Download a PDF version of Dr. Barron’s abstract.
Triage is the process of prioritizing patients based on presenting condition and the resources available, to deal with that condition. Therefore, triage of avian wildlife may necessarily vary to some degree based on individual philosophy, case load, local, state, and federal regulations, and the presenting situation (i.e., oil spill affecting hundreds of animals versus the normal daily load of injury and illness). In any case, the primary objective during the triaging process is the expectation of eventual return of the patient to the wild with 100% function. When triaging birds in a private practice setting, there are some considerations that may help set triage policy. Guidelines about what the practice is willing to accept should be made clear to staff and to the public prior to accepting wildlife cases, including, for example, recognizing when a young animal should be returned to the wild, as is often the case with abducted nestlings and fledglings. It is also important to be aware of reasons why a bird may not be releasable or have a good quality of life in captivity. The goal of wildlife medicine is always eventual release and the reality of euthanasia needs to be kept in mind during triage if chances of release are remote. Realistic resource allocation is especially important in a for-profit clinic that is underwriting wildlife treatment.
Placing intravenous (IV) or intraosseous (IO) catheters is a simple, quick procedure that every clinician should be able to perform in any species of bird. Most patients presented on emergency will warrant venous access. For wild birds, where the patient would be expected to return to eventual flight or hunting performance, IV catheters are usually preferred since IO catheters may cause joint injury/infection. If unable to place an IV, an IO catheter may become a life-saving necessity. These are extremely stable, effective, easily and rapidly placed. It is recommended that peripheral catheters, whether IV or IO, be maintained for no more than 3 days.
In many species of birds, a jugular catheter is often most easily placed using the right (the larger) jugular vein. The basilic vein (wing), also referred to as the cutaneous ulnar vein, and medial metatarsal vein (leg) may also be utilized. The latter is most useful in wading birds, raptors and waterfowl. Pouch veins may be used in some species, such as pelicans.
In birds, IO catheters are generally placed in the distal ulna from the lateral aspect. They may also be placed in the proximal tibiotarsus on either the lateral or medial aspect (try to avoid the patellar ligament). Remember that drugs given IO in the caudal half of the body may go through the renal portal system in birds; thus, this may have some bearing on the choice of catheter placement.
Subcutaneous fluid administration & intramuscular injections
Subcutaneous fluid or drug administration is often a viable choice in avian cases. Particularly in birds, be certain that the needle is subcutaneous, not intramuscular (IM), or in an air sac, lung, or intracoelomic space. Besides visualizing the needle beneath the skin as you begin the administration, you should also notice an immediate bleb. If any resistance is encountered, it is likely that you are against a bone, and it is advisable to retract your needle slightly. Be aware of the patients’ respiration throughout the procedure.
Maintenance volume of crystalloid fluids varies with species and is related to metabolic rate. For birds this may be anywhere from 50-150 ml/kg/day depending on size, species, age, and natural history (i.e. xerophilic vs. aquatic birds).
Again, remember that drugs given IM in the caudal half of the body may go through the renal portal system. Therefore, in many species of birds, the pectoral muscle is a good choice for IM injections.
Fluid support and analgesia are a critical part of early intervention in trauma cases. If the patient is dehydrated or hypothermic, it is important to take a day or so to rehydrate and warm the patient prior to giving any food or medications or metabolism of these substances may be impaired and make the animal worse. Crystalloids, such as Plasmalyte, Normosol-R, LRS or ½ strength LRS + 2.5% dextrose are all acceptable empiric choices prior to obtaining bloodwork. Colloids may also be indicated, especially in cases of hypovolemic shock, common in trauma cases. Hypertonic saline may also be used to address hypovolemia and there has been increasing interest in human and small animal medicine in the use of hypertonic saline in traumatic brain injury (TBI) for its hyperosmotic potential. The author uses a bolus dose of 4 ml/kg IV of 7.2 % NaCl over 10 min in avian species. The increase in intravascular volume after hypertonic saline administration can be transient lasting approximately 15-30 minutes, but these effects can be prolonged by the concurrent administration of colloids. Research suggests, however, that with or without colloids, hypertonic saline-induced reductions in intracranial pressure (ICP) persist for much longer than the vascular effects and significantly longer than mannitol. Hypertonic saline should be avoided in patients with significant sodium derangements or advanced dehydration.
Clinical and research data are gathering which suggests hypertonic saline may reduce ICP faster, more significantly and for longer than mannitol with less undesirable effects. The use of corticosteroids in birds is generally contraindicated, but particularly in cases of TBI. In human medicine the use of glucocorticoids in head-trauma patients has been shown to increase mortality. Corticosteroids will also induce hyperglycemia which has been linked with increased free radical production, excitatory amino acid release, cerebral edema, and altered cerebral vasculature, which could potentiate further neurological injury.
Blood and plasma transfusions
Trauma or disease resulting in hemorrhage is a common emergency presentation in wild birds of any age. Total blood volume in birds is estimated to be approximately 10% of body weight (BW) (i.e. a 1,000 gram bird would have a 100 ml blood volume). Interestingly, birds are less susceptible to shock from blood loss than are mammals. An otherwise healthy bird can lose up to 30% of its blood volume with no apparent ill effects. In one study, the removal of 60% of the blood volume in healthy pigeons did not cause significant clinical affects and resulted in a return to normal PCV by day 7 without treatment. This is due to a bird’s ability to quickly mobilize large numbers of red blood cells from the bone marrow and also probably from an ability to rapidly replace vascular fluid loss from the extravascular space.
Whether to transfuse an avian patient with acute blood loss should be given careful consideration. Pigeons suffering from acute blood loss had a better response to IV fluids than to heterologous or homologous blood transfusions. The same study showed a single iron dextran given IM (10 mg/kg) to pigeons was associated with a significant increase in packed cell volume within 48 hours. In another study in chickens, birds were divided into four groups: untreated controls, and treated with intravenous hetastarch, with a hemoglobin-based oxygen carrier, or by autotransfusion. No significant differences were found in mortality, respiratory rate, heart rate, PCV, or hemoglobin values among the 4 groups at the end of resuscitation. However, in similar a study in mallard ducks, while there was no statistical difference in mortality rate among three fluid resuscitation groups with crystalloids, hetastarch, or hemoglobin-based oxygen-carrying solution (HBOCS), a trend of decreased mortality rate was observed in the HBOCS group.
If the PCV from acute hemorrhage is less than 20% or in more chronic cases less than 15% (normal PCV for most adult birds is in the 35-55% range; keep in mind that normal PCV and RBC counts tend to be lower in young and female birds) and the patient is demonstrating clinical signs of anemia, a whole blood transfusion may be beneficial. Also, if the patient is hypoproteinemic and has an anemia of less than 20%, a whole blood transfusion may be beneficial for the plasma proteins as well as the RBCs. Whole blood and plasma transfusions should ideally be from same species. However, if a homologous donor is not available, a single heterologous transfusion from a donor of the same genus (i.e. Buteo to Buteo) may be of some benefit. The half-life of whole blood heterologous transfusions may range from 12 hours to 4.5 days (depending on whether the donor is within the same genus) whereas a homologous transfusion may have a half-life of 6-11 days.
There is no information on blood grouping in wild birds. In chickens, at least 28 different blood group antigens have been found. A rough cross match (major and minor) can be performed mixing donor and recipient red cells and serum on a glass slide. An absence of gross agglutination or hemolysis suggests compatibility. The recommended anticoagulant is sodium citrate, although acid citrate dextrose (ACD), citrate phosphate dextrose (CPD), or heparin can also be used. The recommended ratio is 0.1 ml of citrate per 0.9 ml of blood or 2 IU of heparin per ml of blood. The amount transfused is 10-20% of the patient’s blood volume (1-2% BW) and can be given IV or IO. This amount would be expected to increase the PCV by 2-5%. Donor blood should be used immediately; there are no reports on the effects of any time of storage for whole blood in raptors. Long term storage of avian blood is not recommended as mammalian storage media appears to be inadequate to support avian erythrocyte metabolism. An 18 μm blood filter should be used. The patient should be evaluated every 15 minutes for the first hour for signs of a transfusion reaction, such as panting due to increased body temperature. Anaphylactic reactions following a single transfusion are rare, but the patient can be premedicated with diphenhydramine. Transfusion reactions have been reported in birds given multiple heterologous transfusions. Supportive care for anemic patients should include B vitamins and iron.
Birds are probably commonly undertreated for pain. In general, if a procedure or injury would be expected to be painful in other species, it is probably a safe assumption that it is equally painful in a bird. Employing balanced, multimodal analgesia is both a medical and ethical imperative. With the opioid epidemic in the U.S., procurement of appropriate analgesic medications is becoming more challenging, forcing avian practitioners to examine viable alternatives. The Drug Enforcement Administration substantially reduced manufacturing quotas for Schedule II opiates and opioid medications for 2018 and this will continue in 2019. Therefore, some flexibility and knowledge of the current options for pain control in birds is warranted.
Euthanasia and assessing life
Full understanding of pertinent federal, state, and local laws and regulations is important when treating wild birds and considering euthanasia. For the most part, wildlife management regulations are delegated to individual states. However, certain federal regulations apply to specific species and specific situations. Veterinarians generally do not need a special permit to possess, stabilize, and euthanize sick or injured birds protected under the Migratory Bird Treaty Act. However, for some species, Federal or State permission may be required prior to euthanasia. Federal migratory bird rehabilitation regulations (50 CFR 21.31) stipulate that any bird that “cannot feed itself, perch upright, or ambulate without inflicting additional injuries to itself where medical and/or rehabilitative care will not reverse such conditions” must be euthanized. Also, “…any bird that is completely blind and any bird that has sustained injuries that would require amputation of a leg, a foot, or a wing at the elbow or above (humero-ulnar joint)…” must be euthanized unless a licensed veterinarian submits a written recommendation that the bird should be kept alive. Additional regulations regarding admission, euthanasia, and documentation apply for migratory birds, and the reader is referred to 50 CFR 21.12 and 50 CFR 21.31 for further details. Other federal Acts may also apply in certain instances, such as the Endangered Species Act (ESA), Marine Mammal Protection Act (MMPA), Bald and Golden Eagle Protection Act, and Lacey Act. Check with the wildlife authorities in your area if you are not sure what the law may entail.
Euthanasia should be conducted without undue patient stress or restraint. Premedication and a multi-modal drug delivery approach to euthanasia should strongly be considered. For example, birds can be premedicated with drugs such as dexmedetomidine and midazolam prior to induction with a gas anesthetic or before administration of sodium pentobarbital IV. If sodium pentobarbital or other barbiturates are used, proper carcass disposal must be practiced, as fatal secondary barbiturate poisonings have been documented.
Euthanasia solution or potassium chloride (KCl) may be given IV, IO, intraperitoneal (IP), intracardiac (IC), or into the supraoccipital sinus. For the last four of these routes or if KCl is used, general anesthesia or heavy sedation is recommended. The dose for both is 1 ml per 10 pounds of body weight. A stethoscope or Doppler can generally be used to verify cessation of heart beat.
About the presenter
Dr. Heather Barron is Director of the Clinic for the Rehabilitation of Wildlife in Sanibel Island, Florida. Dr. Barron was formerly the Chair of Veterinary Clinical Sciences and Professor of Small and Exotic Animal Medicine at St. Matthew’s University School of Veterinary Medicine in the Cayman Islands. Dr. Barron is an associate editor of the Journal of Avian Medicine and Surgery and the author of numerous publications. Dr. Barron is board certified in avian medicine and she served as the 2010-2011 President of the Association of Avian Veterinarians. [MORE]
Dr. Barron also generously shared this video illustrating intravenous catheter placement in an avian patient at the Clinic for the Rehabilitation of Wildlife (C.R.O.W.).
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Gastrointestinal ulcer treatment/prevention protocol
During the live question and answer session, Dr. Barron pledged to share the gastrointestinal ulcer protocol used at the Clinic for the Rehabilitation of Wildlife (C.R.O.W.).
[The Clinic for the Rehabilitation of Wildlife has] successfully used this protocol with thousands of birds. The only caveat is that you cannot feed whole prey items during this time because this protocol makes the gut pH so basic that they will not be able to digest whole prey. They do fine on liquid diets (like the Lafeber critical care diets) or on “soft” foods. Otherwise, discontinue this protocol 24 hours before you intend to feed whole prey.
|Sucralfate||25 mg/kg PO q 8h||For treatment only; not needed for prophylaxis|
|Omeprazole||20 mg/kg PO q 24h|
|Famotidine||0.5-1.0 mg/kg q12-24h|
Epogen use in reptiles
During the question and answer session, an attendee asked about the use of Epogen (epoetin alfa) in birds. Although Dr. Barron has not found it necessary to use Epogen in birds, she promised to share some information on Epogen use in reptiles.
Dr. Barron wrote: Epogen…use in reptiles is still very controversial. There is very little evidence for its use; what is out there is mostly anecdotal. Remember that you can actually cause a non-regenerative anemia by administering Epogen because of the risk of antibody development, and this has been shown in many animal species. Here is…what we know thus far. In one [case report] in tortoises doses of ESA (erythropoiesis stimulating agent) (EPOGEN) at 100 U/kg SC did NOT seem to have any effect. But Mader felt he did see an effect in sea turtles at 100 U/kg SQ and had this proceedings paper:
Mader D. Clinical use of Epogen (epoetin alfa) in green sea turtles (Chelonia mydas) and loggerhead sea turtles. Proc Annu Conf ExoticsCon 2015.
Epogen (epoetin alfa) is a human synthetic erythropoietin analog used to increase red blood cell numbers in anemic humans and cats suffering from chronic renal failure. Its reported clinical application in reptiles is unknown. From 2011-2015, 5 individuals (3 green sea turtles and 2 loggerhead sea turtles) matched the studies inclusion criteria, additionally, 5 matched control cases (3 green sea turtles and 2 loggerhead sea turtles) were identified during the same time period who were only treated with iron dextran. Animals treated with Epogen underwent therapy for anemia an average of 31 days (standard deviation ± 17 days) compared to animals treated with only iron dextran undergoing treatment for an average of 60 days (standard deviation ± 16 days). Epogen is a safe and effective short term therapeutic to accelerate red blood cell production in anemic sea turtles.
And, since I’m also an aquatic animal vet, I read a lot of fish medicine stuff, so I also came across this article some years ago:
Buemi M, Lacquantiti A, Bolignano D, et al. The erythropoietin and regenerative medicine: a lesson from fish. Eur J Clin Invest 39(11):993-999, 2009.
BACKGROUND: Erythropoietin (EPO), the main haematopoietic growth factor for the proliferation and differentiation of erythroid progenitor cells, is also known for its angiogenic and regenerative properties.
MATERIALS AND METHODS: In this study, we aimed to test the regenerative effects of EPO administration in an experimental model of sea bass (Dicentrarchus labrax) subjected to amputation of the caudal fin.
RESULTS: Erythropoietin-treated fishes (3000 UI of human recombinant EPO-alpha immediately after cutting and after 15 days) showed an increased growth rate of their fins compared with those untreated (anova variance: P: 0.01 vs. P: 0.04). By analysing fin length at established times (15 and 30 days after cut), EPO-treated fishes always showed an increased length compared with untreated ones (T-15: 1.1 +/- 0.2 vs. 0.7 +/- 0.2 cm, P: 0.03; T-30: 1.9 +/- 0.3 vs. 1.2 +/- 0.2 cm, P: 0.01). Moreover, exogenous EPO administration induced an enormous increase in EPO-blood levels at each observation time (T-15: 2240 +/- 210 vs. 16.7 +/- 1.8 mU mL(-1), P < 0.001; T-30: 2340 +/- 190 vs. 17.1 +/- 1.9 mU mL(-1), P < 0.001), whereas these levels remained quite unmodified in untreated fishes. Immunochemical analyses performed by confocal laser scanning microscopic observations showed an increased expression of EPO-receptors and PECAM-1 (an endothelial surface marker of vessels sprout) in the regenerating tissue, whereas no signs of inflammation or fibrosis were recognisable.
CONCLUSIONS: All these findings confirm EPO as a new factor involved in regenerative processes, also suggesting a potential, future utility for new therapeutical applications in the field of human regenerative medicine.
But here is the potential downside:
Woods PR, Campbell G, Cowell R. Nonregenerative anaemia associated with administration of recombinant human erythropoietin to a Thoroughbred racehorse. Equine Vet J 29(4):326-328, 1997.
Piercy RJ, Swardson CJ, Hinchcliff KW. Erythroid hypoplasia and anemia following administration of recombinant human erythropoietin to two horses. J Am Vet Med Assoc 212(2):244-247, 1998.
A Standardbred gelding and a colt were examined because of poor performance and anemia. Each horse had been given recombinant human erythropoietin (rhEPO; 4,000 IU) at least twice within the preceding 2 to 4 months. The horses had an Het of 16 and 24%, serum iron concentrations of 210 and 304 micrograms/dl (reference range, 73 to 140 micrograms/dl), total iron binding capacities of 239 and 321 micrograms/dl (reference range, 266 to 364 micrograms/dl), values for the percentage saturation of transferrin by iron of 87.9 and 94% (reference range, 20 to 52%), and serum ferritin concentrations of 255 and 355 ng/ml (reference range, 43 to 261 ng/ml), respectively. There was no clinical or laboratory evidence of immune-mediated hemolysis or an infectious or inflammatory cause of the anemia. Examination of sternebral marrow biopsy specimens revealed generalized bone marrow hypoplasia; myeloid-to-erythroid ratios were 6.7 and 3.2. Moderate-to-marked erythroid hypoplasia was diagnosed in both horses. Compared with serum from a healthy control horse, serum from the affected horses inhibited rhEPO-induced proliferation of erythroid progenitors in vitro. Results suggested that the horses had developed anti-rhEPO antibodies that cross-reacted with endogenous erythropoietin, thereby inhibiting erythropoiesis. Horses were discharged with instructions that rhEPO administration be discontinued and that dexamethasone be administered. Five months later, both horses were back in training. For 1 horse, Hct had increased to 35%, and the other horse was not available for examination.
So, bottom line is that I think there is a lot of risks and probably low potential benefit. It is also very expensive and can be hard to find. But IF you are going to use it, I would recommend using it somewhere in the neighborhood of 100 U/kg. There may be less risk of antibody formation if you use darbopoietin rather than Epogen or Procrit, but this has not been investigated in reptiles. I would ALWAYS give iron dextran first. Good luck.
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