Nursing Care and Rehabilitation Therapy for Patients with Neurologic Disease

Nursing Care and Rehabilitation Therapy for Patients with Neurologic Disease

Mary Tefend CampbellJanice L. Huntingford

Introduction4, 34

Caring for patients with neurologic disease can be extremely challenging, especially if those patients are nonambulatory or only weakly ambulatory. Management of the recumbent dog or cat involves primarily supportive care. A treatment protocol should be implemented at the onset of paresis or paralysis in order to prevent or lessen the severity of complications such as pulmonary atelectasis or pneumonia, poor gastric motility, fecal retention, urinary bladder damage and urinary tract infections, decubital ulcers, muscle atrophy, joint stiffness, pain, inadequate nutritional intake, and patient depression or lethargy. As many of the nursing techniques applied to stable patients require no special equipment or advanced training (e.g. passive physiotherapy, bladder management), clients can become adept at performing basic nursing care for dogs and cats that are expected to be recumbent for long time periods. Successful nursing care of patients with neurologic dysfunction is dependent upon cooperation and communication between clinicians, nurses, and pet owners.

Respiratory care1, 4, 8–10, 16, 19, 22, 24, 27, 34

A key factor in the treatment of the recumbent patient is prevention of respiratory dysfunction. It is of utmost importance to assess the respiratory patterns of the patient throughout the course of treatment, to auscultate frequently, and to consistently monitor oxygen saturation via pulse oximetry. Subtle changes in the respiratory system can occur within a very short time period in recumbent patients; such changes can subsequently lead to rapid deterioration of pulmonary function. Although pulse oximetry can be a useful tool in monitoring a patient’s oxygenation status, it should never be substituted for a thorough physical examination. Patients must be turned every 4 hrs or kept in a sternal position when at all possible. Management should also include sling therapy if the patient is orthopedically stable (see “Rehabilitation therapy” section below). Respiratory complications in the recumbent patient can be life threatening and must be addressed immediately. Stress should be minimized in the patient in respiratory distress, and oxygen therapy should be administered via the most effective route. Clinicians should also be aware that severe cervical lesions or lower motor neuron (LMN) conditions such as polyradiculoneuritis may compromise respiratory function. Blood gas analysis and capnography are useful in monitoring the efficacy of ventilation, particularly the pressure of carbon dioxide in arterial blood (PaCO2), as poor intercostal muscle movement may impair exhalation. Neck bandages or external splints placed for cervical support (e.g. atlantoaxial instability) may also impair ventilation (Fig. 20.1).


Figure 20.1 In patients requiring a neck bandage or splint, it is important to monitor ventilatory effort by assessing chest expansion and ETCO2 and SpO2

   Pain may also cause changes in respiratory patterns. Thoracic radiographs should be included periodically for the recumbent patient to monitor pulmonary health. Treatments for respiratory dysfunction secondary to recumbency include oxygen therapy, nebulization and coupage therapy, positioning techniques, including sling therapy, and mechanical ventilation if respiratory complications are severe. The goals of treatment for respiratory complications include prevention of respiratory secretions and accumulation, expansion of atelectatic lungs, improved oxygenation, elimination of CO2, and patient comfort.

  1. Positioning techniques

    Animals that are recumbent for any reason should be turned every 4 hrs from left to right lateral recumbency to prevent atelectasis or accumulation of lung secretions. If atelectasis or pneumonia is present, the patient should be propped sternally or positioned with the most normal functioning lung down to improve ventilation. Postural drainage can also be accomplished by positioning the patient in a head-down posture (20° from horizontal) for 15–30 min every 4 hrs to increase mucus drainage and prevent accumulation of debris within the trachea (tracheal “plugs”). It is important to supervise a patient closely during treatment in order to intervene if the patient becomes stressed or if secretions obstruct the airway. Postural techniques are recommended by the authors in conjunction with nebulization and coupage therapy. In the patient with brain injury, preventative measures to control increased intracranial pressure (ICP) include positional techniques to promote venous drainage (elevating the head). Avoiding jugular venipuncture, providing adequate sedation/analgesia, and preventing hyperglycemia and hyperthermia are also beneficial (see Chapter 8). Patients with head injury may also have cervical trauma; careful movement of the head is recommended during triage.

  2. Nebulization and coupage therapy

    Nebulization and coupage treatment for pneumonia are effective means to move secretions from smaller airways to larger airways and to elicit coughing to remove such secretions. Acetylcysteine may be added to the nebulizer as a mucolytic agent. The recommended dose is 5 mL added to the nebulizer (for a 10% solution) every 12 hrs at 4–6 L/min of oxygen. It is important to practice proper technique and to minimize stress to the patient during therapy. Patients with severe pneumonia may require ventilator therapy with increased positive end-expiratory pressure (PEEP) setting to ensure expansion of collapsed alveoli. Bronchodilators such as theophylline derivatives (e.g. aminophylline) or beta-agonists (terbutaline) can also be used to combat aspiration-induced bronchoconstriction. Nebulizers may be used to deliver bronchodilators (albuterol), antimicrobials (typically aminoglycosides such as gentamicin at a dose of 7 mg/kg with 8 mL saline), or regular saline for supportive therapy (Table 20.1). Aerosolized agents for the treatment of pneumonia are in addition to traditional intravenous antibiotic administration. The nebulizer and nebulizer tubing must be kept clean and not used for multiple patients in order to prevent iatrogenic respiratory infection.

    Table 20.1 Inhalation therapy: Nebulized bronchodilators.

    Drug Species Dosage* Duration and frequency
    Albuterol Cat, small dog 0.5 mL + 4.0 mL saline 1–3 min sid–qid
    Albuterol Dog (10 kg) 1.0 mL + 4.0 mL saline 1–3 min sid–qid
    Albuterol Dog (20 kg) 2.0 mL + 4.0 mL saline 1–3 min sid–qid
    Albuterol Dog (30 kg) 3.0 mL + 4.0 mL saline 1–3 min sid–qid

    *Albuterol supplied as 2.5 mg/3 mL inhalation solution.

    1. Materials needed

      • handheld nebulizer

      • oxygen tubing

      • anesthetic/oxygen machine

      • 5–10 mL sterile saline

      • facemask.

    2. Technique

      • Place the patient in sternal recumbency in a comfortable position. Sling therapy can be implemented (see “Physical therapy” section below) at the time of nebulization.

      • Instill saline +/– medication into the nebulizer compartment; do not invert the chamber.

      • Connect the oxygen tubing to the oxygen system and nebulizer. Turn the oxygen to 4–5 L/min to ensure proper mist flow.

      • Attach a facemask to the nebulizer and apply to the patient.

      • Nebulize for 5–10 min with saline; medications 1–2 min. If the patient does not tolerate the mask, the nebulizer unit may be held closely to the nose or mouth.

      • Coupage the patient after nebulization by cupping the palms of the hand (or contoured to the curvature of the patient’s thorax) and striking the chest rhythmically by alternating between the right and left sides of the chest (Fig. 20.2).


        Figure 20.2 Technique for coupage therapy.

         Coupage should be forceful enough to elicit a cough but not enough to allow the patient to become stressed. Correct hand positioning and location of coupage are more important than the force applied. Coupage should be rhythmic and in a circular motion back and forth over the diseased areas. For puppies, small dogs, and cats, the three middle fingers should be used for coupage rather than the palm of the hand. If stable, deep-chested dogs may be rolled onto their back for coupage, facilitating drainage from ventral lung lobes. It is of the utmost importance to watch respiratory patterns and mucous membrane color during coupage, as it may become necessary to coupage in stages to prevent an increase in respiratory effort. It is also important not to feed a patient shortly before nebulization and coupage therapy, to avoid aspiration of food into the airway. A suctioning device should be readily accessible if the patient has heavy mucus secretions that may clog the airway and compromise ventilation, particularly in the patient with megaesophagus. Contraindications for nebulization and coupage therapy include flail chest, pneumothorax, severe soft tissue trauma to thoracic area, and thoracotomy with chest tube placement. Nebulizing via an oxygen cage is not recommended, as aerosolization is not direct and is usually inadequate. If the patient is oxygen dependent, nasal oxygen can be utilized during therapy, if not already in use. If possible, a short walk immediately after nebulization may help stimulate a cough.

  3. Oxygen therapy

    Oxygen therapy is critical to the patient exhibiting respiratory dysfunction. Oxygen should be administered in the most effective, least stressful route. Flow-by oxygen supplementation should be provided immediately to postsurgical, postseizure, or head trauma patients in distress while the airway is assessed. It is the opinion of the authors that nasal oxygen is far superior to oxygen cages, although oxygen cages can provide the patient immediate respiratory relief until nasal therapy can be instituted. Transtracheal or nasal-tracheal oxygen can also be administered if the patient has stenotic nares, severe facial trauma, or laryngeal paralysis not responding favorably to nasal oxygen. Nasal prongs (intended for human use) can be used for larger dogs. The prongs are placed in the nares and secured behind the dog’s ears. The tubing can be taped or sutured in place. Nasal prongs generally require higher flow rates than nasal catheters.

    1. Nasal oxygen tube placement

      1. Materials needed

        • soft, polyvinyl catheter (red rubber)

        • petroleum jelly for lubrication

        • local anesthetic

        • suture material

        • oxygen tubing

        • oxygen canister

        • distilled water.

             For facilitation of catheter placement, the patient should be restrained in a comfortable position. The size of the catheter should be large enough to provide adequate oxygen delivery. The authors recommend using a size 5 French tube for a cat or small dog and a size 8–10 French tube for larger dogs. Transparent feeding tubes are not recommended, as the length is excessive and the tube can be mistaken for an intravenous line.

      2. Technique

        • Instill two to three drops of anesthetic solution into the designated nare (2% lidocaine or proparacaine).

        • Lubricate the end of the red rubber catheter, and slide the catheter into the nasal cavity via the ventral meatus to the level of the medial canthus. In the cat, insert the catheter in a ventromedial direction. In the dog, initially insert the catheter in a dorsomedial direction, then ventromedially.

        • The catheter should be brought up over the head either between the eyes or (preferably in cats) along the mandibular area and anchored in place beginning at the nostril by sutures.

        • Oxygen tubing is connected to the catheter by either an adapter or secured with a Chinese finger lock suture.

           Oxygen tubing (the proximal end) is attached to an infusion bottle filled with distilled water. Oxygen is then delivered through the water and into the tubing connected to the catheter. If in-house oxygen is not available, a portable oxygen machine can be utilized by connecting the tubing to the gas inlet valve. An oxygen flow rate of 100 mL/kg/min will provide an oxygen concentration of approximately 40%. Gastric distension may occur if the conflation rate is too high. Nasal oxygen catheters should be replaced and changed to the opposite nare every 48 hrs due to mucus accumulation. Elizabethan collars should be placed on the patient to keep the catheter from being removed.

    2. Transtracheal oxygen catheter placement

      Transtracheal oxygen may be provided if nasal oxygen is not suitable (e.g. nasal occlusion due to hemorrhage). Transtracheal catheters are less irritating than nasal catheters, and lower flow rates can be used to achieve equivalent oxygen concentrations achieved with nasal catheters; however, placement is technically more difficult. Long-term use of transtracheal oxygen should be avoided, as damage to tracheal mucosa may result. Aseptic technique is critical in placing intratracheal oxygen catheters. The bandage should be changed every 24 hrs and the catheter insertion site monitored for inflammation or irritation.

      1. Materials needed

        • a long, flexible, sterile intravenous over-the-needle catheter with a large-gauge needle

        • local anesthetic (2% lidocaine)

        • suture material

        • sterile gloves

        • scalpel blade

        • bandage material

        • oxygen tubing

        • oxygen canister

        • distilled water.

      2. Technique

        • Position the patient in a comfortable, sternal position with the head and neck positioned upward (Fig. 20.3).


          Figure 20.3 Transtracheal oxygen catheter placement.

        • Shave and prepare the tracheal area using sterile technique.

        • Desensitize the overlying skin area with the local anesthetic agent.

        • Put on surgical gloves and remove the catheter from the sleeve; fenestrate the end with a scalpel blade to minimize tracheal trauma.

        • Insert the needle end of the catheter percutaneously into the trachea at the cricothyroid membrane or between tracheal rings just below the larynx.

        • Slide the catheter into the trachea until the tip of the catheter reaches to approximately the level of the carina.

        • Remove the stylet slowly from the catheter.

        • Pull the needle slowly from the trachea and secure it into the needle guard.

        • Connect the oxygen tubing to the catheter and insufflate oxygen and humidification setup as in nasal oxygen administration.

        • Suture the transtracheal catheter both at the insertion site and around the needle guard.

        • Bandage the neck with loose cotton gauze and veterinary tape, and label oxygen lines appropriately.

    3. Nasal-tracheal oxygen catheter placement

      Nasal-tracheal oxygen catheter may be used for laryngeal paresis/paralysis, collapsing trachea, or in brachycephalic patients not responding favorably to nasal oxygen.

      1. Materials needed

        • a soft, polyvinyl catheter (red rubber, 3.5–5F for small patients, 5–8 F for medium size dogs, 8–10F tubes for large dogs

        • petroleum jelly for lubrication

        • laryngoscope with long Miller blade

        • tissue or sponge forceps

        • suture material

        • oxygen tubing

        • oxygen canister

        • distilled water

        • note that patient may require mild sedation for catheter placement.

      2. Technique

        • Sedate the patient for visualization of the oral cavity.

        • Premeasure the tube to the fifth intercostal space and mark with a permanent marker; this length is slightly cranial to the tracheal bifurcation.

        • Repeat the technique for nasal oxygen placement, keeping the head hyperextended as the catheter reaches the level of the pharynx.

        • Open the mouth and visualize the catheter tip, using a laryngoscope with a long Miller blade.

        • Using tissue or sponge forceps, grasp the catheter tip and insert it into the trachea.

        • Slide the catheter to the predetermined length, and suture it in place.

        • Place an Elizabethan collar on the patient.

        • Deliver O2 at 50 mL/kg/min (always humidify).

        • Radiographs may be required to confirm location.

    4. Mechanical ventilation

      Mechanical ventilation is a challenging and labor-intensive therapy in any critical care setting. Indicated for patients who cannot effectively ventilate on their own, mechanical ventilation improves gas exchange, decreases the work of breathing, and assists in the recruitment of alveoli. Neurologic diseases for which mechanical ventilation may be beneficial include secondary pneumonia, severe neurogenic pulmonary edema, ARDS (acute respiratory distress syndrome), thromboembolic disease, head trauma, post-cardiopulmonary arrest, and LMN disease. Modes of mechanical ventilation therapy include controlled ventilation, assisted ventilation, and intermittent mandatory ventilation (IMV). Mechanical ventilators themselves operate as either pressure-limited, volume-limited, or time-limited systems. Depending on the ventilator employed, respiration is controlled by some combination of inspiratory pressure, tidal volume, inspiratory time, respiratory rate, and minute ventilation. Most patients require continuous or intermittent sedation. In some cases, sedative therapy is used in conjunction with paralytic agents. Complications of positive pressure ventilation (PPV) include oxygen toxicity, barotrauma, decreased cardiac output, and pneumonia. Protocols for ventilated patients should be well established and communication between all team members kept active.

         Because positive pressure ventilators achieve lung inflation by applying either continuous or intermittent positive pressure to the airway, airtight seals between the patient and the ventilator must be maintained either by a cuffed tracheal or endotracheal tube. Ensuring tight seals should always be the first priority when ventilating a patient. Oxygen and compressed air lines should be monitored every hour to ensure proper supply and delivery. Electrical cords and the power source should be anchored securely and not in the flow of traffic. The ventilated patient should be in a sternal position, level with the ventilator, and on comfortable bedding or pads to prevent sores or ulcerations. It is recommended that all ventilated patients have sterile tracheotomy or endotracheal tubes placed on setup, with the endotracheal tube clearly marked with a visible line at the upper canine tooth to prevent tube migration. Patients expected to need ventilatory support for longer periods (or brachycephalic breeds) may benefit from a temporary tracheostomy tube for airway management. Tracheostomy tubes also help to decrease anesthetic drug requirements and may facilitate successful weaning from the ventilator. Whether a tracheostomy tube or an endotracheal tube is used, the cuff needs to be inflated to protect the airway and to allow PEEP and PPV to be delivered.

         Also recommended at setup are arterial catheters for serial blood gas sampling and direct blood pressure monitoring, central catheters for central venous pressure monitoring and serial electrolyte analysis, and ECG telemetry for detecting cardiac arrhythmias. Closed urinary catheters are also helpful in calculating fluid output. Body temperature should be monitored frequently, and circulating water heating pads should be available for hypothermia.

         Modes of application are generally determined by patient need and clinician experience. Wherein the goals of PPV are to restore oxygenation and correct ventilation (carbon dioxide levels), barotrauma can result if pressure settings are too aggressive. In addition, patients with diseased or noncompliant lung parenchyma are often placed on higher-than-normal pressure settings in order to prevent small airway and alveolar collapse. Clinical signs of barotrauma can include rapid, shallow voluntary respirations, absence of lung sounds, abnormally high trends in the central venous pressure, and abnormal SpO2 and ETCO2 readings. Pneumothorax is a common complication, occurring more frequently in patients requiring high airway pressures (> 30 cm H20) or large tidal volumes. The presence of a pneumothorax should be considered in any patient with a sudden decline in oxygen saturation and/or tidal volume, or an elevation in ETCO2. Management of a pneumothorax includes immediate thoracocentesis; the placement of unilateral or bilateral chest tubes may be required if PPV is continued.

         Pulse oximetry should be used on every ventilated patient. If pulse oximeter readings are less than 92%, verify data by first ensuring that an adequate supply of oxygen is present and that the tracheal tube has not migrated or is occluded by a kink or mucous plug. Ensure tight connections of all airway tubing. Second, visibly inspect the patient’s mucous membrane color, pulse quality, and heart rate; auscultate lung fields for a possible pneumothorax. Ensure that the probe has not migrated, or change location of the probe. Until the problem can be identified, increase the oxygen flow (FiO2) to 100%. Verify low oxygen content by running an arterial blood gas, and notify the emergency room clinician of possible respiratory deterioration.

         Capnography should be used on the ventilated patient. The ETCO2 values generally should be kept between 30–45 mmHg; high CO2 values should initially prompt a search for tube occlusion, migration, or pneumothorax. Verify high settings with an arterial blood gas analysis; ventilator settings (breaths per minute) or endotracheal tube length may be altered to decrease dead space. Low ETCO2 values should be monitored for tube occlusion, leaking airway tubing, or a nonpatent cuff. Verification is also recommended by analyzing arterial blood gas, as ventilator settings may need to be altered. The ventilated patient should be monitored for leaking airway sounds, such as bubbling or fluid sounds on inspiration, around the cuff or oral area.

         Other considerations with patients on PPV include possible blood pressure abnormalities due to impaired intrathoracic blood flow. As PPV increases pleural pressure, venous return can become compromised to both sides of the heart, affecting diastolic filling. Mechanisms to control such impedance include manipulation of inspiratory time (the length of time pressure is applied) and manipulating respiratory rate (increasing the rate could prevent cardiovascular recovery time). Changes in central venous pressure trends, diastolic and systolic blood pressure trends, pulse quality, mucous membrane color, and heart rate may denote compromise of cardiac performance in the ventilated patient. Any abnormal trends following setup on the ventilator warrant changes in mechanical settings. Cardiovascular compromise may also be particularly severe in patients with intravascular volume depletion or pre-existing cardiovascular instability, or when particularly aggressive ventilator settings are required to maintain adequate arterial blood gas results.

         Note that it is always recommended to use minimal ventilator settings to prevent barotrauma and altered blood flow. Consequently, monitoring the arterial blood gases on setup and within the first few hours of PPV is critical. Minimal acceptable PaO2 values should be about 90 mmHg, with maximum PaCO2 values of 60 mmHg, although every patient should be evaluated on an individual basis.

         As most patients will not voluntarily be mechanically ventilated, “bucking” the ventilator (patient–ventilator asynchrony) can become problematic. Changing ventilator mode to IMV can improve patient–ventilator synchrony and sedation protocols can be modified to control patient–ventilator asynchrony. Opioids should be used with caution, as abnormal breathing patterns (panting) may surface as a result of their use. Injectable anesthetic agents, such as propofol, should be kept readily available in the event of tube migration or tube lumen occlusion, requiring re-intubation.

         Preventing tube occlusion is critical. If an endotracheal tube is used, sterile suctioning after sterile saline administration is recommended every 4 hrs. Note that there are complications of tracheal suctioning, such as hypoxemia, traumatic airway ulceration, cardiac dysrhythmias secondary to hypoxia and pain, and infection from improper technique. High FiO2 levels (100%) are recommended both immediately before and after suctioning, with strict sterile technique applied. Suctioning should be brief and thorough, with the suction catheter fed to the bifurcation only.

         If a tracheotomy tube is used on the ventilated patient, ensure that the tracheotomy tube used has both a cuff and an inner and outer cannula, if at all possible. The inner cannula can be removed for more thorough cleansing, and can be suctioned with the aforementioned technique. If a tracheotomy tube is used without an inner cannula, cleaning must again be accomplished by cautious aseptic technique. Endotracheal tubes should be removed and replaced with a sterile tube every 24 hrs. In addition, nebulizing through the tracheal tube is recommended to maintain hydration of the pulmonary parenchyma, particularly if pneumonia is present.

         Patient hydration should be assessed by monitoring urinary output, daily weighing, electrolyte analysis, and central venous pressure monitoring. As multiple organ dysfunction can occur in any critical patient, renal values should be monitored with consistency. PPV and the use of narcotic sedatives can also increase the secretion of antidiuretic hormone, thereby decreasing urine output. As a result, fluid retention and subsequent edema formation may occur. Intermittent diuretic therapy may be required during the period of ventilation. Nutrition should also be addressed either by nasogastric or gastrostomy tube, depending on the length of proposed ventilation. Note that if nutrition is delivered to the ventilated patient gastrointestinal (GI) motility may be impaired; consequently, aspirating stomach tubes is warranted to monitor residual volume. Facilitating colonic emptying may be required by either an enema or rectal palpation.

         It is recommended that patients on PPV be kept in sternal recumbency. Passive range of motion, flexion/extension, and massage/effleurage should be performed every 4–6 hrs. In addition, gentle coupage is recommended to prevent lung parenchyma atelectasis and to help mobilize secretions to central areas reached by routine suctioning. Oral care should also be addressed in order to prevent nosocomial pneumonia. The oral cavity should be flushed with dilute chlorhexidine and suctioned every 12 hrs. The tongue can be kept moist by wrapping it with moist gauze and kept inside the oral cavity. Pressure sores on the tongue can be reduced by the use of mouth bridges or gags. The eyes should also be kept moist by saline flushes and eye lube every 4–6 hrs. Fluorescein staining should be performed regularly to check for ulceration and treatment instituted if necessary.

         All patients on mechanical ventilation are at a high risk of acquiring nosocomial infections. Minimizing risk by being conscientious, practicing sterility with handling of catheters and tubes, and cleaning circuitry between patients are highly recommended. Development of fever, or inflammatory changes on complete blood count, may be indicative of sepsis. Thoracic radiographs along with culture and susceptibility testing of airway samples from a bronchoalveolar lavage are recommended. In order to limit the development of bacterial resistance, antimicrobial therapy should not be routinely started on mechanically ventilated patients, unless an underlying infectious process was previously identified.

         Established protocols for patient setup, maintenance, and recovery are paramount for a successful outcome. Communication between the nursing staff and clinicians should be concise, and team members need to be ready for any emergency situation, such as re-intubation, hand ventilation (from power failure or machine malfunction), or tube occlusion.

         Weaning a patient from mechanical ventilation can be difficult. For weaning to be successful, the patient must have a sufficient respiratory drive, as well as adequate neuromuscular function to achieve a sufficient tidal volume. The weaning process typically involves a gradual reduction in mechanical ventilation with a proportional increase in the work performed by the patient. The patient should be closely monitored at every stage of the process for any sign of respiratory insufficiency (hypoxemia, hypercapnia, hypertension, or patient distress). In addition, oxygen supplementation (e.g. nasal oxygen) should be provided in every patient coming off mechanical ventilation. Successful weaning off the ventilator is largely dependent on the primary disease process leading to the initiation of ventilation.

    5. Placement of arterial catheters

      Arterial catheters are placed in order to check serial blood gas during respiratory difficulty, hypercapnia, hypoxia, and mechanical ventilation. Catheters are also used for direct blood pressure monitoring.

         Arterial catheters should not be used in thrombolytic disease or coagulopathic disease patients.

      1. Materials needed

        • clippers

        • surgical scrub

        • suture material

        • heparinized saline flush

        • catheter cap

        • arterial catheter kit or cephalic catheter

        • bandage material

        • saline for infusion if continuous blood pressure is monitored.

      2. Technique

        • Lay the patient in lateral recumbency.

        • Give the patient oxygen if the body position compromises respiratory effort.

        • Clip and prep dorsal pedal area (down limb).

        • Palpate the pulse with a single digit.

        • Use a stab incision for a “cut down” at a 45° angle and at least 1 inch away from a palpable pulse.

        • Hold the catheter like a dart; insert the catheter with the bevel up through a tunneled area.

        • Use “baby-steps” toward the pulse (superficial); back out and redirect if there is no flash.

        • Once a flash is obtained, be careful not to move the catheter; slide the entire length of wire via the black tab handle.

        • “Pop” the catheter off the wire and into the arterial space.

        • Remove the wire and ensure blood flow (should be fast and in a spurting or jetting motion).

        • Cap and flush the catheter; aspirate back to ensure blood flow and re-flush.

        • Suture the catheter in place.

        • Place a small amount of antibiotic cream over the insertion site.

        • Cover with a sterile gauze square.

        • Place one layer of gauze bandage.

        • Cover the catheter with elasticon and label “ART.”

           Change the catheter cap, bandage, and sterile gauze daily. Flush the catheter every 2 hrs with heparinized saline, if you are not using it as a continuous rate infusion (CRI) for direct blood pressure monitoring.

        Important note: Do not give any medications through the arterial catheter.

    6. Pulse oximetry and capnography

      1. Pulse oximetry

        Patient monitoring using pulse oximetry should be utilized for every patient undergoing anesthesia or as a monitoring tool in the ICU. A noninvasive method of continually measuring hemoglobin oxygen saturation (SpO2), the pulse oximeter will display an oxygen waveform or an oxygen saturation percentage useful in evaluating lung function. Pulse oximetry does not assess ventilation, and should not take the place of lung auscultation or arterial blood gas analysis. Pulse oximeters commonly use a tongue, earlobe, or toe digit as the cuvette within which SpO2 is measured. Pulse oximetry must be used in conjunction with other hemodynamic markers and not used solely to determine respiratory stability. Invalid pulse oximetry readings are common occurrences on the poorly perfused, hypothermic, jaundiced, or patients with cardiovascular compromise. Low pulse oximeter values (less than 94%) in conjunction with other abnormal exam findings should result in oxygen therapy (or increased FiO2 if already receiving oxygen) and an arterial blood gas analysis.

      2. Capnography

        Capnography is a noninvasive method for the continuous assessment of ventilation by the measurement of ETCO2. Capnography is superior over pulse oximetry for the prompt identification of apnea and airway mishaps, as there are instantaneous changes in ETCO2 as opposed to slower changes in the percentage of hemoglobin saturated with oxygen (SpO2). Clinical indications for capnography include ensuring correct endotracheal tube placement, detection of apnea, and in monitoring adequacy of ventilation and pulmonary perfusion during cardiopulmonary resuscitation. ETCO2 monitoring can be critical in detecting potentially catastrophic anesthetic complications during common neurodiagnostics such as the collection of CSF fluid during neck flexion, when the endotracheal tube may be inadvertently kinked (Fig. 20.4).


        Figure 20.4 Respiratory changes are closely monitored during cervical positioning for CSF collection.

           An instrument called the capnograph displays both numerical and waveform imaging. During anesthesia, respiratory depression secondary to drugs and inhalants can be effectively monitored using a capnograph. There are two types of monitors available for assessing end-tidal carbon dioxide: the capnometer or capnograph. Capnometers provide only minimum and maximum ETCO2 values, while capnographs display graphic representation (waveforms) of exhaled carbon dioxide.

           Capnometers and capnographs may be categorized as mainstream or sidestream, based on the location of the sensing device. Mainstream (nondiverting) monitors analyze the respiratory gases locally (at the endotracheal tube-breathing system interface), while sidestream (diverting) monitors employ sensing tees placed at the endotracheal tube-breathing system interface and pump respiratory gases for analysis up into the measurement chamber via a length of tubing. Although mainstream monitors provide rapid results and have less mechanical problems caused by condensation, sidestream monitors can be utilized for remote patient monitoring such as during an MRI. In addition, sidestream monitors can detect expired gases in nonintubated patients through nasal tubes, a critical monitoring tool for patients susceptible to hypercapnia (e.g. cervical lesions or LMN conditions).

           Normal ETCO2 values are approximately 35–45 mmHg; abnormal ETCO2 trends should be verified by an arterial blood gas sample. Persistently increased ETCO2 values indicate inadequate ventilation, necessitating ventilatory assistance. Prolonged periods of hypercapnia can lead to myocardial depression and respiratory acidosis. Elevated ETCO2 levels may also occur as a result of airway obstruction, pneumothorax, body positioning, or lung parenchymal disease. Hyperventilation (purposefully decreasing ETCO2) during anesthesia for diagnostic or therapeutic procedures can be beneficial to decrease ICP through vasoconstriction of cerebral vasculature.

Recumbency and pressure sores2, 4, 15, 34, 45

Other complications of the recumbent animal include development of pressure sores or decubital ulcers. These are local areas of skin necrosis. Pressure sores are often localized to bony prominences; localized pressure over these areas leads to tissue ischemia of variable severity. The extent of tissue damage is often graded from least severe (grade I: darkened area of thickened skin, no exposure of subcutaneous tissue) to most severe (grade IV: deep tissue loss with exposure of bone). Grade II decubital ulcers involve exposure of subcutaneous fat, and grade III ulcers involve tissue defects to the level of deep fascial layers. Ulcerations on extremities and the pads of the feet can also occur with wheelchair use. Protective boots can prevent damage to paws and help dogs walk on slippery floors. Velcro pads can be applied either onto the bars of the wheelchair or to affected areas to help prevent ulcerations that may form from constant rubbing.

   Frequent turning of the patient and appropriate bedding represent the most important preventative measures of a nursing-care protocol. Increased skin moisture and irritation contribute to the development of decubital ulcers; therefore, patients should be kept clean and dry and should be bathed frequently. Since decubital ulcers are primarily caused by pressure, they can be avoided or minimized by using bedding such as sheepskin, foam or air mattresses, trampolines, or bandaging techniques. Sheepskin is advantageous in that it is inexpensive and can be laundered for multiple uses. The sheepskin minimizes friction and can absorb moisture, which is particularly important in preventing urine scalding. Sheepskin may make patients hot, so rectal temperatures should be monitored frequently when this bedding material is used. Air mattresses are also inexpensive and allow pressure distribution to avoid decubital ulceration. Disadvantages associated with air mattresses include puncture holes from the patient’s nails and the inability to launder air mattresses for long-term use. Urine scalding can also occur with the use of air mattresses. Trampoline beds are an excellent choice for the recumbent patient. The trampolines are constructed from plastic piping and fiberglass netting, allowing air to circulate underneath. The trampoline also distributes a patient’s weight evenly, helping to prevent pressure sores. Urine scalding is also avoided by the fiberglass netting, as the urine falls underneath the patient onto plastic trays. Bandaging techniques in the form of doughnuts (Fig. 20.5) can also be placed over bony prominences to prevent decubital ulcers or over existing decubital ulcers to prevent further pressure damage. Such devices can effectively relieve pressure while allowing for the adequate aeration of tissues.


Figure 20.5 Preparing a “doughnut” to place over a decubital ulcer in a dog.

   Nonsurgical vertebral fractures and/or spinal luxation involve external splints and strict cage rest for 6–8 wks to allow healing (see Chapter 15); sternal positioning is recommended when at all possible. As splints are held in place using bandage material, bandages should be evaluated daily to ensure there is no respiratory compromise caused by excessive constriction. Bandages should be evaluated daily for evidence of soiling, and limbs examined for decubital ulcer formation or abrasions caused by external splints.

   Treatment of decubital ulcers may involve medical and/or surgical therapies. Specific medical therapy depends upon the individual case, but may involve frequent wound lavage, systemic antibiotics, wet-to-dry bandaging, and application of topical drugs. These topical agents include antibacterial preparations (e.g. triple antibiotic, gentamicin, nitrofurazone, silver sulfadiazine ointments), enzymatic debriding agents, and hydrophilic agents (maltodextrin powder wound dressing). Additionally, Preparation H is believed to stimulate wound healing when applied to decubital ulcers. Surgical intervention and wound culture/susceptibility testing may be required for decubital ulcers, especially if they are grade III or IV in severity. Such intervention may include debridement and primary closure, delayed wound closure, or use of cutaneous or myocutaneous flaps.

Bladder management2–4, 18, 19, 25, 34, 36

Urinary complications are common in dogs and cats with neurologic dysfunction. Overdistension of the urinary bladder and urinary tract infections are typical sequelae, both of which are avoidable with attentive nursing care. Proper technique in both expressing and catheterizing the bladder is important to prevent urethral and bladder wall trauma, to prevent introduction of bacteria into the urinary tract, and to measure urinary output in the oliguric or anuric patient as a guideline for appropriate fluid therapy. Poor nutrition and decreased water intake can also affect the patient’s urinary system and should be corrected. Overdistension of the bladder can result in permanent atony of the detrusor muscle. The bladder should be palpated to estimate size, even if there is urine present in the cage. The presence of urine in the patient’s cage is not a reliable indicator of the ability to urinate voluntarily; the patient could have urinary overflow as a result of distension. The bladder should be expressed every 4–6 hrs as a general rule, but the urodynamics of each patient should be assessed on an individual basis (e.g. prednisone use or intravenous fluids could warrant bladder evacuation more frequently). If the bladder cannot be expressed without minimal stress to the patient, a urinary catheter should be placed. Whether placing a closed urinary collection system or intermittently catheterizing the urinary bladder, proper sterile technique must be followed in order to avoid nosocomial urinary tract infections. In the postoperative patient, detrusor dysfunction may occur following systemic (CRI) or epidural use of opiates; therefore, bladder size should be evaluated every 4–6 hrs and expressed or catheterized as necessary.

  1. General guidelines for urinary bladder expression

    It is important to distinguish between upper motor neuron (UMN) bladder and LMN bladder dysfunction to best determine which pharmacologic agents will be most efficacious in improving bladder function (see Chapter 16). Before expressing the bladder, it is advisable to first allow the patient to try to urinate voluntarily by walking or carting the animal outside. If the patient does voluntarily urinate, it is still necessary to palpate the bladder after urination to ensure complete evacuation. Catheterization may be necessary in order to determine the amount of residual urine left after urinating if the bladder still palpates as large. Normal residual urine volume in the dog bladder is between 0.2 and 0.4 mL/kg body weight; this is believed to be similar for cats. Normal urine output for the dog and cat is approximately 1–2 mL/kg/hr. Note that the foremost objective in bladder expression is to avoid overdistension and detrusor muscle atony. Stress should be minimized during bladder expression by using proper technique. Diazepam given intravenously or orally can help relax the external sphincter tone, as well as reduce anxiety in dogs requiring frequent bladder expression.

    1. Technique (Fig. 20.6)


      Figure 20.6 Sequential steps (A–C) for bladder expression in the recumbent patient.

      • Place the patient in a comfortable position, either standing or in lateral recumbency.

      • If the patient is in lateral recumbency, gently place one hand on the upper abdominal wall and one hand underneath the position in a symmetrical fashion. If the patient is standing, place one hand on each side of the abdomen. In either case, the hands should be initially placed just caudal to the last rib.

      • Gently palpate the abdomen, slowly advancing the hands medially toward each other. If the urinary bladder is not palpable, simultaneously move both hands caudally until the bladder is contacted. Since the bladder is somewhat mobile within the abdomen, and bladder size is variable, starting cranially and manipulating the bladder caudally will force the bladder into the pelvic inlet region, preventing it from escaping the clinician’s grasp.

      • Once the bladder wall is palpable, steady, even pressure is applied with both hands. The direction of the pressure should be medial and caudal. Gently express the bladder until it feels empty. The clinician should be able to feel the bladder “deflate” as urine is expressed.

      It is important to note that with frequent bladder expressions the abdomen may become tense as a result of patient anxiety, consequently making the bladder difficult to express. In addition, as the abdomen tenses, the bladder may become displaced in an otherwise abnormal position (cranially). Evacuation of the bladder should never be forceful or aggressive. Catheterization may become necessary until pharmacologic agents are of benefit.

  2. General guidelines for urinary bladder catheterization

    Urinary catheterization can introduce microbes into both the bladder and the kidneys, traumatize the urethra, and cause patient discomfort. Catheterization of the neurologic patient should be performed only if attempts at bladder expression are unsuccessful or contraindicated (i.e. entrapped bladder, suspected bladder trauma from automobile trauma), or for recumbent, critical patients with spinal injuries. The type and length of the urinary catheter is important to facilitate efficient bladder evacuation and to measure urinary output. If a closed system for long-term use is desired, a softer, less-irritating catheter with a ballooning device for anchorage should be used, particularly in the female dog. If less frequent catheterization is warranted, bladder decompression can be obtained by intermittent catheterization two to three times daily using a soft, polyvinyl or red rubber catheter or feeding tube. It is of the utmost importance to ensure adequate length of the urinary catheter. The catheter should always reach into the neck of the bladder for either system. The catheter should have a visible marker in order to determine whether it is backing out during patient use. If a Foley catheter is used, adequate dilation of the balloon should be maintained with at least half the amount of air or fluid suggested by the manufacturer (the amount should be indicated on the sleeve of the balloon).

    1. Materials needed

      • appropriate length and gauge of urinary catheter and stylet

      • sterile gloves

      • sterile lube

      • mild cleansing agent (e.g. dilute chlorhexidine)

      • suture material and medical tape (for indwelling catheters)

      • sterile closed collection system (for indwelling catheters)

      • clippers for sterile preparation (female)

      • sterile syringes for urine sample containment

      • sterile fluid-filled syringes (saline preferred).

    2. Technique

      When positioning the patient for placement of a urinary catheter, it should be kept in mind that, while restraint is necessary, the comfort of the patient is of the utmost importance. The patient should be placed in a position that will facilitate both sterile technique and successful catheterization. In the male dog, lateral recumbency is preferred, with the prepuce retracted and the penis aligned parallel with the long axis of the body (Fig. 20.7). A stylet in the catheter for the male patient is usually not required, unless urethral stones are present or suspected. When catheterizing male cats, the penis must be straightened before passing the catheter. This is accomplished by applying caudal traction to the preputial region, directing the penis in a caudal direction, parallel with the long axis of the body. Female patients can be placed either in sternal (usually preferable) or lateral recumbency (whichever is more comfortable for the patient yet optimal for the visualization of the urethral papilla). Due to the curvature and size of the papilla, a stylet in the urinary catheter is often useful when catheterizing the female patient. Mild sedation should be considered for the comfort of the patient and to facilitate catheterization via relaxation of the urethral musculature. Urethral catheterization may be facilitated in some cases by using a syringe attachment and pulsating fluid as the catheter is being advanced.


      Figure 20.7 Placement of a urinary catheter in the male dog. (Courtesy of Dr. Catherine Langston, The Ohio State University.)

      • Position the patient for catheter placement (lateral for males, sternal for females).

      • Prepare the catheter insertion site with antiseptic solution. Shave the perivulvar area in females prior to skin preparation.

      • Wearing sterile gloves, inspect the balloon on the Foley catheter, lubricate the catheter, and measure the estimated length of catheter to be passed by marking the distal end of the catheter with a permanent marker. Insert the stylet into the catheter (females).

      • Pass the catheter while an assistant retracts the prepuce or vaginal folds.

      • For female patients, visualization of papillae may be best accomplished with a laryngoscope light and/or a vaginal speculum (Fig. 20.8).


        Figure 20.8 Visualization of the urethral papilla in the female dog to facilitate urinary catheter placement. (Courtesy of Dr. Stephen DiBartola, The Ohio State University.)

      • Pass the catheter to the desired measured length.

      For closed indwelling systems:

      • Inflate the balloon of the Foley catheter with the recommended amount of saline (written on the side of the balloon arm).

      • Pull the Foley catheter out of the urethra until the balloon catches on the bladder neck (females only).

      • Wipe the catheter dry, and fasten it with tape and suture it in place (Fig. 20.9).


        Figure 20.9 Anchoring the urinary catheter. (Courtesy of Dr. Catherine Langston, The Ohio State University.)

      • Obtain a urine sample and attach closed system.

      It is recommended to place stay sutures both around the catheter and prepuce/vulva region, with the remainder of the urinary catheter fastened around either the tail or abdominal area to prevent dislodgement. The outer portion of the urinary catheter should be labeled with permanent marker in order to monitor the optimal insertion length for the duration of use. The patient should be observed closely for licking, chewing, or biting at the urinary collection system. An Elizabethan collar should be placed if a patient displays such behavior, to prevent premature catheter removal. Excessive force is contraindicated in the passage of any urinary catheter. Urethral trauma, including tears, can result from aggressive catheterization attempts; such trauma can lead to life-threatening consequences. The type of catheter used is paramount; acceptable indwelling catheters include those made of polyvinyl chloride or silicone Foley catheters. Polypropylene catheters should not be used for indwelling catheters, as they are stiff and uncomfortable to the patient and may cause uroepithelial damage. The length of the catheter is also important. Inserting excessive catheter length may result in the catheter looping around itself and kinking, cutting off the flow of urine. Excessive catheter length may also traumatize the bladder wall. Placing the catheter in the urethra and not into the bladder can also hinder bladder evacuation or drainage. The tip of the catheter should ideally be inserted to reach the caudal aspect of the bladder lumen, in the trigone area. Radiographs may be indicated to ensure proper placement if urine production is questionable.

  3. Indwelling versus intermittent urinary bladder catheterization and catheter care

    Indwelling catheters are advantageous for patients who run the risk of repeat urethral blockage (e.g. feline urolithiasis). Repeated catheterization of such patients may lead to further urethral trauma and tissue inflammation, patient discomfort, and introduction of microbes. Indwelling catheters are also beneficial for patients receiving large volumes of crystalloid fluids as a tool to properly gauge renal function (e.g. “ins and outs” fluid therapy). Indwelling systems are also useful for recumbent patients who cannot be moved outdoors to urinate (e.g. spinal fracture patients) and patients with atonic bladders or historical/clinical evidence of bladder trauma. The importance of aseptic technique cannot be overemphasized; antibiotic therapy to avoid a urinary tract infection is not recommended, as this practice may facilitate antibiotic-resistant urinary tract infections. It is important to maintain a sterile collection system if an indwelling urinary catheter is used. Caution should be exercised when moving either the urine collection bag or the patient to avoid urinary flow from the bag back into the patient. Specialized, anti-reflux urinary collection bags can be utilized to prevent urinary back-flow. If an intravenous drip set is used for the collection device, the roll clamp should be removed to avoid the mistake of restricting urine flow. The urine collection bag should be positioned at a level below the patient to ensure proper urine flow. If the patient is experiencing hematuria or urolithiasis, intermittently flushing the bladder with sterile saline is recommended. The urine collection bag should be emptied every 4 hrs and urine production carefully recorded. If urine production appears inadequate, accurate placement and patency of the urinary catheter should be verified before increasing fluid administration.

       In summary, to minimize urinary tract infections from an indwelling catheter, sterile technique should be observed. However, the best way to avoid urinary tract infection is to remove the catheter as soon as it is feasible to do so. Approximately one-half of dogs catheterized for 4 days or longer will develop a urinary tract infection. If a closed urinary system is used, the prepuce or vaginal area around the urinary catheter should be swabbed every 12 hrs with dilute chlorhexidine to minimize bacterial growth. Urinary collection bags should be changed every other day.

       Current recommendations for urine culture and susceptibility testing does not include culturing the urinary catheter tip. The urinary catheter should be removed and a urine sample collected via cystocentesis with sterile technique 1–2 hrs after catheter removal. Administration of antibiotics as prophylaxis in patients with an indwelling urinary catheter is not recommended.

       Intermittent urinary catheterization must also follow strict aseptic technique. Intermittent catheterization may be used to obtain a urine sample if a diagnosis is dependent on urinalysis and cystocentesis is contraindicated or unsuccessful. Intermittent catheterization may also be used for patients experiencing contractility difficulty; manual expression may be difficult in such patients and may cause discomfort. It should be kept in mind that frequent intermittent catheterization may lead to patient discomfort, urethral trauma, and urinary tract infection. The practice of intermittent catheterization should only be used for the patient who does not need long-term bladder care. Finally, it is very important to keep the patient clean and housed with dry bedding in order to prevent urine scalding if the patient is recumbent and does not have a closed urinary system. As discussed previously, trampolines are available to prevent the patient from lying in urine if 24-hr care is not provided.

       Unlike urination, defecation usually proceeds without assistance in animals with neurologic disease. However, constipation may be caused by opioid use, changes in diet, or stress due to hospitalization. Diarrhea may also occur due to medications such as corticosteroids. In addition, dogs with LMN disease may develop urinary and fecal incontinence. Regular expression of feces is recommended to help keep the patient clean. Low-residue diets may be beneficial in patients with fecal incontinence; however, prognosis for functional recovery is often poor. Stools should be evaluated for blood, as glucocorticoids are frequently prescribed for many neurologic disorders. Clinical signs may be diarrhea, black or tarry stools, or pain with defecation. Treatment often includes the administration of gastric protectants.

Rehabilitation therapy2, 4–7, 11–15, 17, 18, 20, 21, 23, 26, 28, 29, 31–35, 37, 39–44, 46, 47

Although neurologic rehabilitation is commonplace in humans following neurosurgery or after neurologic emergencies such as strokes, veterinary neurologic rehabilitation is not as widely accepted or applied. The major goals of rehabilitation therapy are to attain or maintain full range of joint motion, minimize muscle atrophy, and prevent or ameliorate patient discomfort. Traditional therapies of hot packing, cold packing, massage, and simple stretching exercises in veterinary medicine have been supplemented by more advanced treatments, such as hydrotherapy, laser therapy, sling therapy, ultrasound, electrical stimulation, sling-supported exercise, and acupuncture. There is an increasing demand for prolonged postoperative care in dogs and cats, reflective of advances in veterinary neurosurgery. Emphasis on such physical therapeutics can result in shorter hospitalization periods and improved patient wellbeing. A plan for rehabilitation therapy should be discussed between veterinarian, technician, and owner, in order to provide the best rehabilitation program. Benefits of rehabilitation therapy include improved circulation, increased production of collagen, decreased inflammation, decreased muscle atrophy, and prevention of joint stiffness. Pain management should always be an important consideration for the patient undergoing rehabilitation therapy. Assessment by the veterinary team and owner includes watching out for pain behaviors, posture, muscle imbalances, and gait impairments before, during, and after rehabilitation therapy.

   It is extremely important to have protocols of rehabilitation therapy discussed between veterinarian and technician, as patients with identical diagnoses may require different treatments. For example, animals with intervertebral disc disease (IVDD) may have varying degrees of neurologic impairment and will require differing degrees of rehabilitation therapy. In addition, patients recovering from vertebral fractures may receive varying stabilization techniques and will consequently receive a rehabilitation therapy regime dependent upon the surgical technique. Soft tissue trauma, such as is often encountered in automobile accidents, may be complicated by delayed wound healing if rehabilitation therapy is instituted prematurely.

   Rehabilitation therapy emphasizes a return to function rather than treatment of a specific disease or diagnosis.

   Managing pain in these patients can be a challenge to any clinician. Anti-inflammatory drugs, analgesics, herbal medications, and alternative modalities such as acupuncture may all be needed to treat painful conditions. Analgesic drugs and alternative therapies are discussed in Chapters 21 and 22, respectively. Therapeutic modalities are used to augment medications and reduce pharmaceutical dosages that are required by the patient. The most common modalities used for pain relief include heat, cryotherapy, as well as manual therapies such as massage, joint mobilizations, and stretching, laser therapy, transcutaneous electrical nerve stimulation (TENS), and therapeutic ultrasound (TUS). Muscle strengthening and re-education is accomplished through the use of neuromuscular electrical stimulation (NMES), therapeutic exercises, hydrotherapy, and proprioceptive neuromuscular facilitation (PNF). Assistive devices, such as harnesses, slings, boots, carts and incontinence aids are often essential to the wellbeing and recovery of the patient.

   Therapeutic cold packing, or cryotherapy, is an important nursing technique for the acutely injured or postoperative patient. Cold packing is efficacious in producing local vasoconstriction and in preventing interstitial bleeding and should be the first form of therapy instituted. Owners can be instructed to apply a cold pack to the injured animal en route to the hospital. Benefits of cold therapy include reduction of enzymatic tissue activity (thereby reducing tissue destruction) and a reduction of pain perception (Box 20.1). Typically, cold packing is performed several times a day for 20 min, up to 3 days after injury. Ice packs can also be applied directly over an incision immediately after surgery (e.g. hemilaminectomy) to minimize local swelling and reduce the modulators of pain.

Apr 7, 2020 | Posted by in SMALL ANIMAL | Comments Off on Nursing Care and Rehabilitation Therapy for Patients with Neurologic Disease

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