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Fluid Therapy
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CHAPTER
32
Fluid Therapy Christopher Cebra
Indications Fluid therapy is an important part of the treatment plan for most sick camelids. As a rule of thumb, when camelids are recognized to be ill, they often have moderately to severely advanced disease. They are often dehydrated to some degree and may be azotemic and have electrolyte or acid–base imbal ances as well. Camelids recognized as being ill often respond best to an aggressive treatment approach, which includes sup plemental fluids. That being said, the volume of fluids given under these circumstances need not always be “aggressive.” In many cases, modest volumes of fluids evoke the desired response, and greater volumes are unnecessary and potentially detrimental. The goals of fluid therapy in camelids are similar to those in other species. The major indications are to correct dehydra tion; to increase cardiac output, arterial blood pressure, and the perfusion of various organs; to correct electrolyte, mineral, and acid–base imbalances; to affect diuresis and increase uri nation; to provide a vehicle for other treatments; rarely to increase GI, pulmonary and other secretions; and ultimately to maintain a corrected patient at an acceptable level of the above functions. Generally, three main indications exist for replacement fluids in camelids and a myriad of minor applications. The three major indications are (1) rapidly dehydrating con ditions, where large-volume replacement may be indicated; (2) slowly dehydrating conditions, where replacement
volumes are modest, but continued need is likely; and (3) conditions of vasculopathy or decreased cardiac output. Minor indications exclusive of these include whole blood or plasma transfusions to treat anemia or hypoproteinemia and the use of crystalloid fluids to administer or decrease the toxic effects of other medications. Rapidly dehydrating conditions are the least common of the three main applications. Examples include acute C1 aci dosis, diarrhea, loss of ingesta and secretions because of esophageal obstruction or toxic gastritis (rhododendron and related plants), and intestinal strangulations. A neonate that has never nursed could also fall into this category. These animals are characterized by having acute copious loss of water and usually electrolytes, without necessarily losing or catabolizing much protein in the process. Thus, they may have hyperalbuminemia or hyperproteinemia and packed cell volume (PCV) values at the higher end of the reference range. Slowly dehydrating conditions are common and numer ous. Thus, a variety of other abnormalities may accompany dehydration, and the animal may have adapted somewhat to compensate for the gradual fluid loss. Affected camelids frequently have low-grade anemia of chronic disease and evi dence of protein loss or catabolism, and this may not have blood evidence of hemoconcentration. Vasculopathies and processes that depress cardiac output without specifically causing dehydration are also relatively common. Examples include septic shock caused by neonatal
Chapter 32 • Fluid Therapy bacterial infection, a ruptured ulcer, bacterial enteritis, or streptococcal peritonitis, heat stress, and ingestion of cardio toxic compounds. Affected camelids may or may not be dehydrated but may display evidence of lack of adequate per fusion. These conditions may also complicate fluid therapy protocols because leaky vessels and poor heart function may negate some of the benefits of the additional intravascular volume.
Assessment Determining the need for fluid therapy and formulating a plan for routes, rates, and components involve a combination of clinical intuition, physical examination findings, and labora tory analyses. Physical examination is most useful for finding signs of overt dehydration or subtler signs of poor peripheral perfusion. This may be complicated in camelids because, as a race, they appear to be well adapted to intermittent feed and water deprivation and may endure dehydration better com pared with other domestic hoofstock species. Sunken, dull eyes, dry mucous membranes, and prolonged skin tent may all be found. Neck and eyelid skin appear to be the best places to assess skin tent time. Jugular fill is difficult to assess, but lack of filling after a long period of holding off the vein may provide some indication of severe dehydration. Heart rate may also increase, but it is important to note that dehydrated or adult camelids in shock frequently have heart rates ranging from 72 to 90 beats per minute (beats/min), which are within some published reference ranges. All heart rates greater than 72 beats/min should be considered potentially abnormal in adult camelids, as they may be a sign of dehydration, low cardiac output, or stress. Behavioral evidence of dehydration or poor cardiac output may include water-seeking behavior, lethargy, inappetence, and decreased urination. Clinicopathologic evidence of dehy dration includes hemoconcentration, evidenced by a high PCV, high blood protein or albumin concentrations, or both. It is crucial to keep in mind that these abnormalities are fre quently masked by concurrent protein loss, catabolism, blood loss, anemia, or chronic disease and that refractometer read ings of protein may be confounded and artificially high in camelids with hyperlipemia. Other laboratory evidence of dehydration includes the presence of azotemia, lactic acidosis, and electrolyte abnormalities. Azotemia may be masked or difficult to interpret in some cases; with acute inappetence, blood urea moves into the forestomach and is deaminated by gastric microbes. This effect lasts approximately 3 days, after which microbial die-off diminishes this catabolic pathway. Blood creatinine may also be low in camelids with chronically low muscle mass, as in neonates or chronic poor-doers. To avoid being confounded by one of these processes, measure ment of both blood urea nitrogen (BUN) and creatinine con centrations is recommended. These confounders may also affect the rule of thumb, that a BUN-to-creatinine ratio (both in milligrams per deciliter [mg/dL]) of 20 : 1 or greater reflects prerenal azotemia, so this interpretation should be made with some caution. Lactic acidosis may reflect global (hypovolemia, poor cardiac output) or focal (strangulation, thrombosis) anaerobic metabolism. Both are indications for fluid therapy and may be caused by overt dehydration or other forms of
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shock. Electrolyte abnormalities are suggestive of pathologic conditions associated with dehydration and thus may be helpful. Hypernatremia is common in camelids and usually reflects loss of water from hyperglycemia and glucose diuresis. Moderate to severe hypochloremia, especially when accompa nied by metabolic alkalosis, often reflects a small intestinal obstruction with decreased gastrointestinal absorption and pooling of water in the intestinal tract. Hyponatremia, espe cially when accompanied by hypochloremia and metabolic acidosis, often indicates loss of fluid and electrolyte through the digestive tract, especially when it is caused by enteritis or esophageal obstruction. Decreased pulse strength, poor mucous membrane color, and slow capillary refill time may all indicate dehydration or poor cardiac output. These may be further confirmed by diagnostic tests, including measurement of central venous pressure or cardiac function. In addition to the difficulty in recognizing the need for fluid therapy, a number of other factors must be considered. Sick camelids are often fragile. They may not be as tolerant of procedures as healthy camelids would be, and undergoing too much stress may cause a rapid decline in their condition. Catheterization attempts should be carefully planned for success, other invasive or semi-invasive treatments or diagnos tic tests should be performed gradually or delayed until the patient has had some time to recover, and some invasive interventions should not be attempted at all. Also, the need for fluids and concerns of edema are closely related. The high prevalence of hypoalbuminemia (61% of our hospi tal population of camelids) and hypoproteinemia (35%) in sick camelids dictates conservative rates of fluid administra tion or use of a colloid solution. Even normoproteinemic camelids frequently develop hypoproteinemia during hospi talization, so monitoring this aspect of the patient must be an ongoing process. In addition to dictating the frequent use of colloids, camelid physiology and pathophysiology affect the value of other components of fluids. Sick camelids are often hyperglycemic and assimilate exogenous glucose poorly, so it should not be given without indication, and concurrent use of insulin or insulinotropic medications should be considered. Lactate is also likely to accumulate in sick camelids, so it should likewise be avoided in fluids, when possible.
Routes Once the decision has been made to administer replacement fluids, the route, rate, and components of the fluid must be selected. Of the possible routes, oral and intravenous routes are the major routes used to correct problems of hydration. Subcutaneous, intraperitoneal, and intraosseous routes have specialty applications but are not useful or necessary in most situations.
Oral Fluids and Orogastric Intubation Allowing a thirsty camelid to drink water is the simplest way to provide oral fluids. Overhydration by this route is extremely uncommon, and polydipsia is rarely encountered. The amount that can be offered at one time is unknown. It is likely that water flows aborad relatively quickly in camelids with good gastrointestinal function, but this cannot be ensured in sick
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camelids. Electrolyte solutions may be used in place of water if replacement is necessary, but the high prevalence of hyper natremia in sick camelids mandates that these solutions not be used without evidence of need. If the thirst response is inadequate, which is regrettably the case in many sick camelids, oral fluids are best administered via an orogastric tube. Small diameter tubes may also be passed through the nose, but camelids have narrow nasal pas sages and often resist this procedure more than the passing of a tube through the mouth. However, in crias, small-bore feeding tubes may be passed nasally and left in place. Passing an orogastric tube in an adult camelid may induce struggling and a stress response, so it is important to assess whether the patient can tolerate the procedure, to make a plan to perform the procedure quickly and competently, and to abort the procedure, if necessary. Alpacas may be manually restrained, whereas llamas may be better restrained in a llama chute. Light sedation may help, for example, with butorpha nol (0.022 to 0.05 milligrams per kilogram [mg/kg], intramus cularly [IM]), which has fewer deleterious effects on airway protective responses or cardiac output compared with many other sedatives. To prevent damage to the tube because of chewing by adolescents or adults, a piece of PVC pipe or a block of wood with the edges rounded and a hole drilled through the center for the tube should be used as a speculum (Figure 32-1); crias may be tubed without a speculum. When a block speculum is used, it is introduced from the side of the mouth into the interdental space and seated on the molars. The tube is passed through the hole, over the base of the tongue, and into the esophagus. Negative pressure when sucking back on the tube, palpating the tube adjacent to the cervical trachea, or hearing fluid bubbles over the abdomen when blowing on the tube confirm correct placement. The camelid may respond by struggling, regurgitating, or going into respiratory distress. Any of these may lead to reassessment of the adequacy of the restraint or the safety of the procedure and potentially cancelling or delaying it. Without knowing the amount of fiber fill, it is difficult to estimate gastric fluid capacity. Giving a maximum of 3% of
Figure 32-1 A stomach tube and block speculum appropriate for orogastric intubation of an adult llama or alpaca. Adult llamas usually tolerate tubes with an external diameter of up to 5/8” well (top), and 1 2” tubes are commonly used in adult alpacas (bottom).
body weight by volume to an adult alpaca, 4% to an adult llama, and 3.3% to neonates of either species appears to be safe and causes only minimal regurgitation. That amount may be given every few hours, unless gastric distention develops, but to avoid stress and trauma, most patients should be given no more than three tubings per day. The easiest thing to give by tube is water. In dehydrated camelids that have a tendency toward hypernatremia and hyperchloremia, this may be acceptable or even preferable to salt solutions, but if indicated, electrolyte solutions may be used. Except in neonates, oral sugar is usually contraindicated because it is rapidly fermented by gastric microbes. In neo nates, milk or a milk replacer often makes the best oral replacement fluid.
Intravenous Catheterization Because the thirst response is unreliable in many sick came lids, and many require some degree of rapid fluid resuscita tion, the intravenous route is often preferable to others. Much has been made over the years of the difficulties of jugular catheterization in camelids. Although inherently more diffi cult in adult camelids than in many other domestic species because of the thick skin, possible presence of thick fleece, lack of a visible vein, and somewhat different course of the jugular vein, it is a skill that is quickly mastered using proper land marks, restraint, and technique. Mature intact male llamas have the thickest skin, often over 0.5 centimeters (cm) wide. Late-neutered males are similar. Alpacas, females, and crias have increasingly thinner skin, and often have a visible jugular vein. An exception occurs in some family lines of Suri alpacas, in which the skin of all ages may have thick folds (rolling or wrinkled skin). Some form of restraint is usually necessary to hold the neck straight and prevent movement and contamination of the site during the procedure. Neonates may be manually restrained in the standing position or in sternal or lateral recumbency (Figure 32-2). Adult alpacas may be restrained manually, usually in the standing position or in sternal recumbency
Figure 32-2 Restraint of a neonatal cria in lateral recumbency for jugular catheterization. Placement of a towel over the eyes decreases activity.
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Figure 32-3 Manual restraint of an adult alpaca for jugular catheter
Figure 32-4 Making the guide incision with a #15 scalpel blade after
ization. The neck should be held upright with minimal twisting.
injection of a local anesthetic agent. Note the elevation of the skin lower on the neck to facilitate a full-thickness incision without punc turing the vein and the incision made at a similar angle as is planned for the catheter.
(Figure 32-3). Manual restraint of some adult llamas is pos sible as well, and restraint chutes often facilitate the procedure, especially when restrainers are in short supply. If the animal is uncooperative and moving excessively during the procedure, restraint should be modified to compensate for this, rather than pressing forward and engaging in a frustrating and fre quently unsuccessful exercise. Butorphanol may be used as a sedative. Xylazine and similar agents are less desirable because they may decrease cardiac output and slow jugular filling. As with all procedures with sick camelids, if done efficiently, the risk is minimal, but if the camelid starts to display a stress response such as displacement of the soft palate and dyspnea, regurgitation, violent struggling, or arching the head over the back with strabismus, the procedure should be discontinued until the patient has recovered. As a longstanding practice and to avoid damage to the esophagus, the right jugular is more commonly used. If neces sary, the left vein may be used instead. The jugular vein runs dorsolateral to the trachea and ventromedial to the transverse processes of the cervical vertebrae. It is somewhat more ven tromedial than in horses and often approached best from closer to the front of the patient and not the side to avoid accidental puncture of the carotid artery. The vein runs most linearly and most superficially in the cranial half of the neck. In the caudal half, it dives slightly and curves with the neck toward the thorax. In neonates, some alpacas, and the rare llama, the vein may be seen to bulge visibly upon holding it off near the base of the neck. If the vein cannot be visualized by holding it off, often a palpable bulge is still formed and may be found by bouncing a finger on the site while releasing the occluding pressure. If necessary, the neck may be scanned ultrasonographically to find the held-off vein, or if nothing else works, the point midway between the trachea and the ventral extent of the transverse processes of the cervical verte brae approximately one third of the distance from the ramus on the mandible toward the shoulders is a good place to try.
It is desirable to clip the fleece and aseptically prepare the site. Some owners may be reluctant to allow the clipping in show animals, so some conversation on the trade-off between a cosmetic result and an efficient procedure may be required. The site is then infiltrated with a small volume (0.5 milliliter [mL] in adult and 0.25 mL in neonates) of local anesthetic. Using larger volumes in one place may lead to compression of the vein. The skin is lifted and punctured to the full thick ness in the anesthetized area with a #15 scalpel blade or a hypodermic needle with a diameter larger than that of a cath eter (Figure 32-4). Ideally, this puncture should be directly over the vein at a 25-degree angle to the skin surface or shal lower, directed toward the base of the neck, providing a course for the catheter to follow without resistance. If the angle is too steep, the catheter will be deflected or even kink during passage. If the skin is not punctured to full thickness prior to catheterization, often substantial resistance to the catheter tip and inward dimpling of the skin would occur. The extra force required to overcome this resistance may propel the tip of the catheter through the vein without the operator realizing it, causing accidental puncture of the carotid artery. Recognizing the skin drag and repeating or deepening the skin incision is usually wiser than simply ignoring the resistance and proceed ing with catheter placement. A variety of catheters is available and has been used. I prefer polypropylene over-the-needle catheters for most indications because they have little tissue drag and appear to be suffi ciently durable and antithrombogenic for most applications. For most adults, 14- or 16-gauge catheters are ample and 18or 20-gauge are adequate for neonates. For crias or adults that require a catheter for extended periods, a 16-gauge polyure thane over-the-wire catheter may be more appropriate; they are less thrombogenic and also tend to kink or be dislodged
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Figure 32-6 Three catheters placed through guide incisions of differ ent angles. Note the kinks in the two catheters with the steeper angles (top and middle), and the lack of a kink in the catheter with the shal lowest angle (bottom).
Figure 32-5 Introducing the catheter through the guide incision. Note the angle and the catheter being advanced parallel to the course of the vein.
to a lesser degree. The latter are technically more demanding to place and require strict attention to sterility during place ment. In well-hydrated adult camelids, these catheters may be flushed with heparinized saline and introduced as in many other species, but camelids in shock may not have enough jugular pressure to cause the blood flash in the hub upon suc cessful introduction into the jugular lumen. For that reason, although it requires more hands, it is often preferable to attach an extension set and half-filled 20-mL or larger flush syringe to the hub of the catheter during its introduction. An assistant controlling this syringe gently aspirates as soon as the tip of the catheter enters the skin. Especially with larger-bore cath eters, blood should come back easily once the catheter tip enters the vein. This procedure is not always preferable to straight catheterization in neonates, in whom the diameter of the catheter is often too small to appreciate ease of flow and in which the pressure of aspiration may collapse the vein. Additionally, in crias with hypovolemia, blood pressure may be so poor that it is not able to displace a column of heparin ized saline within the catheter. Therefore, in crias, it may be advisable to flush the catheter and then let the flush drain through prior to placement. The placer must then be patient, slowly advancing the catheter and pausing to observe if blood fills the catheter hub. For introduction, the vein should be held off well down the neck. In dehydrated patients, holding off for up to a minute may be necessary to allow sufficient jugular filling. The catheter stylet is held at the hub and directed at a 45- to 25-degree angle downward, following the course of the guide hole and vein (Figure 32-5). Steeper angles are permissible if the stab incision is wide enough to accommodate the subse quent reduction of angle without kinking the catheter (Figure 32-6). The smaller fingers of the hand may be used to steady it against the animal’s neck and maintain the proper angle. If an extension set is attached, the assistant begins to aspirate.
The stylet is slowly advanced until the hub fills with blood or the assistant reports good flow. The tip of the stylet is advanced an additional 0.5 to 1 cm and rechecked for flow. If it is still good, the angle of the hub to the skin is reduced to 20 to 25 degrees, and the stylet is advanced an additional 1 to 2 cm to seat the catheter in the vein. If the flow is still good, the stylet is held immobile, and the catheter is advanced its full length into the vein. A slight twisting motion decreases skin drag and may allow the tip of the catheter to roll off the valves within the jugular. If severe resistance is encountered, it is possible that the catheter is not in the vein and the procedure must be stopped and attempted again. If the catheter seats well, the hub may be secured to the skin with tissue glue or suturing, potentially wrapped, and then used. If catheterization is unsuccessful but the jugular is punc tured, pressure should immediately be released on the lower vein and pressure applied to the puncture site for 2 minutes. If the carotid artery is punctured, the site should be held with firm pressure for a minimum of 5 minutes. In either case, timely recognition of the problem minimizes ensuing compli cations. Significant cervical hemorrhage makes subsequent catheterization attempts difficult both by compressing the jugular vein and by creating a confounding, flashlike effect when the tip of the catheter enters the hematoma. If a poten tial site is occluded by a hematoma, another site should be selected. In all cases, the first catheterization attempt is usually the easiest, so every effort should be made to make the first attempt a successful one. A small number of camelids will have significant acute complications to an unsuccessful jugular catheterization attempt. These likely result from subcutaneous hemorrhage and the pressure it generates on underlying structures. Clinical signs commonly include weakness, lethargy, and recumbency and may also include upper airway obstruction with stridor, nystagmus, and dorsoflexion of the head and neck. Little can be done at the time beyond maintaining the head and neck in an extended position to open the airway, and minimizing other stressors. These complications are best avoided through good catheterization technique. In some cases, use of the jugular vein is impossible or undesirable. In camels, the lateral thoracic veins are often more accessible, but these are difficult to use in smaller
Chapter 32 • Fluid Therapy camelids and even in large llamas. Cephalic vein catheters have sometimes been used, especially in crias; medially, leg veins, mainly the medial saphenous veins, are occasionally accessible in recumbent animals. Ear veins may be used for very-fine-gauge catheters, but they are hard to maintain for very long.
Intravenous Fluids In general, intravenous fluid therapy may be divided into two phases: (1) replacement and (2) maintenance. Replacement in most species involves estimation of the fluid deficit and delivery of that deficit over a short period. In camelids, estima tion of the deficit may be difficult, as many of the disorders lead to dehydration gradually, allowing the animal time to adapt, and the high prevalence of hypoproteinemia increases the risk of edema after bolus fluids. Since camelids appear to be relatively tolerant of dehydration, they are probably also more tolerant of slow replacement of deficits than of bolus correction. Unless dehydration is clear and lack of rapid cor rection has life-threatening implications, initial boluses of more than 2% of body weight to sick camelids are seldom necessary and occasionally detrimental. If dehydration is clear and life threatening, boluses up to 4% of body weight may be indicated, and in extremely rare cases such as in neonates with hyperosmolar disorder and hyperalbuminemia, a bolus of up to 10% of body weight may be necessary, but these are the exceptions. The risk of larger boluses is edema. In hypoproteinemic camelids, edema may rapidly impair pulmonary function, and the camelid may die of respiratory distress. This easily out weighs the risk of prolonged underperfusion in most cases. The major risks of underperfusion relate to other medications: both aminoglycoside antibiotics and nonsteroidal antiinflam matory drugs (NSAIDs) are more toxic to the dehydrated animal and therefore must be used with caution, or not at all, in camelids with questionable circulatory volume and function. Maintenance fluids come with the same warning. Hypo proteinemia and edema are the two biggest risks. Very little information about maintenance requirements in camelids is available: rates from 2% to 10% of body weight per day have been reported in the literature. Except maybe for neonates, 10% is clearly too high, and the risk of edema formation is significant. Two percent may be adequate, but hardly justifies the catheter. Coupled with a conservative replacement phi losophy, administration of maintenance fluids at a slightly higher rate than the bare minimum to correct remaining defi cits appears to be safe and effective. Roughly 2 mL/kg/hr or a volume equal to 5% percent of body weight over 24 hours appears to have little risk, promotes renal perfusion and urina tion such that iatrogenic toxicities are rare, and does not affect blood protein concentrations too much. Blood protein con centration should be monitored at least twice a day during fluid therapy. In some instances, as when the patient cannot be observed frequently or is co-housed with other animals, it may be desir able to give intermittent boluses instead of a constant infu sion. Boluses work well when given every 3 to 4 hours, but additives such as glucose or potassium may be more toxic or less well assimilated. These are relatively minor concerns, so
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the convenience of boluses frequently outweighs these disad vantages in many field settings or when treating a nursing cria. The next question is what to give. Hypernatremia and hyperchloremia are far more common than hyponatremia and hypochloremia, and hypokalemia is common. Acidosis is a rare but an important finding. Alkalosis is more common but is usually mild and benign. Mineral abnormalities are likewise usually mild and benign. Fluids may be adjusted on the basis of individual abnormalities, but the important point to note is that a plain, balanced electrolyte solution is well suited to replacement therapy, and the same fluid with supplemental potassium (and maybe calcium) is suited to maintenance therapy. Because of issues with hyperlactemia and poor glucose assimilation, fluids with acetate as the buffer base may be preferable to those that use lactate. In cases of metabolic alka losis, 0.9% saline is often preferable to pH-balanced fluids. In some cases, additional supplementation is indicated. Many sick camelids have hypokalemia; on the basis of the recognition that measurement of blood potassium is a poor estimator of whole body stores of this principally intracellular cation, supplemental potassium is often added to mainte nance intravascular fluids to bring the final concentration to 20 to 30 milliequivalents per liter (0.5 mEq/kg/hr). At the rate of 2 mL/kg/hr, this provides supplementation well within the margin of safety. Hypocalcemia may be addressed without elaborate calculation by giving 0.5 to 1 mL/kg of 23% calcium gluconate solution by slow injection or by adding this to intravenous fluids. Hypoglycemia may likewise be addressed by bolus administration of 0.5 to 1 mL/kg of 50% dextrose solution, or more gradually by supplementing fluids to a final concentration of 5% dextrose. Up to two thirds of the camelids at our clinic show evi dence of fat mobilization. If this is severe enough to warrant concern, we often use a partial parenteral nutrition formula for our maintenance fluids. Our most common recipe for this is as follows: 5000 mL Acetate-containing electrolyte solution 1000 mL 8.5% amino acid solution 500 mL 50% dextrose 130 mEq Potassium (in potassium chloride [KCl]) 20 mL B vitamins We administer this solution at 5% of body weight/day (2 mL/kg/hr) using infusion pumps. Both B vitamins and amino acid solutions are light sensitive, so the intravenous bags must be covered. We also administer insulin during the course of peripheral parenteral nutrition (PPN) treatment. For most camelids, we use an ultralente form (0.2 to 0.4 units per kilogram [units/kg], subcutaneously [SC], q24 hours). For overtly lipemic camelids or for ones with very high β-hydroxybutyrate or nonesterified fatty acid (NEFA) values, we sometimes use ultralente insulin as often as twice a day (it lasts 12 to 18 hours) or supplement it with intravenous regular insulin at 0.2 units/kg as often as hourly. The rationale for the PPN is as follows: We administer the insulin to arrest fat mobilization and promote triglyceride clearance. We administer the dextrose to prevent hypoglyce mia and also to provide some alternate energy to the patient. We administer the amino acid solution to provide some energy substrate and hopefully spare the protein-scavenging, cachexia effect. The B vitamins might also increase the usabil ity of the glucose. We are able to usually arrest fat mobilization
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BOX 32-1
Examples of Practical Fluid Therapy
CASE #1 A 4-year-old, 50-kg alpaca dam with a high Haemonchus load refuses to stand or eat. Physical examination reveals a heart rate of 96 beats per minute (beats/min), a respiratory rate of 20 breaths per minute (breaths/min), and a rectal temperature of 100.4°F (38°C). Blood analysis reveals the following results: packed cell volume (PCV) = 12%, total plasma protein concentration = 4 g/dL, albumin = 1.5 g/dL, blood urea nitrogen (BUN) = 73 mg/dL, creatinine = 3.6 mg/dL, sodium = 136 mEq/L, and lactate = 2.2 mg/dL. Clear evidence of shock and dehydration suggests the need to do something, most likely by the intravenous route. However, given the low PCV and plasma protein concentration, use of whole blood or a colloid would be ideal. In the meantime, this animal might benefit from conservative bolus treatment (2% of body weight = 1 L) of a pH-balanced, polyionic solution, followed by 100 mL/hr of the same solution for a few hours, until a colloid or blood is available. Large boluses and high rates should be avoided to prevent edema formation. CASE #2 A 2-year-old, 40-kg male alpaca is suspected of having acute C1 acidosis. Physical examination reveals a heart rate of 120 beats/min, a respiratory rate of 30 breaths/ min, and a rectal temperature of T = 104°F (40°C). Blood analysis reveals the following results: PCV = 35%, total plasma protein concentration = 6.7 g/dL, albumin = 4.1 g/dL, BUN = 95 mg/dL, creatinine = 4 mg/dL, sodium = 152 mEq/L, and lactate = 7.4 mg/dL. Evidence of shock and dehydration, including relatively high protein concentrations, is clear. Given these and the likelihood that the dehydration is caused by an acute process, this camelid would be more likely to need and tolerate a larger fluid bolus than average. A 2-L bolus (5% of body weight) would be a good start, followed by 83 mL/hr for 4 to 24 hours, depending on the severity. Fluid should be alkalinizing in this case.
and clear hypertriglyceridemia within 48 hours, using this treatment scheme.
Colloids Because of the high prevalence of hypoproteinemia and the somewhat lower prevalence of anemia in sick camelids, the use of colloids and whole blood as replacement fluids is relatively common. The use of whole blood is discussed in Chapter 36. General indications for the use of colloids are when total plasma protein concentration is less than 4 g/dL or albumin is less than 2 g/dL. Animals with failure of passive transfer or sepsis may also benefit from the antibodies provided in plasma. Colloids may come from natural plasma products or synthetic compounds. Camelid plasma is commercially avail able in many countries and is generally the preferred form of colloid. Transfusion reactions are rare, and plasma from one New World camelid species appears suitable for transfusion to another species. In Australia, dromedary plasma is commer cially available and marketed for use in alpacas. In a 50-kg sick camelid, 1 L of donor plasma usually increases blood protein concentrations by approximately 2 g/dL after the additional water volume is excreted. Plasma transfusions are administered at a slow rate for 15 to 20 minutes through a filtered blood administration set while the animal is observed for evidence of a reaction, includ ing restlessness, straining, hyperthermia, aberrant chewing, and skin reactions. If no reaction is noted, the remainder of the plasma is given over 2 to 4 hours. More rapid administra tion has been associated with respiratory distress. Hetastarch represents an alternative to plasma. It has been used safely in camelids at dose rates of 10 to 20 mL/kg of a 6% solution.1 Hetastarch is not detected by biochemical tests of blood protein but is detected by using a refractometer (Box 32-1).
REFERENCE 1. Carney KR, et al: Evaluation of the effect of hetastarch and lactated Ringer’s solution on plasma colloid osmotic pressure in healthy llamas, J Am Vet Med Assoc 238:768-772, 2011.
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CHAPTER
32
Fluid Therapy Christopher Cebra
Indications Fluid therapy is an important part of the treatment plan for most sick camelids. As a rule of thumb, when camelids are recognized to be ill, they often have moderately to severely advanced disease. They are often dehydrated to some degree and may be azotemic and have electrolyte or acid–base imbal ances as well. Camelids recognized as being ill often respond best to an aggressive treatment approach, which includes sup plemental fluids. That being said, the volume of fluids given under these circumstances need not always be “aggressive.” In many cases, modest volumes of fluids evoke the desired response, and greater volumes are unnecessary and potentially detrimental. The goals of fluid therapy in camelids are similar to those in other species. The major indications are to correct dehydra tion; to increase cardiac output, arterial blood pressure, and the perfusion of various organs; to correct electrolyte, mineral, and acid–base imbalances; to affect diuresis and increase uri nation; to provide a vehicle for other treatments; rarely to increase GI, pulmonary and other secretions; and ultimately to maintain a corrected patient at an acceptable level of the above functions. Generally, three main indications exist for replacement fluids in camelids and a myriad of minor applications. The three major indications are (1) rapidly dehydrating con ditions, where large-volume replacement may be indicated; (2) slowly dehydrating conditions, where replacement
volumes are modest, but continued need is likely; and (3) conditions of vasculopathy or decreased cardiac output. Minor indications exclusive of these include whole blood or plasma transfusions to treat anemia or hypoproteinemia and the use of crystalloid fluids to administer or decrease the toxic effects of other medications. Rapidly dehydrating conditions are the least common of the three main applications. Examples include acute C1 aci dosis, diarrhea, loss of ingesta and secretions because of esophageal obstruction or toxic gastritis (rhododendron and related plants), and intestinal strangulations. A neonate that has never nursed could also fall into this category. These animals are characterized by having acute copious loss of water and usually electrolytes, without necessarily losing or catabolizing much protein in the process. Thus, they may have hyperalbuminemia or hyperproteinemia and packed cell volume (PCV) values at the higher end of the reference range. Slowly dehydrating conditions are common and numer ous. Thus, a variety of other abnormalities may accompany dehydration, and the animal may have adapted somewhat to compensate for the gradual fluid loss. Affected camelids frequently have low-grade anemia of chronic disease and evi dence of protein loss or catabolism, and this may not have blood evidence of hemoconcentration. Vasculopathies and processes that depress cardiac output without specifically causing dehydration are also relatively common. Examples include septic shock caused by neonatal
Chapter 32 • Fluid Therapy bacterial infection, a ruptured ulcer, bacterial enteritis, or streptococcal peritonitis, heat stress, and ingestion of cardio toxic compounds. Affected camelids may or may not be dehydrated but may display evidence of lack of adequate per fusion. These conditions may also complicate fluid therapy protocols because leaky vessels and poor heart function may negate some of the benefits of the additional intravascular volume.
Assessment Determining the need for fluid therapy and formulating a plan for routes, rates, and components involve a combination of clinical intuition, physical examination findings, and labora tory analyses. Physical examination is most useful for finding signs of overt dehydration or subtler signs of poor peripheral perfusion. This may be complicated in camelids because, as a race, they appear to be well adapted to intermittent feed and water deprivation and may endure dehydration better com pared with other domestic hoofstock species. Sunken, dull eyes, dry mucous membranes, and prolonged skin tent may all be found. Neck and eyelid skin appear to be the best places to assess skin tent time. Jugular fill is difficult to assess, but lack of filling after a long period of holding off the vein may provide some indication of severe dehydration. Heart rate may also increase, but it is important to note that dehydrated or adult camelids in shock frequently have heart rates ranging from 72 to 90 beats per minute (beats/min), which are within some published reference ranges. All heart rates greater than 72 beats/min should be considered potentially abnormal in adult camelids, as they may be a sign of dehydration, low cardiac output, or stress. Behavioral evidence of dehydration or poor cardiac output may include water-seeking behavior, lethargy, inappetence, and decreased urination. Clinicopathologic evidence of dehy dration includes hemoconcentration, evidenced by a high PCV, high blood protein or albumin concentrations, or both. It is crucial to keep in mind that these abnormalities are fre quently masked by concurrent protein loss, catabolism, blood loss, anemia, or chronic disease and that refractometer read ings of protein may be confounded and artificially high in camelids with hyperlipemia. Other laboratory evidence of dehydration includes the presence of azotemia, lactic acidosis, and electrolyte abnormalities. Azotemia may be masked or difficult to interpret in some cases; with acute inappetence, blood urea moves into the forestomach and is deaminated by gastric microbes. This effect lasts approximately 3 days, after which microbial die-off diminishes this catabolic pathway. Blood creatinine may also be low in camelids with chronically low muscle mass, as in neonates or chronic poor-doers. To avoid being confounded by one of these processes, measure ment of both blood urea nitrogen (BUN) and creatinine con centrations is recommended. These confounders may also affect the rule of thumb, that a BUN-to-creatinine ratio (both in milligrams per deciliter [mg/dL]) of 20 : 1 or greater reflects prerenal azotemia, so this interpretation should be made with some caution. Lactic acidosis may reflect global (hypovolemia, poor cardiac output) or focal (strangulation, thrombosis) anaerobic metabolism. Both are indications for fluid therapy and may be caused by overt dehydration or other forms of
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shock. Electrolyte abnormalities are suggestive of pathologic conditions associated with dehydration and thus may be helpful. Hypernatremia is common in camelids and usually reflects loss of water from hyperglycemia and glucose diuresis. Moderate to severe hypochloremia, especially when accompa nied by metabolic alkalosis, often reflects a small intestinal obstruction with decreased gastrointestinal absorption and pooling of water in the intestinal tract. Hyponatremia, espe cially when accompanied by hypochloremia and metabolic acidosis, often indicates loss of fluid and electrolyte through the digestive tract, especially when it is caused by enteritis or esophageal obstruction. Decreased pulse strength, poor mucous membrane color, and slow capillary refill time may all indicate dehydration or poor cardiac output. These may be further confirmed by diagnostic tests, including measurement of central venous pressure or cardiac function. In addition to the difficulty in recognizing the need for fluid therapy, a number of other factors must be considered. Sick camelids are often fragile. They may not be as tolerant of procedures as healthy camelids would be, and undergoing too much stress may cause a rapid decline in their condition. Catheterization attempts should be carefully planned for success, other invasive or semi-invasive treatments or diagnos tic tests should be performed gradually or delayed until the patient has had some time to recover, and some invasive interventions should not be attempted at all. Also, the need for fluids and concerns of edema are closely related. The high prevalence of hypoalbuminemia (61% of our hospi tal population of camelids) and hypoproteinemia (35%) in sick camelids dictates conservative rates of fluid administra tion or use of a colloid solution. Even normoproteinemic camelids frequently develop hypoproteinemia during hospi talization, so monitoring this aspect of the patient must be an ongoing process. In addition to dictating the frequent use of colloids, camelid physiology and pathophysiology affect the value of other components of fluids. Sick camelids are often hyperglycemic and assimilate exogenous glucose poorly, so it should not be given without indication, and concurrent use of insulin or insulinotropic medications should be considered. Lactate is also likely to accumulate in sick camelids, so it should likewise be avoided in fluids, when possible.
Routes Once the decision has been made to administer replacement fluids, the route, rate, and components of the fluid must be selected. Of the possible routes, oral and intravenous routes are the major routes used to correct problems of hydration. Subcutaneous, intraperitoneal, and intraosseous routes have specialty applications but are not useful or necessary in most situations.
Oral Fluids and Orogastric Intubation Allowing a thirsty camelid to drink water is the simplest way to provide oral fluids. Overhydration by this route is extremely uncommon, and polydipsia is rarely encountered. The amount that can be offered at one time is unknown. It is likely that water flows aborad relatively quickly in camelids with good gastrointestinal function, but this cannot be ensured in sick
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camelids. Electrolyte solutions may be used in place of water if replacement is necessary, but the high prevalence of hyper natremia in sick camelids mandates that these solutions not be used without evidence of need. If the thirst response is inadequate, which is regrettably the case in many sick camelids, oral fluids are best administered via an orogastric tube. Small diameter tubes may also be passed through the nose, but camelids have narrow nasal pas sages and often resist this procedure more than the passing of a tube through the mouth. However, in crias, small-bore feeding tubes may be passed nasally and left in place. Passing an orogastric tube in an adult camelid may induce struggling and a stress response, so it is important to assess whether the patient can tolerate the procedure, to make a plan to perform the procedure quickly and competently, and to abort the procedure, if necessary. Alpacas may be manually restrained, whereas llamas may be better restrained in a llama chute. Light sedation may help, for example, with butorpha nol (0.022 to 0.05 milligrams per kilogram [mg/kg], intramus cularly [IM]), which has fewer deleterious effects on airway protective responses or cardiac output compared with many other sedatives. To prevent damage to the tube because of chewing by adolescents or adults, a piece of PVC pipe or a block of wood with the edges rounded and a hole drilled through the center for the tube should be used as a speculum (Figure 32-1); crias may be tubed without a speculum. When a block speculum is used, it is introduced from the side of the mouth into the interdental space and seated on the molars. The tube is passed through the hole, over the base of the tongue, and into the esophagus. Negative pressure when sucking back on the tube, palpating the tube adjacent to the cervical trachea, or hearing fluid bubbles over the abdomen when blowing on the tube confirm correct placement. The camelid may respond by struggling, regurgitating, or going into respiratory distress. Any of these may lead to reassessment of the adequacy of the restraint or the safety of the procedure and potentially cancelling or delaying it. Without knowing the amount of fiber fill, it is difficult to estimate gastric fluid capacity. Giving a maximum of 3% of
Figure 32-1 A stomach tube and block speculum appropriate for orogastric intubation of an adult llama or alpaca. Adult llamas usually tolerate tubes with an external diameter of up to 5/8” well (top), and 1 2” tubes are commonly used in adult alpacas (bottom).
body weight by volume to an adult alpaca, 4% to an adult llama, and 3.3% to neonates of either species appears to be safe and causes only minimal regurgitation. That amount may be given every few hours, unless gastric distention develops, but to avoid stress and trauma, most patients should be given no more than three tubings per day. The easiest thing to give by tube is water. In dehydrated camelids that have a tendency toward hypernatremia and hyperchloremia, this may be acceptable or even preferable to salt solutions, but if indicated, electrolyte solutions may be used. Except in neonates, oral sugar is usually contraindicated because it is rapidly fermented by gastric microbes. In neo nates, milk or a milk replacer often makes the best oral replacement fluid.
Intravenous Catheterization Because the thirst response is unreliable in many sick came lids, and many require some degree of rapid fluid resuscita tion, the intravenous route is often preferable to others. Much has been made over the years of the difficulties of jugular catheterization in camelids. Although inherently more diffi cult in adult camelids than in many other domestic species because of the thick skin, possible presence of thick fleece, lack of a visible vein, and somewhat different course of the jugular vein, it is a skill that is quickly mastered using proper land marks, restraint, and technique. Mature intact male llamas have the thickest skin, often over 0.5 centimeters (cm) wide. Late-neutered males are similar. Alpacas, females, and crias have increasingly thinner skin, and often have a visible jugular vein. An exception occurs in some family lines of Suri alpacas, in which the skin of all ages may have thick folds (rolling or wrinkled skin). Some form of restraint is usually necessary to hold the neck straight and prevent movement and contamination of the site during the procedure. Neonates may be manually restrained in the standing position or in sternal or lateral recumbency (Figure 32-2). Adult alpacas may be restrained manually, usually in the standing position or in sternal recumbency
Figure 32-2 Restraint of a neonatal cria in lateral recumbency for jugular catheterization. Placement of a towel over the eyes decreases activity.
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Figure 32-3 Manual restraint of an adult alpaca for jugular catheter
Figure 32-4 Making the guide incision with a #15 scalpel blade after
ization. The neck should be held upright with minimal twisting.
injection of a local anesthetic agent. Note the elevation of the skin lower on the neck to facilitate a full-thickness incision without punc turing the vein and the incision made at a similar angle as is planned for the catheter.
(Figure 32-3). Manual restraint of some adult llamas is pos sible as well, and restraint chutes often facilitate the procedure, especially when restrainers are in short supply. If the animal is uncooperative and moving excessively during the procedure, restraint should be modified to compensate for this, rather than pressing forward and engaging in a frustrating and fre quently unsuccessful exercise. Butorphanol may be used as a sedative. Xylazine and similar agents are less desirable because they may decrease cardiac output and slow jugular filling. As with all procedures with sick camelids, if done efficiently, the risk is minimal, but if the camelid starts to display a stress response such as displacement of the soft palate and dyspnea, regurgitation, violent struggling, or arching the head over the back with strabismus, the procedure should be discontinued until the patient has recovered. As a longstanding practice and to avoid damage to the esophagus, the right jugular is more commonly used. If neces sary, the left vein may be used instead. The jugular vein runs dorsolateral to the trachea and ventromedial to the transverse processes of the cervical vertebrae. It is somewhat more ven tromedial than in horses and often approached best from closer to the front of the patient and not the side to avoid accidental puncture of the carotid artery. The vein runs most linearly and most superficially in the cranial half of the neck. In the caudal half, it dives slightly and curves with the neck toward the thorax. In neonates, some alpacas, and the rare llama, the vein may be seen to bulge visibly upon holding it off near the base of the neck. If the vein cannot be visualized by holding it off, often a palpable bulge is still formed and may be found by bouncing a finger on the site while releasing the occluding pressure. If necessary, the neck may be scanned ultrasonographically to find the held-off vein, or if nothing else works, the point midway between the trachea and the ventral extent of the transverse processes of the cervical verte brae approximately one third of the distance from the ramus on the mandible toward the shoulders is a good place to try.
It is desirable to clip the fleece and aseptically prepare the site. Some owners may be reluctant to allow the clipping in show animals, so some conversation on the trade-off between a cosmetic result and an efficient procedure may be required. The site is then infiltrated with a small volume (0.5 milliliter [mL] in adult and 0.25 mL in neonates) of local anesthetic. Using larger volumes in one place may lead to compression of the vein. The skin is lifted and punctured to the full thick ness in the anesthetized area with a #15 scalpel blade or a hypodermic needle with a diameter larger than that of a cath eter (Figure 32-4). Ideally, this puncture should be directly over the vein at a 25-degree angle to the skin surface or shal lower, directed toward the base of the neck, providing a course for the catheter to follow without resistance. If the angle is too steep, the catheter will be deflected or even kink during passage. If the skin is not punctured to full thickness prior to catheterization, often substantial resistance to the catheter tip and inward dimpling of the skin would occur. The extra force required to overcome this resistance may propel the tip of the catheter through the vein without the operator realizing it, causing accidental puncture of the carotid artery. Recognizing the skin drag and repeating or deepening the skin incision is usually wiser than simply ignoring the resistance and proceed ing with catheter placement. A variety of catheters is available and has been used. I prefer polypropylene over-the-needle catheters for most indications because they have little tissue drag and appear to be suffi ciently durable and antithrombogenic for most applications. For most adults, 14- or 16-gauge catheters are ample and 18or 20-gauge are adequate for neonates. For crias or adults that require a catheter for extended periods, a 16-gauge polyure thane over-the-wire catheter may be more appropriate; they are less thrombogenic and also tend to kink or be dislodged
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Figure 32-6 Three catheters placed through guide incisions of differ ent angles. Note the kinks in the two catheters with the steeper angles (top and middle), and the lack of a kink in the catheter with the shal lowest angle (bottom).
Figure 32-5 Introducing the catheter through the guide incision. Note the angle and the catheter being advanced parallel to the course of the vein.
to a lesser degree. The latter are technically more demanding to place and require strict attention to sterility during place ment. In well-hydrated adult camelids, these catheters may be flushed with heparinized saline and introduced as in many other species, but camelids in shock may not have enough jugular pressure to cause the blood flash in the hub upon suc cessful introduction into the jugular lumen. For that reason, although it requires more hands, it is often preferable to attach an extension set and half-filled 20-mL or larger flush syringe to the hub of the catheter during its introduction. An assistant controlling this syringe gently aspirates as soon as the tip of the catheter enters the skin. Especially with larger-bore cath eters, blood should come back easily once the catheter tip enters the vein. This procedure is not always preferable to straight catheterization in neonates, in whom the diameter of the catheter is often too small to appreciate ease of flow and in which the pressure of aspiration may collapse the vein. Additionally, in crias with hypovolemia, blood pressure may be so poor that it is not able to displace a column of heparin ized saline within the catheter. Therefore, in crias, it may be advisable to flush the catheter and then let the flush drain through prior to placement. The placer must then be patient, slowly advancing the catheter and pausing to observe if blood fills the catheter hub. For introduction, the vein should be held off well down the neck. In dehydrated patients, holding off for up to a minute may be necessary to allow sufficient jugular filling. The catheter stylet is held at the hub and directed at a 45- to 25-degree angle downward, following the course of the guide hole and vein (Figure 32-5). Steeper angles are permissible if the stab incision is wide enough to accommodate the subse quent reduction of angle without kinking the catheter (Figure 32-6). The smaller fingers of the hand may be used to steady it against the animal’s neck and maintain the proper angle. If an extension set is attached, the assistant begins to aspirate.
The stylet is slowly advanced until the hub fills with blood or the assistant reports good flow. The tip of the stylet is advanced an additional 0.5 to 1 cm and rechecked for flow. If it is still good, the angle of the hub to the skin is reduced to 20 to 25 degrees, and the stylet is advanced an additional 1 to 2 cm to seat the catheter in the vein. If the flow is still good, the stylet is held immobile, and the catheter is advanced its full length into the vein. A slight twisting motion decreases skin drag and may allow the tip of the catheter to roll off the valves within the jugular. If severe resistance is encountered, it is possible that the catheter is not in the vein and the procedure must be stopped and attempted again. If the catheter seats well, the hub may be secured to the skin with tissue glue or suturing, potentially wrapped, and then used. If catheterization is unsuccessful but the jugular is punc tured, pressure should immediately be released on the lower vein and pressure applied to the puncture site for 2 minutes. If the carotid artery is punctured, the site should be held with firm pressure for a minimum of 5 minutes. In either case, timely recognition of the problem minimizes ensuing compli cations. Significant cervical hemorrhage makes subsequent catheterization attempts difficult both by compressing the jugular vein and by creating a confounding, flashlike effect when the tip of the catheter enters the hematoma. If a poten tial site is occluded by a hematoma, another site should be selected. In all cases, the first catheterization attempt is usually the easiest, so every effort should be made to make the first attempt a successful one. A small number of camelids will have significant acute complications to an unsuccessful jugular catheterization attempt. These likely result from subcutaneous hemorrhage and the pressure it generates on underlying structures. Clinical signs commonly include weakness, lethargy, and recumbency and may also include upper airway obstruction with stridor, nystagmus, and dorsoflexion of the head and neck. Little can be done at the time beyond maintaining the head and neck in an extended position to open the airway, and minimizing other stressors. These complications are best avoided through good catheterization technique. In some cases, use of the jugular vein is impossible or undesirable. In camels, the lateral thoracic veins are often more accessible, but these are difficult to use in smaller
Chapter 32 • Fluid Therapy camelids and even in large llamas. Cephalic vein catheters have sometimes been used, especially in crias; medially, leg veins, mainly the medial saphenous veins, are occasionally accessible in recumbent animals. Ear veins may be used for very-fine-gauge catheters, but they are hard to maintain for very long.
Intravenous Fluids In general, intravenous fluid therapy may be divided into two phases: (1) replacement and (2) maintenance. Replacement in most species involves estimation of the fluid deficit and delivery of that deficit over a short period. In camelids, estima tion of the deficit may be difficult, as many of the disorders lead to dehydration gradually, allowing the animal time to adapt, and the high prevalence of hypoproteinemia increases the risk of edema after bolus fluids. Since camelids appear to be relatively tolerant of dehydration, they are probably also more tolerant of slow replacement of deficits than of bolus correction. Unless dehydration is clear and lack of rapid cor rection has life-threatening implications, initial boluses of more than 2% of body weight to sick camelids are seldom necessary and occasionally detrimental. If dehydration is clear and life threatening, boluses up to 4% of body weight may be indicated, and in extremely rare cases such as in neonates with hyperosmolar disorder and hyperalbuminemia, a bolus of up to 10% of body weight may be necessary, but these are the exceptions. The risk of larger boluses is edema. In hypoproteinemic camelids, edema may rapidly impair pulmonary function, and the camelid may die of respiratory distress. This easily out weighs the risk of prolonged underperfusion in most cases. The major risks of underperfusion relate to other medications: both aminoglycoside antibiotics and nonsteroidal antiinflam matory drugs (NSAIDs) are more toxic to the dehydrated animal and therefore must be used with caution, or not at all, in camelids with questionable circulatory volume and function. Maintenance fluids come with the same warning. Hypo proteinemia and edema are the two biggest risks. Very little information about maintenance requirements in camelids is available: rates from 2% to 10% of body weight per day have been reported in the literature. Except maybe for neonates, 10% is clearly too high, and the risk of edema formation is significant. Two percent may be adequate, but hardly justifies the catheter. Coupled with a conservative replacement phi losophy, administration of maintenance fluids at a slightly higher rate than the bare minimum to correct remaining defi cits appears to be safe and effective. Roughly 2 mL/kg/hr or a volume equal to 5% percent of body weight over 24 hours appears to have little risk, promotes renal perfusion and urina tion such that iatrogenic toxicities are rare, and does not affect blood protein concentrations too much. Blood protein con centration should be monitored at least twice a day during fluid therapy. In some instances, as when the patient cannot be observed frequently or is co-housed with other animals, it may be desir able to give intermittent boluses instead of a constant infu sion. Boluses work well when given every 3 to 4 hours, but additives such as glucose or potassium may be more toxic or less well assimilated. These are relatively minor concerns, so
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the convenience of boluses frequently outweighs these disad vantages in many field settings or when treating a nursing cria. The next question is what to give. Hypernatremia and hyperchloremia are far more common than hyponatremia and hypochloremia, and hypokalemia is common. Acidosis is a rare but an important finding. Alkalosis is more common but is usually mild and benign. Mineral abnormalities are likewise usually mild and benign. Fluids may be adjusted on the basis of individual abnormalities, but the important point to note is that a plain, balanced electrolyte solution is well suited to replacement therapy, and the same fluid with supplemental potassium (and maybe calcium) is suited to maintenance therapy. Because of issues with hyperlactemia and poor glucose assimilation, fluids with acetate as the buffer base may be preferable to those that use lactate. In cases of metabolic alka losis, 0.9% saline is often preferable to pH-balanced fluids. In some cases, additional supplementation is indicated. Many sick camelids have hypokalemia; on the basis of the recognition that measurement of blood potassium is a poor estimator of whole body stores of this principally intracellular cation, supplemental potassium is often added to mainte nance intravascular fluids to bring the final concentration to 20 to 30 milliequivalents per liter (0.5 mEq/kg/hr). At the rate of 2 mL/kg/hr, this provides supplementation well within the margin of safety. Hypocalcemia may be addressed without elaborate calculation by giving 0.5 to 1 mL/kg of 23% calcium gluconate solution by slow injection or by adding this to intravenous fluids. Hypoglycemia may likewise be addressed by bolus administration of 0.5 to 1 mL/kg of 50% dextrose solution, or more gradually by supplementing fluids to a final concentration of 5% dextrose. Up to two thirds of the camelids at our clinic show evi dence of fat mobilization. If this is severe enough to warrant concern, we often use a partial parenteral nutrition formula for our maintenance fluids. Our most common recipe for this is as follows: 5000 mL Acetate-containing electrolyte solution 1000 mL 8.5% amino acid solution 500 mL 50% dextrose 130 mEq Potassium (in potassium chloride [KCl]) 20 mL B vitamins We administer this solution at 5% of body weight/day (2 mL/kg/hr) using infusion pumps. Both B vitamins and amino acid solutions are light sensitive, so the intravenous bags must be covered. We also administer insulin during the course of peripheral parenteral nutrition (PPN) treatment. For most camelids, we use an ultralente form (0.2 to 0.4 units per kilogram [units/kg], subcutaneously [SC], q24 hours). For overtly lipemic camelids or for ones with very high β-hydroxybutyrate or nonesterified fatty acid (NEFA) values, we sometimes use ultralente insulin as often as twice a day (it lasts 12 to 18 hours) or supplement it with intravenous regular insulin at 0.2 units/kg as often as hourly. The rationale for the PPN is as follows: We administer the insulin to arrest fat mobilization and promote triglyceride clearance. We administer the dextrose to prevent hypoglyce mia and also to provide some alternate energy to the patient. We administer the amino acid solution to provide some energy substrate and hopefully spare the protein-scavenging, cachexia effect. The B vitamins might also increase the usabil ity of the glucose. We are able to usually arrest fat mobilization
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BOX 32-1
Examples of Practical Fluid Therapy
CASE #1 A 4-year-old, 50-kg alpaca dam with a high Haemonchus load refuses to stand or eat. Physical examination reveals a heart rate of 96 beats per minute (beats/min), a respiratory rate of 20 breaths per minute (breaths/min), and a rectal temperature of 100.4°F (38°C). Blood analysis reveals the following results: packed cell volume (PCV) = 12%, total plasma protein concentration = 4 g/dL, albumin = 1.5 g/dL, blood urea nitrogen (BUN) = 73 mg/dL, creatinine = 3.6 mg/dL, sodium = 136 mEq/L, and lactate = 2.2 mg/dL. Clear evidence of shock and dehydration suggests the need to do something, most likely by the intravenous route. However, given the low PCV and plasma protein concentration, use of whole blood or a colloid would be ideal. In the meantime, this animal might benefit from conservative bolus treatment (2% of body weight = 1 L) of a pH-balanced, polyionic solution, followed by 100 mL/hr of the same solution for a few hours, until a colloid or blood is available. Large boluses and high rates should be avoided to prevent edema formation. CASE #2 A 2-year-old, 40-kg male alpaca is suspected of having acute C1 acidosis. Physical examination reveals a heart rate of 120 beats/min, a respiratory rate of 30 breaths/ min, and a rectal temperature of T = 104°F (40°C). Blood analysis reveals the following results: PCV = 35%, total plasma protein concentration = 6.7 g/dL, albumin = 4.1 g/dL, BUN = 95 mg/dL, creatinine = 4 mg/dL, sodium = 152 mEq/L, and lactate = 7.4 mg/dL. Evidence of shock and dehydration, including relatively high protein concentrations, is clear. Given these and the likelihood that the dehydration is caused by an acute process, this camelid would be more likely to need and tolerate a larger fluid bolus than average. A 2-L bolus (5% of body weight) would be a good start, followed by 83 mL/hr for 4 to 24 hours, depending on the severity. Fluid should be alkalinizing in this case.
and clear hypertriglyceridemia within 48 hours, using this treatment scheme.
Colloids Because of the high prevalence of hypoproteinemia and the somewhat lower prevalence of anemia in sick camelids, the use of colloids and whole blood as replacement fluids is relatively common. The use of whole blood is discussed in Chapter 36. General indications for the use of colloids are when total plasma protein concentration is less than 4 g/dL or albumin is less than 2 g/dL. Animals with failure of passive transfer or sepsis may also benefit from the antibodies provided in plasma. Colloids may come from natural plasma products or synthetic compounds. Camelid plasma is commercially avail able in many countries and is generally the preferred form of colloid. Transfusion reactions are rare, and plasma from one New World camelid species appears suitable for transfusion to another species. In Australia, dromedary plasma is commer cially available and marketed for use in alpacas. In a 50-kg sick camelid, 1 L of donor plasma usually increases blood protein concentrations by approximately 2 g/dL after the additional water volume is excreted. Plasma transfusions are administered at a slow rate for 15 to 20 minutes through a filtered blood administration set while the animal is observed for evidence of a reaction, includ ing restlessness, straining, hyperthermia, aberrant chewing, and skin reactions. If no reaction is noted, the remainder of the plasma is given over 2 to 4 hours. More rapid administra tion has been associated with respiratory distress. Hetastarch represents an alternative to plasma. It has been used safely in camelids at dose rates of 10 to 20 mL/kg of a 6% solution.1 Hetastarch is not detected by biochemical tests of blood protein but is detected by using a refractometer (Box 32-1).
REFERENCE 1. Carney KR, et al: Evaluation of the effect of hetastarch and lactated Ringer’s solution on plasma colloid osmotic pressure in healthy llamas, J Am Vet Med Assoc 238:768-772, 2011.