Emergency Nursing

THE EMERGENCY CARE STATION AND RESUSCITATION AREA

Animals frequently come to the veterinarian with emergent and often life-threatening injuries or illnesses. Such demands require that the veterinary facility be set up in a manner in which quick assessment and immediate therapies are possible. Every veterinary practice should contain a centrally located emergency care station and resuscitation area devoted to crisis management. This area should be designed to facilitate rapid triage and treatment. It should be easy to access and have adequate space to accommodate multiple staff members responding to a patient emergency. Emergency drugs and equipment should be stored within easy reach and in designated areas. Equipment and drug inventory of the emergency care station should be checked at each shift change and following each use to ensure that all items are in working order and in adequate supply.

As a minimum, this area should have a source of oxygen, a suction unit, adequate electrical capability, and sufficient lighting. The emergency care station should have a sufficient number of electrical outlets to supply monitoring equipment without the use of excessive extension cords, which can be clumsy, unsafe, and impede the movements of staff. Standard fluorescent lighting can be augmented by well-positioned overhead surgery or examination lights.

In many veterinary practices, oxygen is supplied via anesthetic equipment. Whereas an anesthetic machine provides a familiar means of ventilation and access to sedation (if needed), it can also be a source of catastrophe when errors of anesthetic depth or pop-off valve closure occur. If an anesthetic machine is used, waste anesthetic gas scavenging systems should be available. To prevent the potential problems associated with anesthetic equipment, an oxygen source with a flowmeter and Ambu bag (Figure 33-1) is preferred. Ambu bags are especially useful because they are an easily transported, easily stored, and inexpensive source of artificial ventilation.

Many small practices have suction units with adjustable suction pressure, which are used in surgery (Figure 33-2). The emergency area should have its own suction unit designated and supplied with a variety of suction tips. Suction equipment is often used in the emergency area to clear the airway or endotracheal tube of fluid or debris (mucus, blood, exudates, vomitus, etc.).

1. Maintaining hydration

2. Replacing fluid losses

3. Maintaining IV access and delivering other medications

4. Treatment of shock or hypoproteinemia

5. Increasing urine output

6. Correcting acid-base or electrolyte disturbances

7. Providing nutritional support

PAIN

Detection and assessment of pain is often a challenge in veterinary patients because animals cannot directly communicate their physical condition and there are no pathognomonic signs of pain. Common signs frequently associated with pain include vocalization, depression, anorexia, tachypnea, tachycardia, hypertension, hypotension, pale mucous membranes, aggression, abnormal postures, excess salivation, and dilated pupils. Abdominal pain in particular may be expressed by a classic “praying” or “play bowing” position in which the forequarters are crouched with the abdomen and hindquarters elevated from the ground (Figure 33-22). It is important to note that animals who do not exhibit any of these signs are not necessarily pain free. All trauma victims should be assumed to experience some degree of pain. Many critical illnesses, such as pancreatitis, meningitis, and cancer, are clearly painful problems. Obtunded animals, such as those with severe head trauma, may be unaware of or simply unable to express their pain. Pain management is an important component of veterinary care. Untreated pain causes stress and harmful physiologic changes that prolong recovery.

Staff members trained in the recognition and treatment of pain can help ensure that appropriate analgesia is provided in a compassionate and preemptive manner as part of sound medical care. Many analgesic drugs (i.e., opioids) have cardiac and respiratory depressant effects, which can be dangerous in systemically unstable animals. Nonsteroidal antiinflammatory drugs (NSAIDs) have little effect on the cardiopulmonary system, but may have side effects that affect the gastrointestinal and renal systems. With few exceptions, the systemic administration of analgesics may be safely considered. Regional or local analgesia techniques are useful and should also be considered. For instance, flail chest segments may be treated with a local anesthetic nerve block to reduce pain and allow comfortable breathing.

DISSEMINATED INTRAVASCULAR COAGULATION

Trauma causes tissue and/or vessel injury that is a normal trigger for coagulation (blood clot formation). In most cases, the body's natural homeostatic mechanisms prevent widespread abnormal clotting by balancing clot formation with clot resolution. However, in animals with massive injuries and severe inflammation, the natural balance between clot formation and clot prevention and resolution may be disrupted. When this happens, massive activation of coagulation overwhelms the body's normal regulatory functions and systemic clot formation begins on a widespread scale (instead of confined to a small site of injury). This phenomenon of systemic clot formation and loss of regulatory control is called disseminated intravascular coagulation. In DIC, microclots form throughout the body's capillary network disrupting blood flow to vital organs and causing organ failure (especially in the kidneys, brain, heart, and lungs). Widespread and uncontrolled clotting and clot lysis consume platelets, clotting factors, and regulatory protein, which paradoxically leads to bleeding tendency. DIC is often described as a vicious cycle where clotting and bleeding are occurring spontaneously and simultaneously. Importantly, DIC is always a secondary complication of some other severe disease. Common veterinary causes of DIC include trauma, pancreatitis, heatstroke, cancer, liver disease, immune-mediated hemolytic anemia, and snake envenomation. Clinical signs of DIC are often masked by those of the primary disease or trauma.

In the early stages of DIC, the body is clotting excessively (hypercoagulable), and signs of thrombosis and poor blood flow are prevalent. These signs include unexplained edema, cold extremities, tachypnea, pale mucous membranes, hypotension, and neurologic signs. In later stages of DIC (once clotting factors have been consumed), bleeding tendencies are prevalent. In this phase, clinical signs include unexplained hemorrhage or bruising (hematoma, intraocular bleeding, hemoabdomen or thorax, and excessive bleeding from venipuncture sites). Tiny pinpoint bruises, known as petechiae, commonly appear on the skin (especially along the ventrum and inguinal areas [Figure 33-23]) and mucous membranes (especially the gums and sclera [Figure 33-24]). Large petechial hemorrhages, known as ecchymoses, may form in similar areas (Figure 33-25).

There is no single specific sign or test for the diagnosis of DIC. Instead, the diagnosis is based on supportive lab findings and clinical signs in animals with severe underlying diseases. Decreased platelet count (as a result of platelet consumption) is a consistent and early finding in DIC. Another reliable indicator of DIC is red blood cell (RBC) morphology. Schistocytes or fragmented RBCs are often seen in cases of DIC because fibrin strands span small blood vessels and “rough up” the red cells. This results in distorted borders and red cell fragments that may be seen on the blood smear.

In later stages of DIC, a coagulation profile may detect clotting factor deficiency. A coagulation profile often includes several tests of clotting function, including the prothrombin time (PT), partial thromboplastin time (PTT), and the activated clotting time (ACT). The advent of in-house coagulation time analyzers allows practitioners to conveniently monitor trends in all bleeding times. Measurement of anticoagulant protein (i.e., antithrombin levels) and by-products of clot breakdown (i.e., fibrin degradation products and d-dimers) are also used in the diagnosis of DIC. In cases of DIC, antithrombin levels decrease, whereas fibrin degradation products (FDP) and d-dimers increase. These changes occur as a result of consumption of regulatory anticoagulant protein and accumulation of products of clot breakdown.

Successful treatment of DIC requires resolution of the primary disease. Supportive care with fluid therapy and oxygen supplementation are important. Adjunctive therapy with blood products and anticoagulants may interrupt the self-propagating cycle of coagulation and blood loss. In addition, plasma products and fresh whole blood transfusions help replenish depleted clotting factors, provide anticoagulant regulatory protein, and manage anemia caused by blood loss. Whole blood may also provide some fresh platelets, but it should be noted that these platelets are generally few in number and survive only a short time. Heparin is an anticoagulant commonly used in conjunction with plasma. If heparin is used at moderate to high doses, it should be tapered before discontinuing therapy. Unfortunately, treatment of DIC is often unsuccessful because many underlying conditions cannot be rapidly resolved and it is exceedingly difficult to restore the body to a state of healthy equilibrium once compensatory mechanisms are overwhelmed. DIC carries a poor to grave prognosis.

SHOCK AND THE SYSTEMIC INFLAMMATORY RESPONSE SYNDROME

Animals with poor blood flow and impaired oxygen delivery to tissues are said to be in a state of “shock.” This clinical syndrome is caused by circulatory failure (despite its name, “shock” has no association with electrocution). Untreated shock is rapidly fatal because imbalance between tissue oxygen demand and oxygen delivery causes tissue injury, organ failure, and death.

In the early stages of shock, impaired perfusion triggers natural compensatory mechanisms (vasoconstriction, increased heart rate, increased cardiac contractility) that maintain blood pressure and increase cardiac output. This early or compensated phase of shock is known as the hyperdynamic phase or compensatory phase. Clinical signs during this phase relate more to adaptive physiologic responses than to perfusion failure. The clinical signs in this phase include increased heart rate and respiratory rate, rapid CRT, injected mucous membranes, and increased pulse pressure. These findings can be subjective and easily missed. During this phase, the weakness, depression, and altered consciousness associated with later stages of shock are either absent or mild. The most recognizable clinical signs of hyperdynamic shock include brick red mucous membranes and bounding pulses.

Close monitoring performed by alert staff members is important because early recognition and treatment may prevent further progression of shock.

Uncompensated or hypodynamic shock ensues if there is progressive underlying disease or if compensatory mechanisms and treatment fail to restore normal blood flow and oxygen delivery to the body. During uncompensated shock, cardiac output and systemic blood pressure are inadequate. Blood flow is preferentially distributed to vital organs (brain, heart) at the expense of other tissues. This shunting of blood exacerbates the oxygen deficit and fluid imbalance in other tissues and can lead to organ failure. The commonly recognized clinical signs of uncompensated shock are associated with circulatory failure and include hypotension (low blood pressure), rapid heart rate, weak pulses, prolonged CRT, pale mucous membranes, hypothermia, overt weakness, depression, and loss of consciousness. Eventually, prolonged hypoxia results in vascular paralysis, systemic vasodilation, and fulminant cardiovascular collapse. This terminal phase of shock is irreversible and rapidly fatal.

Shock is a state of emergency that is associated with many causes. When subdivided according to underlying cause, general categories of shock are hypovolemic shock, distributive shock, cardiogenic shock (including obstructive shock), and septic shock. Hypovolemic shock is the most common form of shock in small animals. In this form of shock, perfusion failure results from a reduction in circulating blood volume caused by bleeding, dehydration, or effusive fluid loss (i.e., abdominal fluid accumulation). Distributive shock is associated with maldistribution of blood flow associated with pathologic vasodilation. In this syndrome, pooling of blood in capillaries and veins results in a decrease in effective blood volume (regardless of intravascular volume or cardiac output). Common causes of distributive shock include trauma, heatstroke, envenomation, and anaphylaxis. As its name implies, cardiogenic shock is associated with decreased cardiac output. Cardiogenic shock can occur from heart failure resulting from many primary heart diseases, such as cardiomyopathy, valvular disease, and cardiac arrhythmias. A subset of cardiogenic shock known as obstructive shock is associated with obstruction of blood flow. Common causes of obstructive shock include pericardial disease, heartworm disease, pulmonary hypertension, and pulmonary thromboembolism.

Shock that is caused by infection is referred to as septic shock or sepsis. Septic shock can be triggered by primary infectious diseases, but it can also occur with opportunistic infections. Extensive tissue damage associated with severe disease (such as trauma, heatstroke, envenomations, and pancreatitis) creates areas of poorly perfused and devitalized tissue that provide a setting for opportunistic bacterial growth. Infection in such areas is difficult to combat because disruptions in blood flow prevent systemically administered antibiotics from reaching the site of infection. The local inflammatory response triggered by some bacterial toxins may help clear infections. However, in animals with widespread injury or tissue damage, an exaggerated inflammatory response develops that leads to uncontrolled systemic inflammation. This systemic inflammation precipitates a state of shock by inducing vasodilation, vascular permeability, poor cardiac function, and activation of coagulation. Hyperglycemia may occur in the early phase of septic shock as a result of the effects of stress hormones on metabolism. In later stages, hypoglycemia is predominant as glucose is consumed by both bacteria and body demands.

In the absence of infection, a systemic inflammatory response syndrome (SIRS), which parallels septic shock, can be triggered by any critical illness where systemic inflammation is a problem. As is the case with other forms of shock, SIRS patients may go through an early hyperdynamic phase followed by an uncompensated or hypodynamic phase. In the hyperdynamic phase, circulatory collapse is temporarily held at bay by compensatory mechanisms that increase cardiac output and maintain blood pressure. During this phase, bounding pulses and brick red mucous membranes may be noted. Clinical manifestations of SIRS include circulatory changes, thermoregulatory dysfunction (fever or hypothermia), depression, tachypnea, and DIC. Recognition of septic shock and SIRS in individual patients is based on clinical findings, supportive history, and laboratory data. Fulminant septic shock and SIRS are both associated with a syndrome of multiple-organ dysfunction (MODS). Kidney failure and liver failure are particularly common.

TREATMENT OF SHOCK

Treatment of underlying diseases is important in the management of animals in shock. For instance, animals with septic shock should receive appropriate antibiotics, and animals with traumatic bleeding may need a blood transfusion. Unfortunately, management of underlying conditions takes time and is often only partially effective. Therefore animals diagnosed with SIRS or other states of shock should receive treatment based on triage priorities similar to those animals in cardiac arrest. Treatments are focused on restoring oxygen delivery and perfusion to the tissues. Oxygen supplementation should be provided immediately via face mask during initial resuscitation efforts. This allows a staff member to continually monitor the patient during treatment. Nasal oxygen catheters or an oxygen cage (or canopy) may also be used. Preventing circulatory collapse is the highest treatment priority during management of shock syndromes. This is accomplished primarily with aggressive fluid therapy to restore effective vascular volume and blood pressure. Shock dosages of crystalloid solutions are 90 ml/kg/hr in the dog and 45 to 60 ml/kg/hr in the cat. When using crystalloid therapy in the dog, shock doses of fluid can be administered in quarter-dose increments. For dogs, a quick formula to calculate a “quarter shock dose” is to take the body weight in pounds and multiply by 10. For example, a volume of 400 ml is an appropriate quarter shock dose for a 40-lb dog. This “quarter shock dose” is given over 15 minutes, and the animal is reevaluated. For cats, the same formula can be used; however, you must divide the dose in half (or use the body weight in kilograms instead of pounds).

A pressure bag may be helpful for administering large fluid volumes over a short period of time (Figure 33-26). Colloid fluids are also used to restore vascular volume to patients in shock. Colloid doses depend on the type of colloid used. Blood products, such as whole blood and plasma, may also be used in some cases during resuscitation efforts. Hemoglobin-based oxygen carrying solutions made from polymerized bovine hemoglobin sometimes have limited commercial availability. When administered IV, these solutions act like a colloid, but also have oxygen-carrying capacity. These features make them popular resuscitation fluids for some clinicians.

As noted before in the section on fluid therapy, all fluid doses are simply guidelines. Fluid therapy should be administered “to effect,” which means that shock fluids are administered until monitoring parameters indicate that treatment has had the desired effect. Monitoring may indicate that higher or lower doses may be necessary. Appropriate fluid therapy often results in normalization of heart rate, blood pressure, and mucous membrane color. Additional monitoring parameters include CVP, urine output, blood lactate levels, hematocrit, and TP. It should be noted that unusually large crystalloid and colloid fluid volumes may be necessary to maintain vascular volume in unstable (“shocky”) animals. Intensive patient monitoring is critical.

Animals with refractory hypotension, despite appropriate fluid therapy, may be candidates for use of vasopressor drugs (dopamine, dobutamine).

Veterinarians generally give these drugs as a constant rate infusion to increase vascular tone and cardiac output. At low dose levels in dogs, dopamine selectively increases renal perfusion, which can be helpful in restoring urine output. At moderate doses, beneficial effects on systemic blood pressure are prevalent. However, at high doses, dopamine has an adverse effect on renal perfusion and may worsen kidney failure. Dobutamine primarily increases blood pressure by enhancing cardiac contractility and cardiac output. Both dopamine and dobutamine can be associated with cardiac arrhythmias, and ECG monitoring and careful auscultation may be helpful.

PERFUSION FAILURE AND REPERFUSION INJURY

Reperfusion injury is a cellular injury that develops as blood flow returns to an area or tissue previously deprived of perfusion. During cardiopulmonary arrest and shock, oxygen-starved tissues develop an anaerobic metabolism and are depleted of cellular energy stores. These conditions alter certain enzyme systems and destabilize white blood cell (WBC) membranes. Upon reestablishment of oxygenation and perfusion (as occurs with successful resuscitation and fluid therapy), the altered enzyme systems generate harmful molecules called oxygen free radicals. At the same time, membrane-damaged WBCs release inflammatory mediators that contribute to the reactive environment. The oxygen free radicals and inflammatory mediators cause inflammation and vessel injury leading to thrombosis and edema. These effects are collectively called “reperfusion injury.” Reperfusion injury may result in systemic disorders, such as DIC, SIRS, and multiorgan dysfunction. Following resuscitation from shock or cardiopulmonary arrest, all vital organ systems can be affected by reperfusion injury and inflammation. For this reason, all systems must be monitored closely and supported, with special attention paid to basic life support systems.

BASIC LIFE SUPPORT

In basic life support, A is for airway. Staff members responding to a potentially arrested animal should note if the animal is breathing. If respirations are absent or weak, the mouth should be opened and the oropharynx examined for possible obstruction. Common sources of airway obstruction include respiratory secretions, aspirated vomitus, blood, ingested foreign material, and mass lesions (hematomas, neoplasia, etc). If obstruction is noted, the airway should be cleared with suction or manual removal of foreign material. Caution is indicated to prevent being bitten, although most animals in partial or full states of arrest have limited or no capacity to bite. A sponge forceps and gauze may be helpful in clearing some exudates. Once the airway is cleared, staff members should note whether these steps have stimulated the animal to breathe.

If the animal does not begin to breathe, the patient must receive ventilation assistance. B is for breathing. Mouth-to-nose resuscitation may be performed by sealing the lip margins and blowing into the animal's nose. This method requires no special equipment and will deliver about 16% oxygen. This level of oxygenation is inadequate and should only be done temporarily until a higher supply of oxygen can be provided. Mouth-to-nose resuscitation efforts carry some risk to caregivers treating animals with potentially zoonotic diseases. Endotracheal intubation and ventilation with an Ambu bag in room air provides 21% oxygen. The best method of assisted ventilation is endotracheal intubation and delivery of 100% oxygen from an oxygen source. Ideally, animals should be intubated in lateral recumbency to prevent elevation of the head and positional changes that impair cerebral blood flow in arrested animals. Many times, however, intubation is performed more rapidly and accurately using a laryngoscope with the animal in sternal recumbency. Rapid intubation is imperative, and delays or failure can be catastrophic to further resuscitation. Following intubation, the tube should be secured with a gauze tie. If intubation is not possible, a narrow orotracheal catheter or transtracheal cannula is sometimes useful. However, a large-bore airway is preferred and may require a surgical tracheostomy. During assisted ventilation, the first two breaths administered should be long breaths lasting a full 2 seconds, followed by patient assessment. In some instances, restoring an open airway will lead to recovery of spontaneous respirations by the patient. If not, the animal should be manually ventilated at a rate slightly higher than the expected normal. Assisted ventilation should expand the chest by 30%, with a slightly longer expiration than inspiration. If the breathing circuit contains a manometer (i.e., most anesthetic machines), a pressure of 10 to 20 cm of water should be obtained with each breath.

Acupuncture is a method that can be attempted to treat respiratory arrest when other efforts have failed. The acupuncture point is Governor Vessel 26 (VG 26) of Jen Chung. This point is located at the nasal philtrum at the level of the ventral edge of the nares (Figure 33-27). A 25-gauge needle is applied to the bone at this point and then twirled to induce breathing.

C is for circulation. Once the airway is established and ventilation provided, circulation must be assessed by palpation of pulses (or apex heartbeat) and auscultation of the heart. Peripheral pulses are nonpalpable when the mean blood pressure is less than 60 mm Hg. An apex heartbeat may be indistinguishable when the pressure is less than 40 mm Hg. It is important to note that some animals suffer respiratory arrest without cardiac arrest. Improper chest compressions can stress the patient and precipitate cardiac arrest in some of these cases. However, once cardiac arrest has been confirmed, chest compressions should be started immediately.

Positioning of the animal during compressions depends on the animal's size, the shape of the chest (barrel chest versus deep and narrow chest), and the caregiver's ability to deliver adequate compressions. There are two theoretic models to explain forward motion of blood during CPCR. In small animals or those with a narrow chest conformation, chest compressions (during closed-chest CPCR) and direct cardiac massage (during open-chest CPCR) apply forces to the heart that mimic the normal heart mechanics. This is known as the “cardiac pump.” In large animals and those with barrel chests, changes in chest conformation limit the direct effect of chest compressions on the heart. In these cases, increased intrathoracic pressure during compression results in forward blood flow from the heart, which serves as a passive blood reservoir. This is referred to as the “thoracic pump” model and is thought to play a significant role in medium- to large-size animals during CPCR. To optimize the cardiac and thoracic pumps, animals less than 15 lb (7 kg) should be placed in lateral recumbency. Animals greater than 15 lb may be placed in either lateral or dorsal recumbency. The point of compression (hand placement) for the cardiac pump is located directly over the heart (Figure 33-28). For the thoracic pump, the point of compression is located at the widest part of the chest.

Effectiveness of CPCR should be assessed by palpating for a pulse and evaluating the mucous membrane color. If available, an ECG can be extremely beneficial at this point to assess the heart and also to evaluate the effectiveness of resuscitation efforts. Traditionally a pulse and an electrical waveform on ECG should be generated with each compression. If available, end-tidal carbon dioxide (capnography) is a reliable monitor of ventilation and perfusion. Other monitoring equipment (ECG, pulse oximeter, venous blood gas analysis, and lactate levels) can provide useful quantitative information. If monitoring suggests the CPCR effort is inadequate, a change in technique may be required to improve effectiveness. Common changes to the CPCR effort include changing places with another team member, changing the animal's position, and/or altering your compression technique. Ventilation and chest compressions may be interposed or administered simultaneously. Administering ventilations and chest compressions simultaneously enhances the thoracic pump by increasing intrathoracic pressure during the compression (systole) and increasing venous return and atrial filling during relaxation (diastole). Once compressions have begun, a solid rhythm can develop if the breaths are administered simultaneously to the compressions. This rate should be 120/min for animals less than 15 lb and 80 to 100/min for animals greater than 15 lb. Interposed abdominal compressions assist in directing blood in the lower half of the body back toward the heart (via increased intraabdominal pressure) and may be administered by another team member (Figure 33-29).

Open-chest CPCR is mostly indicated in animals with chest trauma (flail chest, pneumothorax, and diaphragmatic hernias) because of the interference of such injuries on closed-chest compressions. In this method of CPCR, the chest is surgically opened on the left side at the fifth intercostal space, and compressions are applied to the heart from the apex to the base. Care should be taken not to twist the heart, which can occlude major vessels. Nontraumatic occlusion of the descending aorta during open-chest CPCR may improve coronary and cerebral blood flow. Open-chest CPCR is only beneficial if initiated early in the resuscitation effort. The decision to perform an emergency thoracotomy for open-chest CPCR should be made within 2 minutes of cardiopulmonary arrest.

ADVANCED LIFE SUPPORT

Advanced life support includes interpretation of an ECG and administration of drugs based on cardiac output, blood pressure, and the presence of arrhythmias. These steps in resuscitation are important only after basic life support has been established (i.e., the animal is being ventilated, and adequate circulation is provided). If the animal has responded to resuscitation efforts and has a perfusing rhythm, advanced life support may not be necessary. Unfortunately, in many cases, life-threatening arrhythmias and hypotension are common and require treatment.

Common drugs used in CPCR include atropine, epinephrine, naloxone, lidocaine, and magnesium chloride or sulfate. Proper use of an ECG allows recognition of specific arrhythmias so that appropriate drugs may be administered and patient response to therapy can be gauged. There are three basic arrhythmias seen during an arrest. These include asystole (“flat line”) (Figure 33-30), nonperfusing rhythms (electromechanical dissociation or pulseless electrical activity) (Figure 33-31), and ventricular fibrillation (Figure 33-32). IV drug doses are listed in Table 33-1. In many cases, asystole and nonperfusing rhythms are preceded by progressive bradycardia. Bradycardia can be a sign of imminent arrest and should prompt notification of other staff members. Sinus bradycardia may be treated with atropine as the animal is monitored. During asystole, both electrical and mechanical cardiac activity has stopped. Asystole is treated with atropine and/or epinephrine with repeated doses administered if no response is observed. Electromechanical dissociation (EMD) is a common terminal arrhythmia in cats and dogs. The hallmark of this arrhythmia is the presence of ECG complexes with no cardiac contractions to generate a pulse (hence the synonym, pulseless electrical activity [PEA]). This rhythm can have a diverse appearance, but often mimics a ventricular arrhythmia with wide bizarre QRS complexes occurring at a slow rate. EMD is treated with naloxone, megadose atropine, or epinephrine.

Ventricular fibrillation is a common arrhythmia in people suffering myocardial infarction. It also occurs in cats and dogs during cardiopulmonary arrest. In dogs, ventricular fibrillation may be preceded by rapid ventricular tachycardia (Figure 33-33), especially when multifocal ventricular beats or R on T phenomenon are present. When ventricular fibrillation is diagnosed, it must be converted as soon as possible for resuscitation to be successful. Conversion of this arrhythmia may be attempted before initiation of basic life support. Treatment of choice for this arrhythmia is electrical defibrillation using an electrical defibrillator. If this is not available, chemical defibrillation may be attempted using drugs such as magnesium chloride. A strong precordial thump is potentially effective as a last resort. Electrical defibrillators should only be used by specially trained personnel (Figure 33-34). Tips for appropriate use of an electrical defibrillator include:

Alcohol should never be used near defibrillator paddles because of the risk for fire. The recommended dose is 2 to 4 J/kg. Initially, use a setting at the lower end of the dose range, and repeat or double the dose if no response is seen. Open-chest defibrillation requires specific paddles and a modified dose (usually one tenth of the transthoracic dose).

Drugs administered during CPCR may be ineffective as a result of poor perfusion and failure of the drugs to reach their target tissues (primarily the heart). A central vein catheter (i.e., jugular catheter [Figure 33-35]) is the CPCR drug administration route of preference during closed-chest CPCR. These catheters facilitate delivery of the drug(s) directly to the heart or its close proximity.

The next best route is intratracheal administration. This route of administration takes advantage of alveolar membranes, which have a large surface area and receive a high blood flow separated by a narrow diffusion barrier. An acronym to remember which drugs can be administered by the intratracheal route is LEAN (lidocaine, epinephrine, atropine, and naloxone). Intratracheal drugs are administered via a catheter passed through the endotracheal tube (Figure 33-36). Insertion of drug into the intratracheal catheter must be followed by a flush of air or saline to ensure drug deposition into the airways. A deep breath administered via manual ventilation helps to further distribute the drug. Drug doses administered by the intratracheal route may be rapidly estimated by doubling the IV dose. Small drug volumes may require dilution for effective administration. Drug uptake by the pulmonary circulation is impaired by conditions such as pulmonary edema, which negates this as a suitable route. Good communication between team members performing CPCR is imperative because a vigorous chest compression delivered at an inopportune moment may result in exhalation of medications administered intratracheally.

Drugs may be administered via a peripheral IV catheter (i.e., cephalic or saphenous vein catheters) if a central line or intratracheal access is unavailable or contraindicated. All drugs administered peripherally must be followed with a good flush of saline solution to ensure delivery into the circulation and toward the heart. Intraosseous catheters and intralingual injection are another means of peripheral administration. Placement of an intraosseous catheter (or intralingual injection) in the arrested patient can be rapid and does not need to interrupt the CPCR attempt. Intralingual drug doses are usually double the standard IV drug dose. The last route for drug administration is intracardiac. This is chosen last because of the challenge of hitting a flaccid heart, the need to stop CPCR to administer drugs, and the risk of damaging the heart. However, in open-chest CPCR, intracardiac drug administration is preferred. Usually, one tenth of the IV dose is injected directly into the left ventricle.

Crystalloid fluids or colloids may be beneficial if hypovolemia was a predisposing factor of the arrest or to compete with peripheral vasodilation and support blood pressure. However, fluid support of peripheral circulation is not a high priority during CPCR. Fluids may be contraindicated during CPCR because fluid volumes are poorly tolerated by a failed cardiac pump and volume overload or overhydration easily develops.

The decision to initiate CPCR is made on a case-by-case basis according to the wishes of an informed pet owner. Many critically ill animals face an already grave prognosis. Following an arrest and successful resuscitation, the risk of rearrest and subsequent death is high. “Do not attempt resuscitation” orders may be indicated after discussion with the clinician and pet owner. Assessing the patient's risk of arrest and addressing the desires of the client are important early on. If a patient does not respond to CPCR within 20 minutes, continuation of the resuscitation effort is unlikely to succeed. Successful resuscitation rates in veterinary medicine are approximately 10%. Fortunately, certain patients do respond to basic and advanced life support. Many of these cases have a reversible disease process and/or a treatable cause of arrest. A written record of everything done during the CPCR should be made for the team to learn from and for the client record.

Regardless, the real work begins following successful resuscitation.

PROLONGED LIFE SUPPORT

Proper postresuscitation management has two primary focuses. First, primary factors leading up to the arrest should be identified and treated. Second, problems caused by the arrest and the trauma of the resuscitation effort should be recognized and managed.

The central nervous system is particularly sensitive to injury during states of shock or cardiopulmonary arrest. In health, cerebral blood flow is locally controlled and maintained by reflexes that maintain blood flow and protect the brain from hypertension and volume overload. This autoregulation of cerebral blood flow is lost during arrest, leaving the central nervous system susceptible to further injury during and after resuscitation. Cerebral ischemia and reperfusion injury resulting in nerve cell death is a serious complication of cardiopulmonary arrest and resuscitation. Initially, neurologic examinations should be done hourly. PLR, responsiveness to stimulation, respiratory pattern, motor responses, and motor postures should be noted. Normal pupil size and PLRs are positive signs. Slow PLRs, anisocoria, and pinpoint pupils that are nonresponsive to light are progressively guarded neurologic indicators. Nonresponsive bilaterally dilated pupils indicate severe brain damage and a poor prognosis. Recent administration of atropine during the arrest should be ruled out as a cause of dilated nonresponsive pupils. Brainstem damage should be suspected in patients lacking a corneal reflex or swallow, or gag, reflex. Breathing patterns also reflect brainstem function, and erratic breathing patterns and periods of apnea (breathlessness) are poor prognostic indicators.

The cardiovascular system is clearly “ground zero” during cardiac arrest. In addition, abnormalities associated with other organ failures may further impact heart rhythm and blood pressure. Changes in heart rhythm, vascular tone, and cardiac output predispose to systemic hypotension and rearrest. Consequently, it is imperative to continuously monitor the ECG and blood pressure in the postresuscitation period. Accurate blood pressure measurements may be obtained using direct and indirect methods. Direct blood pressure measurement is ideal, but requires an arterial catheter and specialized equipment. Indirect blood pressure measurements may be obtained with Doppler or oscillometric methods. Arrhythmias causing clinical signs or hemodynamic compromise should be treated. Oxygen therapy has relatively few complications in the short term and is a useful treatment.

Acute kidney failure is a common problem associated with states of shock and cardiac arrest because the kidneys are particularly susceptible to damage caused by hypovolemia and hypotension. Consequently, monitoring of kidney and electrolyte parameters should be performed frequently in the postresuscitation period. Decreased urine production is associated with severe kidney failure. Urine output should be monitored hourly for at least 24 hours after an arrest.

Strict attention should be paid to maintaining adequate hydration and blood pressure. Hypotension is a common complication, and the mean arterial blood pressure should be maintained within normal limits to guarantee adequate kidney blood flow. Veterinarians sometimes use an “ins and outs” fluid therapy plan to maintain fluid balance when kidney function is in question. In such cases, administered fluid doses are balanced with calculated fluid losses. Appropriate fluid therapy will maintain hydration without causing volume overload. CVP, repeated PCV/TP, and body weight measurements are useful indicators of volume status. Hemodialysis and peritoneal dialysis are available at certain specialized treatment centers.

Primary respiratory disease is a common factor leading up to cardiopulmonary arrest. Respiratory complications of arrest and resuscitation include pulmonary edema as a result of congestive heart failure, noncardiogenic edema associated with hypoxia, and pulmonary thromboembolism. Vigorous chest compressions during resuscitation efforts may also result in pulmonary contusions, rib fractures, atelectasis, and/or edema. These injuries must be addressed in the postresuscitation treatment plan. Optimal respiratory therapy may require ongoing oxygen supplementation, ventilation support, and monitoring of arterial blood gas analysis. If blood gas analysis is not available, monitoring should include pulse oximetry and/or capnography.

Blood glucose concentrations should be monitored frequently because many patients that arrest develop hypoglycemia. Normal blood glucose concentrations should be maintained by adequate supplementation when indicated. Hyperglycemia should be prevented.

MEDICAL TREATMENTS IN THE POSTARREST PERIOD

Many variables affect the outcome and management of an arrested patient. These variables include the underlying diseases and reasons for arrest and the experience and equipment available to the resuscitation team. Difficulties facing the arrested patient and the syndromes associated with the postarrest period are well known. However, disagreement exists regarding treatment methods, treatment priorities, and which therapies result in the best outcome. All therapies of postarrest patients are somewhat controversial because of an inability to create a standardized model for conclusive studies. Many drugs may be useful in the postresuscitation patient, and the list continues to grow. The following is a brief review of medical therapies and the rationale behind their use.

Mannitol is an osmotic diuretic sometimes used in the management of acute renal failure and cerebral edema. The mannitol molecule resides in the vascular space and draws water from the interstitial spaces between cells, thereby decreasing edema and expanding the vascular volume. As mannitol is excreted by the kidneys, its osmotic effects “pull” water with it into the urine. Mannitol is also a free-radical scavenger, which may aid in the treatment of reperfusion injury. Some caution is advised in the use of mannitol because it can exacerbate volume overload. At high doses, mannitol may be nephrotoxic, and it is contraindicated in hypovolemic animals.

Furosemide (Lasix) is a potent diuretic that is commonly used in the treatment of pulmonary edema and acute kidney failure. The diuretic effects of furosemide increase urine output and may enhance the effects of mannitol. In cases of pulmonary edema, furosemide causes volume contraction, which decreases edema formation and hastens resolution of edema.

Glucocorticosteroids (i.e., dexamethasone sodium phosphate, prednisolone sodium succinate, and methylprednisolone sodium succinate) are extremely controversial as to appropriate use. They may be beneficial in stabilizing cellular membranes, thereby decreasing the release of membrane-derived inflammatory mediators. Methylprednisolone sodium succinate is a free-radical scavenger. As such, it is one of a few drugs capable of rapid action against the oxygen free radicals created during reperfusion injury.

Dobutamine, a synthetic catecholamine, improves the contractility of heart muscle and may be administered to maintain mean arterial blood pressure. The related drug, dopamine, can be used to increase renal perfusion in canine patients at low doses and to increase systemic blood pressure at higher dosages. The use of dopamine in feline patients is controversial and limited to the systemic blood pressure effects of the drug.

Sodium bicarbonate is used in the treatment of severe life-threatening acidosis. However, caution is warranted because overzealous bicarbonate therapy is both harmful and easy to accomplish. Unpredictable acid-base disturbances and other problems can be attributed to overdoses. Blood pH must be monitored via serial venous or arterial blood gas analyses.

Lidocaine is an antiarrhythmic drug used to treat ventricular tachycardia. Care should be taken to accurately interpret the ECG tracing because this drug may be contraindicated in ventricular escape and isolated premature ventricular complexes. The latter are common arrhythmias observed in the immediate postresuscitation period. Although abnormal, these rhythms do provide functional blood perfusion in many cases. Abolishing a perfusing ventricular rhythm with lidocaine may result in development of a nonperfusing rhythm and death.

Because all body systems are affected by cardiopulmonary arrest, a whole body approach to monitoring and therapy is required. For this reason, successfully resuscitated animals are among the most critical and labor-intensive patients. The prognosis for long-term survival is often poor. Those patients successfully resuscitated require constant monitoring and support for several days.

• Hemotrophic feline mycoplasma (formally hemobartonella)

• Feline leukemia virus (FeLV)

• Feline immunodeficiency virus (FIV)

• Bartonella spp (cat scratch disease, et al)

• Heartworm disease

• Brucellosis

• Common tick-borne pathogens (Ehrlichia, anaplasmosis, neorickettsiosis)

• Babesia canis and Babesia gibsoni

• Leishmania

RESPIRATORY SYSTEM

Care should be taken to minimize stress in animals with respiratory difficulty (dyspnea). Oxygen supplementation is an important treatment in some animals and can be provided via blowby, nasal cannulae, face mask, oxygen canopy, or oxygen cage or incubator. In all patients (especially those with known respiratory disease), it is important to note the rate, pattern, and effort of breathing. Obstructive airway disease (i.e., laryngeal paralysis or tracheal foreign material) is often characterized by a slow and deep respiratory pattern with harsh or whistling upper airway noise. Restrictive respiratory disease (i.e., pleural effusion, chest wall injuries) often manifests as rapid and shallow breathing. Respiratory disease may be detected by monitoring respiratory rate, mucous membrane color, and respiratory effort. A significant abdominal component or effort during inspiration is abnormal. All lung fields and the upper airways should be ausculted. Additional sounds with inspiration or expiration should be noted and significant changes recorded in the medical record and reported to the veterinarian. Fluid and air in the pleural space often result in muffled heart and lung sounds. Percussing the thoracic wall during auscultation may aid in determining whether air or fluid is in the pleural space. Certain lower airway disorders, such as pneumonia and pulmonary edema, are characterized by crackles during each phase of respiration.

Monitoring equipment, such as a pulse oximeter (Figure 33-48) and capnograph, can provide additional information to the veterinary team. These monitors are frequently used for anesthesia monitoring, but can also be used for continuous monitoring of critically ill patients. Pulse oximetry uses an infrared sensor to count the pulse rate and provide a noninvasive measure of arterial hemoglobin-oxygen saturation (usually in percent). Infrared sensors are made to clip onto the tongue, lip margin, or flank. Rectal probes (with sanitary covers) for recumbent patients are also available. It is important to note that the pulse oximeter reading is a hemoglobin-oxygen saturation value and is not identical to the blood oxygen content or arterial oxygen partial pressure (PaO₂). The structure and biochemistry of hemoglobin allows saturation to remain high despite some significant changes in blood oxygen content. The pulse oximeter is a good crisis prevention tool, but does not provide fine details about the oxygenation and ventilation status of the patient.

If the pulse oximeter detects a problem with oxygen saturation, a blood gas analysis and oxygen supplementation may be ordered. In normal patients, the oxygen saturation should remain above 95%. Poor perfusion at the probe site, hypothermia, and movement will impede the sensor's ability to obtain an accurate reading. If the pulse oximeter sensor triggers an alarm, manual palpation of the pulse, auscultation of the heart, and check of the mucous membranes should be performed immediately before adjusting the equipment.

Capnography also uses infrared technology to estimate the carbon dioxide concentration of expired air. By incorporating a capnometer into the breathing circuit of intubated animals, the end-tidal carbon dioxide (carbon dioxide concentration at the end of expiration) can be measured (Figure 33-49). The end-tidal carbon dioxide is an estimate of PaCO₂ (arterial carbon dioxide levels), which can be useful in monitoring the efficiency of mechanical ventilation. Capnometry can also be used to confirm endotracheal intubation and to detect airway occlusion. If an endotracheal tube is misplaced into the esophagus or occluded by exudates, end-tidal carbon dioxide will be negligible. Additionally, capnographs have been used to assess the efficacy of resuscitation efforts during CPR. Successful resuscitation is expected to result in progressively increased end-tidal carbon dioxide values as forward blood flow returns carbon dioxide from the tissues.

Arterial blood gas analysis provides specific data on the oxygenation, ventilation, and acid-base status of the pet. Normal ranges for arterial blood gas values are published elsewhere. Values for oxygen and carbon dioxide are expressed as a pressure (PaO₂ or PaCO₂) in units of millimeters of mercury (mm Hg). Arterial blood samples may be drawn from the femoral artery, dorsal pedal artery, or lingual artery by palpating the pulse and guiding the needle into the artery by tactile sensation (Figure 33-50). The femoral artery and dorsal pedal arteries are the most common sites of arterial blood sampling. Lingual artery samples may be obtained in anesthetized or comatose animals, but they should be avoided because of the risk for inducing a large oral hematoma, which can obstruct the airway and interfere with swallowing. Acquiring arterial blood samples may be too stressful for some patients. By definition, ventilation is determined by the carbon dioxide level (PCO₂) on an arterial blood gas analysis. Hypoventilation (failure to “blow off” carbon dioxide) causes an increase in blood carbon dioxide. Hypoventilation occurs with certain respiratory problems and can cause acidosis. Hyperventilation (“blowing off carbon dioxide”) results in decreased carbon dioxide. This situation may occur with respiratory disease or as a compensatory response to metabolic acidosis. Oxygenation is indicated by the PaO_2, and a low value indicates hypoxemia. Signs of hypoxemia include cyanotic (blue) mucous membranes (Figure 33-51), weakness, rapid heart rate, increased respiratory rate, and increased respiratory effort.

The difference between alveolar (lung) oxygenation and arterial oxygenation is a calculated indicator of the efficiency of oxygen transfer to the blood by the lung. In patients that are hypoxemic, this alveolar-arterial gradient (A-a gradient) calculation can provide a single value to assess and monitor the animal's oxygen exchange. The alveolar oxygen content (A) is calculated by the alveolar gas equation: A = (BP − 47) 0.21 − PCO₂/0.8, where BP is the barometric pressure (760 mm Hg at sea level), 47 is the vaporization pressure of water (a constant physical property of water), and 0.21 is the oxygen percent of room air. This formula is only used when the patient is breathing room air. In the formula above, PCO₂ is obtained from the arterial blood gas analysis, and 0.8 is the respiratory exchange quotient (a mathematic constant). At elevations near sea level, the formula simplifies to A = 150 − PCO₂/0.8. The arterial oxygen content (a) is the PaO₂ obtained from the arterial blood gas analysis (in mm Hg). To finish calculating the A-a gradient, the arterial oxygen content (a) is subtracted from the alveolar oxygen content (A). In animals with normal oxygen exchange, the A-a gradient is less than 10 mm Hg. As a general rule, an A-a gradient more than 30 mm Hg is an indication for oxygen therapy.

To determine how responsive the patient is to oxygen therapy, an arterial sample is collected while the patient is on supplemental oxygen, and a second arterial blood gas analysis is performed. The original A-a calculation is invalid for patients receiving oxygen therapy. Instead, oxygenation is assessed by evaluating the ratio of arterial oxygen to inspired oxygen (PaO₂/FIO₂). PaO₂ is obtained from the blood gas analysis. FIO₂ is the fraction (percent) of inspired oxygen being supplemented in decimal form. For example, a dog receiving 100% oxygen would have an FIO₂ of 1.0. Most of the time, animals receiving oxygen therapy by mask or nasal catheters have an FIO₂ of close to 40% (FIO₂ = 0.4). By definition an animal's disease is responsive to oxygen if the PaO₂/FIO₂ value is greater than 250. As a general rule of thumb, the PaO₂ should be approximately five times the FIO₂.

CARDIOVASCULAR SYSTEM AND PERFUSION/HYDRATION

Monitoring of common physical examination findings can be key to detecting serious health problems. Technicians should be comfortable using a stethoscope and familiar with the basic principles of cardiac auscultation and pulse palpation. The detection of a new heart murmur, irregular heartbeats (arrhythmia), and/or changes in heart or lung sounds can be signs of an impending crisis. For example, animals that are “overhydrated” may develop crackles characteristic of pulmonary edema. Factors that decrease sound intensity include obesity, pleural or pericardial effusion, and hypovolemia. Likewise, pulse character and quality are important clues to the hydration status and stability of the critically ill pet. If the mean arterial blood pressure is less than 60 mm Hg, pulse strength will be diminished and it will be difficult to palpate pulses. Common sites to palpate a pulse are the femoral, dorsal pedal, or lingual arteries. Pulse rates should parallel the heart rate, rhythm, and quality. Comparing the pulse rate and heart rate during auscultation can aid in detecting “pulse deficits” that may be produced by irregular heartbeats.

Monitoring of hydration status and peripheral perfusion is crucial in sick animals. Helpful monitoring parameters include texture and color of mucous membranes, skin turgor (Figure 33-52), CRT, quantitation or close estimation of urine output, thoracic auscultation, pulse rate and character, serial body weights, and temperature (both core body temperature and peripheral limb temperature). Physical signs of overhydration include serous nasal discharge, chemosis (conjunctival edema; see Figure 33-10), peripheral edema, body cavity effusion, and weight gain. Noninvasive monitoring equipment, such as an ECG and indirect blood pressure equipment, can provide additional information regarding the pet's cardiovascular status. Patients at greater risk for fluid imbalance, such as those with heart disease, pneumonia, and renal disease, may require more invasive monitoring techniques. Invasive monitoring techniques include PCV, TP, direct blood pressure, and CVP measurement. Trends in observations of both subjective and objective parameters provide the most information. Diligent record keeping and consistency in monitoring techniques are imperative.

COAGULATION STATUS

Assessment of an animal's coagulation status may be helpful in assessing unexplained bleeding and detecting DIC. Evaluation of a blood smear may detect RBC morphology changes and allow an estimation of platelet numbers. The presence of schistocytes or fragmented RBCs along with a decrease in platelet numbers may be seen with DIC, splenic tumors, and severe heartworm disease. A high percentage of echinocytes can be seen in animals suffering from snake envenomation.

A normal platelet count is between 200,000 and 500,000 platelets per microliter in most species. Although a complete platelet count is advised, rapid assessment or confirmation of platelet numbers may be obtained by estimating the platelet numbers on a blood smear. This can be done by counting the number of platelets per high-power field and multiplying by 15,000. A better estimate is obtained if several high-power fields are assessed and averaged. Platelet clumping, which is common at the edge of blood smears, can make platelet estimates invalid.

The buccal mucosal bleeding time (BMBT) evaluates platelet function and is often used as an in-house screening test for von Willebrand's disease (an inherited disease effecting platelet function). A commercially available lancet device (Figure 33-57) is used to make a standardized incision (specific size and depth) into the buccal mucosa on the inside of the cheek (Figure 33-58). Following this incision, the blood flowing from the cut is blotted away with filter or absorbent paper to permit visualization of the site without disturbing the incision (Figure 33-59). The time required until bleeding stops is the BMBT, which represents the time for formation of an initial platelet plug. Normal time for the dog is less than 4 minutes. Following completion of the test, renewed bleeding at the site can occur if the incision is disturbed or if the animal has other coagulation problems, but such bleeding does not affect the reported time.

RENAL/URINARY SYSTEM

Ideally, all hospitalized animals should have a urine sample obtained for urinalysis. Urine specific gravity is important information in determining urinary tract health and in interpreting changes in kidney parameters and the urine sediment. Because urine specific gravity and blood values for blood urine nitrogen (BUN) and creatinine can be affected by fluid therapy and medications (diuretics, such as furosemide and mannitol); pretreatment blood and urine samples should be obtained. However, if such samples cannot be readily obtained, fluid therapy should not be delayed, particularly if the animal is dehydrated or unstable. Urine output should be estimated in all animals and recorded in the medical record. In critical cases, close monitoring of urine production may be performed via urine collection through indwelling urinary catheters or specifically designed cages that allow urine to drain through a grate to be collected. An indwelling urinary catheter attached to a sterile urine collecting bag is a comfortable, clean, and accurate way to monitor urine output (Figure 33-60). Because catheters may clog, kink, or change position, urine collection equipment should be regularly inspected and replaced if faulty. Minimum urine production is 2 to 4 ml/kg/hr. Inadequate urine production may be an indication for changes in fluid therapy and should be interpreted with other patient data (i.e., body weight, CVP, etc.) and reported to the veterinarian.

CENTRAL NERVOUS SYSTEM

Unfortunately, few objective parameters are available to monitor the central nervous system. Serial physical and neurologic examinations are perhaps the best means of detecting new or ongoing problems. Changes in mentation, responsiveness, level of consciousness, and respiratory patterns may indicate changes in neurologic status and should be reported.

Likewise, pupillary size and responsiveness to light are important signs to monitor. Increased intracranial pressure results in a syndrome of brainstem and cerebral herniation whereby intracranial pressure changes cause compression of the brain against solid connective tissue and bone. Early signs of increased intracranial pressure include mental dullness, tachypnea, tachycardia, and dilated pupils. Later signs include bradycardia, fixed pinpoint pupils (Figure 33-61), seizures, coma, and death. Early recognition and intervention are key to the management of this syndrome. Animals at risk for brain herniation include those with head trauma, hydrocephalus, brain tumors, and encephalitis. Herniation can also be a complication of some diagnostic procedures, such as spinal fluid collection and myelogram.

ABDOMINAL CAVITY

Each patient should receive a daily physical examination. Palpation of the abdomen may detect abdominal distention resulting from fluid accumulation, organ enlargement, and intestinal gas. Abdominal pain can be detected by the presence of discomfort, splinting, or vocalization. Animals with unexplained abdominal pain or distention warrant further investigation and monitoring.

Abdominocentesis is a procedure to confirm the presence of abdominal fluid and to collect samples for fluid analysis. During this procedure, the abdomen is clipped of hair and aseptically cleaned with surgical scrub. A needle attached to a syringe is inserted into the abdomen near the umbilicus, and gentle suction is applied to withdraw fluid (Figure 33-62). Care is taken not to redirect the needle blindly within the abdomen because this may result in unnecessary pain and potential injury to abdominal organs. Fluid collected via abdominocentesis may be submitted for culture and cytology. Abdominocentesis is commonly performed to detect active hemorrhage, infection (peritonitis), ascites, uroabdomen, and neoplastic effusions. If only a small amount of fluid is suspected, a four-quadrant tap may be performed in which the four areas of the abdomen centered around the umbilicus are sampled. If no sample is obtained and abdominal fluid is still suspected, a diagnostic peritoneal lavage (DPL) may be performed. The DPL is a modified procedure for collecting small amounts of abdominal fluid. In this procedure, the animal is prepared as described earlier. Before fluid withdrawal, a volume of 10 to 20 ml/kg of sterile warm isotonic saline is instilled into the abdomen. After a brief moment to adjust the animal's position and allow the instilled fluid to mix with abdominal contents, standard abdominocentesis is performed. The added fluid may dislodge adherent debris and mix with isolated fluid pockets that can then be collected during standard abdominocentesis.

THORACIC CAVITY

Animals with cardiac and respiratory disease often have similar clinical signs (namely, coughing, labored breathing, exercise intolerance, and collapse). Unfortunately, these signs can occur with many different causes, including upper airway obstruction, pulmonary edema, pleural effusion, pneumonia, primary heart disease, and thoracic neoplasia. A thorough physical examination often directs diagnostic tests to differentiate respiratory from cardiac problems. Muffled heart and respiratory sounds often indicate fluid or air in the pleural space (the space between the lungs and chest wall). The presence of pleural fluid or air causes respiratory problems as a result of physical interference with normal lung expansion. Crackles heard during the different phases of breathing usually indicate pulmonary disease, such as pneumonia or pulmonary edema. In general, crackles indicate relatively severe pulmonary disease.

Chest x-rays are commonly used to confirm, classify, and document the presence of cardiac and respiratory diseases. If thoracic fluid is documented on x-rays or clinically suggested during examination, thoracocentesis to remove the fluid is indicated. Thoracocentesis is usually performed with the animal comfortably restrained in sternal recumbency. During this procedure, one or both sides of the chest are clipped of hair (in the area of rib spaces 7 through 9) and cleaned with surgical scrub. A butterfly catheter (or needle with extension set) is attached to a three-way stopcock and syringe (Figure 33-63). The needle is advanced into the chest along the cranial aspect of a rib to prevent discomfort and bleeding associated with nerve and vessel bundles located behind each rib. As the needle is inserted into the chest, an assistant controls the syringe and maintains the stopcock in the closed position to prevent environmental air from entering the chest during inspiration. Once the needle is positioned in the pleural space, the stopcock is opened to allow the assistant to suction fluid from the pleural space into the syringe. If a large amount of fluid must be removed, the stopcock is closed to the patient as the syringe is repeatedly emptied. Samples for culture, cytology, and fluid analysis are often obtained early during the procedure. Thoracocentesis may be performed as an emergency diagnostic procedure in animals with severe respiratory distress and suspected pleural fluid (Figure 33-64). Air can also accumulate in the pleural space. This condition is referred to as a pneumothorax and often results from traumatic rupture of airways or from severe lung disease. In such cases, air leaks outside of the lung, but is trapped inside the chest. Thoracocentesis (as described earlier) may be used to remove air from the pleural space. If fluid or air continues to accumulate in the chest, “chest drains,” or thoracostomy tubes, may be placed that allow repetitive or continuous drainage of air and/or fluid from the pleural space (Figure 33-65).

URINARY OBSTRUCTION

Urethral obstruction is a serious and common emergency in both cats and dogs. Causes of such obstruction include urinary stones, tumors, trauma, and/or inflammation. Animals that are unable to urinate develop metabolic abnormalities quickly and may develop secondary kidney damage. In some cases, unrecognized urinary obstruction results in permanent kidney failure and even rupture of the urinary system and leakage of urine into the abdomen. Consequently, untreated urinary obstruction is fatal.

Diagnosis of urinary obstruction is based on history, physical examination, laboratory tests, and diagnostic imaging (x-rays or ultrasound). Complete urinary obstruction is often preceded by signs of lower urinary tract disease, such as straining to urinate, painful urination, blood in the urine, or lack of urination. Obstruction of urine flow results in a large, firm, swollen, and painful bladder that can be palpated during physical examination. Affected animals often have pelvic or abdominal pain, lethargy, and dehydration. Veterinarians and technicians should be familiar with the historical and physical examination findings associated with urethral obstruction so that a rapid diagnosis and intervention may be made.

Failure to excrete urine results in rapid onset of severe fluid and electrolyte imbalances. Common laboratory abnormalities include elevated potassium (hyperkalemia), elevated kidney parameters (azotemia; increased BUN and creatinine) and metabolic acidosis. Acidosis in animals with urinary obstruction is due to the accumulation of lactic acid (from poor perfusion during dehydration) and from accumulation of uremic acids (metabolic by-products usually excreted by the kidneys). Metabolic acidosis may be recognized by closely scrutinizing the blood pH, bicarbonate levels, and total carbon dioxide levels in lab reports. Venous blood gas analysis is helpful in determining the acid-base status and may be needed to obtain some of this information. Hyperkalemia is often severe and can be acutely life threatening because it triggers cardiac arrhythmias and can be a cause of cardiac arrest. Significant elevation of serum potassium results in fairly unique ECG changes that may allow early recognition of the problem. These changes include: (1) bradycardia, (2) “spiked” T waves, (3) shortened R-wave amplitude, (4) blunted or missing P waves, and (5) prolonged QRS interval (wide, short complexes). Astute veterinarians and technicians should be aware of these classic ECG features that may allow for early recognition of hyperkalemia (Figure 33-66).

X-rays and ultrasound can be used to confirm the presence of urinary obstruction, but are more important for determining the cause of obstruction. Care should be taken to include all of the urinary tract in the x-ray study. In male animals in particular, the x-rays should include the perineal and penile urethra.

X-rays are a common and readily available means to detect most stones in the bladder or urethra. X-rays must be of sufficient quality to detect small stones. However, not all stones are easily detected by radiography because some stones appear less opaque than others. In some cases, nonradiopaque (“lucent”) stones may only be detected by contrast x-ray procedures or ultrasound. Ultrasound provides a noninvasive detailed evaluation of the urinary tract. In particular, ultrasound is useful in the detection of bladder masses and in the evaluation of the upper urinary tract. Evaluation of the upper urinary tract (kidneys and ureters) is important because stones and tumors may occur in this area. In addition, ultrasound may detect congestion and dilation of the ureters (hydroureter) and the urine collecting system of the kidneys (hydronephrosis), both of which may be caused by lower urinary obstruction.

Regardless of the cause, emergency treatment of urinary obstruction has two priorities: (1) relieve the urinary obstruction and (2) correct the fluid and electrolyte changes.

Relief of urinary obstruction generally requires passage of a urinary catheter. Urinary catheterization can be difficult in obstructed patients. Repetitively draining urine via cystocentesis should be avoided or used as a therapy of last resort because the turgid and inflamed urinary bladder is at risk for rupture. In some instances, cystocentesis may be chosen as an emergency procedure to relieve the immediate tension on the bladder. This temporary relief may facilitate later catheterization. In patients where urinary catheterization is not successful, urinary diversion via urethrostomy or a cystostomy tube may be considered.

Placement of urinary catheters in male and female dogs and cats is described later. Proper urinary catheterization generally requires two persons. One person places the catheter in a sterile manner while an assistant is present to restrain and properly position the animal. A catheter of appropriate length and diameter should be selected. The catheter must be long enough to reach the bladder. Generally a large diameter catheter is desirable to allow for higher flow rates during drainage or flushing of the catheter. In some instances, a small diameter catheter may be chosen to bypass an area of partial urinary obstruction.

TOXIN INGESTION

Dogs are well known for randomly ingesting things in their environment during play or from curiosity. Cats are normally more discriminate and fastidious in their eating habits, but will still ingest noxious substances in their environment. These behaviors make toxin ingestion a common presenting complaint or telephone call to veterinary offices. See also Chapter 35. A brief clinical review of common toxin ingestions seen in emergency practice is presented here.

Veterinary technicians are often the first point of contact when pet owners call about toxin ingestion. Although we should be knowledgeable about the most common toxins, we should also be careful about what information and advice is presented by telephone. Information gleaned from telephone conversations is notoriously incomplete, and the consequences of error can be severe. When in question, the best advice to pet owners is to bring the pet to the veterinary practice for examination. Consequently the real purpose of a client telephone call is to gather information relevant to the animal's treatment before an appointment. The following information should be obtained:

Manufacturer labels contain important information and should be saved and brought to the veterinary practice with the pet.

The product label may be used to identify any harmful ingredients and may have instructions to follow after unintended ingestion or overdose. Many products have a toll-free number for questions. This type of information helps the veterinary team anticipate the needs of the patient and plan for the animal's arrival. Additional information may be obtained from local government-sponsored poison control centers. Information from these centers is species specific for humans and may not be entirely applicable to animals. Because of similarities in mammalian physiology, however, information about human poison exposure is still helpful in most cases of small animal toxin ingestion. Fortunately, animal-specific poison control centers are readily available and are a wealth of species-specific information. Animal poison control centers often charge a reasonable fee for this service.

Induction of vomiting (also called emesis) is a common treatment for the ingestion of harmful substances. Timely induction of vomiting may eliminate the toxin from the body before it can be absorbed and exert any harmful effects. In most instances, vomiting should be induced within 4 hours of ingestion. This is a flexible time limit that roughly corresponds to normal gastric emptying time. Gastric emptying time may be prolonged in some instances. Once a substance has exited the stomach and is being absorbed in the small intestine, induction of vomiting is unlikely to have a therapeutic benefit.

Importantly, induction of vomiting is not a cure-all for toxin ingestion and should be cautiously considered like any other treatment. As a general rule, ingestion of caustic substances and petroleum products should not be treated by vomiting because of the risk of further injury or aspiration. Emesis is contraindicated after ingestion of many household products, including bleach, lye, gasoline, and oil (and many other acids and strong alkali agents). In addition, induction of vomiting is strongly contraindicated in animals with altered awareness because sedated, comatose, or tranquilized animals are at high risk for developing aspiration pneumonia. Vomiting may be induced by a variety of means. In dogs, apomorphine is a potent emetic agent that directly stimulates the vomiting centers of the brain to cause vomiting. The drug can be compounded into a variety of forms. The drug is absorbed across the conjunctival membranes of the eye, and small tablets of the drug may be placed in the conjunctival sac until vomiting has occurred. Subsequently the drug can be flushed from the conjunctiva to terminate the vomiting episode. Apomorphine is not effective in cats. Because they do not respond to apomorphine, induction of vomiting in cats is challenging. Xylazine (a sedative drug) often stimulates vomiting in cats. This emetic effect must be balanced with the risk of sedation-related aspiration when xylazine or similar drugs are used for this purpose. A number of household remedies, such as hydrogen peroxide and syrup of ipecac, have been used to induce vomiting in both cats and dogs. Hydrogen peroxide administered orally will reliably result in vomiting as a result of bitter taste and gastric irritation. The widespread use of hydrogen peroxide as a topical antiseptic makes it a convenient over-the-counter and generally available emetic. However, hydrogen peroxide can be dangerous in sensitive or overdosed animals because careless use of hydrogen peroxide can result in severe hemorrhagic gastroenteritis. Syrup of ipecac induces vomiting by inciting gastric irritation and by stimulating the vomiting centers of the brain. This once common home remedy has become less popular because induction of vomiting at home has some risk and may delay other more appropriate medical treatments.

When vomiting is not possible but gastric emptying is desired, gastric lavage is sometimes an option. Unfortunately, gastric lavage does not always result in effective or timely gastric emptying and carries some inherent risks. Therefore whenever possible, gastric emptying via induction of vomiting is nearly always preferred over gastric lavage. For the gastric lavage procedure, animals are sedated and an endotracheal tube is placed to secure the airway and prevent aspiration. Subsequently a stomach tube is placed, and the stomach contents are removed via lavage and drainage. This procedure is time consuming (which may delay other treatments), and the sedation required for gastric lavage adds risk. Additionally, residual lavage fluid sometimes merely adds to gastric volume and raises the risk of vomiting and aspiration. If gastric lavage is indicated, it is best performed by experienced personnel.

In addition to gastric emptying, treatments for nonspecific toxin ingestion may also include administration of adsorbents and cathartics. Adsorbents bind toxins to prevent absorption and facilitate their excretion. Cathartics are medications that affect GI transit and speed fecal elimination of toxins. The most common adsorbent is activated charcoal. Activated charcoal comes in a variety of commercially available formulations. Oral suspension liquid preparations are easy to administer directly by mouth (alone or mixed with food) or by stomach tube (Figures 33-69 and 33-70). Activated charcoal may bind intestinal nutrients and other orally administered medications, but does not appear to have any common clinically significant side effects. Activated charcoal is excreted in the feces and will result in dark or black discolored stool that may be mistaken for melena. Some formulations of activated charcoal contain sorbitol as a cathartic. Sorbitol is a sugar alcohol that draws water into the large bowel, resulting in a laxative effect.

The treatment and clinical evaluation of specific toxins is beyond the scope of this chapter. However, a few common toxicities warrant brief discussion.

INTRODUCTION

An emergency is defined as a sudden, urgent, usually unexpected event or occurrence requiring immediate action. In equine practice, an emergency is usually the result of the patient's response to an environmental stimulus (fight or flight), the curious nature of horses, obstacles in the environment, or the unique nature of the patient's anatomy (natural areas of narrowing of the horse's gastrointestinal tract). Less commonly a veterinary emergency may be the result of human malice, negligence, or neglect. When an emergency arises, the veterinarian and the staff must act with the appropriate amount of care, compassion, and in an expeditious manner. In some cases, such as long bone fractures or severe abdominal pain, the decision to treat or to humanely euthanize the patient is the first issue to be addressed. When treatment is attempted, it is often initiated with minimal objective data regarding the patient's condition. These are circumstances where the veterinarian and the staff must react rather than ponder the situation. Guess and reassess is often the way in which emergencies are initially approached until the patient, client, and situation are stabilized. An experienced, well-trained veterinary technician is essential in the organization of instruments and material, initial assessment, and triage of the horse seen on an emergency basis. Depending on the presenting complaint and the patient's condition, several criteria must be assessed and met:

Each of these criteria will be addressed in detail repeatedly throughout the remainder of this chapter.

The purpose of this chapter is to review the veterinary technician's role in the management of emergencies specific to equine veterinary medicine. This includes the initial examination and determination of the patient's condition, the equipment, instruments, and materials needed to manage equine emergencies, and finally, this chapter will review the use of IV fluids, blood, and blood products in equine emergency medicine.

EMERGENCY PREPARATION

The chief complaint or reason the patient is seen will dictate what preparations are made before the patient's arrival or what equipment is to be carried into the field if examination is undertaken in an ambulatory situation. It is essential that all needed equipment be organized before arrival of the patient. This includes needles, syringes, catheters, IV fluids, fluid delivery systems, oxygen tanks and regulators, IV fluids, and any medications, such as NSAIDs and sedation. In addition, any equipment that may need to warm up, such as x-ray processes, should be turned on before arrival of the patient. Emergency and first-aid equipment can be stored conveniently in already prepared carts or emergency kits that can be organized based on emergency needs (Figure 33-75), or the material can be organized onto carts before arrival of the patient (Figure 33-76).

Emergency drugs that are stored in carts or tots should be arranged in alphabetic order, and a list of the drugs, their indications, dosage, and typical volumes administered to an average 450-kg adult or 50-kg foal can be laminated and stored with the drugs for quick reference (Figure 33-77). Table 33-2 is a list of emergency drugs commonly used in equine practice.

TRANSPORT OF THE INJURED OR CRITICALLY ILL HORSE

Considerable misconceptions exist as a result of the lack of specific published recommendations on the transport of critically ill or injured equine patients. Too often veterinarians are reluctant to recommend and clients are apprehensive about transporting patients to the hospital for definitive care because of the fear of injury during transport. These fears are often unfounded, and the risk of transport seldom if ever outweighs the benefit of appropriate treatment. Although trailering is an athletic event with the horse needing to balance its weight, sometimes on three legs through turns and sudden stops, most horses withstand transport well as long as appropriately prepared and hauled.