[Murat KUTAY]

Intramuscular Injections

the dog’s head is to your right

Several sites can be used for IM injections in dogs, including the triceps muscle, quadriceps group, dorsal lumbar muscles and the semimembranous and semi- tendinosis muscles. Needle sizes used for IM injections range from 25 to 20 gauge. Muscle is dense tissue and can only accommodate small volumes of fluid (a few ml).

The injection site is swabbed with alcohol to wet down the hair coat to assure IM rather than SQ needle placement. The amount of time the alcohol contacts the skin is inadequate for kill of surface bacteria.

the dog’s head to to your right

The triceps muscle belly located caudal to the humerus is one IM injection site. The left thumb is placed on the humerus, isolating the muscle belly in the left hand. The needle is placed in the muscle belly. The plunger is withdrawn to create negative pressure.

If blood is aspirated, the needle should be removed as this indicates needle placement in a blood vessel. Reinsert the needle at a different site. Some pharmacologic substances can cause an anaphylactic reaction if given IV. If no blood is aspirated, the injection is given, the needle is withdrawn and the muscle belly is massaged to facilitate dispersion of the injected material.

the dog’s head to to your right

The quadriceps muscle is located anterior to the femur. The left thumb is on the femur. The needle is inserted at a right angle to the muscle belly.

the dog’s head to to your right

When administering an injection into the semimembranous/semitendinosis muscle group, the tip of the needle (white arrow) should be directed toward the caudal aspect of the limb so if the patient moves, the needle will not advance toward the sciatic nerve. Notice the left hand is being used to isolate the muscle group caudel to the femur.

the dog’s head to to your right

The dorsal lumbar muscles on either side of the midline can be used for IM injections. The thumb of the left hand is on the transverse processes of the lumbar vertebrae.

[Murat KUTAY]

Subcutaneous Administration (Hypodermoclysis)
Dogs and cats have an extensive potential subcutaneous space that can be used for the administration of drugs or fluids. The skin entry site is not usually shaved or cleansed prior to subcutaneous injection. The hair may be wetted with alcohol to better define the skin surface so the injection is not given in the haircoat.

Needle size for subcutaneus injection ranges from 18 to 25 gauge. Use a needle size appropriate to the patient size and viscosity and volume of the fluid being injected. Subcutaneous injections can be given anywhere over the dorsal cervical, thoracic or lumbar regions. If giving large volumes of fluid, deposit 10 to 20 ml/kg at each site. In states of extreme dehydration, blood flow to and subsequently absorption from the subcutaneous site is diminished. Do not use this route in severely dehydrated animals.

To enter the subcutaneous space, pick up a fold of skin and insert the needle under the skin, parallel to the long axis of the skin fold. If the needle is inserted across the fold of skin, the needle may penetrate both folds of skin and the injection will be deposited on the haircoat (or sprayed at the person holding the animal).

After inserting the needle into the subcutaneous space, aspirate by pulling back the plunger of the syringe. If you aspirate air, the needle has penetrated both skin folds and needs to be repositioned. Once the needle is in the subcutaneous space, you can release the skin fold or you can keep the skin tented as you administer the fluid. You can deposit the fluid in the subcutaneous space as rapidly as it can be ejected from the syringe.

When administering a large volume of subcutaneous fluids, use a flexible delivery system instead of a needle rigidly attached to a syringe. Pictured is a hypodermic needle attached to a fluid extension set and then to the syringe containing the fluids to be administered.

A butterfly catheter can also be used

If more than one syringeful of fluid is to be administered, use different needles to aspirate fluids from the sterile fluid container and to administer fluids to the patient to prevent contamination of the fluid container.

[Murat KUTAY]

Placement (yerleşim )of a Jugular Catheter

There are several types of jugular catheters available. Most catheters are threaded through a needle, therefore, the hole made in the vessel wall by the needle is larger than the diameter of the catheter so for a short time after placement, blood tends to leak back around the catheter. Catheters demonstrated in the slide are Intrafusor (a) which has its own extension set and a separate needle guard, the Venocath (b) that has an attached needle guard and the Intracath (c) that has a detached needle guard.

The venipuncture models located in the psychomotor skills lab on the second floor of McCoy are available to you, any time that you want to practice. See Lynn Duncan to obtain a code to access the room.


A jugular catheter can be placed with the patient in sternal recumbency, with the neck extended upwards.

or in lateral recumbency. The catheter insertion site should be widely shaved of hair and aseptically prepared using an antiseptic solution such as BetadineTM or Chlorhexadine.
Digital pressure is applied at the thoracic inlet to cause the jugular vein to distend with blood. This is usually done by the person placing the catheter if the dog is in sternal recumbancy, but is done by the holder if the dog is in lateral recumbancy. The jugular vein is clearly visable in this dog.

The following series of slides will show placement of a jugular catheter using an IntracathTM by Deseret.

Insert the catheter through the skin and through the wall of the vein in one motion, if possible. Make the puncture through skin and vein wall rapidly and forcefully. If you are too delicate and slow when inserting the needle, you will push the vein away from the needle instead of puncturing it. When the vein is punctured, blood will “flash” up the catheter along the blue marker line.

Once it is clear that the vein has been punctured, the catheter is threaded (pushed) through the needle by pinching through the plastic behind the catheter which pushes the catheter forward. The green plastic end of the catheter (a) is securely forced into the metal hub of the needle (b). The white plastic cap that still covers the needle hub in this slide will be removed.


The Intrafusor is packaged with a rigid plastic cylinder (a) covering the catheter. The white y-shaped part of the catheter (b) is pushed forward until it locks into the yellow hub of the needle (c). This results in the catheter being pushed through the needle. The catheter is the gray “stripe” between c and b. The white plastic cylinder (a) is pulled off and discarded. Other parts of this catheter are the extension tubing (d) and the needle guard (e).

When using an Intrafusor style catheter, wait for blood to “flash” back into the extension set (a), to assure a good venipuncture, before pushing the catheter through the needle into the vein.

The needle guard is folded over the needle like a clamshell. Make sure that the needle is in the groove of the guard so that the catheter is not pinched by the needle guard when it is closed. Make sure that the tip of the needle (a) is within the needle guard so the needle does not lacerate the catheter.

The wire stylet (a) is removed. Make sure that the hub of the catheter (b) is firmly locked into the hub of the needle (c) when the stylet is removed or you may remove the entire catheter along with the stylet.

An injection cap (a) is attached to the catheter and the catheter is flushed with heparinized saline to assure the catheter is properly placed before it is bandaged. If the catheter is properly placed, you should be able to aspirate blood from the catheter and solutions can be injected without resistance.

The external portion of the catheter should be positioned just behind the ear. Notice the small amount of blood leaking around the catheter at the puncture site (arrow). Bleeding around the catheter occurs as the catheter is slightly smaller than the hole made in the vein wall by the catheter needle.

Although the needle guard has two holes for suturing (the second hole is under the tape), a butterfly of tape offers better stability in securing the catheter in place.

The butterfly wings of tape are sutured to the skin using a 20 gauge hypodermic needle and nonabsorbable suture such as medium weight vetafil.

Antiseptic or antibiotic ointment is applied on a gauze square to the site that the catheter penetrates the skin. BetadineTM ointment is being applied in this picture.

A 1/3 to 1/2 inch thick pad of gauze is placed behind the catheter to elevate it away from the neck. This change in position of the external portion of the catheter “pushes” the catheter under the skin at the level of the skin puncture. If a segment of the flexible part of the catheter is visable at the puncture site, that segment of catheter may be bent and thus occluded.


One inch tape is placed completely around the neck of the animal on either side of the catheter, catching the butterfly wings and the gauze pad with the antiseptic ointment.

This is followed by a layer of stretchy wrap such as cast padding or cling gauze. The wrap is placed around the neck both in front of, and behind, the external segment of catheter, two to three times on each side. The weave of the gauze can be loosened and placed over the top of the catheter, allowing the catheter to protrude through the gauze.

A piece of 1 inch tape split lengthwise can be wrapped around the catheter for added security.

This is followed by an outer layer of Vet Wrap. This wrap is placed in front of, and behind the catheter. The wrap encircles the neck twice on each side of the catheter. A small hole is cut in the wrap and it is passed over the catheter. The tip of the catheter and injection cap protrude through the hole in the wrap

Another piece of 1 inch tape (~4 inches long) split lengthwise, is wrapped around the top of the catheter to make it more secure.
Jugular catheters can be used for obtaining blood samples. They are especially useful for repetitive sampling such as occurs when monitoring a diabetic animal. When using an Intrafusor, which has an attached extension set, an extra piece of tape should be applied to the extension set. If the animal pulls on the catheter the chances of it being dislodged are reduced if the catheter is securely taped in place.

[Murat KUTAY]

Placement of a Butterfly Catheter

Butterfly catheters have a steel needle (a) attached to flexible plastic wings (b) and a short piece of extension tubing (c). A 3 way stopcock (d) is attached to the catheter in this slide but is not used when the butterfly catheter is placed IV. The 3 way stopcock is used when the butterfly catheter is used for thoracocentesis (see thoracocentesis from selection menu for details).
Butterfly catheters are used for short term administration of drugs (drugs administered over a few minutes). For example, the anticancer drug vincristine or thiacetarsimide which is used to kill adult heartworms. Both of these drugs are very caustic to tissues. To assure that the injection is being administered IV rather than SC, sterile fluid is flushed through the catheter before and after drug administration.
Placement of a butterfly catheter allows you to easily change between syringes containing drug or saline. It usually takes less time to place a butterfly catheter compared to other cephalic catheter types. Butterfly catheters are not taped in place for longer term use as the needle will lacerate the vein as the animal moves about.

The catheter can be filled with saline before placement or air can be left in the line, depending upon blood to displace the air. The wings are folded up and held between thumb and index finger. The skin and vein are punctured with one thrust of the needle. The needle is beveled and should be placed with the opening of the bevel facing up. Notice the thumb of the hand holding the leg is placed adjacent to the vein to stabilize it.

The needle is inserted into the vein to the level of the plastic wings. Notice that the catheter has been placed at the junction of the cephalic vein with the accessary cephalic vein.

The thumb of the hand holding the leg is used to hold the catheter in place during drug administration. The catheter and leg will move as a unit when held with the same hand, preventing the needle from accidently withdrawing from the vein during drug administration.

[Murat KUTAY]

(KEMİK İLİĞİ ASPİRASYONU)

The sites that are most accessible for bone marrow aspiration in the dog are the proximal humerus (a), proximal femur (b) and the wing of the ilium (c), approached either from the dorsal crest or lateral face. The easiest sites from which to obtain bone marrow in the cat are the proximal femur and proximal humerus.

The most common needle types used for aspiration of bone marrow are the Illinois sternal-iliac and the Rosenthal needles (pictured). Needles are available in 16 and 18 gauges, 1 or 1-15/16 inch long. Aspiration-style needles have an inner stylet that is used to penetrate the cortex of the bone, then removed to aspirate marrow. Before introducing the needle into the bone, check to make sure the stylet totally occludes the lumen of the needle. If the lumen is not filled by the stylet, pieces of cortical bone may enter the lumen of the needle as it is driven into the bone. These pieces of cortical bone will plug the needle, preventing aspiration of marrow.

If you have relatively short fingers, the Rosenthal needle is held with the back of the needle pressed at the junction between fingers and palm. This gives you maximum driving force and stability. The stylet does not lock in place and must be held in the needle with digital pressure as the needle is advanced into the bone to prevent the lumen of the needle from becoming obstructed with cortical bone

For individuals with longer fingers, the needle can be held between the index and middle finger, and the thumb can be used to hold the stylet in place.

Bone marrow aspiration is performed in the dog with lidocaine local anesthesia and in cats, under short duration general anesthesia. The skin surface is clipped of hair and aseptically prepared as for surgery. Sterile gloves are usually worn. If local anesthesia is being used, the agent is placed in the skin, muscle and on the periostium of the bone.

A small skin incision is made with a scalpel blade. The bone marrow needle is used to bluntly cut through the muscle. Once the needle is in contact with the surface of the bone it is rotated into the bone with a clockwise/counter-clockwise motion. It takes a considerable amount of force to drive the needle through the cortex of the bone.

To obtain marrow from the lateral aspect of the wing of the ilium the animal is restrained in lateral recumbency.

Palpate the dorsal crest of the wing of the ilium and move down approximately 1 cm (* marks the site of bone penetration on photo below). The marrow cavity is shallow in this location (see below). The needle cannot be seated very deeply or it will pass through the marrow cavity and into or through the cortical bone on the opposite side of the wing. This technique is not recommended for use in small dogs (~25 lb or less) or cats due to the tendency to penetrate both cortices. Obese or very large dogs may have too much paralumbar fat &/or muscle for the needle to pass through before reaching the surface of the bone.

Bone marrow can be aspirated from the dorsal crest of the wing of the ilium. The animal is restrained in sternal recumbancy or standing. Hold the crest of the ilium between your fingers. The dorsal surface of the crest is rounded, and the needle tends to slide off the bone into soft tissues if you don’t maintain control of it. The needle is directed parallel to the long axis of the wing of the ilium. The needle can be firmly seated in this location.

Bone marrow can be obtained from the proximal humerus of most animals, including those of small body size or obese condition. The bone marrow needle can be seated firmly in this location as the marrow cavity is thick. Run your finger down the spine of the scapula. The first prominance you feel is the acromion (a). The next prominance is the greater tubercle of the humerus (b). The needle is inserted at the distal end of this bony prominance (c). Maintain control of the needle as you are seating it into the bone. There is a tendency for the needle to slide down the surface of the bone instead of penetrating the cortex, causing damage to adjacent soft tissues.

The needle should be inserted at an angle ~450 from a line parallel to the long axis of the humerus. Needle insertion too proximal may result in entry into the scapulohumeral joint. Needle insertion at an angle perpendicular to the humerus may result in entry into the bicipital bursa that communicates with the scapulohumeral joint on the medial side of the limb.

To obtain bone marrow from the proximal humerus, the animal is restrained in lateral recumbency. The elbow (a) is rotated inward such that the shoulder joint is turned outward. This positions the greater tubercle (b) in a location that is easier for you to get the bone marrow needle seated in the bone.

[img]http://courses.vetmed.wsu.edu/samdx/images/bm_femur_linedraw.JPG[/img]

The proximal femoral shaft usually contains red marrow and is relatively accessible for aspiration in the cat. The needle may not be of sufficient length to reach this site in large dogs. The needle is seated in the trochanteric fossa between the lesser and greater trochanters.

Hold the stifle in one hand with your thumb laying along the long axis of the femur and the thumb nail over the greater trochanter. Make a small skin incision with a scalpel blade and insert the needle under your thumb until the tip of the needle contacts the periostium of the intertrochanteric fossa. Keep the needle parallel to your thumb. If you do not remain parallel to your thumb (and hence the femur) the needle may exit the cranial or caudel cortex of the bone. Remember that the sciatic nerve is caudel to the femur and can be injured if the needle slips caudel to the femur.

A 12 or 20 ml syringe is used to aspirate bone marrow. It may be necessary to pull back on the syringe plunger to 10-15 ml to create enough negative pressure to break marrow particles loose from the endosteal surface. Aspirate only a small amount of marrow. Aspiration of large volumes results in dilution of the marrow sample with peripheral blood. When marrow just enters the barrel of the syringe (arrow), stop aspirating. Larger volumes of marrow can be drawn into an EDTA solution and the marrow particles (spicules) picked out with a needle or pipette.

A small drop of marrow is placed on each of several slides. Marrow clots very rapidly.

It is important to make the smears immediately after obtaining the marrow sample.

If the marrow sample was heavily contaminated with peripheral blood during sampling it still may be possible to “salvage” such a sample. Smearing a drop of marrow that has heavy blood contamination results in a cytology sample of low cellularity with regard to marrow elements. The sample may be salvaged using the following technique.

Procurement of a core of marrow

A Jamshidi needle is used to obtain a core of marrow for histologic evaluation. The stylette (a) is locked into the hollow part (b) of the needle with a threaded cap, not shown in the photo. (c) is a thin wire used to dislodge the core of marrow from the needle.

A core of marrow can be obtained from the proximal humerus, proximal femur or dorsal crest of the wing of the ilium. The landmarks for starting the needle are the same as described for aspiration of marrow from the proximal humerus. Once the needle penetrates the cortex of the bone, the stylette is removed. The needle is advanced 1 to 2 cm. At that point, the entire needle is “stirred” in the marrow to break loose the core of marrow. Do this by rotating the blue cap of the needle in a circle which will also rotate the tip of the needle in a circle in the marrow, breaking loose a core in the lumen of the needle. The needle is removed from the marrow using a clockwise or counterclockwise (not back and forth) motion. Rotating the needle out in one direction allows a second opportunity to break loose the core. If the core is still attached and you withdraw the needle, rotaing back and forth, you may pull the needle off the core and leave the core in the marrow cavity.

The stylette is inserted into the tapered tip of the needle and the core is pushed out of the needle.

[Murat KUTAY]

Cystocentesis

Cystocentesis can be performed with the animal in dorsal recumbency (shown below), lateral recumbancy, standing or being held standing on its hindlimbs by elevating its forequarters. The position of the animal is primarily based on personal preference. Cystocentesis is usually performed with a 25-22 gauge needle.

Although some veterinarians perform cystocentesis without preparation of the puncture site, (they just wet the site with alcohol) it is the preference of the author to clip the puncture site of hair and clean the area with an antiseptic solution such as BetadineTM.

The bladder is palpated and immobilized. The bladder should not be squeezed tightly as the puncture is being made as this can cause urine to leak from the puncture site into the abdominal cavity.

The needle should be inserted at a 450 angle, a short distance cranial to the junction of bladder and urethra (a) If the needle is inserted at the apex of the bladder (b), as urine is removed, the bladder gets smaller and moves away from the needle. The needle should be inserted into the bladder while creating negative pressure by pulling back on the plunger of the syringe. The needle should not be redirected if urine is not obtained, due to the risk of penetrating a bowel loop and subsequently taking the contaminated needle into the bladder. If a sample is not obtained on the first attempt, change the needle before making another attempt. If a sample is not obtained on 3 attempts, the bladder is probably small and in the pelvic canal. If a sample is obtained, the syringe plunger is released, and the needle is removed from the abdomen.

If you cannot palpate the bladder, you can perform “blind” cystocentesis with the dog in dorsal recumbancy. Pick a point on the midline, midway between the umbilicus (u) and brim of the pelvis (p) . Notice when you are preparing the puncture site, this is the same point at which the antiseptic solution will pool when the patient is in dorsal recumbancy. This is the starting point of cystocentesis. If urine is not obtained with the first puncture, two additional punctures can be attempted from 1 to 2 centimeters cranial and 1 to 2 centimeters caudal to the initial puncture site. If urine is not obtained on the first attempt, change the needle before making another attempt. If a sample is not obtained on 3 attempts, the bladder is probably small and in the pelvic canal. Try again later to collect a sample

Cystocentesis can be performed on the midline in male dogs by retracting the penis and prepuce off the midline.

[Murat KUTAY]

Collection of Cerebral Spinal Fluid
SEREBRO SPİNAL SIVI ALINMASI

[img]http://courses.vetmed.wsu.edu/samdx/images/csf_needle.JPG[/img]

Spinal needles for use in small animals are available in 20 to 22 gauge and 1- 1/2 to 3- 1/2 inches in length. All spinal needles have a stylet.
Look at the tip of the needle. The bevel of the needle is shorter than the bevel of a hypodermic needle. Spinal fluid is obtained from the subarachnoid space which is a shallow space. If you use a needle with a long bevel, part of the opening into the lumen of the needle may be outside the subarchnoid space.
[img]http://courses.vetmed.wsu.edu/samdx/images/csf_landmarks.JPG[/img]
First method: The site of CSF collection from the cisterna magna (a) is between the occipital crest (b) and the most prominent points of the wings of the atlas (c). Although some textbooks recommend insertion of the needle at the halfway point between these two landmarks, I find one tends to be just anterior to the cisterna at the halfway point. Instead, I divide this distance into thirds and make the puncture 2/3 of the way back from the occipital crest.

Second method: Instead of the above mentioned landmarks, use your fingertips to trace the wings of the atlas to their most anterior margins. Palpate the occipital crest to define the midline. The cisterna is located on the midline, at the level of the anterior aspect of the wings of the atlas.

The animal should be placed in lateral recumbency with the nose flexed and ears pulled ventrally. The dorsum should be close to the edge of the table

[img]http://courses.vetmed.wsu.edu/samdx/images/csf_collect.JPG[/img]

The puncture site is clipped of hair and aseptically prepared for the procedure. The person performing the procedure is wearing sterile gloves. The puncture is made on the midline. The needle is advanced slowly and carefully, removing the stylet at intervals to check for the presence of spinal fluid in the needle. If you are unsure if there is fluid in the needle, wipe the stylet across your sterile glove and look for a streak of fluid. If the needle is dry, replace the stylet and continue to slowly advance the needle. If the needle hits bone, and fluid has not been obtained, angle the needle cranial or caudel (which direction depends upon initial point of needle entry) and try to “walk” the needle off the bone into the cisterna. If you are unsure whether to “walk” cranial or caudel, remove the needle, revaluate landmarks and try again. If you obtain blood, let a few drops flow. If the fluid remains heavily bloody, remove the needle. You entered a lateral venous sinus. Try again. If the fluid becomes clear, collect it for analysis.

Although a manometer can be applied to the needle to measure the opening pressure of CSF, I find that application of the manometer often results in loss of the puncture. Therefore I do not routinely measure CSF pressures. You can collect CSF using a small (3ml) syringe or by gravity into a sterile container. If you use a syringe to collect CSF, do not attach the syringe to the hub of the needle. Rather use the syringe to aspirate drops of CSF that form in the hub of the spinal needle. Continue to hold the hub of the spinal needle as spinal fluid is collected so the tip of the needle is not dislodged from the subarachnoid space. Compression of the jugular veins will increase the rate of flow of CSF. The sample for cytology and fluid analysis is placed into an EDTA tube. Samples for bacterial or viral culture are not anticoagulated.

[Murat KUTAY]

Gastric Intubation

The indications for gastric intubation include:
emptying and gavage of the stomach in cases of suspected or known poisoning
relief of gaseous distension in animals with gastric dilatation volvulus
short-term administration of nutrients
administration of diagnostic solutions such as barium
[img]http://courses.vetmed.wsu.edu/samdx/resize/dogSternal.JPG[/img]
General gastric intubation is performed using a large bore gastric tube. The diameter of the tube should be approximately the same size as an endotracheal tube that would be used in the same animal.

The length of the tube should be measured from the tip of the nose to approximately the 9th intercostal space in order to assure that the tip of the gastric tube is within the lumen of the stomach. Place a tape marker (orange tape pointed to by blue arrow) to mark the proper distance.

[img]http://courses.vetmed.wsu.edu/samdx/images/lubricate.JPG[/img]
Over-insertion can result in the tube hitting the gastric wall, flipping 180 degrees, and exiting back into the esophagus. Large diameter tubes are by nature very stiff. Care should be used in their placement, as gastric perforation can occur. The tube should be well lubricated. Notice that the tube pictured has an end hole as well as side holes (indicated by blue arrow).

Orogastric intubation is usually performed in the awake animal with physical restraint when the purpose is to relieve gas distention or administer barium. If the purpose is to gavage to remove toxic contents, heavy sedation or anesthesia is needed.

If awake, the animal is restrained in sternal recumbency with the head in a neutral position.
[img]http://courses.vetmed.wsu.edu/samdx/resize/tapeRoll2.JPG[/img]
A speculum is placed to prevent the animal from chewing the gastric tube. Commercial speculums are available. A 1 inch roll of tape makes an excellent speculum and is minimally traumatic.

A large bore (12 ml) syringe casing cut off at the end can also be used as a speculum, but the rough plastic is more traumatic to the oral mucosa. The plastic speculum can be wrapped with adhesive tape to reduce trauma to the oral membranes. The speculum is placed between the dental arcades and the muzzle is held to prevent the animal from spitting out the speculum.

[img]http://courses.vetmed.wsu.edu/samdx/resize/passtube2.JPG[/img]
The tube is introduced into the oral cavity with the head in a normal position, not extended, not flexed.

You may be able to observe the tube passing on the left side of the animal’s cervical region as it passes through the cervical esophagus or you may palpate the gastric tube in the cervical region, dorsal to the trachea.
[img]http://courses.vetmed.wsu.edu/samdx/images/infuseFluids2.JPG[/img]
When the tube meets the cardia of the stomach there may be some degree of resistance. This is especially true if the stomach is distended with solid matter or air. A gentle rotation of the gastric tube may be necessary to pass through the lower esophageal sphincter into the stomach.

Depending upon the gastric contents, when the tube enters the gastric lumen, air may rush out, fluid may rush out, or you may have no identifiable sensation that the tube is in the gastric lumen. The tube should be inserted to the previously measured length (orange tape).

[img]http://courses.vetmed.wsu.edu/samdx/resize/checkCuff.JPG[/img]
Effective gastric lavage generally requires the animal be sedated or anesthetized. The animal should have a cuffed endotracheal tube in place, and the insufflation of the cuff should be checked immediately before passing the gastric tube.
[img]http://courses.vetmed.wsu.edu/samdx/resize/infuseFluids.JPG[/img]
The animal’s head should be lowered.
Water, saline or a fluid solution containing activated charcoal should be instilled in volumes of 5-10 ml/kg. After each fluid installation, the fluid should either be drained out by gravity, or aspirated out by using a syringe or suction pump. The gavage should be repeated 8-10 times.

The animal needs to be carefully monitored when recovering from the anesthesia so that it does not vomit and aspirate residual gavage solution.

[Murat KUTAY]

Placement of a Thoracic Drain

Chest drains are usually of large diameter (10 to 32 French depending upon patient size). A large diameter tube is necessary if being used to remove viscous fluid from the pleural space.

A flexible, rubber feeding tube with additional fenestrations (arrow) can be used as a chest drain. Fenestrations should be in the most distal portion of the catheter (~ last 4 -5 cm). When the tube is in place, all fenestrations should be within the pleural space rather than the subcutaneous space. The white roller clamp is used to occlude the lumen of the tube when it is not being aspirated. This clamp should be closed when placing the tube in the pleural space.

Pictured is an argyle styleted thoracic drain. The stylet is placed inside the flexible plastic catheter. The end of the flexible catheter is fenestrated for several centimeters. The sharp point is used to drive the catheter through the chest wall.

Pictured is a Jackson-Pratt drain which is generally used for abdominal drainage. It also functions well as a pleural space drain. The part of the drain labeled (a) is made of teflon and is very nonirritating to tissues. The small black spots are multiple drainage holes. The teflon portion and part of the clear plastic tubing will be positioned in the pleural space. The other end of the clear tubing is attached to the bulb. The bulb (b) can be compressed to create a source of constant suction to enhance fluid removal from the pleural space.

Sedation and local anesthetic infiltration are generally adequate for drain placement. The chest wall should be clipped of hair and scrubbed with antiseptic solutions. The animal is placed in lateral recumbancy. The drain should enter the skin surface at the 10th intercostal space, tunnel subcutaneously, 2 to 3 spaces, and penetrate the chest wall at the 7th or 8th intercostal space. The insertion site should be at the junction of the dorsal and middle thirds of the chest. The dog’s head is to your right.

A skin incision is made with a scalpel blade in the 10th intercostal space, caudel to the 10th rib, at the junction of the dorsal and middle thirds of the chest wall.

Placement of a flexible, rubber feeding tube is demonstrated. Before placing the chest tube, estimate the length of tubing that will be inserted into the pleural space. Excessive length of tubing in the pleural space can lead to the tubing becoming bent or twisted. The ideal position of the tip of the tube is to lie along the sternum, anterior to the heart. Be sure to have ready an adapter that can be firmly attached to the chest drain to allow attachment of a syringe. The tubing should be clamped closed during placement. The feeding tube is held between the jaws of a curved hemostat. Make sure that the jaws of the hemostat extend beyond the tip of the catheter. The tip of the hemostat is used to puncture through the muscle of the chest wall. It is difficult to puncture the chest wall if the tube is at the end of the hemostat.

Pass the hemostat in an anterior direction in the subcutaneous space for a distance of 2 or 3 rib spaces, then firmly drive the hemostat through the chest wall at the 7th or 8th intercostal space. Keep the hemostat close to the cranial aspect of the rib, in order to avoid the intercostal vessels located on the caudel border of the ribs. It takes a considerable amount of force to drive the hemostat through the chest wall. The hemostat should be at a 90 degree angle to the chest wall when being driven through it. When the hemostat enters the pleural space, open the jaws of the instrument and push the tube into the pleural space. Remove the hemostat. Before the thoracic drain is sutured in place, attach a syringe to the catheter and remove fluid and/or air to improve the patient’s breathing ability. Then the catheter is clamped closed and sutured in place.

You can see the subcutaneous tunnel traversing 3 intercostal spaces anterior to the skin incision and entering the pleural space at the 7th intercostal space.

Another technique that can be used during insertion of any of the catheter types presented is to pull the skin in an anterior direction so the skin over the 10th intercostal space is located over the 7th or 8th intercostal space. With the skin pulled forward, you can incise the oblique musculature of the chest wall to make it easier to bluntly force the catheter through the rest of the chest wall musculature. The catheter is driven through the chest wall at ~ 80 to 90 degree angle. The operator is pushing the catheter through the chest wall with her right hand and is using her left hand as a “stop” to control the depth of insertion into the pleural space.The subcutaneous tunnel is created when the skin is released. This slide demonstrates placement of a styletted catheter. After penetrating the chest wall, the catheter and stylet are angled forward and parallel to the chest wall, to direct the catheter forward when it is slid off the stylet. (the dog’s head is to your right)

After the catheter is inserted into the pleural space, a purse string suture is placed in the skin incision around the catheter. The ends of the suture are left long and wrapped up and along the length of the tubing in a finger trap or friction type suture.

The subcutaneous tunnel is collapsed around the tube with a suture. A 20 gauge hypodermic needle is passed underneath the subcutaneous segment of the thoracic tube, taking care not to penetrate the tube with the needle. Suture is threaded through the hypodermic needle, the needle is removed, and the suture tied around the subcutaneous segment of the tube.

[Murat KUTAY]

Urinary Catheterization of the Dog

There are several types of urinary catheters available. The top catheter is a stainless steel catheter used only in female dogs. It is sometimes called a “bitch” catheter. The second catheter from the top of the screen is a Foley catheter.

Foley catheters have an inflatable bulb at the end that can be filled with air or fluid (~3-5 ml) to retain the tip of the catheter within the bladder. Foley catheters are too short to reach the bladder of male dogs. The third catheter is a semi-rigid plastic (polypropylene) urinary catheter. This catheter type can be used in either sex. The bottom catheter is a red rubber feeding tube that can be used as a urinary catheter in either sex dog. The smaller diameter red rubber catheters (<10 Fr)may not be long enough to reach the bladder of large male dogs.

Foley Catheter:
The syringe is attached to an adapter which is attached to a thin tube within the catheter. Fluid or air is injected into this tube which terminates in a thin walled balloon. The balloon capacity is printed on the white plastic adapter (a) (usually 3 to 5 ml). The balloon is inflated when the catheter is in the lumen of the bladder (b). The intent is to retain the catheter in the bladder. The balloon is deflated prior to catheter removal by aspirating back the fluid or air. The adaptor contains a valve which prevents loss of fluid or air.

Urinary catheters are sized using French (fr) units. The French number divided by 3 is the outer diameter of the catheter in milimeters.

The rounded tip of the catheter reduces urethral trauma as the catheter is passed. Urine enters the lumen of the catheter through two “eye” holes.

Polypropylene and rubber catheters can be gas sterilized. The autoclave packaging can be used to maintain sterility of the catheter while it is being passed into the urethra. A finger tab about 1 inch wide is cut or torn from the end of the package closest to the rounded tip of the catheter, ~2 inches from the end of the package.

The finger tab is cut free of the reminder of the package. The end of the package closest to the rounded end of the catheter is removed, taking care not to contaminate the exposed end of the catheter. The finger tab is used to guide the catheter into the urethra.

The tip of the catheter should be lubricated with a water soluble lubricant to minimize urethral trauma. The individual packages of lubricant are prefered over multiple use tubes. The multiple use tubes may become contaminated and be a source of contamination of the urinary tract during catheterization.

The penis is extruded from the prepuce and is cleansed with an antiseptic solution such as BetadineTM or NovalsanTM. The finger tab is used to hold the catheter as it is being introduced into the urethra.

The urethra is limited in distensibility by the os penis surrounding it’s dorsal surface. The catheter tends toward taking a straight course at the ischial arch . The catheter can be palpated in the perineal region. You can gently push on the catheter through the skin in the perineal region to guide the catheter around the arch.

A sterile hemostat can be used to feed the catheter out of the sterile package.

Female dogs can be catheterized by digitally palpating the urethral orifice, then passing the catheter under your finger…

or a speculum can be used to visualize the urethral orifice. These are human nasal specula but work well as canine vaginal specula. The one on the bottom has blades approximately 3/4 of an inch long. The blades are too short to aid in visualization of the urethral orifice in most bitches. The blades of the top speculum are approximately 2 inches long.

When using a vaginal speculum, the blades are initially directed vertically until the clitoral fossa is passed and then they are redirected horizontally. The handles of the speculum should be pointed towards the tail so they are not in the operator’s way as they are passing the urethral catheter.

This is an anal scope that works well to visualize the urethral orifice. The speculum (a) and rounded tip insertion guide (b) are lubricated and inserted into the vagina. The insertion guide is removed by pulling on the metal ring (c) attached to the guide. The light source (d) is turned on. The speculum is slowly retracted until the urethral orifice is visualized. The urinary catheter is passed into the bladder and then the speculum is removed. If the urethral orifice is not visualized, reinsert the insertion guide into the speculum before advancing the speculum into the vagina. Note the edge of the speculum is sharp.

This is what the view looks like through the scope. The arrow points to the urethral orifice. The other two lines in the vaginal mucosa are folds in the tissue. Some bitches have a prominant mound of raised tissue around the urethral orifice. Gently probe the urethral orifice with the urinary catheter to confirm that it is the urethral orifice and not a fold of tissue.

An otoscope with a large (wide but short) cone works well as a vaginal speculum to visualize the urethral orifice in the bitch. This is a Foley catheter being placed throughthe otoscope. Unfortunately, the flare end of the Foley will not fit through the otoscope cone and you need to disconnect the cone (a) from the light source (b) and leave the cone with the catheter until the catheter is removed from the patient. The letter (c) points to a magnifying lens attached to the light source. Magnification makes it easier to see the urethral orifice.

A 3 1/2 French polypropylene catheter will fit through the lumen of the Foley catheter to increase its rigidity and make it easier to pass.

[Murat KUTAY]

Nasogastric Tube Placement

Supplies needed include:

* Soft rubber feeding tubes*
* Topical anesthetic **
* Aqueous lubricant

*A 3.5 or 5 Fr tube can be placed in most cats. An 8 Fr tube is difficult to place in all but large cats.

**The same topical anesthetic that is used in performance of ophthalmic exams can be placed in the nose.

Instill several drops of local anesthetic into one nostril. The cat’s head is slightly tilted back to allow the topical anesthetic to run into the nasal cavity. Only a few drops of anesthetic agent are needed. Large amounts of anesthetic agent may run back into the pharynx and anesthetize the arytenoid cartilages, possibly resulting in tracheal intubation rather than gastric intubation.

The tube should be liberally lubricated with aqueous lubricant. The nasal turbinates are very friable and traumatic passage of the nasal tube will result in hemorrhage.

The nasal cavity is divided by the turbinates into several spaces called meatuses. The nasogastric tube is passed into the ventral meatus which communicates with the nasopharynx. The tube is inserted into the nostril as close to the midline and as ventral as possible. If the tube initially passes and then meets resistance, it is hitting against the ethmoid turbinates; draw back the tube and gently advance until it passes into the nasopharynx and subsequently into the esophagus.

[Murat KUTAY]

Urinary Catheterization of the Male Cat

Several styles of urinary catheters are available. The catheter identified by the letter (a) is a flexible rubber feeding tube. A 3 1/2 Fr size is used to catheterize a male cat. Because it is flexible it is more difficult to pass than the other two catheters that are pictured. It is longer than needed to reach the urinary bladder. (b) is an open-ended polypropylene catheter. (c) is a close-ended polypropylene catheter. Both (b) and (c) are 3 1/2 Fr.

To prevent urethral trauma, the penis should be extruded from the prepuce and held parallel to the vertebral column. The cat’s head is to your right.
Passing a urinary catheter in a male cat requires two people. One person exteriorizes the penis and the second person passes the catheter. A finger tab cut from the sterile package that the catheter was wrapped in, can be used to manipulate the catheter without touching it. (If this is not clear, see urinary catheterization of the dog from the selection menu) The catheter should be lubricated with aqueous lubricant. The penis and prepuce should be cleansed with antiseptic solution. In the slide on the video screen, the person passing the catheter has a syringe filled with saline that is slowly being infused to dilate the urethra ahead of the catheter. If you are passing the catheter to obtain a urine sample, do not infuse fluid during passage of the catheter. The infusion technique is used to dislodge debris from the urethra of an obstructed cat.

[Murat KUTAY]

Venipuncture of the Cat
“Cats are not just small dogs”

The same techniques for placing cephalic and jugular catheters as described for the dog, are applicable to the cat. In addition, there exist some procedures that are more commonly used to venipuncture the cat.
[img]http://courses.vetmed.wsu.edu/samdx/resize/cat_veni_bag.JPG[/img]
A canvas or nylon, zippered bag can be used to restrain a cat for jugular venipuncture or catheter placement. Some bags have multiple zippers to allow access to limbs.

The cat is held in dorsal recumbancy. The holder places a finger in the thoracic inlet to impair venous return from the head and cause the vein to distend with blood. The venipuncturist holds the head with one hand and makes the puncture with the other hand.

The medial saphenous vein of the cat has a long straight course and is very superficial. The red arrows point to the vein. It is a good vein from which to collect small volumes of blood or to insert indwelling catheters. The cat is restrained in lateral recumbancy. The holder applies pressure in the inguinal region to occlude venous return and cause the vein to engorge with blood.
cat’s head to right

Blood samples can be obtained from the medial saphenous vein. Because the vein has a small diameter, vigorous aspiration results in collapse of the vein. Therefore only slight suction can be applied to the syringe when aspirating blood. The blood flows slowly so only a small volume (up to ~1ml) of blood can be obtained before the sample clots. At the completion of the venipuncture, as the needle is removed from the vein, the holder should release pressure from the inguinal region and place firm digital pressure at the puncture site for several minutes to prevent hematoma formation. Hematomas form readily and spread from the puncture site when venipuncture is unsuccessful. Therefore, start distal on the limb in case it is necessary to make more than one attempt at venipuncture.

The medial saphenous vein can be catheterized with short indwelling catheters or catheters marketed for jugular vein catheterization. (See the sections on jugular catheter placement and cephalic catheter placement for information on catheter types). A jugular catheter placed in the medial saphenous vein will have its tip in the posterior vena cava. Such catheters can be used for repetitive blood sampling (eg. serial blood glucose sampling used in diabetic regulation). Because the tip of the catheter is in a large vein, hypertonic solutions such as hypertonic dextrose and total parenteral nutrition (TPN) solutions can be administered through catheters in this location. The picture demonstrates an Intrafuser jugular catheter placed in the medial saphenous vein. Note the bruising around the needle puncture site. This may occur when placing a through-the-needle catheter. The hole in the vein created by the needle is larger than the diameter of the catheter so when the needle is retracted from the vein, blood may leak from the puncture site into surrounding tissues.

Correct bandaging of a medial saphenous catheter is crucial to its function. The procedure for bandaging the catheter in place is similar to the procedure for bandaging a jugular catheter in place, except that the bandage is applied in a direction parallel to the needle guard, rather than perpendicular to the needle guard as with the jugular catheter. A butterfly of one inch tape is applied to the needle guard. A piece of one inch tape is wrapped around the leg, catching both wings of the butterfly at the same time. A second piece of one inch tape is wrapped around the limb at the level of the hub of the needle. Antiseptic or antibiotic ointment on a piece of gauze is applied to the puncture site. Stretchy wrap such as cling is wrapped around the limb several times, around both pieces of tape. This is followed by an outer wrap such as Vetwrap. The gauze pad used to elevate the needle guard from the neck when bandaging a jugular catheter is not used when taping in place a medial saphenous catheter. The use of an Intrafuser catheter with an extension set, allows you to place the extension set on the lateral side of the limb so the catheter is easily accessible. It may be necessary to splint the leg to keep the catheter functioning.

[Murat KUTAY]

Cephalic and Saphenous Vein Catheterization

Any of the catheters that can be placed in the jugular vein can also be placed in peripheral veins. Short catheters sold for placement in peripheral veins such as (a) and (b), have the catheter on the outside of the needle so the hole in the vein wall made by the needle is smaller than the catheter. The catheter fills the hole in the vein wall and there is minimal leakage of blood around the catheter.
c) is a butterfly catheter (no, butterflies are not free, they cost ~$2.00). The “catheter” portion that goes in the vein is a rigid needle. Butterfly catheters are used for short term, small volume, infusions such as the administration of CaparsolateTM used to treat heartworm disease.

Because of its rigidity, it is not useful for long term fluid administration as the needle may lacerate the vein. (d) is a catheter that has the conformation of a butterfly catheter but the catheter is flexible teflon rather than metal. The catheter has a wire stylet that is removed after placement. This style of catheter is very useful for placement in peripheral veins in small dogs and cats and breeds of dogs with short, crooked legs. It can also be used to drain fluid or air from body cavities. The flexibility of the catheter after the stylet is removed, reduces the chance of organ laceration.

Have all necessary supplies ready before placing the catheter. Supplies needed in addition to the catheter include one inch tape, (2 pieces of sufficient length to encircle the limb), an injection cap and a 10-12 ml syringe filled with saline or heparinized saline. If the catheter is to be left in place for several days, a small amount of antiseptic or antibiotic ointment on a gauze pad should be placed over the catheter puncture site before bandaging the catheter in place.

The catheter placement site should be widely shaved and the skin scrubbed with antiseptic solutions, using the same technique as for preoperative skin preparation.

Notice that the holder is standing on the side of the dog opposite the leg that is being catheterized. The dog is restrained close to the body of the holder. The muzzle is held away from the face of the holder and the person placing the catheter. She is reaching over the dog to hold off the vein and can apply downward pressure over the dog’s back, if needed to keep the dog in sternal recumbancy. The dog’s leg is being held at the elbow to prevent her from pulling back her leg. The individual holding the leg places the thumb of the same hand across the dorsum of the limb to occlude venous blood returning from the leg, causing the vein to distend with blood. In some cases the vein will be clearly visable, in other cases you may palpate the distended vein.

The holder places her thumb firmly on the medial side of the most proximal aspect of the limb. The thumb is “dragged” to the dorsal aspect of the leg which will “roll” the cephalic vein to the dorsum of the leg. Pressure is applied with the thumb to restrict blood flow returning from the distal limb, causing the vein to engorge with blood. The catheter should be placed as distal in the vein as possible. If the catheter is too proximal, its tip will lay at the elbow. As the animal withdraws its leg, flow through the catheter may cease. You can catheterize the cephalic vein on the medial side of the limb, at a location distal to the junction of the cephalic and accessory cephalic veins. Before making the puncture, the venipuncturist can lay the thumb of the hand that is holding the leg, adjacent to the vein to reduce vein movement when it is being punctured.

f the catheter you are using has a solid cap, remove it. Some catheters have “flashback” caps with holes in the center to allow air in the catheter to be displaced by blood when the venipuncture is made. “Flashback” caps may be left on the catheter when the puncture is made.

Puncture the skin and vein in one swift movement. If you are too gentle, the vein moves away from the catheter. Once the vein is punctured, blood will flow through the needle that is inside the catheter.
Move the thumb and forefinger of the hand holding the leg toward each other and grasp the needle of the catheter, still holding the leg in the same hand. By holding the needle this way, if the patient pulls his leg away from you, the catheter will not be pulled out. Using the other hand (right hand for a right handed person) gently rotate the catheter off the needle, advancing the catheter into the vein. As the needle is removed, blood will flow from the catheter. At this time the holder should remove their thumb from the dorsum of the leg (continuing to hold the leg). If the holder presses firmly over the vein just proximal to the tip of the catheter, less blood will flow from the catheter, making the taping procedure less bloody.

After the stylette is removed, an injection cap is placed and the catheter is flushed well with saline or heparinized saline, to assure patency. Dry the leg and the catheter with gauze before applying tape.

The injection cap should not be taped into the bandage so that it can be easily removed later.

Taping a cephalic catheter in place:


Fold over the end of the tape to create a tab for easier removal.

A second piece of tape placed under the cap will allow easier removal and replacement of the cap and prevent hair from touching the tip of the catheter.

The injection cap can be removed for direct connection of a fluid administration set (a) to the IV catheter, or the cap can be left on the catheter and the fluid administration set attached using a 20 gauge hypodermic needle inserted through the cap. The administration set is “looped” and taped to the leg (b) . This reduces the chance of accidental removal of the catheter if the administration set is pulled.


A hypodermic needle with plastic cover, attached to an injection cap can be taped to the IV pole or the fluid bag or bottle. When fluids are temporarily stopped, the injection cap is placed on the end of the catheter and the needle with plastic cover is placed on the end of the IV administration set to keep the tip of the tubing sterile.

Movement of the leg can occlude fluid flow through the catheter. If needed, a splint such as the pictured Mason-meta splint can be used to keep the limb extended to maintain a constant fluid flow rate. The leg and splint can then be bandaged or the splint can be taped to the leg at both ends, leaving the catheter exposed. The use of IV infusion pumps reduces the need to keep the limb extended. Fluid pumps can often overcome the resistance created by positional changes.

Saphenous Vein:


To place a catheter or obtain blood from the lateral saphenous vein, the animal is positioned in lateral recumbancy. The holder is holding off the vein with her right hand. Some dogs have a prominant medial saphenous vein which can be catheterized or sampled. To access the medial saphenous vein, the animal is held in lateral recumbancy but the holder applies pressure on the medial aspect of the leg closest to the table. *

The technique for placing a catheter in the lateral saphenous vein of the dog is similar to the technique for cephalic placement. Notice the venipuncturist has placed her thumb adjacent to the vein to stabilize it.

[Murat KUTAY]

Intramuscular injections (i.m.)

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