Diagnostic and Therapeutic Procedures
Identifieur interne : 000248 ( Pmc/Corpus ); précédent : 000247; suivant : 000249Diagnostic and Therapeutic Procedures
Auteurs : Richard B. FordSource :
- Kirk and Bistner's Handbook of Veterinary Procedures and Emergency Treatment ; 2006.
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DOI: 10.1016/B0-72-160138-3/50005-9
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PubMed Central: 7158358
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<record><TEI><teiHeader><fileDesc><titleStmt><title xml:lang="en">Diagnostic and Therapeutic Procedures</title><author><name sortKey="Ford, Richard B" sort="Ford, Richard B" uniqKey="Ford R" first="Richard B." last="Ford">Richard B. Ford</name></author></titleStmt><publicationStmt><idno type="wicri:source">PMC</idno><idno type="pmc">7158358</idno><idno type="url">http://www.ncbi.nlm.nih.gov/pmc/articles/PMC7158358</idno><idno type="RBID">PMC:7158358</idno><idno type="doi">10.1016/B0-72-160138-3/50005-9</idno><idno type="pmid">NONE</idno><date when="2006">2006</date><idno type="wicri:Area/Pmc/Corpus">000248</idno><idno type="wicri:explorRef" wicri:stream="Pmc" wicri:step="Corpus" wicri:corpus="PMC">000248</idno></publicationStmt><sourceDesc><biblStruct><analytic><title xml:lang="en" level="a" type="main">Diagnostic and Therapeutic Procedures</title><author><name sortKey="Ford, Richard B" sort="Ford, Richard B" uniqKey="Ford R" first="Richard B." last="Ford">Richard B. Ford</name></author></analytic><series><title level="j">Kirk and Bistner's Handbook of Veterinary Procedures and Emergency Treatment</title><imprint><date when="2006">2006</date></imprint></series></biblStruct></sourceDesc></fileDesc><profileDesc><textClass></textClass></profileDesc></teiHeader></TEI><pmc article-type="chapter-article"><pmc-dir>properties open_access</pmc-dir>
<front><journal-meta><journal-id journal-id-type="nlm-ta">Kirk and Bistner's Handbook of Veterinary Procedures and Emergency Treatment</journal-id><journal-title-group><journal-title>Kirk and Bistner's Handbook of Veterinary Procedures and Emergency Treatment</journal-title></journal-title-group></journal-meta><article-meta><article-id pub-id-type="pmc">7158358</article-id><article-id pub-id-type="publisher-id">B0-7216-0138-3/50005-9</article-id><article-id pub-id-type="doi">10.1016/B0-72-160138-3/50005-9</article-id><article-categories><subj-group subj-group-type="heading"><subject>Article</subject></subj-group></article-categories><title-group><article-title>Diagnostic and Therapeutic Procedures</article-title></title-group><contrib-group><contrib contrib-type="author" id="au1"><name><surname>Ford</surname><given-names>Richard B.</given-names></name><degrees>DVM, MS</degrees></contrib></contrib-group><aff id="aff1">Professor of Medicine, Diplomate, Diplomate (Honorary), Department of Clinical Sciences, College of Veterinary Medicine, North Carolina State University, Raleigh, North Carolina</aff><aff id="aff2">American College of Veterinary Internal Medicine</aff><aff id="aff3">American College of Preventive Medicine</aff><contrib-group><contrib contrib-type="author" id="au2"><name><surname>Mazzaferro</surname><given-names>Elisa M.</given-names></name><degrees>MS, DVM, PhD</degrees></contrib></contrib-group><aff id="aff4">Diplomate, Director of Emergency Services, American College of Veterinary Emergency and Critical Care</aff><aff id="aff5">Wheat Ridge Veterinary Specialists, Wheat Ridge, Colorado</aff><pub-date pub-type="pmc-release"><day>21</day><month>5</month><year>2009</year></pub-date><pmc-comment> PMC Release delay is 0 months and 0 days and was based on .</pmc-comment>
<pub-date pub-type="ppub"><year>2006</year></pub-date><pub-date pub-type="epub"><day>21</day><month>5</month><year>2009</year></pub-date><fpage>449</fpage><lpage>572</lpage><permissions><copyright-statement>Copyright © 2006 Elsevier Inc. All rights reserved.</copyright-statement><copyright-year>2006</copyright-year><copyright-holder>Elsevier Inc.</copyright-holder><license><license-p>Since January 2020 Elsevier has created a COVID-19 resource centre with free information in English and Mandarin on the novel coronavirus COVID-19. The COVID-19 resource centre is hosted on Elsevier Connect, the company's public news and information website. Elsevier hereby grants permission to make all its COVID-19-related research that is available on the COVID-19 resource centre - including this research content - immediately available in PubMed Central and other publicly funded repositories, such as the WHO COVID database with rights for unrestricted research re-use and analyses in any form or by any means with acknowledgement of the original source. These permissions are granted for free by Elsevier for as long as the COVID-19 resource centre remains active.</license-p></license></permissions></article-meta></front><body><sec id="cesec5"><title>ROUTINE PROCEDURES</title><sec id="cesec6"><title>ADMINISTRATION TECHNIQUES FOR MEDICATIONS AND FLUIDS</title><sec id="cesec7"><title>Oral Administration: Tablets/Capsules—Canine</title><p id="para192">Perhaps the simplest and easiest method of administering tablets or capsules to dogs is to hide the medication in food. Offer small portions of <italic>unbaited</italic> cheese, meat, or some favorite food to the dog initially. Then offer one portion that includes the medication. For anorectic dogs or when pills must be given without food, give medications quickly and decisively so that the process of administering the medication is accomplished before the dog realizes what has happened. With cooperative dogs, insert the thumb of one hand through the interdental space, and gently touch the hard palate. This will cause the dog to open the mouth (<xref rid="f1" ref-type="fig">Figure 4-1</xref>). Using the opposite hand (the one holding the medication), gently press down on the mandibular (lower) incisors to open the mouth further (<xref rid="f2" ref-type="fig">Figure 4-2</xref>). Position the tablet or capsule onto the caudal aspect of the tongue as close to the larynx as possible. Quickly withdraw the hand and close the dog's mouth. When the dog licks its nose, the medication likely has been swallowed.<fig id="f1"><label>Figure 4-1</label><caption><p>Use of the thumb only to open a cooperative dog's mouth.</p></caption><graphic xlink:href="gr1"></graphic></fig><fig id="f2"><label>Figure 4-2</label><caption><p>Use of the opposite hand to place a tablet or capsule on the caudal aspect of the tongue.</p></caption><graphic xlink:href="gr2"></graphic></fig></p><p id="para193"><boxed-text id="cetextbox22"><caption><title>Note:</title></caption><p id="para194">Oral medication frequently is dispensed to owners without regard for the client's knowledge of how to administer a pill/tablet or without asking whether the client is even physically able to administer medications.</p><p id="para195">Clear instructions, including a demonstration, and having the client perform the technique in the hospital will improve compliance.</p></boxed-text></p><p id="para196">Dogs that offer more resistance can be induced to open their mouths by compressing their upper lips against their teeth. As they open their mouth, roll their lips medially so that if they attempt to close their mouth, they will pinch their own lips.</p><p id="para197">Dogs that struggle and slash with their teeth are the most difficult, especially if they show aggression toward the individual attempting to administer mediation. They often can be medicated by placing the tablet over the base of the tongue with a 6-inch curved Kelly hemostat or special pill forceps. Cubes of canned food or dried meat often can be “pushed down” a placid but anorectic patient by using the thumb as a lever. The fingers are kept out of the mouth, but the thumb is inserted behind the last molar of the open mouth and pushes the bolus down.</p></sec><sec id="cesec8"><title>Oral Administration: Tablets/Capsules-Feline</title><p id="para198">Two methods of pill administration are used in cats. In both methods the cat's head is elevated slightly. Success in administering pills/tables to a cat entails a delicate balance between what works well and what works safely. In cooperative cats, it may be possible to use one hand to hold and position the head (<xref rid="f3" ref-type="fig">Figure 4-3</xref>) while using the opposite hand (the one holding the medication) to open the mouth gently by depressing the proximal aspect of the mandible (<xref rid="f4" ref-type="fig">Figure 4-4</xref>). Press the skin adjacent to the maxillary teeth gently between the teeth as the mouth opens, thereby discouraging the cat from closing its mouth. With the mouth open, drop the medication (try lubricating the tablet or capsule with butter) into the oral cavity as far caudally on the tongue as possible. The cat can be tapped under the jaw or on the tip of the nose to facilitate swallowing if you really think this works. If the cat licks, administration was probably successful.<fig id="f3"><label>Figure 4-3</label><caption><p>Head restraint technique used while administering a tablet/capsule to a cat.</p></caption><graphic xlink:href="gr3"></graphic></fig><fig id="f4"><label>Figure 4-4</label><caption><p>Use of the opposite hand gently to depress a cat's mandible before dropping a tablet into the caudal aspect of the oral cavity.</p></caption><graphic xlink:href="gr4"></graphic></fig></p><p id="para199">CAUTION: Only experienced individuals should attempt this technique of administering tablets/capsules to cats. Even cooperative cats that become intolerant will bite. Therefore, this is NOT a technique recommended for inexperienced owners to try at home, even if specific instructions have been given.</p><p id="para200">Alternatively, some cats will tolerate a specially designed “pilling syringe” in an attempt to administer a tablet or capsule. The pilling syringe works well as long as it is inserted cautiously and atraumatically into the cat's mouth. However, if resistance ensues, the rigid pilling syringe may injure the hard palate during the ensuing struggle. Subsequent attempts to use the syringe may be met with increasing resistance and increasing risk of injury. Success with a pilling syringe depends largely on the cat.</p><p id="para201">When dispensing oral medications for home administration to cats, do <italic>not</italic> expect clients to force a tablet or capsule into a cat's mouth. Although some clients are remarkably capable and confident with their ability to administer oral medications to cats, the risk of injury to the client can be significant. Whenever feasible, liquid medications or pulverized
tablets should be mixed with the diet or an oral treat readily accepted and consumed (see the following discussion).</p></sec><sec id="cesec9"><title>Oral Administration: Liquids</title><sec id="cesec10"><title>Without a stomach tube</title><p id="para202">Small amounts of liquid medicine can be given successfully to dogs and cats by pulling the commissure of the lip out to form a pocket (<xref rid="f5" ref-type="fig">Figure 4-5</xref>). Hold the patient's head level so that the medication will not ooze into the larynx. Deposit the liquid medication into the “cheek pouch” where it subsequently flows between the teeth as the head is held slightly upwards. Patience and gentleness, along with a reasonably flavored medication, are needed for success.<fig id="f5"><label>Figure 4-5</label><caption><p>Use of a syringe to administer liquid medication into the oral cavity of a cat.</p></caption><graphic xlink:href="gr5"></graphic></fig></p><p id="para203">Spoons are ineffective because they measure fluids inaccurately and materials spill easily. A disposable syringe can be used to measure and administer liquids per os. Depending on the liquid administered, disposable syringes can be reused several times, assuming they are rinsed following each administration. In addition, disposable syringes can be dispensed legally to clients for home administration of liquid mediation. Mixing of medications in the same syringe is <italic>not</italic> recommended. However, dispensing of a separate, clearly marked syringe for each type of liquid medication prescribed for home administration is recommended.</p></sec><sec id="cesec11"><title>With an administration tube</title><p id="para204">Administration of medications, contrast material, and rehydrating fluids can be accomplished with the use of a feeding tube passed through the nostrils into the stomach or distal esophagus. Today, the general recommendation is to <italic>avoid</italic> passing the tip of a feeding tube beyond the distal esophagus. This is particularly true when a feeding tube is placed for long-term and repeated use (described in Gastrointestinal Procedures in this section). The reason for recommending nasoesophageal intubation <italic>over</italic> nasogastric intubation is based on the additional risk of irritation and even ulceration of the esophageal mucosa at the level of the cardia. Reflex peristalsis of the esophagus against a tube passing through the cardia has resulted in significant mucosal ulceration within 72 hours when feeding tubes were left in place. In patients receiving a single dose of medication or contrast material, nasogastric intubation is likely to be as safe as nasoesophageal intubation.</p><p id="para205">The narrow lumen of tubes passed through the nostril of small dogs and cats limits the viscosity of solutions that can be administered through a tube directly into the gastrointestinal tract. Nasoesophageal intubation can be done with a variety of tube types and sizes (<xref rid="cetable1" ref-type="table">Table 4-1</xref>). Newer polyurethane tubes, when coated with a lidocaine lubricating jelly, are nonirritating and may be left in place with the tip at the level of the distal esophagus. When placing the nasogastric tube, instill 4 to 5 drops of 0.5% proparacaine in the nostril of the cat or small dog; 0.5 to 1.0 mL of 2% lidocaine instilled into the nostril of a larger breed dog may be required to achieve the level of topical anesthesia needed to pass a tube through the nostril. With the head elevated, direct the tube dorsomedially toward the alar fold (<xref rid="f6" ref-type="fig">Figure 4-6</xref>). After inserting the tip 1 to 2 cm into the nostril, continue to advance the tube until it reaches the desired length. If the turbinates obstruct the passage of the tube, withdraw the tube by a few centimeters. Then readvance the tube, taking care to direct the tube ventrally through the nasal cavity. Occasionally, it will be necessary to withdraw the tube completely from the nostril and repeat the procedure. In particularly small patients or patients with obstructive lesions (e.g., tumor) in the nasal cavity, it may not be possible to pass a tube. Do not force the tube against significant resistance through the nostril.<table-wrap position="float" id="cetable1"><label>TABLE 4-1</label><caption><p>The French Catheter Scale Equivalents<xref rid="cetablefn1" ref-type="table-fn">*</xref></p></caption><table frame="hsides" rules="groups"><thead><tr><th></th><th colspan="2" align="center">Size<hr></hr></th></tr><tr><th align="center">Scale</th><th align="center">(mm)</th><th align="center">(inches)</th></tr></thead><tbody><tr><td align="center">3</td><td align="center">1</td><td align="char">0.039</td></tr><tr><td align="center">4</td><td align="char">1.35</td><td align="char">0.053</td></tr><tr><td align="center">5</td><td align="char">1.67</td><td align="char">0.066</td></tr><tr><td align="center">6</td><td align="center">2</td><td align="char">0.079</td></tr><tr><td align="center">7</td><td align="char">2.3</td><td align="char">0.092</td></tr><tr><td align="center">8</td><td align="char">2.7</td><td align="char">0.105</td></tr><tr><td align="center">9</td><td align="center">3</td><td align="char">0.118</td></tr><tr><td align="center">10</td><td align="char">3.3</td><td align="char">0.131</td></tr><tr><td align="center">11</td><td align="char">3.7</td><td align="char">0.144</td></tr><tr><td align="center">12</td><td align="center">4</td><td align="char">0.158</td></tr><tr><td align="center">13</td><td align="char">4.3</td><td align="char">0.170</td></tr><tr><td align="center">14</td><td align="char">4.7</td><td align="char">0.184</td></tr><tr><td align="center">15</td><td align="center">5</td><td align="char">0.197</td></tr><tr><td align="center">16</td><td align="char">5.3</td><td align="char">0.210</td></tr><tr><td align="center">17</td><td align="char">5.7</td><td align="char">0.223</td></tr><tr><td align="center">18</td><td align="center">6</td><td align="char">0.236</td></tr><tr><td align="center">19</td><td align="char">6.3</td><td align="char">0.249</td></tr><tr><td align="center">20</td><td align="char">6.7</td><td align="char">0.263</td></tr><tr><td align="center">22</td><td align="char">7.3</td><td align="char">0.288</td></tr><tr><td align="center">24</td><td align="center">8</td><td align="char">0.315</td></tr><tr><td align="center">26</td><td align="char">8.7</td><td align="char">0.341</td></tr><tr><td align="center">28</td><td align="char">9.3</td><td align="char">0.367</td></tr><tr><td align="center">30</td><td align="center">10</td><td align="char">0.393</td></tr><tr><td align="center">32</td><td align="char">10.7</td><td align="char">0.419</td></tr><tr><td align="center">34</td><td align="char">11.3</td><td align="char">0.445</td></tr></tbody></table><table-wrap-foot><fn id="cetablefn1"><label>*</label><p id="cenotep1">Mutiple types of pediatric polyurethane nasogastric feeding tubes are available in sizes ranging from 8F to 12F that easily accommodate administration of liquids medications and fluids to kittens, cats, and small dogs.</p></fn></table-wrap-foot></table-wrap><fig id="f6"><label>Figure 4-6</label><caption><p>Initial dorsomedial placement of a nasoesophageal tube before complete insertion.</p></caption><graphic xlink:href="gr6"></graphic></fig></p><p id="para206">CAUTION: The tip of the tube possibly can be introduced inadvertently through the glottis and into the trachea. Topical anesthetic instilled into the nose can anesthetize the arytenoid cartilages, thereby blocking a cough or gag reflex. I prefer to check the tube placement with a dry, empty syringe. Attach the test syringe to the end of the feeding tube. Rather than inject air or water in an attempt to auscultate borborygmus over the abdomen, attempt simply to aspirate air from the feeding tube. IF THERE IS NO RESISTANCE DURING ASPIRATION AND AIR FILLS THE SYRINGE, THE TUBE LIKELY HAS BEEN PLACED IN THE TRACHEA. Completely remove the tube and repeat the procedure. However, if repeated attempts to aspirate are met with immediate resistance and NO AIR ENTERS THE SYRINGE, the tube tip is positioned properly within the esophagus. If there is any question regarding placement, a lateral survey radiograph is indicated.</p><p id="para207">Gavage, or gastric lavage/feeding, in puppies and kittens can be accomplished by passing a soft rubber catheter or feeding tube through the nose and into the stomach. A 12F catheter is of an adequate diameter to pass freely, but it is too large for dogs and cats less than 2 to 3 weeks of age. Mark the tube with tape or a pen at a point equal to the distance from the tip of the nose to the last rib. Merely push the tube into the pharynx and down the esophagus to the caudal thoracic level (into the stomach). Attach a syringe to the flared end, and slowly inject medication or food. Use the same dry syringe aspiration technique to ensure that the tube is positioned in the esophagus/stomach rather than the trachea before administration.</p><p id="para208">A less desirable but effective technique for one-time tube administration of medications, food, or fluids entails passing the administration tube directly through the oral cavity and into the esophagus or stomach. However, this technique requires the use of a speculum to ensure that the patient does not bite or sever the tube with its teeth. A variety of speculums are available, ranging from hard rubber bite-blocks with a centrally positioned hole for passing the tube to improvised speculums such as a roll of 1- to 2-inch adhesive tape positioned between the mandible and maxilla. A well lubricated 22F rubber catheter, up to 30 inches long, is an ideal tube. Attach the catheter to a syringe that delivers the medication.</p><p id="para209">When the patient swallows, advance the catheter into the esophagus to the level of the eighth or ninth rib. Measure this distance on the tube first, and mark it with a ballpoint pen or a piece of tape. To pass the tube into the trachea in a conscious dog with its head held in a normal position is almost impossible. It may be possible to palpate the neck to feel the tube in the esophagus.</p><p id="para210">Nasoesophageal intubation in cats is generally much better tolerated that orogastric intubation. The cat can be restrained in a bag or cat stocks or by rolling it in a blanket. The cat is held in a vertical position by an assistant. Position a mouth speculum between the mandible and maxilla. This is where the fun begins. The operator then grasps the cat's head, as for pilling, and quickly passes the prelubricated tube 6 to 10 inches down the esophagus. A 12F to 16F soft rubber catheter, 16 inches long, makes a suitable tube.</p><p id="para211">Depending on the feeding tube type, the end of the tube may or may not accommodate a syringe. For example, soft, rubber urinary catheters are excellent tubes for single administration use. However, the flared end may not accommodate a syringe. To affix a syringe to the outside end of a tapered feeding tube or catheter, insert a plastic adapter (<xref rid="f7" ref-type="fig">Figure 4-7</xref>) into the open end of the tube.<fig id="f7"><label>Figure 4-7</label><caption><p>Use of a plastic adaptor (“Christmas tree”) to affix a syringe to a nasoesophageal feeding tube.</p></caption><graphic xlink:href="gr7"></graphic></fig></p></sec></sec><sec id="cesec12"><title>Topical Adminstration</title><sec id="cesec13"><title>Ocular</title><p id="para212">There are numerous ways to apply medication to the eyes, including the use of drops, ointments, subconjunctival injections, and subpalpebral lavage. The route and frequency of medication depend on the disease being treated.</p><p id="para213">If more than 2 drops of aqueous material are administered, the fluid will wash out of the conjunctival cul-de-sac and be wasted. Most drops should be applied every 2 hours (or less) to maintain effect. Ointments should be applied sparingly, and their effect may last a maximum of 4 to 6 hours.</p><p id="para214">Place drops on the inner canthus <italic>without touching the eye with the dropper tip.</italic> Place ointment (1/8-inch-long strip) on the upper sclera or lower palpebral border so that as the lids close, they form a film across the cornea.</p></sec><sec id="cesec14"><title>Otic</title><p id="para215">Medicated powders generally are contraindicated in the external ear canal. Thin films of ointments or propylene glycol solutions are more effective vehicles and are recommended. A few drops generally suffice, and the ear should be massaged gently after instillation to spread the medication over the external ear canal.</p></sec><sec id="cesec15"><title>Nasal</title><p id="para216">Isotonic aqueous drops are used for nasal application and should be applied without touching the dropper to the nose. Oily drops are not advised because they may damage the nasal mucosa or may be inhaled. There is little indication for routine instillation of medication into the nostrils of dogs and cats.</p></sec><sec id="cesec16"><title>Dermatologic</title><p id="para217">Several objectives should be considered when treating dermatologic disorders: (1) eradication of causative agents; (2) alleviation of symptoms, such as reduction of inflammation; (3) cleansing and debridement; (4) protection; (5) restoration of hydration; and (6) reduction of scaling and callus. Many different forms of skin medications are available, but the vehicle in which they are applied is a critical factor (<xref rid="cetextbox1" ref-type="boxed-text">Box 4-1</xref>). In all cases, apply topical medications to a clean skin surface in a very thin film, because only the medication in contact with the skin is effective. In most cases, clipping hair from an affected area enhances the effect of medication.<boxed-text id="cetextbox1"><label>BOX 4-1</label><caption><title>VEHICLES USED IN THE ADMINISTRATION OF TOPICAL SKIN MEDICATIONS</title></caption><p id="para1"><italic>Lotions</italic> are suspensions of powder in water or alcohol. They are used for acute, eczematous lesions. Because they less easily are absorbed than creams and ointments, lotions need to be applied 2 to 6 times a day.</p><p id="para2"><italic>Pastes</italic> are mixtures of 20% to 50% powder in ointment. In general, they are thick, heavy, and difficult to use.</p><p id="para3"><italic>Creams</italic> are oil droplets dispersed in a continuous phase of water. Creams permit excellent percutaneous absorption of ingredients.</p><p id="para4"><italic>Ointments</italic> are water droplets dispersed in a continuous phase of oil. They are very good for dry, scaly eruptions.</p><p id="para5"><italic>Propylene glycol</italic> is a stable vehicle and spreads well. It allows good percutaneous absorption of added agents.</p><p id="para6"><italic>Adherent dressings</italic> are bases that dry quickly and stick to the lesion.</p><p id="para7"><italic>Shampoos</italic> are usually detergents designed to cleanse the skin. If shampoos are left in contact with the skin for a time, added medications may have specific antibacterial, antifungal, or antiparasitic effects.</p></boxed-text></p></sec><sec id="cesec17"><title>Note on compounding pharmacies</title><p id="para218">With the widespread availability of compounding pharmacies, prescribing compounded medications for topical and oral administration recently has become a popular dispensing technique for dogs and cats requiring long-term, daily mediation. Caution is warranted. Some compounding pharmacies that serve the veterinary profession are using inappropriate or ineffective vehicles in which the drug has been compounded, or the drug itself, purchased in bulk, is a lower grade and possibly an ineffective product once compounded. Studies on the quality and efficacy of compounded drugs for use in veterinary patients are limited. However, of those studies that have been performed, serious questions are being raised over the bioavailability of the drug administered.</p></sec></sec><sec id="cesec18"><title>Administration By Injection (Parenteral Administration)</title><p id="para219">Before aspirating medications from multiple-dose vials, carefully wipe the rubber diaphragm stopper with the same antiseptic used on the skin. Observe this basic rule with all medication vials, even with modified live virus vaccines.</p><p id="para220">It would be admirable to prepare the skin surgically before making needle punctures to administer medications. Because such preparation is not practical, carefully part the hair and apply a high-quality skin antiseptic such as benzalkonium chloride in 70% alcohol. Place the needle directly on the prepared area, and thrust the needle through the skin. Although the use of antiseptics on the vial and skin is not highly effective, the procedure removes gross contamination and projects an image of professionalism.</p></sec><sec id="cesec19"><title>Subcutaneous Injection</title><p id="para221">Dogs and cats have abundant loose alveolar tissue and easily can accommodate large volumes of material in this subcutaneous space. The dorsal neck is seldom used for subcutaneous injections because the skin is somewhat more sensitive, causing some patients to move abruptly during administration. A wide surface area of skin and subcutaneous tissue over the dorsum from the shoulders to the lumbar region makes an ideal site for subcutaneous injections.</p><p id="para222">Administration of drugs, vaccines, and fluids by the subcutaneous route represents the most commonly used route of parenteral administration in dogs and cats. For small volumes (<2 mL total), such as vaccines, a 22- to 25-gauge needle generally is used. The site most often used is the wide area of skin over the shoulders. The large subcutaneous space and the relative lack of sensitivity of skin at this location make it an ideal injection site. Cleaning of the skin with alcohol or other disinfectant generally is performed before injection. Several injection techniques are used. A common technique entails grasping a fold of skin with two fingers and the thumb of one hand. Gently lift the skin upward. Using the opposite hand, place the needle, with syringe attached, through the skin at a point below the opposite thumb. <italic>Aspiration before injection is not typically necessary when using this route of administration.</italic> Following administration and on removal of the needle from the skin, gently pinch the injection site and hold it for a few seconds to prevent backflow of medication or vaccine onto the skin.</p><p id="para223">When larger volumes are to be administered—fluids in dehydrated dogs and cats—the skin directly over the shoulders is the injection site most commonly selected. Generally, only isotonic fluids are administered by the subcutaneous route. Depending on the patient's size, needles ranging from 16 to 22 gauge can be used. Because of the larger volumes of fluid involved, warming of the fluids before administration is recommended. Doing so can enhance significantly the patient's tolerance for the displacement of skin during the period of administration. Depending on the rate of administration and breed of dog, relatively large volumes of fluid generally can be given in one location. Cats typically tolerate 10 to 20 mL/kg body mass in a single location. Large dogs can tolerate volumes greater than 200 mL of fluid in a single location. When administering large volumes, it is usually <italic>not</italic> necessary to use <italic>multiple</italic> injection sites for purposes of distributing the total fluid volume. Doing so actually may increase the risk of introducing cutaneous bacteria under the skin. Because the administration time required to deliver larger volumes is longer, and the injection needle will be placed in the skin for extended periods, it is appropriate to cleanse and rinse the skin carefully before actually inserting the needle. Isotonic, warmed fluids may be administered by large syringe or through an administration tube attached to a bag. Monitor skin tension and the patient's comfort tolerance throughout the procedure.</p><p id="para224">Although fluid absorption begins almost immediately on subcutaneous administration of fluids, significant pressure caused by the bolus of fluid delivered can develop within the fluid pocket. On removal of the needle, firmly grasp the injection site with the thumb and forefinger for several seconds. The procedure is <italic>not complete</italic> until one has verified that back-leakage of fluid from the subcutaneous space onto the skin is not occurring.</p><p id="para225"><boxed-text id="cetextbox23"><caption><title>Note:</title></caption><p id="para226">Not all parenteral medications can be administered safely by the subcutaneous route. When administering any compound by the subcutaneous route, verify that the product to be administered is approved for subcutaneous administration. Serious reactions, including abscess formation and tissue necrosis, can occur.</p></boxed-text></p><p id="para227">Depending on the patient's hydration status and physical condition, fluid absorption may take from 6 to 8 hours.</p><p id="para228">NOTE: The rate of absorption of fluid administered by the subcutaneous route largely depends on the patient's hydration state and vascular and cardiac integrity. For that reason, the subcutaneous route is not recommended to manage patients in hypovolemic shock. Exceptions to this do exist, for example, when in a life-or-death situation access to a vein is simply not possible. Subcutaneous or intraosseous (see the following discussion) fluid administration may be the only option available.</p><sec id="cesec20"><title>Implanted subcutaneous fluid ports</title><p id="para229">In clinical practice, it has become increasingly popular to dispense bags of sterile, isotonic fluids, with appropriate administration tubing and needles, to pet owners for home administration of subcutaneous fluids, such as for especially long-term management of chronic renal failure in cats. Although some owners are comfortable administering subcutaneous fluids through a needle, others are not. Recently, an implantable subcutaneous port<xref rid="fn1" ref-type="fn">*</xref>has been introduced for use in patients requiring regular administration of subcutaneous fluids at home. A 9-inch silicon tube is preplaced under the skin and is sutured in place by a veterinarian. Objectively, this offers easy access to the subcutaneous space without need for needle penetration. Owners simply attach a syringe or extension tube tip to the port and administer the appropriate volume of fluids at an appropriate rate and frequency. Because of the usual requirement for long-term placement of an implantable fluid administration tube, there is risk of infection under the skin and around the incision site. Some cats do not tolerate the device.</p></sec></sec><sec id="cesec21"><title>Intramuscular Injection</title><p id="para230">Because the tightly packed muscular tissue cannot expand and accommodate large volumes of injectables without trauma, medications given by this route should be small in volume. These medications are often depot materials that are poorly soluble, and some may be mildly irritating. Never give intramuscular injections in the neck because of the fibrous sheaths there and the complications that may occur. I also believe that injections in the hamstring muscles may cause severe pain, lameness, and occasionally peroneal paralysis because of local nerve involvement. Unless the animal is extremely thin, give injections into the lumbodorsal muscles on either side of the dorsal processes of the vertebral column.</p><p id="para231">After proper preparation of the skin, insert the needle through the skin at a slight angle (if the animal is thin) or at the perpendicular (if the animal is obese). When injecting any medication by a route other than the intravenous one, it is <italic>imperative</italic> to retract the plunger of the syringe before injecting to be certain that a vein was not entered by mistake. This is especially crucial with oil suspension, microcrystalline suspension, or potent-dose medications.</p></sec><sec id="cesec22"><title>Intradermal Injection</title><p id="para232">Intracutaneous (or intradermal) injections are used for testing purposes. Prepare the skin by carefully clipping the hair with a No. 40 clipper blade. If the skin surface is dirty, gently clean it with a moist towel. Scrubbing and disinfection are contraindicated because they may produce iatrogenic trauma and inflammation, which interfere with the test. Stretch the skin by lifting a fold, and use a 25- to 27-gauge intradermal needle attached to a 1-mL tuberculin syringe. Insert the point of the needle, bevel up, in a forward lifting motion as if to pick up the skin with the needle tip. Advance the needle while pushing the syringe (levered) downward until the bevel is completely within the skin. Inject a bleb of 0.05 to 0.10 mL of fluid. If the procedure is done correctly, the small bleb will appear translucent. Intradermal injections generally are used in patients subjected to intradermal skin testing for allergenic antigens. Administration of compounds by the intradermal technique is not necessarily simple. Inadvertent administration of medications into the subcutaneous tissues is easy when attempting intradermal injection. For that reason, specific training/experience is recommended before attempting intradermal skin testing of allergic patients.</p></sec><sec id="cesec23"><title>Transdermal (Needle-Free) Administration</title><p id="para233">Intradermal administration of vaccine and drugs in veterinary and human medicine largely has been limited to the complexities of accurately delivering the desired dose into, and not under, the skin. In 2004 a transdermal administration system<xref rid="fn2" ref-type="fn">†</xref>was introduced that was designed after a similar device used in human medicine. This system consistently delivers a precise volume of vaccine into the skin, subcutaneous tissues, and muscle of vaccinated cats. The advantage of delivering vaccine into the skin of animals is the enhanced processing of antigen by the abundant dendritic cells. In addition to using this delivery system for other vaccines, potential application exists for other medications, such as precise delivery of very small quantities of insulin to cats.</p></sec><sec id="cesec24"><title>Intravenous Injection</title><sec id="cesec25"><title>Cephalic venipuncture</title><p id="para234">To restrain a dog or cat for venipuncture of the cephalic vein, place the dog or cat on the table in sternal recumbency. If the right vein is to be tapped or catheterized, the assistant should stand on the left side of the animal and place the left arm or hand under the animal's chin to immobilize the head and neck. The assistant should reach across the animal and grasp the leg just behind and distal to the right elbow joint. The assistant should use the thumb to occlude and rotate the cephalic vein laterally while the palm of the hand holds the elbow in an immobilized and extended position. Make sure that the animal stays on the table if struggling occurs. The person performing the venipuncture then grasps the leg at the metacarpal region and begins the venipuncture on the medial aspect of the leg, just adjacent to the cephalic vein proximal to the carpus.</p></sec><sec id="cesec26"><title>Jugular venipuncture</title><p id="para235">For a jugular venipuncture in the dog, place the patient in sternal recumbancy, with the hands of the assistant placed around the patient's muzzle to extend the neck and nose dorsally toward the ceiling. In short-coated dogs, the jugular vein usually can be seen coursing from the ramus of the mandible to the thoracic inlet in the jugular furrow. The vessel may be more difficult to visualize in dogs with long-haired coats or if excessive subcutaneous fat or skin is present. The person performing the venipuncture should place the thumb of the nondominant hand across the jugular vein in the thoracic inlet or proximal to the thoracic inlet to occlude venous drainage from the vessel and allow it to fill. With the dominant hand, the person performing the venipuncture should insert the needle and syringe or Vacutainer (BD, Franklin Lakes, New Jersey) into the vessel at a 15- to 30-degree angle to perform the venipuncture.</p><p id="para236">For smaller and very large animals, the jugular vein also can be tapped by placing the patient in lateral recumbancy. The assistant should pull the animal's front legs caudally and extend the head and neck so that the jugular vein can be visualized. The venipuncture then can be performed as previously described. A jugular venipuncture is contraindicated in patients with thrombocytopenia or vitamin K antagonist rodenticide intoxication.</p><p id="para237">Place cats in sternal recumbancy. The assistant should stand behind the patient so that the patient cannot back away from the needle during the venipuncture. The assistant should extend the cat's head and neck dorsally while restraining the cat's front legs with the other hand. The cat's fur can be clipped or moistened with isopropyl alcohol to aid in visualization of the jugular vein as it stands up in the jugular furrow. The person performing the venipuncture should occlude the vessel at the thoracic inlet and insert the needle or Vacutainer apparatus into the vessel as previously described to withdraw the blood sample. Alternately, place the cat in lateral recumbancy as described in the previous paragraph.</p></sec><sec id="cesec27"><title>Lateral saphenous venipuncture</title><p id="para238">To perform a lateral saphenous venipuncture, place the patient in lateral recumbancy. The lateral saphenous vein can be visualized on the lateral portion of the stifle, just proximal to the tarsus. The assistant should extend the hind limb and occlude the lateral saphenous vein just proximal and caudal to the tarsus. The person performing the venipuncture should grasp the distal portion of the patient's limb with the nondominant hand and insert the needle or Vacutainer apparatus with the dominant hand to withdraw the blood sample.</p></sec><sec id="cesec28"><title>Medial saphenous venipuncture</title><p id="para239">To perform a medial saphenous venipuncture, place the patient in lateral recumbancy. Move the top hind limb cranially or caudally to allow visualization of the medial saphenous vein on the medial aspect of the tibia and fibula. The assistant should scruff the patient, if the patient is small, or should place the forearm over the patient's neck to prevent the patient from getting up during the procedure. With the other hand, the assistant should occlude the medial saphenous vein in the inguinal region. The person performing the medial saphenous venipuncture should grasp the paw or hock of the limb and pull the skin taught to prevent the vessel from rolling away from the needle. The fur may be clipped or moistened with isopropyl alcohol to aid in visualization of the vessel. The needle or Vacutainer apparatus can be inserted into the vessel at a 15- to 30-degree angle to withdraw the blood sample.</p></sec></sec><sec id="cesec29"><title>Intraosseous Administration</title><p id="para240">Intraosseous infusion of blood, fluids, or medications is useful whenever rapid, direct access to the circulatory system is required and peripheral or central access is impossible or too time-consuming. This technique can be set up rapidly (3 minutes), is certain, and is especially useful for unusually small patients, especially kittens and puppies. (NOTE: This procedure also is described in Section 1.)</p><p id="para241">Intraosseous infusion is particularly indicated in shock or circulatory collapse syndromes, edematous states, severe burns, and obesity, and when peripheral veins are thrombosed. This method is contraindicated in birds (because their bones contain air), for infusion into fractured bones, or in cases of sepsis, because osteomyelitis may develop.</p><p id="para242">Substances injected into the bone marrow reach the general circulation at about the same rate as those injected directly into peripheral veins. Blood and blood components and solutions of colloids, crystalloids, electrolytes, drugs, and nutrients can be given—even in large volumes.</p><sec id="cesec30"><title>Technique</title><p id="para243">The two easiest and most desirable sites for marrow access are (1) the flat medial side of the proximal tibia but distal to the tibial tuberosity and the proximal growth plate and (2) the trochanteric fossa of the proximal femur.</p><p id="para244">To perform intraosseous administration, follow this procedure:<list list-type="simple" id="celist15"><list-item id="celistitem81"><label>1.</label><p id="para245">Prepare the skin site aseptically, and inject 1% lidocaine into the skin and periosteum.</p></list-item><list-item id="celistitem82"><label>2.</label><p id="para246">Stabilize the leg, and make a small stab incision through the skin. Needles of 18- to 20-gauge are preferred and can be ordinary hypodermic needles (short bevel desired) or special stylet needle sets, such as a spinal needle or an Illinois bone marrow needle (see <xref rid="f10" ref-type="fig">Fig. 4-10</xref>). A needle with a stylet is preferred so that the needle is not occluded with cortical bone or marrow during introduction.</p></list-item><list-item id="celistitem83"><label>3.</label><p id="para247">Point the needle slightly distally and rotate with firm pressure until it enters the near cortex. A properly seated needle will feel stable and firm. Use a 10-mL syringe to aspirate marrow, fat, and bony debris. Prefill the needle before administering fluids.</p></list-item><list-item id="celistitem84"><label>4.</label><p id="para248">Attach a regular fluid infusion set, and start fluid administration. The rate should not exceed 11 mL/minute by gravity or 24 mL/minute with pressure up to 300 mm Hg. Gravity flow through a single catheter may be adequate for patients up to 16 lb. For larger animals, multiple catheters in separate bones or pressurized flow, or both, may be needed for rapid infusions.</p></list-item><list-item id="celistitem85"><label>5.</label><p id="para249">Encase the needle hub in a butterfly tape, and suture the tape in place. Place antibiotic ointment around the skin incision, and protect and immobilize the whole apparatus with a bulky bandage wrap.</p></list-item><list-item id="celistitem86"><label>6.</label><p id="para250">Manage intraosseous catheters in the same way as intravenous catheters. Flush the catheter every 6 hours with heparinized saline, and place the catheter in a new bone every 72 hours. The same bone can be reused at another location if 25 to 36 hours is allowed for occlusion and healing of the original site.</p></list-item></list></p></sec><sec id="cesec31"><title>Complications</title><p id="para251">Infection is the primary concern. Fat embolism and damage to the growth plates are other concerns. Extravasation of fluid from the bone marrow into the subcutaneous tissue may occur if the needle punctures both cortexes or if more than one hole is made in the cortex. In such cases, remove the needle and select another bone.</p><sec id="cesec32"><sec id="cesec33"><title>Additional Reading</title><p id="para252">Crow S, Walshaw S: <italic>Manual of clinical procedures in the dog, cat, and rabbit,</italic> ed 2, Philadelphia, 1997, Lippincott-Raven.</p><p id="para253">Kirby R, Rudloff E: Crystalloid and colloid fluid therapy. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p><p id="para254">Marks S: The principles and practical application of enteral nutrition, <italic>Vet Clin North Am Small Anim Pract</italic> 28:677, 1998.</p><p id="para255">Wingfield WE: <italic>Veterinary emergency medicine secrets,</italic> Philadelphia, 1997, Hanley & Belfus.</p></sec></sec></sec></sec></sec><sec id="cesec34"><title>BANDAGING TECHNIQUES (SEE SECTION 1)</title><p id="para256"> </p></sec><sec id="cesec35"><title>BLOOD PRESSURE MEASUREMENT: INDIRECT</title><p id="para257">Indirect measurement of blood pressure (BP) in dogs and cats is a convenient, noninvasive technique for establishing whether an individual patient's BP is increased (systemic hypertension) or decreased (hypotension). Today, multiple techniques are available; none are perfect. In human medicine, BP measurement is performed routinely and is (relatively) reliable. In veterinary medicine, BP measurement typically is reserved for patient's determined to have diseases most likely to be associated with serious, potentially injurious alterations in BP, such as shock (hypotension) or chronic renal failure (hypertension). Also in veterinary medicine, it is important to note that most of the BP measuring equipment is designed to provide maximum sensitivity in <italic>hypertensive</italic> patients. Sensitivity of the equipment for accurately detecting hypotension is low.</p><p id="para258">Generally, two techniques are used. Oscillometric BP measurement entails use of an automated recording system. A cuff is applied to the base of the tail or a distal limb for access to an artery. This technique generally is regarded as being most accurate in dogs. When oscillometric BP measurements are performed in dogs, the patient should be in lateral recumbency. This places the cuff at approximately the same level as the heart. In cats the patient generally remains in sternal recumbency (and minimally restrained). Most patients experience a brief acclimation period to the cuff placement. For this reason, at least 3 to 5 separate readings are obtained at 1- to 2-minute intervals. This technique can be used on awake or anesthetized patients (<xref rid="f8" ref-type="fig">Figure 4-8</xref>).<fig id="f8"><label>Figure 4-8</label><caption><p>Oscillometric blood pressure measurement in a cat.</p></caption><graphic xlink:href="gr8"></graphic></fig></p><p id="para259">The Doppler-ultrasonic flow detection system is most accurate in cats for measuring systolic BP. Again, the ventral tail base or a dorsal pedal artery (hind limb) or the superficial palmar arterial arch (forelimb) can be used. Apply and inflate an occluding cuff. The readings are obtained by a transducer as the pressure on the cuff is reduced. Caution is recommended in interpreting results from dogs that are reported as hypertensive but have no overt clinical disease. The higher reported occurrence of falsely elevated BP in normotensive dogs measured by this method justifies the additional scrutiny when interpreting Doppler BP results in dogs.</p><p id="para260">Clinically, the most common use of indirect BP measurement is in assessing cats for the presence (or absence) of systemic hypertension caused by renal insufficiency or hyperthyroidism (thyrotoxicosis). A common finding among untreated hypertensive cats is retinal detachment and blindness. Early detection and therapeutic intervention (e.g., enalapril and or amlodipine) is critical. In dogs, BP measurement is indicated in patients with chronic renal insufficiency and/or protein-losing nephropathy, hyperadrenocorticism, and diabetes mellitus. In veterinary medicine, interpretation of BP centers on the <italic>systolic BP reading,</italic> not the diastolic reading (<xref rid="cetable2" ref-type="table">Table 4-2</xref>).<table-wrap position="float" id="cetable2"><label>TABLE 4-2</label><caption><p>Systolic Blood Pressure</p></caption><table frame="hsides" rules="groups"><thead><tr><th></th><th align="left">Normal</th><th align="left">Hypertension</th><th align="left">Hypotension</th></tr></thead><tbody><tr><td align="left" rowspan="2">Dog and cat</td><td align="left" rowspan="2">100–150 mm Hg</td><td align="left">>160 mm Hg</td><td align="left" rowspan="2"><100 mm Hg</td></tr><tr><td align="left">>180 mm Hg (high risk)</td></tr></tbody></table></table-wrap></p><sec id="cesec36"><sec id="cesec37"><sec id="cesec38"><sec id="cesec39"><title>Additional Reading</title><p id="para261">Stepien RL: Blood pressure assessment. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p><p id="para262">Stepien RL: Diagnostic blood pressure measurement. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p></sec></sec></sec></sec></sec><sec id="cesec40"><title>CENTRAL VENOUS PRESSURE MEASUREMENT</title><p id="para263">Central venous pressure (CVP) is the blood pressure within the intrathoracic portions of the cranial or caudal vena cava. Measurement of CVP in the dog provides an excellent index for determining circulation efficiency. The CVP is controlled by interaction of the circulating blood volume, cardiac pumping action, and alterations in the vascular bed. The CVP is not a measure of blood volume but an indication of the ability of the heart to accept and pump blood brought to it. The CVP reflects the interaction of the heart, vascular tone, and circulatory blood volume. When the heart action and vascular tone remain constant, CVP reflects blood volume. When blood volume and vascular tone are constant, CVP reflects heart action. When blood volume and heart action are constant, CVP can be used to measure vascular tone.</p><p id="para264">In addition, the placement of a jugular catheter can be helpful in long-term fluid management and in parenteral alimentation of critically ill animals.</p><p id="para265">Measurement of CVP is indicated (1) in acute circulatory failure that has not responded to initial treatment; (2) in administration of large volumes of blood or fluids, as may occur in acute shock; (3) as part of the monitoring procedure in poor-risk surgical patients; and (4) in patients with reduced urinary output for which fluids are being administered (e.g., acute renal failure).</p><p id="para266">For CVP measurement, a catheter must be placed in the external jugular vein such that the catheter is in direct fluid continuity with the right atrium (see Percutaneous Jugular Vein Catheterization). Place the patient in lateral recumbency, and clip the hair over the jugular vein. Surgically prepare the skin in the clipped area.</p><p id="para267">Make a percutaneous puncture of the jugular vein with the Intracath catheter needle, and advance the tip to approximately the third intercostal space (tip of the catheter at the right atrium). Fasten the catheter securely to the neck of the patient by passing adhesive tape around the neck and the hub of the catheter needle so that the hub of the needle comes to lie at the base of the ear. Connect a three-way stopcock to the catheter. Connect an intravenous setup of isotonic sodium chloride to one end of the stopcock, and to the other end of the stopcock attach a piece of intravenous tubing, which should be taped vertically to a pole or a piece of doweling (<xref rid="f9" ref-type="fig">Figure 4-9</xref>). The metric rule is placed so that the 0 level is aligned with the midpoint of the trachea at the thoracic inlet, and the rule is taped to the vertical pole.<fig id="f9"><label>Figure 4-9</label><caption><p>Central venous manometer. <bold>A,</bold> Standard intravenous infusion tube. <bold>B,</bold> Central venous pressure level. <bold>C,</bold> Thirty-inch intravenous extension tube. <bold>D,</bold> Centimeter scale. <bold>E,</bold> Plastic tube in great veins in thorax or right atrium via jugular vein. <bold>F,</bold> Three-way stopcock set in measuring position (open from manometer to catheter). Note: This procedure should be performed with the dog in right lateral recumbency.</p></caption><graphic xlink:href="gr9"></graphic><attrib>(From Slatter FP: Shock. In Kirk RW, editor: <italic>Current veterinary therapy III,</italic> Philadelphia, 1968, WB Saunders.)</attrib></fig></p><p id="para268">To fill the CVP manometer, turn the three-way stopcock so that fluid will flow from the bottle of saline into the manometer and will exceed the 15-cm mark. Next, turn the stopcock so that a column of fluid exists from the superior vena cava to the manometer. The fluid in the manometer will fall until it reflects the level of the CVP.</p><p id="para269">It is desirable to allow fluid to flow frequently through the catheter so that the catheter tip does not become plugged with a blood clot. Periodic flushing with heparinized saline will help maintain the patency of the catheter. This setup allows easy intravenous administration of fluids and medication to the patient and collection of blood, if necessary.</p><p id="para270">There is no absolute value for a normal CVP. The CVP for the normal dog is −1 to +5 cm H<sub>2</sub>O. Elevations of +5 to +10 cm H<sub>2</sub>O are borderline; however, values greater than 10 cm H<sub>2</sub>O may indicate an abnormally expanded blood volume, and those greater than 15 cm H<sub>2</sub>O may indicate congestive heart failure. The trend of the CVP is what should be monitored and correlated with the regimen of treatment. One must be aware constantly of the interrelationship between blood volume, cardiovascular function, and vascular tone. If the CVP is at levels of 10 to 15 cm H<sub>2</sub>O, the pulmonary venous pressure is approaching 20 to 22 mm Hg, and additional intravenous fluids should not be administered.</p><sec id="cesec41"><sec id="cesec42"><sec id="cesec43"><sec id="cesec44"><title>Additional Reading</title><p id="para271">Haskins SC: Monitoring the critically ill patient, <italic>Vet Clin North Am Small Anim Pract</italic> 19:1059-1078, 1989.</p><p id="para272">Wingfield WE: <italic>Veterinary emergency medicine secrets,</italic> Philadelphia, 1997, Hanley & Belfus.</p></sec></sec></sec></sec></sec><sec id="cesec45"><title>DIAGNOSTIC SAMPLE COLLECTION TECHNIQUES</title><sec id="cesec46"><title>Bacterial Culture</title><p id="para273">Before actually collecting and submitting a sample for bacterial culture, it is appropriate (whenever feasible to do so) to prepare, stain, and examine a direct smear of the suspect material or tissue. After collecting material on a sterile cotton swab, roll the specimen onto a clear glass slide and allow it to dry completely. Staining with a rapid Romanowsky's-type stain (e.g., Diff-Quik stain) may reveal evidence of neutrophilic inflammation (neutrophilia, especially with a left shift) and occasionally degenerative neutrophils with intracellular bacteria visible. These findings greatly facilitate patient management by documenting the immediate need for interventive empiric antimicrobial therapy until definitive culture and antimicrobial susceptibility results are obtained. The absence of cytologic evidence of bacterial infection does <italic>not</italic> rule out the possibility that the patient is bacteremic.</p><sec id="cesec47"><title>Routine culture</title><p id="para274">Inoculate material for culture on blood agar plates or in cystine lactose-electrolyte-deficient (CLED) medium as an acceptable alternative. The CLED medium stimulates growth, detects lactose fermentation, and prevents spreading of <italic>Proteus.</italic> The CLED medium serves as a basis for the isolation of most aerobic microorganisms. Selective media may be necessary for the isolation and identification of specific microorganisms. Biopsy material may be ground in sterile sand and placed in sterile broth.</p></sec><sec id="cesec48"><title>Multiple-media plates</title><p id="para275">Multiple-media plates have been developed commercially to facilitate direct antibiotic sensitivity and tentative identification of common pathogenic bacteria. These prepackaged, relatively inexpensive plates help the small laboratory identify pathogenic bacteria by their characteristic behavior on selective media. Some companies have different kits for different suspected infections. In general, kits are most useful for evaluating conjunctivitis, otitis, pyoderma, wound infections, uterine or anterior vaginal infections, fresh necropsy material, and urinary tract infections. Multiple-media plates are not recommended for culturing areas that have a large population of normal microbial organisms (such as the respiratory tract, throat, and vulva), for fecal samples, or for blood cultures to determine bacteremia (<xref rid="cetable3" ref-type="table">Table 4-3</xref>).<table-wrap position="float" id="cetable3"><label>TABLE 4-3</label><caption><p>Common Bacterial Culture Results</p></caption><table frame="hsides" rules="groups"><thead><tr><th align="left">Site</th><th align="left">Commensals</th><th align="left">Pathogens</th></tr></thead><tbody><tr><td colspan="3" align="left"><bold><italic>External ear canal</italic></bold></td></tr><tr><td align="left">Dog</td><td align="left"><italic>Malassezia, Clostridium, Staphylococcus</italic> (a few), <italic>Bacillus</italic> (a few); never <italic>Streptococcus, Pseudomonas,</italic> or <italic>Proteus</italic></td><td align="left">Many <italic>Staphylococcus</italic> and <italic>Malassezia</italic> together; <italic>Pseudomonas, Proteus, Streptococcus, Escherichia coli</italic></td></tr><tr><td align="left">Cat</td><td align="left">Not documented</td><td align="left"><italic>Staphylococcus aureus,</italic> β-hemolytic streptococcus, <italic>Pasteurella, Pseudomonas, Proteus, E. coli, Malassezia</italic></td></tr><tr><td colspan="3" align="left"><bold><italic>Skin</italic></bold></td></tr><tr><td align="left">Dog</td><td align="left"><italic>Micrococcus, Clostridium,</italic> diphtheroids, <italic>Staphylococcus epidermidis, Corynebacterium, Malassezia</italic></td><td align="left"><italic>S. aureus</italic> (coagulase positive), <italic>Proteus, Pseudomonas, E. coli</italic></td></tr><tr><td align="left">Cat</td><td align="left"><italic>Micrococcus, Streptococcus, S. aureus, S. epidermidis</italic></td><td align="left"><italic>S. aureus, Pasteurella multocida, Bacteroides, Fusobacterium,</italic> haemolytic streptococci</td></tr><tr><td align="left"><bold><italic>Conjunctiva</italic></bold></td><td align="left"><italic>Staphylococcus, Streptococcus, Bacillus, Corynebacterium,</italic> diphtheroids, <italic>Neisseria, Pseudomonas</italic></td><td align="left"><italic>S. aureus, Bacillus, Pseudomonas, E. coli, Aspergillus</italic></td></tr><tr><td align="left"><bold><italic>Vagina</italic></bold></td><td align="left"><italic>Staphylococcus, Streptococcus, Enterococcus, Corynebacterium, E. coli, Haemophilus, Pseudomonas, Peptostreptococcus, Bacteroides</italic></td><td align="left"><italic>Brucella canis;</italic> pure culture of organisum (esp. <italic>E. coli, Staphylococcus, Pseudomonas)</italic> when accompanied by tissue reaction at vaginal cytology</td></tr><tr><td align="left"><bold><italic>Urine</italic></bold></td><td align="left"><1000<xref rid="cetablefn2" ref-type="table-fn">*</xref> organisms/mL; presence of several organisms suggests contamination</td><td align="left">More than 100,000<xref rid="cetablefn2" ref-type="table-fn">*</xref> organisms/mL and often pure culture. <italic>E.coli,</italic> enterobacteria, <italic>klebsiella, Proteus, Pseudomonas aeruginosa, Pasteurella multocida, Staphylococcus, Streptococcus</italic></td></tr></tbody></table><table-wrap-foot><fn id="cetablefn2"><label>*</label><p id="cenotep2">Absolute numbers of bacteria depend on the collection technique.</p></fn></table-wrap-foot></table-wrap></p></sec><sec id="cesec49"><title>Direct smears</title><p id="para276">Cell scrapings taken from conjunctiva during the phase of inflammation (first 10 to 14 days) and stained with Giemsa stain may show typical intracytoplasmic inclusions of initial and elementary bodies accompanied by a polymorphonuclear inflammatory cellular reaction.</p></sec><sec id="cesec50"><title>Transport of samples</title><p id="para277">Because most diagnostic specimens collected for bacterial culture are submitted to commercial laboratories for bacterial isolation, identification, and antimicrobial susceptibility testing, it is important to prepare the sample properly for shipping.</p><p id="para278">No special transport media are required for routine aerobic culture specimens as long as the sample can remain moist and relatively cool <italic>and</italic> the sample can be inoculated onto culture medium within 3 to 4 hours only. For samples that must be shipped overnight to a laboratory, it is imperative that the specimen be kept cool (not frozen) and moist. Elevated temperatures during shipping contribute to bacterial overgrowth of nonpathogenic bacteria, making isolation and identification of disease-producing organisms difficult. Special transport media may be required. Contact the individual laboratory regarding information pertaining to shipping of specimens for bacterial culture.</p><p id="para279">Specimens submitted for anaerobic culture need to be inoculated onto culture media within minutes following collection. Although special anaerobic transport media are available, they may not be well suited for extended shipping times (>24 hours).</p></sec><sec id="cesec51"><title>Isolation and identification</title><p id="para280">For isolation, obtain columnar epithelial cells (not exudate). Use calcium alginate (not wooden) swabs. Place swabs directly into liquid-holding medium on wet ice. The most commonly used transport medium is 2-SP, composed of 0.2M sucrose and 0.02M phosphate (pH is 7.2) with added antibiotics. This can be supplied by the laboratory that is doing the isolations. Monolayers of McCoy and HeLa cells are best for isolation of <italic>Chlamydophila</italic> spp. Egg (yolk sac) inoculation of embryonated eggs has been abandoned. <italic>Chlamydophila felis</italic> (formerly <italic>Chlamydia psittaci</italic>) inclusions are detected by fluorescent antibody techniques.</p></sec><sec id="cesec52"><title>Puncture fluids</title><p id="para281">Aspirate material using aseptic technique. Centrifuge the aspirated material at high speed, and stain a smear of the sediment with Gram stain. Culture the sediment on blood agar, in thioglycolate medium, on Sabouraud dextrose agar, or on one of the multiple-media plates. Also consider anaerobic cultures.</p></sec><sec id="cesec53"><title>Wounds and ulcers</title><p id="para282">In dealing with an abscess (except those of the eye), clip and clean the abscess site. Aspirate material from the abscess into a sterile syringe and culture in blood agar and thioglycolate broth or on one of the multiple-media plates. In open wounds, use a sterile cotton swab and obtain fresh exudate from the deeper portion of the lesion. Also consider anaerobic cultures.</p></sec><sec id="cesec54"><title>Spinal fluid</title><p id="para283">If the spinal fluid is cloudy, make a direct smear and stain with Gram and Giemsa stains. If the fluid is fairly clear, centrifuge for 10 minutes, make a smear, and stain the sediment with Gram stain. Make cultures of the sediment on blood agar, in thioglycolate medium, or on one of the multiple-media plates, and on Sabouraud dextrose agar.</p></sec><sec id="cesec55"><title>Ear cultures</title><p id="para284">Collect material on sterile cotton swabs, make a smear, and stain it with Gram stain. Place the swab on blood agar or Columbia colistin-nalidixic acid blood agar and eosin-methylene blue agar. Look for star-shaped colonies (yeasts) after 48 hours on eosin-methylene blue agar.</p></sec><sec id="cesec56"><title>Eye cultures</title><p id="para285">Use a sterile cotton swab moistened with sterile saline or broth, and pass it over the conjunctiva of the inferior fornix of each eye. Use one half of a blood agar plate and one half of a mannitol plate for each eye. Also place material into thioglycolate medium. Alternatively, use one of the commercial multiple-media plates. Make two conjunctival scrapings and stain one with Gram stain and the other with Giemsa stain.</p></sec><sec id="cesec57"><title>Skin cultures</title><p id="para286">Cultures made from the surface of the epidermis or open ulcers are of little significance because they usually grow a mixture of nonpathogenic organisms. A culture made from the deep tissue of a biopsy specimen may be helpful in the diagnosis of a bacterial, atypical mycobacterial, or subcutaneous mycotic infection. Diagnostic isolates may be obtained from cultures of tissue sections from ulcers, fistulas, abscesses, enlarged nodes, or granulomatous lesions. Smears and cultures made from exudates of deep fistulas and node aspirates may be useful in some cases.</p><p id="para287">Intact pustules are satisfactory lesions for making smears and cultures. After the skin surface has been sterilized carefully, gently aspirate the fluid content of the pustule with a sterile needle and syringe for inoculation into appropriate media; alternatively, open the pustule roof and take a culture (by swab) from the fluid inside. In all these procedures, take the utmost care to prevent contamination from tissues outside the area of primary involvement.</p><p id="para288">When any fluid material or tissue is cultured, it is always desirable to use a portion of the sample to make stained smears. Stained smears often provide immediate clues to the diagnosis (organisms present [yeast, bacteria, or fungi] and indications as to the host response [cell types, phagocytosis, or eosinophils]). Examine the slides for the presence of bacteria and for cell morphology.</p></sec><sec id="cesec58"><title>Urine culture</title><p id="para289">Urine, as it is secreted by the kidneys, is sterile unless the kidney is infected. Most urinary tract infections are ascending infections by organisms introduced through the urethra. The most common sites of infection in female animals are the urethra and urinary bladder. Chronic prostatitis is common in male dogs and often is associated with relapsing urinary tract infections.</p><p id="para290">Urine specimens can be collected by catheterization, by collecting a clean voided midstream sample, or by cystocentesis (<xref rid="cetable4" ref-type="table">Table 4-4</xref>). Cystocentesis is the preferred method for qualitative and quantitative bacterial culture. To calibrate bacterial counts in urinary cultures, use a standard platinum milk dilution loop calibrated to deliver 0.001 mL of urine to one half of a blood agar plate. The initial loop of urine is streaked onto the plate. One hundred colonies or more signifies a bacterial count in the original specimen greater than or equal to 10<sup>5</sup> cells/mL. The number of bacteria that is significant varies with the method of collection. With cystocentesis, a bacterial count greater than 10<sup>3</sup> cells/mL of urine is significant; with catheterization, greater than 10<sup>5</sup> cells/mL is significant. A MacConkey agar plate can be used in addition to a blood agar plate.<table-wrap position="float" id="cetable4"><label>TABLE 4-4</label><caption><p>Interpretation of Quantitative Urine Cultures in Dogs and Cats<xref rid="cetablefn3" ref-type="table-fn">*</xref></p></caption><table frame="hsides" rules="groups"><thead><tr><th></th><th colspan="6" align="center">Colony-forming units/mL urine<hr></hr></th></tr><tr><th></th><th colspan="2" align="center">Significant<hr></hr></th><th colspan="2" align="center">Suspicious<hr></hr></th><th colspan="2" align="center">Contaminant<hr></hr></th></tr><tr><th align="center">Collection method</th><th align="center">Dogs</th><th align="center">Cats</th><th align="center">Dogs</th><th align="center">Cats</th><th align="center">Dogs</th><th align="center">Cats</th></tr></thead><tbody><tr><td align="left">Cystocentesis</td><td align="left">≥1000</td><td align="left">≥1000</td><td align="center">100–1000</td><td align="center">100–1000</td><td align="left">≤100</td><td align="left">≤100</td></tr><tr><td align="left">Catheterization</td><td align="left">≥10,000</td><td align="left">≥1000</td><td align="center">1000–10,000</td><td align="center">100–1000</td><td align="left">≤1000</td><td align="left">≤100</td></tr><tr><td align="left">Voluntary voiding</td><td align="left">≥100,000<xref rid="cetablefn4" ref-type="table-fn">†</xref></td><td align="left">≥10,000</td><td align="center">10,000–90,000</td><td align="center">1000–10,000</td><td align="left">≤10,000</td><td align="left">≤1000</td></tr><tr><td align="left">Manual compression</td><td align="left">≥100,000<xref rid="cetablefn4" ref-type="table-fn">†</xref></td><td align="left">≥10,000</td><td align="center">10,000–90,000</td><td align="center">1000–10,000</td><td align="left">≤10,000</td><td align="left">≤1000</td></tr></tbody></table><table-wrap-foot><fn id="cetablefn3"><label>*</label><p id="cenotep3">The data represent generalities. On occasion, bacterial urinary tract infections may be detected in dogs and cats with the fewer organisms (i.e., false-negative results).</p></fn></table-wrap-foot><table-wrap-foot><fn id="cetablefn4"><label>†</label><p id="cenotep4"><italic>Caution:</italic> Because contamination of midstream samples may result in colony counts of 10,000/mL or more in some dogs (i.e., false-positive results), they should not be used for routine diagnostic culture of urine from dogs.</p></fn></table-wrap-foot><attrib>From Osborne CA, Finco DR: Canine and Feline Nephrology and Urology, Baltimore, Williams & Wilkins, 1995.</attrib><permissions><copyright-statement>© 2006 Williams & Wilkins</copyright-statement><copyright-year>2006</copyright-year><license><license-p>Since January 2020 Elsevier has created a COVID-19 resource centre with free information in English and Mandarin on the novel coronavirus COVID-19. The COVID-19 resource centre is hosted on Elsevier Connect, the company's public news and information website. Elsevier hereby grants permission to make all its COVID-19-related research that is available on the COVID-19 resource centre - including this research content - immediately available in PubMed Central and other publicly funded repositories, such as the WHO COVID database with rights for unrestricted research re-use and analyses in any form or by any means with acknowledgement of the original source. These permissions are granted for free by Elsevier for as long as the COVID-19 resource centre remains active.</license-p></license></permissions></table-wrap></p><p id="para291">Cystocentesis samples collected from animals that have received antimicrobial therapy should have 5 mL of urine centrifuged at 2500 rpm for 5 minutes, and the sediment should be streaked onto blood agar and MacConkey agar.</p><p id="para292">MacConkey agar and eosin-methylene blue agar are selective and differential media that are used to identify urinary tract organisms. MacConkey agar prevents early growth of <italic>Proteus,</italic> inhibits growth of gram-positive bacteria, and allows separation of gram-negative bacteria in lactose-positive and lactose-negative subgroups.</p><p id="para293">Several commercial methods for urinary culture are available for screening urine for bacterial infection. Bayer Microstix (Fisher Scientific International, Inc., Hampton, New Hampshire) has proved 92% accurate in detecting bacteriuria of greater than 10<sup>5</sup> cells/mL. If urine is collected by cystocentesis, significant bacteriuria may not be observed. Reculture samples that are positive by Microstix using calibrated loop or pour plate techniques.</p><p id="para294">Use catheterization with aseptic technique or antepubic cystocentesis to collect urine for culture. Refrigerate specimens of urine within a few minutes after collection if culture is not done immediately. Perform bacterial culture of the specimen within 2 hours of collection. Becton Dickinson supplies a Vacutainer urine transport kit for urine culture.
The Vacutainer tube can hold 5 mL of urine, which can be taken from a midstream catch or cystocentesis. The collection tube has a bacteriostatic fluid that preserves unrefrigerated urine specimens for up to 24 hours for culture.</p></sec><sec id="cesec59"><title>Prostatic fluid culture</title><p id="para295">Bacterial infection of the prostate may result in a nidus of infection that can cause recurrent urinary tract infection and prostatomegaly in male dogs. An effective way to evaluate the prostate for bacterial infection is to examine the prostatic fraction (the third fraction) of the male ejaculate; if separation proves to be too difficult, use the whole ejaculate specimen for culture. To better interpret the results of the prostatic culture, obtain urethral cultures before the ejaculate sample (see also Prostatic Wash).</p><p id="para296">Collect the ejaculate fraction into a sterile side-mouth container (such as a 12-mL sterile plastic syringe container). Make subcultures with 0.1 mL of ejaculate onto differential media as for urethral swabs. The prostatic ejaculate culture shows significant bacterial infection if the number of bacteria in the prostatic culture is greater than 2 logs of growth compared with the bacteria in the urethral culture.</p></sec><sec id="cesec60"><title>Stool cultures</title><p id="para297">Acute infectious diarrhea can be caused by bacteria, viruses, and protozoa. The major bacteria in feces are non–spore-forming anaerobic bacilli, but gram-negative facultative anaerobic bacteria such as <italic>Escherichia coli</italic> and other members of the Enterobacteriaceae family are usually present. The clinical picture in acute infectious diarrhea is frequent loose stools containing pus or blood, abdominal pain, and fever. Damage to the intestinal tract may be produced by an enterotoxin, as with <italic>Staphylococcus aureus</italic> or <italic>E. coli,</italic> or by invasion of the mucosa of the small intestine and colon. The most common bacterial pathogens of the intestinal tract in small animals are <italic>E. coli, Salmonella</italic> spp., and <italic>Campylobacter jejuni.</italic></p></sec></sec><sec id="cesec61"><title>Blood Culture</title><p id="para298">Bacteria can enter the blood from extravascular sites by way of the lymphatic circulation. Direct entry of bacteria into the bloodstream can be observed in the presence of endocarditis, suppurative phlebitis, infected intravenous catheters, dialysis cannulas, and osteomyelitis. Bacteremia can be transient, intermittent, or persistent. Transient bacteremia is produced by manipulation of an abscess, dental procedures, urethral catheterization, or surgery on contaminated areas. Intermittent bacteremia is associated with undetected and undrained abscesses. Most dogs with bacteremia, especially gram-negative bacteremia, are febrile and have an abnormal peripheral blood picture with an increased white blood cell count, increased number of band and segmented neutrophils, increased number of monocytes, and lymphopenia. An exception to this is osteomyelitis, in which dogs with bacteremia associated with staphylococci have basically normal hemograms. Large-breed male dogs with valvular insufficiency, congestive heart failure, or thromboembolism should be suspects for infectious endocarditis. The mitral valve most often is involved, followed by the aortic, tricuspid, and pulmonary valve.</p><p id="para299">The material for culture must be collected under aseptic conditions. Clip and surgically prepare the skin over the cephalic, recurrent tarsal, or jugular vein. Do not draw blood for culture through an indwelling intravenous or intraarterial catheter. Collection vials are available for aerobic and anaerobic bacterial culture. Add the required volume of blood (usually 8 to 10 mL) to the enriched culture medium. Immediately after collection, mix the contents of bottles or tubes to prevent clotting.</p><p id="para300">Take blood for cultures 1 hour before temperature spikes if intermittent fevers are present (<xref rid="cetextbox2" ref-type="boxed-text">Box 4-2</xref>). Take three separate blood culture specimens over a 24-hour-period. With a 1:10 dilution of blood in broth, antibiotics that may have been administered systemically usually are diluted to noninhibitory concentrations. The addition of sodium polyanethole sulfonate to commercial culture media inactivates aminoglycosides present in clinical concentrations.<boxed-text id="cetextbox2"><label>BOX 4-2</label><caption><title>INDICATIONS FOR PERFORMING A BLOOD CULTURE</title></caption><p id="para8">Any acute illness with fever (fever of unknown origin)</p><p id="para9">Hypothermia</p><p id="para10">Leukocytosis, particularly with a left shift</p><p id="para11">Neutropenia</p><p id="para12">Unexplained tachycardia</p><p id="para13">Undiagnosed hypoglycemia</p><p id="para14">Unexplained tachypnea or dyspnea</p><p id="para15">Undiagnosed anuria or oliguria</p><p id="para16">Unexplained icterus</p><p id="para17">Thrombocytopenia</p><p id="para18">Disseminated intravascular coagulation</p><p id="para19">Intermittent shifting leg lameness</p><p id="para20">Sudden development of, or change in, a murmur</p></boxed-text></p><p id="para301">Other media that may be used as selective agents include MacConkey agar, brain-heart infusion agar, mannitol salt agar, Streptosel agar, urea agar, blood agar, and eosin-methylene blue agar. Special techniques make it possible to determine total bacteria counts and whether an organism is coagulase-positive or coagulase-negative.</p><sec id="cesec62"><title>Anaerobic culture of blood</title><p id="para302">Because anaerobes may be present in significant numbers in positive cultures from blood, abscesses, wounds, and urine, it may be advisable to make these special examinations. Anaerobes are present in the normal flora in fecal, throat, and bronchial swabs, so the anaerobic culture of these samples may be difficult to evaluate.</p><p id="para303">Specimens for anaerobic examination should be protected from air, held at room temperature, and inoculated directly onto culture media as soon as possible. Specimens should not be inoculated onto transport or enrichment media. Specimens can be held for short periods in sterile, carbon dioxide–filled, tightly stoppered tubes or bottles. Inoculate the sample onto prereduced anaerobically sterilized medium under oxygen-free gas. Specimens can be inoculated deep into thioglycolate medium for transfer and subculture. With anaerobic organisms, it is especially important to make a smear and a Gram stain and to record all morphotypes present and the relative numbers of each (<xref rid="cetextbox3" ref-type="boxed-text">Box 4-3</xref>).
<boxed-text id="cetextbox3"><label>BOX 4-3</label><caption><title>INDICATIONS FOR SUBMITTING SPECIMENS FOR ANAEROBIC CULTURE</title></caption><p id="para21">Any focal pain and swelling with fever</p><p id="para22">Nonhealing bite or puncture wound</p><p id="para23">Foul-smelling wounds with persistent discharge</p><p id="para24">Presence of gas in tissue, especially if associated with a penetrating injury</p><p id="para25">Abscess, especially if recurrent</p><p id="para26">Necrotic or devitalized tissue</p><p id="para27">Dark, discolored discharge from the site of a penetrating injury</p><p id="para28">Visible sulfur granules in any discharge</p><p id="para29">Identification of filamentous bacteria during routine microscopy of exudates</p><p id="para30">Failure to obtain bacterial growth using aerobic techniques</p></boxed-text></p><sec id="cesec63"><sec id="cesec64"><title>Additional Reading</title><p id="para304">Dow S: Diagnosis of bacteremia in critically ill dogs and cats. In Bonagura J, editor: <italic>Current veterinary therapy XII. Small animal practice,</italic> Philadelphia, 1995, WB Saunders.</p><p id="para305">Greene CE: <italic>Infectious diseases of the dog and cat,</italic> ed 3, St Louis, 2006, Elsevier-Saunders.</p><p id="para306">Osborne C: Three steps to effective management of bacterial urinary tract infections: <italic>Compend Contin Educ Pract Vet</italic> 17:1233-1248, 1995.</p><p id="para307">Osborne CA, Finco DR: <italic>Canine and feline nephrology and urology</italic>, Baltimore, 1997, Williams & Wilkins.</p><p id="para308">Scott DW, Miller WH Jr, Griffin CE: <italic>Muller and Kirk's small animal dermatology,</italic> ed 5, Philadelphia, 1997, WB Saunders.</p></sec></sec></sec></sec><sec id="cesec65"><title>Fungal Culture</title><p id="para309">Diagnostic fungal cultures depend on selection of the most appropriate culture site, proper collection of specimens, and appropriate use of selective media. Culture specimens from patients suspected of having superficial fungal infections (dermatophytosis) are made from hair, skin, nails, and biopsy tissues. Test patients suspected of having deep mycoses (e.g., blastomycosis and histoplasmosis) by cytopathologic or diagnostic serologic testing (see Section 5).</p><sec id="cesec66"><title>Hair</title><p id="para310">If the hair is grossly dirty, clean it with soap and water; if not, wash it carefully with alcohol. Allow the hair to dry thoroughly. Select a site at the edge of an active lesion, and look for broken or stubby hairs. Use a forceps (curved Kelly or mosquito hemostats), and depilate hair from these areas by pulling parallel to the direction of the hair growth. It is important to get the hair root and not break off the hair shaft. Pluck many hairs and implant (push) the roots of the hair into the selected agar. Then gently lay the hair shaft down to contact the surface of the medium. Hairs for inoculation often can be selected by choosing those that fluoresce with a Wood's light.</p><p id="para311">Examination of some of the plucked hairs with a potassium hydroxide or wet-mount preparation for spores and hyphae is desirable. Never take specimens from areas that have been treated within 1 week. If samples are to be sent to a laboratory, the dry hair can be placed in a clean, tightly sealed envelope and mailed.</p></sec><sec id="cesec67"><title>Skin</title><p id="para312">Dermatophyte or yeast infections may affect glabrous skin. If necessary, cleanse culture sites with <italic>alcohol gauze swabs</italic> (cotton will leave excess fibers) and allow to dry. Using a fine scalpel blade, collect superficial scrapings of scales, crusts, and epidermal debris at the periphery of typical lesions. Dermatophytes live in a dry state for several weeks, but yeast infections should be cultured immediately or placed in transport medium to prevent drying.</p></sec><sec id="cesec68"><title>Nails</title><p id="para313">Although hard keratin fungal infections are rare in animals, diseased nails should be avulsed, scraped, or ground into fine pieces for collection in a sterile Petri dish. Pieces can be examined directly for arthrospores or hyphae and placed on appropriate media for culture.</p></sec><sec id="cesec69"><title>Tissue biopsy</title><p id="para314">Tissue core or excision samples can be sliced and the newly exposed surface used for impression smears or inoculation of medium. Samples also may be chopped or ground and placed in medium. Place small amounts in sterile saline or broth for referral to an appropriate laboratory for further processing.</p></sec><sec id="cesec70"><title>Dermatophyte media</title><p id="para315">Sabouraud dextrose agar has been used traditionally in veterinary mycology for isolation of fungi; however, other media are available with bacterial and fungal inhibitors, such as dermatophyte test medium (DTM), potato dextrose agar, and rice grain medium. Mycosel and mycobiotic agar are formulations of Sabouraud dextrose agar with cycloheximide and chloramphenicol added to inhibit fungal and bacterial contaminants. If a medium with cycloheximide is used, fungi sensitive to it will not be isolated. Organisms sensitive to cycloheximide include <italic>Cryptococcus neoformans,</italic> many members of the Zygomycota, some <italic>Candida</italic> spp., <italic>Aspergillus</italic> spp., <italic>Pseudallescheria boydii,</italic> and many agents of phaeohyphomycosis. Dermatophyte test medium is essentially a Sabouraud dextrose agar containing cycloheximide, gentamicin, and chlortetracycline as antifungal and antibacterial agents. The pH indicator phenol red has been added. Dermatophytes use protein in the medium first, and alkaline metabolites turn the medium red. When the protein is exhausted, the dermatophytes use carbohydrates and give off acid metabolites, and the color of the medium returns to yellow. Most other fungi use carbohydrates first and protein later, so they too may produce a red change in DTM, but only after a prolonged incubation (10 to 14 days or more). Consequently, examine DTM cultures daily for the first 10 days. Fungi such as <italic>Blastomyces dermatitidis, Sporothrix schenckii, Histoplasma capsulatum, Coccidioides immitis, P. boydii,</italic> some <italic>Aspergillus</italic> spp., and others may cause a red change in DTM, so microscopic examination is essential to avoid an erroneous presumptive diagnosis. Because DTM may (1) depress development of conidia, (2) mask colony pigmentation, and (3) inhibit some pathogens, fungi recovered on DTM should be transferred to plain Sabouraud dextrose agar for identification.</p><p id="para316">Potato dextrose agar is useful for promoting sporulation and observing pigmentation. On potato dextrose agar, <italic>Microsporum canis</italic> has a lemon-yellow pigment, whereas <italic>M. audouinii</italic> has a salmon- or peach-colored pigment. Rice agar medium promotes conidia formation in some dermatophytes, especially <italic>M. canis</italic> strains, which produce no conidia on Sabouraud dextrose agar.</p><p id="para317">Inoculate skin scrapings, nails, and hair onto Sabouraud dextrose agar, DTM, mycosel, or mycobiotic agar. Incubate cultures at 30° C with 30% humidity. A pan of water in the incubator usually will provide enough humidity. Check cultures every 2 to 3 days for fungal growth. Cultures on DTM may be incubated for 10 to 14 days, but cultures on Sabouraud dextrose agar should be allowed 30 days to develop.</p><p id="para318">Diagnosis should depend on characteristic gross identification of cultures and careful inspection of elements from those cultures using slide preparations and slide cultures for microscopic examination. Cultures of fungi other than dermatophytes should be made by commercial or institutional laboratories with appropriate equipment and special expertise.</p></sec><sec id="cesec71"><title>The Wood's light</title><p id="para319">Ultraviolet light filtered through nickel oxide produces a beam called <italic>Wood's light.</italic> If an animal is taken into a dark room and its hair and skin are exposed to a Wood's light, fluorescence may show for several reasons. Hair shafts affected by some species of <italic>Microsporum</italic> fluoresce a bright yellow-green (like the color of a fluorescing watch face). However, iodide medications, petroleum, soap, dyes, and even keratin may produce purple-, blue-, or yellow-colored fluorescence. The positive fungal fluorescence is a valuable aid in selecting affected hairs for culture inoculation. Remember, a negative fluorescence does not preclude a possible diagnosis of fungal infection. False negatives and false positives may occur.</p><sec id="cesec72"><sec id="cesec73"><title>Additional Reading</title><p id="para320">Dow. S: Diagnosis of bacteremia in critically ill dogs and cats. In Bonagura J, editor: <italic>Current veterinary therapy XII. Small animal practice,</italic> Philadelphia, 1995, WB Saunders.</p><p id="para321">Greene CE: <italic>Infectious diseases of the dog and cat,</italic> ed 3, St Louis, 2006, Elsevier-Saunders.</p><p id="para322">Scott DW, Miller WH Jr, Griffin CE: <italic>Muller and Kirk's small animal dermatology,</italic> ed 5, Philadelphia, 1997, WB Saunders.</p></sec></sec></sec></sec><sec id="cesec74"><title>Virus Isolation</title><p id="para323">Several techniques are currently available for the identification of viral infections in dogs and cats. Among the most convenient are antigen-detection systems available as point-of-care tests for feline leukemia virus antigen in blood and canine parvovirus antigen in feces. These tests identify infected patients with excellent accuracy. Additionally, and seldom considered for their value, point-of-care tests for viral infections are especially capable of identifying patients that have not been exposed, allowing the clinician reliably to rule out infection by the organism for which the animal is tested.</p><p id="para324">In addition, many commercial and point-of-care serologic assays are available that detect antibody to many of the viruses affecting dogs and cats. However, the positive predictive value of antibody tests is considerably lower than that for antigen tests. For example, a single positive antibody titer in a dog for canine distemper is evidence of prior exposure or vaccination. A POSITIVE antibody titer to a particular viral pathogen does <italic>not</italic> constitute a diagnosis of infection, especially in the absence of clinical signs. However, a NEGATIVE antibody titer generally does denote no prior exposure or failure to respond to vaccination. Obtaining acute and convalescent titers in a patient suspected of having an acute viral infection can be a reliable diagnostic tool if a fourfold or greater increase in titer can be demonstrated over 3 to 4 weeks. Acute and convalescent viral titers in individual patients are rarely performed in veterinary medicine.</p><p id="para325">Virus isolation, however, is a valuable diagnostic tool that is underutilized in veterinary medicine, perhaps because of the limited number of commercial and university laboratories that provide viral isolation services.</p><p id="para326">Diagnosis of viral upper respiratory infection in cats (herpesvirus-1 and/or calicivirus) is perhaps among the situations for which virus isolation can be most useful, especially in cluster households where many cats and kittens may be at risk. Quickly insert a sterile cotton swab into the oral cavity to the level of the tonsil or oropharynx. By rolling the swab across the epithelium, it is possible to harvest cells and virus from infected cats. Immediately place the swab into a virus transport medium (usually provided by the laboratory). Antibiotics added to the solution prevent bacterial overgrowth of the sample. For short-term transit (5 days or less), hold specimens for viral isolation at 4° C rather than frozen. On reaching the laboratory, the specimen will be inoculated into a suitable tissue culture. Within a few days it is usually possible to establish, based on the cytopathic effect on the tissue culture, whether a virus infection is present. Fluorescent antibody test can be done subsequently to confirm the isolate.</p><p id="para327">Some laboratories offer direct assessment of specimens (e.g., feces for canine parvovirus or canine coronavirus) by electron microscopy. These methods can be useful for infections in which the virus titer in the specimen reaches 10<sup>6</sup> to 10<sup>7</sup> organisms/mL. Specimens such as feces, vesicle fluid, brain tissue, urine, or serum can be negatively stained for electron microscopy.</p><p id="para328">Recently, considerable interest has surfaced regarding the availability of virus testing by polymerase chain reaction. Polymerase chain reaction is an exquisitely sensitive test for viral (or bacterial or fungal) DNA in patients known or suspected to have an infection. Objectively, only trace amounts of DNA (or in some cases, RNA) of the infecting virus are required in the specimen. These amounts are amplified millions of times to facilitate identification of the target sequence (and the infecting organism). However, the disadvantages to polymerase chain reaction testing are price (prices are sufficiently high today to preclude routine diagnostic testing for viral infection by polymerase chain reaction) and the sensitivity of the test. So sensitive is polymerase chain reaction, that extraneous, non–target DNA (in the specimen or in the laboratory) may contaminate the sample easily, making diagnosis difficult to impossible.</p></sec><sec id="cesec75"><title>Blood Collection Techniques</title><p id="para329">Venipuncture of the dog and cat may be accomplished by using the cephalic, jugular, femoral, or recurrent tarsal veins. In large dogs and cats, the cephalic and jugular veins are preferred. Performance of venipuncture as atraumatically as possible is essential to preserve the integrity of the vein, particularly when multiple venipunctures must be performed and a central venous catheter cannot be placed to facilitate the collection of multiple blood samples. For aesthetic reasons and in show cats and dogs, it is undesirable to clip the fur over the vessels, if this can be avoided. However, clipping the hair can aid in identification and visualization of the vein. After clipping the overlying fur, aseptically scrub the skin or place a quantity of isopropyl alcohol over the vessel. If the fur is not clipped, part the fur with alcohol to improve visualization.</p><p id="para330">In most instances, a 3- to 5-mL (2-mL minimum according to most commercial laboratories) blood sample is adequate for routine hematologic and biochemical analyses. Plan ahead which samples are required to prevent the need for further venipuncture at a later time, whenever possible. In small dogs and cats, use the jugular veins to allow adequate sample collection. If smaller samples are required, the cephalic, lateral saphenous, or medial saphenous veins can be used for sample collection. Do not use the jugular vein if a coagulopathy is suspected because hemorrhage may be difficult to control following venipuncture.</p><p id="para331">For successful venipuncture, proper restraint of the animal is important. Details for the proper restraint for various venipuncture locations are discussed with each specific topic throughout this text. The patient must remain comfortable yet relatively motionless to avoid iatrogenic vessel laceration. Stretch the skin tightly over the selected vessel without causing vascular occlusion to help anchor the vessel in place during penetration by the needle.</p><p id="para332">Use a dry, sterile needle and syringe. Grasp the syringe tightly in between the thumb and forefingers. Place the index finger near the hub of the syringe to help guide it in place. In most cases, it is best to penetrate the skin just lateral to the vessel. Further advance the needle to puncture the vessel from the side. Blood usually will enter the hub of the needle spontaneously but can be encouraged to enter by pulling on the plunger of the syringe. After the skin and underlying fascia have been punctured, some clinicians apply constant gentle negative pressure on the vessel by pulling back on the plunger of the syringe. Too much suction will collapse the vein and cause inadequate flow of blood into the syringe. Decreased blood flow also may be caused by inadequate tissue perfusion, hypothermia, circulatory failure, hematoma formation, and piercing the vessel wall without penetrating into the vessel lumen. Occasionally, the tip of the needle will become snagged on the opposite wall of the vessel, or by a valve within the vessel. Repositioning the needle with slight rotation and retraction from the vessel may correct the problem. The flow of blood can be improved by alternating occlusion and release of the vein, combined with passive motion of the leg. Complications of venipuncture include hematoma formation, minor hemorrhage, vascular trauma, and thrombophlebitis.</p><p id="para333">The use of a Vacutainer has simplified the collection of blood samples from small animals, particularly when larger vessels are catheterized for blood collection. Do not use the larger-sized Vacutainer containers in small veins because the veins will collapse from the negative pressure within the tube. To ensure that the proper ratio of anticoagulant-to-blood is obtained, fill all tubes that contain anticoagulants until the vacuum is exhausted.</p><p id="para334">When handling blood samples, use clean and dry syringe, needles, or evacuated blood collection tubes. Avoid hemolysis by using clean, dry equipment and avoiding trauma to the red blood cells. Trauma to the red blood cells occurs because of application of too much or fluctuating suction during aspiration, excessive force when expelling the blood sample into the blood collection tube, or excessive agitation of the sample once within the tube. Hemolysis can interfere with a number of diagnostic tests and should be avoided.</p><p id="para335">Make an effort to have the animal fast for a minimum of 12 hours before the collection of blood samples to avoid postprandial lipemia. Lipemia is attributable to metabolic disorders (pancreatitis, diabetes mellitus) and to recent meals and adversely and artifactually can affect blood test results. Total protein, albumin, glucose, calcium, phosphorus, and bilirubin are examples of tests that can be affected greatly by sample lipemia. The clinician should be aware of the tests that are affected by lipemia and sample hemolysis and various anticoagulants.</p><sec id="cesec76"><title>Routine hematologic testing (see also Section 5)</title><p id="para336">The anticoagulant of choice for hematologic testing is EDTA. Heparin is especially to be avoided if blood films are to be made from blood mixed with anticoagulant because contact with whole blood will distort the morphology of cells significantly. Heparin is acceptable for most procedures requiring blood plasma. The anticoagulant effect of heparin is transitory. Specimens still may clot after 2 to 3 days.</p><p id="para337">Make blood films immediately after collection because cell morphology rapidly deteriorates after sample collection. Although blood films can be made after introducing blood to EDTA, a better practice is to make blood smears (films) immediately from the collection needle before the blood comes in contact with any anticoagulant. <italic>Never use blood exposed to heparin to make blood smears.</italic></p><p id="para338">Incorrect proportions of blood to anticoagulant may result in water shifts between plasma and red blood cells. Such shifts may alter the packed cell volume, especially when small amounts of blood are added to tubes prepared with volumes of anticoagulant sufficient for much larger volumes of blood. Erroneous laboratory results also may be obtained when small volumes of blood are placed in a relatively large container. Evaporation of plasma water and adherence of the cells to the surface of the container can produce artifactual changes in hematologic results.</p><p id="para339">Refrigerate liquid blood mixed with anticoagulant after collection if there is a delay in making the laboratory determinations. White blood cell (WBC) and red blood cell counts, packed cell volume, and hemoglobin level can be measured within 24 hours of sample collection. Platelet counts, however, should be done within 1 hour of collection. Dried, unfixed blood smears can be stained with most conventional stains 24 to 48 hours after being made. If a considerable delay is anticipated between the time that the blood smear is made and the staining process, the blood smear should be fixed by immersion in absolute methanol for at least 5 minutes. Blood smears fixed by this method are stable indefinitely. Never place unfixed blood smears in a refrigerator because condensation forming after the smear is removed from the refrigerator will ruin the blood smear and make it unusable for cytologic evaluation. Take care to leave unfixed blood smears face down on a countertop or in a closed box. Special stains, such as peroxidase, may require fresh blood films.</p></sec><sec id="cesec77"><title>Routine biochemistry testing (see also Section 5)</title><p id="para340">Most clinical chemistry procedures are performed on serum. The serum is obtained by collecting blood without any anticoagulant and allowing the blood to clot in a clean, dry tube. Separate serum from cells within 45 minutes of sample collection (venipuncture). Special vacuum vials are available that produce a strong barrier between the clot and the serum so that it is not necessary to draw off the serum into a separate vial. Clotting of the blood and retraction of the clot occur best and maximum yields of serum are obtained at room or body temperatures. Refrigeration of the sample impairs clot retraction. When the blood is firmly clotted, free the clot from the walls of the container by rimming with an applicator stick or by tapping sharply on the outside of the tube. After the clot is freed, allow clot retraction to occur, and then centrifuge and draw off the clear supernatant serum using a pipette or suction bulb. Serum yield is usually one third of the whole blood volume, unless severe hypovolemia or intravascular dehydration is present.</p><p id="para341">Many clinical chemistry procedures can be performed on plasma and on serum. The advantage of using plasma is that separation of cells can be accomplished immediately after centrifugation or sedimentation, without the need to wait for clot formation and retraction. The disadvantage of plasma is that the presence of the anticoagulant interferes with many of the chemistry assay procedures. Plasma is less clear than serum, which may be an additional disadvantage for colorimetric assays. Plasma and serum are virtually identical in chemical composition except that plasma has fibrinogen and the anticoagulant. For many procedures in which plasma or whole blood is to be used, heparin is the anticoagulant of choice. Heparinized blood is the only acceptable specimen for blood pH and blood gas analyses. Although blood containing EDTA is acceptable for certain chemical procedures, it cannot be used for determination of plasma electrolytes because it contributes to and sequesters them from the specimen. In addition, EDTA can interfere with alkaline phosphatase levels, decrease total carbon dioxide, and elevated blood nonprotein nitrogen.</p><p id="para342">Separate serum or plasma and remove it from the cells as soon as possible after blood is collected, because many of the constituents of plasma exist in higher concentrations in red blood cells. With time, these substances leak into the plasma and cause falsely elevated values (positive interference) and falsely lower values (negative interference) (<xref rid="cetable5" ref-type="table">Table 4-5</xref>). Under no circumstances should whole blood be sent via the mail; serum derived from such specimens usually is hemolyzed, and results are often inaccurate. Separate serum and transfer it to a clean, dry tube for shipment.<table-wrap position="float" id="cetable5"><label>TABLE 4-5</label><caption><p>Examples of Positive and Negative Interference on Biochemistry Analytes Induced by Sample Hemolysis</p></caption><table frame="hsides" rules="groups"><thead><tr><th align="left">Analyte</th><th align="left">Effect of hemolysis<xref rid="cetablefn5" ref-type="table-fn">*</xref></th></tr></thead><tbody><tr><td align="left">Alanine transaminase</td><td align="left">Minimal effect</td></tr><tr><td align="left">Alkaline phosphatase</td><td align="left">Increased</td></tr><tr><td align="left">Bilirubin</td><td align="left">Increased</td></tr><tr><td align="left">Chloride</td><td align="left">Decreased</td></tr><tr><td align="left">Creatinine</td><td align="left">Increased</td></tr><tr><td align="left">Inorganic phosphate</td><td align="left">Increased</td></tr><tr><td align="left">Lipase</td><td align="left">Decreased</td></tr><tr><td align="left">pH</td><td align="left">Decreased</td></tr><tr><td align="left">Potassium</td><td align="left">No detectable effect</td></tr><tr><td align="left">Total calcium</td><td align="left">Increased</td></tr><tr><td align="left">Total protein</td><td align="left">Increased</td></tr><tr><td align="left">Urea nitrogen</td><td align="left">Increased</td></tr></tbody></table><table-wrap-foot><fn id="cetablefn5"><label>*</label><p id="cenotep5">Type and degree of interference varies among different testing modalities unique to individual laboratories or in-hospital biochemistry analyzers.</p></fn></table-wrap-foot></table-wrap></p><sec id="cesec78"><sec id="cesec79"><title>Additional Reading</title><p id="para343">Meyer DJ, Harvey JW: <italic>Veterinary laboratory medicine,</italic> ed 3, St Louis, 2004, Elsevier-Saunders.</p><p id="para344">Raskin R, Meyer DJ (eds): Update on clinical pathology, <italic>Vet Clin North Am Small Anim Pract</italic> 26:5, September 1996.</p><p id="para345">Thomas JS: Introduction to serum chemistries: artifacts in biochemical determinations. In Willard MD, Tvedten H, editors: <italic>Small animal clinical diagnosis by laboratory methods,</italic> ed 4, St Louis, 2004, Elsevier-Saunders.</p></sec></sec></sec></sec><sec id="cesec80"><title>Bone Marrow Aspiration</title><p id="para346">Collection of bone marrow may prove valuable in diseases of the blood in which examination of the peripheral blood reveals abnormal cells or cell counts. Conditions such as leukopenia, thrombocytopenia, nonregenerative anemias, agranulocytosis, pancytopenia, and leukemias may be present because of pathologic changes within the bone marrow.</p><p id="para347">Bone marrow in the young animal is cellular and exists in the flat bones (sternum, ribs, pelvic bones, and vertebrae) and in the long bones (humerus and femur). As the animal ages, the cellular content of the marrow decreases, especially in the long bones. In older animals, bone marrow cells still exist in the flat bones; however, in conditions of stress in which new blood cells must be produced in large numbers, primitive cells in the bone marrow of the long bones again become active. Interpretation of the bone marrow smear may be limited by (1) technique used to obtain a bone marrow specimen or (2) the specialized knowledge necessary to interpret bone marrow cells.</p><p id="para348">Bone marrow aspiration is much underused in clinical practice. The procedure does require some degree of skill to obtain high-quality samples, but the procedure is low risk to the patient and can be highly valuable in establishing a diagnosis or prognosis.</p><sec id="cesec81"><title>Canine</title><p id="para349">The biopsy techniques that may be used in the examination of bone marrow are aspiration, core, and incisional biopsy. The most frequently used technique is aspiration biopsy. When aspiration biopsy fails to produce bone marrow cells (as in advanced myelofibrosis, neoplasia, or marrow aplasia), a core biopsy of bone marrow is indicated. Bone marrow aspiration needles may be a 16-gauge Rosenthal needle or Illinois needle for medium-sized dogs; an 18-gauge Rosenthal needle for small dogs and cats; or a Jamshidi (pronounced yam-she-dee) bone marrow biopsy needle, 12 gauge for most adult dogs and 14 gauge for small dogs and cats.</p><p id="para350">The selection of needles for aspiration biopsy of bone marrow is based on the biopsy site, the depth of the biopsy site, and the density of cortical bone. For bone marrow aspiration, the modified disposable Illinois sternal-iliac bone marrow aspiration needle works well (<xref rid="f10" ref-type="fig">Figure 4-10</xref>). For a core biopsy of bone marrow, the Jamshidi bone marrow biopsy-aspiration needle (pediatric, 3.5 inches, 13 gauge) can be used (<xref rid="f11" ref-type="fig">Figure 4-11</xref>).<fig id="f10"><label>Figure 4-10</label><caption><p>Illinois iliac-sternal bone marrow needle used for aspiration of bone marrow from the humerus, ileum, or femur of dogs and cats.</p></caption><graphic xlink:href="gr10"></graphic></fig><fig id="f11"><label>Figure 4-11</label><caption><p>Jamshidi bone marrow biopsy needles.</p></caption><graphic xlink:href="gr11"></graphic></fig></p><p id="para351">The iliac crest is a commonly used site for marrow aspiration in dogs. A short-acting anesthetic occasionally may be needed, but tranquilization together with local anesthesia is usually sufficient. Place the animal in lateral recumbency, and clip the hair over the area of the iliac crest. Surgically prepare the site. To aspirate marrow, have the needle enter the widest part of the iliac crest and stop the needle just after penetration of the bone. Remove the stylet, place a 12-mL syringe on the needle, and aspirate 0.2 mL of marrow.</p><p id="para352">Alternatively, the head of the humerus offers easy access to abundant bone marrow. Sedation may be required. With the patient in lateral recumbency and the humerus flexed (the humerus is positioned parallel to the patient's thorax), instill local anesthetic into the skin and subcutaneous tissues to the level of the head of the humerus. The site of needle insertion is on the most proximal facet of the humoral head (<xref rid="f12" ref-type="fig">Figure 4-12</xref>). Direct the needle into the bone toward the elbow and parallel to the humeral shaft. If the needle is positioned too far medially over the humeral head, it is easy to penetrate the joint capsule. Although this is a common occurrence, it does not pose a risk of injury to the patient (assuming the skin was surgically prepared). However, in the event joint fluid contaminates the bone marrow aspirate, the sample will be rendered useless.<fig id="f12"><label>Figure 4-12</label><caption><p>Illinois bone marrow needle positioned in the humeral head of dog prepared for bone marrow aspiration.</p></caption><graphic xlink:href="gr12"></graphic></fig></p><p id="para353">Contamination of the bone marrow with peripheral blood results if (1) the marrow is not aspirated immediately after the needle enters the marrow cavity or (2) if aspiration time is sustained and a large volume of blood enters the syringe subsequent to the rupture of small blood vessels in the bone marrow.</p><p id="para354">Perhaps the least desired technique is to obtain marrow from the proximal end of the femur by insertion of the bone marrow needle into the trochanteric fossa. Make a small skin incision over the trochanteric fossa just medial to the summit of the trochanter major. Insert the bone marrow aspiration needle medial to the trochanter major, and place the long axis of the needle parallel to the long axis of the femur.</p><p id="para355">Once the site is selected, grasp the needle firmly. Apply steady, slight pressure while alternately rotating the needle tip against the bone (fast, 180-degree clockwise and then counterclockwise movements). Begin with gentle pressure until the needle begins to seat into the bone. Gradually increase the pressure as the needle penetrates into the bone. Insert the bone marrow needle ½ inch into the femoral canal. Remove the stylet from the needle, and aspirate using a 12- or 20-mL syringe that contains a small volume (approximately 0.1 mL) of 4% EDTA. Use significant negative pressure, for example, by withdrawing the plunger of a 12-mL syringe to the 8- or 9-mL mark. COLLECTION OF MORE THAN 1 ML OF BONE MARROW IS UNNECESSARY. Collection of larger volumes may cause greater amounts of peripheral blood to enter the syringe, leading to hemodilution of the sample. Once collected, immediately transfer the aspirate to a watch glass containing approximately 0.25 mL of 4% EDTA. Immediately mix the sample well using the end of the syringe. This is also a good time to remove the bone marrow needle from the patient.</p><p id="para356">For accurate bone marrow interpretation, it is important that smears contain marrow particles. Marrow particles (also called spicules) appear as tiny grains within the sample and can be visualized in the watch glass. Prepare slides in a manner similar to that used for peripheral blood smears. Preparation of 5 to 8 quality slides for submission is customary. Smears are air-dried. Slides may be stained using the same stains used for peripheral blood smears.</p></sec><sec id="cesec82"><title>Feline</title><p id="para357">Accessible sites for bone marrow sampling in the cat are the iliac crest, the head of the humerus, and the proximal end of the femur via the trochanteric fossa. The techniques described for the dog can be used; however, caution is advised against using vigorous restraint with a severely anemic cat because such restraint may precipitate severe cyanosis, apnea, and cardiac arrest. Adequate sedation with supplemental oxygen administration and local anesthesia may be indicated.</p><p id="para358">Make smears of bone marrow immediately after aspiration of material. Extrinsic thromboplastin present in bone marrow tissue will cause the marrow to clot within 30 seconds. In addition, small pieces of marrow can be fixed in formalin for histologic preparation. Staining with new methylene blue, Wright's, May-Gru¨nwald, or Giemsa stain may be used. A peroxidase stain may be helpful in differentiating granulocytic elements from lymphocytes.</p><p id="para359">Another method is to aspirate the bone marrow into a syringe containing 0.25 mL of 4% EDTA solution. Expel the aspirate, up to 1 mL, into a sterile Petri dish, from which the marrow particles can be isolated easily by aspirating an aliquot with a glass pipette, placing an appropriate volume onto several glass slides, making the smear, and then staining.</p><sec id="cesec83"><sec id="cesec84"><title>Additional Reading</title><p id="para360">Crow S, Walshaw S: <italic>Manual of clinical procedures in the dog, cat, and rabbit,</italic> ed 2, Philadelphia, 1997, Lippincott-Raven.</p><p id="para361">McSherry LJ: Techniques for bone marrow aspiration and biopsy. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p><p id="para362">Meyer D, Harvey J: <italic>Veterinary laboratory medicine,</italic> ed 3, St Louis, 2004, Elsevier-Saunders.</p></sec></sec></sec></sec><sec id="cesec85"><title>Cytopathologic Specimen Collection Techniques</title><p id="para363">(See also Section 5 for additional information on slide preparation of samples to be submitted for cytopathologic examination.)</p><p id="para364">Cytopathology involves a simple, direct, and inexpensive technique that can yield significant diagnostic information within a short time at minimal direct cost. Cytologic examination can be made of material obtained from pustules, vesicles, or the raw, ulcerated, or cut surfaces of a lesion. To make the smear, press a clean microscope slide firmly against a raw or ulcerated lesion to transfer cellular material to the slide. Exudates may be collected by sterile swab or may be aspirated into a sterile syringe. Roll the swab gently across the slide, or place a drop of fluid from the syringe onto the slide and carefully spread the fluid in a uniform film. Transfer material from a block of tissue to the slide by gently pressing the tissue onto the slide in several locations. Use various stains for different conditions.</p><p id="para365">Rapid stains such as new methylene blue or a quick Romanowsky's-type stain (e.g., Diff-Quik) are useful and convenient for office procedures. Even Wright's and Gram stains for evaluation of bacteria in tissues/fluids are easy to use. The presence of many bacteria, especially mixed types, may mean only surface contamination, whereas single types of bacteria, abundant polymorphonuclear WBCs, and especially phagocytosis support the diagnosis of infection and the host response to it. A <italic>few</italic> acantholytic cells (loose epidermal cells) in the smear may be compatible with infectious processes, but large numbers, or “rafts,” of acantholytic cells are highly suggestive of pemphigus and imply the need for more complex tests for positive diagnosis.</p><p id="para366">Large numbers of eosinophils sometimes are found in stained smears. Contrary to popular opinion, they usually do not mean allergy. These cells are seen most commonly with furunculosis and may be associated with the eosinophilic granulomas, eosinophilic plaques, sterile eosinophilic pustulosis, pemphigus complex, and ectoparasites. Yeasts (usually <italic>Malassezia,</italic> rarely <italic>Candida</italic>) commonly are found as budding cells in masses of wax and debris from ear smears.</p><p id="para367">Tumor cells may be recognized in some impression or aspiration samples where Giemsa is a preferred stain. Although special expertise is needed, cases of mastocytoma, histiocytoma, and lymphoma are recognized most easily. Always prepare formalin-fixed tissues for histologic diagnosis in tumor evaluations (<xref rid="cetextbox4" ref-type="boxed-text">Box 4-4</xref>).<boxed-text id="cetextbox4"><label>BOX 4-4</label><caption><title>CYTOLOGIC FEATURES OF MALIGNANCY</title></caption><p id="para31">Enlargement of nucleus/nuclei larger than 10 nm</p><p id="para32">Decreased nuclear/cytoplasmic ratio</p><p id="para33">Multinucleation because of abnormal mitosis</p><p id="para34">Abnormal or frequent mitosis</p><p id="para35">Variations in size and shape of nuclei</p><p id="para36">Increase in size and number of nucleoli</p><p id="para37">Increased basophilia of cellular cytoplasm-increased RNA content</p><p id="para38">Anisokaryosis or pleomorphism</p><p id="para39">Multinucleated giant cells</p></boxed-text></p><sec id="cesec86"><title>Fine-needle aspiration</title><p id="para368">The ability to aspirate cells from normal and abnormal tissue, apply them to a glass slide, stain the smear, and review the results is among the most useful, cost-effective diagnostic procedures available in clinical practice. The most significant limiting factors are (1) the technical ability to prepare high-quality slides and (2) the ability to interpret the cytologic findings. Some experience is needed to obtain the skills needed to aspirate cells and make diagnostic preparations. Significant training is required to interpret the slides adequately. However, access to cytopathologists affiliated with diagnostic laboratories today makes fine-needle aspiration a highly useful diagnostic tool. The lymph node aspiration technique, described next, illustrates the finer points of the fine-needle aspiration technique.</p><p id="para369">Lymph node aspiration is a procedure that can, and should, be performed routinely in clinical practice. Follow proper technique to maximize the diagnostic use of this procedure. Lymph node aspiration typically is indicated (1) in patients with generalized lymphadenomegaly, (2) to evaluate abnormally enlarged solitary lymphnodes, and (3) in suspected instances of tumor metastases to lymph nodes. Surgically prepare the skin over the node from which a biopsy specimen is to be taken. With one hand, localize and immobilize the lymph node; with the other hand, guide the aspiration biopsy needle into the affected node. Affix a 6-mL syringe onto a 22- to 20-gauge needle (a 25-gauge needle can be used when the site to be aspirated is particularly small), and advance the needle into the lymph node. Withdrawal of the syringe to approximately 0.5 mL <italic>before inserting it into the tissue</italic> is recommended. Doing so helps to prevent expelling material when removing the sample from the tissue. When the needle is in position in the approximate center of the node, gradually draw negative pressure on the syringe to a level of 4 to 5 mL. Hold the negative pressure in place for a few seconds. Release, and then repeat 2 to 3 times. Before removing the needle from the tissue, release the negative pressure in the syringe (this is why it is recommended to have 0.25 mL of air prepositioned inside). DO NOT REMOVE THE SYRINGE FROM THE TISSUE WHILE MAINTAINING NEGATIVE PRESSURE because this can result in the aspiration of significant amounts of blood from the skin, thereby significantly diluting the sample with peripheral blood. Eject cellular material within the needle onto clean glass slides. Handle all aspirates gently. To make slides, place two slides together and pull the slides apart to avoid shearing the cells. Do not compress or force slides together. In addition, a biopsy of the lymph node can (and usually should) be performed as a means of confirming or supporting diagnostic decisions made on aspirates. Lymph node biopsies can be obtained easily and safely by punch (core) techniques (e.g., 4-mm skin biopsy punch) or Tru-Cut biopsy needle.</p></sec><sec id="cesec87"><title>Exfoliative cytologic procedure</title><p id="para370">Also called “touch impression cytology,” this technique entails preparing cytologic slides directly from the cut surface of incisional and excisional biopsy samples. Use a scalpel blade to make a full-thickness linear cut through the biopsy specimen. A fresh surface of the tissue of interest is exposed. Using forceps or a sterile needle, gently lay the tissue on a clean glass slide. DO NOT FORCE THE TISSUE ONTO THE SLIDE because this can significantly damage cells. Several imprints can be made from the same surface. As needed, make new cuts to obtain a fresh surface from which to exfoliate cells. Allow the slide to air dry completely. Apply conventional staining, and examine the specimen when it is dry. The remaining tissue, if not significantly damaged, can be submitted for histopathologic examination (recommended).</p></sec><sec id="cesec88"><title>Scrapings and swabs</title><p id="para371">Depending on the tissue type and lesion, it may be possible to obtain diagnostic cytologic samples from scrapings (e.g., conjunctival epithelium for virus inclusions), brushes (e.g., material obtained during endoscopy), and swabs (e.g., ear and vaginal swabs). The cells, once harvested, can be applied delicately directly to a clean glass slide by carefully rolling or even by just touching the material to the slide to create a thin layer. Allow the sample to air-dry thoroughly before staining.</p></sec><sec id="cesec89"><title>Fluids</title><p id="para372">Cytologic examination of fluids obtained with needle and syringe from body cavities, cysts, and urine typically require additional preparation in obtaining adequate cell concentration to make diagnostic decisions. Analyze fluid specimens with respect to protein and nucleated cell count and a morphologic description of the cells. If overall cell counts are low, centrifugation will be required to concentrate cellular material for analysis. After centrifugation, remove the supranatant (and save it). Resuspend the cells in 2 to 3 drops of the supranatant. Apply a single drop of the mixture to a glass slide and allow it to air dry. I prefer NOT to smear the liquid onto the slide; instead, I allow the liquid to run, by gravity, from one end of the slide to the other. After the liquid is thoroughly air-dried, it can be stained and reviewed.</p></sec></sec><sec id="cesec90"><title>Ectoparasites</title><sec id="cesec91"><title>Skin scraping</title><p id="para373">Skin scrapings frequently are obtained to find and identify microscopic parasites or fungal elements in the skin. Material required includes mineral oil in a small dropper bottle, a dull scalpel blade, glass slides, coverslips, and a microscope.</p><p id="para374">Select undisturbed, untreated skin for a scraping site. The best method is to scrape the periphery of skin lesions and avoid the excoriated or traumatized center areas. In scraping for demodectic mange, pinch a small fold of affected skin firmly and collect the surface material for examination. This procedure forces the mites out of the hair follicles and onto or near the skin surface. For sarcoptic mange, scrape large areas. Select sites on the elbows, hocks, and ear margins when searching for sarcoptic mange. Many or frequent scrapings may be necessary to demonstrate sarcoptic mange mites or their fecal pellets or eggs.</p><p id="para375">Place the accumulated material on a microscope slide and mix it with mineral oil. Examine the entire area with a ×10 objective thoroughly and carefully. Dry keratin and dead hairs also may be accumulated by scraping without mineral oil for inoculation of fungal cultures.</p></sec><sec id="cesec92"><title>Acetate tape preparation</title><p id="para376">This is one of the simplest diagnostic procedures. Use clear (not frosted) acetate tape. Bend the tape into a loop around the fingers with the sticky side facing out. Part the animal's hair coat, and press the tape firmly onto the skin and hair around suspect lesions. The sticky tape picks up loose particles with which it makes contact. Cut the loop of tape and place the strip of tape sticky side down on a clean microscope slide. Use a low-power microscope to look through the tape at the collected particles. This technique is excellent for trapping and identifying biting and sucking lice, <italic>Otodectes</italic> and <italic>Cheyletiella</italic> mites, flea dirt and larvae, fly larvae, or dandruff scales.</p><p id="para377">Acetate tape also is useful for studying hair abnormalities. Use a strong hemostat to securely clamp and quickly avulse a group of 10 to 20 hair shafts. Press the pointed distal ends onto sticky acetate tape (lined up like pickets in a fence), and cut the hair shafts off in the middle with a scissors. Likewise, press the butt ends with the hair roots onto another piece of tape. Then press the tape holding the hair onto a microscope slide to allow low-power examination of the hairs through the clear tape. The tips of the hairs will be well oriented and controlled; thus, it is easy to evaluate whether the hairs are split, broken, or bitten off and whether the hair roots are in the anagen or telogen growth stage.</p><sec id="cesec93"><sec id="cesec94"><title>Additional Reading</title><p id="para378">Baker R, Lumsden JH: <italic>Color atlas of cytology of the dog and cat,</italic> St Louis, 2000, Mosby.</p><p id="para379">Burkhard MJ, Meyer DJ: Invasive cytology of internal organs: cytology of the thorax and abdomen, <italic>Vet Clin North Am Small Anim Pract</italic> 26:1203, 1996.</p><p id="para380">Cowell R: <italic>Diagnostic cytology of the dog and cat,</italic> St Louis, 1998, Mosby-Year Book.</p><p id="para381">Ehrhart N: Principles of tumor biopsy, <italic>Clin Tech Small Anim Pract</italic> 13:1998.</p></sec></sec></sec></sec><sec id="cesec95"><title>Urine Collection Techniques</title><p id="para382">Urine can be removed from the bladder by one of four methods: (1) naturally voided (aka, the “free catch”), (2) manual compression of the urinary bladder (aka, expressing the bladder), (3) catheterization, or (4) cystocentesis.</p><p id="para383">For routine urinalysis, collection of urine by natural micturition is often satisfactory. The major disadvantage is the contamination of the sample with cells, bacteria, and other debris located in the genital tract. Discard the first portion of the stream because it contains the most contamination and debris. Voided urine samples are not recommended when bacterial cystitis is suspected.</p><p id="para384">Manual compression of the bladder may be used to collect urine samples from dogs and cats. Do not use excessive digital pressure; if moderate digital pressure does not induce micturition, discontinue the technique. The technique can be difficult to use in male dogs and male cats.</p><p id="para385">Urinary catheters are hollow tubes made of rubber, plastic, nylon, latex, or metal and are designed to serve four purposes:<list list-type="simple" id="celist16"><list-item id="celistitem87"><label>1.</label><p id="para386">To relieve urinary retention</p></list-item><list-item id="celistitem88"><label>2.</label><p id="para387">To test for residual urine</p></list-item><list-item id="celistitem89"><label>3.</label><p id="para388">To obtain urine directly from the bladder for diagnostic purposes</p></list-item><list-item id="celistitem90"><label>4.</label><p id="para389">To perform bladder lavage and instillations</p></list-item></list></p><p id="para390">The size of catheters (diameter) usually is calibrated in the French scale; each French unit is equivalent to roughly 0.33 mm. The openings adjacent to the catheter tips are called “eyes.” Human urethral catheters are used routinely in male and female dogs; 4F to 10F catheters are satisfactory for most dogs (<xref rid="cetable6" ref-type="table">Table 4-6</xref>). Catheters should be individually packaged and sterilized by autoclaving or ethylene oxide gas.<table-wrap position="float" id="cetable6"><label>TABLE 4-6</label><caption><p>Recommended Urethral Catheter Sizes for Routine Use in Dogs and Cats</p></caption><table frame="hsides" rules="groups"><thead><tr><th align="left">Animal</th><th align="left">Urethral catheter type</th><th align="left">Size (French units<xref rid="cetablefn6" ref-type="table-fn">*</xref>)</th></tr></thead><tbody><tr><td align="left">Cat</td><td align="left">Flexible vinyl, red rubber, or Tom Cat catheter (polyethylene)</td><td align="left">3.5</td></tr><tr><td align="left">Male dog (≤25 lb)</td><td align="left">Flexible vinyl, red rubber, or Polyethylene</td><td align="left">3.5 or 5</td></tr><tr><td align="left">Male dog (≥25 lb)</td><td align="left">Flexible vinyl, red rubber, or Polyethylene</td><td align="left">8</td></tr><tr><td align="left">Male dog (>75 lb)</td><td align="left">Flexible vinyl, red rubber, or Polyethylene</td><td align="left">10 or 12</td></tr><tr><td align="left">Female dog(≤10 lb))</td><td align="left">Flexible vinyl, red rubber, or Polyethylene</td><td align="left">5</td></tr><tr><td align="left">Female dog (10–50 lb)</td><td align="left">Flexible vinyl, red rubber, or Polyethylene</td><td align="left">8</td></tr><tr><td align="left">Female dog(>50 lb)</td><td align="left">Flexible vinyl, red rubber, or Polyethylene</td><td align="left">10,12, or 14</td></tr></tbody></table><table-wrap-foot><fn id="cetablefn6"><label>*</label><p id="cenotep6">The diameter of urinary catheters is measured on the French (F) scale. One French unit equals roughly 0.33 mm.</p></fn></table-wrap-foot><attrib>From Crow S, Walshaw S: Manual of Clinical Procedures in the Dog, Cat and Rabbit, ed 2, Philadelphia, Lippincott-Raven, 1997.</attrib><permissions><copyright-statement>© 2006 Lippincott-Raven</copyright-statement><copyright-year>2006</copyright-year><license><license-p>Since January 2020 Elsevier has created a COVID-19 resource centre with free information in English and Mandarin on the novel coronavirus COVID-19. The COVID-19 resource centre is hosted on Elsevier Connect, the company's public news and information website. Elsevier hereby grants permission to make all its COVID-19-related research that is available on the COVID-19 resource centre - including this research content - immediately available in PubMed Central and other publicly funded repositories, such as the WHO COVID database with rights for unrestricted research re-use and analyses in any form or by any means with acknowledgement of the original source. These permissions are granted for free by Elsevier for as long as the COVID-19 resource centre remains active.</license-p></license></permissions></table-wrap></p><sec id="cesec96"><title>Catheterization of the male dog</title><p id="para391">Equipment needed to catheterize a male dog includes a sterile catheter (4F to 10F, 18 inches long, with one end adapted to fit a syringe), sterile lubricating jelly, povidine-iodine soap or benzalkonium chloride, sterile rubber gloves or a sterile hemostat, a 20-mL sterile syringe, and an appropriate receptacle for the collection of urine.</p><p id="para392">Proper catheterization of the male dog requires two persons. Place the dog in lateral recumbency on either side. Pull the rear leg that is on top forward, and then flex it (<xref rid="f13" ref-type="fig">Figure 4-13</xref>). Alternatively, long-legged dogs can be catheterized easily in a standing position.<fig id="f13"><label>Figure 4-13</label><caption><p>Technique for catheterizing the urinary bladder of a male dog.</p></caption><graphic xlink:href="gr13"></graphic></fig></p><p id="para393">Next, retract the sheath of the penis and cleanse the glans penis with a solution of povidone-iodine 1%, triclosan (Septisol), benzalkonium chloride, or bichloride of mercury solution diluted 1:1000. Lubricate the distal 2 to 3 cm of the appropriate-size catheter with sterile lubricating jelly. Never entirely remove the catheter from its container while it is being passed because the container enables one to hold the catheter without contaminating it. The catheter may be passed with sterile gloved hands or by using a sterile hemostat to grasp the catheter and pass it into the urethra. Alternatively, cut a 2-inch “butterfly” section from the end of the thin plastic catheter container. This section can be used as a cover for the sterile catheter, and the clinician can use the cover to grasp and advance the catheter without using gloves.</p><p id="para394">If the catheter cannot be passed into the bladder, the tip of the catheter may be caught in a mucosal fold of the urethra or there may be a stricture or block in the urethra. In small breeds of dogs, the size of the groove in the os penis may limit the size of the catheter that can be passed. One also may experience difficulty in passing the catheter through the urethra where the urethra curves around the ischial arch. Occasionally, a catheter of small diameter may kink and bend on being passed into the urethra. When the catheter cannot be passed on the first try, reevaluate the size of the catheter and gently rotate the catheter while passing it a second time. Never force the catheter through the urethral orifice.</p><p id="para395">Effective catheterization is indicated by the flow of urine at the end of the catheter, and a sterile 20-mL syringe is used to aspirate the urine from the bladder. Walk the dog immediately following catheterization to encourage urination.</p></sec><sec id="cesec97"><title>Catheterization of the female dog</title><p id="para396">Equipment needed to catheterize a female dog includes flexible human ureteral or urethral catheters identical to those used in the male dog. Sterile metal or plastic female catheters also can be used; however, they tend to traumatize the urethra. The following materials also should be on hand: a small nasal speculum, a 20-mL sterile syringe, lidocaine 0.5%, sterile lubricating jelly, a focal source of light, appropriate receptacles for urine collection, and 5 mL of povidone-iodine solution.</p><p id="para397">Use strict asepsis. Cleanse the vulva with a solution of povidone-iodine, benzalkonium chloride, or bichloride of mercury diluted 1:1000. Instillation of lidocaine 0.5% into the vaginal vault helps to relieve the discomfort of catheterization. The external urethral orifice is 3 to 5 cm cranial to the ventral commissure of the vulva. In many instances, the female dog may be catheterized in the standing position by passing the female catheter into the vaginal vault, despite the fact that the urethral tubercle is not visualized directly.</p><p id="para398">In the spayed female dog in which blind catheterization may be difficult, the use of an otoscope speculum and light source (<xref rid="f14" ref-type="fig">Figure 4-14</xref>) or an anal speculum with a light source will help to visualize the urethral tubercle on the floor of the vagina. In difficult catheterizations, it may be helpful to place the animal in dorsal recumbency (<xref rid="f15" ref-type="fig">Figure 4-15</xref>, <xref rid="f16" ref-type="fig">Figure 4-16</xref>). Insertion of a speculum into the vagina almost always permits visualization of the urethral tubercle and facilitates passage of the catheter. Take care to avoid attempts to pass the catheter into the fossa of the clitoris because this is a blind, possibly contaminated cul-de-sac.<fig id="f14"><label>Figure 4-14</label><caption><p>An otoscope speculum with attached light source provides excellent visualization of the urethral orifice in a female dog. Note the position of the otoscope handle (see <xref rid="f15" ref-type="fig">Figure 4-15</xref>).</p></caption><graphic xlink:href="gr14"></graphic></fig><fig id="f15"><label>Figure 4-15</label><caption><p>Visualization of the urethral orifice and catheterization of the urinary bladder in a female dog is accomplished using an otoscope with a sterile speculum attached. Note: the patient is in dorsal recumbency with the otoscope handle positioned upwards.</p></caption><graphic xlink:href="gr15"></graphic></fig><fig id="f16"><label>Figure 4-16</label><caption><p>Technique for catheterizing the urinary bladder of a female cat using an otoscope speculum and light source.</p></caption><graphic xlink:href="gr16"></graphic></fig></p></sec><sec id="cesec98"><title>Catheterization of the male cat</title><p id="para399">Before attempting urinary bladder catheterization of the male cat, administer a short-term anesthetic (e.g., ketamine, 25 mg/kg IM), but only after a careful assessment of the cat's physical status. Males cats with urethral obstruction (partial or complete) may have underlying renal or hepatic disease that precludes the use of a dissociative anesthestic. In such patients, administration of an inhalation anesthetic only may be required. In addition, a topical anesthetic such as a topical ophthalmic anesthetic solution (proparacaine) can be administered directly into the urethral orifice to minimize discomfort. Place the anesthetized patient in dorsal recumbency. Gently grasp the ventral aspect of the prepuce and move it caudally in such a manner that the penis is extruded. Withdraw the penis from the sheath and gently pull the penis backward. Pass a sterile, flexible plastic or polyethylene (PE 60 to 90) catheter or 3- to 5-inch, 3.5F urethral catheter into the urethral orifice and gently into the bladder, keeping the catheter parallel to the vertebral column of the cat. Never force the catheter through the urethra. The presence of concretions within the urethral lumen may require the injection of 3 to 5 mL of sterile saline to back-flush urinary “sand” or concretions so that the catheter can be passed.</p></sec><sec id="cesec99"><title>Catheterization of the female cat</title><p id="para400">Urinary bladder catheterization of the female cat is not a simple procedure. However, when indicated, only attempt the technique in the anesthetized cat using the same technique described for the male cat. Take care to assess the health of the patient before administering the anesthetic. Urinary bladder catheterization can be accomplished with the use of a rubber or plastic, side-hole (blunt-ended) urinary catheter. The same catheter type used in male cats is effective in female cats. Instilling lidocaine 0.5% has been recommended as a means of decreasing sensitivity to the required manipulations and catheter insertion. However, if the cat is anesthetized adequately, this additional step is neither helpful nor necessary. Cleanse the vulva with an appropriate antiseptic. Catheterization can be accomplished with the cat in dorsal or ventral recumbency. Experience and size of the cat dictates which technique works best. Because the likelihood of successfully catheterizing a female cat's urinary bladder using a blind technique is low, a speculum is strongly recommended. However, there are only limited types of small-diameter speculums that are suitable. An inexpensive technique entails use of a sterilized otoscope speculum, with attached light source to facilitate visualization of the urethral orifice (<xref rid="f16" ref-type="fig">Figure 4-16</xref>). Insert the catheter through the speculum and into the urethra. The most significant drawback to this technique comes when using the male cat, polyurethane urinary catheter. The diameter of the otoscope speculum may not allow withdrawal of the speculum over the expanded end of the urinary catheter. A soft, rubber catheter, however, can be pulled through even the smallest otoscope speculum and is recommended when necessary to remove the speculum completely and leave the catheter in place. Once the urethral orifice can be visualized, pass the catheter into the orifice until urine flow is established.</p></sec><sec id="cesec100"><title>Indwelling urethral catheter</title><p id="para401">For continuous urine drainage, use a closed collection system to help prevent urinary tract infection. A soft urethral or Foley catheter can be used, and polyvinyl chloride tubing should be connected to the catheter and to the collection bottle outside the cage. The collection bottle should be below the level of the animal's urinary bladder. Place an Elizabethan collar on the animal to discourage chewing on the catheter and associated tubing. Apply antibacterial ointment to the urethral orifice. Despite care of the catheter, urinary tract infection still may develop in any patient fitted with an indwelling urinary catheter. Ideally, replace catheters after 48 to 72 hours. Prophylactic antimicrobial therapy is indicated in any dog or cat in which a urinary catheter is in place for more than 48 hours. Observe the animal for development of fever, discomfort, pyuria, or other evidence of urinary tract infection. If infection is suspect, remove the catheter and submit the catheter tip for culture and sensitivity or determination of minimum inhibitory concentration (MIC).</p></sec><sec id="cesec101"><title>Cystocentesis</title><p id="para402">Cystocentesis is a common clinical technique used to obtain a sample of urine directly from the urinary bladder of dogs and cats <italic>when collecting a voided, or free-catch, aliquot is not preferred.</italic> The procedure is indicated when necessary to obtain bladder urine for culture purposes. Urine that is collected by free catch has passed through the urethra and may be contaminated with bacteria, thereby making interpretation of the culture results difficult. Cystocentesis also is performed as a convenience when it is desirable to obtain a small sample of urine but the patient is not ready or cooperative.</p><p id="para403">Generally, cycstocentesis is a safe procedure, assuming the patient is cooperative and the bladder can be identified and stabilized throughout the procedure. However, injury and adverse reactions can occur. In addition to lacerating the bladder with the inserted needle (patient moves abruptly), the needle can be passed completely through the bladder and into the colon (improper technique) risking bacterial contamination of the bladder or peritoneal cavity.</p><p id="para404">Cystocentesis involves insertion of a needle, with a 6- or 12-mL syringe attached, through the abdominal wall and bladder wall to obtain urine samples for urinalysis or bacterial culture. The technique prevents contamination of urine by urethra, genital tract, or skin and reduces the risk of obtaining a contaminated sample. Cystocentesis also may be needed to decompress a severely overdistended bladder temporarily in an animal with urethral obstruction. In these cases, cystocentesis should be performed only if urethral catheterization is impossible. WARNING: Penetration of a distended urinary bladder with a needle could result in rupture of the bladder.</p><p id="para405">To perform cystocentesis, clip and surgically prepare the skin over the cystocentesis site on the ventral abdomen. Perform cystocentesis by placing the needle in the ventral abdominal wall slightly (3 to 5 cm) cranial to the junction of the bladder with the urethra. Insert the needle at a 45-degree angle (<xref rid="f17" ref-type="fig">Figure 4-17</xref>). The bladder must contain a sufficient volume of urine to permit palpation through the abdominal wall before cystocentesis. Use one hand to hold the bladder steady within the peritoneal cavity while the other guides the needle.<fig id="f17"><label>Figure 4-17</label><caption><p>Technique for performing cystocentesis in a dog.</p></caption><graphic xlink:href="gr17"></graphic></fig></p><p id="para406">Although this procedure is relatively safe, the bladder must have a reasonable volume of urine, the procedure should not be made without first identifying and immobilizing the bladder. For the procedure to be performed safely and quickly, the patient <italic>must</italic> be cooperative. If collection of a urine sample by cystocentesis is absolutely necessary, sedation may be indicated to restrain the patient adequately for the procedure.</p><sec id="cesec102"><sec id="cesec103"><title>Additional Reading</title><p id="para407">Crow S, Walshaw S: <italic>Manual of clinical procedures in the dog, cat, and rabbit,</italic> ed 2, Philadelphia, 1997, Lippincott-Raven.</p><p id="para408">Osborne CA, Finco DR: <italic>Canine and feline nephrology and urology,</italic> Baltimore, 1995, Williams & Wilkins.</p></sec></sec></sec></sec></sec><sec id="cesec104"><title>DERMATOLOGIC PROCEDURES</title><sec id="cesec105"><title>Skin Biopsy</title><p id="para409">Obtaining a skin biopsy from abnormal skin only to receive a nondiagnostic result as reported from a pathologist suggests that improved biopsy technique may culminate in collecting a specimen with higher diagnostic value. The following guidelines apply when performing skin biopsies:<list list-type="simple" id="celist17"><list-item id="celistitem91"><label>•</label><p id="para410">Consider obtaining multiple samples from multiple sites, which is especially useful when different stages of similar lesions are identifiable.</p></list-item><list-item id="celistitem92"><label>•</label><p id="para411">Do <italic>not</italic> perform a surgical scrub before collecting the sample; shaving the hair away is fine, but surgically prepared skin removes superficial lesions that, had they been left in place, might have been diagnostic.</p></list-item><list-item id="celistitem93"><label>•</label><p id="para412">Biopsy of lesions that are <italic>depigmenting</italic> should be done before they have turned white; the absence of color usually denotes absence of active skin lesions; biopsies from completely depigmented skin are less likely to demonstrate active lesions.</p></list-item><list-item id="celistitem94"><label>•</label><p id="para413">Biopsy of lesions associated with alopecia should be done in the center of the most alopecic area.</p></list-item><list-item id="celistitem95"><label>•</label><p id="para414">Also, biopsy of lesions associated with alopecia should be done at junctional (between normal- and abnormal-appearing) skin.</p></list-item><list-item id="celistitem96"><label>•</label><p id="para415">Consider submitting biopsies from completely nonaffect, normal-appearing skin.</p></list-item><list-item id="celistitem97"><label>•</label><p id="para416">Avoid biopsies from ulcerated skin areas.</p></list-item></list></p><p id="para417">Biopsies may be made with a scalpel blade (incisional or excisional) or a dermatologic punch biopsy. Punch biopsy instruments are circular blades available in 4-mm, 6-mm, and 8-mm diameter sizes (<xref rid="f18" ref-type="fig">Figure 4-18</xref>). Hold the punch perpendicular to the skin site of interest. A back-and-forth motion that rotates the circular blade cuts through the skin. When the skin no longer moves as the punch is rotated, the biopsy is complete and the skin sample may be removed (from the skin or from the biopsy instrument). Avoid grasping the dermis/epidermis of the sample with any instrument to prevent crushing of the sample and causing artifact. If the sample must be lifted, use the attached subcutaneous fat only.<fig id="f18"><label>Figure 4-18</label><caption><p>Disposable skin biopsy punches: 4-, 6-, and 8-mm sizes are available.</p></caption><graphic xlink:href="gr18"></graphic></fig></p><p id="para418">If the lesion of interest is deep, the punch biopsy technique may not be effective. In this situation, an incisional or excisional biopsy using a sterile No. 10 or No. 15 surgical blade is indicated. Biopsies of ulcerated skin and solitary nodules are best done by removing a wedge of skin (incisional biopsy). In some cases, it is possible surgically to remove all visible, palpable parts of the lesion (excisional biopsy). Place each sample of skin in buffered formalin, using a volume that is at least 10 times that of the sample size. If particularly large areas of skin are harvested during biopsy, cut these into 1-cm thick pieces before placing them into formalin.</p><p id="para419">Alternatively, it is possible, and in many cases important, to evaluate a biopsy of skin or subcutaneous tissue at the time of collection. When the lesion of interest is suspected to be neoplastic, quickly differentiating between inflammatory cells and neoplasia may be possible by simply performing an exfoliative cytologic examination (see Section 5) on one of the biopsy samples <italic>in addition to</italic> fixing a separate sample in formalin and submitting it for histopathologic examination. Exfoliated cells on a glass slide are air-dried and stained with a quick Romanowsky's-type stain (e.g., Diff-Quick). Generally, biopsy samples that have been subjected to the additional handling required to make impressions on a glass slide are not good candidates for subsequent fixation and histopathologic examination. One strong recommendation is to perform exfoliative cytologic and histopathologic examinations on separate samples.</p></sec><sec id="cesec106"><title>Skin Scraping</title><sec id="cesec107"><title>Superficial skin scraping</title><p id="para420">Among the most common diagnostic procedures carried out on the skin of dogs and cats is a routine skin scraping. Yet despite the frequency this test is used, doing a skin scraping in such a manner that the sample recovered maximizes the opportunity to establish a diagnosis can be anything but routine. A skin scraping, properly done, does require using consistent techniques appropriate to the suspected diagnoses, and as such, superficial or deep scrapings, or both, may be indicated.</p><p id="para421">Skin scraping is indicated whenever ectoparasite infestation is suspected. Superficial scrapings are appropriate for detecting mites that live on the skin surface, such as <italic>Cheyletiella</italic> spp. and <italic>Otodectes cynotis,</italic> as well as those mites that burrow within the outermost layers of skin (stratum corneum), such as <italic>Sarcoptes</italic> spp. and <italic>Notoedres cati.</italic></p><p id="para422">Because the area to be scraped is relatively large (≥2 cm<sup>2</sup>), shave dogs and cats with long-hair coats before attempting the procedure, unless <italic>Chyletiella</italic> infestation is suspected. Make the scraping over healthy-appearing skin. Do <italic>not</italic> cleanse the skin of superficial scale or crusts. The technique for superficial skin scraping entails the use of mineral oil or pyrethrin ear drops applied to a clean scalpel blade <italic>and</italic> directly onto the area of skin to be scraped. Scraping begins as a gentle motion made in the direction of the hair coat. Gradually increase the pressure of the blade against the skin with repetitive scrapings over the same area. Take care not to lacerate the skin, although minor capillary bleeding at the site is common. Transpose material collected on the edge of the blade to a clean glass slide, cover it with a coverslip, and thoroughly examine the material under low magnification for evidence of ectoparasites. Note that for mites such as <italic>Chyletiella</italic> or scabies, finding just one mite or one egg is diagnostic and justifies implementing treatment.</p></sec><sec id="cesec108"><title>Deep skin scraping</title><p id="para423">A slightly different technique is indicated in dogs and cats suspected of having an infestation that includes <italic>Demodex canis</italic> mites. The mites are known to live predominantly in sebaceous glands and hair follicles. They can survive in the skin of animals without manifesting lesions. Hair loss and skin lesions develop where overgrowth of the mite population occurs. <italic>Demodex</italic> infestations can be localized or generalized; infestations can occur in either dogs or cats but the most severe, generalized infestations are much more likely to occur in young dogs.</p><p id="para424">Although both superficial and deep skin scrapings may reveal the presence of mites on the skin, deep scrapings may reveal <italic>Demodex</italic> mites in some patients when superficial scrapings are negative. The technique for deep skin scraping targets a small area of skin (≤2 cm<sup>2</sup>). It may be helpful to apply gentle pressure to the skin or actually to squeeze the area of interest between the thumb and a finger in at attempt to force mites from the deeper to the more superficial skin. In some breeds (e.g., Old English Sheepdogs and shar peis) recovering mites on a skin scraping can be particularly difficult. In such cases, where <italic>Demodex</italic> infestation is highly suspected but the results of repeated skin scrapings are negative, a skin biopsy is appropriate. Alternatively, a procedure called a <italic>trichogram</italic> that involves pulling (plucking) a few hairs from the hair follicles using a hemostat may be diagnostic. Once the hairs are plucked from the skin, place them on a glass slide that has been preprepared with a drop of mineral oil, add a coverslip, and examine the hair shaft under low magnification. Half of all dogs with <italic>Demodex</italic> infestation will have a positive trichogram.</p><sec id="cesec109"><title>Additional Reading</title><p id="para425">Baker R, Lumsden JH: The skin. In Baker R, Lumsden JH, editors: <italic>Color atlas of cytology of the dog and cat,</italic> St Louis, 2000, Mosby.</p><p id="para426">Bettenay SV, Mueller RS: Skin scraping and skin biopsies. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p><p id="para427">Campbell KL: Other external parasites. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine</italic>, ed 6, St Louis, 2005, Elsevier-Saunders.</p></sec></sec></sec></sec><sec id="cesec110"><title>EAR CLEANING: EXTERNAL EAR CANAL</title><p id="para428">Certainly not all dogs and cats with otitis externa require comprehensive ear flushing and debridement before or as part of the therapy. In many cases, home treatment is sufficient to resolve the problems effectively, assuming the underlying diagnosis has been established. However, in patients with chronic or particularly severe infections, topical treatment administered by the owners at home may not be sufficient. In such cases, the external ear canal requires a careful and comprehensive cleaning before administration of topical medications.</p><p id="para429">Properly perfomed, flushing and cleaning of the external ear canals is not a quick procedure. Anesthetize the patient. Attempting to perform a thorough ear cleansing under sedation usually will not be successful. Once the animal is anesthetized, perform a careful otoscopic (or videootoscopic) examination to establish the integrity of the ear canal, such as the presence or absence of tumors and mites. In severe cases, the tympanic membrane may not even be visible.</p><p id="para430">With the patient in lateral recumbency, flush the ear canal (<xref rid="f19" ref-type="fig">Figure 4-19</xref>) or lavage it with warm saline initially, and then aspirate the material from the canal. If this procedure is not successful in removing the debris attached to the epithelium of the ear canal, use ceruminolytic ear solutions to facilitate breakdown and removal of this material. A 5-minute instillation and soak is recommended, followed by thorough flushing to remove debris and the ceruminolytic material. Remove hair growing inside the ear canal with forceps. A suction apparatus is recommended for removal of debris and liquid remaining.<fig id="f19"><label>Figure 4-19</label><caption><p>Low-pressure water jet system used to flush the external ear canal in anesthetized patients.</p></caption><graphic xlink:href="gr19"></graphic></fig></p><p id="para431">Reintroduce an otoscope to examine the integrity of the skin in the ear canal and to look for any evidence of stenosis, foreign body, or tumor. The flushing process is <italic>not</italic> complete until it is possible to visualize the tympanic membrane. Carefully remove any remaining debris with an otologic loop (<xref rid="f20" ref-type="fig">Figure 4-20</xref>), not a cotton-tipped swab.<fig id="f20"><label>Figure 4-20</label><caption><p><bold>A,</bold> Otologic loops used to remove debris from the external ear canal. <bold>B,</bold> Placement of an otologic loop through the specululm of an otoscope facilitates removal of debris deep in the external ear canal.</p></caption><graphic xlink:href="gr20"></graphic></fig></p><p id="para432">Repeat the procedure on the opposite ear as indicated. At the conclusion of the examination, apply appropriate topical medication into the ear canal before allowing the patient to recover from anesthesia. Systemic therapy or surgical intervention may be required in some patients for complete resolution of the problem. However, a thorough examination and cleaning is critical before actually making decisions regarding medical versus surgical intervention.</p><sec id="cesec111"><sec id="cesec112"><sec id="cesec113"><sec id="cesec114"><title>Additional Reading</title><p id="para433">Gortel K: Ear flushing. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p><p id="para434">Taibo RA: <italic>Otology: clinical and surgical issues,</italic> Buenos Aires, 2003, Intermedica.</p></sec></sec></sec></sec></sec><sec id="cesec115"><title>ENDOTRACHEAL INTUBATION</title><p id="para435">In selecting an appropriate-sized endotracheal tube, consider the size of the animal and select the size of tube that has the largest diameter that can be inserted without force (<xref rid="cetable7" ref-type="table">Table 4-7</xref>). The use of high-volume, low-pressure cuffs on endotracheal tubes is better. Overinflation of the endotracheal tube cuff can cause tracheal ulceration, tracheitis, hemorrhage, tracheomalacia, fibrosis, stenosis, and subcutaneous emphysema. Occlusion of high-volume, low-pressure cuffs can be achieved at 25 mm Hg or less.<table-wrap position="float" id="cetable7"><label>TABLE 4-7</label><caption><p>Recommended Sizes for Endotracheal Tubes</p></caption><table frame="hsides" rules="groups"><thead><tr><th></th><th align="center">Body weight (kg)</th><th align="center">Magill size</th><th align="center">French size</th><th align="center">Internal diameter (mm)</th></tr></thead><tbody><tr><td align="left" rowspan="8">Dogs</td><td align="center">2</td><td align="center">2</td><td align="center">22</td><td align="center">6</td></tr><tr><td align="center">4</td><td align="center">4–5</td><td align="center">26–28</td><td align="center">8</td></tr><tr><td align="center">6</td><td align="center">6–7</td><td align="center">28–30</td><td align="center">9</td></tr><tr><td align="center">9</td><td align="center">8</td><td align="center">32</td><td align="center">10</td></tr><tr><td align="center">12</td><td align="center">9–10</td><td align="center">34–36</td><td align="center">11–12</td></tr><tr><td align="center">14</td><td align="center">9–10</td><td align="center">34–36</td><td align="center">11–12</td></tr><tr><td align="center">16</td><td align="center">10–11</td><td align="center">36–38</td><td align="center">11–12</td></tr><tr><td align="center">18–20</td><td align="center">11–12</td><td align="center">38–44</td><td align="center">12</td></tr><tr><td align="left" rowspan="3">Cats</td><td align="center">1</td><td align="center">00</td><td align="center">13</td><td align="center">4</td></tr><tr><td align="center">2</td><td align="center">0</td><td align="center">16</td><td align="center">5</td></tr><tr><td align="center">4</td><td align="center">1</td><td align="center">20</td><td align="center">5</td></tr></tbody></table></table-wrap></p><p id="para436">Always check the cuff of a cuffed tube to ensure that there are no leaks and that the cuff is working properly before intubation. Lubricate the selected endotracheal tube with water or lubricating jelly. Do not intubate the animal until after induction of anesthetic drugs. Intubation in the dog or cat may cause an increase in sympathetic activity or vagal stimulation and result in cardiac dysrhythmias. Administer atropine or glycopyrrolate to canine and feline patients to avoid dysrhythmias associated with intubation.</p><p id="para437">Following appropriate administration of anesthetic and parasympatholytic drugs, place the patient in lateral or sternal recumbancy and elevate the head. Measure the tube from the mouth to the level of the carina. The person inserting the tube should pull out the tongue, holding the tongue with a piece of gauze to improve handling and stability. Use caution to not lacerate the tongue on the lower incisors. Place the tip of a laryngoscope at the base of the tongue at the glossoepiglottic fold. Use small pediatric blades for cats and small dogs and larger blades for larger dogs. Press the tip of the laryngoscope ventrally to move the epiglottis and expose the glottis. Directly visualize the arytenoid cartilages, and then pass the tube through the arytenoid cartilages into the trachea using a slight twisting motion. If the arytenoid cartilage spasms shut during attempt at intubation, place a drop or two of 2% lidocaine on the arytenoid cartilages to help facilitate passing the tube. Once inserted into the trachea, never advance the tube farther than the carina, or else one-lung (endobronchial) intubation can occur. Once the tube is in place, secure it in place with a loop of ½-inch white tape or muzzle gauze.</p><sec id="cesec116"><sec id="cesec117"><sec id="cesec118"><sec id="cesec119"><title>Additional Reading</title><p id="para438">Muir W, Hubbell J, Skarda R, et al: <italic>Handbook of veterinary anesthesia,</italic> ed 3, St Louis, 2000, Mosby.</p></sec></sec></sec></sec></sec><sec id="cesec120"><title>INTRAVENOUS CATHETERIZATION</title><p id="para439">Contamination and infection at the catheterization site are common complications of indwelling venous catheters. Aseptic preparation of the site is therefore paramount. In emergency situations it may be necessary to perform catheterization under less than ideal circumstances and without time for adequate aseptic preparation. When the animal's condition is stabilized, remove the catheter and place it properly elsewhere.</p><p id="para440">Clip the hair or shave over a wide area to facilitate disinfection of the skin surface. Scrub the skin surface with a detergent solution for 1 to 2 minutes. Then remove the detergent with an iodine or alcohol solution, and spray the skin with an iodine-based solution. If it is necessary to maintain the aseptic conditions of the procedure, wear sterile gloves and drape the field with sterile towels or a fenestrated drape.</p><p id="para441">When placing the catheter, fix it in position. Do not allow the catheter to move in and out of the skin because this will predispose to mechanical vessel trauma and the introduction of bacteria. When the puncture site is a large flat surface such as the medial femoral area or neck, apply a tag of tape to the catheter at a point close to the puncture site and then suture the tape to the skin. Pass the adhesive tape around the entire circumference of the catheter, and then tape the catheter to the appendage. Additional tape may be applied to isolate the injection site from the underlying skin and to secure the catheter cap. Place an antibiotic-antifungal ointment at the puncture site and incorporate it into the dressing over the skin.</p><sec id="cesec121"><title>Percutaneous Jugular Vein Catheterization</title><p id="para442">The immobilization procedure for the jugular vein is particularly important because of the tendency of the vein to roll in the loose subcutaneous tissues of the neck. Occlude the vein by digital pressure in the thoracic inlet so that the skin and underlying tissues are retracted toward the body. Extend the head to provide traction on the upper portion of the vein. All of this positioning is accomplished by a second person.</p><p id="para443">A catheter-inside-the-needle system is easy to maintain sterile, but the needle leaves a larger hole in the vessel than is filled by the remaining catheter, and early postcatheterization hemorrhage may be a problem. Locate the position of the vessel with one hand, and insert the needle subcutaneously with the other hand. Place the needle directly over the jugular vein and below the palpating index finger. Advance the needle steeply at first to secure the superficial wall of the vessel and then more parallel to the vein to allow insertion of the needle into the lumen without penetrating the deep wall of the vessel. Ascertain the position of the needle within the vein by feeling the needle “pop” through the vessel wall and seeing the reflux of blood into the catheter. If, after the initial plunge, the needle is not in the vessel lumen, withdraw the needle slowly. Penetration of the needle into the deep wall of the vein is a common occurrence. Constant traction on the syringe plunger will aspirate blood into the catheter if the needle passes back through the vessel lumen during withdrawal.</p><p id="para444">When it has been ascertained that the needle is within the lumen, gently thread the catheter into the vein without moving the vein or the needle. Insert the catheter to its full length. If the catheter cannot be advanced fully, consider that (1) the tip of the needle may not have been entirely within the vessel lumen, (2) the vessel and needle may have moved with respect to one another during the initial threading process, (3) the catheter may be caught at the thoracic flexure (change the position of the head), or (4) the catheter may be caught in one of the tributaries to the front legs (change the position of the front leg). Exercise care when withdrawing the catheter through the needle. The catheter can catch on the needle bevel and be cut off, resulting in catheter embolization.</p><p id="para445">When the catheter has been inserted to the desired length, remove the needle from the skin and place the needle guard. Aspirate air from the catheter, and then flush the catheter with heparinized saline. If blood cannot be aspirated from the catheter, consider that (l) the catheter may be kinked or compressed by the needle guard, (2) the catheter tip may be against a vessel wall (withdraw the catheter slightly), (3) the catheter may be clotted (if the clot cannot be aspirated, remove the catheter), or (4) the catheter may not be in the vein.</p></sec><sec id="cesec122"><title>Surgical Approach to Jugular Vein Catheterization</title><p id="para446">Make a skin incision over or next to the jugular vein. The normal jugular vein is superficial and usually rises with venous occlusion. Location of the jugular vein following repeated unsuccessful attempts at percutaneous catheterization is more difficult because of the resultant hematoma. The vein is always located in the middle of the hematoma, and there is a temptation to skirt the hematoma with the dissection procedure. The vein can be identified during blunt dissection by the appearance of an off-white longitudinal structure against the dark background of the hematoma.</p><p id="para447">Isolate the vein and suspend it by two sutures. Visually introduce the needle into the lumen of the vein, and take care not to penetrate the deep wall of the vessel. Insert the catheter and relax the proximal suture to allow passage of the catheter. Remove both stay sutures (the vein is not tied), and close the incision in a routine manner.</p></sec><sec id="cesec123"><title>Maintenance of Indwelling Catheters</title><p id="para448">Re-dress indwelling catheters and inspect them every day. Discard all soiled bandage material. Clean the puncture site with antiseptic solutions and fresh antibiotic-antifungal ointment, and reapply the occlusive wrap. Inspect the skin puncture site and the vessel at each re-dressing. A very small ring of inflammation at the puncture site is normal. Excess inflammation, diffuse tissue swelling, expulsion of exudate from the puncture site upon palpation, and tenderness or pain upon palpation are signs of untoward effects of the indwelling catheter. Phlebitis may be caused by mechanical, chemical, or infectious irritation of the vein. Phlebitis is recognized as warm, erythematous skin overlying a tender, indurated vessel. Purulent thrombophlebitis is heralded by all of the signs of simple phlebitis plus free exudate, which may drain exteriorly with or without palpation or may drain internally and result in a severe septicemia. Thrombotic occlusion of the vessel is recognized by severe hardening (ropiness) of the vessel. Thromboembolic occlusion is associated with inability to infuse fluids by gravity and may result in severe subcutaneous fluid accumulation if the fluids are administered with a pump. Occult infection of the intravenous site may occur in the absence of local inflammatory, thrombotic, or exudative signs. Unexplained fever and leukocytosis may be the only early signs. The diagnosis may be confirmed by obtaining a culture of material from the catheter tip.</p><p id="para449">Remove the catheter if there is evidence of cellulitis, phlebitis, thrombosis, purulent thrombophlebitis, or catheter-associated bacteremia or septicemia; if the catheter ceases to function properly because of thrombosis or catheter occlusion by a clot or kink; after 3 days if there is another location to which it can be moved; if the patient begins to lick or chew at the bandages; and when it is no longer necessary (<xref rid="cetextbox5" ref-type="boxed-text">Box 4-5</xref>).<boxed-text id="cetextbox5"><label>BOX 4-5</label><caption><title>HEPARINIZED SALINE FLUSH</title></caption><p id="para40">Catheter patency can be maintained with 3-mL syringes preloaded with 2 to 3 mL of heparinized saline kept in a stock solution:</p><p id="para41">Add 100 units of heparin to a single 50-mL bag of sterile saline (2 units heparin/ml saline).</p></boxed-text></p><p id="para450">Infusion fluids and administration tubing must be sterile. Do not disconnect connections unless it is absolutely necessary, and then they must be disconnected aseptically. Clean all injection caps well with an antiseptic solution before needle insertion. Change the fluid bottles and all administration tubing every 1 to 2 days. Change tubing after blood or colloid infusion. Do not use the primary catheter or infusion line for the collection of blood samples except in emergencies. Clearly mark the fluid bottle if any drug or concentrate has been added to the bottle. In-line filters are not necessary for infection control for routine fluid administration.</p></sec></sec><sec id="cesec124"><title>PHYSICAL THERAPY</title><sec id="cesec125"><title>Hydrotherapy</title><p id="para451">Water baths or soaks are among the easiest and most versatile modes of physical therapy in small animal practice. Wet packs, water soaks, or whirlpool baths can be helpful in adding moisture to (hydrating) or removing moisture from (dehydrating) the skin. Cyclic repetitions of moistening and drying the skin many times a day serve to dehydrate (similar to the chapped lips and hands seen in human beings). Constant moisture hydrates and even macerates the skin.</p><p id="para452">Whirlpool baths are the most efficient and popular means of applying hydrotherapy and, often, antiseptic medication to the skin. Whirlpool baths combine moist heat, gentle massage, and the solvent properties of water, with or without the mechanical impact of water from a whirlpool, to remove dirt, pus, and necrotic debris. Dry or scaly skin is softened and moisturized. Whirlpools may increase edema and are contraindicated in acute traumatic and inflammatory conditions or in cases of impaired sensation or circulation (unless treated with extreme caution). Whirlpools are particularly beneficial for skin infections, chronic dryness, open or infected wounds, skin grafts, adhesions, arthritis, postsurgical fractures, amputations, muscle spasms, and stiff joints. This modality is especially useful for cleaning and stimulating the skin of patients that are predisposed to decubitus ulcers.</p><p id="para453">Place the patient in an appropriate water bath at a temperature of 39° to 42° C (102° to 108° F). A low-suds detergent or antiseptic solution (povidone-iodine, chlorine, chlorhexidine) can be added for cleansing or germicidal effects. Allow the water turbine to circulate the water around and against the affected parts for 10 to 15 minutes once or twice daily. Support and reassure the animal during treatment, and never leave an animal unattended. Following therapy, ensure that the tub and turbine are thoroughly cleaned and sanitized.</p><p id="para454">Commercial whirlpool baths or Jacuzzi-type agitators are a good investment for a busy practice. A less expensive alternative is a variable-temperature bath with agitation provided by a pressure hose.</p></sec><sec id="cesec126"><title>Heat Therapy</title><p id="para455">NOTE: In the presence of trauma, swelling, and edema, circulation may be impeded and the application of heat may cause necrosis. Cold is more beneficial in the early acute stages of inflammation and edema (<xref rid="cetextbox6" ref-type="boxed-text">Box 4-6</xref>).<boxed-text id="cetextbox6"><label>BOX 4-6</label><caption><title>THE BENEFITS OF HEAT IN PHYSICAL THERAPY</title></caption><p id="para42"><list list-type="simple" id="celist1"><list-item id="celistitem1"><label>•</label><p id="para43">Hyperemia and dilatation of cutaneous vessels</p></list-item><list-item id="celistitem2"><label>•</label><p id="para44">Increase in pulse, blood pressure, and pulmonary ventilation</p></list-item><list-item id="celistitem3"><label>•</label><p id="para45">Increased metabolite transfer across capillary membranes</p></list-item><list-item id="celistitem4"><label>•</label><p id="para46">General muscle relaxation</p></list-item><list-item id="celistitem5"><label>•</label><p id="para47">Sedative and analgesic effect</p></list-item><list-item id="celistitem6"><label>•</label><p id="para48">Improved extensibility of connective tissue</p></list-item></list></p></boxed-text></p><sec id="cesec127"><title>Superficial heat</title><p id="para456"><italic>Infrared radiation</italic> is produced by long-wave generators that glow red and produce heat that penetrates only 1 to 2 mm. Shortwave generators produce invisible light; their heat penetrates 10 to 12 mm and reaches blood-carrying layers of tissue, where the heat is dispersed by the circulation. Because of their penetration, short waves do not produce a burning sensation; however, one should check the skin meticulously during therapy to ensure that it is not overheated. A warm sensation is normal. If the skin feels hot to the touch, the heat is too intense. Shortwave therapy is contraindicated in acute trauma and inflammation. Superficial heat is beneficial for subacute and chronic problems, skin infections, and abscesses. Treat the animal for 15 to 20 minutes once or twice daily.</p><p id="para457"><italic>Hot packs</italic> or wet towels can be used to apply mild, gentle heat. The indications are the same as for infrared heat, but additional precautions are needed. Hot packs may spread contagious skin disease, and the weight of the packs in an insensitive area is more likely to cause burns or tissue damage. This modality has the advantage of providing moist heat, which is particularly beneficial for chronic soft tissue problems such as arthritis, myositis, and contractures. Apply hot packs for 10 to 15 minutes several times a day. Check the skin under the packs frequently during therapy.</p><p id="para458"><italic>Whirlpool baths</italic> combine moist heat, gentle massage, and the solvent properties of water to remove dirt, pus, and necrotic debris. The moist heat is indicated especially for extensive involvement of musculoskeletal disorders. Warm water baths can be repeated 2 to 3 times daily for 10 to 15 minutes and often can be followed with a gentle massage and passive flexion and extension exercises to improve range-of-motion and soft tissue flexibility.</p></sec><sec id="cesec128"><title>Deep heat</title><p id="para459"><italic>Shortwave diathermy</italic> transmits physical energy deep into tissues; because the body tissues resist the flow of high-frequency current (27 million cps [Hz]), heat is produced. In dissipating heat, there are marked vascular dilatation, sedation, analgesia, and relief of muscle spasm. However, edema may increase. Absolute contraindications are the presence of imbedded metal implants, ischemia, malignancy, and pregnancy. No water can be in the field, and splints and bandages must be removed. This therapy can result in electrical shock to the patient and technician, so all cables, electrodes, and other equipment must be in excellent condition. Safety cannot be overemphasized. Calibrate each unit and adjust units individually to produce only a sensation of warmth. Treatment is applied daily for 15 to 20 minutes.</p><p id="para460"><italic>Microwave techniques</italic> produce about the same effects as diathermy except that more localized heating occurs (one side of a joint may be treated at a time). Microwaves are more readily absorbed by water, so great care must be used around the eye or in the presence of edema to ensure that the effects are not excessive.</p><p id="para461"><italic>Ultrasound</italic> produces the deepest heat. Ultrasound produces mechanical vibrations (1 million/second) in the elastic media of the body. Ultrasonic waves are reflected from boundaries between different types of tissues. The vibrations produce a micromassage that accelerates fluid absorption by increasing permeability. Ultrasonic therapy can be dangerous if the intensity or application is concentrated too long in a small area. Burn or tissue destruction may result. Ultrasonic therapy is contraindicated in neoplasms, the eye, heart, spine, and brain; near growing bony epiphyses; and in acute infections. Otherwise, its beneficial effects are similar to those of other forms of deep heat. Ultrasound is indicated particularly for softening scar tissue and reducing the pain of neuromas and degenerative joint disease. Ultrasound can be used over and around metal implants.</p><p id="para462">Ultrasonic waves are applied via a transducer and using coupling medium such as water or contact gel. The transducer is moved constantly over a small area (usually 6 to 8 sq in, depending on the size of the transducer). Shaving of the skin before therapy is best, to enhance contact (or use a water bath).</p><p id="para463">The dose varies with each patient. Most ultrasonic generators have an output of 700,000 to 1 million Hz at intensities of 0.1 to 1.0 W/cm<sup>2</sup>. Use of the lowest intensity possible is best. The maximum dose should be 1.0 W/cm<sup>2</sup> for 5 minutes of application to the affected tissues. This application can be repeated once daily for 5 days and then every other day for five treatments. Then the treatment should not be repeated for at least 1 month. Do not use ultrasonic therapy in acute injuries, inflammations, or infections.</p><p id="para464">For cervical intervertebral disease, use 0.3 W/cm<sup>2</sup> for 3 minutes daily for 5 days and then every other day for five treatments.</p><p id="para465">For arthritis, bursitis, and myositis, use 0.2 W/cm<sup>2</sup> for 3 minutes for joints of the extremities. Repeat 2 times weekly.</p></sec><sec id="cesec129"><title>Cold therapy</title><p id="para466">Cold can be applied by blowing cold air on the skin, by evaporation of volatile liquids from the skin, or by direct contact of the cooling substance with the skin surface (<xref rid="cetextbox7" ref-type="boxed-text">Box 4-7</xref>).<boxed-text id="cetextbox7"><label>BOX 4-7</label><caption><title>THE BENEFITS OF COLD IN PHYSICAL THERAPY</title></caption><p id="para49"><list list-type="simple" id="celist2"><list-item id="celistitem7"><label>•</label><p id="para50">Decreased tissue temperature</p></list-item><list-item id="celistitem8"><label>•</label><p id="para51">Decreased blood flow, vasoconstriction</p></list-item><list-item id="celistitem9"><label>•</label><p id="para52">Decreased tendency to edema</p></list-item><list-item id="celistitem10"><label>•</label><p id="para53">Decreased delivery of nutrients, phagocytes</p></list-item><list-item id="celistitem11"><label>•</label><p id="para54">Decreased phagocytic action</p></list-item><list-item id="celistitem12"><label>•</label><p id="para55">Transient vasoconstriction followed by vasodilation and increased blood flow (brief cold applications)</p></list-item></list></p></boxed-text></p><p id="para467">Cold reduces extravasation of blood and fluid into tissues after trauma, reduces pain and spasticity, and is indicated in acute traumatic and inflammatory conditions. Overtreatment with cold may produce maceration and frostbite. Prolonged cold produces a vascular response with stasis of blood, occlusion of vessels, and tissue anoxia and necrosis.</p><p id="para468"><italic>Cold packs</italic> over a damp towel applied to the affected area and covered with a folded dry towel to prevent rapid warming can be used for 15 to 20 minutes. Treatment is repeated several times daily. It is important to keep the rest of the patient's body warm, dry, and comfortable during treatment. Cold treatments are often more effective when alternated with heat treatments (immersion bath or moist warm packs). The combined treatment is most effective when heat for 3 to 4 minutes is alternated with 1 to 2 minutes of cold. This regimen can be repeated for 15 to 20 minutes and should always end with the hot phase.</p><p id="para469"><italic>Cold immersion baths</italic> for one or several extremities may be useful. The temperature should be 15.5° to 21° C (60° to 70° F) and can be decreased by adding ice or cold water. Continue treatment until the muscles are relaxed or the animal cannot tolerate the cold, usually 2 to 5 minutes. Modification of this technique can be used for heat stroke, but one must be careful not to overchill such patients.</p></sec></sec><sec id="cesec130"><title>Electrotherapy</title><p id="para470"><italic>Medical galvanism</italic> is the physiologic use of direct current. This treatment will produce the same effects as heat except that it has no tendency to produce edema. The treatment is beneficial for acute, subacute, or chronic traumatic and inflammatory problems such as arthritis, decubitus sores, neuralgia, tenosynovitis, or postfracture repair. Do not use electricity near the brain, heart, or neoplasms.</p><p id="para471">Low-intensity therapy (0.5 to 1.0 mA/sq in of electrode) is desirable. Electrodes should be wet and held in firm contact with the skin. No metal should be in or near the area being treated. Halfway through the treatment, reduce the intensity of current to zero, reverse the polarity, and the return the intensity to starting levels.</p><p id="para472"><italic>Electrical stimulation</italic> of partially or wholly innervated muscle is possible with alternating current. Intact or denervated muscle will contract when stimulated with interrupted pulses of direct current. These kinds of stimulation improve circulation and nutrition of the muscle, promote venous return, remove lymph, relax spasm, reduce edema, and assist in muscle reeducation. Muscle atrophy and weakness can be retarded or controlled. Each muscle or group of muscles can be stimulated 10 to 20 times for one procedure, depending on the condition. Avoid overtreatment.</p></sec><sec id="cesec131"><title>Massage Therapy</title><p id="para473">Massage is the use of the hands and fingers to manipulate soft tissues. Massage usually is used in combination with heat, cold, or whirlpool treatments. Massage improves circulation, reduces edema, loosens and stretches fibrotic or contracted tissue, and has a soothing or sedative effect. Do not use massage in acute, inflammatory, traumatic, and painful lesions or with tumors, hemorrhages, and perhaps contagious conditions. Massage is indicated for tight or contracted tendons, ligaments, or muscles; chronic traumatic or inflammatory problems; and subacute or chronic edema.</p><p id="para474">In performing massage, keep the strokes in the direction of venous flow. Firm, rapid pressure tends to be stimulating, whereas slow, light strokes are soothing. Some type of lubricating powder or oil can be used to reduce friction. The massage can be stroking, kneading, or applied with friction. Stroking and kneading assist circulation, whereas friction and kneading tend to loosen adhesions and scars and to stretch tissues. Massage should last 15 or 20 minutes and can be repeated several times daily if desired.</p></sec><sec id="cesec132"><title>Exercise Therapy</title><p id="para475">Therapeutic exercise should strengthen musculoskeletal function, improve range-of-motion flexibility, improve endurance or coordination, and increase cardiovascular and respiratory capabilities. Never force exercise, but keep it within safe tolerance of the patient's cardiac and respiratory capacity. Active exercise (such as walking, running, or swimming) is most desirable because endurance and strength increase with repetition. Passive exercise is useful when paralysis or traumatic injuries preclude active exercise.</p><p id="para476">Never force movements, but use stabilization of parts and controlled pressure to activate only the structures of concern. When attempting to increase range of motion, use smooth, controlled pressure to move the joint slightly beyond its limited range; hold the stretch for a count of five, and slowly release the traction. Several repetitions can be performed 2 or 3 times daily. Gradual improvement can be expected within several weeks.</p></sec></sec></sec><sec id="cesec133"><title>ADVANCED PROCEDURES</title><sec id="cesec134"><title>ABDOMINOCENTESIS</title><p id="para477">Abdominal paracentesis refers to the surgical puncture of the abdominal cavity for the purpose of removing fluids. Always weigh the animal before and after removing abdominal fluid. Any subsequent gain in weight indicates a reaccumulation of abdominal fluid. Place the animal in left lateral recumbency and restrain it in this position. Clip and surgically prepare a 1- to 3-inch square between the bladder and the umbilicus just lateral to the midline. If the bladder is distended, empty it before performing paracentesis. Infiltrate the paracentesis site with lidocaine 0.5% using a 22- to 25-gauge needle. In most cases, local anesthesia is not necessary. Abdominal puncture can be made with an 18- to 20-gauge needle. When the abdominal puncture has been made, allow the animal to rest quietly to facilitate drainage of the fluid. Some clinicians recommend tapping while the patient is in a standing position in the hope of obtaining more complete drainage. Changing the patient's position after the tapping may result in needle-tip laceration of intraabdominal organs. Aspiration may be easier if a specially adapted needle with multiple holes drilled in the shaft is used because it is less likely to become plugged with omentum. Ideally, tap four quadrants of the abdomen. Single-needle taps are not as accurate as instilling a lavage fluid (warmed lactated Ringer's solution) into the abdomen and examining the lavage fluid. Measure the amount of fluid obtained, and examine the fluid to determine whether it is an exudate or a transudate. Cytologic examination and culture also may be performed. Rather than drain the abdominal fluid completely, it may be better to spare protein loss and mobilize the fluids with diuretics. Paracentesis also can be performed by using a sterile intravenous catheter to enter the abdomen. When performing paracentesis, ultrasonographic guidance can prove valuable in placing the needle into the compartmental space desired and in avoiding complications. The major complications in abdominal paracentesis are perforated hollow viscus, laceration of abdominal organs, and iatrogenic peritonitis.</p><sec id="cesec135"><sec id="cesec136"><sec id="cesec137"><sec id="cesec138"><title>Additional Reading</title><p id="para478">Meyer DJ, Harvey J: <italic>Veterinary laboratory medicine,</italic> ed 3, St Louis, 2004, Saunders.</p><p id="para479">Rudloff E: Abdominocentesis and diagnostic peritoneal lavage. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine</italic>, ed 6, St Louis, 2005, Elsevier-Saunders.</p></sec></sec></sec></sec></sec><sec id="cesec139"><title>BIOPSY TECHNIQUES: ADVANCED</title><p id="para480">Numerous biopsy techniques are available, and the selection of the appropriate technique is based on the tissue to be examined, the condition of the patient, and the skill of the examiner.</p><p id="para481"><italic>Excisional biopsy</italic> refers to the surgical removal of the entire lesion/organ with subsequent histologic examination. Excisional biopsy is used most frequently for skin lesions and cases in which an entire organ may have to be removed (such as an eye or an internal organ that has developed a tumor). <italic>Incisional biopsy</italic> refers to the surgical removal of a <italic>portion</italic> of a lesion with subsequent histologic examination. Choose a representative area of the lesion for biopsy. Include lesion margins, if possible. <italic>Needle aspiration</italic> refers to the use of needle and syringe to remove representative cells from the tissue/organ of interest. Specialized needles are available that allow removal of very small biopsies that can be submitted for histopathologic examination. (See also Cytopathologic Specimen Collection Techniques.)</p><sec id="cesec140"><title>Needle Biopsy Techniques: General Considerations</title><p id="para482">Needle biopsy or aspiration techniques refer to a variety of techniques used to obtain diagnostic tissue or cells from internal organs, including the lung, liver, spleen, pancreas, abdominal lymph nodes and mass lesions within the abdomen and thorax. In contrast, fine-needle aspiration is a technique generally used to recover cytopathologic samples (cells only) from skin or subcutaneous tissues (e.g., superficial lymph nodes). The advantage of needle biopsy is related directly to how well the abnormal tissue has been characterized and how easily it can be identified during the procedure. In addition, depending on patient cooperation, most procedures can be performed safely with the patient sedated only. Short-term intravenous anesthesia and general anesthesia eliminate undesired patient movement during the biopsy technique.</p><p id="para483">Potential lesions or abnormal tissues from which aspirate or biopsy samples are to be taken are located using palpation, radiographs, or ultrasound-guided imaging techniques. Shave the skin over the site of needle penetration and surgically prepare it. The type of sedation or anesthesia depends on the temperament of the animal and the site on which the biopsy will be performed. Attach a 22-gauge needle without stylet to a 12-mL syringe prefilled with 0.5 to 1.0 mL of air. Optionally, affix a flexible extension set to the needle and connect it proximally to the syringe. Needle length may vary from 1 to 3½ inches depending on the required depth of penetration and size of the patient. Guide the needle into the tissue/organ of interest. Stabilize the tip of the needle to avoid random movements through organs, especially highly vascular tissue such as liver and spleen. Once the needle is inserted, the aspiration techniques entails withdrawing the plunger of the syringe to the 7- or 8-mL level. Holding for that position for 1 to 2 seconds, and then releasing. Repeat the procedure. Depending on the nature of the lesion, it may not be indicated to thrust the needle into the tissue at multiple and different angles.</p><p id="para484">Neutralize the pressure in the syringe, and withdraw the needle rapidly. Expel any material within the needle onto glass slides using the air in the syringe. This same procedure can be repeated with a new needle to obtain an additional three to five samples from alternative sites. This technique allows samples to be obtained without applying negative pressure to the syringe, which may damage cells.</p><p id="para485">Ultrasound-guided needle aspirations from abdominal tissues greatly enhance the safety of this technique, especially when obtaining samples from smaller animals. Automatic-trigger needles such as Cook or Temno biopsy needles (14 to 18 gauge) are available for use in human beings but are seldom used in veterinary medicine. The risks associated with fine-needle aspiration include rupture of an encapsulated inflammatory process, dissemination of an infectious agent, seeding of neoplastic cells in the needle tract, and hemorrhage. Larger volumes of fluid and cells can be placed directly into a vial containing EDTA to prevent clot formation. Prepare and examine direct and sedimentation specimens.</p><p id="para486">Needle biopsy of internal organs using the Tru-Cut needle is particularly useful in patients with subcutaneous (<xref rid="f21" ref-type="fig">Figure 4-21</xref>) or cutaneous masses and for localized abdominal and thoracic mass lesions, diffuse liver, kidney, and splenic disease. Serious complications, usually hemorrhage or laceration of the gall bladder (during liver biopsy), can occur when the procedure is performed blindly. Therefore ultrasound-guided needle biopsy is strongly recommended whenever a percutaneous biopsy of internal organs is performed. Additional safety factors provided by ultrasound guidance include the ability to image, and avoid, large aberrant blood vessels.<fig id="f21"><label>Figure 4-21</label><caption><p>Mechanism of action of Tru-Cut biopsy needle for typical nodular biopsy. A small skin incision is made with a No. 11 blade to allow insertion of the instrument. <bold>A,</bold> With the instrument closed, the outer capsule is penetrated. <bold>B,</bold> The outer cannula is fixed in place, and the inner cannula with specimen notch is thrust into the tumor. Tissue then fills the notch. <bold>C,</bold> The inner cannula now is fixed while the outer cannula is moved forward to cut off the biopsy specimen. <bold>D,</bold> The entire instrument is removed. <bold>E,</bold> The inner cannula is pushed ahead to expose tissue in the specimen notch.</p><p>Rights were not granted to include this figure in electronic media. Please refer to the printed book.</p></caption><graphic xlink:href="figure4-21"></graphic><attrib>(From Withrow SJ, Lowes N: Biopsy techniques in small animal oncology, <italic>J Am Anim Hosp Assoc</italic> 14:899-902, 1981.)</attrib><permissions><copyright-statement>© 2006 </copyright-statement><copyright-year>2006</copyright-year><license><license-p>Since January 2020 Elsevier has created a COVID-19 resource centre with free information in English and Mandarin on the novel coronavirus COVID-19. The COVID-19 resource centre is hosted on Elsevier Connect, the company's public news and information website. Elsevier hereby grants permission to make all its COVID-19-related research that is available on the COVID-19 resource centre - including this research content - immediately available in PubMed Central and other publicly funded repositories, such as the WHO COVID database with rights for unrestricted research re-use and analyses in any form or by any means with acknowledgement of the original source. These permissions are granted for free by Elsevier for as long as the COVID-19 resource centre remains active.</license-p></license></permissions></fig></p><p id="para487">Risk of complications associated with needle aspiration of the lung is considerably higher than for most abdominal procedures. Pneumothorax can occur following a single, “clean” aspiration attempt. See RESPIRATORY TRACT PROCEDURES for a detailed description of performing fine-needle aspiration of lung.</p><sec id="cesec141"><sec id="cesec142"><sec id="cesec143"><title>Additional Reading</title><p id="para488">Lumsden JH, Baker R: Cytopathology techniques and interpretation. In Baker R, Lumsden JH, editors: <italic>Color atlas of cytology of the dog and cat,</italic> St Louis, 2000, Mosby.</p><p id="para489">MacNeill AL, Alleman AR: Cytology of internal organs. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p><p id="para490">Menard M, Papgeorges M: Ultrasound-guided liver fine needle biopsies in cats: results of 307 cases, <italic>Vet Pathol</italic> 33:570, 1996.</p><p id="para491">Menard M, Papgeorges M: Fine needle biopsies: how to increase diagnostic yield, <italic>Compend Contin Educ Pract Vet</italic> 19:738, 1997.</p><p id="para492">Meyer D, Harvey J: <italic>Veterinary laboratory medicine,</italic> ed 3, St Louis, 2004, Elsevier-Saunders.</p></sec></sec></sec></sec><sec id="cesec144"><title>Skin Biopsy</title><p id="para493">Histologic examination of diseased skin can serve as a means for diagnosis of cutaneous lesions. The causative agent often is found in acute and chronic skin infections. Punch biopsy of the skin is a quick and accurate means of removing a small sample of diseased skin for histopathologic examination. Select a site that is well developed but not traumatized or excoriated. The sample should include little or no normal tissue. If the lesion (pustule, vesicle) can be identified early in its development and if the biopsy sample is taken only from the lesion, one may obtain a superior specimen. It is best not to take too large a sample that contains much normal skin; by mistake, the technician might take a section that misses the lesion. Proper selection of the biopsy site is crucial to accurate diagnosis. Carefully clip the hair from the lesion. Lightly blot the skin with 70% alcohol. Avoid superficial trauma while cleaning the skin. Inject a small subcutaneous bleb of 2.0% lidocaine to deaden the area. Special equipment needed for the biopsy includes a 4-mm, 6-mm, or 8-mm biopsy punch and 10% buffered formalin solution. After the area has been anesthetized with lidocaine, press and rotate the biopsy punch through the skin until the subcutaneous tissue is penetrated. Remove the biopsy specimen by “spearing” the subcutaneous fat with a fine needle. Do not grasp the specimen with a forceps. Blot the specimen gently between two paper towels. Spread the tissue out gently (like a pancake), place the specimen epidermal side up on a piece of cardboard or tongue depressor, press the specimen gently to cause adhesion, and drop the specimen into the formalin fixative. The skin defect may be closed with one or two simple interrupted sutures. If deep subcutaneous tissue or large biopsy samples are needed, a punch biopsy is inadequate. Use a small (No. 15) scalpel blade to obtain an appropriate sample. In all cases in which skin biopsies are made, take <italic>multiple</italic> samples to increase the odds that at least one will have diagnostic lesions. Specimens submitted to laboratories should be accompanied by extensive, detailed clinical information, including a differential diagnosis. Skin biopsies routinely are stained with hematoxylin-eosin; however, periodic acid–Schiff, Gomori's methenamine silver, and Verhoeff's stains are used for special problems.</p></sec><sec id="cesec145"><title>Liver Biopsy</title><p id="para494">The diagnosis of liver disease can be made based on clinical signs coupled with clinicopathologic results obtained by laboratory finding, radiography, and abdominal ultrasound. The development of a more specific diagnosis and prognosis in liver disease may be aided greatly by information obtained in a liver biopsy. Percutaneous liver biopsies are of much greater value in generalized liver disease such as cirrhosis, generalized acute hepatic necrosis, or amyloidosis than in focal hepatic disease. The major indications for performing a liver biopsy are (l) to explain an abnormal liver profile, (2) to define reasons for abnormal liver size, (3) to identify a possible liver tumor, (4) to obtain a prognosis and rational approach to management, and (5) to identify the cause of ascites.</p><p id="para495">The procedures for obtaining liver tissue are numerous; however, needle biopsy of the liver, when performed properly, can be helpful. Careful physical and clinicopathologic examination should precede a liver biopsy. A normal coagulation profile should be documented on every patient undergoing liver biopsy. Detect and correct abnormalities in normal hemostatic mechanisms, if feasible, before needle biopsy of the liver.</p><p id="para496"><boxed-text id="cetextbox24"><caption><title>Note:</title></caption><p id="para497">The liver biopsy, although a critical diagnostic tool in patients with laboratory evidence of liver disease, can be a fatal event, even in the hands of the experienced clinician. Abdominal ultrasound imaging of the liver before and during biopsy is strongly recommended.</p></boxed-text></p><sec id="cesec146"><title>Technique</title><p id="para498">Percutaneous needle biopsies and fine-needle aspirations of the liver are performed routinely with local anesthesia in the sedated patients. General anesthesia is a reasonable alternative when feasible. Biopsy sites in the liver can be selected best when needle biopsy techniques are used along with laparoscopy or ultrasound techniques. Blind percutaneous needle biopsies of the liver can be performed with relative safety if the liver is significantly enlarged and easily palpated. However, blind biopsies do carry the risk that the operator is unable to determine the impact of penetrating the liver if only an abdominal radiograph and impression of abdominal palpation are available. In cases where the liver is <italic>not</italic> palpable, blind biopsy carries significantly higher risk and should be performed only when no alternative exits.</p><p id="para499">A modified percutaneous liver biopsy can be performed by the following method. Before biopsy, have the animal fast, and remove any ascitic fluid. Place the animal in dorsal recumbency, and place a local block in the midline of the skin and abdomen at the caudoventral aspect of the left hepatic lobe. The incision into the peritoneal cavity should be large enough to accommodate the gloved index finger. Make a separate skin puncture site in the abdominal wall to accommodate the biopsy needle. Use the index finger manually to fix the left hepatic lobe (or other desired hepatic lobe) against the diaphragm or other adjacent structures, and insert the outer cannula and stylet through the abdominal wall in the isolated hepatic lobe. Remove the stylet, and rapidly insert the cutting prongs. If properly placed, the cutting prongs should not go through the entire hepatic lobe. Advance the outer cannula over the blades of the cutting prongs, thus entrapping the hepatic tissue material within the cutting prongs. Remove the biopsy needle. Using a wooden applicator stick, carefully place the biopsy specimen into fixative. Biopsy samples can be used to prepare slides for cytologic examination, and the biopsy needle may be cultured. Close the abdominal incision in the routine manner.</p><p id="para500">Another liver biopsy technique uses the Tru-Cut biopsy needle. Place the dog in dorsal recumbency. Clip a 5-cm<sup>2</sup> area over the triangle formed by the xiphoid cartilage and left costal arch, and prepare the area as for aseptic surgery. Make a small paramedian incision large enough to accommodate a sterile otoscope head 7 mm in diameter. Use a halogen-illuminated otoscope head to visualize the liver. Pass a Tru-Cut biopsy needle through the otoscope cone to obtain a biopsy specimen of the liver.</p></sec></sec><sec id="cesec147"><title>Nasal Biopsy</title><p id="para501">Chronic nasal discharge, with or without sneezing, represents the single most common indication for nasal biopsy. Affected patients should have the benefit of nasal radiographs or a computed tomography study of the nose in advance of endoscopy and/or biopsy. Once the justification for biopsy has been established, the clinician can use one of at least two different techniques to obtain samples.</p><p id="para502">An older technique still in use today entails the use of a polypropylene catheter with an angled (45-degree), pointed cut on the tip to achieve a blind biopsy (<xref rid="f22" ref-type="fig">Figure 4-22</xref>). With the patient anesthetized, measure the distance from the tip of the nose to the eyes externally and mark the catheter at the level of the external nares (this is the “DON'T GO BEYOND HERE” mark). Gently pass the catheter into the nasal cavity until it will not pass further, reaches the “DON'T GO BEYOND HERE” mark, or reaches a depth consistent with that of a lesion demonstrated radiographically. The procedure entails performing a nasal flush and aspiration of fluid for cytopathologic examination or forcing the catheter tip blindly into the tissue, or both. A syringe on the end of the catheter facilitates removal of tissue from the nasal cavity by aspiration. Some of the tissue may be suitable for histopathologic examination.<fig id="f22"><label>Figure 4-22</label><caption><p>Illustration depicting the technique for obtaining a blind biopsy from the nasal cavity of a dog using a polypropylene catheter with an angled tip.</p></caption><graphic xlink:href="gr22"></graphic></fig></p><p id="para503">Alternatively, pass small biopsy forceps (<xref rid="f23" ref-type="fig">Figure 4-23</xref>) into the affected nostril(s). These instruments allow the collection of several small biopsies (usually up to 0.5 cm diameter) from which exfoliative cytologic examination can be performed, or the sample can be submitted for histopathologic examination.<fig id="f23"><label>Figure 4-23</label><caption><p>Small biopsy forceps used to obtain nasal biopsies.</p></caption><graphic xlink:href="gr23"></graphic></fig></p><p id="para504">Rhinoscopy equipment is expensive but does offer the advantage of being able to examine most (sometimes all) of the nasal cavity and actually to visualize an intranasal lesion—sometimes. However, even in the event it is possible to visualize a lesion, simultaneously obtaining the biopsy specimen while observing through the rhinoscope is problematic. Because of the limited amount of working space, most nasal biopsies ultimately are performed blindly.</p></sec><sec id="cesec148"><title>Renal Biopsy</title><p id="para505">Renal biopsies can be valuable in confirming or eliminating a diagnosis of renal disease that is based on history, physical examination, and radiographic and laboratory data (<xref rid="cetextbox8" ref-type="boxed-text">Box 4-8</xref>). In addition, biopsy may be a way of arriving at a prognosis in generalized renal disease and a better means of evaluating the type of treatment to be instituted. Ultrasonographic guidance can prove valuable during renal biopsy for placing the needle into the tissue desired and avoiding complications.<boxed-text id="cetextbox8"><label>BOX 4-8</label><caption><title>CONTRAINDICATIONS TO RENAL BIOPSY</title></caption><p id="para56"><list list-type="simple" id="celist3"><list-item id="celistitem13"><label>•</label><p id="para57">Coagulation abnormalities</p></list-item><list-item id="celistitem14"><label>•</label><p id="para58">A single functional kidney</p></list-item><list-item id="celistitem15"><label>•</label><p id="para59">Marked hydronephrosis</p></list-item><list-item id="celistitem16"><label>•</label><p id="para60">Greatly contracted kidneys</p></list-item><list-item id="celistitem17"><label>•</label><p id="para61">Acute pyelonephritis</p></list-item><list-item id="celistitem18"><label>•</label><p id="para62">Large cysts</p></list-item></list></p></boxed-text></p><p id="para506">Before renal biopsy, the animal should have a baseline coagulation profile that includes, at the very least, an activated coagulation time and platelet count. A buccal mucosal bleeding time may be indicated if there is any history of spontaneous bleeding in a patient with a normal platelet count. Obtain biopsies from the renal cortex. Administer fluids to patients before and after biopsy.</p><sec id="cesec149"><title>Technique</title><p id="para507">Many patients with generalized renal disease are critically ill and debilitated, and general anesthesia is contraindicated. In these cases, a neuroleptanalgesic agent may be used for sedation. If the animal is a good anesthetic risk and renal function will permit it, use inhalation anesthesia.</p><p id="para508">When bilateral renal disease is documented, select the LEFT kidney for biopsy because it is more accessible than the right kidney. With the anesthetized patient in right lateral recumbency, surgically prepare the skin behind and below the junction of the costal arch at the level of the second and third lumbar vertebrae. Make a 2-inch paralumbar incision parallel to, but just behind, the costal arch. Dissect muscle and fascia until the peritoneum is visible. Carefully open the peritoneal cavity. Digitally feel for and examine the caudal pole of the left kidney. Guide the needle toward the posterior pole of the kidney with the index finger. Immobilize the kidney against the body wall and insert the Tru-Cut biopsy needle, with the biopsy notch exposed into the parenchyma of the kidney. Capture the biopsy by sliding the outer sleeve of the needle over the (now imbedded in the kidney) biopsy notch. Remove the needle and gently lift the biopsy from the needle and place it into formalin. Evaluate the site for hemorrhage. Once bleeding is controlled, a second biopsy may be collected. Once bleeding from the biopsy site has stopped, the incision can be closed. In dogs, renal biopsy can be performed under ultrasound guidance using probes with channels for biopsy needle insertion.</p></sec></sec><sec id="cesec150"><title>Bone Biopsy</title><p id="para509">Evaluation of bone marrow is indicated in patients with evidence of persistently diminished cell counts of any or all cell lines (white blood cells, red blood cells, platelets) or evidence of morphologically abnormal cells in peripheral blood. Bone marrow aspiration and bone biopsy are extremely helpful but underused diagnostic procedures. The availability of inexpensive, high-quality biopsy needles makes these procedures safe and easy to perform (once experience is gained).</p><p id="para510">Conventional practice today is to obtain a bone marrow aspirate (cytopathologic examination) <italic>and</italic> a bone biopsy from the same patient during the same procedure when changes in the peripheral blood justify this level of diagnostic testing. Bone marrow aspiration technique is described in this section.</p><p id="para511">Two types of bone biopsy needles are available. The most commonly described procedure involves use of the Jamshidi biopsy needle, an 11- to 13-gauge needle that ranges in length from 5 to 10 cm (<xref rid="f11" ref-type="fig">Figure 4-11</xref>). The needle contains a stylet that extends beyond the needle tip by 3 to 4 mm. Because of the size of the Jamshidi needle, its use is limited to medium and large dogs. For bone biopsies in cats and small dogs, the author prefers to use the Illinois bone marrow aspiration needle (<xref rid="f10" ref-type="fig">Figure 10-10</xref>), which is a 15- to 18-gauge needle available in lengths ranging from 2.5 to 5.0 cm.</p><p id="para512">The patient usually is sedated or anesthetized for the procedure. Although some patients will tolerate this procedure when performed under local anesthesia only, the additional manipulation required to obtain a quality sample justifies sedation. In some cases, the patient is sufficiently obtunded that sedation is neither indicated nor required.</p><p id="para513">The technique is the same regardless of the needle used. Once the site is selected (usually the same sites selected for bone marrow aspiration: head of the humerus, wing of the ileum, ishial tuberosity, proximal femur), clip the hair and surgically prepare the skin. Make a small stab incision in the skin over the site selected. Pass the needle, with stylet in place, through the incision and subcutaneous tissues until the needle tip makes firm contact with bone. Advance the needle using steady, increasing pressure and stable rotation. Rotation, in this case, means rotating the needle back and forth to the left 180 degrees and then to the right 180 degrees. Once the needle is situated in the bone (about 0.5 cm penetration only), STOP. Carefully remove the stylet. Continue the penetration by gradually applying additional pressure and simultaneously rotating the needle.</p><p id="para514">The usual depth of penetration varies from 1 inch to as much as 3 inches. On reaching the desired depth, remove the needle by continuing to rotate as described but gradually withdrawing the needle from the bone. An obturator is provided to push the sample out of the bone. Place the core of bone directly into buffered formalin and submit it for histopathologic examination (decalicification will be required, which takes a little longer).</p><p id="para515">Some authors recommend carefully rolling the bone core across a glass slide (for cytopathologic examination) before placing the bone in formalin. I do not recommend this because additional handling of the biopsy sample can sufficiently disrupt the architecture of the tissue and compromise the quality of the biopsy (besides upsetting the pathologist). Note also, the needle can, with a little gentle manipulation, be reinserted into the hole from which the biopsy sample was obtained. Because the Illinois needle and the Jamshidi needle accommodate a syringe, it is possible to obtain (quickly, to prevent clotting) a bone marrow aspirate from the same site. Place that sample directly onto glass slides or (recommended) into 4% EDTA and mix it before making slides.</p><p id="para516">There are no specific requirements for postbiopsy care of the patient. Clean the blood from the skin using hydrogen peroxide; sutures generally are not required.</p></sec><sec id="cesec151"><title>Prostate Biopsy (See Urinary Tract Procedures)</title><sec id="cesec152"><sec id="cesec153"><sec id="cesec154"><title>Additional Reading</title><p id="para517">Acierno MJ, Labato MA: Rhinoscopy, nasal flushing, and biopsy. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p><p id="para518">Burkhard MJ, Meyer DJ: Invasive cytology of internal organs: cytology of the thorax and abdomen, <italic>Vet Clin North Am Small Anim Pract</italic> 6:103, 1996.</p><p id="para519">Crow S, Walshaw S: <italic>Manual of clinical procedures in the dog, cat, and rabbit,</italic> ed 2, Philadelphia, 1997, Lippincott-Raven.</p><p id="para520">Guilford WG, Center SA, Strombeck DR, et al: <italic>Strombeck's small animal gastroenterology,</italic> ed 3, Philadelphia, 1996, WB Saunders.</p><p id="para521">Kerwin S: Hepatic aspiration and biopsy techniques, <italic>Vet Clin North Am Small Anim Pract</italic> 25:275, 1995.</p><p id="para522">Osborne CA, Finco DR: <italic>Canine and feline nephrology and urology,</italic> Baltimore, 1995, Williams & Wilkins.</p><p id="para523">Stone E: Biopsy: Principles, technical considerations, and pitfalls, <italic>Vet Clin North Am Small Anim Pract</italic> 25:33, 1995.</p></sec></sec></sec></sec></sec><sec id="cesec155"><title>BLOOD GAS: ARTERIAL</title><p id="para524">The femoral and dorsal pedal arteries can be punctured to obtain an arterial blood sample for blood gas and electrolyte analyses. To obtain a sample from the femoral artery, place the patient in lateral recumbency and restrain the patient in a manner similar to that for a medial saphenous venipuncture. A 25-gauge needle affixed to a tuberculin syringe is preferred for arterial venipuncture. Prepare the tuberculin syringe by coating it with heparin and forcing all the heparin out except for that left in the hub of the needle. Pull back on the plunger of the syringe slightly to facilitate visualizing the point at which the artery is entered. Arterial blood initially will enter the syringe without the plunger being drawn back. After the proper equipment is assembled and the patient is sufficiently restrained, the individual collecting the arterial blood sample should palpate the medial aspect of the limb over the proximal medial femur until palpating the femoral pulse. Direct the needle at a 30- to 45-degree angle, inserting the needle slowly, watching for a flash of blood in the hub of the needle (<xref rid="f24" ref-type="fig">Figure 4-24</xref>). Gradually withdraw the plunger to facilitate blood entering the syringe. Collect 0.4 to 0.5 mL and immediately submit it for anaylsis.<fig id="f24"><label>Figure 4-24</label><caption><p>Technique for collecting arterial blood from the dorsal pedal artery of a dog.</p></caption><graphic xlink:href="gr24"></graphic></fig></p><p id="para525">To obtain blood from a dorsal pedal artery, place the patient in lateral recumbency and extend the rear limb as for a medial saphenous blood sample collection. The person obtaining the blood sample should pull the paw of the down leg in the nondominant hand toward his or her body, rotating the limb slightly in a medial direction to palpate the arterial pulse. Palpate the pulse in the dorsal pedal artery on the dorsomedial aspect of the tarsus. Gently insert the needle on a 30-degree angle into the artery, watching carefully for a flash of blood into the syringe. When the necessary amount of blood has filled the syringe, remove the needle and place pressure over the site of arterial puncture for a minimum of 2 minutes.</p><p id="para526">Evacuate excess air from the syringe and needle, and cap the needle with a red rubber stopper to prevent air from entering the needle and syringe. Place the sample on ice until analysis, if arterial blood gas analyses cannot be performed immediately.</p><sec id="cesec156"><title>Surgical Cutdown</title><p id="para527">In the event percutaneous access to a peripheral artery is not possible, the femoral artery can be isolated and prepared for surgical cutdown. Following appropriate aseptic skin preparation, make a 4- to 5-cm incision in the skin over the femoral artery. Find the caudal edge of the sartorius muscle by blunt dissection and then reflect it anteriorly to expose the underlying femoral artery, vein, and nerve. Taking care to avoid tearing any vessel branches, gently isolate up to 2 cm of the femoral artery from the surrounding fascia. Visually direct the needle into the artery at this point. Alternatively, catheterize the artery in the event repeated arterial samples are required. Elevate the femoral artery by preplacing two stay sutures beneath the artery and then elevating the vessel to the level of the skin. Insert a long catheter-over-the-needle system into the lumen of the artery without penetrating the deep wall. Gently insert the catheter into the vessel, remove the needle, and cap and flush the catheter. Close the incision and affix the catheter to the skin via a tape tag sutured to the skin.</p><sec id="cesec157"><sec id="cesec158"><sec id="cesec159"><title>Additional Reading</title><p id="para528">Davis H: Venous and arterial puncture. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p><p id="para529">Crow S, Walshaw S: <italic>Manual of clinical procedures in the dog, cat, and rabbit,</italic> ed 2, Philadelphia, 1997, Lippincott-Raven.</p><p id="para530">Shiroshita Y, Tanaka R, Shibazaki A, et al: Retrospective study of clinical complications occurring after arterial punctures in dogs: 111 cases, <italic>Vet Rec</italic> 146(1):16-19, 2000.</p></sec></sec></sec></sec></sec><sec id="cesec160"><title>CEREBROSPINAL FLUID COLLECTION</title><p id="para531">In the dog and the cat the preferred site for obtaining a diagnostic sample of cerebrospinal fluid (CSF) is the cerebellomedullary cistern (cisterna magna) at the level of the atlantooccipital articulation (between the back of the skull and the first cervical vertebra). Although the procedure is not commonly done in private practice, a number of infectious and noninfectious conditions justify performing the procedure. Collection of CSF is indicated in any patient suspected of having an infection (bacterial, fungal, viral, rickettsial, protozoal, and [uncommonly] parasitic) suspected of reaching the central nervous system. Several types of noninfectious causes of meningitis are described in dogs and cats. In dogs specifically, most causes of meningitis are idiopathic. Defined diseases include granulomatous meningoencephalitis, breed-specific meningitis (e.g., pug encephalitis), neoplasia, and unexplained seizure disorder, especially in a patient with a history of seizure activity that is increasing in frequency.</p><sec id="cesec161"><title>Contraindications</title><p id="para532">Among the reasons <italic>not</italic> to perform CSF collection, lack of experience perhaps ranks at the top (<xref rid="cetextbox9" ref-type="boxed-text">Box 4-9</xref>). Cerebrospinal fluid collection is not without some risk. Inadvertent penetration of the cervical spinal cord can culminate in acute death. Anatomic reasons for <italic>not</italic> performing CSF collection include congenital abnormalities involving malformations of the foramen magnum or suspected neural malformations in the region of the cisterna magna. Patients with fractures, dislocations, or subluxations of the occipital region of the skull or rostral cervical region, resulting in distortion of the brainstem, medulla, cervical cord, and any patient suspected of having brain herniation should not be subjected to the procedure.<boxed-text id="cetextbox9"><label>BOX 4-9</label><caption><title>COMPLICATIONS OF CEREBROSPINAL FLUID COLLECTION</title></caption><p id="para63"><list list-type="simple" id="celist4"><list-item id="celistitem19"><label>•</label><p id="para64">No cerebrospinal fluid obtained</p></list-item><list-item id="celistitem20"><label>•</label><p id="para65">Herniation of the brain</p></list-item><list-item id="celistitem21"><label>•</label><p id="para66">Contamination of the cerebrospinal fluid with blood</p></list-item><list-item id="celistitem22"><label>•</label><p id="para67">Needle penetration of the medulla or the rostral spinal cord</p></list-item><list-item id="celistitem23"><label>•</label><p id="para68">Infection of the central nervous system</p></list-item><list-item id="celistitem24"><label>•</label><p id="para69">Respiratory or cardiac arrest</p></list-item><list-item id="celistitem25"><label>•</label><p id="para70">Vestibular dysfunction</p></list-item><list-item id="celistitem26"><label>•</label><p id="para71">Paresis or paralysis</p></list-item></list></p></boxed-text></p></sec><sec id="cesec162"><title>Technique</title><p id="para533">To perform cisternal puncture, intubation and general anesthesia are required. Place the patient in lateral recumbency. Clip and surgically prepare the area of skin at least from the external occipital protuberance to the wings of the atlas and as wide apart as the distance between the ears. Position the animal with the prepared area at the edge of the table. Flex the head ventrally and maintain it at a right angle to the long axis of the neck. Various recommendations have been made regarding how to position the patient with respect to the individual performing the procedure. In the technique described, right-handed persons should position the patient in right-lateral recumbency. Left-handed persons should position the patient in left lateral recumbency.</p><p id="para534">To identify the site of needle penetration in the skin, the landmarks are a vertical line connecting the <italic>anterior edge</italic> of the first cervical vertebra and a horizontal line from the occipital protuberance that runs parallel to the spine. <italic>The point of intersection of the two lines represents the point of needle penetration.</italic> IMPORTANT: For consistency in performing this technique, as the needle penetrates the skin and muscle overlying the cisterna magna, pass it <italic>perpendicular to the skin</italic> and <italic>parallel to the floor throughout the procedure.</italic></p><p id="para535">Using a 2- to 2 ½-inch, 20- to 22-gauge spinal needle (with a stylet), advance the needle methodically and slowly toward the cisterna. The depth of penetration required to reach the cisterna varies considerably among patients of different sizes but can range from less than 1 inch to just over 2 inches. Generally, if the needle hits bone with superficial penetration, the position of the needle is too far caudal. If the needle hits bone after deep penetration (most common), the needle point is on the occipital bone and too far cranial. Only minor changes of a few millimeters generally are required to correct.</p><p id="para536">On penetration of the cisterna magna, the needle passes through the dura mater. At this point, there is usually a slight degree of resistance felt on the needle, but <italic>not</italic> always. Therefore, checking the position of the needle 3 to 5 times during penetration by withdrawing the sylet to assess for presence of CSF flow is recommended.</p><p id="para537">Once the flow of CSF is established, collect fluid for analysis by allowing the CSF to <italic>flow freely.</italic> DO NOT ASPIRATE FLUID FROM THE CISTERNA MAGNA BECAUSE THIS MAY CREATE SUFFICIENT NEGATIVE PRESSURE THAT HERNIATION COULD RESULT. Collect aliquots of CSF in two sterile collection tubes (e.g., red-topped blood collection tubes). Collect 0.5 to 2.0 mL in each tube and submit it for fluid analysis. The reason for collecting two aliquots of CSF is that if the cytologic examination of one tube suggests neutrophils (or bacteria), the second sample can be submitted for culture.</p><p id="para538">The presence of blood contamination is not unusual, particularly when multiple penetrations of the neck muscles have been required. As the flow of CSF is established, if it has a blood-tinged appearance, delay the collection for several drops. If clearing is apparent within a few drops, the blood likely represents contamination. If clearing does not occur after several drops, the blood may represent active bleeding into the central nervous system. Collecting CSF in EDTA is an option if the sample appears to contain significant amounts of blood. If, however, the flow from the spinal needle appears to be pure, dark blood, the needle likely has penetrated one of the large venous sinuses that course outside of and lateral to the spinal cord. This is a consequence of passing the needle perpendicular to the skin but not exactly parallel to the floor. Withdraw the needle and repeat the procedure using a new needle. If the technique is properly performed, clear CSF still can be obtained despite having collected what appears to be pure blood on a previous attempt.</p><sec id="cesec163"><sec id="cesec164"><sec id="cesec165"><title>Additional Reading</title><p id="para539">Anderson SM: Cerebrospinal fluid collection, myelography, and epidurals. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p><p id="para540">Meyer D, Harvey J: <italic>Veterinary laboratory medicine,</italic> ed 3, St Louis, 2004, Elsevier-Saunders.</p><p id="para541">Oliver J, Lorenz M, Kornegay J: <italic>Handbook of veterinary neurology,</italic> ed 4, St Louis, 2004, Elsevier-Saunders.</p></sec></sec></sec></sec></sec><sec id="cesec166"><title>ELECTROCARDIOGRAPHY</title><p id="para542">The electrocardiogram provides a fast, efficient way to obtain considerable data about a patient's cardiovascular status. Electrocardiography is a clinical test and must be correlated with clinical findings (<xref rid="cetextbox10" ref-type="boxed-text">Box 4-10</xref>). Keep in mind that an electrocardiogram measures only electrical activity of the heart as seen on the body surface at any one instant. Electrical disorders of the myocardium can be transient or intermittent and, as such, can be missed on a single electrocardiogram.<boxed-text id="cetextbox10"><label>BOX 4-10</label><caption><title>INDICATIONS FOR PERFORMING AN ELECTROCARDIOGRAM</title></caption><p id="para72"><list list-type="simple" id="celist5"><list-item id="celistitem27"><label>•</label><p id="para73">Detect enlargement of any of the cardiac chambers</p></list-item><list-item id="celistitem28"><label>•</label><p id="para74">Diagnose cardiac arrhythmia</p></list-item><list-item id="celistitem29"><label>•</label><p id="para75">Identify effects of electrolyte imbalances, especially potassium</p></list-item><list-item id="celistitem30"><label>•</label><p id="para76">Monitor response to and direct cardiac drug therapy</p></list-item><list-item id="celistitem31"><label>•</label><p id="para77">Develop prognoses (degree of change in heart function over time)</p></list-item></list></p></boxed-text></p><sec id="cesec167"><title>Interpretation of the Electrocardiogram</title><p id="para543">Read each electrocardiogram using a definite system. Begin by examining the lead II rhythm strip: Is there a P wave for every QRS complex? Is there a QRS complex for every P wave? Do all the P waves look alike? Do all the QRS complexes look alike? Are the P wave and QRS complex consistently related to each other?</p><p id="para544">If the answer to any of these questions is no, proceed to identify the abnormality. Next, determine the rate, rhythm, and wave character; that is, evaluate measurements of the P wave, PR interval, and QRS complex. Evaluate the ST segment and T wave and the QT interval. Use all leads to determine the axis and any miscellaneous criteria.</p></sec><sec id="cesec168"><title>Heart Rate</title><p id="para545">Depending on the type of electrocardiographic equipment used, there are several methods for determining <italic>heart rate</italic> from the electrocardiographic tracing. Many electrocardiographs compute the heart rate and print that on the tracing. However, in patients with a significant dysrhythmia, these calculations can be flawed and should be verified manually when a question exists. Small linear lines or demarcations at the top of the electrocardiogram paper can be used to determine the heart rate. At a paper speed of 50 mm/second, the time between adjacent marks is 1.5 seconds. By counting the number of QRS complexes (or R waves) between just two of these divisions and multiplying by 20 equals the heart rate in beats/minute (<xref rid="f25" ref-type="fig">Figure 4-25</xref>). For those inclined to higher mathematics, the heart rate also may be determined by counting the number of small squares between R waves (at a paper speed of 50 mm/second) and then dividing into 3000 (<xref rid="cetextbox11" ref-type="boxed-text">Box 4-11</xref>).<fig id="f25"><label>Figure 4-25</label><caption><p>Using the electrocardiogram to determine heart rate. The distance between R waves is 20 small boxes: 3000/20 = 150 beats/minute. (Paper speed is 50 mm/second.)</p></caption><graphic xlink:href="gr25"></graphic></fig><boxed-text id="cetextbox11"><label>BOX 4-11</label><caption><title>NORMAL HEART RATE</title></caption><sec id="cesec1"><title>Dog</title><p id="para78"><list list-type="simple" id="celist6"><list-item id="celistitem32"><label>•</label><p id="para79">Large dogs: 60 to 100 beats/minute</p></list-item><list-item id="celistitem33"><label>•</label><p id="para80">Medium-size dogs: 80 to 120 beats/minute</p></list-item><list-item id="celistitem34"><label>•</label><p id="para81">Small dogs: 90 to 140 beats/minute</p></list-item><list-item id="celistitem35"><label>•</label><p id="para82">Puppies: Up to 220 beats/minute</p></list-item></list></p></sec><sec id="cesec2"><title>Cat</title><p id="para83"><list list-type="simple" id="celist7"><list-item id="celistitem36"><label>•</label><p id="para84">Domestic cats: 140 to 250 beats/minute</p></list-item></list></p></sec></boxed-text></p></sec><sec id="cesec169"><title>Heart Rhythm</title><p id="para546">The normal heart rhythm is sinus in origin. For every QRS complex there is a P wave (<xref rid="f26" ref-type="fig">Figure 4-26</xref>). The P waves are related to QRS complexes (P-P interval is constant). Sinus arrhythmia, sinus arrest, and wandering pacemaker are normal rhythm variations. In sinus arrhythmia, the P-P interval is irregular. The pauses are never longer than twice the usual P-P interval (<xref rid="f27" ref-type="fig">Figure 4-27</xref>). A <italic>wandering pacemaker</italic> means that the P waves vary in height and may even be negative temporarily (<xref rid="f28" ref-type="fig">Figure 4-28</xref>). <italic>Sinus arrest</italic> is defined as a prolongation of the P-R interval longer than twice the usual P-P interval.<fig id="f26"><label>Figure 4-26</label><caption><p>Normal lead II QRS complexes in an adult dog.</p></caption><graphic xlink:href="gr26"></graphic></fig><fig id="f27"><label>Figure 4-27</label><caption><p><bold>A,</bold> Mild sinus arrhythmia in a dog. There are P waves for every QRS complex, and P waves are related to the QRS complexes, which make this a sinus rhythm. The variation of the R-R intervals also is visible. An irregular sinus rhythm is a sinus arrhythmia. <bold>B,</bold> Sinus arrhythmia in the cat.</p></caption><graphic xlink:href="gr27"></graphic><attrib>(From Edwards NF: <italic>Bolton's handbook of canine and feline electrocardiography,</italic> ed 2, Philadelphia, 1987, WB Saunders.)</attrib></fig><fig id="f28"><label>Figure 4-28</label><caption><p><bold>A,</bold> The wandering pacemaker in this recording is suggested by the slightly negative P waves in some of the complexes. Negative P waves of this nature result from vagal depression of the sinoatrial node and the development of a junctional atrioventricular nodal rhythm. <bold>B,</bold> Marked sinus arrhythmia and a wandering pacemaker result in a decreased heart rate (increased R-R interval) and negative P waves in the fifth complex. As the pacemaker returns to the sinoatrial node, the rate increases, and positive P waves of varying amplitude result in the sixth and seventh complexes.</p></caption><graphic xlink:href="gr28"></graphic></fig></p></sec><sec id="cesec170"><title>NORMAL ELECTROCARDIOGRAM MEASUREMENTS</title><sec id="cesec171"><title>P wave</title><p id="para547">The normal P wave is 0.04 second × 0.4 mV (two boxes wide × four boxes tall) for the dog and 0.04 second × 0.2 mV for the cat. In P mitrale (left atrial enlargement), the P wave is wider than 0.04 second. In P pulmonale (right atrial enlargement), the P wave is taller than 0.4 mV for the dog and 0.2 mV for the cat.</p></sec><sec id="cesec172"><title>PR interval</title><p id="para548">The PR interval is measured from the beginning of the P wave to the beginning of the QRS complex. The normal interval is 0.06 to 0.13 second (3 to 6.5 boxes wide) for the dog and 0.06 to 0.08 second for the cat. In first-degree atrioventricular heart block, the PR interval is prolonged. The PR interval is sometimes useful in monitoring the effects of digitalis therapy.</p></sec><sec id="cesec173"><title>QRS complex</title><p id="para549">The QRS complex duration is measured from the beginning of the Q wave to the end of the S wave. Normal duration is up to 0.04 second in cats, 0.05 second in small dogs, and 0.06 second in large dogs. A QRS complex that is too wide indicates left ventricular enlargement (<xref rid="f29" ref-type="fig">Figure 4-29</xref>). An R wave that is too tall indicates left ventricular enlargement. The amplitude is measured from the baseline to the top of the R wave (<xref rid="f30" ref-type="fig">Figure 4-30</xref>). The normal R wave can be up to 0.8 mV tall in cats, 2.5 mV in small dogs, and 3.0 mV in large dogs.<fig id="f29"><label>Figure 4-29</label><caption><p>In these two examples of left ventricular enlargement the QRS complexes have normal configuration but are too wide. <bold>A,</bold> This QRS complex from a miniature poodle is 0.07 second (three boxes) wide. <bold>B,</bold> This QRS complex from a Doberman Pinscher is 0.09 second (four boxes) wide. A small dog such as the Poodle should not have a QRS complex wider than 0.05 second (wider), and the larger dog's QRS complex should not exceed 0.06 second (three boxes). Because each dog's QRS complex is too wide, left ventricular enlargement is diagnosed in both cases. The Doberman Pinscher has no P waves because he is in atrial fibrillation. (Paper speed is 50 mm/second; 1 cm equals 1 mV.)</p></caption><graphic xlink:href="gr29"></graphic><attrib>(From Edwards NF: <italic>Bolton's handbook of canine and feline electrocardiography,</italic> ed 2, Philadelphia, 1987, WB Saunders.)</attrib></fig><fig id="f30"><label>Figure 4-30</label><caption><p><bold>A,</bold> In this tracing the R wave averages 3.8 mV (38 boxes). The R wave should not be taller than 3.0 mV (30 boxes) in any dog. A tall R wave indicates left ventricular enlargement. The measurement is made from the baseline (not from the bottom of the Q wave) to the top of the R wave. Two other criteria that indicate left ventricular enlargement are present. The QRS complex is 0.07 second (three boxes) wide and ST segment slurring is present, because the ST segment moves into the T wave without straightening out along the baseline. (Paper speed is 50 mm/second; 1 cm equals 1 mV.) <bold>B,</bold> Left ventricular enlargement in a cat. This lead II electrocardiogram was recorded from an aged cat suffering from hyperthyroidism. Thyroxine levels were 9.9 mg/dL. Note the tall R waves (>0.9 mV). (Paper speed is 50 mm/seconds; 1 cm equals 1 mV.) ST segment slurring is characterized by the slurring of the downstroke of the R wave into the T wave, with no discernible ST segment. This occurs because of ischemia resulting from wall strain in cardiac enlargement.</p></caption><graphic xlink:href="gr30"></graphic><attrib>(Courtesy of NS Moise, New York State College of Veterinary Medicine, Cornell University, Ithaca, NY. From Edwards NF: <italic>Bolton's handbook of canine and feline electrocardiography,</italic> ed 2, Philadelphia, 1987, WB Saunders.)</attrib></fig></p></sec><sec id="cesec174"><title>ST segment</title><p id="para550">The ST segment is between the end of the S wave and the beginning of the T wave. Normally, the ST segment lies on the baseline and then dips into the T wave. Slurring of S into T indicates left ventricular enlargement and is seen when the S wave slurs into the T wave and no ST segment is discernible. The ST segment is elevated if it lies more than 0.1 mV (one box) above the baseline (>0.2 mV in CV<sub>6</sub>LL and CV<sub>6</sub>LU). Elevation of the ST segment may occur with hypercalcemia or myocardial hypoxia. The ST segment is depressed if it lies more than 0.1 mV (one box; >0.2 mV in CV<sub>6</sub>LL and CV<sub>6</sub>LU) below the baseline. Depression of ST may be seen with myocardial ischemia, hypoxia, or hypocalcemia.</p></sec><sec id="cesec175"><title>QT interval</title><p id="para551">The QT interval is measured from the beginning of the Q wave to the end of the T wave. The normal interval is 0.14 to 0.22 second (7 to 11 boxes wide) in dogs and up to 0.16 second in cats. A lengthened QT interval may be seen with hypokalemia or hypocalcemia. The QT interval varies with heart rate and tends to be prolonged when bradycardia occurs. A decreased QT interval may be seen with hypercalcemia.</p></sec></sec><sec id="cesec176"><title>Mean Electrical Axis</title><p id="para552">The <italic>mean electrical cardiac axis</italic> measures the direction (vector) of the cardiac ventricular impulse during depolarization. Therefore the QRS complex is examined in leads I, II, III, aV<sub>R</sub>, aV<sub>L</sub>, and aV<sub>F</sub>. These six leads determine the axis. They are arranged in a manner known as <italic>Bailey's hexaxial lead system</italic> (<xref rid="f31" ref-type="fig">Figure 4-31</xref>). The procedure is as follows:<list list-type="simple" id="celist18"><list-item id="celistitem98"><label>1.</label><p id="para553">Find an isoelectric lead; that is, a lead for which the total number of positve (upward) and negative (downward) deflections of the QRS complex is equal to zero (<xref rid="f32" ref-type="fig">Figure 4-32</xref>). When there is no perfectly isoelectric lead, use the one that comes closest.<fig id="f32"><label>Figure 4-32</label><caption><p>In each of these three leads, the total of the positive and negative deflections equals zero. Each is considered an isoelectric lead.</p></caption><graphic xlink:href="gr32"></graphic></fig></p></list-item><list-item id="celistitem99"><label>2.</label><p id="para554">Find the lead that is perpendicular to the isoelectric lead: lead I is perpendicular to aV<sub>F</sub>; lead II is perpendicular to aV<sub>L</sub>; and lead III is perpendicular to aV<sub>R</sub>.</p></list-item><list-item id="celistitem100"><label>3.</label><p id="para555">Determine whether the perpendicular lead is positive or negative on the patient's electrocardiogram. If the perpendicular lead is negative, the axis is at the negativeend of that lead (each lead has a plus and a minus pole marked. If the perpendicular lead is positive, the mean electrical axis is at the positive end of the perpendicular lead. For example, if aVL is isoelectric (normally it is), lead II is its perpendicular. If lead II is positive on the electrocardiogram, the axis is +60 degress. If lead II is negative on the electrocardiogram, the axis is −120 degrees.</p></list-item></list><fig id="f31"><label>Figure 4-31</label><caption><p>Bailey's hexaxial reference system. The lead axes are marked in 30-degree increments from 0 to 180 degrees and from 0 to −180 degrees. The six leads are marked with a plus sign at the positive electrode and a minus sign at the negative electrode. Note that in the leads I, II, III, and aV<sub>F</sub> the polarity and the angle of the leads are positive or negative simultaneously. Leads aV<sub>F</sub> and aV<sub>L</sub> are positive at the positions of −150 degrees and −30 degrees, respectively, because the positive electrodes for those leads lie in the negative 0 to −180 degrees zone.</p></caption><graphic xlink:href="gr31"></graphic><attrib>(From Ettinger SJ, Suter PF: <italic>Canine cardiology,</italic> Philadelphia, 1970, WB Saunders.)</attrib></fig></p><sec id="cesec177"><title>Normal mean electrical axis</title><p id="para556">The normal dog mean electrical axis is +40 to +100 degrees; for the cat it is more variable: ±0 to ±180 degrees. Right axis deviation (axis more than +100) indicates right ventricular enlargement in the dog (<xref rid="f33" ref-type="fig">Figure 4-33</xref>). Left axis deviation (axis 0 to +40 degrees) indicates left ventricular enlargement in the dog. When there is biventricular enlargement, the axis usually remains normal. Axis determinations are of less value in the cat because the normal range is so wide (<xref rid="cetextbox12" ref-type="boxed-text">BOX 4-12</xref>, <xref rid="cetextbox13" ref-type="boxed-text">BOX 4-13</xref>, <xref rid="cetextbox14" ref-type="boxed-text">BOX 4-14</xref>).<fig id="f33"><label>Figure 4-33</label><caption><p>The mean electrical axis in the frontal plane of this electrocardiogram recorded from a wire-haired fox terrier with pulmonic stenosis is approximately +165 degrees.</p></caption><graphic xlink:href="gr33"></graphic><attrib>(From Edwards NJ: <italic>Bolton's handbook of canine and feline electrocardiography,</italic> ed 2, Philadelphia, 1987, WB Saunders.)</attrib></fig><boxed-text id="cetextbox12"><label>BOX 4-12</label><caption><title>ELECTROCARDIOGRAM CRITERIA FOR LEFT VENTRICULAR ENLARGEMENT</title></caption><p id="para85"><list list-type="simple" id="celist8"><list-item id="celistitem37"><label>1.</label><p id="para86">Left axis deviation (dog)</p></list-item><list-item id="celistitem38"><label>2.</label><p id="para87">QRS complex too wide (but has normal configuration)</p></list-item><list-item id="celistitem39"><label>3.</label><p id="para88">R wave too tall</p></list-item><list-item id="celistitem40"><label>4.</label><p id="para89">S-T segment slurring</p></list-item><list-item id="celistitem41"><label>5.</label><p id="para90">May be associated with P mitrale</p></list-item></list></p></boxed-text><boxed-text id="cetextbox13"><label>BOX 4-13</label><caption><title>ELECTROCARDIOGRAM CRITERIA FOR RIGHT VENTRICULAR ENLARGEMENT</title></caption><p id="para91"><list list-type="simple" id="celist9"><list-item id="celistitem42"><label>1.</label><p id="para92">Right axis deviation (dog and cat)</p></list-item><list-item id="celistitem43"><label>2.</label><p id="para93">Presence of an S wave in leads I, II, and III (dog only)</p></list-item><list-item id="celistitem44"><label>3.</label><p id="para94">S wave deeper than 0.7 mV in lead CV<sub>6</sub>LU (V<sub>4</sub>) in the dog</p></list-item><list-item id="celistitem45"><label>4.</label><p id="para95">May be associated with P pulmonale</p></list-item></list></p></boxed-text><boxed-text id="cetextbox14"><label>BOX 4-14</label><caption><title>ELECTROCARDIOGRAM CRITERIA FOR BIVENTRICULAR ENLARGEMENT</title></caption><p id="para96"><list list-type="simple" id="celist10"><list-item id="celistitem46"><label>1.</label><p id="para97">Tall R wave</p></list-item><list-item id="celistitem47"><label>2.</label><p id="para98">Wide QRS complex</p></list-item><list-item id="celistitem48"><label>3.</label><p id="para99">ST segment slurring</p></list-item><list-item id="celistitem49"><label>4.</label><p id="para100">Deep Q waves in lead II (deeper than 0.3 mV for the cat, 0.5 mV for the dog)</p></list-item><list-item id="celistitem50"><label>5.</label><p id="para101">Normal mean electrical axis</p></list-item><list-item id="celistitem51"><label>6.</label><p id="para102">P mitrale or P pulmonale, or both</p></list-item></list></p></boxed-text></p></sec></sec></sec><sec id="cesec178"><title>ENDOSCOPY: INDICATIONS AND EQUIPMENT REQUIREMENTS</title><p id="para557"><boxed-text id="cetextbox25"><caption><title>Note:</title></caption><p id="para558">The discussion that follows centers around indications and capabilities of endoscopy in clinical practice. The discussion is not intended to be used as a “How-To” instruction guide on performing endoscopic procedures in dogs and cats. Today, numerous types of endoscopes and accessory materials are available for use in clinical practice. Specific hands-on training and complete familiarity with the equipment package available <italic>is essential</italic> before attempting to perform any of the procedures outlined.</p><p id="para559">Inappropriate use of endoscopic equipment not only can damage expensive equipment but also can cause serious injury to the patient.</p></boxed-text></p><sec id="cesec179"><title>Upper Respiratory Tract: Laryngoscopy and Pharyngoscopy</title><p id="para560">Endoscopy of the upper respiratory tract is among the most important advanced diagnostic and therapeutic tools used in the evaluation of patients that have stertor (snorting), reverse sneeze, stridor (wheezing), and chronic cough. Laryngoscopy is of value in the diagnosis of upper airway obstructions such as eversion of the lateral ventricles, collapsed arytenoid cartilages, hyperplasia of the vocal cords, nodules on the vocal cords, elongated soft palate, collapsed proximal trachea, and traumatic injuries to the neck. Note also, however, that a careful visual examination of the larynx in the anesthetized patient (only) can be highly valuable even without the use of endoscopic equipment; for example, for assessment of laryngeal movement in patients with laryngeal paralysis. Suspected lesions <italic>inside</italic> the larynx may be difficult to visualize with or without endoscopic equipment. Examination of the trachea and main stem bronchi requires endoscopic evaluation to assess the integrity of the airway for conditions such as collapsed trachea, mediastinal tumors, hilar lymph node enlargement, parasitic nodules <italic>(Filaroides osleri)</italic>, and foreign body aspiration. In addition, tracheobronchoscopy is a valuable technique that permits culturing and cytologic examination of material from bronchi involved in chronic respiratory disease. Upper airway obstruction that is not responsive to conservative therapy is an indication for more extensive diagnostic procedures, such as bronchoscopy.</p><p id="para561">Endoscopes of varying sizes are appropriate for use in examining the larynx and trachea. However, in cats and small dogs, examination of the trachea using equipment as small as a (human) bronchoscope may limit the examination because the endoscope nearly occludes the tracheal diameter. Additional training and/or experience is recommended when performing tracheoscopy in small patients.</p><p id="para562">One of the most important endoscopic techniques performed in dogs and cats involves examination of the nasopaharynx, the upper respiratory compartment above the soft palate. Sometimes called pharyngoscopy, examination entails retroflexion of a small-diameter endoscope (e.g., bronchoscope) 170 to 180 degrees to allow visualization of the space between the posterior nares (choana) and the larynx (<xref rid="f34" ref-type="fig">Figure 4-34</xref>). This is a common location for foreign body entrapment and occasional tumor development in cats and dogs (<xref rid="f35" ref-type="fig">Figure 4-35</xref>). Pharyngoscopy is the <italic>only</italic> effective means of examining this portion of the upper respiratory tract in patients that have a history of stertor (snorting) and so-called reverse sneeze.<fig id="f34"><label>Figure 4-34</label><caption><p><bold>A,</bold> The appearance of the normal choana (posterior nares) in a cat. <bold>B,</bold> The appearance of the choana of cat with a posterior nasal mass diagnosed as lymphoma.</p></caption><graphic xlink:href="gr34"></graphic></fig><fig id="f35"><label>Figure 4-35</label><caption><p>Lateral skull radiograph of a dog depicting the proper endoscopic placement for pharyngoscopy.</p></caption><graphic xlink:href="gr35"></graphic></fig></p></sec><sec id="cesec180"><title>Lower Respiratory Tract: Bronchoscopy</title><p id="para563">Endoscopic examination of the bronchi and lower airways is a highly diagnostic, occasionally therapeutic procedure indicated in patients that present for persistent cough. As in all endoscopic procedures, the patient is anesthetized for the examination. However, examination of the lower respiratory tract requires considerable attention to patient oxygenation and respiratory status during the examination. The requirement for oxygen to be administered throughout the procedure may be a significant limiting factor unless special accessories are used. In the ideal situation, the patient is a medium- to large-sized dog and the endoscope can be passed <italic>through</italic> the endoscope using a T adaptor while oxygen and anesthetic are administered simultaneously.</p><p id="para564">However, in cats and small dogs, it is usually not possible to pass an endoscope through the endotracheal tube. The procedure must be done by passing the endoscope directly into the trachea to the level of the right and left main bronchi and probably not much farther. Supplemental intravenous anesthetic is likely to be required because of the time required to complete the examination. Training and/or experience is essential before performing bronchoscopy, particularly in cats and small dogs.</p><p id="para565">The greatest advantage in performing bronchoscopy is to visualize the integrity of the trachea and, to a limited extent, the lower airways. Airway collapse, not visible on conventional radiography, can be strikingly apparent. Foreign body entrapment, tumors, respiratory parasites and airway trauma also can be identified with bronchoscopy. Additionally, the bronchoscopic examination allows for collection of cytologic samples from discrete areas (airways) within the lower respiratory tract. The ability to perform bronchoalveolar lavage in patients with reactive airway disease, subclinical or clinical infections, and certain types of tumors can be highly diagnostic.</p></sec><sec id="cesec181"><title>Gastrointestinal Endoscopy</title><p id="para566">Flexible fiberoptic endoscopy is a noninvasive, atraumatic means of visualizing the mucosal surfaces of the esophagus, stomach, and colon. Flexible endoscopes are available from several companies at a wide range of prices. To minimize the risk of injury to the animal and to reduce the possibility of damage to the endoscope, place animals undergoing endoscopic examination under general anesthesia after routine preanesthetic preparation. A fast of 12 to 24 hours is recommended for most patients undergoing upper gastrointestinal endoscopy. However, for patients with indications of delayed gastric emptying, a longer fast (24 to 48 hours) may be needed to empty the stomach completely. In preparation for colonoscopy, a 24- to 48-hour fast is recommended. Give a high warm-water enema the evening before and again 2 to 4 hours before the procedure. Give such enemas until the return is clear.</p><sec id="cesec182"><title>Esophagoscopy</title><p id="para567">The clinical signs indicating esophageal disease and a potential benefit of esophagoscopy include repeated regurgitation, excessive drooling, ballooning of the esophagus, anorexia or dysphagia, and recurrent pneumonia. Esophagoscopy allows visualization of the mucosal lining of the esophagus and makes it possible to detect inflammation, ulcerations, dilatations, diverticula, strictures, foreign bodies, tumors, and parasite infestations.</p></sec><sec id="cesec183"><title>Gastroscopy and duodenoscopy</title><p id="para568">Endoscopic examination of the mucosal aspect of the stomach is indicated when the clinical signs or physical findings suggest the presence of gastric disease or when there is a need for confirmation or clarification of radiographic findings. In most cases, persistent vomiting is the chief complaint. Other clinical signs suggestive of serious gastric disease include hematemesis, melena, weight loss, anemia, and abdominal pain. Gastroscopy allows visualization of the mucosal lining of the stomach and enables detection of inflammation, ulceration, foreign bodies, and tumors. In most dogs and cats the endoscope can be passed into the proximal duodenum. Depending on the patient size and length of the scope, it may be possible to evaluate as much as 12 inches or more of the proximal duodenum.</p></sec><sec id="cesec184"><title>Colonoscopy</title><p id="para569">Colonoscopy refers to endoscopic examination of colon, rectum, and anus. The technique is helpful in the definitive diagnosis of lower bowel lesions, such as granulomatous colitis, foreign bodies, tumors, lacerations, and other mucosal abnormalities. The primary indication for colonoscopy is the presence of signs of large bowel disease, which typically include tenesmus and the passage of small, frequent stools containing fresh blood or excess mucus. Endoscopic examination of the colon allows direct visualization of the effects of mucosal inflammation, ulceration, mucosal polyps, malignant neoplasia, and strictures. Histologic examination of mucosal biopsy material will confirm the diagnosis of colonic disease.</p><p id="para570">The large bowel must be empty for the colonic mucosa to be visualized. The bowel can be emptied by withholding food for 24 hours and performing a colonic irrigation the evening before and again 2 hours before the examination. The material used for the enema must be nonirritating and nonoily. Mildly hypertonic saline solutions such as Fleet enemas work well if given 2 hours before examination so that gas and fluid can be passed completely. However, do not use Fleet enemas in cats or small dogs.</p><p id="para571">If the general physical condition of the animal is poor and withholding food is not possible, feeding a low-residue diet for 12 to 18 hours preceding colonoscopy can be helpful. This diet could consist of cooked eggs, small amounts of cooked beef or chicken, and small amounts of carbohydrates, such as a slice of toast or ¼ to ½ cup of moist kibble. Maintain good hydration. If all food is contraindicated, oral electrolyte solutions such as Gatorade (PepsiCo, Purchase, New York) can be used to maintain hydration without moving solids through the intestinal tract.</p><p id="para572">Give the animal a short-acting anesthetic and place the animal on a tilted table in lateral recumbency with the hindquarters elevated. Perform a digital examination of the rectum and pelvic cavity to ensure that there are no strictures, polyps, or other obstructions. Lubricate the proctoscope thoroughly with water-soluble jelly and pass it gently through the anal sphincter. Press the proctoscope forward slowly and carefully with a spiral motion. If any resistance is encountered, stop the motion, remove the obturator, and inspect the bowel to determine the cause of the resistance. If possible, replace the obturator and continue forward motion until the instrument is passed its full length. Withdraw the obturator, and observe the mucosa.</p><p id="para573">The major portion of the examination is conducted as the instrument is withdrawn. To view the colonic and rectal walls completely, one must move the anterior end of the proctoscope around the circumference of a small circle while withdrawing the proctoscope. Occasional insufflation with the inflating bulb is helpful in smoothing out folds of tissue. Repeated instrumentation may produce petechiae and minor hemorrhages that are not pathologic. For examination of the terminal rectum and anus, the Hirshman anoscope provides adequate, convenient visualization.</p><p id="para574">Newer techniques for visualizing the upper and lower gastrointestinal tract are being used in dogs. The flexible fiberoptic endoscope enables one to visualize and photograph the esophagus, colon, and stomach. One is able not only to visualize lesions of the gastrointestinal tract directly but also to assess motility, take biopsies of lesions, and remove foreign bodies.</p></sec><sec id="cesec185"><title>Vaginoscopy</title><p id="para575">The ability to visualize directly the vestibule, the vagina to the level of the cervix, and the urethral orifice in female dogs is of particular value in evaluating patients with known or suspected congenital urinary tract disorders, such as incontinence or ectopic ureters and vaginal strictures (congenital or traumatic). Numerous vaginal malformations and chronic infections have visual changes that are identified easily during endoscopic examination. Frequently, the procedure can be conducted in the standing awake patient. Sedation or general anesthesia is indicated when extensive manipulation, catheterization of the bladder, or a vaginal biopsy are indicated. Position the sedated or anesthetized patient in dorsal or ventral recumbency to facilitate orientation during the procedure. If catheterization of the urinary bladder is required during the procedure, dorsal recumbency seems to facilitate visualization of the urethral papilla and insertion of the catheter.</p><p id="para576">Vaginoscopy entails use of a relatively small, flexible endoscope 4 to 6 mm in diameter or a 2- to 3-mm rigid scope. The flexible scope offer the advantage of a larger biopsy channel and the ability to view the lateral vaginal wall easily. Vaginoscopy is considered an invasive procedure and should be conducted under sterile conditions. Before insertion of the sterilized endoscope, the vulva should be free of obvious debris, should be clipped if necessary, and should be cleaned gently with a surgical soap and rinsed. Insert the scope such that initial position of the tip of the scope is directed toward the anus. As insertion proceeds, the tip of the endoscope reaches the horizontal portion of the vestibule and vagina. When feasible, pass the scope to the level of the cervix. Slight insufflation of the vagina may be useful in dilating the vagina, greatly facilitating the examination. Conducting the examination from the level of the cervix caudally is recommended. This maximizes the ability to visualize critical anatomic features.</p></sec><sec id="cesec186"><title>Cystoscopy</title><p id="para577">The relatively recent introduction of very small (2-mm diameter) flexible and rigid endoscopes into veterinary medicine allows visual examination of the urethra, trigone, urinary bladder, and the right and left ureterovesicular junctions of female dogs and even cats. Such examinations are most useful when obstructive lesions (tumor or calculi) of the urethra or trigone are suspected. Visual examination of the interior surface of the bladder and the capability of collecting biopsy samples makes this a particularly useful diagnostic tool in the hands of the experienced clinician.</p><sec id="cesec187"><sec id="cesec188"><title>Additional Reading</title><p id="para578">Guilford WG, Center SA, Strombeck DR, et al: <italic>Strombeck's small animal gastroenterology,</italic> ed 3, Philadelphia, 1996, WB Saunders.</p><p id="para579">Holt DE: Laryngoscopy and pharnygoscopy. In King LG, editor: <italic>Textbook of respiratory disease in dogs and cats,</italic> St Louis, 2004, Elsevier-Saunders.</p><p id="para580">Jones B: Incorporating endoscopy in veterinary practice, <italic>Compend Contin Educ Pract Vet</italic> 20:307-313, 1998.</p><p id="para581">Kuehn NF, Hess RS: Bronchoscopy. In King LG, editor: <italic>Textbook of respiratory disease in dogs and cats,</italic> St Louis, 2004, Elsevier-Saunders.</p><p id="para582">Tams TR: <italic>Handbook of small animal gastroenterology,</italic> Philadelphia, 1996, WB Saunders.</p></sec></sec></sec></sec></sec><sec id="cesec189"><title>FLUID THERAPY (SEE ALSO SECTION 1)</title><sec id="cesec190"><title>Maintenance Fluids (Management of Normal Fluid Losses)</title><p id="para583">The volume of fluids required to replace the normal ongoing losses can be determined from predictive charts (<xref rid="cetable8" ref-type="table">TABLE 4-8</xref>, <xref rid="cetable9" ref-type="table">TABLE 4-9</xref>). If a chart is not available, the maintenance volume is assumed to be 40 to 60 mL/kg/day (higher values per kilogram for smaller dogs and cats and lower values per kilogram for larger dogs and cats).<table-wrap position="float" id="cetable8"><label>TABLE 4-8</label><caption><p>Approximate Daily Energy and Water Requirements of Dogs Based on Body Weight<xref rid="cetablefn7" ref-type="table-fn">*</xref></p></caption><table frame="hsides" rules="groups"><thead><tr><th></th><th colspan="3" align="center">Total energy (kcal) or water (mL)<hr></hr></th></tr><tr><th align="center">Body weight (kg)</th><th align="center">Per day</th><th align="center">Per kilogram</th><th align="center">Per hour</th></tr></thead><tbody><tr><td align="center">1</td><td align="center">132</td><td align="center">132</td><td align="char">5.5</td></tr><tr><td align="center">2</td><td align="center">222</td><td align="center">111</td><td align="char">9.5</td></tr><tr><td align="center">3</td><td align="center">301</td><td align="center">100</td><td align="char">12.5</td></tr><tr><td align="center">4</td><td align="center">373</td><td align="center">93</td><td align="char">15.5</td></tr><tr><td align="center">5</td><td align="center">441</td><td align="center">88</td><td align="char">18.5</td></tr><tr><td align="center">6</td><td align="center">506</td><td align="center">84</td><td align="center">21</td></tr><tr><td align="center">7</td><td align="center">568</td><td align="center">81</td><td align="char">23.5</td></tr><tr><td align="center">8</td><td align="center">628</td><td align="center">78</td><td align="center">26</td></tr><tr><td align="center">9</td><td align="center">686</td><td align="center">76</td><td align="char">28.5</td></tr><tr><td align="center">10</td><td align="center">742</td><td align="center">74</td><td align="center">31</td></tr><tr><td align="center">11</td><td align="center">797</td><td align="center">72</td><td align="center">33</td></tr><tr><td align="center">12</td><td align="center">851</td><td align="center">71</td><td align="char">35.5</td></tr><tr><td align="center">13</td><td align="center">904</td><td align="center">70</td><td align="char">37.5</td></tr><tr><td align="center">14</td><td align="center">955</td><td align="center">68</td><td align="center">40</td></tr><tr><td align="center">15</td><td align="center">1006</td><td align="center">67</td><td align="center">42</td></tr><tr><td align="center">16</td><td align="center">1056</td><td align="center">66</td><td align="center">44</td></tr><tr><td align="center">17</td><td align="center">1105</td><td align="center">65</td><td align="center">46</td></tr><tr><td align="center">18</td><td align="center">1154</td><td align="center">64</td><td align="center">48</td></tr><tr><td align="center">19</td><td align="center">1201</td><td align="center">63</td><td align="center">50</td></tr><tr><td align="center">20</td><td align="center">1248</td><td align="center">62</td><td align="center">52</td></tr><tr><td align="center">21</td><td align="center">1295</td><td align="center">62</td><td align="center">54</td></tr><tr><td align="center">22</td><td align="center">1341</td><td align="center">61</td><td align="center">56</td></tr><tr><td align="center">23</td><td align="center">1386</td><td align="center">60</td><td align="center">58</td></tr><tr><td align="center">24</td><td align="center">1431</td><td align="center">60</td><td align="char">59.5</td></tr><tr><td align="center">25</td><td align="center">1476</td><td align="center">59</td><td align="char">61.5</td></tr><tr><td align="center">26</td><td align="center">1520</td><td align="center">58</td><td align="char">63.5</td></tr><tr><td align="center">27</td><td align="center">1564</td><td align="center">58</td><td align="center">65</td></tr><tr><td align="center">28</td><td align="center">1607</td><td align="center">57</td><td align="center">67</td></tr><tr><td align="center">29</td><td align="center">1650</td><td align="center">57</td><td align="char">68.5</td></tr><tr><td align="center">30</td><td align="center">1692</td><td align="center">56</td><td align="char">70.5</td></tr><tr><td align="center">35</td><td align="center">1899</td><td align="center">54</td><td align="center">79</td></tr><tr><td align="center">40</td><td align="center">2100</td><td align="center">52</td><td align="char">87.5</td></tr><tr><td align="center">45</td><td align="center">2293</td><td align="center">51</td><td align="char">95.5</td></tr><tr><td align="center">50</td><td align="center">2482</td><td align="center">50</td><td align="char">103.5</td></tr><tr><td align="center">55</td><td align="center">2666</td><td align="center">48</td><td align="center">111</td></tr><tr><td align="center">60</td><td align="center">2846</td><td align="center">47</td><td align="char">118.5</td></tr><tr><td align="center">70</td><td align="center">3195</td><td align="center">46</td><td align="center">133</td></tr><tr><td align="center">80</td><td align="center">3531</td><td align="center">44</td><td align="center">147</td></tr><tr><td align="center">90</td><td align="center">3857</td><td align="center">43</td><td align="center">161</td></tr><tr><td align="center">100</td><td align="center">4174</td><td align="center">42</td><td align="center">174</td></tr></tbody></table><table-wrap-foot><fn id="cetablefn7"><label>*</label><p id="cenotep7">132 kcal/kg<sup>0,75</sup>.</p></fn></table-wrap-foot><attrib>From Nutritional Requirements of the Dog, National Research Council, Bethesda, MD, 1985.</attrib><permissions><copyright-statement>© 2006 National Research Council</copyright-statement><copyright-year>2006</copyright-year><license><license-p>Since January 2020 Elsevier has created a COVID-19 resource centre with free information in English and Mandarin on the novel coronavirus COVID-19. The COVID-19 resource centre is hosted on Elsevier Connect, the company's public news and information website. Elsevier hereby grants permission to make all its COVID-19-related research that is available on the COVID-19 resource centre - including this research content - immediately available in PubMed Central and other publicly funded repositories, such as the WHO COVID database with rights for unrestricted research re-use and analyses in any form or by any means with acknowledgement of the original source. These permissions are granted for free by Elsevier for as long as the COVID-19 resource centre remains active.</license-p></license></permissions></table-wrap><table-wrap position="float" id="cetable9"><label>TABLE 4-9</label><caption><p>Approximate Daily Energy and Water Requirements of Cats Based on Body Weight<xref rid="cetablefn8" ref-type="table-fn">*</xref></p></caption><table frame="hsides" rules="groups"><thead><tr><th></th><th colspan="3" align="center">Total energy (kcal) or water (mL)<hr></hr></th></tr><tr><th align="center">Body weight (kg)</th><th align="center">Per day</th><th align="center">Per kilogram</th><th align="center">Per hour</th></tr></thead><tbody><tr><td align="char">1.0</td><td align="char">80.0</td><td align="center">80</td><td align="center">3</td></tr><tr><td align="char">1.5</td><td align="char">108.4</td><td align="center">72</td><td align="center">5</td></tr><tr><td align="char">2.0</td><td align="char">134.5</td><td align="center">67</td><td align="center">6</td></tr><tr><td align="char">2.5</td><td align="char">159.1</td><td align="center">64</td><td align="center">7</td></tr><tr><td align="char">3.0</td><td align="char">182.4</td><td align="center">61</td><td align="center">8</td></tr><tr><td align="char">3.5</td><td align="char">204.7</td><td align="center">58</td><td align="center">9</td></tr><tr><td align="char">4.0</td><td align="char">226.3</td><td align="center">57</td><td align="center">9</td></tr><tr><td align="char">4.5</td><td align="char">247.2</td><td align="center">55</td><td align="center">10</td></tr><tr><td align="char">5.0</td><td align="char">267.5</td><td align="center">53</td><td align="center">11</td></tr></tbody></table><table-wrap-foot><fn id="cetablefn8"><label>*</label><p id="cenotep8">80 kcal/kg<sup>0,75</sup>.</p></fn></table-wrap-foot><attrib>From Nutritional Requirements of the Cat, National Research Council, Bethesda, MD, 1987.</attrib><permissions><copyright-statement>© 2006 National Research Council</copyright-statement><copyright-year>2006</copyright-year><license><license-p>Since January 2020 Elsevier has created a COVID-19 resource centre with free information in English and Mandarin on the novel coronavirus COVID-19. The COVID-19 resource centre is hosted on Elsevier Connect, the company's public news and information website. Elsevier hereby grants permission to make all its COVID-19-related research that is available on the COVID-19 resource centre - including this research content - immediately available in PubMed Central and other publicly funded repositories, such as the WHO COVID database with rights for unrestricted research re-use and analyses in any form or by any means with acknowledgement of the original source. These permissions are granted for free by Elsevier for as long as the COVID-19 resource centre remains active.</license-p></license></permissions></table-wrap></p><p id="para584">The nature of the fluids used for maintenance is distinctly different from that of fluids used to replace extracellular volume deficits. The average concentration of normal urine and insensible losses is 40 to 50 mEq/L, and the potassium concentration is 15 to 20 mEq/L. Administration of a replacement solution to an animal for its maintenance requirements predisposes to hypernatremia (most animals are able to eliminate the excess sodium) and hypokalemia.</p><p id="para585">Lactated Ringer's solution or equivalent replacement solutions are poor maintenance solutions because they predispose to hypernatremia and hypokalemia. In one version of a homemade maintenance solution, a replacement solution is supplemented with potassium (15 to 20 mEq/L) to accommodate the potassium losses that normally occur. A further modification is dilution of the replacement solution with one to two parts of 5% dextrose in water per one part of replacement solution. The easiest approach is to use commercial maintenance solutions.</p><p id="para586">Maintenance solutions also should not be used to replace extracellular volume deficits because they may cause hyponatremia and hyperkalemia when administered in large volumes. If a patient is receiving a maintenance solution and is noted to be dehydrated or hypotensive, administer a replacement solution.</p></sec></sec><sec id="cesec191"><title>MAINTENANCE FLUIDS (MANAGEMENT OF ABNORMAL FLUID LOSSES)</title><p id="para587">Ongoing losses that occur via transudation into one of the major body cavities, into the tissues, or through burn wounds are similar in composition to extracellular fluid and should be replaced with lactated Ringer's solution or an equivalent replacement solution. Ongoing losses that occur via vomiting, diarrhea, or diuresis should be replaced with lactated Ringer's solution or an equivalent solution that has been supplemented with potassium (10 to 30 mEq/L). One exception is the patient that has been vomiting stomach contents chronically, a situation in which 0.9% sodium chloride supplemented with potassium (10 to 30 mEq/L) is recommended.</p><p id="para588">When the fluid therapy plan is being developed, how much fluid the animal will lose over the day is not known. One can leave this category blank initially and then, as losses occur during the day, add equivalent volumes of the appropriate fluid to the fluid therapy regimen. Alternatively, if the patient has a disease that is known to be associated with severe ongoing fluid losses (e.g., parvovirus gastroenteritis), one can fill in an estimated volume initially and then adjust upward or downward as the day progresses.</p></sec><sec id="cesec192"><title>FLUID THERAPY IN THE NONCRITICAL PATIENT</title><p id="para589">The fluid therapy prescription is the best guess as to the requirements of the patient. Implementation of a fluid therapy regimen that is as close as possible to the prescription is important; however, considering the inherent inaccuracies in the assumptions used to construct the prescription, it is not imperative to administer exactly what has been prescribed. There are many acceptable ways to administer the prescribed fluids. One way is to mix all of the fluids and additives from each category into one large bottle and administer them throughout the day. Another way is to administer the fluids simultaneously, in
parallel, throughout the day in one administration line. A third way is to administer the fluids in series. Administer the prescribed fluids in a manner that is convenient for your practice situation.</p><p id="para590">The following are important points to remember in fluid therapy:<list list-type="simple" id="celist19"><list-item id="celistitem101"><label>1.</label><p id="para591">To determine the rate of intravenous infusion of the fluids and additives, take the total volume of the fluids that have been prescribed and divide the total volume by the total number of hours in the day that are available for safe administration of the fluids. There should be no need to front-load the deficit repair fluid volume as long as the intravascular volume has been stabilized previously.</p></list-item><list-item id="celistitem102"><label>2.</label><p id="para592">Administer the fluids over as many hours as possible to allow the patient as much time as possible to redistribute and fully use the administered fluids and electrolytes. With faster administration, a diuresis will occur and more of the fluids will be excreted in urine. Do not administer fluids continuously intravenously when the patient cannot be observed periodically to ascertain whether the fluids are continuing to run at an appropriate rate and that the administration line has not become disconnected. If the available time is limited (i.e., less than 12 hours) or if extra time is needed for safe administration of the fluids, an alternative plan (e.g., administering some of the required fluids subcutaneously) is indicated.</p></list-item><list-item id="celistitem103"><label>3.</label><p id="para593">Intravenous administration of fluids is the preferred route because the fluid is dispersed rapidly and is immediately available to the patient. Intravenous administration may be inconvenient in some practice settings. The prescribed fluid can be administered subcutaneously in several divided doses. The subcutaneous route is usually well tolerated by patients, and therapy often is efficacious. However, the subcutaneous route is slower in onset than the intravenous route, is less efficacious than the intravenous route because some patients (especially those that are severely dehydrated and vasoconstricted) do not absorb the fluids well or at all, and it carries a slight risk of infection. Fluids can be administered orally or via stomach tube, in several divided doses, as long as the gastrointestinal tract is functional. Fluids also can be administered intraperitoneally. The intraperitoneal route is characterized by the same advantages and disadvantages as the subcutaneous route, but in addition there is a danger of injury or perforation of an abdominal organ. Fluids also can be administered via the intramedullary route. Intramedullary administration is more difficult than subcutaneous or intraperitoneal administration but is associated with much more rapid and reliable systemic uptake. The intramedullary route may be useful when venous access is difficult (e.g., a severely dehydrated kitten or puppy).</p></list-item><list-item id="celistitem104"><label>4.</label><p id="para594">Fluids may be administered through central or peripheral veins. Indwelling catheters must be introduced and maintained aseptically. All fluids and administration sets must be sterile. Fluids with osmolalities less than 600 mOsm/L can be administered safely via a peripheral vein. Administer fluids with osmolalities greater than 700 mOsm/L via a large central vein because hyperosmolar fluids may cause thrombophlebitis if administered via small peripheral veins.</p></list-item><list-item id="celistitem105"><label>5.</label><p id="para595">Control of the infusion rate is difficult when fluids are administered by gravity. Eliminate this problem by using an infusion pump (<xref rid="cetextbox15" ref-type="boxed-text">Box 4-15</xref>).<boxed-text id="cetextbox15"><label>BOX 4-15</label><caption><title>RATES OF FLOW AMONG INFUSION SETS</title></caption><p id="para103">Cutter: 20 drops/mL</p><p id="para104">Abbott and McGaw: 15 drops/mL</p><p id="para105">McGaw: 15 drops/mL</p><p id="para106">Travenol: 10 drops/mL</p><p id="para107">To calculate drops per minute, use the following formulas:</p><p id="para108">Fluid volume to be infused (mL)/No. of hours available (hour) = mL/hour</p><p id="para109">For adult drip sets:</p><p id="para110">Cutter: mL/hour ÷ 3 = drops/minute</p><p id="para111">Abbott and McGaw: mL/hour ÷ 4 = drops/minute</p><p id="para112">Travenol: mL/hour ÷6 = drops/minute</p></boxed-text></p></list-item></list></p><p id="para596">Most pediatric drip sets deliver 60 drops/mL (milliliters per hour equals drops per minute). Write fluid orders so that the volume to be administered is recorded as <italic>mL/day, mL/hour,</italic> and <italic>drops/minute.</italic> This will allow personnel to detect major calculation errors more easily. The clinician should not assume that the animal has received the volume of fluid ordered, and nursing personnel should note in the record the volume actually received. Clearly list all additives on the bottle; adhesive labels for this purpose are available. Attach a strip of adhesive tape to the bottle and mark it appropriately to provide a quick visual estimate of the volume of fluid received.</p><sec id="cesec193"><sec id="cesec194"><sec id="cesec195"><sec id="cesec196"><title>Additional Reading</title><p id="para597">DiBartola SP: <italic>Fluid electrolyte, and acid-base disorders in small animal practice,</italic> St Louis, 2006, Elsevier-Saunders.</p><p id="para598">Mathews KA: The various types of parenteral fluids and their indications, <italic>Vet Clin North Am Small Anim Pract</italic> 28:483, 1998.</p><p id="para599">Phillips S, Polzin D: Clinical disorders of potassium homeostasis: hyperkalemia and hypokalemia, <italic>Vet Clin North Am Small Anim Pract</italic> 28:545, 1998.</p><p id="para600">Schaer M (ed): Advances in fluid and electrolyte disorders, <italic>Vet Clin North Am Small Anim Pract</italic> 28:3, May 1998.</p><p id="para601">Wingfield WE: <italic>Veterinary emergency medicine secrets,</italic> Philadelphia, 1997, Hanley & Belfus.</p></sec></sec></sec></sec></sec></sec><sec id="cesec197"><sec id="cesec198"><title>GASTROINTESTINAL PROCEDURES</title><p id="para602">Numerous techniques are described for administering calories and nutrients to patients that are unable or unwilling to take in, chew, or swallow food. One method, intravenous hyperalimentation, is reserved for patients that are not able to tolerate any food being introduced via the gastrointestinal tract and represents a radical, and ideally transient, departure from normal. However, enteral feeding, which is always preferable to intravenous hyperalimentation, allows the clinician several options for administering food directly into the gastrointestinal tract. However, consideration of several variables is critical when one is initiating enteral feeding programs, such as the patient's diagnosis, attitude, status of the gastrointestinal tract, and the ability of the patient to digest and absorb food once introduced. In addition, consideration of the type and constituency of the diet provided is important. Although the options available for enteral nutrition are much greater that those for intravenous hyperalimentation, the clinician must consider dietary requirements carefully when planning enteral nutritional support.</p><p id="para603">When evaluating enteral feeding for the individual patient, the clinician has four basic options: nasoesophageal tube, pharyngostomy tube (least recommended), esophagostomy tube, and percutaneous gastroscopy tube (which can be performed using an endoscope or performed using the so-called blind technique). All techniques involve use of polyurethane or silicone feeding tube. The nasoesophageal tube placement technique does not require general anesthesia and may be inserted using a topical anesthetic only. Each of the other techniques described requires the patient to be anesthetized to ensure proper and safe placement.</p><sec id="cesec199"><title>Nasoesophageal Intubation</title><p id="para604">For temporary, short-term feeding, nasoesophageal intubation is a simple technique that works well in cats, puppies, and adult dogs. Patients that are comatose; have severe, persistent vomiting; or are unable to swallow are not candidates for this procedure. The objective of the procedure is to place a small-diameter tube (8F to 10F for dogs weighing more than 15 kg and 5F to 8F for small dogs and cats) through the nasal cavity into the distal esophagus. The tube does not have to enter the stomach. When measuring the tube length, measure from the tip of the nose to the eighth or ninth rib (<xref rid="f36" ref-type="fig">Figure 4-36</xref>).<fig id="f36"><label>Figure 4-36</label><caption><p>Technique for measuring the insertion length of nasoesophageal tube in a dog.</p></caption><graphic xlink:href="gr36"></graphic></fig></p><p id="para605">Administer 3 to 5 drops of a topical ophthalmic solution (0.5% proparacaine) directly into one nostril. Hold the head gently upward for a few seconds to allow the solution to reach the back of the nasal cavity. In most patients, it is desirable to wait 1 to 2 minutes and then to repeat the instillation in the same nostril. For larger dogs, 2% lidocaine solution (0.5 to 2.0 mL) gradually instilled into the nostril is an alternative technique to achieve topical anesthesia. Lubricate the tube with a thin coat of a water-soluble lubricant, such a 2% lidocaine lubricating gel. Pass the tube into the nasal cavity while directing the tube tip medial and ventral into the ventral meatus. The anatomic shape of a dog's nostril usually requires directing the tip medially but almost perpendicular to the plane of the nasal cavity to facilitate insertion. Initial resistance (pressure, not pain) usually is perceived, and the patient's head as expected quickly retracts, leaving the operator holding the tube tip some inches away from the patient's nose. Be persistent. Repeat the procedure, as necessary, by <italic>quickly</italic> inserting the first inch or more of the tube into the nostril. Once started, the remainder of the technique is relatively straightforward.</p><p id="para606">As the tube reaches the caudal aspect of the nasopharynx, it should pass directly into the esophagus with little or no resistance. Affix the tube remaining outside the patient to the head or face using a “butterfly” tape, gauze, suture (<xref rid="f37" ref-type="fig">Figure 4-37</xref>), or skin glue (skin glue [Superglue] generally is NOT recommended because this can result in loss of hair and skin pigment when the glue becomes dislodged).<fig id="f37"><label>Figure 4-37</label><caption><p>Nasoesophageal tube secured with stay sutures on the face of a puppy.</p></caption><graphic xlink:href="gr37"></graphic></fig></p><p id="para607">CAUTION: The tip of the tube possibly can be introduced inadvertently through the glottis and into the trachea. Topical anesthetic instilled into the nose can anesthetize the arytenoid cartilages, thereby blocking a cough or gag reflex. I prefer to check the tube placement with a dry, empty syringe. Attach the test syringe to the end of the feeding tube. Rather than inject air or water in an attempt to auscultate borborygmus over the abdomen, simply attempt to aspirate air from the feeding tube (<xref rid="f38" ref-type="fig">Figure 4-38</xref>). IF THERE IS NO RESISTANCE DURING ASPIRATION AND AIR FILLS THE SYRINGE, IT IS LIKELY THAT THE TUBE HAS BEEN PLACED IN THE TRACHEA. Completely remove the tube and repeat the procedure. However, if repeated attempts to aspirate are met with immediate resistance and NO AIR ENTERS THE SYRINGE, the tube tip is positioned properly within the esophagus. If there is any question regarding placement, a lateral survey radiograph is indicated.<fig id="f38"><label>Figure 4-38</label><caption><p>Technique for verifying esophageal placement of the tip of a nasoesophageal tube in a cat.</p></caption><graphic xlink:href="gr38"></graphic></fig></p></sec><sec id="cesec200"><title>Pharyngostomy Tube Placement (Not Generally Recommended)</title><p id="para608">Originally indicated for use in dogs having long-term enteral feeding requirement, the pharyngostomy tube is associated with a number of complications, including laryngeal obstruction, aspiration, reflux, epiglottic entrapment, ulcerative esophagitis, and tube displacement. This technique generally is not recommended today. The technique entails passing a 14F to 20F tube, measured to a length such that the tube tip will not enter the stomach, through the skin overlying the pharynx, caudal to the hyoid apparatus, and into the esophagus. Although the surgery is not particularly complex, the exit point of the tube through the wall of pharynx requires that the skin and tissue of the pharynx be incised in proximity to the external jugular vein, carotid artery, vagosympathetic trunk, and hypoglossal nerve. Alternative options make this technique the least attractive and perhaps highest maintenance.</p></sec><sec id="cesec201"><title>Esophagostomy Tube Placement</title><p id="para609">Less invasive and not requiring endoscopy equipment, esophagostomy tube placement in dogs and cats is an alternative technique to use in patients that have long-term feeding needs. Use a 14F to 20F rubber, polyurethane, or silicone feeding tube placed at the level of the middle of the cervical esophagus to the level of the eighth rib. The technique does require general anesthesia or, in the hands of an experienced individual, short-term intravenous anesthesia. The technique has been described in detail in textbooks (see “Marks SL” under Additional Reading). However, one should observe the technique being performed by someone with experience before attempting to place an esophagostomy tube for the first time. Although postplacement complications generally are limited to local irritation or minor infection at the site of the stoma in the midcervical region, variations to performing the procedure are described, and the placement technique is not intuitive from published descriptions.</p></sec><sec id="cesec202"><title>Percutaneous Gastrostomy Tube Placement</title><p id="para610">Percutaneous gastrostomy tubes are used routinely to administer nutrients and medications orally over days or weeks to cats and dogs that cannot have nutrients administered by mouth or that will not eat (e.g., because of feline hepatic lipidosis, oropharyngeal neoplasms, maxillary or mandibular fractures, oral reconstructive surgery, esophageal masses or foreign bodies, or severe pharyngitis). The percutaneous gastrostomy tube is placed so that it extends through the skin and left cranial abdominal wall of the abdomen into the body of the stomach.</p><sec id="cesec203"><title>Catheter preparation</title><p id="para611">Catheter preparation for a percutaneous gastrostomy tube is as follows:<list list-type="simple" id="celist20"><list-item id="celistitem106"><label>1.</label><p id="para612">Use the French-Pezzar mushroom-tipped catheter.</p></list-item><list-item id="celistitem107"><label>2.</label><p id="para613">Cut off 1.5 cm of the open (distal) end of the catheter with scissors.</p></list-item><list-item id="celistitem108"><label>3.</label><p id="para614">Cut 3-mm holes on either side of the 1.5-cm piece (outer flange).</p></list-item><list-item id="celistitem109"><label>4.</label><p id="para615">Cut the distal end of the catheter to form a sharp bevel point.</p></list-item><list-item id="celistitem110"><label>5.</label><p id="para616">Measure the length of the tube from the mushroom tip to 2 cm below the bevel.</p></list-item></list></p></sec><sec id="cesec204"><title>Preparation of the stomach tube</title><p id="para617">Stomach tube preparation for a percutaneous gastrostomy tube is as follows:<list list-type="simple" id="celist21"><list-item id="celistitem111"><label>1.</label><p id="para618">Use a smooth-ended vinyl stomach tube.</p></list-item><list-item id="celistitem112"><label>2.</label><p id="para619">Measure the length of the tube needed to reach the stomach by laying the tube along the animal's side with the rounded end 1 to 2 cm caudal to the last rib.</p></list-item><list-item id="celistitem113"><label>3.</label><p id="para620">Mark the tube with an indelible marker or adhesive tape at the tip of the muzzle and cut off the excess tube.</p></list-item><list-item id="celistitem114"><label>4.</label><p id="para621">Put the tube in the freezer for 30 minutes before beginning the procedure to stiffen the tube.</p></list-item></list></p></sec><sec id="cesec205"><title>Placement of the percutaneous gastrotomy tube</title><p id="para622">Placement of a percutaneous gastrostomy tube is as follows:<list list-type="simple" id="celist22"><list-item id="celistitem115"><label>1.</label><p id="para623">Clip and surgically prepare the skin over the left abdominal wall.</p></list-item><list-item id="celistitem116"><label>2.</label><p id="para624">Place the mouth speculum between the right canine teeth.</p></list-item><list-item id="celistitem117"><label>3.</label><p id="para625">Place the stomach tube in the esophagus to the level of the cardia.</p></list-item><list-item id="celistitem118"><label>4.</label><p id="para626">Rotate the tube counterclockwise while carefully advancing it through the cardia.</p></list-item><list-item id="celistitem119"><label>5.</label><p id="para627">Turn the tube back clockwise and advance the tube until it can be visualized through the abdominal wall 1 to 2 cm caudal to the last rib (<xref rid="f39" ref-type="fig">Figure 4-39</xref>).<fig id="f39"><label>Figure 4-39</label><caption><p>Locating the end of the rigid stomach tube at the left lateral abdominal wall.</p></caption><graphic xlink:href="gr39"></graphic><attrib>(From Crow S, Walshaw S: <italic>Manual of clinical procedures in the dog, cat, and rabbit,</italic> ed 2, Philadelphia, 1997, Lippincott-Raven.)</attrib><permissions><copyright-statement>© 2006 Lippincott-Raven</copyright-statement><copyright-year>2006</copyright-year><license><license-p>Since January 2020 Elsevier has created a COVID-19 resource centre with free information in English and Mandarin on the novel coronavirus COVID-19. The COVID-19 resource centre is hosted on Elsevier Connect, the company's public news and information website. Elsevier hereby grants permission to make all its COVID-19-related research that is available on the COVID-19 resource centre - including this research content - immediately available in PubMed Central and other publicly funded repositories, such as the WHO COVID database with rights for unrestricted research re-use and analyses in any form or by any means with acknowledgement of the original source. These permissions are granted for free by Elsevier for as long as the COVID-19 resource centre remains active.</license-p></license></permissions></fig></p></list-item><list-item id="celistitem120"><label>6.</label><p id="para628">Rotate the tube so that the tip lies against the stomach and abdominal wall one third of the distance between the epaxial muscles and the ventral midline.</p></list-item><list-item id="celistitem121"><label>7.</label><p id="para629">Make a 2- to 3-mm skin incision directly over the lumen of the stomach tube.</p></list-item><list-item id="celistitem122"><label>8.</label><p id="para630">Use a Sovereign catheter (over the needle) and puncture the abdominal and stomach walls, placing the catheter inside the lumen of the stomach tube. Remove the needle (<xref rid="f40" ref-type="fig">Figure 4-40</xref>).<fig id="f40"><label>Figure 4-40</label><caption><p>Placement of the Sovereign catheter through the abdominal and stomach walls and into the lumen of the somach tube.</p></caption><graphic xlink:href="gr40"></graphic><attrib>(From Crow S, Walshaw S: <italic>Manual of clinical procedures in the dog, cat, and rabbit,</italic> ed 2, Philadelphia, 1997, Lippincott-Raven.)</attrib><permissions><copyright-statement>© 2006 Lippincott-Raven</copyright-statement><copyright-year>2006</copyright-year><license><license-p>Since January 2020 Elsevier has created a COVID-19 resource centre with free information in English and Mandarin on the novel coronavirus COVID-19. The COVID-19 resource centre is hosted on Elsevier Connect, the company's public news and information website. Elsevier hereby grants permission to make all its COVID-19-related research that is available on the COVID-19 resource centre - including this research content - immediately available in PubMed Central and other publicly funded repositories, such as the WHO COVID database with rights for unrestricted research re-use and analyses in any form or by any means with acknowledgement of the original source. These permissions are granted for free by Elsevier for as long as the COVID-19 resource centre remains active.</license-p></license></permissions></fig></p></list-item><list-item id="celistitem123"><label>9.</label><p id="para631">Thread a long, rigid suture through the catheter and advance it through the stomach tube until the end is observed at the mouth end of the tube (<xref rid="f41" ref-type="fig">Figure 4-41</xref>).<fig id="f41"><label>Figure 4-41</label><caption><p>Threading the introduction line retrograde through the Sovereign catheter and stomach tube.</p></caption><graphic xlink:href="gr41"></graphic><attrib>(From Crow S, Walshaw S: <italic>Manual of clinical procedures in the dog, cat, and rabbit,</italic> ed 2, Philadelphia, 1997, Lippincott-Raven.)</attrib><permissions><copyright-statement>© 2006 Lippincott-Raven</copyright-statement><copyright-year>2006</copyright-year><license><license-p>Since January 2020 Elsevier has created a COVID-19 resource centre with free information in English and Mandarin on the novel coronavirus COVID-19. The COVID-19 resource centre is hosted on Elsevier Connect, the company's public news and information website. Elsevier hereby grants permission to make all its COVID-19-related research that is available on the COVID-19 resource centre - including this research content - immediately available in PubMed Central and other publicly funded repositories, such as the WHO COVID database with rights for unrestricted research re-use and analyses in any form or by any means with acknowledgement of the original source. These permissions are granted for free by Elsevier for as long as the COVID-19 resource centre remains active.</license-p></license></permissions></fig></p></list-item><list-item id="celistitem124"><label>10.</label><p id="para632">Carefully remove the plastic catheter from the stomach tube opening and place a hemostat clamp at the end of the suture material.</p></list-item><list-item id="celistitem125"><label>11.</label><p id="para633">Remove the stomach tube over the oral end of the stiff introduction suture line.</p></list-item><list-item id="celistitem126"><label>12.</label><p id="para634">Attach the open, beveled end of the French-Pezzar catheter stomach tube to a plastic Sovereign catheter using a mattress suture (<xref rid="f42" ref-type="fig">Figure 42-42</xref>).<fig id="f42"><label>Figure 4-42</label><caption><p>Suturing the introduction line to the beveled end of the gastrostomy catheter.</p></caption><graphic xlink:href="gr42"></graphic><attrib>(From Crow S, Walshaw S: <italic>Manual of clinical procedures in the dog, cat, and rabbit,</italic> ed 2, Philadelphia, 1997, Lippincott-Raven.)</attrib><permissions><copyright-statement>© 2006 Lippincott-Raven</copyright-statement><copyright-year>2006</copyright-year><license><license-p>Since January 2020 Elsevier has created a COVID-19 resource centre with free information in English and Mandarin on the novel coronavirus COVID-19. The COVID-19 resource centre is hosted on Elsevier Connect, the company's public news and information website. Elsevier hereby grants permission to make all its COVID-19-related research that is available on the COVID-19 resource centre - including this research content - immediately available in PubMed Central and other publicly funded repositories, such as the WHO COVID database with rights for unrestricted research re-use and analyses in any form or by any means with acknowledgement of the original source. These permissions are granted for free by Elsevier for as long as the COVID-19 resource centre remains active.</license-p></license></permissions></fig></p></list-item><list-item id="celistitem127"><label>13.</label><p id="para635">Force the tip of the rubber stomach tube into the large end of the Sovereign catheter.</p></list-item><list-item id="celistitem128"><label>14.</label><p id="para636">Advance the catheter-tube through the mouth and esophagus into the stomach by placing traction on the abdominal end of the introduction line.</p></list-item><list-item id="celistitem129"><label>15.</label><p id="para637">The catheter will emerge through the skin incision followed by the rubber tube. Grasp the tube with forceps and pull it through the incision opening (<xref rid="f43" ref-type="fig">Figure 4-43, A</xref>).<fig id="f43"><label>Figure 4-43</label><caption><p>Catheter-tube assembly being pulled through the mouth and esophagus and the stomach and abdominal walls.</p></caption><graphic xlink:href="gr43"></graphic><attrib>(From Crow S, Walshaw S: <italic>Manual of clinical procedures in the dog, cat, and rabbit,</italic> ed 2, Philadelphia, 1997, Lippincott-Raven.)</attrib><permissions><copyright-statement>© 2006 Lippincott-Raven</copyright-statement><copyright-year>2006</copyright-year><license><license-p>Since January 2020 Elsevier has created a COVID-19 resource centre with free information in English and Mandarin on the novel coronavirus COVID-19. The COVID-19 resource centre is hosted on Elsevier Connect, the company's public news and information website. Elsevier hereby grants permission to make all its COVID-19-related research that is available on the COVID-19 resource centre - including this research content - immediately available in PubMed Central and other publicly funded repositories, such as the WHO COVID database with rights for unrestricted research re-use and analyses in any form or by any means with acknowledgement of the original source. These permissions are granted for free by Elsevier for as long as the COVID-19 resource centre remains active.</license-p></license></permissions></fig></p></list-item><list-item id="celistitem130"><label>16.</label><p id="para638">Remove the catheter by cutting it off 2 cm below the beveled tip. Pull the rubber tube through the abdominal wall until slight resistance is felt (<xref rid="f43" ref-type="fig">Figure 43-43, B</xref>).</p></list-item><list-item id="celistitem131"><label>17.</label><p id="para639">Slide the outer flange over the end of the tube down to the skin level (<xref rid="f44" ref-type="fig">Figure 4-44</xref>).<fig id="f44"><label>Figure 4-44</label><caption><p>Diagram showing the inner and outer flanges in place against stomach mucosa and skin, respectively.</p></caption><graphic xlink:href="gr44"></graphic><attrib>(From Crow S, Walshaw S: <italic>Manual of clinical procedures in the dog, cat, and rabbit,</italic> ed 2, Philadelphia, 1997, Lippincott-Raven.)</attrib><permissions><copyright-statement>© 2006 Lippincott-Raven</copyright-statement><copyright-year>2006</copyright-year><license><license-p>Since January 2020 Elsevier has created a COVID-19 resource centre with free information in English and Mandarin on the novel coronavirus COVID-19. The COVID-19 resource centre is hosted on Elsevier Connect, the company's public news and information website. Elsevier hereby grants permission to make all its COVID-19-related research that is available on the COVID-19 resource centre - including this research content - immediately available in PubMed Central and other publicly funded repositories, such as the WHO COVID database with rights for unrestricted research re-use and analyses in any form or by any means with acknowledgement of the original source. These permissions are granted for free by Elsevier for as long as the COVID-19 resource centre remains active.</license-p></license></permissions></fig></p></list-item><list-item id="celistitem132"><label>18.</label><p id="para640">Apply antimicrobial ointment and a sterile gauze sponge over the skin incision.</p></list-item><list-item id="celistitem133"><label>19.</label><p id="para641">Bandage the gastrostomy tube in place (<xref rid="f45" ref-type="fig">Figure 4-45</xref>).<fig id="f45"><label>Figure 4-45</label><caption><p>Full abdominal bandage showing the plugged end of the gastostomy tube emerging dorsally.</p></caption><graphic xlink:href="gr45"></graphic><attrib>(From Crow S, Walshaw S: <italic>Manual of clinical procedures in the dog, cat, and rabbit,</italic> ed 2, Philadelphia, 1997, Lippincott-Raven.)</attrib><permissions><copyright-statement>© 2006 Lippincott-Raven</copyright-statement><copyright-year>2006</copyright-year><license><license-p>Since January 2020 Elsevier has created a COVID-19 resource centre with free information in English and Mandarin on the novel coronavirus COVID-19. The COVID-19 resource centre is hosted on Elsevier Connect, the company's public news and information website. Elsevier hereby grants permission to make all its COVID-19-related research that is available on the COVID-19 resource centre - including this research content - immediately available in PubMed Central and other publicly funded repositories, such as the WHO COVID database with rights for unrestricted research re-use and analyses in any form or by any means with acknowledgement of the original source. These permissions are granted for free by Elsevier for as long as the COVID-19 resource centre remains active.</license-p></license></permissions></fig></p></list-item></list></p><sec id="cesec206"><sec id="cesec207"><title>Additional Reading</title><p id="para642">Crow S, Walshaw S: <italic>Manual of clinical procedures in the dog, cat, and rabbit,</italic> ed 2, Philadelphia, 1997, Lippincott-Raven.</p><p id="para643">Marks SL: Nasoesophageal, esophagostomy, and gastrostomy tube placement techniques. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p><p id="para644">Mazzaferro EM: Esophagostomy tubes: don't underutilize them!, <italic>J Vet Emerg Care</italic> 11(2): 153-156, 2001.</p></sec></sec></sec></sec></sec></sec><sec id="cesec208"><sec id="cesec209"><title>LAPAROSCOPY</title><p id="para645">Laparoscopy is a procedure for performing a visual examination of the peritoneal cavity and its contents after the establishment of pneumoperitoneum. The advantage of the procedure is that it is minimally invasive, although general anesthesia is required. Laparoscopy does allow limited visualization of several organs, at least in part. Visualization of needle biopsies of kidney, liver, and/or spleen is also possible during the procedure. The disadvantage is that the required equipment can be expensive and specific training and experience are required to become efficient at the procedure. Additionally, insufflation of the abdomen must be monitored in addition to monitoring the patient's quality of respirations and oxygenation (pulse oximetry).</p><p id="para646">The Dyonics Needlescope (Smith & Nephew, Andover, Massachusetts) is a small fiberoptic laparoscope 1.7 or 2.2 mm in diameter. The device does require a high-intensity light source, but because of its small size, it can be inserted readily into the abdomen. Abdominal insufflation and laparoscopy require general anesthesia, neuroleptanalgesia with local anesthesia, or rarely (in the critically ill animal), regional local anesthesia alone. The depth and type of anesthesia or analgesia depend on the condition of the patient and the skill and experience of the examiner.</p><p id="para647">Before laparoscopy, perform a cleansing enema. Surgically prepare the laparoscopy site. To insufflate the abdomen, use a Verees pneumoperitoneum needle. Place the needle 3 to 4 cm below the umbilicus, along the linea alba. Inject 10 mL of saline through the needle, and attempt aspiration to ensure that a blood vessel or hollow viscus has not been penetrated. The intraabdominal pressure created should not be greater than 20 mm Hg. Remove and examine any ascitic fluid that is present.</p><p id="para648">Inject air into the peritoneal cavity through an in-line filter with the Verees needle. Insufflation should be slow, and vital signs should be monitored. Following effective insufflation, remove the needle, make a small skin incision over the needle entry point, and insert the larger trocar and cannula at a 30-degree angle to the animal's longitudinal plane. Take extreme care when placing the trocar into the abdomen. Move the endoscope (Needlescope) cephalad along the abdominal wall while maintaining good insufflation. Rotate the animal into different positions to enable visualization of various internal organs. Biopsy specimens can be obtained through the Needlescope or through a separate incision while observing through the Needlescope. When endoscopic inspection has been completed, remove the Needlescope, allow the insufflated air to escape, and place the skin sutures.</p><p id="para649">Indications for laparoscopy include biopsy, visual diagnosis, follow-up examinations, and research needs. Contraindications to laparoscopy include peritonitis, hernias, coagulation defects, obesity, abdominal adhesions, and inexperience of the clinician.</p><sec id="cesec210"><sec id="cesec211"><sec id="cesec212"><sec id="cesec213"><title>Additional Reading</title><p id="para650">Guilford WG, Center SA, Strombeck DR, et al: <italic>Strombeck's small animal gastroenterology,</italic> ed 3, Philadelphia, 1996, WB Saunders.</p><p id="para651">Richter K: Laparoscopy. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p></sec></sec></sec></sec></sec></sec><sec id="cesec214"><sec id="cesec215"><title>Ophthalmic Procedures</title><sec id="cesec216"><title>EVALUATION OF TEAR PRODUCTION</title><p id="para652">Tear production comes predominantly from the tarsal and conjunctival glands and from the accessory tarsal glands. The reflex tear secretors are the main lacrimal gland and the accessory lacrimal glands. The production of normal lacrimal secretions can be tested by using Schirmer's tear test, a standardized filter paper (<xref rid="f46" ref-type="fig">Figure 4-46</xref>) that effectively measures the rate of tear production in millimeters per minute. Schirmer's tear strips now are impregnated with a blue dye to facilitate visualization of the distance (in millimeters) that the tear migrates during the 1-minute test.<fig id="f46"><label>Figure 4-46</label><caption><p>Schirmer's tear test strips depicting tear production as a function of distance (millimeters).</p></caption><graphic xlink:href="gr46"></graphic></fig></p><p id="para653">Each eye can be tested independently, or both eyes can be tested simultaneously in the cooperative patient. Carefully fold the notched end of the test strip before removing it from the plastic package. Insert the folded end into the lower conjunctival cul-de-sac (<xref rid="f47" ref-type="fig">Figure 4-47</xref>) and begin the timing. Maintain the Schirmer's test strip in position by gently holding the eyelids closed but not touching the paper. At the end of 1 minute, note the degree (distance) of wetting that occurred and record it in the medical record. The normal dog and cat should produce wetting over 10 to 25 mm in 1 minute for each eye. Amounts less than that are consistent with keratoconjunctivitis sicca. Amounts greater than 25 mm may be normal or may be consistent with excessive tear production, or epiphora.<fig id="f47"><label>Figure 4-47</label><caption><p>Placement of a Schirmer's tear test strip into the lower conjuctival cul-de-sac of a dog; the test strip is held in place for 60 seconds only.</p></caption><graphic xlink:href="gr47"></graphic></fig></p></sec><sec id="cesec217"><title>Fluorescein Staining of the Cornea</title><p id="para654">The cornea is composed of various layers of specialized avascular epithelium and stroma. The outer layer, the corneal epithelium, is a highly sensitive, thin layer overlying the corneal stroma, the thickest layer. Descemet's membrane is a distinct, thin layer of tissue beneath the stroma. The innermost layer of the cornea is the endothelium. Damage to the corneal epithelium occurs frequently in dogs and cats. Clinical presentation typically is characterized by blepharospasm of the affected eye with or without a visible ocular discharge or conjunctivitis.</p><p id="para655">Whenever superficial corneal injury is suspected, assessment of the integrity of the corneal epithelium is indicated. Fluorescein dye-impregnated test strips can be used to determine whether the epithelial barrier overlying the corneal stroma has been disrupted and, as such, can establish the presence or absence of a corneal ulcer (<xref rid="f48" ref-type="fig">Figure 4-48</xref>).<fig id="f48"><label>Figure 4-48</label><caption><p>Fluorescein sodium–impregnated test strip used to enhance visualization of a corneal ulcer.</p></caption><graphic xlink:href="gr48"></graphic></fig></p><p id="para656">The test is simple to accomplish. Moisten the dye-impregnated tip of the test strip with a drop of balanced saline solution (or commercial ocular irrigation solution). Gently allow the tip of the test paper to touch the cornea, or sclera, of the affected eye. (In patients with particularly painful, sensitive eye, use a topical anesthetic to moisten the test strip or apply the anesthetic directly to the cornea before testing.) Immediately rinse the eye with a sterile irrigation solution to remove the excess dye (the test strip has a lot of dye; be prepared to catch the excess fluid with a 2 × 2-inch gauze).</p><p id="para657">Promply examine the eye with a direct, focal light source. Evidence of green dye uptake in the stroma indicates that an ulcer is present. The absence of staining generally indicates that the corneal integrity is intact. One exception exists. Descemet's membrane will not take up fluorescein dye. A patient with a deep corneal ulcer that penetrates through the corneal stroma and allows herniation of Descemet's membrane (descemetocele) will not demonstrate a positive stain. Careful visualization of the cornea, however, is likely to reveal the presence of a such a serious, deep ulcer.</p></sec><sec id="cesec218"><title>Assessment of Nasolacrimal Duct Patency</title><p id="para658">Fluroescein dye can be used to assess patency of the nasolacrimal duct. To perform this examination, place a drop of fluorescein dye from a sterile fluorescein strip into the eye and add 1 to 2 drops of a sterile eye wash. After 2 to 5 minutes, examine the external nares with the aid of a cobalt blue filter or Wood's light for the presence or absence of fluorescence. A clean, 2 × 2-inch white gauze touched against the nasal planum also will pick up the green-colored dye if the duct is patent. If dye is present, the lacrimal excretory system is patent and functioning. If epiphora exists but the primary dye test indicates that the lacrimal excretory system is patent, hypersecretion of tear fluid may be implicated as the cause of the epiphora.</p><p id="para659">Irrigation of the nasolacrimal system is indicated if the primary dye test is negative. In the dog, the nasolacrimal puncta are located 1 to 3 mm from the medial canthus on the mucocutaneous border of the upper and lower lids. In the dog, use a 20- to 22-gauge (in the cat, a 23-gauge) nasolacrimal cannula (<xref rid="f49" ref-type="fig">Figure 4-49</xref>). Topical anesthesia often is required. Fill a 2-mL syringe with saline, and attach the lacrimal cannula and pass it into the lacrimal puncta of the upper lid.<fig id="f49"><label>Figure 4-49</label><caption><p>Lacrimal canulas used to flush the nasolacrimal ducts.</p></caption><graphic xlink:href="gr49"></graphic></fig></p><p id="para660">Several points should be made about evaluating the nasolacrimal system. Brachycephalic breeds of dogs and cats occasionally may have a negative primary dye test, although no blockage in the nasolacrimal system exists. In flushing the nasolacrimal system of some animals, fluid may not appear at the nose; however, the animal may gag and exhibit swallowing movements, indicating that the fluid has entered the mouth and the system is patent.</p></sec><sec id="cesec219"><title>Conjunctival Smears, Scrapings, and Cultures</title><p id="para661">In performing conjunctival scrapings, use a platinum spatula (Kimura spatula) the tip of which has been sterilized previously in the flame of an alcohol lamp and has been allowed to cool. Scrape the inferior conjunctival cul-de-sac, preferably without prior topical anesthesia, because anesthetics may distort the cells (<xref rid="f50" ref-type="fig">Figure 4-50</xref>). Place the material on two glass slides. Fix one slide in acetone-free 95% methanol for 5 to 10 minutes; then stain the slide with Giemsa stain. Heat-fix the other slide, and apply Gram stain.<fig id="f50"><label>Figure 4-50</label><caption><p>Spatula used for performing conjunctival scrapings.</p></caption><graphic xlink:href="gr50"></graphic></fig></p><p id="para662">To culture the conjunctiva, use sterile cotton-tipped applicators, fluid thioglycollate medium, and blood agar medium. Evert the palpebral conjunctiva of the lower lid, and pass one side of a sterile cotton applicator, previously moistened with sterile broth or thioglycollate medium, over the palpebral conjunctival surface. Streak the swab onto a sterile blood agar plate; then place the plate in a tube of thioglycollate broth. No topical anesthesia is used before culturing because preservatives present in anesthetics can inhibit the growth of bacteria.</p></sec><sec id="cesec220"><title>Tonometry</title><p id="para663">Glaucoma is an increase in intraocular pressure incompatible with normal ocular and visual functions. One method used to measure intraocular pressure is tonometry, in which the tension of the outer coat of the eye is assessed by measuring the impressibility, or applanability, of the cornea. Because the measurements based on tonometry involve calculations that have a wide base of variations, tonometry readings are always approximations.</p><sec id="cesec221"><title>Schiøtz tonometry</title><p id="para664">The Schiøtz tonometer consists of a corneal footplate, plunger, holding bracket, recording scale, and 5.5-, 7.5-, 10.0-, and 15.0-g weights (<xref rid="f51" ref-type="fig">Figure 4-51</xref>). The principle of the Schiøtz tonometer is that the amount that the plunger protrudes from the footplate is related to the indentability of the cornea, which in turn is related to the intraocular pressure. The plunger is connected to a scale so that 0.05-mm protrusion of the plunger equals 1 scale unit (<xref rid="cetable10" ref-type="table">Table 4-10</xref>).<fig id="f51"><label>Figure 4-51</label><caption><p>Footplate (for corneal contact) of the Schiøtz tonometer.</p></caption><graphic xlink:href="gr51"></graphic></fig><table-wrap position="float" id="cetable10"><label>TABLE 4-10</label><caption><p>Schiøtz Tonometer—Calibration Table for the Canine Eye</p></caption><table frame="hsides" rules="groups"><thead><tr><th></th><th colspan="3" align="center">Intraocular pressure (mm Hg)<hr></hr></th></tr><tr><th align="center">Schiøtz scale reading</th><th align="center">5.5 g Weight</th><th align="center">7.5 g Weight</th><th align="center">10.0 g Weight</th></tr></thead><tbody><tr><td align="char">0.5</td><td align="char">52.6</td><td align="char">71.2</td><td align="char">93.6</td></tr><tr><td align="char">1.0</td><td align="char">49.3</td><td align="char">67.0</td><td align="char">88.3</td></tr><tr><td align="char">1.5</td><td align="char">46.3</td><td align="char">63.1</td><td align="char">83.3</td></tr><tr><td align="char">2.0</td><td align="char">43.4</td><td align="char">59.4</td><td align="char">78.6</td></tr><tr><td align="char">2.5</td><td align="char">40.8</td><td align="char">55.9</td><td align="char">74.1</td></tr><tr><td align="char">3.0</td><td align="char">38.3</td><td align="char">52.6</td><td align="char">69.6</td></tr><tr><td align="char">3.5</td><td align="char">36.0</td><td align="char">49.6</td><td align="char">66.0</td></tr><tr><td align="char">4.0</td><td align="char">33.9</td><td align="char">46.7</td><td align="char">62.2</td></tr><tr><td align="char">4.5</td><td align="char">31.9</td><td align="char">44.0</td><td align="char">58.7</td></tr><tr><td align="char">5.0</td><td align="char">30.1</td><td align="char">41.6</td><td align="char">55.4</td></tr><tr><td align="char">5.5</td><td align="char">28.4</td><td align="char">39.2</td><td align="char">52.3</td></tr><tr><td align="char">6.0</td><td align="char">26.9</td><td align="char">37.1</td><td align="char">49.4</td></tr><tr><td align="char">6.5</td><td align="char">25.5</td><td align="char">35.1</td><td align="char">46.7</td></tr><tr><td align="char">7.0</td><td align="char">24.2</td><td align="char">33.2</td><td align="char">44.2</td></tr><tr><td align="char">7.5</td><td align="char">23.0</td><td align="char">31.5</td><td align="char">41.8</td></tr><tr><td align="char">8.0</td><td align="char">21.9</td><td align="char">29.9</td><td align="char">39.6</td></tr><tr><td align="char">8.5</td><td align="char">21.0</td><td align="char">28.5</td><td align="char">37.5</td></tr><tr><td align="char">9.0</td><td align="char">20.1</td><td align="char">27.1</td><td align="char">35.6</td></tr><tr><td align="char">9.5</td><td align="char">19.3</td><td align="char">25.9</td><td align="char">33.8</td></tr><tr><td align="char">10.0</td><td align="char">18.6</td><td align="char">24.8</td><td align="char">32.1</td></tr><tr><td align="char">10.5</td><td align="char">18.0</td><td align="char">23.8</td><td align="char">30.6</td></tr><tr><td align="char">11.0</td><td align="char">17.4</td><td align="char">22.8</td><td align="char">29.1</td></tr><tr><td align="char">11.5</td><td align="char">17.0</td><td align="char">22.0</td><td align="char">27.8</td></tr><tr><td align="char">12.0</td><td align="char">16.6</td><td align="char">21.3</td><td align="char">26.6</td></tr><tr><td align="char">12.5</td><td align="char">16.3</td><td align="char">20.6</td><td align="char">25.5</td></tr><tr><td align="char">13.0</td><td align="char">16.0</td><td align="char">20.0</td><td align="char">24.5</td></tr><tr><td align="char">13.5</td><td align="char">15.8</td><td align="char">19.5</td><td align="char">23.6</td></tr><tr><td align="char">14.0</td><td align="char">15.7</td><td align="char">19.1</td><td align="char">22.8</td></tr><tr><td align="char">14.5</td><td align="char">15.7</td><td align="char">18.8</td><td align="char">22.0</td></tr><tr><td align="char">15.0</td><td align="char">15.7</td><td align="char">18.5</td><td align="char">21.4</td></tr><tr><td align="char">15.5</td><td align="char">15.8</td><td align="char">18.3</td><td align="char">20.8</td></tr><tr><td align="char">16.0</td><td align="char">15.9</td><td align="char">18.1</td><td align="char">20.3</td></tr><tr><td align="char">16.5</td><td align="char">16.1</td><td align="char">18.0</td><td align="char">19.9</td></tr><tr><td align="char">17.0</td><td align="char">16.4</td><td align="char">18.0</td><td align="char">19.5</td></tr><tr><td align="char">17.5</td><td align="char">16.8</td><td align="char">18.1</td><td align="char">19.2</td></tr><tr><td align="char">18.0</td><td align="char">17.2</td><td align="char">18.2</td><td align="char">19.0</td></tr><tr><td align="char">18.5</td><td align="char">17.7</td><td align="char">18.4</td><td align="char">18.8</td></tr><tr><td align="char">19.0</td><td align="char">18.3</td><td align="char">18.7</td><td align="char">18.7</td></tr><tr><td align="char">19.5</td><td align="char">19.0</td><td align="char">19.0</td><td align="char">18.6</td></tr><tr><td align="char">20.0</td><td align="char">19.7</td><td align="char">19.4</td><td align="char">18.7</td></tr></tbody></table></table-wrap></p><p id="para665">Place the dog in the sitting or dorsal recumbent position. Instill topical anesthesia, and hold open the eyelids with the fingers, which are placed far away from the lid margins. Place the footplate vertically on the central aspect of the cornea (<xref rid="f52" ref-type="fig">Figure 4-52</xref>). Take three readings in each eye and then average the readings. Normal intraocular tension with the Schiøtz tonometer in dogs is 15 to 25 mm Hg.<fig id="f52"><label>Figure 4-52</label><caption><p>Technique for measuring intraocular pressure using the Schiøtz tonometer.</p></caption><graphic xlink:href="gr52"></graphic></fig></p></sec><sec id="cesec222"><title>Applanation tonometry</title><p id="para666">In applanation tonometry, a very small area of the cornea is flattened by a known force, usually a calibrated burst of air. The advantage of this technique over the indentation (Schiøtz) method is that the errors resulting from ocular rigidity and corneal curvature are greatly reduced. Special equipment is required to perform applanation tonometry.</p></sec></sec><sec id="cesec223"><title>Gonioscopy</title><p id="para667">Glaucoma can be caused by many different disorders that elevate intraocular pressure. In many types of glaucoma, there is an abnormality in the anterior angle of the eye (filtration angle). Gonioscopy permits one to visualize and examine the iridocorneal angle, which cannot be seen without the use of a special contact lens.</p><p id="para668">The Koeppe gonioscopic lens seems to be well-suited to dogs and cats. The lens is available in 17-, 19-, and 21-mm sizes. The lens can be inserted into the eye following the application of topical anesthesia. The gonioscopic lens can be filled with 1% methylcellulose or saline. The inside of the lens is illuminated with a Barkan lamp, otoscope head, or binocular indirect ophthalmoscope. Magnification suitable for visualization of the angle can be provided by an otoscope head, indirect ophthalmoscope, or Haag-Streit goniomicroscope.</p><sec id="cesec224"><sec id="cesec225"><sec id="cesec226"><title>Additional Reading</title><p id="para669">Barnett KC, Sansom J, Heinrich C: <italic>Canine ophthalmology</italic>, Philadelphia, 2002, WB Saunders.</p><p id="para670">Barnett KC, Crispin SM: <italic>Feline ophthalmology</italic>, Philadelphia, 1998, WB Saunders.</p></sec></sec></sec></sec></sec></sec><sec id="cesec227"><sec id="cesec228"><title>RADIOGRAPHY: ADVANCED CONTRAST STUDIES</title><sec id="cesec229"><title>Gastrointestinal Studies</title><p id="para671">When considering a contrast study of the gastrointestinal tract, it is not unreasonable to question the value of doing this procedure. At issue is the fact that abdominal ultrasound and/or gastrointestinal endoscopy have largely replaced contrast radiography of the gastrointestinal tract and for good reason. Diagnostic modalities such as ultrasound (in the hands of an experienced individual) and endoscopy have a much greater diagnostic yield than the less sensitive contrast study. So, why even try? Endoscopes are not available in every practice, and access to an ultrasound, much less someone who is qualified to use the equipment, puts routine use of advanced diagnostic modalities out of reach for many practices. However, it must be appreciated that the diagnostic value of a radiographic contrast study of the gastrointestinal tract is a far less sensitive diagnostic modality than abdominal ultrasound or endoscopy. However, the procedure for gastrointestinal radiographic constrast study is outlined next.</p><p id="para672">Contrast agents available for gastrointestinal studies include barium suspension preparations or Micropaque (Guerbet, Villepente, France); and water-soluble agents (Gastrografin [Bracco Diagnostics Inc., Princeton, New Jersey], which is 60% meglumine and 10% sodium diatrizoate). Water-soluble agents are used if bowel perforation is suspected. Undiluted water-soluble agents are hypertonic and should be diluted at a ratio of one part Gastrografin to two parts water. No single procedure is appropriate for all gastrointestinal cases. The clinician must select procedures based on the clinical history and physical findings, apparent location of the lesion within the gastrointestinal tract, endoscopic findings, and results from other imaging studies, such as abdominal ultrasound.</p><sec id="cesec230"><title>Contrast esophagram</title><p id="para673">The contrast esophagram also is called barium swallow. The decision to perform a contrast esophagogram is based on physical evidence of dysphagia (difficulty or pain while attempting to swallow) and/or persistent regurgitation (reflux of swallowed food without effort). The procedure necessitates that the animal fast for 12 hours before radiography. Remove all leashes from around the animal's neck, and obtain survey radiographs of the thorax. In esophageal contrast studies, administer barium suspension contrast medium, 2 to 5 mL/kg body mass. <italic>Administration of barium as a contrast material is contraindicated if a perforation of the esophagus is suspected.</italic> When the esophagus has been coated with radiopaque material, take lateral, ventrodorsal, and right ventrodorsal oblique thoracic radiographs to visualize the esophagus.</p><p id="para674">Properly prepared, the barium should be thick and pasty (like marshmallow fluff). Position the patient and cassette, and have the radiographic technique set up. Give a tablespoonful of barium orally. Make the exposure when the animal takes its second swallow after the barium has been given.</p><p id="para675">For esophageal studies and barium swallows, sedation with acepromazine 0.1 mg/kg and buprenorphine 0.015 mg/kg will produce no adverse alteration in gastrointestinal motility. For cats, ketamine 10 mg and midazolam 0.2 mg/kg (combined) can be administered intramuscularly with no significant effect in esophageal motility.</p><p id="para676">In some cases of incomplete esophageal stricture, barium liquid will pass through the esophagus unobstructed, whereas food will not. Veterinarians have to be particularly helpful in these patients to mix kibbled food with the barium and allow the patient to eat the mixture just before making the radiograph.</p><p id="para677">Ideally, however, contrast esophagrams are performed using fluoroscopy, rather than conventional radiographs. In this manner, it is possible not only to identify strictures and dilatations, if present, but also to obtain a dynamic study of the esophagus that provides valuable information pertaining to swallowing and esophageal motility and function and an opportunity to evaluate sphincter activity at the level of the cardia.</p></sec><sec id="cesec231"><title>Upper gastrointestinal tract (stomach, pylorus, and small intestine)</title><p id="para678">Contrast studies of the upper gastrointestinal tract are used to facilitate diagnosis of persistent vomiting, hematemesis, unexplained and chronic diarrhea, suspected enteric foreign bodies, suspected neoplasms and obstructions, and for confirmation of displaced intestinal organs, as may be seen in diaphragmatic hernias.</p><p id="para679">That said, the availability of abdominal ultrasound has largely replaced the upper gastrointestinal series. At the hands of an experienced ultrasonographer, the diagnostic value of abdominal ultrasound far exceeds that derived from evaluating sequential radiographs of a patient after oral administration of contrast medium such as barium. In the event ultrasound capability is <italic>not</italic> available, contrast study of the upper gastrointestinal tract still can be used. However, the clinician must appreciate that a barium contrast study of the stomach, duodenum, jejunum, and ileum has a low sensitivity as a diagnostic test. That is, negative findings are not expected to correlate well with the absence of clinical disease. A negative study does not rule out disease. Likewise, a contrast study of the upper gastrointestinal tract is not recognized for its ability to confirm a diagnosis gastrointestinal tract disease, even when disease is present. Perhaps the greatest value in performing the upper gastrointestinal series in a dog or cat today centers on the need to identify a displacement of the stomach and/or small intestine because of an extraluminal mass lesion or congenital defect in the patient. In addition, the use of a microfine barium suspension may facilitate identification of intestinal ulcers, irregularities (e.g., intraluminal neoplasia), and radiolucent foreign bodies. However, variable-diameter, solid-phase radiopaque markers called BIPS (barium-impregnated polyethylene spheres) can be used to assess gastric emptying time, gastrointestinal transit times, and to some extent, obstructive disorders.</p></sec><sec id="cesec232"><title>Technique for upper gastrointestinal study</title><p id="para680">If an upper gastrointestinal study is indicated, follow the technique described:<list list-type="simple" id="celist23"><list-item id="celistitem134"><label>1.</label><p id="para681">Ensure that the hair of the animal is free from dirt, paint, and foreign material. Bathe the animal if necessary.</p></list-item><list-item id="celistitem135"><label>2.</label><p id="para682">Withhold food for 18 to 24 hours.</p></list-item><list-item id="celistitem136"><label>3.</label><p id="para683">If the colon is filled with feces, administer a cleansing (Fleet) enema the evening before performing the procedure. In dogs, give a second enema 3 to 5 hours before the start of the gastrointestinal series.</p></list-item><list-item id="celistitem137"><label>4.</label><p id="para684">At the start of an upper gastrointestinal series, obtain survey radiographs of the abdomen. Administer a barium sulfate (micropulverized) preparation by stomach tube, or induce the animal to swallow the fluids. Flavored, prepared barium suspensions are available, but they taste bad (personal experience). Dosage levels vary, but for barium suspensions, give approximately 10 mL/kg. As an alternative to barium, use an organic iodide liquid preparation. Administer 0.5 mL/kg by stomach tube. Obtain lateral and dorsoventral radiographs of the abdomen immediately following administration of the contrast material and at 30-minute, 1-hour, and 2-hour intervals. Water-soluble contrast material passes through the gastrointestinal tract in 30 to 90 minutes. Barium suspensions take 60 to 180 minutes to traverse the intestine. The colon usually is filled with barium 6 hours after oral administration and may contain barium for 2 to 3 days following administration.</p></list-item></list></p><p id="para685">Barium contrast radiography is contraindicated if perforation of the stomach or upper gastrointestinal tract is suspected. In these cases, use water-soluble contrast media such as the oral diatrizoates because leakage into the abdomen will produce no foreign body granuloma. In addition, do not administer barium sulfate when an obstruction of the lower bowel may be present. In these cases, barium may only contribute to the obstipation.</p><p id="para686">The following radiographic views are recommended following administration of radiographic contrast material:<list list-type="simple" id="celist24"><list-item id="celistitem138"><label>1.</label><p id="para687">Immediately following administration of contrast material obtain a ventrodorsal, right lateral, and left lateral views. The right lateral view shows the pylorus of the stomach filled with barium, and the left lateral view shows the cardia and fundic portion filled with barium. The objective is to evaluate the distended stomach and initial gastric emptying.</p></list-item><list-item id="celistitem139"><label>2.</label><p id="para688">Twenty to 30 minutes following administration of contrast material, obtain ventrodorsal and right lateral views to assess the stomach, pyloric emptying, and the proximal duodenum.</p></list-item><list-item id="celistitem140"><label>3.</label><p id="para689">Sixty minutes following administration of contrast material, repeat the ventrodorsal and right lateral recumbency views to assess the small intestine.</p></list-item><list-item id="celistitem141"><label>4.</label><p id="para690">Two hours following administration of contrast material, repeat the ventrodorsal and right lateral views to evaluate passage of contrast material into the colon and complete emptying of the stomach; contrast material should be in the terminal portion of the small intestine.</p></list-item></list></p></sec><sec id="cesec233"><title>Guidelines for passage of contrast material through the gastrointestinal tract</title><p id="para691">The passage of contrast material through the normal gastrointestinal tract is variable; however, the following guidelines have been suggested:<list list-type="simple" id="celist25"><list-item id="celistitem142"><label>1.</label><p id="para692">Contrast material is in the duodenum within 15 minutes in most patients. Excitement can delay gastric emptying time to 20 to 25 minutes.</p></list-item><list-item id="celistitem143"><label>2.</label><p id="para693">Contrast material reaches the jejunum within 30 minutes and is within the jejunum and ileum at 60 minutes.</p></list-item><list-item id="celistitem144"><label>3.</label><p id="para694">Contrast material reaches the ileocecal junction in 90 to 120 minutes.</p></list-item><list-item id="celistitem145"><label>4.</label><p id="para695">At 3 to 5 hours after administration, contrast material has cleared the upper gastrointestinal tract and is within the ileum and the large intestine.</p></list-item></list></p><p id="para696">In evaluation of gastrointestinal contrast studies, consider the following criteria: (1) the size of the intestinal mass, (2) the contour of the mucosal surface, (3) thickness of the bowel wall, (4) flexibility and motility of the bowel wall, (5) position of the small intestine, (6) continuity of the opaque column, and (7) transit time.</p></sec><sec id="cesec234"><title>The barium enema</title><p id="para697">Clinical disorders for which the barium enema is indicated in dogs include ileocolic intussusception and cecal inversion (intussusception), mechanical and functional large bowel obstruction, invasive lesions of the large bowel, a mass outside the large bowel compressing the bowel, and inflammation of the lower intestinal tract. Barium sulfate enemas are contraindicated in suspected obstruction of the colon and rupture or perforation of the colon. However, these same disorders also can be identified by ultrasonic examination or colonoscopy, either of which is the preferred diagnostic modality over a barium enema.</p><p id="para698">Use the following procedure when giving barium enemas:<list list-type="simple" id="celist26"><list-item id="celistitem146"><label>1.</label><p id="para699">Twenty-four hours preceding radiographs, administer a liquid diet only, preferably water or broth.</p></list-item><list-item id="celistitem147"><label>2.</label><p id="para700">During the 18 to 24 hours before the radiographs, administer a mild high colonic enema or give a saline laxative orally.</p></list-item><list-item id="celistitem148"><label>3.</label><p id="para701">Do not give any irritating enemas within 12 hours of the scheduled radiographic examination; however, administer isotonic saline solution or plain water enemas before the examination to ensure that the bowel is clear.</p></list-item><list-item id="celistitem149"><label>4.</label><p id="para702">Obtain survey radiographs of the abdomen, and examine the colon to ensure that this portion of the bowel is clear.</p></list-item><list-item id="celistitem150"><label>5.</label><p id="para703">Do not force barium into the colon under pressure. Do not elevate the enema bag more than 18 inches above the animal.</p></list-item><list-item id="celistitem151"><label>6.</label><p id="para704">Do not perform a proctoscopic examination on the same day that the barium enema is given.</p></list-item></list></p><p id="para705">Cuffed rectal catheters (Bardex cuffed rectal catheters, 24F to 38F, and the Bardex cuffed pediatric rectal catheter, 18F [C.R. Bard, Inc., Murray Hill, New Jersey]) can be used in dogs (<xref rid="f53" ref-type="fig">Figure 4-53</xref>). For very small dogs and cats, use smaller catheters. A plastic catheter adapter and a three-way stopcock are needed. Various barium sulfate preparations can be used; however, the final concentration should be 15% to 20% w/v. A commercially available barium enema kit is helpful.<fig id="f53"><label>Figure 4-53</label><caption><p>Bardex catheter depicting the expanded balloon tip used to facilitate infusion of barium into the colon.</p></caption><graphic xlink:href="gr53"></graphic></fig></p><p id="para706">To perform a barium enema effectively, sedate or anesthetize the animal. Place the cuffed rectal catheter so that the inflated bulb is cranial to the anal sphincter. Place the animal in right lateral recumbency and fill the colon with contrast material at a dose of 20 to 30 mL/kg body mass. Take the radiographs after infusion of a two-thirds dose of barium. If the colon is not filled, infuse more contrast agent. Obtain radiographs in the ventrodorsal and lateral positions, and determine whether the colon is distended adequately. Remove as much of the contrast material as possible from the colon, and repeat the radiographs. Then insert air at 2 mL/kg into the colon, and repeat the radiographs. Deflate the cuff on the catheter, and remove the catheter from the rectum. Throughout the procedure of filling the colon with contrast material or air, take care not to overdistend the colon, which may lead to rupture.</p><p id="para707">When reviewing individual radiographs, look for the following radiographic lesions: (1) irregularity of the barium-mucosal interface; (2) spasm, stricture, or occlusion of the bowel lumen; (3) filling defects; (4) outpouching of the bowel wall caused by diverticulum or perforation; and (5) displacement of the bowel.</p><sec id="cesec235"><sec id="cesec236"><title>Additional Reading</title><p id="para708">Burk RL, Ackerman N: <italic>Small animal radiology and ultrasonography: a diagnostic atlas and text,</italic> ed 3, St Louis, 2003, Saunders.</p><p id="para709">Hall EJ, German AJ: Diseases of the small intestine. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p><p id="para710">Thrall DE: <italic>Textbook of veterinary diagnostic radiology,</italic> ed 4, Philadelphia, 2002, Saunders.</p><p id="para711">Washabau RJ, Holt DE: Diseases of the large intestine. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p></sec></sec></sec></sec></sec><sec id="cesec237"><sec id="cesec238"><title>Excretory Urography</title><p id="para712">Intravenous administration of organic iodinated compounds in high concentrations permits visualization in four phases: (1) the arteriogram, (2) the nephrogram, (3) the pyelogram, and (4) the cystogram (<xref rid="cetextbox16" ref-type="boxed-text">Box 4-16</xref>). The arterial phase demonstrates renal blood flow; the nephrogram demonstrates the accumulation of contrast agent in the renal tubules and is used to evaluate renal parenchyma; the pyelogram phase evaluates the urinary collecting system, including the ureters; and the cystogram reveals the collection of contrast agent in the urinary bladder. Excretory urography does not result in any quantitative information about renal function and is not a substitute for renal function tests. The degree of visualization of contrast material within the renal excretory system depends on the concentration of iodine in the contrast medium, the technique of excretory urography performed, the state of hydration of the patient, renal blood flow, and the functional capacity of the kidneys.<boxed-text id="cetextbox16"><label>BOX 4-16</label><caption><title>PATIENT PREPARATION FOR EXCRETORY UROGRAPHY</title></caption><p id="para113"><list list-type="simple" id="celist11"><list-item id="celistitem52"><label>1.</label><p id="para114">Have the patient fast for 12 to 18 hours.</p></list-item><list-item id="celistitem53"><label>2.</label><p id="para115">Administer a cleansing enema or give a saline laxative orally 12 to 18 hours before radiography.</p></list-item><list-item id="celistitem54"><label>3.</label><p id="para116">Ensure that the animal's hair is free of dirt and debris.</p></list-item><list-item id="celistitem55"><label>4.</label><p id="para117">Try to limit the animal's fluid intake in the 12 hours preceding radiography.</p></list-item><list-item id="celistitem56"><label>5.</label><p id="para118">Empty the animal's bladder immediately before taking radiographs.</p></list-item><list-item id="celistitem57"><label>6.</label><p id="para119">Take survey radiographs before administering contrast media.</p></list-item></list></p></boxed-text></p><p id="para713">The contrast medium most commonly used is a diatrizoate or iothalamate compound. Rapidly administer 850 mg/kg of an iodine compound intravenously. Obtain a ventrodorsal radiograph at 10 seconds after injection, and repeat ventrodorsal and lateral radiographs 1, 3, 5, 15, 20, and 40 minutes following injection. This method is the current standard technique. If the patient's blood urea nitrogen level is greater than or equal to 50 mg/dL or the creatinine level is greater than 4 mg/dL, double the dose rate.</p><p id="para714">Lesions that can be detected by using intravenous urography are renal mass lesions; neoplasia; renal cysts; renal and ureteral traumatic lesions; pyelonephritis; hydroureter; hydronephrosis; renal agenesis; hypoplasia; pelvic and ureteral obstructions (calculi, blood clots); renal parasites; ectopic ureter; and duplication of the collecting system.</p></sec><sec id="cesec239"><title>Retrograde Contrast Urethrography</title><p id="para715">Retrograde urethrography is a diagnostic tool used to localize diseases of the lower urinary tract of dogs and cats. This method can reveal conditions such as urethral neoplasms, strictures, trauma, calculi, or other anomalies.</p><p id="para716">The technique involves the injection of an aqueous iodine contrast medium into the urethra through a ureteral or balloon-tipped catheter. The radiopaque contrast material is mixed to a three- to fivefold dilution with sterile lubricating jelly to increase the viscosity. A dilution of 1:3 contrast medium with sterile distilled water or saline also can be used. Before retrograde contrast urethography is performed, give the animal a cleansing enema. Sedation or anesthesia may be necessary. Inject 5 to 10 mL of contrast medium. Near the end of the injection, while the urethra is still under pressure, obtain a lateral radiograph.</p><p id="para717">If the urinary bladder is to be distended with contrast material or air, remove urine from the bladder. In the male dog, position the catheter so that the tip of the catheter is distal to the os penis. Inject lidocaine 1 to 2 mL into the urethral lumen to anesthetize the urethra adjacent to the balloon-tipped catheter. Take extreme care in the amount of fluid placed in the bladder if the urethra is occluded by a balloon catheter. Overdistention of the bladder results in hematuria, pyuria, urinary bladder rupture, and mild to severe bladder inflammation. Palpate the bladder carefully during distention, and note the backpressure on the syringe used in filling the bladder.</p><p id="para718">Retrograde contrast urethrography is a definite aid in defining the extent of urethral damage (stricture) or in demonstrating urethral calculi in male cats. In male cats, use a 4F balloon catheter or a 3.5F Tomcat open-ended urethral catheter. Insert the catheter 1.5 cm into the penile urethra. If the urethra is patent, 2 to 3 mL of contrast material will enable visualization of the urethra, but increased amounts of contrast material (2 to 3 mL/lb) injected into the bladder are needed for maximum distention of the preprostatic urethra. A voiding positive contrast urethrogram is necessary to visualize the distal (penile) urethra. Apply external pressure to the bladder (using a wooden spoon or other external compression device), and radiograph the distal urethra.</p></sec><sec id="cesec240"><title>Cystography</title><p id="para719">Cystography refers to contrast radiographic procedures that facilitate visualization of the lumen and/or contents of the urinary bladder and trigone (<xref rid="cetextbox17" ref-type="boxed-text">Box 4-17</xref>). Three procedures can be used to image the urinary bladder: positive contrast cystography, negative contrast cystography (also called pneumocystography), and double contrast cystography (combination of positive and negative cystography performed in the same patient). NOTE: Many of the indications for performing contrast cystography are also indications for ultrasound examination. Contrast cystography is most useful for characterizing congenital and acquired alternations in the normal anatomy and function of the ureters and lower urinary tract, such as ectopic ureter. Abdominal ultrasound, when available, remains the preferred method for imaging abnormalities within the bladder lumen (e.g., calculi and tumors) and changes within the bladder wall.<boxed-text id="cetextbox17"><label>BOX 4-17</label><caption><title>INDICATIONS FOR CYSTOGRAPHY</title></caption><p id="para120"><list list-type="simple" id="celist12"><list-item id="celistitem58"><label>•</label><p id="para121">Incontinence unresponsive to medical treatment, especially in young dogs</p></list-item><list-item id="celistitem59"><label>•</label><p id="para122">Persistent hematuria (WARNING: <italic>Pneumocystography is contraindicated.</italic>)</p></list-item><list-item id="celistitem60"><label>•</label><p id="para123">Stranguria</p></list-item><list-item id="celistitem61"><label>•</label><p id="para124">Pyuria</p></list-item><list-item id="celistitem62"><label>•</label><p id="para125">Persistent crystalluria</p></list-item><list-item id="celistitem63"><label>•</label><p id="para126">Significant proteinuria</p></list-item><list-item id="celistitem64"><label>•</label><p id="para127">Dysuria</p></list-item><list-item id="celistitem65"><label>•</label><p id="para128">Persistent or recurrent urinary tract infection</p></list-item></list></p></boxed-text></p><sec id="cesec241"><title>Pneumocystography</title><p id="para720">Pneumocystography, also called negative-contrast cystography, involves the insufflation of a soluble gas into the lumen of the urinary bladder to facilitate imaging of any material or tissue within the bladder lumen that otherwise would be obscured by the presence of urine or positive contrast material. Prepare the patient as described previously. Once a urinary catheter is placed and the urethra is occluded, use a syringe and a three-way valve to inject 4 to 10 mL/kg of carbon dioxide or nitrous oxide. Palpate the bladder while filling it with gas to avoid overdistention or rupture. Inject air until there is pressure on the syringe barrel or leakage of air around the catheter. Replace any air that escapes during the procedure. Take lateral and ventrodorsal views of the abdomen.</p><p id="para721"><boxed-text id="cetextbox26"><caption><title>Note:</title></caption><p id="para722">Room air is the most accessible contrast material and generally can be found in most practices. <italic>However,</italic> an increased risk of air emboli is associated with the placement of room air into the bladder under positive pressure.</p></boxed-text></p><p id="para723">Pneumocystography is not an innocuous procedure; fatal venous air emboli have occurred in dogs and cats. This complication is seen most commonly in cases of severe hematuria. Ultrasound or positive contrast cystography is preferred over pneumocystography in such cases if a soluble gas is not available. If possible, use a gas that is readily soluble in blood (such as carbon dioxide or nitrous oxide) for bladder insufflation.</p></sec><sec id="cesec242"><title>Positive contrast cystography</title><p id="para724">The injection of radiographic contrast material into the urinary bladder is referred to as contrast cystography or positive contrast cystography. When ultrasound examination is not available or not feasible, the clinical and radiographic findings noted in <xref rid="cetextbox18" ref-type="boxed-text">Box 4-18</xref>justify the use of a contrast radiography to image the bladder.<boxed-text id="cetextbox18"><label>BOX 4-18</label><caption><title>INDICATIONS FOR PERFORMING CONTRAST CYSTOGRAPHY</title></caption><p id="para129"><list list-type="simple" id="celist13"><list-item id="celistitem66"><label>•</label><p id="para130">Frequent urination</p></list-item><list-item id="celistitem67"><label>•</label><p id="para131">Intermittent or chronic hematuria or small volumes of voided urine</p></list-item><list-item id="celistitem68"><label>•</label><p id="para132">Hematuria that is seen throughout or in the later stages of voiding</p></list-item><list-item id="celistitem69"><label>•</label><p id="para133">Dysuria</p></list-item><list-item id="celistitem70"><label>•</label><p id="para134">Persistent posttraumatic hematuria</p></list-item><list-item id="celistitem71"><label>•</label><p id="para135">Areas of increased or decreased density associated with the urinary bladder</p></list-item><list-item id="celistitem72"><label>•</label><p id="para136">Nonvisualization of the urinary bladder after trauma</p></list-item><list-item id="celistitem73"><label>•</label><p id="para137">Evaluation of abnormal caudal abdominal masses and structures adjacent to the urinary bladder</p></list-item><list-item id="celistitem74"><label>•</label><p id="para138">Evaluation of abnormal bladder shape or location</p></list-item></list></p></boxed-text></p><p id="para725">The same principles of preparation apply as for performing a pneumocystogram. Use a urethral catheter with a three-way valve or a small Foley catheter with an inflatable cuff. Organic iodides are the contrast material of choice and should be used in 5% to 10% concentrations.</p></sec><sec id="cesec243"><title>Double contrast cystography</title><p id="para726">Double contrast cystography also can be performed in patients for which a positive contrast study is not diagnostic, yet there is reasonable indication for an intraluminal lesion. In this case, using the same urinary catheter as used for the contrast study, remove all remaining urine and contrast material. If necessary, inject 2 to 5 mL of an aqueous organic iodine contrast material into the bladder. Gently roll the patient over in an attempt to coat the bladder with contrast material. Then distend the bladder with air in the same manner as described for pneumocystography.</p><p id="para727">Some of the routine lesions diagnosed with the aid of cystography are calculi (<xref rid="cetable11" ref-type="table">Table 4-11</xref>); neoplasia; cystitis, if proliferative changes are present; muscle hypertrophy; bladder diverticula; duplications; adhesions, especially uterine stump infection; persistent urachus; ruptures; and atonic bladder.<table-wrap position="float" id="cetable11"><label>TABLE 4-11</label><caption><p>Radiopacity of Cystic Calculi on Plain Addominal Radiographs</p></caption><table frame="hsides" rules="groups"><thead><tr><th align="left">Calculus composition</th><th align="left">Density</th></tr></thead><tbody><tr><td align="left">Calcium oxalate</td><td align="left">Radiopaque</td></tr><tr><td align="left">Calcium carbonate</td><td align="left">Radiopaque</td></tr><tr><td align="left">Triple phosphate</td><td align="left">Radiopaque—small calculi may be nonradiopaque</td></tr><tr><td align="left">Cystine</td><td align="left">Variable density—may have radiopaque stippling</td></tr><tr><td align="left">Uric acid and urates</td><td align="left">Nonradiopaque</td></tr><tr><td align="left">Xanthine</td><td align="left">Nonradiopaque</td></tr><tr><td align="left">Matrix concretions</td><td align="left">Nonradiopaque</td></tr></tbody></table><attrib>From Park RD: Radiology of the urinary bladder and urethra. In O’Brien TR, ed: <italic>Radiographic diagnosis of abdominal disorders in the dog and cat</italic>. Philadelphia, WB Saunders, 1978.</attrib></table-wrap></p><sec id="cesec244"><sec id="cesec245"><title>Additional Reading</title><p id="para728">Burk RL, Ackerman N: <italic>Small animal radiology and ultrasonography: a diagnostic atlas and text,</italic> ed 3, St Louis, 2003, Saunders.</p><p id="para729">Osborne CA, Finco DR: <italic>Canine and feline nephrology and urology,</italic> Baltimore, 1995, Williams & Wilkins.</p><p id="para730">Thrall DE: <italic>Textbook of veterinary diagnostic radiology,</italic> ed 4, St Louis, 2002, Saunders.</p></sec></sec></sec></sec></sec><sec id="cesec246"><sec id="cesec247"><title>Myelography</title><p id="para731">Myelography is the study of the spinal cord and vertebral canal made possible by the use of contrast media in the subarachnoid space. Ideally, the contrast material should be relatively nontoxic and absorbable, should provide good contrast, and should be distributed evenly throughout the subarachnoid space. Indications for myelography are progressive neurologic disease in which survey radiographs have failed to reveal substantive findings.</p><p id="para732">The nonionic water-soluble agents currently are preferred for myelography. The agents are iopamidol, iohexol, and ioxaglate. These agents are stable in solution and much more convenient than metrizamide and have low toxicity and low epileptogenic activity, are inert to nervous tissue, have no long-term side effects, and are resorbed and excreted rapidly from the CSF. These agents can be injected into the subarachnoid space at the cerebellomedullary cistern or at the caudal lumbar spine (preferred) at a dose level of 0.22 mL/kg. Five minutes after cisternal injection, if there is no obstruction, the cervical and thoracic cord segments are outlined, and after 10 to 15 minutes the entire cord is outlined.</p><p id="para733">Patients undergoing myelography can be pretreated with diazepam at a dose of 0.25 mL/kg, not to exceed 10 mL in larger patients. Diazepam has a short biologic half-life of 30 to 45 minutes and may be better used after myelography if seizures do occur.</p><p id="para734">Myelography is a relatively high-risk procedure and should be performed only by individuals with additional training and experience. The following steps are involved:<list list-type="simple" id="celist27"><list-item id="celistitem152"><label>1.</label><p id="para735">Have the patient fast for 18 to 24 hours preceding myelography if the procedure is elective.</p></list-item><list-item id="celistitem153"><label>2.</label><p id="para736">Anesthetize the patient with a short-acting anesthetic agent, and maintain the animal on the gas anesthetic of choice. Maintain an intravenous catheter and good hydration through fluid support.</p></list-item><list-item id="celistitem154"><label>3.</label><p id="para737">Clip and surgically prepare the skin over the cisterna magna or in the lumbosacral area, depending on where one wishes to enter the spinal canal. When fluoroscopic visualization is available, myelograms can be done for all animals from a lumbar subarachnoid tap. Lumbar puncture can be made between L1 and L6 or between L4 and L5. Use a short-bevel spinal needle (1.5-inch, 22-gauge needle for dogs under 20 lb, and a 2.5-inch, 22-gauge needle for larger dogs). The average dose for any of the four agents is 0.22 mL/kg. More is needed per kilogram for small dogs and less per kilogram for large dogs (for cisterna magna puncture).</p></list-item><list-item id="celistitem155"><label>4.</label><p id="para738">Inject the contrast medium through a flexible extension tube attached to the needle. The recommended dose of iohexol 300 mg/mL is for cervical myelogram (lumbar tap), 0.45 mL/kg; for cervical tap, 0.30 mL/kg; for thoracolumbar myelogram (lumbar tap), 0.30 mL/kg; for thoracolumbar myelogram (cervical tap), 0.30 mL/kg; and for cervicothoracolumbar myelogram (cervical or lumbar), 0.45 mL/kg.</p></list-item></list></p><sec id="cesec248"><sec id="cesec249"><sec id="cesec250"><title>Additional Reading</title><p id="para739">Burk RL, Ackerman N: <italic>Small animal radiology and ultrasonography: a diagnostic atlas and text,</italic> ed 3, St Louis, 2003, Saunders.</p><p id="para740">Thrall DE: <italic>Textbook of veterinary diagnostic radiology,</italic> ed 4, St Louis, 2002, Saunders.</p></sec></sec></sec></sec></sec><sec id="cesec251"><sec id="cesec252"><title>Cholecystography</title><p id="para741">Cholecystography refers to the use of contrast material to facilitate imaging of the gall bladder and the common bile duct. Use of this technique depends on the ability of selected radiopaque compounds to be removed from the blood and excreted via active transport by hepatocytes. Contrast agents for cholecystography can be administered either orally or intravenously. Oral cholecystographic examination requires that the contrast agent (1) enter the small bowel, (2) be absorbed and enter the portal circulation, and (3) be excreted into the bile and concentrated in the gallbladder. Intravenous cholecystography eliminates some of the variables within the digestive tract and may be a more reliable technique in dogs and cats. For that reason, and others, cholecystography is <italic>not</italic> a particularly popular procedure. The reality is that abdominal ultrasound, in the hands of an experienced ultrasonographer, will provide more reliable diagnostic information on the liver and biliary tract than will contrast radiography.</p></sec></sec></sec><sec id="cesec253"><sec id="cesec254"><title>REPRODUCTIVE TRACT: FEMALE</title><sec id="cesec255"><title>Culture Of the Vagina</title><p id="para742">Vaginal examination is indicated for collection of material from the mucosal wall for culture and exfoliative cytologic examination and for vaginoscopic examination of vaginal and cervical mucosa (<xref rid="cetextbox19" ref-type="boxed-text">Box 4-19</xref>).<boxed-text id="cetextbox19"><label>BOX 4-19</label><caption><title>EQUIPMENT TO USE FOR EXAMINATION OF THE CANINE VAGINA</title></caption><p id="para139">Sterile vaginal speculum (e.g., adjustable-spreading, stainless steel, or disposable plastic; cylindric; glass, plastic, stainless steel, or nylon)</p><p id="para140">Sterile otoscope heads of variable size for small dogs</p><p id="para141">Sterile protected culture swabs (Tiegland type or other)</p><p id="para142">Sterile culture swabs (Culturettes)</p><p id="para143">Amies transport medium with charcoal</p><p id="para144">Viral transport media</p><p id="para145">Glass slides and coverslips</p><p id="para146">Sterile proctoscope (Welch Allyn, human pediatric type) or other endoscope, flexible or rigid</p><p id="para147">Sterile offset biopsy punch</p></boxed-text></p><p id="para743">Examination of the vagina for culture and cytologic or vaginoscopic examination occasionally can be performed in the cooperative patient without the use of sedation or anesthesia. An assistant is used to restrain the patient on an examination table. Bitches that can be restrained for other minor examinations (ears, teeth, toenails, anal sacs, and blood samples) often will tolerate vaginal examinations. Those that need further restraint may require sedation or administration of a short-acting barbiturate anesthetic.</p><p id="para744">If the vulva appears clean, no preparation before examination is needed. If the vulva is soiled or vulvar hair is matted or soiled, trim the hair and wash the area with a germicidal or surgical scrub such as povidone-iodine, or give a general grooming and bath a few hours before examination. Water and germicidal soap usually will not control surface contamination by <italic>Pseudomonas</italic> and <italic>Proteus</italic> spp., which frequently contaminate culture swabs. Bitches with long hair should have the leg hair pinned to one side with clips and the tail bandaged before examination.</p><p id="para745">Obtain a deep vaginal culture FIRST to avoid contamination during the general examination. Pass a sterile, warm vaginal speculum with only a thin coating of lubricating gel into the posterior vagina while an assistant spreads the vulva. Guide the speculum into the vagina by placing the speculum into the vulva just at the dorsal commissure of the vulva and applying pressure up and out against the commissure. Direct the speculum <italic>dorsally</italic> toward the rectum until meeting resistance, and then direct it horizontally into the cranial vagina. This procedure bypasses the clitoral fossa and enables visualization of the urethral opening and pelvic arch.</p><p id="para746">Take a guarded culture swab (swab covered by a protective plastic pipette) from its individual sterile bag and pass it inside the vaginal speculum to the anterior vagina or cervical area. Then expose the swab from the protective plastic tubing and rotate it against the mucosa. Retract the swab into the protective plastic tubing and carefully remove it from the vagina. The protected swab then may be placed back in its original sterile bag until it is processed for culture (30 minutes) or placed in Amies transport medium with charcoal. Amies transport medium with refrigerator packs and a styrofoam-insulated mailing box will retain fastidious organisms for 72 to 96 hours. Process bacterial, <italic>Mycoplasma,</italic> and <italic>Ureaplasma</italic> cultures for potential infectious agents. Viral transport medium can be used for a separate sterile swab if viral agents such as the genital form of canine herpesvirus are suspected.</p><p id="para747">Immediately after the swabbing for culture, while the vaginal speculum is still in place, advance a clean or sterile swab moistened with sterile physiologic saline solution carefully into the anterior vagina to make a smear for cytologic examination. Gently scrape vaginal epithelial cells from the ceiling of the vagina at or cranial to the region of the external urethral orifice. Collect samples from the region of the clitoral fossa, which is lined by stratified squamous epithelium at all stages of the estrous cycle. Gently rub the swab on the vaginal mucosa. Remove the swab and roll it smoothly onto two or three clean glass slides. The smears may be fixed immediately in 95% alcohol, sprayed with a commercial fixative or hair spray, or left to dry in air.</p><p id="para748">A drop of new methylene blue stain placed on a coverslip and inverted on the smear can be used to examine a wet mount preparation immediately. This stain is not permanent and precipitates when it dries, and new methylene blue–stained smears cannot be used for comparison with other smears made later in the cycle. A better and quick method that provides a permanent record is the use of the Diff-Quik or Leukostat stain. Giemsa stain, toluidine blue, Wright's stain, Shorr's stain, or phase-contrast microscopy also can be used. Examine the smear for stage of estrous cycle and evidence of active inflammation. Compare these findings with culture results and vaginoscopic findings to interpret evidence for an active genital tract infection, a carrier state of a potential infectious agent, or a possible contaminant at culture. A diagnostic laboratory with the ability to isolate specific infectious agents should indicate the number of organisms (few, moderate, many, or heavy) and report whether the isolates are pure or mixed and their significance.</p></sec><sec id="cesec256"><title>Examination of the Vagina</title><p id="para749">The vagina of the bitch is long in comparison to that of other domestic animals, hence digital examination of the cervix, and in many cases, the urethral orifice, is simply not possible. The mucosa forms longitudinal folds. The clitoris is in a well-developed fossa in the floor of the vestibule. The vagina can be visualized completely with a small, sterile proctoscope or flexible endoscope. Lubricate the warmed, sterile instrument, and pass it to the region of the cervix. Examine first without insufflation for true color and vaginal fluids or discharge. When insufflation is performed while the vulva is compressed around the sterile proctoscope, the vagina expands and its entire wall can be viewed completely as the instrument is withdrawn.</p><p id="para750">The normal canine vagina has a uniform light pink color and longitudinal folds. During proestrus and estrus, the folds become more prominent and cross-striations give the surface a cobblestone appearance. This cobblestone appearance remains smooth when estrogen levels are high but quickly becomes angular (worn cobblestone appearance) when estrogen levels drop during the luteinizing hormone peak (ovulation), and progesterone levels increase. This change can be used to indicate ovulation and the ideal time for breeding. The hyperemia causes the vagina to appear reddish and congested. The pressure of air insufflation balances the mucosa. The canine vulva has a large cranial dorsal median fold that may obscure the cervix. In fact, ridges near the dorsal fold may give a false impression that this fold is the cervix. During estrogen stimulation, the cervix may be open and uterine blood may be escaping. In the management of dystocia, the vaginoscope can be used to detect puppies in the birth canal and to diagnose malpositions and aid in the correction of these conditions.</p><p id="para751">During the endoscopic examination, small tumors or polyps can be removed or large masses can be sampled with the biopsy punch. Ulcers or erosions can be cauterized, and foreign bodies can be removed.</p><p id="para752">A complete vaginal examination must include careful palpation of the vaginal wall and pelvic canal. This palpation is accomplished by digital examination through the vulva (using a sterile glove) and is assisted by palpation through the posterior abdominal wall. Incomplete hymen rings, vaginal fibrous stenotic rings, or pelvic malformation can be diagnosed. A digital rectal examination may be needed for vaginal masses or pelvic deformities.</p></sec><sec id="cesec257"><title>Estrous Cycle: Staging and Cytologic Findings—Canine</title><p id="para753">The canine reproductive cycle begins at the age of 6 to 12 months and repeats at intervals of 4 to 12 months. In the average bitch, ovulation occurs spontaneously 1 to 3 days after the onset of estrus; in normal bitches ovulation may occur between 3 days before and 11 days after the onset of estrus. Sperm live in the uterus of the estrous bitch up to 11 days, and the ovum lives up to 5 days after ovulation. The fertilized ovum takes 4 to 10 days to reach the uterus, and implantation takes place 18 to 20 days after ovulation. The gestation period from the first breeding is 57 to 72 days and from the luteinizing hormone peak is 64 to 66 days.</p><sec id="cesec258"><title>Anestrus</title><p id="para754">Anestrus is characterized by dryness of the mucosa and a thin vaginal wall with stratified squamous epithelial cells a few cells to several layers thick but without cornification. Noncornified epithelial cells and WBCs are present in a ratio of 1:5 in the vaginal smear. The WBCs are polymorphonuclear. The noncornified epithelial cells are 15 to 51 nm in diameter and have round free edges, granular cytoplasm, and large nuclei with distinct chromatin granules. The period of anestrus is 2 to 3 months or longer in some breeds.</p></sec><sec id="cesec259"><title>Proestrus</title><p id="para755">In proestrus the vaginal wall is thicker than in anestrus, and the mucosa shows prominent cornified squamous epithelium (20 to 30 cells thick) with rete pegs. The longitudinal and transverse vaginal folds are thick, smooth, and round. The vaginal wall becomes impervious to WBCs, but there is extravasation of red blood cells to the surface epithelium. The red blood cells are discharged. Vaginal smears show predominantly red blood cells and noncornified epithelial cells, which become cornified as proestrus progresses. White blood cells are present, but their numbers decrease as estrus approaches. Debris and bacteria are abundant for 7 to 10 days.</p></sec><sec id="cesec260"><title>Estrus</title><p id="para756">The vagina is thick with longitudinal and transverse folds that become angular as estrogen levels decrease and progesterone levels increase. Fluid is abundant, often tinged with blood. Noncornified epithelial cells and WBCs are absent. Cornified epithelial cells, which are polyhedral and contain pyknotic nuclei or no nucleus, are predominant; their presence seems to be related to the appearance of flirting by the bitch and acceptance of the stud. White blood cells reappear about 36 to 96 hours after ovulation. Bacteria and debris are absent during estrus, but they are seen again in the smears after ovulation when WBCs reappear 7 to 10 days later.</p></sec><sec id="cesec261"><title>Diestrus</title><p id="para757">The number of WBCs increases rapidly, the number of cornified epithelial cells decreases, and the number of noncornified epithelial cells increases. After 5 to 7 days, the number of WBCs may decrease to 10 to 30 per field.</p><p id="para758">Following parturition, much cellular debris, WBCs, red blood cells, and a few epithelial cells are present for several days, until placental sloughing is complete. The presence of masses of degenerate WBCs (and bacteria) indicates metritis or endometritis. The continued presence of blood-tinged fluids containing abundant red blood cells, a few noncornified epithelial cells, and occasional WBCs (nontoxic) plus necrotic cells for months postpartum is evidence of subinvolution of placental sites.</p></sec></sec><sec id="cesec262"><title>Estrous Cycle: Staging and Cytologic Findings—Feline</title><p id="para759">Most of the characteristics just discussed that apply to bitches also pertain to queens. However, the small size of the feline vagina precludes palpation and early vaginoscopy. A sterile, warm, small-animal otoscope speculum enables fairly good visualization of the vaginal mucosa and can be used with a small, 4-mm-diameter sterile swab to obtain smears for culture procedures. Use of the speculum is easiest following parturition or during estrus.</p><p id="para760">Vaginal cells for cytologic examination can be obtained with a moistened 3-mm cotton swab (Calgiswab) inserted 2 cm into the vagina. In some cases, flushing the vagina with sterile saline injected and aspirated with a clean glass eyedropper is more successful. Use of an eyedropper may trigger ovulation, as it simulates coitus.</p><p id="para761">Unlike the bitch, the queen shows no diapedesis of red blood cells during proestrus or throughout the estrous cycle. Cytologic examination of feline vaginal smears reveals the following by stage of the estrous cycle.</p><sec id="cesec263"><title>Anestrus or prepuberty</title><p id="para762">Cytologic examination reveals scarce debris and numerous small, round epithelial cells with a high nuclear/cytoplasmic ratio, frequently in groups (seasonal: from September to January in the Northern Hemisphere).</p></sec><sec id="cesec264"><title>Proestrus</title><p id="para763">Cytologic examination reveals increased debris and fewer but larger nucleated epithelial cells with a low nuclear/cytoplasmic ratio (0 to 2 days).</p></sec><sec id="cesec265"><title>Estrus</title><p id="para764">Cytologic examination reveals markedly less debris and numerous large polyhedral cornified cells with curled edges and small dark pyknotic nuclei or loss of nuclei (6 to 8 days) following coitus or induced ovulation.</p></sec><sec id="cesec266"><title>Early diestrus</title><p id="para765">Cytologic examination reveals hazy, ragged-edged cornified cells and zero to numerous WBCs with numerous bacteria and increased debris.</p></sec><sec id="cesec267"><title>Late diestrus</title><p id="para766">Cytologic examination reveals increasing numbers of small basophilic cells with WBCs still present (total period of metestrus, 7 to 21 days). If ovulation does not occur, the smear will return to an anestrous stage with few to no WBCs.</p><p id="para767">The feline estrous cycle is continuous every 14 to 36 days if 12 to 14 hours of light is present daily. Ovulation is induced 24 to 30 hours after coitus. Sperm require 2 to 24 hours for capacitation in the uterus. Implantation is expected 13 to 14 days after coitus.</p><sec id="cesec268"><sec id="cesec269"><title>Additional Reading</title><p id="para768">Baker R, Lumsden JH: The reproductive tract: vagina, uterus, protate, and testicle. In Baker R, Lummsden JH, editors: <italic>Color atlas of the cytology of the dog and cat,</italic> St. Louis, 2000, Mosby. (NOTE: Textbook contains exceptional color plates of normal and abnormal reproductive tract cytologic findings of the dog and cat.)</p><p id="para769">Feldman EC, Nelson RW: <italic>Canine and feline endocrinology and reproduction,</italic> ed 3, St Louis, 2004, Elsevier-Saunders.</p><p id="para770">Grundy SA, Davidson AP: Feline reproduction. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p><p id="para771">Schaefers-Okkens AC: Estrous cycle and breeding management of the healthy bitch. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p></sec></sec></sec></sec></sec><sec id="cesec270"><sec id="cesec271"><title>Artificial Insemination: Canine</title><p id="para772">The procedure for artificial insemination in dogs includes the following steps:<list list-type="simple" id="celist28"><list-item id="celistitem156"><label>1.</label><p id="para773">Determine the correct time to inseminate by test-teasing with a stud, by cytologic examination of vaginal smears, or by vaginoscopic examination to determine the day when vaginal folds change from round to angular. Breed the day after the bitch first stands staunchly to accept service and “flags” her tail or during cytologic indications of estrus (complete cornification of vaginal epithelial cells) but before WBCs reappear in the smears. Breed at 48-hour intervals until the female dog goes out of heat or for three or four inseminations.</p></list-item><list-item id="celistitem157"><label>2.</label><p id="para774">If the vulva is soiled, clean it thoroughly with alcohol swabs (<xref rid="cetextbox20" ref-type="boxed-text">Box 4-20</xref>).<boxed-text id="cetextbox20"><label>BOX 4-20</label><caption><title>MATERIALS USED FOR PERFORMING ARTIFICIAL INSEMINATION</title></caption><p id="para148"><list list-type="simple" id="celist14"><list-item id="celistitem75"><label>•</label><p id="para149">Dry, warm, sterile 5- or 10-mL syringes</p></list-item><list-item id="celistitem76"><label>•</label><p id="para150">Rubber adapter tubing, inch long</p></list-item><list-item id="celistitem77"><label>•</label><p id="para151">A 6- to 9-inch plastic or polypropylene inseminating pipette</p></list-item><list-item id="celistitem78"><label>•</label><p id="para152">A sterile examination glove</p></list-item><list-item id="celistitem79"><label>•</label><p id="para153">Alcohol</p></list-item><list-item id="celistitem80"><label>•</label><p id="para154">Cotton</p></list-item></list></p><p id="para155">Do not use lubricating materials.</p></boxed-text></p></list-item><list-item id="celistitem158"><label>3.</label><p id="para775">Gently aspirate semen through the inseminating pipette into the warm syringe.</p></list-item><list-item id="celistitem159"><label>4.</label><p id="para776">Using a gloved left index finger (not lubricated) as a guide, insert the pipette through the vulva and dorsally into the vagina and forward to the cervix. Elevate the bitch's rear quarters to a 45-degree angle by having an assistant pick up the bitch by holding the hock region so that no pressure is applied to the ventral abdomen and uterus. Eject the semen gently and slowly. Eject a bubble of air to push all the semen through the pipette. Deposit the semen in the anterior vagina.</p></list-item><list-item id="celistitem160"><label>5.</label><p id="para777">Remove the pipette, and hold the bitch in an elevated position for 5 minutes. During this time, use the finger encased in a sterile glove to “feather” the ceiling of the vagina to stimulate constrictor activity. This may be important to simulate a “tie” and transport semen into the uterus.</p></list-item><list-item id="celistitem161"><label>6.</label><p id="para778">Lower the bitch to the normal position, and immediately walk her for 5 minutes so that she does not sit down or jump up on a person and allow semen to run back out of the vagina.</p></list-item><list-item id="celistitem162"><label>7.</label><p id="para779">For best conception, inseminate undiluted fresh semen immediately.</p></list-item><list-item id="celistitem163"><label>8.</label><p id="para780">Refrigerated extended semen is best used within 24 to 48 hours if possible. However, refrigerated semen has been kept viable for up to 9 days with proper care.</p></list-item></list></p><p id="para781">Skim milk has been used as an economical and adequate extender. Heat milk to 92° to 94° C for 10 minutes, cool it, and skim it at room temperature. To each milliliter, add 1000 units of crystalline penicillin. If <italic>Pseudomonas</italic> spp. affect the semen, polymyxin B may be added at 200 units/mL of extender. Dilute semen with extender at a semen/extender ratio of 1:1 to 1:4. Extend canine semen for freezing with a diluent containing 11% lactose, 4% glycerin, and 20% egg yolk. Refrigerate the 1:4 diluted semen; then pipette 0.05-mL portions into depressions in a block of dry ice and hold them for 8 minutes to freeze. Store the frozen pellets in liquid nitrogen. Frozen semen can be thawed in buffered saline at 30° to 37° C. Good semen may be stored in liquid nitrogen for many years without significant loss of motility. Conception is best when large numbers of thawed motile sperm are deposited in the cervix or uterine cavity. Conception is poor when thawed semen is placed in the anterior vagina, as done in artificial breeding with raw semen.</p><sec id="cesec272"><sec id="cesec273"><sec id="cesec274"><title>Additional Reading</title><p id="para782">Memon MA, Sirinarumitr K: Semen evaluation, canine male infertility, and common disorders of the male. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p></sec></sec></sec></sec></sec></sec><sec id="cesec275"><sec id="cesec276"><title>REPRODUCTIVE TRACT: MALE</title><sec id="cesec277"><title>Semen Collection: Canine</title><p id="para783">Semen is collected for examination for breeding soundness, for investigation of infertility or prostatic disease, and for artificial insemination (<xref rid="cetextbox21" ref-type="boxed-text">Box 4-21</xref>).<boxed-text id="cetextbox21"><label>BOX 4-21</label><caption><title>EQUIPMENT USED TO COLLECT SEMEN FROM A MALE DOG</title></caption><sec id="cesec3"><title>Sterile</title><p id="para156">Sterile rubber cone (artificial vagina) connected to a semen collection tube</p><p id="para157">Glass, polytef (Teflon), or plastic test tubes</p><p id="para158">Saline solution, 0.9%</p><p id="para159">Sterile aqueous lubricant</p></sec><sec id="cesec4"><title>Nonsterile</title><p id="para160">Microscope slides and coverslips (warmed)</p><p id="para161">A quick Romanowsky's stain, buffered formalin</p><p id="para162">Hemocytometer-counting chamber and 1:100 white blood cell dilutor pipette or Unopette</p><p id="para163">Microscope with oil immersion objective (×1000) and light</p><p id="para164">Muzzle gauze</p></sec></boxed-text></p><p id="para784">The following steps outline the procedure for collecting semen from a male dog:<list list-type="simple" id="celist29"><list-item id="celistitem164"><label>1.</label><p id="para785">Take the stud and an estrous teaser bitch (if available) to a quiet room where there will be no distractions and where there is good traction (rubber mats or rug) for mounting by the stud.</p></list-item><list-item id="celistitem165"><label>2.</label><p id="para786">Hold the bitch, and allow the stud to “flirt” (become aroused) for several minutes. If the bitch is in heat, a brief period of foreplay (I am not really certain if “foreplay” is an appropriate word to use when describing the mating behavior of dogs, but you get the point) with both dogs unrestricted will help the process.</p></list-item><list-item id="celistitem166"><label>3.</label><p id="para787">If necessary, have assistants restrain the muzzled bitch and control the stud by a collar and leash. Bring the stud up to the rear end of the bitch, and allow him to mount her or keep his nose in the region of her perineal area.</p></list-item><list-item id="celistitem167"><label>4.</label><p id="para788">Attach the artificial vagina to the semen collection tube, and apply a scant amount of lubricant to the opening of the artificial vagina.</p></list-item><list-item id="celistitem168"><label>5.</label><p id="para789">If mounting occurs, allow the stud to grasp the bitch and start to thrust his pelvis in an attempt to copulate. Gently, from the side of the sheath, grasp the penis by the prepuce and move the prepuce back over the engorged bulbus glandis; while applying the artificial vagina to the shaft of the penis, apply pressure with the thumb and forefinger proximal to the exposed glandis. This usually can be done with one motion as the stud is thrusting. If the stud is shy and not interested, massage the penis slightly in the prepuce or in the artificial vagina to cause erection. When erection of the bulbus is felt, reflect the prepuce posteriorly to free the bulbus. Apply pressure with the thumb and forefinger behind the bulbus, circling the shaft of the penis. After completion of the most rapid pelvic thrusting and ejaculation of the sperm-rich fractions of semen (1 to 3 mL), twist the penis 180 degrees backward in a horizontal plane, between the hind legs, so that the penis remains in the same plane as in the forward position with the thumb and forefinger still applying pressure around the circumference of the penis proximal to the bulbus. The penis cannot be twisted unless the prepuce is reflected posterior or proximal to the bulbus glandis. Twisting the penis in this position simulates a natural “tie” and allows the person collecting the semen to better visualize the collection (artificial vaginas are widely available now and are much preferred because they simulate the natural pressure of the vagina). The first drops of ejaculate may be discarded, especially if any urine is present. Collect the sperm-rich fraction separately. A clear ejaculate is prostatic fluid, which may be collected separately for examination.</p></list-item><list-item id="celistitem169"><label>6.</label><p id="para790">After semen collection, place the penis in the forward position, straighten out the prepuce to avoid paraphimosis, and remove the bitch from the room. Allow the stud to lick the erect penis and lose the erection. Check the stud for evidence of paraphimosis before he is released or caged. The ejaculate consists of three fractions:</p><p id="para791"><italic>First fraction:</italic> Urethral secretion (usually clear fluid)—0.1 to 2 mL within 50 seconds, pH 6.3. If evidence of urine is present, discard this fraction and do not add it to the sperm-rich fraction. In most ejaculates collected from dogs, the first and second semen fractions are collected together.</p><p id="para792"><italic>Second fraction:</italic> Sperm-containing secretion (milky opaque fluid)—0.5 to 3 mL within 1 to 2 minutes, pH 6.1.</p><p id="para793"><italic>Third fraction:</italic> Prostatic secretion (usually clear fluid)—2 to 20 mL within 30 minutes, pH 6.5. The total specimen is 0.3 to 20 mL, pH 6.4. Because the first and third fractions are clear, waterlike material and the second fraction is milky-opaque, the clinician can separate them by changing collecting tubes as each fraction is ejaculated. Collection of only enough prostatic fluid to rinse the sperm fraction into the test tube is best. Too much prostatic fluid may be detrimental to the longevity of sperm in storage. Collecting individual fractions may be important in determining the site of an inflammatory reaction, but for artificial insemination only the sperm-rich, low-volume ejaculate is needed for insemination, dilution, or freezing.</p></list-item><list-item id="celistitem170"><label>7.</label><p id="para794">Return the stud to his cage. Retain the bitch until the semen is examined, if insemination is to be performed.</p></list-item></list></p></sec><sec id="cesec278"><title>Evaluation of Semen</title><p id="para795">Immediately after semen collection, slowly invert the tube several times to mix the semen gently. Determine the <italic>motility of sperm</italic> by placing 1 drop of semen on a warmed microscope slide. Cover the slide with a coverslip, and observe the specimen under low power for progressive motility. There will be no “waves,” but general vigorous forward motion should be evident. If the sample is too concentrated for individual sperm to be found, mix 1 drop of semen with 1 drop of saline at body temperature on a warmed microscope slide. Using high power, count 10 different groups of 10 sperm, observing the numbers of motile and nonmotile sperm. Total motility for a suitable sample should be 80% or greater. Motility less than 60% is not satisfactory.</p><p id="para796">Determine the <italic>number of sperm</italic> in the total ejaculate. Sperm concentration may be determined in a hemocytometer with a 1:100 blood cell dilutor kit (Unopette), and concentration then is multiplied by volume to determine sperm numbers per ejaculate. Remember that more dilute samples will be obtained when prostatic fluid is collected, but total sperm numbers in the ejaculate will be only marginally influenced by dilution with prostatic fluid. Total sperm per ejaculate should exceed 300 million in a normal male dog and may approach 2 billion in large dogs. A minimum number of 200 million sperm per insemination is needed on average for conception.</p><p id="para797">Determine morphology. Make a smear of a drop of semen like a blood smear and allow it to air dry. Then stain the smear with Diff-Quik stain; dip the slide into the fixative and solutions 1 and 2 for 2 to 3 minutes each. Then count 100 sperm at ×1000 magnification, noting normal and abnormal sperm. If there is any question about abnormality, examine 500 sperm cells.</p><p id="para798">Normal canine sperm are 63 nm long; the heads are 7 nm long. The percentage of abnormal sperm should be less than 20%. Differential abnormality is important, and the following abnormalities should not be exceeded in any sperm count: abnormality of the head, 10% to 12%; midpiece abnormalities, 3% to 4%; tail abnormalities, 3% to 4%; and retained protoplasmic droplets, 3% to 4%. <xref rid="f54" ref-type="fig">Figure 4-54</xref>shows abnormalities that should be counted and recorded. The presence and location of distal or proximal protoplasmic droplets, which may indicate cell immaturity, is important to note.<fig id="f54"><label>Figure 4-54</label><caption><p>Chart of abnormal sperm.</p></caption><graphic xlink:href="gr54"></graphic></fig></p><p id="para799">Defects of the cells within the testes are generally more serious than defects that occur in the sperm during epididymal transport or after ejaculation (such as fractured heads, retained protoplasmic droplets, or bent tails). Usually, a biopsy should not be done on material from testes unless the testes are azoospermic. Damage produced after the sperm have left the testes may indicate epididymal disease or may be the result of cold, trauma, or osmotic or urinary contamination. When abnormalities are found, it is wise to obtain two or three semen samples within a few days for baseline evaluation and then repeat the studies in 4 to 6 weeks to determine whether there is a healing or regressing trend. There are usually 64 days from the date of sperm formation to the date of ejaculation: 54 days in the testes and 10 days in transport and maturation in the epididymis.</p><p id="para800">Normal male dogs can be used at stud once every other day indefinitely or once every day for 7 to 9 days, after which sperm numbers in the ejaculate will decline but not to less than the numbers needed to achieve conception.</p></sec><sec id="cesec279"><title>Semen Collection: Feline</title><p id="para801">Semen can be collected by means of electroejaculation in the anesthetized male cat (and, after reviewing the following procedures for use of an artificial vagina, this technique may be preferred). For semen collection with an artificial vagina, the male cat must be trained for several weeks and even then, not all male cats will effectively ejaculate; thus the artificial vagina method generally is not used except in larger catteries. Teaser queens can be produced by injecting spayed females with 0.25 mg of estradiol cyclopentyl propionate, or normal queens in heat can be used.</p><p id="para802">An artificial vagina can be made by cutting off the bulb end of a 2-mL rubber bulb pipette and inserting a 3 × 44-mm test tube into the cut end. Place the apparatus into a 60-mL plastic bottle filled with warm water (52° C). Stretch the rolled end of the rubber pipette over the rim of the bottle. Sparingly lubricate the opening of the pipette (“vagina”) with sterile aqueous lubricant. Place a teaser queen in a quiet cage with the tom. As the tom mounts the queen and develops an erection, place the artificial vagina over the penis. The ejaculation takes 1 to 4 minutes, the semen volume is 0.05 mL (range is 0.03 to 0.3 mL), and the semen contains 50 million to 100 million sperm. Motility is normally 80% to 90%, and pH is 7.4. There should be less than 10% abnormal sperm. The semen can be diluted for insemination to contain 10 million sperm in 0.1 mL of saline; then each 0.1 mL is an adequate insemination volume. With such small samples, microequipment is essential to avoid losing semen by surface absorption. Toms can undergo this collection procedure 3 times weekly and maintain excellent semen quality and libido.</p><p id="para803">Queens can be detected in estrus by stroking their backs and necks daily and noting the arching of the back and the extended and treading action of the rear feet when they are in receptive heat. Estrus is verified by examination of vaginal smears, which show epithelial cell cornification with pyknotic nuclei. Insemination is carried out with a 0.25-mL syringe and a bulb-tipped 9-cm spinal needle. Ovulation is induced by intramuscular injection of 25 μg of gonadotropin-releasing hormone or 250 IU of human chorionic gonadotropin. If insemination is repeated 24 hours later, the conception rate improves from 60% to 75%.</p><sec id="cesec280"><sec id="cesec281"><sec id="cesec282"><title>Additional Reading</title><p id="para804">Baker R, Lumsden JH: The reproductive tract: vagina, uterus, protate, and testicle. In Baker R, Lummsden JH, editors: <italic>Color atlas of the cytology of the dog and cat,</italic> St Louis, 2000, Mosby. (NOTE: This textbook contains exceptional color plates of normal and abnormal reproductive tract cytologic findings of the dog and cat.)</p><p id="para805">Feldman EC, Nelson RW: <italic>Canine and feline endocrinology and reproduction,</italic> ed 3, St Louis, 2004, Elsevier-Saunders.</p><p id="para806">Memon MA, Sirinarumitr K: Semen evaluation, canine male infertility, and common disorders of the male. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p><p id="para807">Wright PJ, Parry BW: Cytology of the canine reproductive system, <italic>Vet Clin North Am Small Anim Pract</italic> 19:851-874, 1989.</p></sec></sec></sec></sec></sec><sec id="cesec283"><sec id="cesec284"><title>Prostatic Wash</title><p id="para808">Although castration is a common first recommendation for any male dog with known or suspected prostatic disease, a number of prostatic disorders are recognized for which cytopathologic and histopathologic examination, rather than castration, is indicated. Benign prostatic hyperplasia is recognized as the most common prostatic disorder of male dogs. In half of the dog population, changes consistent with benign prostatic hyperplasia are present by 4 to 5 years of age, especially older intact dogs. Because benign prostatic hyperplasia is androgen dependent, routine castration is the recommended treatment. However, at least three differential diagnoses justify additional diagnostic tests: prostatic neoplasia (usually adenocarcinoma), acute and chronic bacterial prostatitis, and prostatic cysts (septic and nonseptic).</p><p id="para809">In male dogs presenting with prostatomegaly and associated signs (dysuria and/or dyschezia), further evaluation of the prostate is indicated. Several techniques have been described. Abdominal ultrasonography is the preferred technique for evaluating prostate size, shape, and consistency. Distention retrograde contrast urethrocystogram has been described as a means for evaluating the internal integrity of the prostate and is moderately effective in distinguishing normal from abnormal. However, this technique is not known to distinguish among various types of prostate disease.</p><p id="para810">Cytologic examination and quantitative bacterial culture of the ejaculate (especially the third fraction) of a male dog is recommended in any patient with prostatomegaly. However, sample collection can be difficult and is frequently not successful. In addition to lumbar radiographs and abdominal ultrasonography, performing a prostatic wash is a simple, noninvasive technique that may yield diagnostic information.</p><p id="para811">Using aseptic technique, place a conventional urinary catheter into the bladder and remove all urine. Lavage of the urinary bladder with up to 5 mL of sterile saline is recommended. Recover the saline and save it (sample No. 1). Subsequently, retract the catheter tip, but only to the level of the prostate gland (immediately caudal to the trigone). Position of the tip usually can be verified by tactile placement and the detection of increased resistance to catheter movement during retraction. Position can be confirmed with a lateral radiograph of the pelvis.</p><p id="para812">With the catheter in place, identify the prostate on a digital rectal examination and gently massage for approximately 1 minute to force prostatic fluids into the urethra. Infuse 5 mL of sterile saline through the catheter. The objective is to wash prostatic fluids and cells into the urinary bladder and recover the saline from the bladder (sample No. 2).</p><p id="para813">Examine fluid from both samples cytologically by distributing a drop of fluid across a glass slide, air drying, and staining; submit a small aliquot (0.5 mL) for bacterial culture. Cytologic examination is used to detect the presence of inflammatory cells versus neoplastic cells. Low numbers of neutrophils (<5 cells per high power field) are present in ejaculates and prostatic washes from normal dogs. Quantitative bacterial culture, with a yield of greater than 2 log<sub>10</sub> of one or more bacterial species in sample No. 2 confirms bacterial prostatitis.</p><p id="para814">Complications from this procedure are unlikely, but conceivably a patient with septic prostatitis and prostatic abscesses could become bacteremic following this procedure, which in some patients could lead to sepsis.</p></sec><sec id="cesec285"><title>Prostate Biopsy and Fine-Needle Aspiration</title><p id="para815">Ultrasound examination is an important first step, when available, in assessing the size, shape, and internal integrity of the canine prostate gland and for detecting any changes in structures adjacent to the prostate. However, ultrasonography generally will <italic>not</italic> distinguish between different types of prostatic disease. Further diagnostic tests are especially indicated in castrated, middle-aged to older male dogs with evidence of prostatomegaly. Percutaneous fine-needle aspiration and/or prostatic biopsy are indicated.</p><p id="para816">Fine-needle aspiration of the prostate is performed through a ventral abdominal approach. Use aseptic technique, and surgically prepare the skin at the level of needle insertion. Because needle movement, once the needle inserted, could damage the urethra or adjacent structures, perform the procedure in the sedated or anesthetized patient. Use an approach similar to that used for cystocentesis in a male dog with the exception that needle entry is at a point caudal to that used to enter the urinary bladder but is cranial to the pubis. The procedure can be performed with or without ultrasound guidance. In the absence of ultrasound guidance, determine needle position by tactile placement and detection of resistance as the needle enters the prostate. Multiple needle penetrations and aspirations are attempted without withdrawing the needle from the skin. Relieve negative pressure in the syringe before removing the needle. Apply any material collected to a glass slide and allow it to air dry before staining. Any conventional stain used for peripheral blood is appropriate.</p><p id="para817">A transrectal approach to fine-needle aspiration of the prostate has been used in dogs and is performed routinely in men. However, the distance from the anus to the prostate, visualization, and the risk of infection generally are cited as reasons for not performing this technique in dogs.</p><p id="para818">Fine-needle aspiration may not be diagnostic, particularly in patients with isolated, discrete lesions (cysts or neoplastic nodules) within the prostatic parenchyma. In such cases, ultrasound-guided needle (Tru-Cut) biopsy of the prostate is indicated. Specific training and experience are indicated when performing this procedure because significant complications can result.</p><p id="para819">Complications associated with prostate biopsy and fine-needle aspiration are not insignificant. Hematuria and periprostatic hemorrhage are described. Postaspiration/biopsy abscess also is described. Consider the risk of urethral penetration and subsequent stricture at the site of penetration.</p><sec id="cesec286"><sec id="cesec287"><sec id="cesec288"><title>Additional Reading</title><p id="para820">Kutzler MA, Yeager A: Prostatic diseases. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p></sec></sec></sec></sec></sec></sec><sec id="cesec289"><sec id="cesec290"><title>RESPIRATORY TRACT PROCEDURES</title><sec id="cesec291"><title>Upper Respiratory Tract</title><p id="para821">For purposes of this discussion, the anatomic limits of the upper respiratory tract of the dog and cat extend caudally from the nasal planum to the first tracheal ring. Key anatomic structures that principally can cause clinical signs include the anterior (external) nares, nasal cavity, nasal turbinates, frontal sinuses, maxillary recesses, upper dental arcade (especially the roots of the maxillary canine teeth), the choana (posterior nares), nasopharynx, soft palate, arytenoid cartilages, glottis, larynx, and the vocal folds.</p><p id="para822">Clinical signs related to the upper respiratory tract in dogs and cats are among the most common presenting complaints encountered in small animal practice and interestingly are frequent reasons for referral to specialty practices and veterinary teaching hospitals. The oral and nasal cavities are important portals of entry for foreign body entrapment and infectious agents. In addition to the occurrence of nasal neoplasia and trauma, it is not surprising that upper respiratory tract diseases in dogs and cats are common presentations. However, upper respiratory signs can be associated with significantly different underlying causes. Localizing the problem amid a variety of clinical signs in an anatomically complex area represents significant diagnostic and therapeutic challenges to even the most astute clinician. The presentation addresses upper respiratory disease in the dog, with specific emphasis on clearly defining the presenting clinical signs, localizing the problem, and establishing the diagnosis.</p><sec id="cesec292"><title>Anatomic limits</title><p id="para823">Strictly speaking, the anatomic limits of the upper respiratory tract are not defined. For this presentation, the upper respiratory tract begins at the level of the external nares and ends at the level of the first tracheal ring. In the clinical setting, however, it is practical to establish anatomic limits, or compartments, around the various clinical signs attributable to upper respiratory disease. For example, using the foregoing anatomic limits, the upper respiratory tract can be categorized into three distinct compartments. EACH compartment is associated with a defining clinical sign (<xref rid="cetable12" ref-type="table">Table 4-12</xref>). The importance of this categorization is to facilitate the search for the underlying problem and to establish a diagnosis quickly. Properly characterizing the clinical signs is the first step in establishing the cause and defining the diagnosis of upper respiratory tract disease.<table-wrap position="float" id="cetable12"><label>TABLE 4-12</label><caption><p>Anatomic Limits of the Upper Respiratory Tract and Defining Clinical Signs</p></caption><table frame="hsides" rules="groups"><thead><tr><th align="left">Compartment</th><th align="left">Anatomic limits</th><th align="left">Defining clinical sign(s)</th></tr></thead><tbody><tr><td align="left">I</td><td align="left">Nose, nasal cavity, and paranasal sinuses</td><td align="left">Sneezing and/or nasal discharge</td></tr><tr><td align="left">II</td><td align="left">Nasopharynx, posterior nares (choana), and soft palate</td><td align="left">Stertor (snort) and reverse sneeze</td></tr><tr><td align="left">III</td><td align="left">Larynx</td><td align="left">Stridor (wheeze)</td></tr></tbody></table></table-wrap></p></sec><sec id="cesec293"><title>Clinical signs</title><p id="para824">The first and most important step in establishing a diagnosis of canine upper respiratory disease is to define the presenting sign. Experience has shown that an owner's ability to describe the patient's clinical signs accurately, particularly when signs are <italic>not</italic> present at the time of examination, is usually inconsistent and inaccurate, although it can be most entertaining. The four localizing clinical signs characteristically associated with upper respiratory diseases are sneezing and/or nasal discharge, stertor, stridor, and cough. Each sign, considered independently, will focus the examination to the appropriate anatomic region of the upper respiratory tract.</p><sec id="cesec294"><title>Sneezing and/or nasal discharge</title><p id="para825">Definition of the clinical signs sneezing and nasal discharge may seem intuitive. This is the most common presenting sign in dogs with upper respiratory disease. Owners that present a dog for <italic>sneezing</italic> are likely to be accurate in their description of the problem. However, the presence or absence of a nasal discharge may be more difficult to establish. Volume, character, and frequency of the discharge ultimately determine whether the owner will have even observed this sign. The astute owner will report whether the discharge is unilateral or bilateral. In the patient that has a history of sneezing and nasal discharge, instillation of a topical nasal decongestant into each nostril occasionally will provoke sneezing and elicit the nature of any discharge that is present.</p><p id="para826">Sneezing and/or nasal discharge localize the problem to the nose, nasal cavity, and paranasal sinuses. However, thorough examination of the nose and nasal cavity can be difficult, even with the availability of appropriate endoscopy equipment. In addition to careful examination of facial symmetry, the first part of the examination begins in the oral cavity, with emphasis on the maxilla, the hard palate, and the canine teeth. Examine the hard palate for evidence of trauma (penetrating or nonpenetrating) and congenital cleft palate (puppies). Carefully probe the medial aspect of the maxillary canine teeth for evidence of oronasal fistulas. Despite normal-appearing teeth and gingiva, severe, occult periodontal disease with resulting necrosis of bone does result in a septic communication between the oral and nasal cavity. The owner characteristically describes paroxysms of sneezing associated with a sanguineous nasal discharge or spray.</p><p id="para827">If these findings are negative, radiographs of the skull are indicated. Three views, obtained in the anesthetized patient, are indicated: lateral, ventrodorsal, and occlusal (open mouth) view. Radiographic interpretation of the nasal cavity and sinuses dictates that the clinician has a thorough understanding of the anatomy of the upper respiratory tract. Subsequently, with the patient still anesthetized, attempt a visual examination of the nasal cavity. Radiographs are always performed before visual examination of the nasal cavity. Manipulation of the tissue may result in intranasal bleeding, which will significantly complicate radiographic interpretation. A simple otoscope speculum placed into each nostril allows an adequate examination of the proximal 20% to 25% of the nasal cavity in most dogs. Visual examination of the caudal 75% of the nasal cavity can be attempted only with a small-diameter endoscope. Flexible and rigid scopes are available; each has advantages and disadvantages that will be discussed. Computed tomography and magnetic resonance imaging are important alternative diagnostic tools; however, expense and availability are significant limiting factors.</p><p id="para828">Understanding the most commonly diagnosed causes of sneezing and nasal discharge is especially helpful in patient management. In no particular order, the most common differential diagnoses for sneezing and/or nasal discharge include the following:<list list-type="simple" id="celist30"><list-item id="celistitem171"><label>1.</label><p id="para829"><italic>Oronasal fistulas:</italic> Especially common in middle-aged to older dogs, despite a history of recent dental prophylaxis. Empiric treatment with an orally administered antibiotic typically results in rapid and complete resolution of clinical signs, but only during the time the patient is receiving the antibiotic. Diagnosis is confirmed by probing the gingival sulcus of the upper canine teeth.</p></list-item><list-item id="celistitem172"><label>2.</label><p id="para830"><italic>Nasal neoplasia:</italic> Most commonly reported in dogs between 8 and 10 years of age (range: 1 to 15 years of age). No breed is predisposed, but the condition is uncommon in brachycephalic breeds. Persistent nasal discharge, sneezing, and intermittent epistaxis are common presenting signs. Nasal radiographs may demonstrate lytic bone lesions. Lysis of the vomer strongly supports neoplasia versus mycotic rhinitis. Exposure to tobacco smoke has been associated with 2.5 times greater risk in long-nosed dogs. No or minimal response of the discharge to antibiotics occurs. Eighty percent of nasal tumors are malignant. Adenocarcinoma is most common, followed by squamous cell carcinoma. Sarcomas account for small number of nasal tumors.</p></list-item><list-item id="celistitem173"><label>3.</label><p id="para831"><italic>Mycotic rhinitis:</italic> Difficult to distinguish from neoplasia. Persistent and voluminous mucoid nasal discharge, with or without sneezing, and nasal pain are reported. Erosion of external nares is an important physcial finding. Discharge is NOT responsive to antimicrobial treatment. Occlusal view radiographs of the nasal cavity may demonstrate evidence of turbinate destruction and/or increased fluid density on the affected side. Forty percent of patients are 3 years or younger; 80% are 7 years and younger. The diagnosis is uncommon in brachycephalic breeds. Localized <italic>Aspergillus fumigatus</italic> infection is reported most commonly.</p></list-item><list-item id="celistitem174"><label>4.</label><p id="para832"><italic>Lymphoplasmacytic rhinitis:</italic> Poorly described clinical syndrome associated with chronic sneezing and nasal discharge (bilateral or unilateral). Affected dogs are typically young to middle-aged, large-breed dogs. Signs are NOT usually responsive to antibiotics or steroids (topical or systemic). Diagnosis is based on ruling out other causes and nasal biopsy.</p></list-item></list></p></sec><sec id="cesec295"><title>Stertor</title><p id="para833">The second most common clinical sign associated with upper respiratory disease in dogs, stertor refers to intermittent, yet persistent, or continuous snorting, also called stertorous breathing. Paroxysms of stertor, typically called reverse sneezing, characterize rapid, consecutive inspiratory bursts through the nose. Seldom actually seen during examination, reverse sneezing is likely to result from the patient's attempt to displace matter trapped in the nasopharynx and move it into the oropharynx, where it can be swallowed.</p><p id="para834">Visualization of the nasopharynx and choana is essential in the patient that has chronic or persistent stertor. The examination can be accomplished only in the anesthetized patient. Sedation is NOT sufficient to conduct the examination. A flexible endoscope with the ability to flex approximately 170 to 180 degrees is recommended. Examination allows visualization of the nasopharynx and associated mucosa, the choana (posterior nares), and the top of the soft palate (see <xref rid="f34" ref-type="fig">Figure 34-34</xref>).</p><p id="para835">Nasopharyngeal foreign bodies are by far the most common finding. Sticks, plant material (grass and juniper twigs), peas, cotton balls, and thread are just a few examples. Neoplasia is the second most common finding. In cats, lymphoma (feline leukemia virus–related) obstructing the choana most commonly is observed (see <xref rid="f35" ref-type="fig">Figure 35-35</xref>). In dogs, neoplasia is uncommon, but (in my experience) sarcomas in young dogs have been seen most frequently.</p></sec><sec id="cesec296"><title>Stridor</title><p id="para836">The least commonly encountered of the upper respiratory signs is stridor, or stridulous breathing. Stridor is audible wheezing and is associated with restriction to airflow, usually at the level of the larynx. Therefore stridor is <italic>the most critical and potentially life-threatening upper respiratory sign.</italic>This is especially true when stridor is continuous. The patient that has continuous stridor deserves immediate attention. Make every effort to discern the cause once the clinical sign is characterized. In obtaining the history, owners generally describe wheezing accurately; however, some patients actually may present for severe dyspnea or orthopnea. Careful questioning of the client is indicated to determine whether wheezing is associated with the additional effort to breath. The clinician also should make an effort to discern whether the owner has observed any change in the ability of the dog to vocalize or bark.</p><p id="para837">Simply listening to the patient breath in a quiet room is the first step in assessing stridor. A stethoscope is not required to hear wheezing but should always be used to examine the cervical trachea, the larynx, and the lungs. Any restriction to airflow in the larynx or cervical trachea can cause stridor. However, in the majority of cases, the stridor will be significantly louder at the level of the larynx, indicating a restrictive lesion at that level.</p><p id="para838">If any indication of respiratory distress is reported or is manifest during the examination, subject the patient to a visual examination under general anesthesia. Sedation is NOT sufficient to conduct the examination. Be prepared. These patients are NOT routine. Emergency resuscitation may be required on induction of anesthesia, including the need to perform a tracheostomy.</p><p id="para839">On induction, <italic>carefully</italic> place an endotracheal tube. If there are no complications associated with inserting the tube, once anesthesia is effectively induced and the patient is stable, a lateral and dorsoventral radiographs of the larynx and cervical trachea are indicated. Metallic objects (e.g., fish hooks) can become buried in the mucosa and may not be observed during a visual examination.</p><p id="para840">Remove the endotracheal tube in order to conduct a visual examination. A focal, hands-free light source directed into the oropharynx is strongly recommended. Carefully examine the epiglottis, arytenoid cartilages, glottis and vocal folds using a cotton-tipped applicator. Careful observation of the symmetry and function of the arytenoid cartilages is essential. The left and right cartilages normally respond to tactile stimuli when the patient is in a light plane of anesthesia; both sides should move to the medial plane rapidly and at the same time. They may not close, depending on the depth of anesthesia. It should be possible to visualize the cartilage on the inside of the tracheal rings while looking through the glottis.</p><p id="para841">In large breed, middle-aged and older dogs, laryngeal paralysis is the most common cause of stridor. Associated signs may include exercise intolerance and collapse during exertion. Laryngeal paralysis and stridor also may be observed in young breeds as a congenital disorder (Dalmatian, Rottweiler, Bouvier des Flandres, Siberian Husky, and Bull Terrier). Foreign body penetration of the laryngeal tissues can cause serious and life-threatening obstruction because of infection and swelling. Neoplasia may cause obstructive mass lesions involving the larynx, especially squamous cell carcinoma and lymphoma. Granulomatous laryngeal disease and fungal mycetoma have been reported.</p><p id="para842">The presence of a mass lesion, assuming there is no foreign body detected, warrants biopsy of the lesion. Additional effort to control postbiopsy bleeding is important. I use a cotton-tipped applicator saturated with a 1:10,000 dilution of epinephrine held against the biopsy site for 30 to 60 seconds. This is time well spent. Postbiopsy administration of systemically effective dexamethasone has been suggested to control laryngeal swelling, but I have not found this to be effective or important.</p><sec id="cesec297"><title>Additional Reading</title><p id="para843">Holt DE: Upper airway obstruction, stertor, and stridor. In King LG, editor: <italic>Textbook of respiratory disease in dogs and cats,</italic> St Louis, 2004, Elsevier-Saunders.</p><p id="para844">Van Pelt DV, McKiernan BC: Pathogenesis and treatment of canine rhinitis, <italic>Vet Clin North Am Small Anim Pract</italic> 24:789, 1994.</p><p id="para845">Van Pelt DV, Lappin MR: Pathogenesis and treatment of feline rhinitis, <italic>Vet Clin North Am Small Anim Pract</italic> 24:807, 1994.</p><p id="para846">Withrow SJ: Tumors of the gastrointestinal system: cancer of the oral cavity. In Withrow SJ, MacEwan EG, editors: <italic>Small animal clinical oncology,</italic> ed 3, Philadelphia, 2001, Saunders.</p></sec></sec></sec></sec></sec><sec id="cesec298"><sec id="cesec299"><title>Lower Respiratory Tract</title><p id="para847">NOTE: Cats and dogs with acute, severe dyspnea must be regarded as having a life-threatening condition until proved otherwise. Immediate therapeutic and diagnostic intervention is indicated. Section 1 describes appropriate interventive procedures for the management of these patients.</p><p id="para848">The following diagnostic procedures are elective and are indicated in patients with chronic disorders of the lower respiratory tract that are not considered life-threatening.</p><sec id="cesec300"><title>Transtracheal aspiration</title><p id="para849">Transtracheal aspiration is a safe and clinically useful method for obtaining material for cytologic and bacteriologic examination from the lower respiratory tract of medium-sized to large dogs without invading the oval cavity. This procedure is <italic>not</italic> indicated in cats. The technique can be performed on the unanesthetized animal, although some sedation may be necessary in fractious animals. In small dogs and cats, tracheal aspirates are collected by passing the catheter through sterile tracheal tubes. Light levels of anesthesia are used to accommodate coughing and tracheal intubation.</p><p id="para850">Place the animal in sternal recumbency, and elevate and extend the head. Clip the area around the larynx, and surgically prepare the skin. Locate the cricothyroid membrane by moving the finger cranially along the trachea until the large ventral ridge of the cricoid cartilage is felt. Use a 16-gauge, ½-inch intravenous catheter to collect material through the trachea (<xref rid="f55" ref-type="fig">Figure 4-55</xref>). Puncture the cricothyroid membrane with the 16-gauge needle, and pass the catheter into the trachea until it reaches the distal trachea or main stem bronchus. (Alternatively, in large dogs, insert the catheter between the tracheal rings at the junction of the middle third and distal third of the cervical trachea.) Withdraw the needle, and leave the catheter in place. Attach a 12-mL syringe containing sterile saline solution to the catheter. Expel 1 to 2 mL of saline from the syringe. When the animal coughs, aspirate with the syringe to collect cells and mucus for bacteriologic and cytologic examination. When material has been collected, remove the catheter and bandage the animal's neck. Culture material present in the syringe in blood agar and in thioglycollate medium. Prepare material from aspiration for cytologic examination. Press large plugs of mucus between two clean glass slides, and stain thin smears with Wright's or Giemsa stain.<fig id="f55"><label>Figure 4-55</label><caption><p><bold>A,</bold> Diagramatic representation of anatomic structures involved with transtracheal aspiration technique. The best landmark for percutaneous puncture is the cricothyroid ligament of the larynx, although the tracheal lumen also can be entered between cervical tracheal rings. <bold>B,</bold> The needle is advanced and directed slightly caudad until the trachea is entered. Once the needle is positioned within the tracheal lumen, the catheter is advanced through the needle and down the trachea.</p></caption><graphic xlink:href="gr55"></graphic><attrib>(From Kirk RW: Current veterinary therapy VIII: Small Animal Practice, Philadelphia, 1983, WB Saunders.)</attrib></fig></p><p id="para851">Complications of transtracheal aspiration biopsy include catheter trauma to the lower airway or needle trauma to the larynx, resulting in bleeding, subcutaneous emphysema, pneumomediastinum, pneumothorax, or airway obstruction.</p></sec><sec id="cesec301"><title>Endotracheal wash</title><p id="para852">In cats and small dogs and in dogs for which general anesthesia is <italic>not</italic> contraindicated, tracheal aspiration (or tracheal wash) is a relatively safe, easy-to-perform procedure that can yield excellent diagnostic cytologic and culture specimens. The procedure has some advantages over transtracheal aspiration in that it allows sample collection from airways beyond the bifurcation of the trachea (carina) and avoids complications associated with patient discomfort and movement during the procedure. However, cough reflexes are eliminated completely, thereby decreasing potential sample yields from deep in the airway structure. In either case, transtracheal and tracheal aspirations provide the best diagnostic material from large airways, not small airways and alveoli.</p><p id="para853">The anesthetized dog or cat usually is placed in sternal recumbency. Lateral recumbency (affected side down) may facilitate recovery of specimens from patients with focal or regional lung disease. Use a sterile endotracheal tube to administer the anesthetic and oxygen. Introduce a sterile red rubber catheter (long enough to extend beyond the carina) through the endotracheal tube (<xref rid="f56" ref-type="fig">Figure 4-56</xref>). (NOTE: Disposable adapters for use with endotracheal tubes are available that allow continuous administration of anesthetic gases while passing the rubber catheter through the tube [<xref rid="f57" ref-type="fig">Figure 4-57</xref>].) Introduce the catheter blindly until resistance is met as the tube attempts to enter smaller airways.<fig id="f56"><label>Figure 4-56</label><caption><p>Endotracheal wash performed directly through a prepositioned endotracheal tube in a cat.</p></caption><graphic xlink:href="gr56"></graphic></fig><fig id="f57"><label>Figure 4-57</label><caption><p>An endotracheal tube adapter can be used in medium to large dogs to enable administration of anesthetic gases and oxygen throughout the endotracheal wash procedure.</p></caption><graphic xlink:href="gr57"></graphic></fig></p><p id="para854">Use aliquots of warmed, sterile saline in prepared syringes to wash and retrieve samples. Aliquots of 3 to 5 mL can be used per collection attempt in small dogs and cats, whereas volumes up to 10 and 20 mL are appropriate for larger dogs. With the catheter positioned as deep as practical in the airway, infuse the entire volume of saline. <italic>Gentle agitation</italic> (intermittent aspiration/injection) may facilitate sample collection. If 10 mL is infused, retrieval of only 1 to 2 mL as a final volume per collection attempt is not unusual. The remaining fluid is rapidly (seconds) absorbed into the pulmonary vasculature. IMPORTANT: When performing this procedure, DO NOT withdraw the rubber catheter while maintaining a high negative pressure on the syringe. Doing so actually may tear mucosa away from the airway and could lead to pneumothorax or pneumomediastinum.</p><p id="para855">The procedure can be repeated safely in the same patient several times. Collection of three to five samples is routine. More samples may be indicated depending on the patient's condition and response to the procedure. Monitoring of patients undergoing a tracheal wash procedure for oxygen saturation (pulse oximetry) throughout the procedure is recommended. In some patients with reactive airways, infusion of saline may cause significant bronchoconstriction, detected by a rapid decline in oxygen saturation.</p><p id="para856">Process samples collected immediately. Submit at least one sample of liquid (<italic>not</italic> a swab of the liquid) for bacterial culture and sensitivity or MIC. Quantitative cultures are impractical because specimens will be diluted. If the sample appears to be highly cellular (characterized by turbidity), place aliquots into tubes containing EDTA.</p><sec id="cesec302"><sec id="cesec303"><title>Additional Reading</title><p id="para857">Syring RS: Tracheal washes. In King LG, editor: <italic>Textbook of respiratory disease in dogs and cats,</italic> St Louis, 2004, Elsevier-Saunders.</p></sec></sec></sec></sec></sec><sec id="cesec304"><sec id="cesec305"><title>Bronchoalveolar Lavage</title><p id="para858">Bronchoalveolar lavage (BAL) is an alternative diagnostic procedure to transtracheal aspiration and endotracheal wash. Bronchoalveolar lavage has the advantage of retrieving fluid samples from distal airways and alveoli. This is a highly diagnostic procedure indicated in patients with generalized lung and regional (interstitial and/or airway) disease that are <italic>not</italic> in respiratory distress. Patients suspected of having allergic or infectious respiratory disease or neoplasia are candidates for BAL. Although BAL is used as a therapeutic procedure in human beings with chronic lung disease associated with accumulations of surfactant in the alveoli, there is no therapeutic indication for BAL in dogs or cats.</p><p id="para859">The technique for BAL entails instilling sufficiently large volumes of fluid into the distal airways to reach, and recover, reasonable cytological samples representative of small airways and alveoli. Several variations on the technique are described, but all recommend blind or visual placement of a catheter or bronchoscope into an airway of a lung lobe such that the airway is occluded. Sterile, nonbacteriostatic 0.9% saline, warmed to approximately body temperature and drawn into prepared syringes, is the fluid of choice. The volume of fluid varies with the size of the patient. Defined doses of saline per kilogram of body mass have not been described. In large dogs, two 25-mL aliquots (50 mL total) can be infused into each lobe sampled. In small dogs and cats, total volumes per lobe generally are restricted to 10-mL aliquots. Recovery may be as low as 2 to 5 mL with each attempt.</p><p id="para860">For dogs undergoing BAL, particularly when reactive (allergic) airway disease is suspected, pretreatment with a bronchodilator is appropriate and is recommended. Aminophylline can be administered at 5 mg/kg (cats) or 11 mg/kg (dogs) orally 1 to 2 hours before the procedure. Alternatively, terbutaline, 0.01 mg/kg, can be administered subcutaneously to cats 30 minutes before the procedure.</p><p id="para861">Bronchoscopic BAL allows direct visualization of the airway/lobe of interest. In medium to large dogs, place the bronchoscope directly through a sterile endotracheal tube. Use of an inexpensive, disposable endotracheal tube adaptor permits simultaneous administration of oxygen and anesthetic throughout the procedure. Saline can be infused from a syringe directly through the biopsy channel of the endoscope. The bronchoscope serves as the infusion catheter. Using this technique, samples can be collected effectively from multiple lobes. Blind placement (nonbronchoscopic) BAL using a rubber end-hole catheter is required in cats and small dogs. Blind placement is also appropriately used in patients with generalized lung or airway disease when discrete placement of bronchoscope cannot be accomplished reliably.</p><p id="para862">As with the endotracheal wash procedure described before, <italic>gentle agitation with the syringe</italic> (intermittent aspiration/injection) may facilitate sample collection. DO NOT withdraw the bronchoscope or catheter while maintaining significant negative pressure because this may lacerate the airway, leading to pneumothorax or pneumomediastinum.</p><p id="para863">Bronchoalveolar lavage is an invasive diagnostic procedure that is not without risk of injury or death. Following completion of BAL, administration of 100% oxygen for 5 to 10 minutes via endotracheal tube is recommended for all patients. Evaluate the patient carefully for breathing effort and oxygen saturation (pulse oximetry) during recovery. Although significant quantities of fluid remain in the airways following BAL, most of the volume is absorbed rapidly. Residual amounts of fluid, however, can be retained for 24 to 48 hours after the procedure. During this time, some patients will manifest cough. Crackles may be auscultated.</p><sec id="cesec306"><sec id="cesec307"><sec id="cesec308"><title>Additional Reading</title><p id="para864">Hawkins EC: Bronchoalveolar lavage. In King LG, editor: <italic>Textbook of respiratory disease in dogs and cats</italic>, St Louis, 2004, Elsevier-Saunders.</p><p id="para865">Hawkins EC, DeNicola DB, Plier ML: Cytological analysis of bronchoalveolar lavage fluid in the diagnosis of spontaneous respiratory tract disease in dogs, <italic>J Vet Intern Med</italic> 9:386-392, 1995.</p></sec></sec></sec></sec></sec><sec id="cesec309"><sec id="cesec310"><title>Fine-Needle Aspiration of Lung</title><p id="para866">Percutaneous aspiration needle biopsy can be helpful in establishing a diagnosis in conditions such as (l) chronic inflammatory disease of the lung—for example, granulomatous lung disease caused by mycotic organisms; (2) chronic inflammatory disease; (3) metastases to the lung; and (4) primary lung tumors. The biopsy may provide enough diagnostic information to preclude performing an exploratory thoracotomy. Lung biopsy is contraindicated in animals with hemorrhagic disease or thoracic disease that produces forceful breathing and coughing.</p><p id="para867">Clip and surgically prepare the biopsy site. Infiltrate the skin, subcutaneous tissue, muscle, and parietal pleura with 1% to 2% lidocaine.</p><p id="para868">In diffuse parenchymal lung disease, taking biopsy material from the diaphragmatic lobes is recommended. The dorsal portions of the seventh to ninth intercostal spaces are preferred for percutaneous biopsies. In diffuse lesions, take biopsy material from the right or left thorax.</p><p id="para869"><italic>Understanding of the risks associated with performing fine-needle aspiration of lung is important.</italic> Lung aspirates will yield only cells, fluid, and trace amounts of tissue, yet there is a significant risk of inducing pneumothorax following the procedure, even when performed without difficulty or complications.</p><p id="para870">When performing the procedure, a 22- to 25-gauge disposable needle (such as a 1-inch spinal needle) with stylet is preferred. Leave the stylet within the needle until the lung has been penetrated. Then quickly remove the stylet and immediately attach a sterile 6- to 12-mL syringe. The amount of air that might enter the lung between the time the stylet is removed and the syringe attached is negligible. Holding the syringe carefully and steadily against the patient's thorax, establish negative pressure in the same manner as when obtaining an aspiration from a lymph node. As much as the patient will permit, attempt three to four aspirations without withdrawing the needle.</p><p id="para871">Alternatively, insert a conventional 25-gauge needle, attached to a 6-mL syringe, <italic>subcutaneously</italic> over the area of interest. Then establish significant negative pressure while the tip of the needle is still positioned in the subcutaneous tissues <italic>outside the parietal pleura.</italic> While maintaining the same amount of negative pressure in the syringe, direct the needle into the lung, leave it in place for 1 to 2 seconds, and withdraw it completely. Apply any material collected directly to glass slide. This procedure is best conducted in patients that are awake. Attempting the procedure in anesthetized dogs or cats could result in an unsuccessful aspirate or, if the lungs were under positive pressure (ventilation or bagging), the risk of causing pneumothorax could be increased. Other reported complications include hemothorax (always exciting), lung laceration caused by patient movement during the procedure, pulmonary hemorrhage, and hemoptysis. Contraindications to fine-needle aspiration include patients with a known bleeding diathesis and coagulopathy, thrombocytopenia, uncontrolled coughing, pulmonary hypertension, pulmonary cysts, and bullous emphysema.</p><p id="para872">Ultrasound guided techniques for fine-needle aspiration or biopsy of lung recently have been described and generally are associated with fewer procedural complications. However, additional training and experience, in addition to having access to the proper size and type of ultrasound probe, is critical.</p><sec id="cesec311"><sec id="cesec312"><sec id="cesec313"><title>Additional Reading</title><p id="para873">Cole SG: Fine needle aspirates. In King LG, editor: <italic>Textbook of respiratory disease in dogs and cats,</italic> St Louis, 2004, Elsevier-Saunders.</p></sec></sec></sec></sec></sec><sec id="cesec314"><sec id="cesec315"><title>Nebulization and Aerosol Therapy</title><p id="para874">Inhalation therapy can be defined as nebulization (humidification of the inspired air) and aerosol therapy (the process whereby drugs are vaporized in a solution and delivered directly into the respiratory tract). In companion animals, inhalation therapy is most useful for humidifying air in the respiratory tract and moistening the mucous membranes (nebulization). Sustained inspiration of dry air/gases causes irritation to the respiratory epithelium, which in turn results in swelling, bronchial gland hypertrophy, goblet cell proliferation, and loss of ciliary epithelium over time. Respiratory secretions become thick and tenacious, and efficient bronchial drainage is impaired.</p><p id="para875">The objectives of inhalation therapy include the following:<list list-type="simple" id="celist31"><list-item id="celistitem175"><label>1.</label><p id="para876">Humidification of bronchial mucous membranes</p></list-item><list-item id="celistitem176"><label>2.</label><p id="para877">Deposition of miniscule amounts of potent drugs in smaller airways to obtain optimal topical therapeutic effects with minimal systemic side effects (e.g., bronchodilators)</p></list-item><list-item id="celistitem177"><label>3.</label><p id="para878">Deposition of moderate amounts of potent agents or agents that are only effective topically (e.g., antibiotics and mucolytics)</p></list-item><list-item id="celistitem178"><label>4.</label><p id="para879">Deposition of relatively large quantities of bland substances that promote bronchial drainage with minimal irritation (e.g., saline, propylene glycol, glycerine, and detergents)</p></list-item></list></p><p id="para880">Nebulization is used (l) in combination with oxygen therapy; (2) in tracheostomy care; (3) in acute respiratory diseases such as tracheobronchitis, bronchiolitis, upper respiratory disease of cats, pneumonia, and postoperative atelectasis and pneumonia; and (4) in chronic respiratory diseases such as chronic bronchitis, bronchopneumonia, collapsed trachea with secondary tracheobronchitis, emphysema, and bronchiectasis.</p><p id="para881">Aerosol therapy, however, is a limited-use therapeutic technique used in dogs and cats to administer antimicrobials, bronchodilators (aminophylline, 100 mg), or corticosteroids. The advantage of doing so is to achieve relatively high levels of drug in the respiratory tract in patients with <italic>defined</italic> lower respiratory tract disease. In addition, administration of potentially toxic antimicrobials (aminoglycosides) by this route has been shown to be associated with minimal or insignificant uptake into the general circulation, thereby minimizing (or eliminating) any risk of renal toxicity.</p><sec id="cesec316"><title>Principles of action</title><p id="para882">Large water particles (10 to 60 μm) in the high-velocity air flow of the nose and throat settle on the mucosa of the larynx, nose, and throat. Particles smaller than 10 μm (2 to 10 μm) are deposited in the bronchi, but only the smallest particles reach the bronchioles. Ultrasonic aerosol generators are the most effective machines for nebulization. The mists can be directed into a cage or a face mask. If nebulization is used with an endotracheal or tracheostomy tube, warm inspired gases to body temperature.</p><p id="para883">Dense mist from an unheated jet nebulizer (<xref rid="f58" ref-type="fig">Figure 4-58</xref>) contains only slightly more water than is needed to humidify air with temperature increasing from 22° to 37° C. Evaporation of the aerosol solution can be prevented by stabilization; that is, by heating it to 35° C or by reducing the vapor pressure by adding 10% propylene glycol. Because distilled water and hypertonic solutions are irritating to the mucosa, use only isotonic or half-strength isotonic saline.<fig id="f58"><label>Figure 4-58</label><caption><p>Disposable jet nebulizer used to administer humidified air and/or medication directly into the respiratory tract.</p></caption><graphic xlink:href="gr58"></graphic></fig></p><p id="para884">Although continuous, low-level humidification of the oxygen tent atmosphere is necessary, periodic medication by aerosol spray is permissible. High levels of water can be introduced several times daily for 10 to 15 minutes per treatment, or drugs can be added to the solution during these times. Many drugs have been used; isoproterenol, epinephrine, and phenylephrine are some of the drugs that may cause bronchodilation and decreased airway resistance.</p><p id="para885">Differentiation of obstruction of the bronchi caused by pulmonary edema from that caused by bronchial secretions is important. In both cases the patient cannot breath because of fluids or semifluid liquids in the bronchi. In pulmonary edema, the fluid turns to a frothy, bubbly material that produces a rattling sound in the trachea.</p><p id="para886">Thick, inflammatory exudates, however, must be thinned by detergent materials that liquefy bronchial secretions. However, these agents increase frothing and, indirectly, anoxia if used in pulmonary edema. In patients with acute, severe bronchial inflammation, mucolytic agents such as acetylcysteine have been administered <italic>intravenously</italic> to dogs at doses as high as 144 mg/kg followed by 70 mg/kg 12 hours later. Aerosolization of acetylcysteine currently is used in human medicine but has not been validated in dogs.</p><p id="para887">Heated aerosol units (vaporizers) produce large water droplets that do not penetrate small bronchioles and may overheat the patient. These units should not be used for intensive therapy.</p></sec><sec id="cesec317"><title>Drug delivery by aerosolization</title><p id="para888">Drugs that can be applied by jet nebulizer (<xref rid="f59" ref-type="fig">Figure 4-59</xref>, <xref rid="f60" ref-type="fig">Figure 4-60</xref>) include the following:<list list-type="simple" id="celist32"><list-item id="celistitem179"><label>1.</label><p id="para889"><italic>Bronchodilators:</italic> Always use bronchodilators when administering drugs that may be irritating and constricting, such as isoetharine hydrochloride 1% and phenylephrine 0.25%, 0.5 to 1.0 mL in 2 to 3 mL of saline 3 to 4 times daily.</p></list-item><list-item id="celistitem180"><label>2.</label><p id="para890"><italic>Antibiotics:</italic> Antibiotics are poorly absorbed from the respiratory mucosa. Systemic administration of most antibiotics produces adequate pulmonary concentration for antibacterial effect. For <italic>Bordetella</italic> spp. that are located at the tips of bronchial cilia, topical contact via nebulization may be useful. Antibiotics that have been used successfully and safely include kanamycin (250 mg in 5 mL saline twice daily); gentamicin (50 mg in 5 mL saline twice daily); and polymyxin B (333,000 IU in 5 mL saline twice daily).</p></list-item><list-item id="celistitem181"><label>3.</label><p id="para891"><italic>Bland solutions:</italic> Use these in large volume for prolonged mist effect: 0.9% sterile saline (5 to 200 mL as needed); glycerine (5% in saline); and propylene glycol (10% to 20% solution in saline).</p></list-item><list-item id="celistitem182"><label>4.</label><p id="para892"><italic>Detergents and mucolytics:</italic> These compounds are irritating and currently are not recommended by most authors.</p></list-item><list-item id="celistitem183"><label>5.</label><p id="para893"><italic>Antifoaming agents:</italic> Administer ethyl alcohol (70% solution 5 to 10 mL twice daily).</p></list-item></list><fig id="f59"><label>Figure 4-59</label><caption><p>Disposable jet nebulizer attached to a face mask for administering aerosol therapy to dogs.</p></caption><graphic xlink:href="gr59"></graphic></fig><fig id="f60"><label>Figure 4-60</label><caption><p>A disposable jet nebulizer attached to a Plexiglas anesthesia induction box for administration of aerosol therapy to cats.</p></caption><graphic xlink:href="gr60"></graphic></fig></p><sec id="cesec318"><sec id="cesec319"><title>Additional Reading</title><p id="para894">Boothe DM: Drugs affecting the respiratory system. In King LG, editor: <italic>Textbook of respiratory disease in dogs and cats,</italic> St Louis, 2004, Elsevier-Saunders.</p><p id="para895">Tseng LW, Drobatz KJ: Oxygen supplementation and humidification. In King LG, editor: <italic>Textbook of respiratory disease in dogs and cats,</italic> St Louis, 2004, Elsevier-Saunders.</p></sec></sec></sec></sec></sec></sec><sec id="cesec320"><sec id="cesec321"><title>URINARY TRACT PROCEDURES</title><sec id="cesec322"><title>Urohydropropulsion</title><p id="para896">Removal of uroliths from dogs and cats is a commonly performed, yet critically important, clinical procedure. Several techniques are available for removal of calculi and obstructive concretions in male and female animals. Cystotomy is performed routinely to remove calculi from the lumen of the bladder but especially in male dogs may not be an effective approach for removing obstructing urethral calculi. Advanced and expensive techniques recently have been described: laparoscopic-assisted cystotomy, Ellik evaculator, use of stone “baskets” (through a cystoscope), and lithotripsy are examples. However, for removal of uroliths from dogs and cats with partial or complete urinary obstruction, urohydropulsion is among the more effective yet inexpensive techniques available.</p><p id="para897">Urohydropulsion is a therapeutic procedure for removal of foreign material, namely, uroliths, from the bladder and/or urethra of dogs. Two techniques are described: <italic>voiding</italic> urohydropulsion and <italic>retrograde</italic> urohydropulsion. Both procedures have advantages and disadvantages.</p><sec id="cesec323"><title>Voiding urohydropulsion</title><p id="para898">The objective of voiding urohydropulsion is to induce forceful voiding of urine by manually compressing the bladder to facilitate removal of cystic uroliths in <italic>female</italic> dogs. Do not perform this procedure until it can be confirmed by catheterization or cystoscopy that the urethra is patent. With the bladder filled with urine or saline (via catheterization), lift the patient (preferably sedated or anesthetized, although this procedure can be done in the awake patient) into a position such that the tail and perineum are ventral and head is upright. The spine should be approximately perpendicular to the working surface. Using one or both hands, gradually increase pressure on the bladder to induce and maintain a forceful stream of urine. Objectively, small uroliths will be extruded. If the procedure is only partially successful, it can be repeated as necessary. Obviously, voiding urohyropulsion has limitations and cannot be used in male dogs or in dogs with urethral obstructions or strictures.</p></sec><sec id="cesec324"><title>Retrograde urohydropulsion</title><p id="para899">This procedure is indicated for male dogs and cats with partial or complete urethral obstruction caused by uroliths or accumulations of “sand.” Preferably perform the procedure in the anesthetized patient.</p><p id="para900">NOTE: There are discrepancies in the literature regarding whether to empty the urinary bladder of urine before performing this procedure. Because patients with urethral obstructions may have a significant volume of urine in the bladder at the time of presentation, some authors recommend performing cystocentesis to relieve the internal pressure before attempting urohydropulsion. However, in patients that have had a profoundly distended bladder for several hours (even days), penetrating the urinary bladder with a needle presents significant risk of rupturing a fragile bladder. The next step, of course, is abdominal surgery. I recommend avoiding cystocentesis whenever possible. The volume of saline required to flush uroliths into the bladder is inconsequential considering the total volume already present.</p><p id="para901">With the patient positioned in lateral recumbency, retract the prepuce and expose the penis as for conventional bladder catheterization technique. Use sterile technique to pass an appropriately sized flexible catheter, which is advanced to the point of obstruction. Attach a catheter-tipped 60-mL syringe filled with warmed (my preference) sterile saline and a water-soluble lubricant mixture (approximately 2 parts saline to 1 part lubricant) to the urinary catheter. An assistant places a gloved (always preferred) finger into the rectum to identify and occlude the lumen of the pelvic urethra at the level of the pubis. Subsequently, infuse saline forcefully into the catheter to dilate the urethra proximal to the obstructing urolith. At that point, release the digital pressure on the proximal urethra while the solution continues to be infused through the catheter. Objectively, the pressure within the urethra forces small stones retrograde into the urinary bladder, thereby relieving the obstruction.</p><p id="para902">NOTE: The objective of this procedure is NOT to push the calculi into the bladder with the catheter, because this can substantially injure the urethral mucosa, nor to force the calculi around the catheter and move it antegrade.</p><sec id="cesec325"><sec id="cesec326"><title>Additional Reading</title><p id="para903">Adams LG, Syme HM: Canine lower urinary tract diseases. In Ettinger SJ, Feldman EC, editors: <italic>Textbook of veterinary internal medicine,</italic> ed 6, St Louis, 2005, Elsevier-Saunders.</p><p id="para904">Osborne CA, Finco DR: <italic>Canine and feline nephrology and urology,</italic> Baltimore, 1995</p></sec></sec></sec></sec></sec></sec></body><back><fn-group><fn id="fn1"><label>*</label><p id="cenotep9">GIF-Tube Kit (Greta Implantable Fluid Tube); VSM, Phoenix, Arizona, <ext-link ext-link-type="uri" xlink:href="http://www.practivet.com" id="interref1">www.practivet.com</ext-link>.</p></fn><fn id="fn2"><label>†</label><p id="cenotep10">Vet-Jet Transdermal Administration System for delivery of the recombinant feline leukemia vaccine; Merial Ltd., Duluth, Georgia.</p></fn></fn-group></back></pmc></record>Pour manipuler ce document sous Unix (Dilib)
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