gauge needle for cattle, 16- to 18-gauge needle for small ruminants) through the skin and into the rumen. Rumen fluid is aspirated into the syringe.

• Sterile gloves

• Ample collection tubes:

• EDTA (for cytology)

• Serum (for microbiology)

• Heparin (for pH, lactate, and glucose) or fluoride (for pH and glucose), depending on specific laboratory analyzer requirements

• Instrument of veterinarian's choice:

• Needle (minimum 3 inches) with large gauge

Or

• 14- to 16-gauge IV catheter

Or

• Sharp trocar and cannula

Or

• Teat cannula

Or

• Bitch catheter

• Two percent lidocaine 3 to 5 ml

• Six-milliliter syringe with 20- to 22-gauge × 1-inch to 1.5-inch needle

• Scalpel blade #15

• Sterile 35- to 60-ml luer tip syringe

• Three-way stopcock

• IV extension tubing

• Suture material, needle drivers, and scissors

• Ultrasound machine, if available and requested by veterinarian

Transtracheal Wash (Tracheal Wash, Trach Wash)

Transtracheal aspiration is the collection of fluid from the lower respiratory tract (bronchi, bronchioles, and alveoli) for a cytologic and microbiologic analysis and is performed to aid in the diagnosis of lower airway and lung disease. The fluid is a mixture of secretions and cellular material that has collected in the distal portion of the trachea.

Percutaneous Method

Restrain the horse appropriately and identify the location for the procedure. A horse may require mild sedation (heavy sedation should be avoided because it may suppress the cough reflex). Select a site on the midline of the neck directly over the trachea about one third of the way down the neck. Clip or shave an area approximately 4 inches × 4 inches, perform a sterile prep, and set up necessary materials.

Materials:

Inject approximately 1 to 2 ml of 2% lidocaine ID and SQ over the selected site on the trachea and apply a final prep.

Wearing sterile gloves, tie the canine urinary catheter in a loose half hitch knot. This will allow the catheter to stay completely on the sterile field and makes it easier to control the distal tip to prevent contamination by accidentally touching it on something before or during the insertion. Using the scalpel blade, cut off the distal tip (approximately 1 inch) of the canine catheter and place it back on the sterile field. This step makes aspirating thick mucus possible.

Make a stab incision with the scalpel blade through the skin between the tracheal rings. Grasp the 12-gauge Medicut with one hand. Stabilize the trachea with the other hand, palm side up, use fingers and thumb placed on each side of the trachea and hold firmly. Insert the distal tip of the needle with bevel side down through the incision and advance it into the tracheal lumen. If resistance is felt, redirect the tip of the needle between the tracheal ring spaces, then advance into trachea. A burst of air will exit the needle when it penetrates the lumen of the trachea. A 3-ml syringe comes attached to the needle with the Medicut. The syringe can be kept attached to the needle and plunger pulled back to aspirate for the presence of air.

Remove the Medicut stylet. This prevents possible laceration and loss of the canine catheter tubing. Place on sterile field in case it is needed again. Insert the canine urinary catheter until it reaches the thoracic inlet. Attach the 60-ml syringe and retract the plunger. Air is aspirated if the catheter is within the tracheal lumen. If no air is aspirated, reposition the catheter because it may be bent or occluded against the tracheal wall. Once air is aspirated, infuse 30 ml of NaCl and immediately try to aspirate a 5-ml or greater fluid sample (only a small portion of the fluid that is instilled will be retrieved). A 5-ml sample should be sufficient for microbiology and cytology. Continue aspirating while slowly withdrawing the canine catheter. Stop withdrawing when fluid for aspiration is evident and collect as much as possible with a catheter at that location. Withdraw again as needed to continue the collection of fluid (do not reinsert the catheter). If fluid is not obtained, remove the canine catheter, insert a second sterile canine catheter, and infuse another 20 ml of NaCl and try again (Figure 20-73). The total volume of saline infused (first and second attempt) should not exceed 50 ml. Some horses will cough while the saline is infused, which may have the positive effect of increasing the yield of mucopurulent material. Unfortunately, coughing may also cause the catheter to kink cranially, which may prevent the collection of any sample. With a sample syringe, remove the 5F catheter while keeping the Medicut in place (this helps to prevent an SQ infection). Remove the Medicut, apply pressure to the site, control any bleeding, and place a 4 inch × 4 inch gauze with antiseptic ointment over the incision for 24 hours. Use a sterile needle to cap the syringe containing the sample for transport to the laboratory.

An alternate to using a Medicut catheter is to use a 12- to 14-gauge needle and insert the canine catheter through the needle. The needle is removed after the canine catheter is inserted into the tracheal lumen. Using the needle alone rather than an insertion catheter (Medicut) makes laceration of the canine catheter more likely and increases the likelihood of SQ contamination of the insertion site because the contaminated canine catheter will be in direct contact with the tissue at the insertion site when removed.

Cellulitis or SQ abscessation at the tracheal puncture site is the most common complication. If swelling occurs, warm compresses are applied. Other complications include SQ emphysema around the trachea, pulmonary foreign body as a result of the presence of a catheter piece in the airway, acute dyspnea, tracheal laceration, minor SQ hemorrhage, and iatrogenic infection.

Endoscopic Method

The use of an endoscope is considered noninvasive and allows for the visual examination of the upper airways, trachea, carina, and primary and secondary bronchi, but the presence of the endoscope leads to the questionable accuracy of the microbial samples recovered with this technique. An endoscope is inserted through the nasal cavity to the tracheal lumen. Special tubing (polyethylene tubing or endoscopic microbiology aspiration catheter) is placed through the biopsy channel of the endoscope and fluid injected and aspirated to collect the sample. The use of these specialized sample collection items decreases the likelihood of contamination from the pharynx or the endoscope that would otherwise occur with the endoscopic approach. It is usually necessary to infuse saline, as described for the percutaneous approach, to aspirate a fluid sample from the trachea.

Bronchoalveolar Lavage (BAL)

Bronchoalveolar lavage is a procedure done to collect fluid samples from the lower airway. BAL provides fluid samples that are better for a cytologic assessment than samples obtained by transtracheal aspiration, but the fluid samples are representative of only a limited area of the lung and are subject to contamination from passing the tube through the nares. The BAL is done by the insertion of sterile tubing through the nares and down the trachea as far as possible. To prepare for the procedure, the technician should have sterile BAL tubing, a syringe containing 50 ml of 2% lidocaine, and three syringes each containing 60 ml of sterile saline (not bacteriostatic saline). The veterinarian may choose to use more or less saline, so the technician should check before the procedure to be sure of the desired amount of saline to have ready in syringes.

The horse should be sufficiently restrained and may require sedation. While the tube is passed into the trachea, 2% lidocaine is injected into the tube whenever the horse coughs. This acts as a local anesthetic to the bronchi and is done to decrease the cough reflex. It is not uncommon to use the entire 50 ml of lidocaine before the procedure is complete. Sterile saline (previously drawn up into three separate 60-ml syringes) is injected and aspirated (as is done with the transtracheal aspiration procedure) (Figure 20-74).

The procedure can also be done with the use of an endoscope, with sterile tubing passed through the biopsy channel of the scope so that saline can be injected and fluid samples aspirated. This method is the preferred method for a sample collection when bronchial and/or alveolar disease is suspected. The use of the endoscope allows for sampling from specific areas of the lung, but a limited sample is obtained rather than the pooled secretions that are obtained when a transtracheal aspiration is done.

If the veterinarian intends to perform both a percutaneous transtracheal aspiration and a BAL, the transtracheal aspiration should be performed first. This will allow the fluid samples to be obtained before any contamination that may be introduced with the passage of the BAL tubing (or endoscope). Often the cough reflex is intentionally suppressed for the BAL, whereas a cough may be desirable during the transtracheal aspiration procedure.

Teat Cannula or Female Canine Urinary Catheter (Bitch Catheter) Method

Using a blunt-tipped bovine teat cannula or stainless steel female canine urinary catheter reduces the risk of bowel penetration, therefore this method may be chosen over the needle method for animals with abdominal distention or bowel distention (as identified with a rectal examination by veterinarian). This method requires the use of a local anesthetic.

Shave an area approximately 2 inches square. Perform a sterile prep. Aspirate 2 ml of 2% lidocaine into a 3-ml syringe. Perform local anesthesia by infusing the skin and SQ tissue with approximately 1 ml of the lidocaine using a 25-gauge needle. Insert the needle into the center of the shaved area (avoid any obvious cutaneous vasculature), aspirate to check for blood, and then infuse the lidocaine. Remove the 25-gauge needle and place a 19-gauge × 1.5-inch needle directly into the center of the SQ bleb. Insert the needle completely to the hub and inject the remaining 1 ml of lidocaine while slowly removing the needle from the patient. This is to block the parietal peritoneum.

Complete the sterile prep with a final swab of Betadine solution (or alternate antiseptic).

Assemble the equipment:

(Figure 20-75)

Create a sterile field by opening a pair of sterile gloves and placing the teat cannula (or bitch catheter or needle), scalpel blade, and sterile gauze 4 inches × 4 inches on the field (Figure 20-76).

Put on sterile gloves.

Puncture the center of one gauze 4-inch × 4-inch sponge with the scalpel blade and put the teat cannula through the gauze. This will help prevent contamination of the sample with blood or dust.

Using the scalpel blade, make a stab incision through the skin. Hold the blade with about inch exposed. Using the back of the gloved hand, gently touch the horse's belly then insert the scalpel blade straight in the bleb and then pull it straight out. Avoid cutting the musculature. Slowly but firmly insert the teat cannula through the incision perpendicular to the musculature. The muscle will feel “gritty,” and a slight “pop” will be felt when the peritoneum is punctured. A decrease in resistance is felt when the abdomen has been entered. If firm resistance is felt at this point, it may indicate contact with an organ, and care must be taken if further manipulation on the cannula is required. Once the cannula is in, expect to wait while the horse takes a few breaths before fluid begins to flow from the cannula. If fluid does not immediately flow, the cannula can be gently “flicked” with a finger in an effort to encourage the flow, and the cannula can be moved around slightly, rotated, or redirected. If necessary, a syringe may be attached to the cannula and aspiration may be attempted (Figure 20-77).

The first few drops of fluid may contain contaminants, so these drops should not be included in the sample. Collect the sample by gravity flow into collection tubes. The EDTA tube should be collected first since most of the desired laboratory tests will require this sample. The tube should be filled as much as possible to ensure the correct ratio of EDTA to abdominal fluid. If less than 1 ml of abdominal fluid is collected, the results of the laboratory analysis may be inaccurate. A common practice is to remove the rubber stopper from the EDTA tube and shake the tube to remove a bit of the EDTA before collecting the fluid sample. This is useful when a small sample volume is obtained. When refractometry is performed, excessive EDTA in relation to sample size will falsely elevate the protein reading. EDTA samples are used for cytologic analysis, protein measurement, and PCV (if fluid appears bloody). A serum (plain, clot) tube should be collected for bacterial culture (as little as 1 drop of fluid is sufficient for this purpose). Some facilities have the capability to perform pH, gas, lactate, and glucose analyses. The technician should be familiar with the laboratory capabilities and be familiar with the appropriate anticoagulant requirements. For many analyzers that perform these additional tests, heparin is the choice of anticoagulant. Some facilities will require a sodium fluoride tube for glucose and lactate measurements.

When removing the cannula, the technician should be aware that some omentum may have attached to the cannula and can follow the cannula out when the cannula is pulled from the site. To prevent it from being exteriorized, care should be taken to use fingers to guard close to the site when removing the cannula. As a result of the incidental perforation of skin vessels, slight bleeding is common after the removal of the cannula. Manual pressure applied to the site usually stops the bleeding. If necessary, the veterinarian will suture or staple the site. The centesis site can be cleaned gently and antibiotic ointment applied daily for a couple of days.

18- to 22-gauge, 1.5-inch needle method

A local anesthetic is usually not required for this method, but that will vary depending on the horse.

The needle is held midway between the thumb and forefinger and inserted through the skin while avoiding superficial veins. The fingers are then moved slightly to grasp the hub of the needle for gradual advancement. The needle should be inserted in slight intervals, pausing before each interval of advancement to notice (by feel) if a scratching sensation is present. The scratching sensation indicates that bowel is rubbing over the tip of the needle. The fingers should be removed from the needle periodically to watch for rotary or “flicking” movement of the needle, which is also indicative of bowel contact. Periodic back and forth movement of the needle in time with respiration is normal. The needle is advanced slowly to the hub if no bowel is encountered or until fluid is obtained. If abdominal fluid is not seen in the needle hub, the needle can be repositioned and rotated, and a syringe may be used to try and aspirate a sample. If fluid is not obtained, 1 to 2 ml of air (in a sterile syringe) may be injected in an effort to dislodge any material that might be occluding the needle. Another option to encourage the flow of fluid is to insert a second or third needle a few centimeters from the first (while leaving the first needle in place) to release the negative pressure in the abdomen.

18-Gauge, 3.5-Inch Spinal Needle Method

This long needle may be required in very large horses, draft horses, and obese individuals. Some clinicians report that this long needle is also useful in Arabian horses to facilitate penetration beyond the abdominal wall and subperitoneal fat layer.

A local anesthetic is not usually necessary for this method, but that will vary depending on the horse.

With the stylet in place, the needle is inserted to a depth of approximately ¼ inch. The stylet is removed before further advancement and the procedure continued as is done with a standard needle.

Atlanto-Occipital Site (AO Tap)

This site is located on the dorsal midline just caudal to the poll. Collecting spinal fluid from this site requires a general anesthetic. With the animal anesthetized, it is placed in lateral recumbency. The area is clipped, shaved, and a complete sterile prep performed. When all preparations have been made, the nose is directed down toward the front feet to flex the head. The head should be at a right angle to the neck. The veterinarian will insert an 18-gauge × 3-inch spinal needle into the atlanto-occipital space (about 5 to 7 cm depth) and once in, will remove the trocar (stylet) and place it on a sterile field. A sterile syringe is attached to the needle and a sample gently aspirated. Alternately the fluid may be collected directly into a tube by free flow. If the fluid is blood tinged, aspirate a few milliliters and then attach a new syringe. The technician should be prepared with additional syringes in case multiple samples can be obtained. The trocar (still sterile) is replaced in the needle, and the needle is withdrawn. Following the removal of the needle, any blood present can be cleaned from the site, and a Betadine-soaked gauze sponge can be placed on the site.

The AO tap in neonates is done with a 20-gauge × 1.5-inch needle directed at the mandible. Fluid should drip from the needle hub. Normally, 3 to 6 ml of fluid is obtained.

Lumbosacral Site (LS Tap)

This site can be located by making an imaginary line across from the caudal edge of each tuber coxae and another line on the dorsal midline. A slight depression can be palpated using firm pressure at the intersection of these imaginary lines, just caudal to the sixth lumbar spinal process (L6). A large area is clipped, shaved, and a sterile prep performed. Sedation is required, and the use of a twitch for restraint is indicated because the animal must remain very still for the procedure. The horse should be placed in stocks. A local anesthetic is injected into the skin and SQ. The patient should be standing as squarely as possible (the best that its condition will allow) because an asymmetric stance makes collection more difficult (Figure 20-79, A-F). If sufficient personnel are available, it is helpful to have someone standing behind the horse to comment to the veterinarian as to the placement of the needle so that small errors in the direction of insertion can be noted and corrected. The veterinarian will insert a 6-inch (15 cm) × 18-gauge spinal needle perpendicular to the midline (about 11 to 15 cm, or 4.5 to 6 inches). For some draft horses and warmblooded animals, longer needles (up to 8 inches) may be needed. When the needle reaches the subarachnoid space, the patient may respond with movement. The trocar is removed from the needle and placed on a sterile field. The technician will place a sterile syringe in the veterinarian's still sterile, gloved hands. The initial sample may be contaminated with blood, so a second or third syringe may be collected if sufficient fluid is aspirated. The veterinarian may instruct assistants to occlude both jugular veins in an attempt to increase the intracranial pressure. The trocar (still sterile) is replaced into the needle and the needle removed.

The lumbosacral CSF tap in neonates can be done with the foal standing, in sternal recumbency, or in lateral recumbency using a 3-inch × 20-gauge spinal needle.

Complications that may result from CSF taps include trauma to the spinal cord during needle placement, herniation of the cerebellum (can occur with high intracranial pressure or from aggressive aspiration), and infection of the meninges. These can result in the death of the patient. The chances of these complications occurring are minimized by having the patient sufficiently restrained to remain still and by following strict sterile technique throughout the procedure.

Camelids

A CSF fluid collection from llamas and alpacas follows the same procedure guidelines as described for horses.

The atlanto-occipital site is located midline as it intersects the wings of the atlas. In adults, a 20-gauge × 2.5-inch spinal needle is used with the subarachnoid space usually reached at a depth of 4 cm.

The lumbosacral site is midline about 2 cm caudal to the dorsal spinal process of the seventh lumbar vertebra. The landmarks used to locate the site are the tuber sacrale of the pelvis and dorsal spinal process of the last lumbar vertebra. The site is cranial to the tuber sacrale. An 18- or 19-gauge × 3.5-inch spinal needle is appropriate for most adult llamas and alpacas.

Equine

With most horses, minimal restraint (halter and lead rope) can be employed to successfully administer medication using this route. If the patient is uncooperative, aggressive, or sensitive to needles, placing the animal in the stocks or using a second person and a twitch may make the procedure easier and reduce the likelihood of injury to personnel and complications with misdirected puncture.

Equine IV Catheterization

For repeated IV drug injections or when large volumes of IV fluids are required, an indwelling catheter should be placed in the jugular vein. If either jugular vein is not accessible as a result of thrombosis or trauma or if the horse pesters the jugular catheter, it may be necessary to catheterize the cephalic vein or the lateral thoracic vein. There are many different options for catheters, and the choice should be made based on the length of time the catheter will be in place and the number of ports that may be necessary. For adult equine patients, when using OTN catheters, 14 gauge × 5.25 inches is sufficient, but for neonates, small ponies, or miniature horses, 16 gauge × 3.25 inches may be preferred. Miniature horse neonates may require a smaller catheter, and an 18- to 20-gauge × 3-inch catheter may be used, with the realization that the size of the catheter will dictate the fluid administration rates that can be achieved.

The site chosen is shaved and surgically scrubbed to remove all debris. A final wipe with Betadine solution over the area is then added and left on to dry. A “bleb” of lidocaine (ID administration using a 25-gauge needle and approximately 2 ml of lidocaine) should be administered over the intended injection site, including above the site, to desensitize an area for suturing the catheter in place. A sterile field is created by opening up a package of sterile gloves and aseptically laying the desired catheter on the gloves. All necessary items need to be either placed on the sterile field or readily accessible nearby.

Wearing sterile gloves, the technician grasps the catheter with the dominant hand and uses the other gloved hand to apply digital pressure to the jugular furrow. The catheter is inserted through the skin (into the “bleb”) at approximately a 45-degree angle and should be inserted toward the heart, with the direction of blood flow. Once the lumen of the vein has been accessed (identified by the “flash” of blood at the hub of the catheter), the catheter is aligned more perpendicular to the vein and advanced 1 cm more. If the catheter is still in the vein (as indicated by flash back blood coming from catheter), the hand applying the digital pressure releases and then grasps the top of the stylet portion of the catheter. The catheter is then slid all the way into the vessel, removing the stylet at the same time. Recheck to make sure that the vein is still catheterized by applying digital pressure yet again (below the tip of the catheter) and watch for blood to drip from the hub. Once this has been identified, attach a PRN or T-port and suture the catheter into place (Figure 20-80, A-E).

If the carotid artery is catheterized, bright red blood will forcibly pulse out of the catheter. If this should occur, immediately remove the catheter and apply digital pressure over the sight for a minimum of 5 minutes to prevent the formation of a hematoma. Neck wraps or stents can be placed over catheters to stabilize them and to prevent the patient from rubbing them out. A common practice is to apply a small amount of antibacterial ointment before placing a wrap over the catheter. If the catheter remains in place long term, the ointment used can be alternated (e.g., Nolvasan, Betadine, triple antibiotic) in an attempt to reduce the chances of a resistant staphylococcus infection taking hold. Foals and adult horses that spend a considerable amount of time in recumbency should have wraps placed over the catheter to protect the site from bedding, urine, and manure. When placing IV catheters in recumbent neonatal foals, it is helpful to place a rolled-up towel under the neck to enhance the view of the jugular vein and stretch the skin. Making a small nick in the skin at the insertion site with a needle or blade will facilitate the insertion of the catheter. It is also helpful to have an assistant stretch the skin while applying digital pressure and thus maintaining distention while the catheter is advanced.

When administering fluids or medications into an IV catheter, the technician should always first clean the injection port with isopropyl alcohol to prevent bacterial contamination.

If the technician wishes to use a guidewire-type catheter, the same steps for preparing and placing the catheter are used, but the guidewire-type catheters entail a few extra steps (refer to Seldinger guidewire technique described for small animals or read directions on individual packages). The most important thing to remember when placing these types of catheters is to NEVER let go of the guidewire until the catheter is successfully placed and the guidewire is fully removed.

Bovine

Bovine IV Catheter

When administering large volumes of fluids or repeated IV administrations, a catheter should be placed in the jugular vein to perform these tasks successfully and without the potential for causing problems or damage to the vessels. To place an IV catheter, the same restraint techniques should be employed as outlined previously. The site that is chosen needs to be clipped and surgically prepared, and an antiseptic, such as Betadine solution, is wiped onto the site and left to dry. Because of the skin thickness of cattle, a cutdown using a #15 scalpel blade or a puncture with the same size needle as the catheter will allow for easier insertion of the catheter. If a blade is used to cut the skin before the catheter insertion, a local anesthetic bleb should be placed. A 12- to 16-gauge × 5¼-inch catheter is used on adult cattle, but a smaller size, such as 15- or 18-gauge × 3¼-inch, can be used for calves (Figure 20-83).

The cephalic vein can also be used to place a catheter if the jugular veins are inaccessible. The caudal auricular vein (ear vein) may be used for small-gauge catheters, but these are hard to maintain because it is difficult to stabilize the catheter and prevent the animal from rubbing the catheter (Box 20-6).

Camelid

Although the jugular vein is the most common site for IV injections for llamas and alpacas, it is not as easily accessed as the other large animal species. Injections can be given either high on the neck or low on the neck, but each has potential complications (see previous section of chapter). The high neck position is identified by landmarks as described earlier in the chapter for venous blood sampling. Using the same technique employed with all IV injections, the needle should be detached from the syringe and inserted to make sure that the vein has been punctured and not the artery. Once identified that you have successfully accessed the vein, the syringe is attached, aspirate back, and then gently administer the medication.

The same process is followed for the administration of the low neck site. The landmarks for jugular access on the lower neck are the space between the fifth and sixth cervical vertebrae. Locate the enlarged transverse process of the sixth cervical vertebra, and the jugular vein lies just medially to it. The advantage to choosing the low neck site is the thinner skin, better visualization, and fewer problems with the animal moving or flinching away from the needle and technician.

Ovine and Caprine

As with the other large animal species listed above, the most common route for IV administration in the goat or sheep is the jugular vein. Proper restraint is essential to administer the medication efficiently and effectively. It is possible to perform the procedure with one person, but having additional help will make the procedure smoother. For both sheep and goats, backing the animal up against a wall, the technician then straddles the patient over the shoulders facing the head and gently grasps under the mandible and lifts the head up and away from the vein to be punctured. For sheep, the wool should be parted for the visualization of the skin. Sheep can be set up on their rump and veins accessed as described for blood collection. The area is to be wiped down with isopropyl alcohol, digital pressure applied in the jugular furrow to distend the vein, and then a needle is inserted into the vessel (20-gauge × 1 inch for adults; 22-gauge × 1 inch for kids and lambs). Once the vein has been punctured and it is established that it is venous blood, the syringe with medication can be attached, aspirate back for confirmation, then slowly inject substance. After all medication is delivered, remove the needle, and apply digital pressure for a couple of minutes to prevent hematoma formation.

Porcine

IV administration to pigs is accomplished using the auricular veins, located on the dorsal aspect of the pinna. There are three veins, and the most common one for injection is the lateral vein. The pig should be restrained by using a snare or chute, or if small enough, an assistant can hold the animal against their body. Pigs have very sharp teeth and strong jaws. Without sufficient restraint, a bite is a serous possibility. The ear should be cleaned with alcohol-soaked gauze and the base of the ear occluded with digital pressure. A small-gauge needle is inserted into the vessel at a very shallow angle. The needle should be attached to the syringe at the time of insertion because of the very fragile nature of these vessels. Aspirate very gently to confirm venous placement, release occlusion, and then inject the substance with steady pressure. Remove needle and syringe and apply digital pressure. The cephalic vein can also be used, but in small piglets, the jugular vein should be used because of the small diameter and difficultly accessing the cephalic and ear veins at that age.

Equine IM Administration

There are several locations that can be used on the equine patient for the IM administration of therapeutics. These sites include the lateral cervical (neck), semimembranosus and semitendinosus, and pectoral and gluteal muscles on both the left and right side of the animal. When choosing the location for IM injections, the technician should consider: the volume to be delivered, the viscosity of the solution, the potential injury to personnel, and the potential for complications in the muscle chosen. Restraint must be employed whenever giving an injection, with a halter and lead rope as the minimum. Placing a horse in stocks will greatly reduce the likelihood of the animal moving once the needle is introduced and will also prevent injury to personnel, although it can be detrimental to a patient that has a reaction to the medication given if they get caught in the stocks. Individual horses respond differently to needles. Some respond only slightly or not obviously at all. Others can respond violently. A method for desensitizing the injection site just before a needle insertion is to rub the site very firmly and rapidly back and forth 100 times using an alcohol-soaked cotton swab or piece of gauze and then immediately insert the needle. Some people like to tap the horse firmly with the edge of the fist just before inserting a needle into the muscle. This is also thought to desensitize the area before the insertion of the needle. Other people are convinced that tapping in this manner before a needle insertion simply lets the horse know what is about to happen. The technician should use whatever method proves to be a good approach for him or her.

Bovine IM Administration

Because most cattle are eventually consumed, IM injections are highly discouraged to prevent muscle damage. If it is absolutely necessary to give an IM injection, the muscles of the neck should be used. The animal should be restrained in a head gate or squeeze chute. The technician should approach the animal from the forequarters, stay close, and leaning in to the animal, place a halter on the head and tie it securely to the side. The borders are the same as described for equine patients (the spine, nuchal ligament, and the scapula). The needle should be inserted with a quick thrust into the muscle. Following beef quality assurance guidelines, the needle must be clean and sharp, the injection smooth so as not to cause too much muscle damage, and no more than 10 ml of substance is to be administered at any one time.

The semitendinosus, semimembranosus, and gluteal muscles should not be used for IM injections in cattle.

Ovine and Caprine IM Administration

Sheep and goats have small muscle masses. The most common muscle groups used for IM injections are the semitendinosus and semimembranosus muscles (Figure 20-89). The technician must avoid the sciatic nerve, which runs down the caudomedial aspect of the hind legs. The neck, gluteals, and triceps can be used, but only for very small volumes (Figure 20-90). IM injections into the neck may cause significant soreness, and the animal may be reluctant to raise its head. This can be particularly problematic in kids and lambs because they may become too sore to nurse. Once the muscle to be used is identified, the standard procedure described for large animals is followed. For adult sheep and goats, an 18- to 20-gauge × 1-inch needle should be used. A 20- to 22-gauge × 1-inch needle is appropriate for lambs and kids.

Porcine IM Administration

IM injections in pigs can prove complicated because of the thickness of the skin, the tendency to store a thick layer of SQ body fat, the difficulty in restraining them, and potential damage to muscle (meat). Generally the cervical neck muscles are used just caudal and ventral to the ear. In adults, a maximum volume of 5 to 10 ml per site is recommended. Piglets can receive 1 to 2 ml per site. For adults, a long needle (at least 1.5 inches) should be used to avoid the fat because injecting into the fat will delay drug absorption. Depending on the size of the animal, the needle gauge can be anywhere from 20 gauge for piglets to 16 gauge for larger stock. Drug residues in various muscles will reduce the market value of the animal. The gluteal, semimembranosus, and semitendinosus muscles can be used, but not in animals destined to be used for meat. Following the same meat quality assurance guidelines as identified for cattle, the technician will grasp the skin in this area and pull cranially. With a firm motion, insert the needle at a perpendicular angle to the skin. Once the needle is in, attach the syringe, aspirate back, and inject the medication.

Camelid IM Administration

IM injection sites for llamas and alpacas are generally the same as in other large animal species. They do not have a large muscle mass in any one place, so SQ is the preferred route for the administration of large volumes or potentially irritating substances. The neck should not be used because of the potential for causing soreness in the area. The semimembranosus and semitendinosus muscles are good choices for IM injections in these animals (Figure 20-91). For adults an 18- to 20-gauge × 1-inch needle is appropriate. Twenty-gauge to 22-gauge × 1-inch needles are recommended for crias.

Equine Administration

In the equine patient, intraperitoneal (IP) administration of fluids and medication is usually accomplished through an abdominal lavage system. The drain system is inserted surgically by the veterinarian either during abdominal surgery or in a standing position when peritonitis is suspected and general anesthesia is not necessary. The technician will not be involved with the surgical procedure, but will be responsible for the care and maintenance afterward. To lavage the abdomen, latex tubing is attached to the desired fluids and connected to the drain system using a 5 in 1 connector. The desired amount of fluid (routinely 10 L for an adult equine patient, adjusted for smaller patients) is administered along with any medication (heparin, antibiotics, etc.), the latex tubing is clamped off, and the patient walked. This is done to attempt to distribute the fluid and medication throughout the abdominal cavity, washing internal organs, and breaking up any adhesions. The latex tubing is then unclamped, and the fluid is allowed to drain out of the abdomen back into the original fluid bag (Figure 20-99, A-C). Ideally the amount returned is equal to the amount originally administered. This process can be repeated several times per day. When handling the drain system, gloves should be worn, and the technician should pay careful attention to keeping the system clean and not introducing any contaminants during the administration.

Bovine Administration

IP injections may be indicated if IV administration is not possible and for treatment of peritonitis. If an IP injection is administered in cattle, the site selected is usually in the paralumbar fossa. Care must be taken when on the left side to prevent puncturing the rumen and on the right side to prevent puncturing the intestine or dilated or displaced internal organs.

Caprine and Ovine Administration

IP injections are usually reserved for neonatal kids and lambs with umbilical infections or hypoglycemia. The neonate is lifted by its front legs, and using a 20-gauge needle attached to the syringe filled with medication, the technician inserts it just to the left of the umbilicus up to a depth of 1 cm. Aspirate back to verify that you have not hit a vessel or internal organ. Once placement is confirmed, inject medication into the peritoneal cavity. Remove the needle and syringe.

Porcine Administration

In neonatal pigs, fluids are generally administered IP because of the impracticality of placing IV catheters and administering fluids using that route. Fluids should be body temperature, nonirritating, and isotonic. The site used needs to be prepared using aseptic technique to ensure that contaminants are not introduced. The piglet is held up by the rear legs, and an 18-gauge × ¾- to 1-inch needle is inserted paramedially between the midline and flank. The needle is stabilized to prevent damage to internal organs. To do this procedure in a mature, standing pig, follow the preparation guidelines and use a 16- to 18-gauge × 3-inch needle inserted through the paralumbar fossa.

Complications from IP administration include peritonitis, abscess, and injury to internal organs.

Syringes

Many commercially available products come in a paste form in premeasured dosing syringes, which make accurate dosing and administration easy. Other oral medications come in tablet form that need to be both crushed and mixed with water or simply dissolved in water over time. The easiest way to do this is by placing the tablets in a 60-ml catheter tip syringe, adding water, and letting the tablets dissolve. If the medication is unpalatable to the patient, molasses, Karo, or maple syrup or thin applesauce can be added to the mixture. To use the syringe method, proper restraint of the head should be used, with the free arm cradling the head, reaching around so that the hand is up over the muzzle. The technician then inserts the syringe into the mouth at the commissure of the lips near the interdental space, between the cheeks and teeth, and advances it as far back into the mouth as possible (Figure 20-100). With firm pressure, the medication is given, and the syringe can be withdrawn. Lifting the head slightly may encourage the animal to swallow instead of spit the medication out. The medication should not be injected too rapidly because it may be lost from the other side of the mouth or may be aspirated. The technician should be conscious of the probability of the patient spitting the material out and getting it on the skin or mucous membranes of the personnel. Always wash hands and skin that has been in contact with the medication.

Drench (Dose Syringe)

Liquid medication or small volumes of fluid can be administered using a dose syringe. For calves, sheep, and goats, a catheter tip syringe can be used, whereas a large dose syringe is appropriate for adult cattle. Drenching in sheep and goats should be limited to small volumes of fluid (no greater than 30 ml). Holding the animal with the nose slightly elevated and pulled toward the handler, the tip of the dose syringe is inserted into the interdental space, and the fluid is slowly dribbled onto the tongue (Figure 20-101).

Balling Guns (Pilling)

For cattle, sheep, and goats, balling guns are commonly used to accomplish oral tablet administration. There are several sizes available, and the appropriate size should be picked in accordance with the size of your patient. Using a large balling gun on a small calf, goat, or sheep can result in a splitting of the soft palate and rupture of the pharynx. The balling gun should have a smooth end, preferably made of rubber to limit the trauma that can be caused to the back of the mouth, including the soft palate, pharynx, and esophagus. It should be inspected before each use to make sure that no sharp edges have formed. All food should be removed from the patient's mouth before dosing. For cattle, placing the patient in a head gate will help restrain the animal for the procedure. The technician should stand next to the animal's head facing the same direction as the animal. The arm nearest the animal reaches over and grasps the mouth at the interdental space and pressing on the hard palate, opens the animal's mouth. Alternately, placing a finger in one nostril and the thumb in the other and then pulling the nose dorsally will encourage cattle to open their mouth making insertion of the gun easier. The balling gun is inserted into the mouth and gently worked back to the pharynx. Once the thumb rings of the gun are at the commissure of the lips, the plunger is depressed (Figure 20-102). The animal's head should be kept down to prevent the loss of the medication. Cattle will lick their nostrils once they swallow the pill.

For sheep and goats, back them up to a wall and straddle the shoulders as stated before. Insert a hand into the interdental space at the commissure of the lips, open the mouth, and insert the balling gun to the back of the throat before depressing the plunger. If the balling gun is not inserted far enough, the medication may be chewed by the animal and spit out. If the balling gun is inserted too far, there is a risk for serious damage to the pharynx and larynx. To decrease the chance of aspiration of the medication, the head of the animal should not be overly elevated and the neck should not be overextended.

Nasogastric and Orogastric Intubation

When large volumes of an oral medication need to be given (mineral oil, fluids, bismuth), oral fluid therapy or enteral feedings for extended periods of time are required, passage of an OGT (cattle, sheep, goats, pigs, and camelids) or an NGT (horses, cattle, adult sheep and goats, neonatal camelids) should be used.

Nasogastric intubation is a commonly used procedure in equine patients. Cattle, sheep, goats, and camelids have small nasal passages and although nasogastric intubation (with small diameter tubes) can be done (Figure 20-103), the usual method for these species is to place an OGT.

Equine Epidural Administration

The horse should be restrained in stocks and a twitch applied. Some horses will require the administration of a sedative. The technician will clip, shave, and aseptically prepare a 3-inch square (approximately) area over the first and second coccygeal vertebrae. This site can be determined by lifting the tail up and down with one hand while feeling for the vertebral space with the other hand (Figure 20-111). This area is usually close to where the coarse tail hairs originate. An SQ bleb of local anesthetic is placed. The technician should attempt to have the horse stand still and squarely upon its legs. If the animal is not standing squarely, there will be an uneven distribution of the drug because more will run into one side of the epidural space.

The technician should prepare the supplies necessary, including sterile gloves, local anesthetic, a sterile 12-ml syringe, 19-gauge × 1.5-inch needle (3.5 inches for very large horses), or an 18-gauge epidural catheter with stylet.

Directing the needle slightly cranially and ventrally into the epidural space at an approximately 45-degree angle to the rump, an 18- to 19-gauge × 1.5-inch needle is inserted about 1 inch in adult horses. When the needle enters the epidural space, a slight “pop” is felt, and there is a loss of resistance to the passage of the needle. The drug is injected, and the needle may be left in place to facilitate an additional injection. An epidural catheter may be placed and secured to the skin to facilitate repeated drug administration. When the needle or catheter is removed, antibiotic ointment is applied to the site.

The technician should be aware that hind leg instability may occur following epidural anesthesia. Lidocaine, Carbocaine, xylazine, and morphine are commonly administered as epidurals.

Bovine Epidural Administration

The technician must first ensure that the animal is sufficiently restrained. To locate the site, the tail is moved up and down using one hand while the other hand feels the top of the vertebrae to find the first moveable joint (S1 and S2). Clip, shave, and aseptically prep a 3 inch × 3 inch area (approximately). An 18-gauge × 1.5 to 3-inch needle is inserted perpendicular to the spine and into the vertebral space between S1 and S2 (Figure 20-112). A “pop” is felt when the space is entered. The epidural space is a relative vacuum compared with atmospheric conditions, and the medication will be sucked into the space. (A drop from the syringe into the hub of the needle should quickly be drawn into the needle.) Three milliliters of 2% lidocaine is commonly used for bovine epidurals.

Camelid Epidural Administration

The site is clipped and surgically prepared. Owners may object to the clipping of the fiber, so efforts should be made to shave only a small site and secure the surrounding fibers away from the site with adhesive tape. The tail is moved up and down to locate the intervertebral space between S5 and C1. In most llamas and alpacas, this will be the first moveable joint because the five sacral vertebrae are usually fused. A 20-gauge × 1.5-inch needle is inserted. The veterinarian will determine a successful insertion as described for other large animals.

Ovine and Caprine Epidural Administration

Following the procedure guidelines for cattle, an 18- to 21-gauge × 1 to 1.5-inch needle is inserted at a 45-degree angle. Sheep and goats are very sensitive to local anesthetics.

Porcine Epidural Administration

The site for epidural injections in pigs varies from the site described earlier for other large animal species. The lumbosacral junction (between L6 and S1) is accessible for porcine epidurals. This is considered a cranial epidural, whereas the common sites described for other species are caudal epidurals. To locate the site, an imaginary line is drawn vertically up from the patella to the back and a dorsal midline site clipped and surgically prepared. An 18- to 20-gauge spinal needle may be used, with the length determined by the size of the animal.

Rectal Medications

To deliver medication per rectum, the appropriate size tube should be selected based on the size of the patient. A Harris enema tube (24F) or fenestrated tube (multiple holes along the distal end) are appropriate for adult animals, foals, and calves, whereas smaller-diameter soft rubber tubes can be used for lambs, kids, and crias. The fenestrated tube may provide better distribution of the medication, but the fenestrations on some tubes may be rough and may cause irritation to the rectal mucosa. The technician should always check the tube for any rough edges before using and avoid using anything that is not smooth. The distal end of the tube is lubricated with a water-soluble solution (such as KY jelly), and the tube is inserted from 1 to 12 inches into the rectum. This distance is determined by the size of the patient. Appropriate restraint is used, tailored to the individual species and age of the animal. For standing animals, the technician should take precautions to stand to the side of the animal so as not to get kicked. Medications are dissolved in a small amount of water (or at the veterinarian's request, in another solution, such as DMSO) and injected gently via a catheter tip syringe into the tube followed by a small “chaser” of water (or air) to ensure that all of the medication is administered and none remains in the tube. The tube is then gently removed. It may be necessary to remove feces from the rectum before the administration of medications. The technician must clarify this with the veterinarian in advance because it may or may not be necessary, and there is an increased risk for injury to the animal when a hand is inserted into the rectum. Rectal tears can be fatal. If instructed to do so, the technician must have fingernails clipped short and wear no rings or watches. With a well-lubricated rectal sleeve, the technician will gently insert the hand a short distance into the rectum and gently remove obvious feces present before the insertion of the tube.

The veterinarian may administer 2% lidocaine per rectum to facilitate performance of a rectal examination by reducing the straining of the patient. A 60-ml syringe containing lidocaine is attached to rubber tubing or IV extension tubing that is inserted into the rectum and the drug injected. Sedation may also be given via the epidural route for patients that are straining to prevent potential problems, such as rectal tears, during the examination.

Enema Administration

Enemas are administered to constipated animals to encourage defecation. They can be administered to animals of any age or species. The tube used, volume, and composition of fluid administered will vary with the size and condition of the animal. Fluids should be nonirritating and warmed to room temperature, but not above body temperature.