C
Cardiopulmonary Cerebral Resuscitation
EQUIPMENT
• Fluids: crystalloids (e.g., lactated Ringer’s solution, 0.9% NaCl) and colloids (e.g., Hetastarch)
• Point of care testing for blood glucose, electrolytes, packed cell volume, total solids, lactate, blood gases
• Other items that should be nearby: lidocaine jelly, suction tubing and means of generating suction, IV and urethral catheters, thoracotomy tubes, laceration packs
• Items for doing open thoracic cardiac massage and vascular cutdown procedures, self-retaining retractors
The data collection sheet for record keeping should include:
• Animal name and medical record number, body weight, signalment and underlying disease, as well as assumed cause of the arrest
• A table or another quick and clear way to list which drugs are given, at what time they are given and at what dosage, and their effects
PREPARATION: IMPORTANT CHECKPOINTS
• The owners of any critically ill animal should be consulted at the time of the animal’s admission regarding their resuscitation wishes for the pet. In many cases of advanced or aggressive disease, advanced age, and financial limitations of the family, CPCR may not be appropriate, and the pet should be allowed to pass away peacefully.
• Successful CPCR is a team effort. The team consists of the clinician leading the arrest and individuals who are keeping a record of the event, acquiring the requested drugs or materials, administering the drugs and therapies, providing the chest compressions, and communicating with the animal’s family. The team may contain any number of people, from few to many, with some having multiple roles.
• A senior knowledgeable individual must take charge in fostering a calm environment during CPCR, directing the other team members in their tasks, and keeping the team informed of progress. One responsible team member should be in communication with the owners to keep them apprised of progress and inform the team of changes in the family’s wishes.
• All of the equipment that was previously listed (e.g., endotracheal tubes, breathing circuits, IV catheters, defibrillation paddles, sterile gloves, etc.) should be available in sizes appropriate for a wide variety of animals.
• Drug boxes should be checked weekly at least to ensure all drugs are available and have not reached their expiration dates.
• Equipment should be checked regularly for malfunctions and cleaned immediately after each use so that each piece is ready to be used again without notice.
• Data sheets should be kept nearby so that events, drug administration, animal information, and responses to treatments can be recorded in a timely and accurate fashion.
• After every CPCR event, the team should review the process to assess whether changes in plan, arrangement, or protocols are necessary for improvement of efforts in the future.
Calcium gluconate*: (10% = 100 mg/mL [calcium: 27.2 mg/mL or 1.36 mEq/mL])
Dextrose*: 5% (50 mg/mL) or 50% (500 mg/mL)
Dobutamine*: 12.5 mg/mL
Epinephrine*1: 1 : 1000 (1 mg/mL) or 1 : 10,000 (0.1 mg/mL)
Furosemide*: 5% (50 mg/mL)
Magnesium sulphate*: 10% (100 mg/mL, 0.8 mEq/mL), 12.5% (125 mg/mL, 1 mEq/mL), 50% (500 mg/mL, 4 mEq/mL)
Mannitol*: 5% (50 mg/mL, 275 mOsm/L), 10% (100 mg/mL, 550 mOsm/L)
Norepinephrine*1: 1 mg/mL
Procainamide*: 100 mg/mL
Sodium bicarbonate*: 8.4% (1 mEq/mL)
Prednisolone sodium succinate: (10 or 50 mg/mL)
Methylprednisolone sodium succinate: (62.5 mg/mL)
Flumazenil: 0.1 mg/mL Naloxone: 0.4 or 1 mg/mL
* Indicates drugs that should be in all basic emergency kits. The others may be beneficial, and their use is dependent on clinician knowledge and familiarity.
1 Indicates that the drug can be administered intratracheally using a long catheter, generally at double the dose and followed by several deep breaths.
POSSIBLE COMPLICATIONS AND COMMON ERRORS TO AVOID
• Incorrect assessment of ECG, especially false interpretation of bad contact between ECG machine and animal as “ventricular fibrillation”
• Delayed defibrillation. The success of defibrillation in an experimental setting approaches 100% when instituted within 1 minute of onset of fibrillation and falls precipitously to virtually 0% after 10-15 minutes.
• Acute renal failure, diarrhea, liver dysfunction, and altered mentation and reflexes are common sequelae to inadequate oxygen delivery and tissue perfusion. Organ function must be closely monitored to take preemptive action should parameters deteriorate.
PROCEDURE
Most of the initial tasks listed here should be occurring simultaneously:
• Initiate chest compressions at 80-100 compressions/minute, compressing approximately 30% of chest diameter, with a 1:1 ratio of compression to relaxation. Hands should be placed over the widest part of the chest for patients > 10 kg. Hands should be placed over the heart for patients < 10 kg. Interruption of chest compressions should be minimized to <10 seconds until ROSC occurs.
• Begin assisted ventilation with 100% inspired oxygen at 10-12 breaths/minute, higher if pulmonary disease preventing effective oxygen transfer exists, or the patient was already hypoxemic.
• If anesthetized, turnoff inhalant anesthetics, and reverse any injectable drugs that have reversal agents.
• If thoracic compromise exists (e.g., pleural effusion, pneumothorax) or arrest is due to pericardial effusion, open-chest CPR must be instituted immediately. With pericardial effusion, open the pericardium ventral to the phrenic nerve to improve diastolic function of the heart and remove effusion.
• Fluid administration: IV (or intraosseous [IO], especially in pediatric animals). Shock rate administration (90 mL/kg in dogs, 45 mL/kg in cats, with constant monitoring and adjustments as needed) is appropriate if the patient was hypovolemic prior to the CPA. Otherwise, consider an initial bolus of 20 mL/kg in dogs, 10 mL/kg in cats if the patient was euvolemic; use caution with cardiac or renal disease. Hypertonic saline can also be considered (4-6 mL/kg of 3% saline IV slowly over 5 minutes).
• Administer epinephrine (low dose: 0.02 mg/kg IV; high dose: 0.2 mg/kg IV). Consider starting with the low dose, repeat with high dose if initial dose is unsuccessful.
• Defibrillate electrically (e.g., starting with 2-5 J/kg if closed chest, 0.2-0.5 J/kg if open chest) if ventricular defibrillation is present. Repeat defibrillation if ventricular fibrillation persists.
• Additional medications to consider to treat arrhythmias: atropine (0.04 mg/kg IV) followed by epinephrine again if asystole unresponsive to initial epinephrine; amiodarone (5 mg/kg IV or IO over 10 minutes) for refractory ventricular fibrillation, atrial fibrillation, supraventricular tachycardia (SVT), ventricular tachycardia (VT); lidocaine (2-3 mg/kg IV in dogs) for ventricular tachycardia or flutter; atropine for persistent bradycardia; magnesium (0.15-0.3 mEq/kg IV slowly over 10 minutes) for refractory ventricular arrhythmias.
• Start vasopressors (e.g., dopamine 2-10 mcg/kg/min IV) if BP not adequate once appropriate fluid resuscitation is administered.
• Assess metabolic condition with repeated blood gas analysis and electrolyte and blood glucose monitoring: treat abnormalities accordingly.
POSTPROCEDURE
Clinicians should also consider the following during the monitoring period:
• Administration of mannitol at 1-2 g/kg IV over 30 minutes is useful to reduce cerebral edema secondary to hypoxic brain injury, as well as to support renal perfusion and improve circulation to tissues.
• Until the animal is conscious and mobile, recumbent animal care protocols must be used. Frequently turning the animal, lubricating its eyes, keeping it clean of eliminations, and keeping the oropharynx clean and/or suctioning the endotracheal tube are imperative. Monitor for pressure-induced sores and blood flow impairment.
ECC Committee, Subcommittees and Task Force of the American Heart Association. 2005 American Heart Association guidelines for cardiopulmonary resuscitation and emergency cardiovascular care. Circulation. 2005;112:IV1-IV203.
Hackett TB. Cardiopulmonary cerebral resuscitation. Vet Clin North Am Small Anim Pract. 2001;31:1253-1264. 6
Plunkett SJ, McMichael M. Cardiopulmonary resuscitation in small animal medicine: an update. J Vet Intern Med. 2008;22:9-25.
Catheterization, Urethral
EQUIPMENT, ANESTHESIA
• Sterile polypropylene or red rubber-type catheter (or Foley catheter if indwelling); size ranges from 5-14 Fr, depending on the animal’s size.
POSSIBLE COMPLICATIONS AND COMMON ERRORS TO AVOID
• Incorrect orientation (female): to enter the vulva, the directing (index) finger must be oriented dorsally, not cranially.
• Incorrect catheter position (female): clitoral fossa externally and vagina internally must be avoided in catheterizing female dog.
• Forceful advancement of the catheter: catheter should pass with no resistance and no discomfort. Otherwise, consider free catch, cystocentesis, or sedation/anesthesia (if placing indwelling catheter).
PROCEDURE
• The dog stands on an exam table or on the ground (giant-breed dogs). The procedure is also easily performed on the laterally recumbent (e.g., anesthetized) dog.
• While an assistant holds the dog’s head, the examiner opens the sterile catheter wrapper, applies sterile lubricant to the catheter tip, and ensures that the catheter remains in a sterile field (e.g., the opened catheter wrapper).
• The right-handed examiner puts on sterile gloves, lubricates the right index finger, and takes the urinary catheter under his or her right index finger (palmar surface of finger), catheter tip at fingertip.
• The examiner lifts the dog’s tail with the left hand and places the index finger on the vulva, pointing dorsally, and then enters the vulva by gently everting the vulvar lips and pressing cranially.
• The examiner advances the index finger and catheter directly dorsally (i.e., toward the ceiling in the standing dog) to a narrowing, which is the vestibulovaginal junction. The ischial arch also is palpable cranially.
• The urethral opening is on the cranial/ventral surface in this region, immediately adjacent to the ischial arch. Therefore, the catheter is advanced under the index finger such that the index finger blocks off the vestibulovaginal route and directs the catheter ventrally into the urethra. There should be no palpable resistance when advancing the catheter.
• Once urine flows from the catheter, it is advanced 2-3 cm (small dog) to 10-15 cm (large dog) into its final position.
• An assistant positions himself or herself dorsal to the laterally recumbent dog, elevates the dog’s nondependent leg, and retracts the prepuce to expose the penis (see figure). A second assistant is usually needed to restrain the dog’s head.
• The sterile catheter is kept within its sterile wrapper except for the distal end of the wrapper, which is cut off.
• The next 3-5 cm of sterile wrapper is then cut and left overlying the catheter; it will serve as a sliding sterile cuff for advancing the catheter out of the wrapper and up the urethra, while keeping the clinician from touching the catheter directly with the fingers. This approach minimizes contamination without requiring sterile gloves.
Central Venous Pressure Monitoring
INDICATIONS
• Monitoring the administration of large volumes of fluids in animals that are in shock or have impaired urine production
CONTRAINDICATIONS
Only those of jugular catheter placement (e.g., hypercoagulable or hypocoagulable states, skin infection over jugular site; see p. 1293)
EQUIPMENT, ANESTHESIA
• CVP set (Universal CVP set [Abbott Laboratories]). Alternatively, if a purpose-made kit is not available, use the following: a sterile three-way stopcock, sterile IV fluid-type tubing, and a ruler (centimeter grading and at least 30 cm long). The sterile tubing is taped to the ruler, and it is held vertically, creating a simple pressure manometer.
PREPARATION: IMPORTANT CHECKPOINTS
• The prerequisite is the placement of an IV jugular catheter (see p. 1293):
○ The distal tip of this catheter should be in the cranial vena cava, just cranial to the right atrium, which allows accurate measurement of CVP without inducing premature atrial contractions. Confirm position radiographically.
PROCEDURE
Before connecting CVP set to patient’s jugular catheter:
• One end (female end) of the CVP set goes to a suspended IV fluid bag. The line on that end should be closed using a clamp.
• A second end connects to tubing that is fixed to a scale (e.g., manometer scale or ruler). This line also should be closed using a clamp.
• Using adhesive strips, fix the manometer on a vertical surface, such as the wall of a cage or a vertical stand, next to the animal and at a height such that the 0 on the scale of the manometer/ruler is at the level of the animal’s heart (right atrium).
• Then unclamp the manometer tubing and turn the three-way stopcock such that flow exists between the fluid line from the fluid bag and the fluid line of the manometer.
Connecting CVP set to patient:
• Next, connect the remaining end (male end) of the CVP set to the jugular catheter. The stopcock is usually integral to the set.
• Finally, turn the stopcock so that a column of fluid exists between the jugular catheter and the manometer.
POSSIBLE COMPLICATIONS AND COMMON ERRORS TO AVOID
• One must allow fluid to flow frequently through the IV line to avoid plugging of the catheter by a blood clot. Periodic flushing with heparinized saline is desirable.
• Cardiac function needs to be factored in when interpreting results. For example, animals with tricuspid regurgitation may have elevated CVP irrespective of intravascular volume loads.
• The CVP level represents mostly loading conditions of the right side of the heart (right ventricular preload), not afterload.
(Reprinted with permission from Sattler FP: Shock. In Kirk RW, editor: Current veterinary therapy III, Philadelphia, 1968, Saunders.)
INTERPRETATION
• Values above +10 cm H2O may indicate too much blood volume expansion. IV fluids should no longer be administered.
ALTERNATIVES AND THEIR RELATIVE MERITS
• Blood pressure (BP) monitoring: easier to use and represents another way to monitor cardiovascular status and the impact of IV fluids. This is the best alternative to CVP monitoring. It does not require an IV jugular catheter.
• Weight: can be used for documenting rehydration; excessive weight gain could be a signal to slow/stop IV fluids.
• Urine specific gravity: a drop in the specific gravity could document successful restoration of plasma volume but is very dependent on kidney function.
• Pulmonary capillary wedge pressure/Swan-Ganz catheterization: accurately measures left-sided heart pressures, which CVP does not; however, this technique is much more technically demanding.
Cerebrospinal Fluid Collection
OVERVIEW AND GOALS
• To safely collect an uncontaminated sample of cerebrospinal fluid (CSF) from an animal with suspected central nervous system (CNS) disease.
CONTRAINDICATIONS
• Increased intracranial pressure (depressed mental status, bradycardia, hypertension, miosis, anisocoria)
EQUIPMENT, ANESTHESIA
• A few 20- or 22-gauge, 1½-inch spinal needles (2½ – or 3½-inch needle may be needed for large dogs or for lumbar approach).
PREPARATION: IMPORTANT CHECKPOINTS
• Performed preferably after advanced imaging has ruled out both noninflammatory processes (neoplasia, malformation, vascular) and obvious increased intracranial pressure (coning and caudal displacement of the cerebellum, flattening of gyri, loss of sulci).
• Make arrangements for laboratory transport within 30 minutes of collection, or have in-house analysis equipment prepared and calibrated.
• If immediate analysis of the CSF is not possible, special preservation techniques may be required (prior laboratory consultation is recommended).
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