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Supplemental Oxygen Delivery and Feeding Tube Techniques
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Indwelling tubes that enter the nose and stop in the ventral nasal meatus (nasal), pharynx (nasopharyngeal), or trachea (nasotracheal) are effective for the delivery of supplemental oxygen (O2). Those that continue on through the ventral nasal meatus and pharynx and stop in the caudal thoracic esophagus (nasoesophageal [NEO]) are useful for the delivery of fluids and nutritional supplements or for the aspiration of air and fluids to provide decompression of the esophagus in conditions causing megaesophagus. Tubes that continue on into the stomach and either stop there (nasogastric [NG]) or continue into the duodenum or jejunum (nasoenteric [NET]) are useful for delivery of fluids and nutrients or for removal of accumulated air and fluids. All these tubes are placed initially into the nasal passage and are passed into the ventral meatus using the same technique. The type of tube selected depends on its intended use. Placement of each of the types of tube is simple to perform. In rare instances, placement under fluoroscopic guidance may be required (i.e., placing an NG tube past an esophageal stricture or placing an NET tube). After insertion, all indwelling tubes are generally well tolerated by most patients, even patients that are completely alert. On occasion, an Elizabethan collar is recommended to prevent the patient from dislodging the tube. Sedation is not necessary in most patients. The nose generally accommodates up to three to four types of tubes at the same time. When more than one type of tube is placed in the nose, the tubes must be labeled appropriately to avoid complications.
Supplemental oxygen (O2) should be provided as a first line of treatment to dogs and cats in shock (septic, traumatic, cardiogenic) and cardiac failure and those with respiratory compromise. This supplementation is also a useful treatment in postoperative critically ill patients during the anesthetic recovery period and in anemic animals.
The use of O2 cages has been helpful in providing an O2-enriched atmosphere for animals. However, these cages are expensive, and available sizes often cannot house large to giant breed dogs adequately. They also are inefficient to operate because a considerable amount of O2 is dissipated into the room each time the door is opened. Furthermore, once a patient is placed into an O2 cage, careful evaluation, continued monitoring, and treatment are difficult in the “forced” isolation that this form of O2 therapy requires. Much time is also required to generate the higher levels of O2 recommended in patients placed in O2 cages. The law of displacement dictates the time required. The cubic volume of commercial O2 cages varies from 300 to 500 L. If O2 is provided at a flow rate of 20 L/minute into the cage, and no leakage occurs, it will take a minimum of 12 minutes to achieve the O2 concentration of near 100% that is recommended in patients suffering from life-threatening conditions. O2 cages are also inefficient at providing sustained concentrations of O2 higher than 50% because of unavoidable leaks. In investigations with one O2 cage, the O2 concentration could not be held above 40%.
Other available means of providing supplemental O2 therapy include the use of face masks, O2 hoods, bilateral human nasal cannulas, and transtracheal catheters. Difficulties with the use of a mask in nervous and apprehensive animals are all too familiar. O2 hoods are well tolerated and provide up to 80% O2 concentrations, but access to the face is restricted, and the animal is unable to drink or eat (Figure 6-1). These collars can, however, be used in conjunction with nasal catheters or short nasal cannulas to increase tracheal O2 concentration.
Short human nasal cannulas are inserted into the nares and are secured around the neck using a drawstring. These devices are well tolerated, but they frequently dislodge if the patient is active. Complications with transtracheal catheters have been reported. Nasal O2 administration is an efficient and effective means of providing high inhalational concentrations of O2 (up to 85 to 95%). The deeper the placement of the end of the tube in the respiratory tract, the more efficient the device is in elevating the concentration of O2. Nasal tubes are not as effective as nasopharyngeal tubes in raising the inhaled tracheal O2 concentration. The highest concentrations of O2 are achieved with the use of nasotracheal tubes.
The animal’s head is held gently restrained upward, and 1 mL of 2% lidocaine (dogs) (Animal Health Associates, Kansas City, MO) or 5 drops of 0.5% proparacaine ophthalmic Solution (dogs and cats) (Ophthaine, ER Squibb & Sons, Princeton, NJ) are administered into either nostril. The right nostril generally is preferred for right handed operators and the left nostril for left handed operators. The local anesthetic solution is allowed to run down the nasal passage. This procedure is repeated after 10 to 20 seconds. After another short waiting period to allow for desensitization, the tip of the selected catheter is lubricated on its outer surface with a commercial water soluble lubricant (Xylocaine Jelly 2%, Astra Pharmaceutical Products, Inc., Worcester MA). The catheter can be a 3.5- to 8-French red rubber (Sovereign, Sherwood Medical Products, St. Louis, MO) or polyvinyl chloride (Cook Critical Care, Bloomington, IN) tube, or for extremely small patients, a long flexible 17-gauge polyethylene intravenous catheter. The addition of small side holes helps to disperse the stream of O2 more evenly within the nasal passage; however, these holes are not usually required.
For nasal O2 tube placement, the tube is premeasured alongside the patient’s face so the tube’s tip, after placement, extends into the nasal cavity to the level of the first or second premolar. This facilitates flow through the ventral nasal meatus. This tube can be measured alongside the animal’s teeth or by measuring from the tip of the nose to the medial canthus of the eye. After premeasuring, the tube is introduced into the nasal orifice while the patient’s head is held firmly. Cats have a straight nasal passage, and the tubes generally pass easily. In the dog, pushing the tip of the nose upward allows the tube to be passed more easily into the ventral meatus. The tip is directed ventromedially (Figure 6-2). In the cat, the tube can be simply inserted straight in most cases. After this initial introduction, the tip, in both the dog and the cat, is directed ventromedially until the desired length has been inserted (Figure 6-3). Most animals object to the initial passage of the tube by sneezing and trying to shake their heads, but then they remain quiet after tube passage has been completed. If an animal objects to the insertion of the tube, slight sedation is recommended using low doses of intravenous neuroleptanalgesia (e.g., butorphanol [Torbugesic], 0.1 to 0.4 mg/ kg, and diazepam [Valium], 0.05 to 0.2 mg/kg, or acepromazine .02 to .04 mg/kg).
After insertion to the level required, the tube is fixed to the skin using 3-0 or 2-0 silk suture with a swaged-on cutting needle. The most critical area requiring initial fixation is the first 0.5 cm after the tube exits from the nostril. This suture is usually preplaced to facilitate securing the tube immediately after it is placed. Several sutures are used to secure the tube (Figure 6-4). Each suture is placed through the skin in a “quick pass” fashion without hair clipping, aseptic preparation, or local anesthesia. After a loose simple interrupted suture is tied, the ends are wrapped around the tube and are tied again. An alternative fixation method is to apply a few drops of cyanoacrylate glue to the tube and tufts of hair on the nose and along the face, or skin staples can be used to secure the tube. Elizabethan collars are only required in patients objecting to the tube.
Oxygen Delivery Protocol
Tubing for O2 administration (Tomac, American Hospital Supply Corp., Chicago) or an intravenous administration set is connected to the external end of the tube. The other end, in turn, is attached to the O2 source with a standard O2 flow meter. If O2 supplementation for more than 24 hours is anticipated, use of a commercial humidification chamber is recommended. Alternatively, a homemade humidifier can be fashioned using a crated intravenous fluid infusion bottle. The O2 source is attached to the vent hole, and O2 is bubbled through warm water. Additional tubing, as necessary, is used between the patient and the humidifying unit to allow the animal freedom to move without fear of tube disconnection. The homemade humidification chamber full of water must not tip over, because this would result in rapid delivery of water into the patient’s nasal passage.
For patients being resuscitated, flow rates that generate at least 60 to 80% O2 concentrations are recommended. In patients that have hemodynamic and pulmonary stability, flow rates are decreased 50% to provide approximately 40% inspired O2. The flow rate to provide 60 to 80% O2 concentrations is approximately 50 mL/kg body weight per minute in small dogs and cats and approximately 100 mL/kg body weight per minute in large dogs when delivering O2 using properly placed nasal catheters. A proportionally greater amount probably is required in large breed dogs because of a concomitant increased amount of anatomic dead space in larger animals.
After O2 administration is begun, the patient should be observed carefully to determine the response to therapy and to identify adverse effects, which are rare. Clinical signs such as decreased anxiety and decreased respiratory rate and effort indicate an improvement in response to the O2. Pulse oximetry can also be used to assess oxygenation. O2 supplementation is indicated whenever O2 saturation is below 92%. Accurate measurements are, however, sometimes difficult to obtain in the awake patient because of probe placement difficulties. In the critically ill patient, arterial blood gases should be monitored whenever possible. Partial O2 pressures considered sufficient should be at least 60 to 65 mm Hg. If hypercapnia exists (PCO2 greater than 50 mm Hg), mechanical ventilation rather than simple O2 supplementation should be performed. Provided sufficient volume exchange is taking place to prevent hypercapnia, the O2 flow rate can be increased to provide greater inspiratory O2 concentrations if no favorable clinical response is observed or arterial PO2 values remain below 65 mm Hg. Permissible flow rates and the corresponding O2 percentages in the inspired air are given in Table 6-1. If after increasing the flow rates arterial O2 values do not increase above 70 mm Hg, intermittent positive pressure ventilation (IPPV) with positive end-expiratory pressure should be instituted. If the patient’s work of breathing does not improve with the high concentration of O2, then control of breathing with IPPV should be provided. The use of mechanical ventilation in these patients is important; otherwise, ventilatory failure and death will ensue.
Complications with the use of nasal O2 administration are uncommon. O2 is dry and cool; therefore, prolonged use (more than 3 to 5 days) may cause rhinitis and sinusitis. When these complications do occur, they usually are mild and become evident as a persistent serous nasal discharge. The discharge usually clears within several days after the nasal tube is removed. The use of nasal O2 in patients with nasal bone fractures may lead to subcutaneous emphysema. If blood is present in the nose, nasal O2 administration is not recommended because bubble formation and foam may interfere with air exchange. In these patients, nasotracheal or transtracheal O2 is recommended.
Tube dislodgment is an infrequent complication if the catheter is placed in the nose for a sufficient distance and if fixation of the tube is performed correctly. Persistent sneezing and continued irritation are rare and necessitate the use of repeated local anesthetic instillation, an Elizabethan collar, or light intravenous chemical sedation (e.g., oxymorphone at 0.02 mg/kg or diazepam at 0.1 mg/kg). Mild epistaxis caused by misdirection of the tube into the maxillary or ethmoid turbinates during placement may occur, but in our experience this occurs rarely and is not severe enough to warrant discontinuation of a tube’s insertion or use.
Patients with severe tracheobronchial froth or fluid accumulation, as observed in animals with severe pulmonary edema, should receive nasotracheal or transtracheal O2 rather than nasal O2. Nasal tubes should be avoided in those patients with severe epistaxis or mucopurulent nasal discharge, suspicion of maxillary or cranial vault fracture after head injury, or head injury or any condition in which elevation of intracranial pressures secondary to sneezing or struggling is contraindicated. Ineffective ventilation requiring other primary care (intubation and positive-pressure ventilation) is also a contraindication to the placement of nasal O2 tubes.
Nasopharyngeal tubes allow delivery of O2 into the nasopharynx. This method can provide high concentrations of O2 and, if flows are high enough, some level of continuous positive airway pressure (CPAP). CPAP is even more effective if bilateral nasopharyngeal tubes are placed. As the patient exhales, it exhales against some force created by the flow of the O2 in a caudal laryngeal direction. The goal is to create an increase in the patient’s functional residual volume. This can be done with CPAP.
A nasopharyngeal tube is placed in a fashion similar to that of a nasal catheter, but the lubricated tip of the tube is continued through the ventral meatus past the maxillary turbinate. The tube is held alongside the face and neck and is premeasured from the external naris to just proximal to the larynx. In dogs, some resistance may be encountered at the maxillary turbinate region because of a narrowing of the ventral meatus in a dorsoventral direction. If the tube cannot be passed farther than the level of the eyes in dogs or cats, the tube is assumed to be in the dorsal meatus with its tip in the ethmoid turbinate. The tube must be withdrawn and redirected ventrally if this occurs. After the tip is past the maxillary turbinate in the ventral meatus, resistance to the tube’s passage decreases, and the tube can be passed into the nasal pharynx and pharyngeal isthmus. The ideal location is just dorsal to the rima glottis (Figure 6-5).
High O2 flow rates (greater than 200 mL/kg per minute) should be administered carefully when providing O2 through nasopharyngeal tubes. Rarely, gastric distension occurs if flow rates are exceedingly high (greater than 200 mL/kg per minute) or if the nasopharyngeal catheter migrates into the esophagus. Bradycardia, believed to be vagally mediated, can also occur.
Nasotracheal tubes provide an effective means of providing O2 to the patient that has laryngeal palsy or a collapsing cervical trachea. These catheters also generate some degree of CPAP when high flow rates are used. Patient tolerance is usually good, with little coughing. In animals that do not tolerate the tubes, mild sedation may be required.
Before placement of a nasotracheal tube, the tube should be premeasured such that the tip will rest at the level of the tracheal bifurcation or fifth intercostal space. A 3.5- to 8-French feeding tube is generally used. The tube is placed in a fashion similar to that of a nasopharyngeal catheter. The tube is passed blindly into the trachea through the larynx by hyperextending the patient’s head and neck and advancing the tube (Figure 6-6). If coughing is noted, another 0.33 mL of local anesthetic is infused through the tubing, with the tubing in the mid distal pharynx. Once the membranes around the larynx are anesthetized, the tube is advanced as inhalation occurs. If the tube does not pass after several attempts, a short-acting neuroleptoanalgesic can be administered to the patient, and the tube can be placed by direct visualization using a laryngoscope and something to grasp the tip of the tube and direct it through the rima glottis into the trachea.
The position of the tube should be confirmed with a radiograph or by aspiration using a 60-mL syringe. If the tube is in the trachea, air should continue to be aspirated easily. If the catheter is in the esophagus, air may be initially aspirated, but it should stop.
The nasotracheal tube is used in a fashion similar to that of nasal and nasopharyngeal tubes. For nasotracheal catheters, flow rates are decreased by 50% from those recommended for nasal O2 tubes to provide equivalent O2 concentrations. Humidification of the O2 is essential with the use of nasotracheal tubes, to prevent mucosal drying and dysfunction of the mucociliary apparatus, which can lead to an inability to clear secretions and possible pneumonia. Infusion of saline through the nasotracheal tube can be used to help loosen secretions in patients with dysfunction of the mucociliary apparatus or pneumonia.
Tubes for Gastrointestinal Access
NEO, NG, and NET tubes can be used for decompression and feeding. Smaller bore NEO, NG, and NET tubes are useful for the administration of water, electrolytes, and liquid enteral support diets. Because dehydration and protein–energy malnutrition frequently are encountered in seriously ill or injured animals, the use of these indwelling tubes for rehydration and nutritional support often is a key component in successful overall patient management. Contraindications to use of NEO or NG fluid and nutritional therapy support include persistent vomiting and high gastric residual volumes. The presence of stupor or coma is a relative contraindication to NEO and NG feeding, particularly if bolus feeding is provided. If slow, continuous-rate infusions result in minimal residual volumes, then the risk of regurgitation and aspiration is low enough that NEO or NG feeding can be used.
Decompression of a dilated esophagus, stomach, or intestinal tract can be accomplished by use of large-bore single lumen or double-lumen (sump) NEO, NG, or NET tubes. Decompression of the esophagus alleviates some of the risk of aspiration in the patient with megaesophagus and actively decreases the stretch in the skeletal muscle that results in dilatation. In the stuporous or comatose patient, or in the patient receiving mechanical ventilation, active decompression helps to prevent aspiration. In the patient having difficulty ventilating, decompression of the stomach improves ventilation because of reduced impedance to diaphragmatic excursions. This is particularly helpful in cats and small dogs because they breathe primarily using the diaphragm. Clinically, NG decompression has been helpful in the temporary management of gastric dilation–volvulus syndrome when the gastric distension has been due primarily to air and fluid. Decompression of the stomach after abdominal surgery helps to decrease the time to return to normal gastric motility. After placement, the NG tube is periodically aspirated (e.g., once every 1 to 2 hours). The tube is left in place until bowel sounds return or the patient is believed to be out of danger of postoperative redistension. Antral dilation is a strong stimulus for vomiting. The use of NG tubes decreases the incidence of vomiting in the patient with gastrointestinal or pancreatic disease and is especially useful in the patient with canine parvovirus infection.
Tube Selection and Insertion
The techniques for inserting an NEO, NG, or NET tube for decompression or feeding are the same. Polyvinyl chloride (Argyle nasogastric feeding tube, Sherwood Medical Products), polyurethane (Cook Critical Care), or red rubber tubes from 3.5 French (cats and small dogs) to 12 French (medium to large dogs) are used. Specially designed tubes that are weighted on their proximal ends with either tungsten or mercury are useful to ensure that the tube will stay in the stomach lumen (Travasorb dualport feeding tube, Baxter Health Care Corp., Deerfield, IL). The smaller the tube, the more difficult it is to use for decompression. A nylon stylet that accompanies commercial polyurethane tubes provides added stiffness necessary for insertion. With smaller polyvinyl chloride tubes, a woven angiographic wire stylet (Wire guide, Cook Critical) is used to provide added stiffness. One or two milliliters of vegetable or mineral oil is injected into the lumen of a tube to facilitate ease of insertion and withdrawal of the woven wire through the lumen.
After selection of the tube and placement of the stylet, the length necessary to reach the distal thoracic esophagus (NEO) or the stomach (NG) is determined by measuring alongside the patient’s neck and body from the tip of the nose to the eighth or ninth rib for NEO tubes or to the thirteenth rib for NG tubes (Figure 6-7). For NET tubes, length is added to ensure that the tip of the proximal end of the tube will reach the area of the bowel lumen selected. Most often, the tube for enteral feeding is a nasoduodenal tube with a tip that ends near the pelvic flexure of the duodenum. The tube in these cases is premeasured to extend from the nose to the wing of the ilium (See Figure 6-7).
The lubricated tip of the tube is introduced into the patient’s nostril in the same manner as described for nasopharyngeal tubes. After the tip is past the maxillary turbinate in the ventral meatus, resistance to the tube’s passage decreases, and the tube can be passed into the nasal pharynx and pharyngeal isthmus. At this point, the patient’s head must be kept in a neutral position, with the neck gently flexed to facilitate passage of the tube into the esophagus (Figure 6-8). If the neck is hyperextended, the tube may enter the larynx and trachea. With continued advancement of the tube, the patient is often observed to swallow several times. Once the tip of the tube has been advanced into the caudal thoracic esophagus (NEO tube) or into the proximal portion of the stomach (NG tube), the lubricated stylet is withdrawn. The use of a stylet also helps to facilitate the passage of the tube into the stomach through the cardia.
Air is injected into the tube while auscultation of the left chest wall and left paralumbar fossa is performed; the presence of gargling sounds during this procedure indicates that the tube is in the distal esophagus or stomach, respectively. In most cases, a lack of coughing during injection of 5 to 10 mL of sterile saline down the tube indicates that the tube is not in the trachea. However, the result of this test may vary with the individual animal, and the position of all tubes should be radiographically confirmed if they are to be used for infusion of fluids or liquid diets.
Special tubes or manipulations are required for placement of NET tubes into the duodenum or jejunum. The tube can be guided by peristaltic action into the duodenum, but this is often difficult to accomplish. The tubes can be guided through the pylorus using endoscopy or fluoroscopy. NET tubes have been most successfully placed at the time of abdominal surgery by the surgeon guiding the tip of the tube, which is palpated and guided through the stomach and intestine into the portion of the bowel intended. Weighted tungsten or mercury tubes have been used to help in guiding tubes through the stomach into the intestine (Travasorb dualport feeding tube, Baxter Health Care Corp.). The weighted tip also may help to ensure that the tube will stay in the bowel lumen and not curl or kick back into the stomach. Passage of the tube into the small intestine through the action of peristalsis has been unreliable, particularly in sick patients with at least some degree of gastroparesis. Metoclopramide, 0.4 mg/kg per day intravenously, has been used to help stimulate gastric motility to facilitate the tube’s passage into the duodenum.
Once the tip of the tube has been placed in the desired location, the tube is secured with several sutures placed at the base of the nostril and around the tube, or with glue as described previously for nasal O2 tubes. If the tube demonstrates a tendency to back out of the nose, 1 to 2 cm of coated copper wire (18 gauge telephone wire) can be used to support the bend in the tube as it exits from the nose. On occasion, the tube may back out of the intestine, or the dog or cat may vomit the tubes into the mouth. In this case, the tube must be removed. A narrow gauge flexible wire can sometimes be left in the tube to help prevent tube migration. Specially designed catheters are also available that allow the delivery of nutrients while the wire is left inside the catheter lumen.
The remaining length of the tube or an attached extension tube (intravenous administration extension set) is secured to the top of the patient’s head or the side of the face. An Elizabethan collar can be applied if necessary. The end of the tube is capped to prevent air from entering the gastrointestinal tract by diaphragmatic movement until its use is required.
Protocol for Using Tubes for Decompression
A 60-mL syringe is attached to the end of the tube, and aspiration is done as often as required to keep a slight amount of negative pressure on the hollow viscus aspirated. For prevention of recurrence of gastric dilation or for decompression of the small intestine, aspiration generally is performed every 1 to 2 hours until a negative pressure is reached each time. If the fluid aspirated is viscous, dilution with sterile water or saline may be required. The tube should be flushed with a small amount of saline or water each time the tube is used, and then the tube should be capped. Holding the column of water in the tube helps to prevent clogging. Maintenance of decompression usually is required only for 24 to 48 hours because most intestinal ileus or gastroparesis is resolved by then.
The efficiency of gastrointestinal decompression achievable with a simple single lumen tube (Argyle stomach tube (Levine Type), Sherwood Medical Products) and intermittent aspiration with a syringe can be improved by the use of a double-lumen sump tube (Salem sump tube, Sherwood Medical Products) with continuous 20- to 30-mm Hg suction or intermittent mechanical 80- to 90-mm Hg suction. This type of suction requires the use of specially designed equipment. Automatic intermittent suction, for example, is often best performed with the use of a thermotic drainage pump that is electronically driven (Thermotic drainage pump, GOMCO, Allied Healthcare Inc., Buffalo, NY). Fortunately, in most clinical patients, this type of special equipment is not necessary, and simple intermittent syringe decompression is sufficient.
Protocol for Using Tubes for Feeding
For the administration of fluids and liquid enteral diets, a syringe is used for slow bolus delivery. Slow bolus delivery of fluids and liquid enteral diets can be done safely through NEO and NG tubes in animals that are conscious. However, bolus feeding is not recommended in unconscious or semiconscious patients because of the higher risk of pulmonary aspiration. Bolus feeding should not be done through an NET tube initially because of the high occurrence of vomiting and diarrhea, which can be caused by the acute overload of hyperosmolar nutrients in the small intestine. Drip infusion is the preferred method of the delivery in these circumstances. A pediatric intravenous fluid administration set and bottle are used for the delivery of enteral diets. The use of an enteral or intravenous infusion pump or a syringe facilitates the delivery of these enteral liquid diets.
Initially, an electrolyte and glucose mixture is administered at a rate of 0.25 to 0.5 mL/kg per hour. This rate can be used in all patients including those that have had gastrointestinal surgery; however, it may be too fast for those patients that have undergone massive bowel resections or have pancreatitis. In such patients, the initial rate infused should be no greater than 0.1 to 0.2 mL/kg per hour. The drip rate is steadily increased until caloric requirements are met. Rates higher than 4 mL/kg per hour are usually associated with severe, osmotically induced diarrhea; therefore, the maximum rate usually used for constant rate infusions is 2.0 to 3.0 mL/kg per hour.
Many monomeric and polymeric liquid diets are available for tube feeding. Monomeric or elemental diets are composed of amino acids (Vivonex, Sandoz Nutrition, Minneapolis MN; Alitraq, Ross Laboratories, Columbus, OH) or dipeptides and tripeptides (Peptamen, Clintec Nutrition Co., Deerfield, IL) and require no digestion before absorption. The amino acid–based diets tend to be hyperosmotic and may require dilution initially to a 50% concentration. They usually are more expensive than polymeric diets, but they may be useful in patients with decreased digestive ability. The dipeptide- and tripeptide- based diets tend to be isosmolar and can generally be given initially at full strength concentration. Polymeric diets (Impact, Sandoz Nutrition; Jevity, Ross Laboratories) are made of complex carbohydrates and proteins and require digestion before absorption, but they are usually isosmotic unless they are flavored. Special polymeric diets designed specifically for cats and dogs (CliniCare and RenalCare, Pet Ag Inc., Hampshire, IL) have been developed and have been clinically effective in providing nutritional support to critically ill or injured dogs and cats. Polymeric diets are usually administered either full strength if plasma proteins are normal and anorexia has not been present for longer than 3 days. If plasma protein levels are below normal or anorexia has been present for longer than 3 days the diets should be initially diluted to a 50% concentration with water. The monomeric diets may require dilution to 25% concentration for initial administration. After the rate of administration is stabilized at 2 to 3 mL/kg per hour and the diet is found to be tolerable (no abdominal pain, vomiting, or diarrhea), the concentration of the diet can be gradually increased.
Complications with feeding and decompression tubes are primarily associated with tube migration, especially dislodgment. Dislodgment is usually caused by vomiting or by the animal’s pawing at the tube or rubbing its face.
When concern exists about the location of the tip of the tube, a radiograph should be taken to ensure that the location is correct. Disaster can occur if a tube is displaced into the trachea and food is administered.
- Crowe DT. Clinical use of an indwelling nasogastric tube for enteral nutrition and fluid therapy in the dog and cat. J Am Anim Hosp Assoc 1986;22:675 678.
- Crowe DT. Use of a nasogastric tube for gastric and esophageal de compression in the dog and cat. J Am Vet Med Assoc 1986; 188:1178 1182.
- Crowe DT. Enteral nutrition for critically ill or injured patients. Part I. Compend Contin Educ Pract Vet 1986;8:603.
- Crowe DT. Enteral nutrition for critically ill or injured patients. Part II. Compend Contin Educ Pract Vet 1986;8:826.
- Fitzpatrick RI, Crowe DT. Nasal oxygen administration in dogs and cats: experimental and clinical investigations. J Am Anim Hosp Assoc 1986;22:293 297.
Esophagostomy tubes provide a simple and effective means of administering fluid and nutritional support to the small animal patient. The tubes can also be used for esophageal or gastric decompression.1 Esophagostomy tubes can be rapidly placed (generally within 5 minutes) and require minimal surgical equipment (a scalpel blade, a pair of curved forceps, and nonabsorbable suture material). Simple red rubber feeding tubes are most frequently used. Patients have been fed for up to 2 years using these tubes. No cases of esophageal stricture or permanent esophagocutaneous fistula have been observed.
Esophagostomy tubes are indicated whenever nutritional support is required and the stomach is functional but the patient is unwilling or unable to ingest food or water. Esophagostomy tubes can also be used to keep the stomach and esophagus decompressed because aspiration of these tubes helps to prevent air or fluid from accumulating. This may be useful in the management of patients with megaesophagus or those that have undergone surgical correction of gastric dilatation–volvulus. Esophagostomy tubes were developed and first used in clinical veterinary medicine by Crowe.2 They were developed and used to avoid the airway difficulties associated with pharyngostomy tubes (Figure 6-9).3 With pharyngostomy tubes, a portion of the tube can interfere with laryngeal function, even after careful placement using modified techniques. The surgical approach for placement of the esophagostomy tube is simpler than that of the pharyngostomy tube, with less likelihood of damage to vital vascular and neurologic structures. Percutaneous gastrostomy tubes require special feeding tubes and because of penetration of the stomach and peritoneal cavity, the risk of leakage and subsequent development of peritonitis always exists. From our experience, the patient does not need to be subjected to these risks, and, whenever possible, an esophagostomy tube should be selected over a gastrostomy tube. Most conditions for which clinicians use percutaneous gastrostomy tubes for feeding can be also managed with esophagostomy tubes. Esophagostomy tubes can be used in patients that have had esophageal surgery; however, care should be taken to ensure that a smaller bore flexible feeding tube is used and that the end of the tube is not rubbing against a wound site or surgical incision.
In general, esophagostomy tubes should not be used for feeding or decompression if the patient 1) is vomiting, 2) has cervical or thoracic esophageal disease that will be worsened by the placement of a tube passing through the affected area, and 3) has an infection involving the cervical region close to the tube exit site. Because placement of esophagostomy tubes requires light general anesthesia, the risks of anesthesia should be weighed against the benefits of the placement of esophagostomy tubes in critically ill animals.
The type and length of tube selected depends on the intended use of the tube. Esophagostomy tubes used for feeding or for esophageal decompression (i.e., for long term management of megaesophagus) should end in the distal thoracic esophagus. Tubes that pass through the lower esophageal sphincter increase the risk of gastroesophageal reflux in some patients. For gastric decompression or feeding, whenever the esophagus needs to be bypassed, an esophagogastric tube is placed with the tip of the tube resting in the midfundic region of the stomach. An esophagoenteral tube can also be placed at the time of abdominal surgery if the stomach needs to be bypassed. The proximal end of the tube should be shortened as required, so only sufficient tubing protrudes from the skin to permit attachment to a syringe for feeding or decompression. Excessive tube length protruding from the skin may be annoying to the animal and may catch on objects.
Esophagostomy tubes used for feeding or decompression should be flexible and in general of as large a bore as possible. This provides less chance for kinking and occlusion. The actual size of each tube selected depends on the size of the animal and on the intended purpose for the tube (Table 6-2). Generally, no tube smaller than 10 French should be used for decompression or if a canned or gruel diet is to be used for feeding. For small cats and dogs, a 10- to 12 French tube is used. For medium sized dogs, a 12- to 18 French tube is used, and for large to giant breed dogs, an 18- to 30 French tube is inserted. When using the tube only for the delivery of liquids, smaller-diameter tubes can be used. Tubes should be flexible yet stiff enough to resist kinking. Commonly, tubes made of red rubber (Sovereign, Sherwood Medical Products, St. Louis, MO), polyvinyl chloride (Argyle feeding catheter, Sherwood Medical Products; Cook Critical Care, Bloomington, IN), polyurethane (Cook Critical Care), Teflon (Cook Critical Care), and silicone (Baxter Health Care Corp., Deerfield, IN) are used. Tubes made of polyurethane or silicone resist the hardening caused by gastric fluids and are recommended if one anticipates that the tube will be used for longer than 1 week. Commercially available tubes frequently require the addition of three to five side holes. These holes can be made carefully using curved scissors. The diameter of the holes should not exceed approximately 20% of the tube’s circumference.
Light general anesthesia is induced and is maintained throughout the procedure. The airway is protected with a cuffed endotracheal tube. The entire lateral cervical region from the ventral midline to near the dorsal midline is clipped and is aseptically prepared for surgery. Usually, the left side is chosen; however, both sides can be used. The procedure is illustrated in Figure 6-10. Curved forceps are inserted into the pharynx and then into the proximal cervical esophagus. Curved Kelly forceps are recommended for use in cats and small dogs. In larger dogs, longer curved Carmalt, Mixter, or Schnidt forceps are recommended. The tips of the forceps are turned laterally, and pressure is applied in an outward direction, thereby tenting up the tissues so the tips can be seen and palpated (Figure 6-10A). A small skin incision (just large enough to accommodate the tube) is made over the tips of the forceps using a scalpel blade, and the tips of the forceps are bluntly forced to the outside (Figure 6-10). In larger animals, as continued pressure is applied, the scalpel blade is used to cut through the thicker esophagus and to allow passage of the forceps.
The selected tube is premeasured and marked using the landmarks listed in Table 6-3. Esophagostomy tubes are usually measured to the level of the xiphoid or ninth intercostal space. Esophagogastrostomy tubes are measured to the thirteenth rib.
The tip of the tube is grasped by the forceps (Figure 6-10C) and is pulled into the esophagus and out through the mouth (Figure 6-10D). The aboral tip of the tube is turned around and is redirected into the esophagus. The tube is then pushed into the esophagus with the aid of the forceps (Figure 6-10E) By retracting the external end of the tube 2 to 4 cm, the tube is felt to “straighten,” and then it passes more easily. The tube is then passed to the premeasured mark. The oropharynx is visually examined to confirm location of the tube in the esophagus. Ideally, the location of the tip should be confirmed with a lateral radio- graph in patients with megaesophagus, esophageal stricture, or any other unusual condition involving the esophagus.
An alternative method of confirming appropriate location of the tube in the distal esophagus involves passing the tube into the stomach. Placement is checked by infusing 30 mL or more of air (using a syringe) and ausculting for bubbles over the stomach region. Once bubbles are heard, the tube is retracted to locate the tip in the distal esophagus. If bubbles are not ausculted in the desired location, a chest radiograph should always be taken to confirm appropriate location.
The tube is secured to the periosteum of the wing of the atlas or deep fascia using nonabsorbable suture (Figure 6-10F). The suture is secured to the tube by using several wraps of the suture around the tube. The tube should also be secured to the skin where the tube exits. Care should be taken not to tighten the suture to the point that it binds the skin to the tube because this may cause irritation and necrosis.
A 4x4 gauze dressing containing chlorhexidine, povidone-iodine, or triple antibiotic ointment is placed over the tube’s exit site in the skin, and a light circumferential wrap is placed. The end of the tube should be capped to prevent spontaneous air or fluid movement through the tube. Commercial feeding tubes are supplied with caps. For most noncommercial tubes, the cap to a hypodermic needle makes a tight fit and easily can be removed.
Care of theTube
A “trap door” is made in the bandage to allow inspection, cleaning, and 4x4 gauze dressing changes (Figure 6-11). The ostomy site should be inspected on a daily basis for the first 5 days after insertion, then every other day for 10 days, then every 3 days thereafter. The ostomy site should be cleaned of exudate with a dilute bactericidal solution suitable for using on wounds or a 50:50 mixture of 3% hydrogen peroxide and sterile saline. Triple antibiotic ointment is then applied, and the 4x4 gauze dressing is replaced.
Procedure for Administration of Fluids and Liquids
Fluids (crystalloids, oral rehydrating solutions, water) and liquid diets can be infused as a constant rate infusion using an administration set and pump similar to that used for intravenous crystalloids. Rates should be set at 1 mL/kg per hour initially. The infusion can be gradually increased by 1 mL/kg per hour until the desired infusion rate is achieved. The infusion rate should not exceed 6 mL/kg per hour.
Fluids, liquid medications, and liquid diets can also be infused slowly using a syringe. The esophagostomy tube should be flushed with water (5 to 60 mL, depending on the size of the tube and patient) after every bolus feeding or every 6 hours in patients fed by constant rate infusions.
Procedure for Administration of Gruel Diets
Gruel diets should be blenderized to ensure that no large particles that may cause an obstruction are infused. If one has any doubt about whether the gruel is liquid enough to pass through the tube, the gruel should be infused through a tube of equivalent diameter first. Boluses should be limited to less than 5 mL/kg initially. Rates can be slowly increased based on patient tolerance. The feeding should be stopped if one sees evidence of salivation, regurgitation, or vomiting or if the animal appears nauseated or uncomfortable. Boluses should not be larger than 25 mL/kg at one time. A bolus should not be given rapidly, and extremely hot or cold materials should not be infused. Immediately after the conclusion of the bolus feeding, the tube should be flushed with water. This helps to prevent the gruel from remaining in the tube where, over time, it may become inspissated and cause an obstruction. A plastic shield or plastic wrap should be used to cover the bandage when infusions are administered to prevent soiling of the dressing.
Procedure for Use for Decompression
Esophagostomy tubes ending in the esophagus can be used to keep the esophagus decompressed in the patient that has poor esophageal motility. Patients with chronic megaesophagus, persistent right aortic arch, or acute megaesophagus are at increased risk for pulmonary aspiration and may benefit from esophageal decompression.2 Decompression is performed by aspirating the tube periodically until all the retained air, fluid, and other material is removed. Esophagostomy tubes ending in the esophagus or stomach can also be used to prevent the recurrence of gastric dilatation in patients recovering from surgery to correct gastric dilatation–volvulus. Studies in human patients have shown that, by preventing passage of air into the stomach, patients return to full oral feeding much more rapidly.5 This finding is assumed, but not proved, to be true in dogs and cats. The tube can be hand suctioned as frequently as needed or connected to a continuous suction device (GOMCO, Allied Healthcare, Buffalo, NY). If viscous or tenacious fluids are suctioned, small volumes of saline or water should be infused into the tube to prevent tube obstruction. An esophagogastric tube can be used for gastric decompression. If gastric secretions are tenacious, saline can be infused initially to break up the secretions before aspiration.
Removal of the Tube
As opposed to gastrostomy tubes, which must remain in place at least several days before removal to allow for a good seal to form between the stomach and the abdominal wall, esophagostomy tubes can be safely removed the same day they are placed. The dressing and the sutures are removed while the tube is held in place. The tube is then occluded and pulled out. The ostomy site should be cleaned, bactericidal ointment should be applied, and a light bandage should be placed around the patient’s neck. The bandage should be removed in 24 hours and the wound inspected. If the ostomy site has not sealed yet, the bandage should be replaced. In patients requiring a new bandage, changes are done every 1 to 2 days until the ostomy site has sealed. This generally takes only a few days.
Long Term Feeding
On occasion, animals require the use of an esophagostomy feeding tube for weeks or months. A fistula usually develops after a few weeks. If the feeding tube needs to be replaced, it is generally a simple procedure because the old tube is removed and a new one is directly fed into the fistula. This usually only requires a local anesthetic block for suture placement. Once these tubes are no longer needed, they are removed as described previously. The fistula closes quickly (within a maximum of a few days), but it may take a week or more to completely heal.
Most complications relate to skin irritation and inflammation. These problems usually can be prevented by ensuring that the skin sutures are not placed too tightly and that the skin is not pinched or folded during suture placement. If the tube is not secured to the periosteum or deep fascia, the tube will retract and move as the animal moves around, leading to possible inadvertent tube removal and significant skin irritation. If mild dermatitis is present, it will usually resolve with time and regular wound cleaning. On occasion, the dermatitis may not resolve until the tube is removed.
By pushing the forceps out in a lateral direction, the esophagus is approximated to the skin. If this maneuver is not performed adequately, the surgeon risks lacerating the external jugular vein as well as creating additional tissue trauma. This complication is rare when proper technique is used. Bleeding from a lacerated jugular vein has occurred in one known patient; this bleeding was controlled easily and definitively using direct pressure.
In extremely debilitated animals, care must be taken to adhere closely to the technique described. Serious complications can result, with dissection of the tube alongside the esophagus, if the tube is not brought out into the patient’s mouth after grasping of the tip of the tube with the forceps. Because the surrounding soft tissues are more easily penetrated, the tube can then course alongside the esophagus instead of in the esophageal lumen. Because the clinician may not be aware of this situation, the tube must be brought out into the patient’s mouth before being passed back into the esophagus.
The use of esophagostomy tubes for both feeding and decom- pression is both a practical and a life saving procedure. More than 500 of these tubes are estimated to have been used to feed dogs and cats since 1988, with beneficial results. The technique has also been used in other mammalian species including the rat, ferret, and monkey. Esophagostomy tubes can also be used effectively in the nutritional support of birds. When comparing the technique with percutaneous gastrostomy tube placement, the use of esophagostomy tubes is less costly, requires no special equipment or special tubes, takes less operative and anesthetic time, is easier to perform, and is associated with fewer complications. No threat of peritonitis exists, and the tube can be removed safely at any time.
- Crowe DT. Use of a nasogastric tube for gastric and esophageal decompression in the dog and cat. J Am Vet Med Assoc 1986;188:1178- 1182.
- Crowe DT. Feeding the sick patient. In: Proceedings of the Eastern States Veterinary Conference. Orlando, FL. 1988;3:95-96.
- Crowe DT, Downs MO. Pharyngostomy complications in dogs and cats and recommended technical modifications: experimental and clinical investigations. J Ain Anim Hosp Assoc 1986; 22:493-496.
- Crowe DT. Nutritional support for the hospitalized patient: an introduction to tube feeding. Compend Contin Educ Pract Vet 1990; 12:1711- 1721.
- Moss G. Maintenance of gastrointestinal function after bowel surgery and immediate enteral full nutrition. ll. Clinical experience, with objective demonstration of intestinal absorption and motility. JPEN J Parenter Enteral Nutr 1981;5:215-220.
Metabolic support has become an integral part of surgical critical care in veterinary medicine.1 Jejunostomy or enterostomy tubes are methods of nutritional supplementation in patients after abdominal surgery. Small animal patients undergoing abdominal surgical procedures are often compromised and are likely in need of nutritional support.
Nutritional support is indicated in patients that are unable to meet nutritional demands by oral consumption of food. Malnutrition can be defined by one or more of the following criteria: anorexia for longer than 5 days, weight loss of more than 10% body weight, increased nutrient loss (i.e., vomiting, diarrhea, protein-losing nephropathy), low albumin, and increased nutrient demands (i.e., surgical stress, sepsis, cancer, chronic infections).
A basic premise “if the gut works, use it” may seem an oversimplification of the benefits of providing nutritional support by physiologic routes (i.e., the gastrointestinal tract versus parenteral administration). In general, the more orad nutrients are placed in the gastrointestinal tract, the better patients are able to assimilate complex diets into essential nutrients. Conversely, bypassing a functional segment of the gastrointestinal tract (i.e., stomach) results in necessary alteration of the dietary composition to accommodate for the loss of the portion of gastrointestinal tract.
Whenever a surgeon enters the abdominal cavity, one question should be answered: Could this patient benefit from a feeding tube? Surgically placed feeding tubes carry little additional operative risk, are economical, and are simple to place and manage; therefore, they pose little risk to the patient while providing a large potential benefit. Special equipment is not required for placement of enteral feeding tubes. The tubes used are 3.5- to 5 French infant feeding tubes at least 36 inches in length. If intestinal surgery is performed, the catheter is placed aboral to the site of surgery. Appropriate diets include commercially available polymeric and monomeric diets. The preferred mode of administration is by slow, continuous rate infusion; however, small frequent boluses can suffice.
Placement of an enterostomy feeding tube may be indicated in any patient undergoing an abdominal operation. The major criteria are a functional small intestine and the need for nutritional support.2,3 Choosing the appropriate method and determining the need for nutritional support are based on applying the least invasive technique that carries the greatest likelihood of success with the least amount of morbidity.
Feeding through an enterostomy tube has induced pancreatic secretion and therefore was previously contraindicated in patients with pancreatitis.4,5 Acute pancreatitis induces a hypermetabolic state with increased caloric and nitrogen demands and at the same time renders the gastrointestinal tract unable to meet these increased needs.4,5 Because the exocrine function of the pancreas is stimulated by the vagus nerve and by release of gastrointestinal hormones in response to food, one can reasonably expect that if the diet is administered into the jejunum, thereby bypassing the cephalic, gastric, and duodenal source of pancreatic stimulation, no significant increase will occur in the exocrine activity of the pancreas.6 Patients with pancreatitis experience modulation of bacterial flora within the intestinal tract and increased bacterial translocation, and they suffer from a negative energy balance. Early alimentation through an enterostomy tube in human patients with pancreatitis results in improved immune status and fewer complications.4,6,7 A jejunostomy tube may allow aggressive nutritional support at an earlier time in the postoperative period. Although these issues are controversial, enteral nutrition is considered an integral part of aggressive treatment of acute pancreatitis in human patients.4,6,7
The major contraindication to the use of a jejunostomy tube is any disorder causing a nonfunctional gastrointestinal tract (i.e., ileus or neoplastic obstruction of the intestine).2,3
From a midline laparotomy incision, a segment of proximal jejunum that is easily approximated to the ventrolateral body wall is isolated. The direction of ingesta flow (orad to aborad) is determined by tracing the bowel segment from a known anatomic landmark (i.e., stomach or duodenum). A 2- to 3 cm longitudinal seromuscular incision is made in the antimesenteric border of the isolated segment of jejunum. At the aboral end of the seromuscular incision, a stab incision is made through the submucosa and mucosa into the lumen of the jejunum (Figure 6-12A). A 5 French Argyle feeding tube (Sherwood Medical Products, St. Louis, MO) is directed through the stab incision aborally into the lumen of the jejunum. Approximately 20 cm of feeding tube is threaded aborally into the small intestine (Figure 6-12B). The seromuscular incision is closed with 3-0 or 4-0 monofilament synthetic absorbable suture in an interrupted Cushing pattern (Figure 6-12C). The surgeon should close this incision in such a manner that the feeding tube is buried in the submucosa of the incision, effectively creating a submucosal tunnel (Figure 6-12, inset). The remaining catheter is exteriorized through a small stab incision in the ventrolateral body wall. Care is taken to select a site that will not result in excessive tension or radial directional changes of the bowel. The enterostomy site is sutured to the peritoneal surface of the adjacent body wall (Figure 6-13). Care is taken to create a watertight jejunopexy on all sides of the enterostomy. The catheter is secured to the skin of the adjacent body wall with a Chinese finger trap friction suture. Abdominal wall closure is routine. A protective bandage is placed on the patient after the procedure, and an Elizabethan collar is used to prevent premature removal of the jejunostomy tube.
Diet Selection, Dose, and Administration
The ideal enteral diet formulation is isotonic, has a caloric density of 1 kcal/mL, a protein content of 4.0 g/100 kcal (16% of total calories), and approximately 30% of calories as fat. Commercially available diets designed for humans are the best diets for small animal patients. Liquid enteral diets can be categorized as polymeric diets or monomeric diets. Polymeric diets contain large molecular weight proteins, carbohydrates, and fats. They require normal intestinal digestion. Most are relatively isotonic, contain about 1 kcal/mL, and are readily available. Monomeric diets are composed of crystalline amino acids as the protein source, glucose and oligosaccharides as the carbohydrate source, and safflower oil as the essential fatty acid source. They are hyperosmolar and expensive. A summary of polymeric and monomeric diets is included in Table 6-4.
For patients with impaired digestive or absorptive function (pancreatitis, enteritis, hepatic disease) or suspected food allergy, a commercial polymeric, enteral liquid diet may be indicated. Patients should be closely monitored for formula intolerance. Jevity (Ross Laboratories, Columbus, OH) is the initial formula of choice, owing to the potential benefits of its fiber content. If the patient becomes intolerant to Jevity, Osmolite HN (Ross Laboratories) should be used. The protein sources of many human products may not provide adequate arginine and sulfur-containing amino acids for cats, and additional protein supplementation is required for long term use.
Monomeric diets are indicated for patients with exocrine pancreatic insufficiency, short bowel syndrome, or inflammatory bowel disease or when polymeric diets are not tolerated. Monomeric diets promote maximal nutrient absorption and minimal digestive and absorptive work. In addition, monomeric diets are less stimulatory for exocrine pancreatic secretion and may have a role in nutritional support of pancreatitis patients.8 To match the caloric density of polymeric formulas, their osmolality must be two to three times higher, a feature that can create disorders of gut motility or fluid balance. Their cost is about seven times more per calorie compared with polymeric formulas. In most cases, a polymeric diet may be tried first, owing to the decreased cost, ease of preparation, and physiologic benefits to enterocyte function.
To determine the dosage of diet to feed, one must first calculate the basal energy requirement (BER, resting energy requirement) based on body weight. The BER is calculated from the following formulas for dogs weighing less than 2 kg: BER (kcal/day) = 70(wtkg0.75)
The following formula is used for dogs weighing more than 2 kg: BER (kcal/day) = 30(wtkg) + 70
After determination of the BER, additional factors can be multiplied depending on the condition of the animal: ER (kcal/day) = BER X 1.25 to 1.5
Protein supplementation should be considered in patients with significant negative nitrogen balance. Commercially available polymeric and monomeric enteral diets are designed for human patients and have significantly lower protein levels. ProMod (Ross Laboratories) is a readily available protein supplement and contains approximately 75% high quality protein (5 g/6.6 g scoop). The guideline for dietary protein requirements in dogs is 5 to 7.5 g/100 kcal, the guideline for cats is 6 to 9 g/100 kcal. Patients with renal or hepatic insufficiency should be reduced to less than 3 g/100 kcal in dogs and less than 4 g/100 kcal in cats.
Feeding can begin immediately in patients with good peristalsis noted at surgery, a secure jejunopexy, and an adequate submucosal tunnel of the feeding tube. However, if uncertainty exists, waiting 18 to 24 hours after placement allows a fibrin seal to form at the jejunostomy site and gut motility to normalize. The calculated volume of diet is gradually administered over 4 days (Table 6-5). These are only guidelines, however, and each patient requires a feeding regimen tailored to fit individual needs.
Complications of jejunostomy tubes include leakage of intestinal contents or diet and are rare; however, they can be devastating.2,3 Therefore, critical placement and monitoring of the tubes in the early postoperative period are imperative. Peritonitis can result from leakage of intestinal contents from the jejunostomy site or from tube displacement into the peritoneal cavity. Clinical signs of peritonitis include vomiting, tachycardia, pyrexia, and abdominal pain. Patients in which a leak is suspected should be evaluated and treated immediately, because progression of clinical signs can be rapid.
Abdominal discomfort, nausea, vomiting, and diarrhea can occur if the diet is infused too rapidly, if a large dose is given, or if the formula is not tolerated by the patient. Decreasing the amount, rate, or concentration of diet infused may alleviate these problems. If gastrointestinal upset persists, one should consider changing the diet or method of nutritional support.
Metabolic complications can occur and include transient hyperglycemia as a result of the insulin resistance present in many critically ill patients. Occasionally, these patients require additional insulin supplementation. Hypophosphatemia has been reported to develop subsequent to enteral alimentation in severely debilitated cats.9 Complications associated with hypophosphatemia include hemolytic anemia and neurologic signs. Investigators have hypothesized that cats in a state of chronic malnutrition have phosphorus depletion despite normal serum phosphorus levels. The institution of enteral alimentation stimulates insulin secretion and cellular uptake of phosphorus and glucose for glycolysis. Phosphorylation of adenosine diphosphate to adenosine triphosphate results in further phosphorus depletion and severe hypophosphatemia. This condition is referred to as the refeeding phenomenon in humans and was first described in World War II victims. One should begin feeding cautiously in debilitated, hypophosphatemic patients.
- Carnevale JM, et al. Nutritional assessment: guidelines to selecting patients for nutritional support. Compend Contin Educ Pract Vet 1991;13:255-261.
- Orton EC. Needle catheter jejunostomy. In: Bojrab MJ, ed. Current techniques of small animal surgery. Philadelphia: Lea & Febiger, 1990:257.
- Moore EE, Moore FA. Immediate enteral nutrition following multisys- temic trauma: a decade perspective. J Am Coll Nutr 1995;10:633 648.
- Marulenda S, Kirby DF. Nutrition support in pancreatitis. NutrClin Pract 1995;10:45-53.
- Freeman LM, et al. Nutritional support in pancreatitis: a retrospective study. J Vet Emerg Crit Care 1995;5:32-41.
- Bodoky G, et al. Effect of enteral nutrition on exocrine pancreatic function. Am J Surg 1991;161:144-148.
- Simpson WG, Marsino L, Gates L. Enteral nutritional support in acute alcoholic pancreatitis. J Am Coll Nutr 1995;14:662-665.
- Guan D, Ohta H, Green GM. Rat pancreatic secretory response to intraduodenal infusion of elemental vs. polymeric defined formula diet. JPEN J Parenter Enteral Nutr 1994;18:335-339.
- Justin RB, Hohenhaus AE. Hypophosphotemia associated with enteral alimentation in cats. J Vet Intern Med 1995;9:228-233.
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