Upper airway procedures are performed to remove, repair, or by-pass areas of obstruction, injury, or disease. Affected animals may show signs of mild to severe respiratory distress. Mild or moderately dyspneic patients should initially be examined from a distance to avoid exacerbating the condition. Open mouth breathing, abducted forelimbs, labored breathing, and restlessness indicate moderate to severe respiratory distress that may require emergency therapy. Minimal restraint should be used in severely dyspneic patients and they should be allowed to maintain the position they feel most comfortable in. Supplemental oxygen may be given via nasal insufflation, tracheostomy tube or catheter, endotracheal intubation, mask, or cage. Corticosteroids, sedation, and/or cooling may relieve distress. Sedation may be beneficial for anxious patients in moderate to severe respiratory distress. Combinations of intravenous drugs are commonly used; oxymorphone or butorphanol plus either acepromazine or diazepam are commonly given in dogs. Alternatively, fentanyl plus droperidol may be used. In cats, acepromazine or diazepam is recommended. To cool dyspneic animals, a fan may be directed at the patient, ice packs may be applied to the head, axilla, inguinal area, and extremities, and/or cooled fluids may be administered intravenously.

Diagnosis of upper respiratory disease is based on history and clinical signs, physical examination findings, hematologic and serum biochemical parameters, radiographs, endoscopy, cytology, culture, and/or biopsy. History and clinical signs may include abnormal respiratory noises (e.g., cough, stridor, wheeze), exercise intolerance, hyperthermia, tachypnea, dyspnea, cyanosis, restlessness, and/or collapse. Gagging and regurgitation of secretions are common with nasopharyngeal, laryngeal, and some tracheal abnormalities. Voice change may occur with laryngeal paralysis and dysphagia may be noted with supraglottic obstructions. Subcutaneous emphysema occurs with penetrating laryngotracheal injuries. Clinical signs may intensify or be precipitated by excitement, stress, eating, drinking, or high ambient temperatures. Laboratory data should be evaluated to determine the presence of underlying metabolic disease and advisability of general anesthesia. Tidal breathing flow volume loops are helpful in classifying obstructions as fixed or non-fixed. Pulmonary function tests, electromyography, and nerve conduction studies are ancillary tests which may support the presence of pulmonary or neuromuscular disease.

Animals with nasal neoplasia, fungal infection, or foreign bodies may be anemic due to profuse epistaxis. Affected animals should be carefully evaluated for clotting abnormalities by assessing platelet numbers, bleeding from venipuncture sites, or the presence of ecchymoses, petechiation, melena, hematuria, or retinal hemorrhages. If available, coagulation ability may be assessed by activated clotting time, prothrombin time, or partial thromboplastin time. Blood transfusions should be given before surgery if the PCV 20%. Bleeding during rhinotomy may be severe, requiring intraoperative blood transfusion and/or carotid artery ligation.

Preoperative anti-inflammatory doses of corticosteroids may reduce nasopharyngeal and/or upper airway edema secondary to surgical or diagnostic manipulations. They are routinely given when nasopharyngeal and intraluminal laryngeal procedures are performed.

Antibiotics

The respiratory tract has a normal bacterial flora; therefore, prophylactic antibiotics (e.g., cefazolin) are frequently given prior to surgery. However, animals with normal immune function undergoing short procedures (e.g., nares resection, laryngeal saccule resection, vocal cordectomy) don't need them. Streptococcus spp, E.coli, Pseudomonas spp, Klebsiella spp., and Bordetella bronchiseptica are most commonly isolated from normal dogs. In one study, approximately 64% of trachea cultures were sterile, whereas 95% of pharyngeal cultures were positive.

Most canine respiratory tract infections are due to gram-negative organisms, many being resistant to commonly used antibiotics. Antimicrobial drug selection should be based on cytologic and culture results of tracheobronchial, pulmonary parenchymal, and/or pleural secretions. Bland aerosol therapy (e.g., sterile 0.9% NaCl) helps loosen secretions and facilitate their clearance in dogs with tracheostomies. Addition of antibiotics to the aerosol is generally unnecessary. Lipid soluble antibiotics that contain a benzene ring reach highest levels in the normal trachea and bronchus; however, increased permeability associated with inflammation allows numerous antibiotics to achieve high levels during infection. Antibiotics commonly recommended for treatment of respiratory disease include ampicillin, potentiated sulfonamides, cephalosporins, aminoglycosides, and fluoroquinolones.

TRACHEAL COLLAPSE

Tracheal collapse is a commonly recognized disease of toy and miniature-breed dogs (e.g., Toy Poodles, Yorkshire Terriers, Pomeranians, Maltese, Chihuahuas) associated with tracheal cartilage flaccidity and flattening. Either sex may be affected and most animals are middle-aged when clinical signs are noted; however, animals as young as one year of age may be diagnosed with this condition. The etiology of tracheal collapse is unknown and probably multifactorial, but may include genetic and nutritional factors, neurologic abnormalities, and cartilage matrix degeneration. In affected animals, the cartilages usually collapse in a dorsoventral direction, with the cervical trachea collapsing during inspiration and the thoracic trachea collapsing during expiration.

With tracheal collapse, clinical signs often progress with age and include abnormal respiratory noise, dyspnea, exercise intolerance, cyanosis, and syncope. Clinical signs are more severe in obese animals. Respiratory noises include wheezing, hacking, coughing, and stridulous breathing, although some dogs do not make abnormal respiratory noises. The cough may be productive or nonproductive but is classically a "goose honk" cough. Coughing often becomes cyclic and paroxysmal and gagging is common. Signs may be elicited or exacerbated by tracheal infections or tracheal compression. Second-hand smoke may also precipitate clinical signs.

Differential diagnoses that should be considered include other causes of chronic coughing or respiratory distress such as brachycephalic syndrome, tonsillitis, laryngeal collapse, laryngeal paralysis or paresis, bronchitis, tracheobronchitis, allergies, heartworm disease, pulmonary disease, chronic mitral valvular disease, hypoplastic trachea, tracheal stenosis, and tracheal neoplasia.

Tracheal hypoplasia is a congenital form of tracheal stenosis that occurs when tracheal cartilages are abnormally small and abnormally shaped. The tracheal cartilages are circular (the ends appose or overlap), rather than C-shaped, causing tracheal rigidity. The dorsal tracheal membrane is narrow or absent. Tracheal hypoplasia (Table -1) primarily affects brachycephalic breeds, especially English bulldogs. Affected dogs may have other congenital abnormalities such as stenotic nares, elongated soft palate, and megaesophagus. They may have continuous respiratory distress, coughing, and recurrent tracheitis, or may have intermittent signs that are mild or moderate. This condition can typically be diagnosed on radiographs and must be differentiated from tracheal collapse. Treatment consists of symptomatic medical therapy (i.e., antibiotics, cough suppressants) and correction of other airway obstructions (e.g., resection of nares, palate, saccules); however, because the entire length of the trachea is involved, surgical correction is not feasible. Traumatic stenosis may occur following trauma or surgery. If only a segment of the trachea is involved, the narrowed area may be treated by balloon dilation or tracheal resection and anastomosis may be performed.

Diagnosis

Palpation of the trachea may reveal flaccid tracheal cartilages with prominent lateral borders and may elicit paroxysmal coughing. A soft end-expiratory noise may be auscultated in some dogs with intrathoracic tracheal collapse and probably represents the walls of the trachea snapping together during expiration. Abnormal heart sounds may be associated with concurrent cardiac disease. Electrocardiography may reveal sinus arrhythmia or evidence of cor pulmonale or left ventricular enlargement.

The first step in diagnosing tracheal collapse is usually to perform inspiratory and expiratory lateral radiographs of the neck and thorax. Because the collapse is dynamic (the cervical trachea collapses on inspiration and the thoracic trachea on expiration), both inspiratory and expiratory films should be taken to evaluate the entire trachea. Radiographs are diagnostic in approximately 60% of patients with moderate to severe tracheal collapse. Fluoroscopy, though seldom available in veterinary practices, facilitates evaluation of the dynamic movement of the trachea and mainstem bronchi through all phases of respiration. Special attention should be paid to evaluating the mainstem bronchi because animals with mainstem bronhial collapse are unlikely to benefit from surgical repair of their collapsing cervical trachea. Presently, there is no clinically used method to stent collapsing mainstem bronchi; however, a recent publication investigated intraluminal stents for mainstem bronchial collapse and concluded that such a technique might be useful in affected dogs. In addition to evaluating the trachea on radiographs, thoracic radiographs should also be evaluated for cardiomegaly and pulmonary disease.

If tracheal collapse is suspected, but radiographs were non-diagnostic, tracheoscopy should be performed. It is also recommended as a procedure to evaluate the trachea prior to surgery in all dogs, irregardless of radiographic findings. Prior to performing tracheoscopy, laryngoscopy should be performed to rule out associated conditions causing upper airway disease. Laryngeal paresis, paralysis, or collapse is present in approximately 30% of dogs with tracheal collapse. Laryngoscopy must be performed under light anesthesia. During tracheoscopy, tracheal conformation should be evaluated as the scope is withdrawn to determine the location and severity of the collapse. The entire trachea is usually collapsed; however, one area of the trachea is often more severely affected and is used for classification purposes. Grade I tracheal collapse is a 25% reduction in lumen diameter with the trachealis muscle being slightly pendulous and the cartilages maintaining a somewhat circular shape (Table-2). Grade II collapse is a 50% reduction in lumen diameter with the trachealis muscle stretched and pendulous and the cartilages beginning to flatten. Grade III collapse is defined as a 75% reduction in lumen diameter with the trachealis more stretched and pendulous and the cartilages nearly flattened. In grade IV collapse the lumen is essentially obliterated; tracheal cartilages are completely flattened and may invert to contact the trachealis muscle. Tracheal cultures and brushings taken during tracheoscopy are useful in selecting antibiotics. Positive tracheobronchial cultures are found in more than 50% of animals with tracheal collapse.

Medical Treatment

Medical therapy is recommended for all animals with mild clinical signs and those with less than 50% collapse. Medical therapy for dogs with tracheal collapse (Table 3) includes antitussives (i.e., butorphanol tartrate, hydrocodone bitartrate), antibiotics (i.e., ampicillin, cefazolin, clindamycin, enrofloxacin), bronchodilators (i.e., aminophylline, oxtriphylline), and/or corticosteroids (i.e., dexamethasone, prednisone). Sedation with acepromazine (0.05 to 0.2 mg/kg [maximum 1 mg] intravenously, intramuscularly, or subcutaneously, TID) and/or diazepam (0.2 mg/kg intravenously BID) and supplemental oxygen may be required in severely dyspneic patients. Weight reduction should be instituted for obese patients. Exercise restriction is recommended. Affected dogs should be maintained in an environment free of smoke and other respiratory irritants or allergens. Response to medical therapy is usually transient and the disease typically progresses.

Surgical Treatment

Surgery is recommended for all dogs with moderate to severe clinical signs, a 50% or greater reduction of the tracheal lumen (without significant mainstem bronchial collapse; see above), and those refractory to medical therapy. Dogs presenting with laryngeal paralysis or collapse, generalized cardiomegaly, and chronic pulmonary disease are poor surgical candidates. Coughing and dyspnea caused by laryngeal, pulmonary, or cardiac disease are unlikely to improve without appropriate therapy. Respiratory distress and death may occur in animals with severe laryngeal dysfunction or bronchopulmonary disease.

The goal of surgery is to support the tracheal cartilages and trachealis muscle, while preserving as much of the segmental blood and nerve supply to the trachea as possible. Many techniques have been described. Currently, the only techniques that meet this goal are placement of individual rings or modified spiral ring prostheses (Figure 1). Generally only the cervical trachea and most proximal portion of the thoracic trachea are supported, even when cervical and thoracic tracheal collapse are present. Readers are referred to surgical textbooks for descriptions of tracheoplasty using individual ring prostheses. Patients with concurrent laryngeal paralysis or laryngeal collapse may also require arytenoid lateralization or permanent tracheostomy, respectively.

The main complication of tracheoplasty is laryngeal paralysis. The segmental blood and nerve supply to the trachea travels in the lateral pedicles on each side of the trachea. The left recurrent laryngeal nerve is located in the lateral pedicle; the right is sometimes located within the carotid sheath.

STENOTIC NARES

Stenotic nares (congenital malformations of the nasal cartilages) are commonly seen in brachycephalic breeds. The cartilages lack normal rigidity and collapse medially causing partial occlusion of the external nares. Airflow into the nasal cavity is restricted and greater inspiratory effort is necessary, causing mild to severe dyspnea. Concurrent soft palate elongation, everted laryngeal saccules, aryepiglottic collapse, and/or corniculate collapse often contribute to the severity of respiratory distress.

"Brachycephalic syndrome" refers to the combination of stenotic nares, soft palate elongation, and laryngeal saccule eversion which is commonly seen in brachycephalic dogs. Concurrent tracheal hypoplasia or advanced laryngeal collapse often contributes to the respiratory distress. Brachycephalic animals exhibit signs of upper airway obstruction due to anatomic and functional abnormalities. They typically have a compressed face with poorly developed nares and a distorted nasopharynx. Their head shape is the result of an inherited developmental defect in the bones of the base of the skull. These bones grow to a normal width but reduced length. The soft tissues of the head are not proportionally reduced and often appear redundant.

Surgical Technique

Resection of a portion of the dorsal lateral nasal cartilage may be performed to widen the nares. Other techniques described include resection of horizontal or lateral tissue wedges. Grasp the margin of the nares with a Brown-Adson thumb forceps (Figure 2). Maintaining this grip, make a "V"-shaped incision around the forceps with a #11 scalpel blade. Make the first incision medially and the second incision laterally. Remove the vertical wedge of tissue. Control hemorrhage with pressure and by reapposing the cut edges. Align the ventral margin of the nares and mucocutaneous junction and place 3 to 4 simple interrupted sutures (e.g., polydioxanone, 3-0 or 4-0) to reappose the tissues. Repeat the procedure on the opposite side being careful to excise the same size tissue wedge.

ELONGATED SOFT PALATE

Elongated soft palate is the most commonly diagnosed respiratory problem in brachycephalic dogs. It is a part of the "brachycephalic syndrome" (in addition to stenotic nares and laryngeal saccule eversion). It is a congenital abnormality. The elongated soft palate is pulled caudally during inspiration, obstructing the dorsal aspect of the glottis. It is sometimes sucked between the corniculate processes of the arytenoid cartilages which increases inspiratory effort and causes more turbulent airflow. The laryngeal mucosa becomes inflamed and edematous further narrowing the airway. The tip of the soft palate is blown into the nasopharynx during expiration. Affected dogs may have trouble swallowing because normal occlusion of the airway during deglutition compromises ventilation.

Resection may be done with scissors, carbon dioxide laser, or electrosurgery, although the latter may increase postoperative swelling. Hemorrhage is generally mild to moderate following resection and can be controlled with gentle pressure. The caudal margin of the soft palate should be shortened so that it contacts the tip of the epiglottis. Resection of too little soft palate will not optimally relieve respiratory distress, while resection of too much soft palate results in nasal regurgitation, rhinitis, and sinusitis. Visually mark the site of proposed resection using the tip of the epiglottis and the caudal or midpoint of the tonsils as landmarks. Handle the soft palate gently and as little as possible to avoid excessive mucosal swelling. Grasp the tip of the soft palate with thumb forceps or Allis tissue forceps. Place stay sutures at the proposed site of resection on the right and left borders of the palate. Place hemostats on these sutures and have an assistant apply lateral traction. Transect across one-third to one-half the width of the soft palate with curved Metzenbaum scissors. Begin a simple continuous suture pattern (4-0 polydioxanone) at the border of the palate apposing the oropharyngeal and nasopharyngeal mucosa. Continue transecting and suturing until the excess palate has been resected.

LARYNGEAL COLLAPSE

Laryngeal collapse occurs secondary to chronic upper airway obstruction or trauma. Trauma may fracture or disrupt the laryngeal cartilages and allow medial collapse. Laryngeal collapse due to chronic upper airway obstruction and cartilage fatigue or degeneration is most common. The obstruction causes increased airway resistance, increased negative intraglottic luminal pressure, and increased air velocity. These forces displace laryngeal structures medially with permanent cartilage deformation and fatigues the cartilages. Increased inspiratory effort irritates the mucosa causing inflammation and edema. This further obstructs the airway causing more airflow resistance and increasing the effort of breathing.

Laryngeal collapse is described in three stages. Stage 1 is commonly referred to as laryngeal saccule eversion. Medial deviation of the cuneiform cartilage and aryepiglottic fold or aryepiglottic collapse is Stage 2 collapse, while Stage 3 collapse is medial deviation of the corniculate process of the arytenoid cartilages or corniculate collapse. Stages 2 and 3 are advanced stages of laryngeal collapse. Diagnosis of laryngeal collapse that occurs concurrently with other upper respiratory abnormalities (i.e., elongated soft palate, stenotic nares) is easily overlooked on oral examination. If response to treatment is less than expected following appropriate surgery for these abnormalities, laryngeal collapse may be present.

LARYNGEAL PARALYSIS

Laryngeal paralysis causes upper respiratory obstruction and mild to severe dyspnea. Because of dysfunction of the laryngeal muscles, recurrent laryngeal or vagus nerves, or cricoarytenoid ankylosis, acquired or congenital neurologic causes are most common. The intrinsic laryngeal abductor and adductor muscles are innervated by the recurrent laryngeal nerves. Subsequent atrophy of the cricoarytenoideus dorsalis muscle causes the cartilages to remain in a paramedian position during inspiration, preventing maximal air intake and increasing airflow resistance. Ineffective laryngeal adduction and closure during swallowing predisposes the patient to aspiration of food and secretions causing subsequent aspiration pneumonia.

Congenital, inherited laryngeal paralysis occurs in the Bouvier de Flanders, Bull Terriers, Siberian Huskies, and Dalmatians. Laryngeal paralysis in Bouvier's is due to degeneration of the nucleus ambiguous. Polyneuropathy associated with dying-back of peripheral nerves causes laryngeal paralysis in Dalmatians. Acquired laryngeal paralysis is usually idiopathic but may occur secondary to trauma, disease (e.g., polyneuropathy, myopathy, Chagas disease [Trypanosomiasis], hypothyroidism, neoplasia), or may be iatrogenic following surgery. It affects one or both sides of the larynx; however, unilateral paralysis is often asymptomatic.

Surgical treatment is recommended for patients with laryngeal paralysis who have moderate to severe signs of respiratory distress. The goal of treatment is to enlarge the glottis without exaggerating aspiration of food or saliva. Many surgical techniques have been described to treat laryngeal paralysis including partial laryngectomy, lateralization, castellated laryngofissures, and muscle nerve pedicle transposition. Vocal fold excision enlarges the ventral aspect of the glottis, is effective in mild-moderate cases, and is relatively easy to perform. However, glottic stenosis occurs in approximately 20% of the cases and is difficult to treat successfully. Partial arytenoidectomy (corniculate process) enlarges the dorsal aspect of the glottis. Its success is dependent on the skill of the surgeon. Serious complications and death occur in up to 50% of the cases following partial arytenoidectomy. The modified castellated laryngofissure technique is a combination of vocal fold excision, lateralization and laryngofissure creation to enlarge the glottis. This technique is effective but technically difficult. Muscle-nerve pedicle transposition can successfully reinnervate the larynx and improve function, but the process takes 5 to 11 months before clinical improvement is seen. Arytenoid lateralization is recommended as it gives consistently good results (> 90%) with few complications.

Unilateral Arytenoid Lateralization

Make a skin incision just ventral to the jugular vein beginning at the caudal angle of the mandible and extending over the dorsolateral aspect of the larynx to 1 to 2 cm caudal to the larynx (Figure 3). Incise and retract subcutaneous tissues, platysma, and parotidoauricularis muscles. Retract the sternocephalicus muscle and jugular vein dorsally and the sternohyoid muscle ventrally to expose the laryngeal area. Palpate the dorsal margin of the thyroid cartilage. Incise the thyropharyngeus muscle along the dorsolateral margin of the thyroid cartilage lamina. Place a stay suture through the thyroid cartilage lamina to retract and rotate the larynx laterally. Identify the cricoarytenoideus dorsalis muscle. Disarticulate the cricothyroid articulation with a # 11 blade or scissors. Palpate, identify, and disarticulate the cricoarytenoid articulation at the muscular process. Using curved Metzenbaum scissors transect the sesamoid band (interarytenoid ligament) between the two corniculate processes being careful not to penetrate the laryngeal mucosa. Place a polypropylene suture (2-0 to #2) through the muscular process of the arytenoid cartilage and the caudal one-third of the cricoid cartilage near the dorsal midline to mimic the direction of the cricoarytenoid muscle. Alternatively, place the suture through the muscular process and the most caudodorsal aspect of the thyroid cartilage. Muscular process-to-thyroid cartilage sutures tend to pull the arytenoid laterally while muscular process-to-cricoid cartilage sutures tend to rotate the arytenoid laterally. Tie the suture with enough tension to moderately abduct the arytenoid cartilage. Have an assistant verify abduction by intraoral visualization of the larynx. If abduction is insufficient the suture can be repositioned to achieve better abduction. Lavage the surgical site. Appose the thyropharyngeus muscle with a cruciate or simple continuous pattern (3-0 polydioxanone). Appose subcutaneous tissues and skin routinely.

Partial Laryngectomy

Partial laryngectomy may be done via an oral approach or a ventral laryngotomy approach. Vocal fold resection and unilateral resection of the corniculate, cuneiform, and vocal processes of the arytenoid cartilage should be performed. Partial laryngectomy via an oral approach is extremely difficult in small dogs because of limited exposure.

Via an oral approach - Grasp the corniculate process and retract it medial with biopsy forceps. Use a long-handled scalpel or scissors to excise the corniculate process and the proximal half and base of the cuneiform process. Do not excise the aryepiglottic fold or the distal half of the cuneiform process. Remove the vocal fold, vocal process, and vocal muscle with biopsy forceps and/or Metzenbaum scissors. Control bleeding by applying pressure with gauze sponges.

Via a laryngotomy approach - Make a ventral midline incision over the larynx. Separate the sternohyoid muscles and incise the cricothyroid membrane and thyroid cartilage on the midline. Retract the edges of the thyroid cartilage with small Gelpi forceps. Visualize the arytenoid cartilages and vocal folds. Have an assistant visualize the larynx per os to help determine how much to remove. After incising the mucosa over the corniculate, cuneiform, and vocal processes of ONE arytenoid cartilage, excise them with scissors or a scalpel. Also excise the vocal fold on that side (if necessary excise the vocal cord and process on the opposite side). Excise redundant mucosa and suture the defect with 4-0 to 6-0 absorbable suture material in a continuous pattern. Suture the thyroid cartilage with interrupted sutures that do not penetrate the laryngeal lumen. Close subcutaneous tissues and skin routinely.

Intestinal biopsy is indicated to diagnose intestinal diseases that have not been defined by other tests. The small intestine may be biopsied during endoscopy, ultrasonography, or laparotomy using general anesthesia (endoscopy, laparotomy) or sedation (ultrasonography). Full-thickness intestinal biopsies are performed in conjunction with a complete abdominal exploratory.

Exteriorize and isolate the diseased or desired intestine from the abdomen by packing with towels or laparotomy sponges. Gently milk intestinal contents from the lumen of the identified intestinal segment. Occlude the lumen at both ends of the isolated segment to minimize spillage of chyme by having an assistant use a "scissor-like" grip with the index and middle fingers a distance of 4 to 6 cm on each side of the proposed enterotomy site. Make a full-thickness stab incision into the intestinal lumen on the antimesenteric border with a #11 scalpel blade. Obtain full-thickness biopsies 2 to 3 mm wide by either making a second longitudinal incision parallel to the first with the scalpel blade or remove an ellipse of intestinal wall at one margin of the incision with Metzenbaum scissors. To help prevent curling of the specimen, place the biopsy on a piece of sterile paper, serosal side down. Following biopsy, if necessary, trim everted mucosa so that it is even with the serosal edge. Suction the isolated lumen. Close the incision with gentle appositional force using simple interrupted sutures. Place sutures through all layers of the intestinal wall, 2 mm from the edge and 2 to 3 mm apart, with extraluminal knots. Angle the needle so the serosa is engaged slightly further from the edge than the mucosa. This helps reposition everting mucosa within the lumen. Tie each suture carefully without cutting through layers of the intestinal wall so as to gently appose all intestinal layers without crushing the tissue. Transverse closure of a longitudinal incision helps maintain adequate lumen size in patients with very small intestine. Simple continuous or crushing sutures may also be used. Use a monofilament absorbable suture material (4-0 or 3-0 polydioxanone or polyglyconate) with a swaged on taper or tapercut point needle. While maintaining luminal occlusion near the enterotomy site, moderately distend the lumen with sterile saline, apply gentle digital pressure, and observe for leakage between sutures or through needle holes. Place additional sutures if leakage occurs between sutures. Lavage the isolated intestine and the entire abdomen if contamination has occurred. Place omentum over the suture line prior to abdominal closure. If intestinal integrity is questionable or leakage occurs from needle holes, use a serosal patch rather than omentum. Replace contaminated instruments and gloves prior to abdominal closure.

INTESTINAL RESECTION AND ANASTOMOSIS

Intestinal resection and anastomosis are recommended for removing ischemic, necrotic, neoplastic, or fungalinfected segments of intestine. Irreducible intussusceptions are also managed by resection and anastomosis. Endtoend anastomoses are recommended.

Make an abdominal incision which allow complete exploration of the abdomen. Perform a thorough examination of the abdomen and collect any nonintestinal specimens. Exteriorize and isolate the diseased intestine from the abdomen by packing with towels or laparotomy sponges. Assess intestinal viability and determine the amount of intestine needing resection. Double ligate and transect the arcadial mesenteric vessels from the cranial mesenteric artery that supplies this segment of intestine. Double ligate the terminal arcade vessels and vasa recta vessels within the mesenteric fat at the points of proposed intestinal transection. Gently milk chyme (intestinal contents) from the lumen of the identified intestinal segment. Occlude the lumen at both ends of the segment to minimize spillage of chyme. Place forceps across each end of the diseased bowel segment. Transect the intestine with either a scalpel blade or Metzenbaum scissors along the outside of the forceps. Make an oblique incision across the intestine if the luminal diameters are the same. When two pieces of intestine are being joined that are of unequal size, use a perpendicular incision across the intestine with the larger luminal diameter and an oblique incision (45- to 60- degree angle) across the intestine with the smaller luminal diameter to help correct size disparity). Make the oblique incision such that the antimesenteric border is shorter than the mesenteric border. Suction the intestinal ends and remove any debris clinging to the cut edges with a moistened gauze sponge. Trim everting mucosa with Metzenbaum scissors just before beginning the end-to-end anastomosis.

Use 3-0 or 4-0 monofilament, absorbable suture (polydioxanone or polyglyconate) with a swaged-on taper or tapercut point needle. Place simple interrupted sutures through all layers of the intestinal wall. Angle the needle so the serosa is engaged slightly further from the edge than the mucosa. This helps reposition everting mucosa within the lumen. Tie each suture carefully so as to gently appose the edges of the intestine with the knots extraluminally. Appose intestinal ends by first placing a simple interrupted suture at the mesenteric border and then placing a second suture at the antimesenteric border approximately 180 degrees from the first (this divides the suture line into equal halves and allows determination of whether the ends are of approximately equal diameter). The mesenteric suture is the most difficult suture to place in the anastomosis because of mesenteric fat. It is also the most common site of leakage. If the ends are of equal diameter, space additional sutures between the first two sutures approximately 2 mm from the edge and 2 to 3 mm apart. If minor disparity still exists between lumen sizes, space the sutures around the larger lumen slightly further apart than the sutures in the intestine with the smaller lumen. To correct luminal disparity that cannot be accommodated by the angle of the incisions or by suture spacing, resect a small wedge (1 to 2 cm long and 1 to 3 mm wide) from the antimesenteric border of the intestine with the smaller lumen. After suture placement inspect the anastomosis and check for leakage. While maintaining luminal occlusion adjacent to the anastomotic site, moderately distend the lumen with sterile saline, apply gentle digital pressure, and observe for leakage between sutures or through needle holes. Close the mesenteric defect with a simple continuous or interrupted suture pattern (4-0 polydioxanone or polyglyconate) being careful not to penetrate or traumatize arcadial vessels near the defect (see Figure 1).

SEROSAL PATCHING

Serosal patching is placement of an antimesenteric border of the small intestine over a suture line or organ defect and securing it with sutures. Serosal patching serves to provide support, a fibrin seal, resistance to leakage, blood supply to the damaged area, and may prevent intussusception. Patches are commonly used following intestinal surgery when closure integrity is questioned or dehiscence is repaired. Patches which span visceral defects are covered with mucosal epithelium within 8 weeks. Most commonly, jejunum adjacent to the defect or area of questionable viability is used for the serosal patch, although other sources could include stomach, other intestinal segments, or urinary bladder.

Use one or more loops of intestine to form the patch. Use gentle loops to avoid stretching, twisting, or kinking the intestine and mesenteric vessels. If using more than one loop of intestine, suture these loops together before securing the patch to the damaged area. All sutures used to create or secure the patch engage the submucosa, muscularis, and serosa; they should not penetrate the intestinal lumen. Place interrupted or continuous sutures in healthy tissue to secure the patch and isolate the damaged area.

BOWEL PLICATION

Bowel plication is performed to prevent recurrence of intussusception; without plication the incidence of repeat intussusception in dogs is relatively high. Serosa-to-serosa adhesions are formed by suturing adjacent loops of intestine together. The small intestine from the duodenocolic ligament to the ileocolic junction should be sutured to decrease the potential for intestinal strangulation. Place small intestinal loops side-by-side to form a series of gentle loops from the distal duodenum to the distal ileum. Secure the loops by placing sutures which engage the submucosa, muscularis, and serosa, 6 to 10 cm apart. Use 3-0 or 4-0 monofilament absorbable or non-absorbable sutures with a taper point swaged-on needle. Avoid positioning the intestinal loops at acute angles lest intestinal obstruction occurs. The adhesions formed by this procedure are probably not permanent adhesions, but they are sufficient to prevent recurrent intussuception.

COLOPEXY

Colopexy is done to prevent caudal movement of the colon and rectum and is especially useful in animals with recurrent rectal prolopse. The procedure creates permanent adhesions between the serosal surfaces of the colon and abdominal wall. Incisional and non-incisional techniques have been described and both are equally effective. A potential complication (but rare if the technique is performed properly) is infection as a result of suture penetration into the colonic lumen.

Expose and explore the abdomen. Locate and isolate the descending colon from the remainder of the abdomen. Pull the descending colon cranially to reduce the prolapse. Verify prolapse reduction by having a non-sterile assistant inspect the anus visually and perform a rectal examination. Make a 3 to 5 cm longitudinal incision along the antimesenteric border of the distal descending colon through only the serosal and muscularis layers. Create a similar incision on the left abdominal wall several centimeters lateral (( 2.5 cm) to the linea alba through the peritoneum and underlying muscle. Appose each edge of the colonic and abdominal wall incisions with 2 simple continuous or simple interrupted rows of sutures using 2-0 or 3-0 monofilament absorbable (e.g., polydioxanone or polyglyconate) or nonabsorbable (nylon, polypropylene) suture material. Engage the submucosa as each suture is placed. Lavage the surgical site and surround it with omentum prior to abdominal closure. Alternatively, scarify an 8 to 10 cm antimesenteric segment of the descending colon by scraping the serosa with a scalpel blade or rubbing it with a gauze sponge. On the left abdominal wall opposite the prepared colon, scarify the peritoneum in the same manner. Preplace, then tie, 6 to 8 horizontal mattress sutures between the two scarified surfaces. Roll the colon toward the midline and place a second row of 6 to 8 sutures. Use 2-0 to 3-0 monofilament absorbable or nonabsorbable sutures which engage the submucosa, but do not penetrate the colonic mucosa. Tie the sutures apposing the scarified surfaces (see Figure 2).

EXPLORATORY LAPAROTOMY

The decision to operate is based on historical and physical examination findings, radiographic/ultrasonographic studies, and/or laboratory analyses. Physical examination can be unreliable in predicting the severity of abdominal trauma. The inaccuracy associated with examining patients with acute abdominal disease (particularly that associated with trauma) may be attributed in part to the patient's condition at the time of examination and delayed development of clinical signs associated with some injuries. Depressed or lethargic animals may not exhibit pain during abdominal palpation. Clinical signs of hemorrhage often are not apparent immediately after trauma; delays of 3 to 4 hours between trauma and development of shock and collapse are common in patients with liver or spleen lacerations. Thus traumatized animals should be closely observed for at least 8 to 12 hours. In most instances lifethreatening hemorrhage will become apparent before this time. However, animals with traumatic bile peritonitis often are without clinical signs for several weeks. Likewise, traumatic mesenteric avulsion is seldom associated with clinical signs until subsequent peritonitis develops (usually several days after trauma occurs). Sensitive diagnostic tests such as diagnostic peritoneal lavage may facilitate identification of patients with significant abdominal trauma, before the development of overt clinical signs.

Preoperative management of most animals undergoing exploratory laparotomy is dictated by their underlying abdominal disease. General observations include noting the animal's attitude and posture, temperature, respiratory rate and effort, and heart rate and rhythm. Additionally, auscultation, percussion, and palpation of the abdomen, plus a rectal examination, should be performed. Serial examinations are important to detect trends or deterioration in patient status. An intravenous catheter should be placed for fluid and drug administration, and blood samples should be drawn. Useful initial blood work in an animal with acute abdomen includes hematocrit, serum total protein, serum glucose concentrations, complete blood count (CBC), platelet count, and blood urea nitrogen. Other laboratory tests (i.e., serum biochemistry profile, clotting parameters) can be performed, depending on the animal's condition and suspected underlying disease. Urine may be collected via cystocentesis or catheterization for urinalysis. An indwelling urinary catheter may be used to quantitate urinary output if necessary. Abdominal radiographs may detect peritoneal fluid (i.e., uroabdomen, peritonitis) or abnormal accumulations of air. Surgery is warranted if free air is present in the abdominal cavity because this usually indicates rupture or perforation of the gastrointestinal tract. Animals with acute abdominal signs of uncertain cause should have diagnostic peritoneal lavage performed if radiographs are nondiagnostic. Electrolyte and hydration abnormalities should be corrected before surgery.

SURGICAL TECHNIQUES

Systematically explore the entire abdomen. Various techniques may be used; however, every surgeon should develop a consistent pattern to ensure that the entire abdominal cavity and all structures are visualized and/or palpated in each animal. Use moistened laparotomy sponges to protect tissues from drying during the procedure. If generalized infection is present or if diffuse intraoperative contamination has occurred, flush the abdomen with copious amounts of warmed, sterile saline solution. Historically, many different antiseptics (i.e., povidone-iodine, chlorhexidine) and antibiotics have been added to lavage fluids. Povidone-iodine is the most widely used antiseptic; however, it has not shown a beneficial effect in repeated experimental and clinical trials and may be detrimental in animals with established peritonitis because the carrier, polyvinylpyrrolidone, inhibits macrophage chemotaxis. Similarly, there is no substantial evidence that adding antibiotics to lavage fluid benefits patients treated with appropriate systemic antibiotics. Remove lavage fluid and blood and inspect the abdominal cavity before abdominal closure to ensure that all foreign material and surgical equipment have been removed. Perform a sponge count and compare it with the preoperative count to ensure that surgical sponges have not been left in the abdominal cavity.

ABDOMINAL WALL CLOSURE

Close the linea alba with simple interrupted sutures or a simple continuous suture pattern. A simple continuous suture pattern does not increase the risk of dehiscence when properly performed (i.e., secure knots, appropriate suture material), and it allows for rapid closure. Preferably strong, absorbable suture material (i.e., polydioxanone or polyglyconate) should be used for continuous suture patterns and six to eight knots placed at each end of the incision line. Monofilament, nonabsorbable suture material (i.e., nylon, polypropylene) has been associated with suture sinus formation in dogs and should be avoided. Surgical gut and stainless steel wire should not be used for continuous suture patterns. On each side of the incision incorporate 4 to 10 mm of fascia in each suture. Place interrupted sutures 5 to 10 mm apart, depending on the animals' size. Tighten sutures sufficiently to appose, but not strangulate, tissue because the latter will adversely affect wound healing. Incorporate full-thickness bites of the abdominal wall in the sutures if the incision is midline (i.e., through the linea alba). Do not incorporate the falciform ligament between the fascial edges. If the incision is lateral to the linea alba and muscular tissue is exposed (i.e., paramedian), close the external rectus sheath without including muscle in the sutures. Do not attempt to include peritoneum in the sutures. Close subcutaneous tissues with a simple continuous pattern of absorbable suture material and reappose preputialis muscle fibers. Use nonabsorbable sutures (simple interrupted or continuous appositional patterns) or stainless steel staples to close skin. Place skin sutures without tension.

POSTOPERATIVE CARE AND ASSESSMENT

The abdominal incision should be checked twice daily for evidence of redness, swelling, or discharge. If the animal licks or chews at the incision, an Elizabethan collar or sidebar should be used to prevent iatrogenic suture removal. Early signs of altered wound healing are inflammation and edema. Serosanguineous drainage from the incision and swelling are consistent signs of acute incisional dehiscence. Dehiscence usually occurs 3 to 5 days postoperatively when minimal healing has occurred and the sutures have weakened; however, it may occur earlier if knots were tied improperly or if fascia was not incorporated into the sutures. Evisceration usually results in sepsis and severe blood loss secondary to mutilation of exposed intestine and must be treated promptly. The abdomen should be bandaged, fluid therapy initiated, and broad-spectrum antibiotics given while the animal is prepared for surgery. If technical failure is suspected (i.e., poor knot tieing, improper suturing), the entire suture line should be removed and replaced. Debridement of the wound edges is not necessary and will delay wound healing. The intestine should be closely inspected for viability and damaged sections resected if appropriate. The abdominal cavity should be lavaged with copious amounts of warmed, sterile saline. Open abdominal drainage should be considered in animals with generalized peritonitis. Wound disruption after 10 to 21 days usually results in hernia formation rather than evisceration. Hernial repair in these animals may require excision of fibrotic tissues. Subsequent closure requires that tissue layers be accurately apposed.

General Considerations and Clinically Relevant Pathophysiology

Secondary generalized peritonitis is the predominant form of peritonitis in dogs and is usually caused by bacteria. Primary generalized peritonitis occurs in cats associated with feline infectious peritonitis. Generalized peritonitis may result from intestinal or gall-bladder perforation, rupture, or necrosis (e.g., gastric or intestinal foreign bodies, intussusception, mesenteric avulsion, gastric dilatation-volvulus, or necrotizing cholecystitis), pancreatic abscessation, prostatic abscesses, or foreign body penetration.

Diagnosis

Signalment. Any age, sex, or breed of dog or cat may develop peritonitis. It is particularly common in young animals that have perforating foreign bodies and in those that receive abdominal trauma (i.e., vehicular trauma and bite wounds).

History. The history is often nonspecific. The animal may not show signs of illness for several days after the traumatic episode. Mesenteric avulsions often do not cause clinical signs of peritonitis for 5 to 7 days after the injury. Animals with traumatic bile peritonitis may be asymptomatic for several weeks after the injury. Most animals are presented for lethargy, anorexia, vomiting, diarrhea, and/or abdominal pain.

Physical Examination Findings

Affected animals are usually painful on abdominal palpation. The pain may be localized but generalized pain is more common and the animal will often tense or "splint" the abdomen during palpation. Vomiting and diarrhea may be noted. Abdominal distension may be noted if sufficient fluid has accumulated. Pale mucous membranes, prolonged capillary refill times, and tachycardia may indicate that the animal is in shock. Dehydration and arrhythmias may also occur.

Radiography/Ultrasonography

The classic radiographic finding in animals with peritonitis is loss of abdominal detail with a focal or generalized "ground-glass" appearance. The intestinal tract may be dilated with air and/or fluid. Free air in the abdomen may be noted with rupture of a hollow organ or sometimes occurs with gas-producing anaerobes, without gut rupture. A more localized peritonitis may occur secondary to pancreatitis and cause the duodenum to appear fixed and elevated. Ultrasonography is useful to localize fluid accumulation and help determine etiology.

Laboratory Findings

The most common laboratory finding in animals with peritonitis is a marked leukocytosis. The predominant cell type is the neutrophil and a left shift is often apparent. Other abnormalities may include anemia, dehydration, and electrolyte and acid-base abnormalities.

Abdominocentesis should be performed (see below) and fluid retrieved for analysis. Inflammatory fluids should have an elevated number of neutrophils, which may appear degenerative. Significant numbers of leukocytes accumulate in the peritoneal cavity within 2 to 3 hours of contamination with blood, bile, urine, feces, or gastric or pancreatic secretions. Leukocyte counts in abdominal fluid of normal dogs are usually < 500 cells/(l. Following peritoneal lavage (see below) in dogs, white blood cell counts of > 1000/(l are indicative of mild to moderate irritation, while counts > 2,000/(l indicate marked peritonitis. The presence of degenerate leukocytes and bacteria in the lavage fluid also suggest intraabdominal infection. However, the presence and number of WBCs should be correlated with other clinical findings when considering abdominal exploration. Elevated leukocyte counts are found in most dogs following abdominal surgery. In animals that have undergone recent surgery > 7,000 cells/(l and > 9,000 cells/(l indicates mild to moderate and marked peritonitis, respectively.

Following abdominocentesis, the amount of blood in the abdominal cavity can be estimated by observing the lavage sample. A red color reflects the presence of RBCs and a deep-red color usually indicates severe hemorrhage. If newsprint cannot be read through the plastic tubing, then hemorrhage is significant. If print can be seen through the tubing, only moderate or minimal hemorrhage is present. Surgical intervention is indicated when there is a substantial increase in the PCV of lavage samples taken within 5 to 20 minutes of each other, or if an animal in shock does not respond to aggressive fluid therapy.

Differential Diagnosis

Advanced peritonitis with significant accumulation of abdominal fluid is not difficult to diagnose. The difficulty usually arise in determining the etiology of the effusion or infection. Early peritonitis, prior to the onset of overt clinical signs, is difficult to diagnose and may require diagnostic peritoneal lavage (see below).

Medical Management

The goals of management of animals with peritonitis are to treat the cause of the contamination, resolve the infection, and restore normal fluid and electrolyte balances. Food should be withheld if the animal is vomiting. Fluid replacement therapy should be initiated as soon as possible, particularly if the animal is dehydrated or appears shocky (up to 90 ml/kg IV, based on the animal's condition). Hypokalemia and hyponatremia may be present and require intravenous supplementation. Hypoglycemia is common if the animal has septic shock (systemic inflammatory response syndrome) and glucose may need to be added to the fluids (i.e., 2.5 - 5% dextrose). Standard shock therapy (i.e., fluid replacement, antibiotics, ± soluble corticosteroids) should be initiated. If severe metabolic acidosis is present, bicarbonate therapy may be indicated.

Broad spectrum antibiotic therapy should be initiated as soon as the diagnosis is made. Ampicillin plus enrofloxacin is an effective combination against most bacteria responsible for peritonitis in dogs. However, amikacin plus clindamycin or amikacin plus metronidazole may be necessary if anaerobic infection is present. A second-generation cephalosporin (e.g., cefoxitin sodium may also be used if gram-negative plus anaerobic infection is suspected. If renal compromise is present in an animal with a resistant bacterial infection, imipenem may be considered. Initial antibiotic therapy should be altered based on results of aerobic and anaerobic culture results of lavage fluid (see below) or cultures obtained at surgery.

Low-dose heparin (50-100 units/kg; SC; BID) increases survival and significantly reduces abscess formation in experimental peritonitis. The inflammatory process in peritonitis is associated with an outpouring of fibrous exudate which causes intraabdominal loculation of bacteria. The loculated bacteria are protected from host defense mechanisms and antibiotics which may not be able to penetrate the fibrin clots. Although the exact mechanism of its beneficial effect is still unknown, there does not appear to be any doubt that heparin is indicated in patients with severe peritonitis. Heparin may also be incubated with plasma and given to animals in DIC (incubate 5 - 10 units/kg heparin with 1 unit fresh plasma for 30 minutes; 10 ml/kg IV).

Flunixin meglumine is beneficial in experimental peritonitis (1 mg/kg IV, once or twice if in septic shock). Dogs treated with banamine in addition to gentamicin sulfate and fluids had higher blood pressures, less hemoconcentration, less metabolic acidosis, and higher survival rates than dogs treated with gentamicin sulfate and fluids alone (Hardie, 1985). Banamine blocks thromboxane and prostacyclin production. It must, however, be used with caution in dogs due to the potential of GI and renal toxicity.

Surgical Treatment

Abdominocentesis is the percutaneous removal of fluid from the abdominal cavity, usually for diagnostic purposes, although it may occasionally be therapeutic. Indications include shock without apparent cause, undiagnosed disease with signs involving the abdominal cavity, suspicion of postoperative GI dehiscence, blunt or penetrating abdominal injuries (i.e., gunshot wounds, dog-bites, automobile accidents), and undiagnosed abdominal pain. A multi-fenestrated catheter should be used to enhance fluid collection. Physical and radiographic examinations should precede abdominocentesis to rule out instances where it may not be safe and to guide needle placement. Four-quadrant paracentesis may be performed if simple abdominocentesis is not successful in retrieving fluid. It is similar to simple abdominocentesis except that multiple abdominal sites are assessed by dividing the abdomen into four quadrants through the umbilicus and tapping each of these four areas. Diagnostic peritoneal lavage should be performed in animals with suspected peritonitis if the above methods are unsuccessful in obtaining fluid for analysis (see below).

Exploratory surgery is indicated when the cause of peritonitis cannot be determined or when bowel rupture, intestinal obstruction (e.g., bowel incarceration, neoplasia), or mesenteric avulsion is suspected. Serosal patching and plication are techniques which decrease the incidence of intestinal leakage, dehiscence, or repeated intussusception. Animals requiring surgery that have peritonitis secondary to intestinal trauma (disruption of mesenteric blood supply, bowel perforation, chronic intussusception, or foreign body) are frequently hypoproteinemic. The role that protein levels play in healing of intestinal incisions is not well understood. However, most surgeons are concerned that hypoproteinemic patients may not heal as quickly as patients with normal protein levels, despite one study that showed similar complication rates between these group for animals undergoing intestinal surgery (Harvey, 1990). Most experimental evidence has shown that retardation of wound healing is not seen with moderate protein-depletion, but only with severe deficiencies (<1.5 - 2 g/dl).

Although whether or not one should lavage the abdominal cavity of animals with peritonitis is controversial, lavage is generally indicated with diffuse peritonitis. Lavage should be done with care in animals with localized peritonitis to prevent causing diffuse dissemination of infection. When lavage is performed, as much of the fluid as possible should be removed because fluid inhibits the bodies ability to fight off infection, probably by inhibiting neutrophil function. Historically, many different agents have been added to lavage fluids, especially antiseptics and antibiotics. Povidone-iodine is the most widely added antiseptic; however, its use may be contraindicated in established peritonitis. Furthermore, no beneficial effect of this agent has been shown in repeated experimental and clinical trials. Although a great many antibiotics have been added to lavage fluids over the years, there is no substantial evidence that their addition is of any benefit to patients who are being treated with appropriate systemic antibiotics. Warmed sterile saline is the most appropriate lavage fluid.

Open abdominal drainage (OAD) is a useful technique for managing animals with peritonitis. Reported advantages are improvement in the patient's metabolic condition secondary to improved drainage, reduced abdominal adhesion and abscess formation, and access for repeated inspection and exploration of the abdomen. With this technique, the abdomen is left open and sterile wraps are placed around the wound. The frequency of the wrap changes is dependent upon the amount of fluid being drained and the amount of external soiling. Experimentally, dogs with peritonitis treated by OAD recover faster than those treated with closed abdomens. Peritoneal bacterial numbers are significantly less in OAD dogs when compared with control dogs and at necropsy there are fewer abdominal adhesions and less peritoneal fluid in the former group. Complications of open abdominal drainage include persistent fluid loss, hypoalbuminemia, weight loss, adhesions of abdominal viscera to the bandage, and contamination of the peritoneal cavity with cutaneous organisms.

Techniques

Abdominocentesis

Insert an 18 or 20-gauge, 1-1/2 inch plastic over-the-needle catheter (with added side holes) into the abdominal cavity at the most dependent part of the abdomen. Do not attach a syringe, instead allow the fluid to drip from the needle and collect it in a sterile tube. If sufficient fluid is obtained, place the fluid in a clot tube, and EDTA tube, submit samples for aerobic and anaerobic culture, and make 4 to 6 smears for analysis. If fluid is not obtained, apply gentle suction using a 3cc syringe. It is difficult to puncture bowel by this method since mobile loops of bowel move away from the tip of the needle as it strikes them. Perforations created by a needle this size usually heal without complications. The major disadvantage of needle paracentesis is that it is insensitive to the presence of small volumes of intraperitoneal fluid and hence a negative result can be meaningless. At least 5 or 6 ml of fluid/kg body weight must be present in the abdominal cavity of dogs to obtain positive results in a majority of cases using this technique.

Diagnostic Peritoneal Lavage

Make a 2-cm skin incision just caudal to the umbilicus and ligate any bleeders to avoid false positive results. Spread loose subcutaneous tissues and make a small incision in the linea alba. Hold the edges of the incision with forceps while the peritoneal lavage catheter (without the trocar) is inserted into the abdominal cavity. Direct the catheter caudally into the pelvis. With the catheter in place, apply gentle suction. If blood or fluid cannot be aspirated, connect the catheter to a bottle of warm sterile saline and infuse 22 ml/kg of fluid into the abdominal cavity. When the calculated volume of fluid has been delivered, roll the patient gently from side to side, place the bottle on the floor, vent it, and collect the fluid by gravity drainage. Do not attempt to remove all the fluid.

Exploratory Laparotomy

Perform a ventral midline incision from the xiphoid to the pubis. Obtain a sample of fluid for culture and analysis. Explore and inspect the entire abdomen. Find the source of infection and correct it. Break down adhesions that may hinder drainage. Lavage the abdomen with copious amounts of warm sterile saline if the infection is generalized. Remove as much necrotic debris and fluid as possible. Close the abdomen routinely or perform open abdominal drainage.

Open Abdominal Drainage

After completing the abdominal procedure, leave a portion of the abdominal incision (usually the most dependent portion) open to drain. Close the cranial and caudal aspects of the incision with monofilament suture using a continuous suture pattern. Place a sterile laparotomy pad over the opening, then place a sterile wrap over the laparotomy pad. Change the wrap at least twice daily initially with the animal standing (sedation is seldom necessary). Break down adhesions to the incision that may interfere with drainage. Abdominal lavage may be attempted, but is seldom necessary. Place a diaper over the wrap to decrease contamination from urine. Assess the fluid daily for bacterial numbers and cell morphology. When bacterial numbers have decreased and normal neutrophil morphology is present (non-degenerative), close the incision (generally in 3 to 5 days). If the opening is small it may be left to heal by second intention.

Classically, the GDV syndrome is an acute condition with a mortality rate of 30% to 45% in treated animals. The gastric enlargement is thought to be associated with a functional or mechanical gastric outflow obstruction. The initiating cause of the outflow obstruction is unknown; however, once the stomach dilates, normal physiologic means of removing air (i.e., eructation, vomiting, and pyloric emptying) are hindered because the esophageal and pyloric portals are obstructed.. The stomach becomes enlarged as gas and/or fluid accumulate within the lumen. The gas probably comes from aerophagia, although bacterial fermentation of carbohydrates, diffusion from the bloodstream, and metabolic reactions may contribute. Normal gastric secretion and transudation of fluids into the gastric lumen as a result of venous congestion contribute to fluid accumulation. The cause of GDV is unknown, but exercise after ingestion of large meals of highly processed foods or water has been suggested to contribute. Epidemiologic studies have not supported a causal relationship between feeding soy-based or cereal-based dry dog food and GDV. Other contributing causes include an anatomic predisposition, ileus, trauma, primary gastric motility disorders, vomiting, and stress.

Generally, the stomach rotates in a clockwise direction when viewed from the surgeon's perspective (with the dog on its back and the clinician standing at the dog's side, facing cranially. The rotation may be 90 to 360 degrees, but is usually 220 to 270 degrees. The duodenum and pylorus move ventrally and to the left of midline and become displaced between the esophagus and stomach. The spleen is usually displaced to the right ventral side of the abdomen. Caudal vena cava and portal vein compression by the distended stomach decreases venous return and cardiac output, causing myocardial ischemia. Decreases in central venous pressure, stroke volume, mean arterial pressures, and cardiac output occur. Obstructive shock and inadequate tissue perfusion affect multiple organs, including the kidney, heart, pancreas, stomach, and small intestine. Cardiac arrhythmias occur in many dogs with GDV, particularly those with gastric necrosis. Arrhythmias may contribute to mortality and require appropriate monitoring and treatment Myocardial depressant factor has also been recognized in affected dogs. Reperfusion injury has been implicated as causing much of the tissue damage that ultimately results in death after correction of GDV. Lazaroids, which inhibit lipid peroxidation, appear to decrease reperfusion injury and may eventually increase survival.

Partial or chronic gastric dilatation-volvulus may occur in dogs and is usually a progressive but non-life-threatening syndrome that may be associated with vomiting, anorexia, and/or weight loss. These dogs may have chronic, intermittent signs and appear normal between episodes. Gastric malpositioning may be intermittent or chronic but without dilatation. Plain or contrast radiographs are diagnostic, but repeat radiographs may be necessary if the stomach is intermittently malpositioned.

Diagnosis

Signalment. GDV primarily occurs in large, deep-chested breeds (i.e., Great Dane, Weimaraner, Saint Bernard, German Shepherd, Irish and Gordon Setters, Doberman Pinscher), but has been reported in cats and small breed dogs. Shar-peis may have an increased incidence compared with other medium-sized breeds. In a recent study, Basset Hounds had a high risk of GDV, despite their relatively small size (Glickman et al, 1994). Large breed size, degree of purity of breed, and increase of weight are significant risk factors for development of this disease. GDV may occur in any age dog, but is most common in middle-aged to older animals. Chest depth/width ratio appears to be highly correlated with the risk of bloat.

History. A dog with GDV may present with a history of a progressively distending and tympanic abdomen, or the owner may simply find the animal recumbent and depressed, with a distended abdomen. The dog may appear to be in pain and may have an arched back. Nonproductive retching, hypersalivation, and restlessness are common.

Physical Examination Findings

Abdominal palpation often reveals various degrees of abdominal tympany or enlargement; however, it may be difficult to feel gastric distension in heavily muscled large-breed or very obese dogs. Splenomegaly is occasionally palpated. Clinical signs associated with shock, including weak peripheral pulses, tachycardia, prolonged capillary refill time, pale mucous membranes, or dyspnea, may be present.

Radiography

Radiographic evaluation is necessary to differentiate simple dilatation from dilatation plus volvulus. Affected animals should be decompressed before radiographs are taken. Right lateral and dorsoventral radiographic views are preferred. In normal dogs, the pylorus is located ventral to the fundus on the lateral view, and on the right side of the abdomen on the dorsoventral view. On a right lateral view of a dog with GDV, the pylorus lies cranial to the body of the stomach and is separated from the rest of the stomach by soft tissue (reverse C sign). On the dorsoventral view, the pylorus appears as a gas-filled structure to the left of midline. Free abdominal air suggests gastric rupture and warrants immediate surgery.

Laboratory Findings

The CBC is seldom helpful unless DIC causes thrombocytopenia. Although normal or increased potassium concentrations may occur, hypokalemia is more common. Vascular stasis may cause increased lactic acid production, resulting in a metabolic acidosis. However, metabolic alkalosis caused by sequestration of hydrogen ions in the gastric lumen can offset the metabolic acidosis, causing the pH to be normal (i.e., a mixed acid-base disorder). Respiratory acidosis may be caused by hypoventilation secondary to gastric impingement on the diaphragm and decreased ventilatory compliance. Hence, routine use of sodium bicarbonate is inappropriate.

Medical Management

Patient stabilization is the initial objective. A large-bore intravenous catheter(s) should be placed in either a jugular or both cephalic veins. Either isotonic fluids (90 ml/kg/hr), hypertonic 7% saline (4-5 ml/kg over 5 to 15 min), or hetastarch (5-10 ml/kg over 10 to 15 min) is administered. If hypertonic saline or hetastarch is given, adjustment of the rate of subsequent crystalloid administration is necessary. Blood should be drawn for blood gas analyses, a CBC, and a biochemical panel. Broad-spectrum antibiotics (e.g., cefazolin, ampicillin plus enrofloxacin) and possibly flunixin meglumine (for septic shock) should be administered. If the animal is dyspneic, oxygen therapy may be given by nasal insufflation or mask.

Gastric decompression should be performed while shock therapy is initiated. The stomach may be decompressed percutaneously with several large-bore intravenous catheters or a small trocar, or (preferably) a stomach tube may be passed. The stomach tube should be measured from the point of the nose to the xiphoid process and a piece of tape applied to the tube to mark the correct length. A roll of tape can be placed between the incisors and the tube passed through the center hole. Attempts should be made to pass the tube to the measured point. Placing the animal in different positions (i.e., sitting, reclining on a tilt-table) may help if it is difficult to advance the tube into the stomach. Do not perforate the esophagus with overly rigorous attempts to pass the tube. If these attempts fail, percutaneous decompression of the stomach should be attempted. This may relieve pressure on the cardia and allow the tube to enter the stomach. Once the air has been removed, the stomach should be flushed with warm water. If blood is seen in the fluid from the stomach, prompt surgical intervention is warranted because this may indicate gastric necrosis. If the stomach tube can still not be passed, and immediate surgical correction is not possible, temporary decompression may be achieved by performing a temporary gastrostomy. Placement of a Foley catheter into the stomach percutaneously should not be done unless the stomach is simultaneously tacked to the body wall because of the high risk of peritonitis if the stomach pulls away from the tube. Disadvantages of a temporary gastrostomy are that the stomach must be closed when the permanent gastropexy is performed, and there is a high risk of peritoneal contamination. However, a temporary gastrostomy will maintain gastric decompression if the animal is being referred, or surgery is delayed. If immediate surgery is not possible in an animal in which a stomach tube was passed but that dilates rapidly after decompression, the stomach tube can be exteriorized through a pharyngostomy approach. This will prevent the animal from chewing on the tube, until definitive surgery can be performed. After the patient has been decompressed and is stable, radiographs may be taken.

Surgical Treatment

Surgery should be performed as soon as the animal has been stabilized, even if the stomach has been decompressed. Rotation of a nondistended stomach interferes with gastric blood flow and may potentiate gastric necrosis.

Preoperative Management

The animal should be given intravenous fluids, antibiotics, and possibly flunixin meglumine before surgery (see discussion of Medical Management). Significant electrolyte and acid-base abnormalities should be corrected. A greatly enlarged stomach may hinder respiration and make it difficult for the animal to ventilate during anesthetic induction. An ECG should be monitored to detect cardiac arrhythmias, which, if significant, should be treated with lidocaine before surgery.

Anesthesia

Numerous anesthetic protocols have been described for dogs with GDV. If the animal has been decompressed and stabilized and cardiac arrhythmias are not present, the animal may be given oxymorphone and diazepam intravenously and induced with etomidate, thiobarbiturates, or propofol. If the animal is depressed, oxymorphone and diazepam alone may be used for induction, or if necessary for intubation, etomidate may be given. Etomidate is a good choice for induction if the animal has not been well stabilized because it maintains cardiac output and is not arrhythmogenic. Alternatively, a combination of lidocaine and thiobarbiturate may be used if arrhythmias are present. For the latter, 9 mg/kg of each is drawn up and half is given initially, intravenously. Additional drug is given to effect to allow the dog to be intubated. Generally, no more than 6 mg/kg of lidocaine is given intravenously to prevent toxicity. If bradycardia occurs, anticholinergics (e.g., atropine or glycopyrrolate) may be given. Nitrous oxide should not be used in dogs with GDV. Isoflurane is the inhalation agent of choice because it is less arrhythmogenic than halothane.

Surgical Anatomy

Normally, when viewed from the surgeon's perspective (i.e., with the animal in dorsal recumbency), the pylorus is located on the dog's right side, and the greater omentum arises from the greater curvature of the stomach and covers the intestines. The gastric (lesser curvature) and gastroepiploic (greater curvature) arteries supply the stomach and are derived from the celiac artery. The short gastric arteries arise from the splenic artery and supply the greater curvature. Rupture of the short gastrics in dogs with GDV is common and may contribute to blood loss and gastric infarction or necrosis. Eighty percent of the arterial flow is to the mucosa and the remainder is to the muscularis and serosa; therefore observation of mucosal color is not a reliable indicator of gastric wall viability. The mucosa often appears darkened due to vascular compromise, even when full-thickness necrosis is not present.

Surgical Technique

The goals of surgical treatment are three-fold: (1) to inspect the stomach and spleen so as to identify and remove damaged or necrotic tissues, (2) to decompress the stomach and correct any malpositioning, and (3) to adhere the stomach to the body wall to prevent subsequent malpositioning. Upon entering the abdominal cavity of a dog with GDV, the first structure noted is the greater omentum, which usually covers the dilated stomach.

Decompress the stomach before repositioning, by using a large-bore needle (i.e., 14 or 16 gauge) attached to suction. If the needle becomes occluded with ingesta, have an assistant pass an orogastric stomach tube and perform gastric lavage. Intraoperative manipulation of the cardia will usually allow the tube to be passed into the stomach without difficulty. If adequate decompression is still not achieved, or an assistant is not available, a small gastrotomy incision can be performed to remove the gastric contents, although this should be avoided if possible. For a clockwise rotation, once the stomach has been decompressed, rotate it counterclockwise by grasping the pylorus (usually found below the esophagus) with the right hand and the greater curvature with the left. Push the greater curvature, or fundus, of the stomach towards the table while simultaneously elevating the pylorus (towards the incision). Check to make sure that the spleen is normally positioned in the left abdominal quadrant. If there is splenic necrosis or significant infarction, perform a partial or complete splenectomy. Remove or invaginate necrotic gastric tissues. Avoid entering the gastric lumen, if possible. If you are uncertain whether gastric tissue will remain viable, invaginate the abnormal tissue. Verify that the gastrosplenic ligament is not torsed, and before closure, palpate the intraabdominal esophagus to ensure that the stomach is derotated. To prevent recurrence of GDV, the stomach must be permanently adhered to the body wall. Gastropexy should always be performed in conjunction with abdominal exploration and derotation of the stomach.

Partial Gastrectomy and Invagination of Gastric Tissue

Partial gastrectomy is indicated when necrosis, ulceration, or neoplasia involves the greater curvature, or middle portion, of the stomach. Necrosis of the greater curvature is primarily associated with gastric dilatation-volvulus (GDV) and may be treated by resection or invagination. Invagination does not require opening of the gastric lumen; however, obstruction from excessive intraluminal tissue is possible (but rare). The extent of necrosis is assessed by observing serosal color, gastric wall texture, vascular patency, and bleeding on incision; however, it is difficult to determine tissue viability in many cases with these techniques. Necrotic tissue may range in color from gray-green to black and often feels thin. A full-thickness incision can be made into the suspected necrotic tissue to assess arterial bleeding. Intravenous fluorescein dye has not proved to be an accurate method of determining gastric viability in dogs with GDV. Generally, if you question the viability of the gastric tissue, remove it or invaginate it. Failure to remove or invaginate necrotic tissue may cause perforation, peritonitis, and death. Melena is commonly observed for a few days after gastric invagination.

To remove the greater curvature of the stomach, ligate branches of the left gastroepiploic vessels and/or short gastric vessels along the section of the stomach to be removed. Excise the necrotic tissue, leaving a margin of normal, actively bleeding tissue to suture. Close the stomach with a two-layer inverting suture pattern, using an absorbable suture (e.g., polydioxanone or polyglyconate suture; 2-0 or 3-0). Incorporate submucosa, muscularis, and serosal layers in a Cushing or simple continuous pattern in the first layer. Then use a Cushing or Lembert pattern to invert the serosa and muscularis over the first layer. Alternatively, you may use a thoracoabdominal (TA) stapling device to close the incision. To invaginate necrotic tissue, use a simple continuous suture pattern followed by an inverting suture pattern. Place sutures in healthy gastric tissue on both sides of the tissue that is to be invaginated, bringing the healthy tissue over the top of the necrotic tissue. Be sure that sutures are placed in healthy tissues to prevent dehiscence.

Permanent Gastropexy

Gastropexy techniques are designed to permanently adhere the stomach to the body wall. The most common indications are GDV (pyloric antrum to right body wall) and hiatal herniation (fundus to left body wall). Numerous gastropexy techniques have been described. Although the strength and extent of adhesions created by these various techniques differ, all of them (when properly performed) prevent movement of the stomach. To create a permanent adhesion, the gastric muscle must be in contact with the muscle of the body wall; intact gastric serosa will not form permanent adhesions to an intact peritoneal surface.

A technique for gastropexy has recently been described in which the stomach is incorporated into the abdominal incision during closure (Meyer-Lindenberg et al, 1993). Although this technique is easy, quick, and decreases recurrence of GDV, it results in the stomach being permanently adhered to the ventral body wall. The main advantage of this procedure is that it can be performed quickly. However, the subsequent abdominal exploration via a midline abdominal incision could perforate the stomach. Therefore, although this technique is preferable to not performing any type of "pexy," it is not generally recommended.

Muscular Flap (Incisional) Gastropexy

Muscular flap (incisional) gastropexy (see Figure 1) is easier than circumcostal gastropexy and avoids potential complications associated with tube gastropexy. Make two hinged flaps in the seromuscular layer of the gastric antrum (similar to that for a circumcostal gastropexy). Then make similar flaps in the right ventrolateral abdominal wall by incising the peritoneum and internal fascia of the rectus abdominis or transverse abdominis muscles. Elevate flaps by dissecting ventral to the muscle layer. Invert the flaps, and suture the edge of the abdominal flaps to the gastric flaps, using a simple continuous suture pattern of 2-0 absorbable or nonabsorbable suture. Ensure that the muscularis layer of the stomach is in contact with the abdominal wall muscle). Suture the cranial margin first, followed by the caudal margin. Be sure to place sufficient sutures so that a loop of bowel cannot become incarcerated between the flaps.

Postoperative Care And Assessment

Electrolyte, fluid, and acid-basis status should be monitored closely postoperatively. Many dogs with GDV are hypokalemic postoperatively and require potassium supplementation. Small amounts of water and soft, low-fat food should be offered 12 to 24 hours after surgery, and these patients should be observed for vomiting. Gastritis secondary to mucosal ischemia is common and may be associated with gastric hemorrhage or vomiting. If vomiting is severe or continuous, a centrally-acting antiemetic may be given. Secondary gastric ulcers may occur and require treatment. H2 receptor blockers (e.g., cimetidine, ranitidine, or famotidine) decrease gastric acidity and may be beneficial. Intravenous fluid therapy should be continued until oral fluid intake is adequate to maintain hydration. Patients should be monitored for hypoproteinemia and anemia in the early postoperative period. Ventricular arrhythmias are common in dogs with GDV and usually begin 12 to 36 hours postoperatively. Their cause is unknown, but myocardial depressant factor, decreased cardiac output, and myocardial ischemia may contribute. Treatment of cardiac arrhythmias includes maintenance of normal hydration and correction of electrolyte imbalances [some antiarrhythmic drugs (i.e., lidocaine) are ineffective when the animal is hypokalemic]. If the arrhythmias: (1) interfere with cardiac output (as noted by poor peripheral pulses), (2) are multiform, (3) have subsequent premature beats inscribed on the wave of the previous complex (R on T), or (4) have a sustained ventricular rate greater than 160 beats per minute, they should be treated; usually with intravenous drugs. A test bolus of lidocaine, given intravenously (2 mg/kg bolus, up to 8 mg/kg total dose), can be used to determine responsiveness to this drug. If the arrhythmias decrease or stop, lidocaine should be given by a continuous intravenous infusion of 50 to 75 µg/kg/min. Low doses should be used initially and only increased if necessary. Signs of lidocaine toxicity include muscle tremors, vomiting, and seizures; lidocaine therapy should be discontinued if these signs occur. Other potentially effective antiarrhythmic drugs include procainamide and sotolol. Procainamide may be given as an intravenous bolus, by continuous infusion, intramuscularly, or orally. Sotolol may be effective in animals that have not responded to lidocaine and procainamide.

Complications

Sepsis and peritonitis may be caused by gastric necrosis or perforation if devitalized tissue is not adequately removed. Diagnostic peritoneal lavage may help diagnose peritonitis. Peritonitis mandates immediate surgical intervention. DIC may occur in dogs with GDV or peritonitis. Assessment of clotting parameters, plus appropriate treatment with fluids and heparin, may be necessary.

Prognosis
With timely surgery, the prognosis is fair; however, mortality rates as high as 45% and greater have been reported (Matthiesen, 1985). A recent study reported a mortality rate of 15% among dogs with GDV; the mortality rate was 0.9% if gastric dilation without volvulus was present (or if GDV could not be verified at surgery) (Brockman, Washabau, Drobatz, 1995). The prognosis is poor if gastric necrosis or perforation occurs or if surgery is delayed. Recurrence rates for GDV differ, depending on techniques used, but most have reported rates of less than 10%. Tube gastropexy has the highest reported recurrence rate, varying from 5% to 29% (Fox et al, 1984; Johnson, Barrus, Greene, 1984). Some dogs with GDV respond to tube decompression and medical stabilization alone. Occasionally, the stomach becomes normally positioned after the air is removed; or, it was only partially rotated (less than 180 degrees) or merely dilated. However, these dogs still have a high likelihood of recurrence. Therefore gastropexy should be recommended, even when conservative management successfully alleviates the gastric malpositioning. The reported recurrence rates of dogs operated on for GDV in which the stomach has been repositioned but gastropexy not performed approaches 80% (Meyer-Lindenberg et al, 1993; Whitney, 1989).

• Reduce the pre-surgical fast to 4-8 hours.
• Administer dextrose containing fluids intravenously in the longer elective procedures and offer the patient a small meal 1-2 hours after surgery.

• Calculate medications carefully and dilute medications 1:1 with sterile saline to allow for slow administration of intravenous medications.
• Protocols used at TAMU
• Puppies less than 4 months of age are typically given glycopyrrolate and butorphanol intramuscularly prior to induction. These patients are normally induced with diluted thiopental and maintained on halothane or isoflurane.
• Kittens less than 4 months of age are given premedications that include glycopyrrolate, butorphanol, acepromazine, and ketamine intramuscularly. The kitten is administered gas anesthesia via face mask, if needed, prior to endotracheal intubation.
• Use a Bain or other coaxial nonrebreathing system for patients weighing less than 6.8 kg. In larger patients use a small semi-closed anesthesia system. (Kitten castrations don't generally require intubation, unless a risk factor is identified.)
• Place an intravenous catheter prior to the procedure to allow for administration of emergency medication.
• Administer warmed dextrose-containing fluids IV (22 ml/kg- first hour, 11 ml/kg- thereafter) during surgery (except for kitten castrations). For kitten castrations administer warmed isotonic fluids (ie. LRS or Normosol) subcutaneously (11 ml/kg). Mechanical syringe pumps or careful monitoring of the drip rate is needed to prevent fluid overload.