Avian Laboratory Diagnostics

Natalie Antinoff, DVM, ABVP (Avian)
Gulf Coast Veterinary Specialists
Gulf Coast Avian & Exotics
Houston, TX

Of any species encountered in companion pet medicine, birds may be the most alien, and therefore most intimidating to practitioners. While it may be possible to adapt the knowledge of one mammal to treat another, the unique anatomy and physiology of birds makes even safe restraint a challenge. Although interpretation of a hemogram and biochemistry profile is learned as a veterinary student, there are many differences in avian laboratory evaluation. Collection techniques vary; all cells have a different appearance than mammals, and different biochemistry values may be more (or less) significant than those of mammals. Even the choice of commercial laboratory can affect the interpretation of laboratory results.

Restraint/examination
In emergency situations, patient assessment must be performed efficiently and effectively, in a manner that is safe to the patient. Excessive handling, even when appropriate, may cause patient stress, resulting in increased catecholamine release and subsequent cardiac compromise. It may be necessary to perform a portion of the physical examination, then return the patient to its cage for a few minutes before completing the exam.

Removing the patient from the cage can be a challenge. Be sure that the room is escape-proof and windows are closed and covered. The weapon is the beak. Use a towel covering your hand to introduce your hand into the bird’s cage. Then cover the head (and body if possible) with the towel and position your hands around the mandible. Wrap the rest of the towel gently around the body. In most cases, avoid the use of leather or heavy gloves, as they will not only interfere with your ability to perform a thorough physical exam, but may also be extremely frightening and distressing to the bird. Turning off the lights may help calm a frantic bird so that it can be more easily caught. It is also helpful if the bird does escape from the cage. It is generally not recommended to remove a bird from its owners’ shoulder or hand; place the bird on a table, floor, or in a cage before restraint.

Proper restraint:
Psittacines: One hand below the mandibles, gently extending the neck to maintain control of the head and beak. The other hand grasps the feet. Depending on the size of the bird, the holder can “cup” the bird in the palm of the hand to stabilize the wings, or hold the wings against the body. Use of a towel can improve control. Do not restrict the movement of the keel, or restrain across the pectoral region, as this must expand for effective respiration. Maintain the patient in an upright position for evaluation.
Alternate technique: Thumb and second finger on either side of the mandibles, gently holding the mandibles behind the beak. The index finger can be placed on top of the head for greater control. Bruising of the face may occur; do not use excessive force. This technique can be more difficult for people with small hands, especially in larger birds.

Passerines/softbills: The head is gently restrained between the index finger and the second finger, cupping the bird in the palm of the hand for support.
Pigeons/doves: Grasp the wings and tail in one hand over the dorsal tail base, wrapping the hand around the body. The head does not require restraint unless a procedure is performed in this area.

Raptors/Birds of prey: Weapons in these birds are primarily the feet, secondarily the beak. For these birds the use of leather gloves is appropriate. Covering the head of the bird with a towel or falconer’s hood generally calms these birds, and the cover should remain over the head for as much of the exam as possible to minimize stress. Once the head is covered, grasp the feet (wearing leather gloves) from the top, and restrain the feet while removing the bird from its cage or enclosure. In most birds, only light restraint of the head is necessary if the head is kept covered. If prolonged handling is required, the feet can be temporarily taped into a ball bandage.

Sample Collection
Total blood volume in a bird is approximately 10% of body weight, and the amount of blood that can be safely collected is 10% of that amount. The maximum blood volume collected should be 1/100 of the body weight. In any sick patient, half that amount is recommended, to allow for anemia, illness, and hematoma formation. Example: in a 380g African grey, 3.8cc could be safely collected; 1.9cc would be half that. It is almost never necessary to collect more than 1cc from a bird unless collecting for blood transfusion. In a 100g cockatiel, 1cc could be taken, and in illness, 0.5cc. This should be enough blood for a full CBC and biochemical profile. However, in a 22g Budgerigar, 0.22cc is the maximum that can be safely collected, and may only provide enough blood for a few tests. In this situation, indicate your testing priority to the laboratory to maximize the value of the small sample. If necessary, more blood can be collected 48-72 hours later.

As in mammals, blood values are most meaningful when collected prior to any treatment. However, many birds are highly compromised from illness, and it may be necessary to treat and stabilize the patient for 24-48 hours prior to blood collection. (A diagnostic test is never worth a patient’s life!)Blood samples should be collected directly from a blood vessel. The (right) jugular vein is easily accessed and provides for rapid collection, usually with minimal patient stress. Other sites for venipuncture are the wing vein (brachial, basilic, or cutaneous ulnar, depending on where it is accessed); and the medial metatarsal vein (leg). Both of these sites are more prone to bruising, and since they are smaller vessels, may take longer for collection. The toenail should not be used as a site for blood collection; samples from this site are prone to artifacts, and contamination with urates in the cage can lead to inaccuracies in uric acid levels. In addition, this is painful to the patient for several days. The one instance in which a toenail clip may be preferable is to collect a PCV in cases where severe anemia or coagulopathies are present or suspected.

Samples should be collected with a small needle and syringe. Insulin syringes with 29-ga needles are preferable for budgies and canaries; TB syringes with 27-or 25-ga needles for cockatiel/conure size; and a 3-cc syringe with 25-ga needle for larger birds. *Remove the needle before putting the blood into the tube to prevent hemolysis. Heparinizing the syringe is NOT recommended, as heparin causes artifacts in the CBC. Rather, capillary tubes and fresh slides without anticoagulant are preferable for the CBC to minimize artifact. Many laboratories prefer a coverslip technique to decrease smudge cells (cellular destruction, common in leukocytes). Heparinized microtainer tubes are preferred for biochemistries, and are available with or without plasma separators. (Beckton-Dickenson, Cockeysville, MD; Monoject, St. Louis, MO).

Complete Blood Count
The PCV is performed as in mammalian species, and is measured after centrifugation in a capillary tube. The HCT represents a calculated value. False decreases in PCV may occur in aged samples. Automated cell counters cannot be used in avian hematology because of the nucleated red blood cells.

The leukocyte count can be determined by a hemocytometer using either the Natt and Herrick or Eosinophil Unopette method. For the direct, or Natt and Herrick, a 1:200 dilution of methylene violet 2B is added to the hematocytometer; this stain is taken up by all blood cells. After 5 minutes (to allow cells to settle), all erythrocytes are counted in the 5 large squares, and multiplied by 10,000 to obtain the result of RBC’s per ml. The leukocytes are counted in all 9 squares; the raw count is added to 10% of the raw count, and the result is multiplied by 200 to determine leukocytes in microliters. Using the indirect, or Eosinophil Unopette method, blood is mixed with phloxine B and the mixture is used in the hemocytometer. The heterophils and eosinophils take up stain, and all cells in the 18 large squares are counted. The total heterophil/eosinophil count obtained is multiplied by 10 x 32/18. The total leukocyte count combines the heterophil/eosinophil count with the leukocyte differential from the slide, as follows: heterophils and eosinophils counted / (%heterophils and %eosinophils) x 1003. Alternately, and perhaps more commonly used, the leukocytes are estimated from slides, but well-prepared slides with a consistent monolayer is essential for interpretation. All leukocytes are counted in 10 high-dry (40x) power fields. The number obtained is divided by the number of high power fields (10), and the result is multiplied by 2000 to obtain the estimated leukocyte count. Anemic patients require the following correction: (est. leukocyte count X actual PCV)/normal PCV.
In addition to the manual methods, laser flow cytometry can be used to count leukocytes from avian blood, although thrombocytes may be counted as leukocytes which leads to inaccuracy.

Identification of cells:
Erythrocytes are nucleated, elliptical cells with pink cytoplasm and basophilic nuclei. They are larger than mammalian RBC’s (130-180 femtoliters). Younger erythrocytes are commonly present in the smear, and are recognizable by their slightly more basophilic cytoplasm. Reticulocytes can be detected with new methylene blue staining. Slight polychromasia (1+) is considered normal, as is slight anisocytosis.
Anemia is associated with pathologic conditions similar to those of mammals; i.e. blood loss, (trauma, parasitism, coagulopathy) hemolysis/destruction (toxin), lack of production (leukemia, nutrition), or chronic disease (Aspergillus, Chlamydia). Polycythemia is most commonly due to dehydration, but can occur in macaws and is often associated with respiratory disease.

Leukocytes:
Heterophils are the avian neutrophil. These cells are round, with rod-shaped pink (or orangish) granules and bluish cytoplasm. Bands are uncommon but are also difficult to identify. Lack of nuclear segmentation and lack of folding of the nucleus are the features of bands. Heterophilia indicates inflammation or infection, stress, and occasionally neoplasia. Mild heterophilia is common in juvenile captive birds as their immune system develops, and in captured wild birds as a response t stress. Heteropenia is uncommon clinically, but is a common artifact from smudge cells. In true heteropenia, there is usually a leukocytopenia, and the most common cause is viral disease or sepsis.
Eosinophils are often confused with neutrophils. They have rounder granules which may stain pink or bluish, with more uniform cytoplasmic color and more distinct nuclear/cytoplasmic borders. These are of unknown clinical significance, and attempts to correlate with parasitism or allergic response have been inconsistent.
Lymphocytes are similar in appearance to mammalian lymphocytes. They are smaller round cells with basophilic round nuclei, high N:C ratio, and a small amount of basophilic cytoplasm. These cells can be variable in size, and are sometimes confused with monocytes. Lymphocytosis is most commonly due to laboratory error (smudged heterophils), but can occur in lymphoid neoplasia. Lymphocytopenia is often associated with stress. Some species (Amazons, canaries) are normally lymphocytic.
Basophils are smaller than heterophils, with colorless cytoplasm and many dark basophilic granules. The clinical significance of basophils is unknown, but counts can increase with respiratory infections, tissue damage, and active Chlamydial infections (Amazons, budgies). Absence of basophils is normal.
Monocytes are large leukocytes with abundant cytoplasm that usually stains darker than lymphocytes. The cytoplasm may have a grainier, blue-gray appearance, often with vacuoles. The shape of both the nucleus and cytoplasm can vary, and the nuclear chromatin is less dense or clumped than the lymphocyte. Monocytosis is associated with chronic infection/inflammation, most often associated with Chlamydia infection, Aspergillus or other fungal infection, or mycobacterium. The absence of monocytes in a peripheral blood sample is considered normal.

Thrombocytes: Thrombocytes are small oval nucleated cells with colorless cytoplasm and pyknotic nuclei. Small numbers of red granules and small clear vacuoles may be present in the cytoplasm. Clumping is common. Thrombocytopenia can occur with viral disease or potentially excessive bleeding and consumption. Thrombocytosis has not been documented in birds.


Biochemical Profile

Plasma protein: In avian samples, the Biuret method is more reliable for determination of plasma protein than a refractometer. At levels below 3.5 g/dL, refractometer readings are less accurate. Protein levels will be higher in plasma than serum because plasma contains fibrinogen. Hyperproteinemia is attributed to dehydration or chronic infection, particularly due to the excess production of gamma globulins. Hypoproteinemia is due to malnutrition, decreased synthesis (hepatic disease), malabsorption (gastrointestinal disease), or increased loss (renal disease).

Hepatic Enzymes
Aspartate Aminotransferase (AST, SGOT): Nonspecific indicator of hepatocellular damage.. Released from liver, muscle, heart, kidney, and brain when damage to any of these organs is present. Most elevations are from liver and muscle. CPK can be used to aid in differentiating muscle breakdown. AST provides no information about liver function.

Lactate Dehydrogenase (LDH): Nonspecific indicator of acute hepatocellular damage only. liver, skeletal muscle, cardiac muscle, kidney, bone, and erythrocyte. Elevations most common with liver disease, but very short half life. May not be elevated in ongoing or chronic liver disease. False elevations common with hemolysis.

Gamma Glutamyl Transferase (GGT): Attempts to correlate elevations with hepatic disease have been inconsistent. May be an indicator of hepatocellular or bile duct carcinoma. Limited usefulness.

Creatine Kinase (CK): Skeletal muscle, cardiac muscle, brain. Used in conjunction with AST to determine if muscle breakdown is occurring. (Example: elevated AST with normal CK supports hepatic etiology; elevated AST and CK provides no information about the liver. )

Bile acids: The only laboratory assay that is an indicator of liver function. Conjugated bile salts are normally secreted into the bile, 90% of which are reabsorbed in the jejunum and ileum. They are then taken up by the liver via enterohepatic circulation. The small portion of bile acids that is not removed by the liver is measured in blood. In diseased states, bile acid absorption from the blood is impaired, creating measurable elevation. Bile acids do not significantly elevate postprandially in birds, probably because of the normal retrograde movement of food from the intestinal tract into the ventriculus, creating difficulty in obtaining a true fasting value. Single bile acids are considered valuable, independent of fasting or food consumption.


Alanine Aminotransferase, Blood Urea Nitrogen, Alkaline Phosphatase and Total bilirubin are not produced in significant amounts from the avian liver and are not useful in assessment of avian hepatic disease.

Renal Analytes:
Uric acid: Synthesized in the liver and renal tubules, 90% excreted by secretion into the tubules. Reliable assessment of renal disease. Minimally affected by glomerular filtration; therefore uric acid levels will not increase markedly with dehydration alone. Normal uric acid does not eliminate the possibility of renal disease, because of the ability of the kidneys to compensate.

Blood Urea Nitrogen and Creatinine are not reliable indicators of renal disease.

Nutrients and Metabolites:
Glucose: Glucose abnormalities are rare. Criteria for diagnosis of diabetes in birds are repeatable glucose levels >800-1000mg/dl (depending on reference). Avian red blood cells do not utilize glucose as mammalian cells do, so prolonged contact is less likely to affect results.

Calcium: Interpret in conjunction with albumen, although mammalian conversion formulas are not accurate. Normal physiologic elevations occur as a response to estrogen during periods of reproductive activity. Decreases are often nutritional, and may be related to lack of vitamin D. Measurement of sample in EDTA can falsely lower calcium.

Phosphorous: Unreliable in many cases. May elevate in severe renal disease, or renal secondary hyperparathyroidism; may be decreased from hypovitaminosis D or malabsorption. Most common reason for elevation is artifact from hemolysis or prolonged contact with RBC’s.

Amylase: Marked elevations may be present in cases of pancreatitis (>1000 U/l), and is commonly associated with zinc toxicity.

Cholesterol: May vary with diet. May elevate with hepatic disease or bile duct obstruction, and in birds with xanthomas. May decrease with liver disease.

Triglycerides: Elevations are predominantly associated with egg-related peritonitis and reproductive disorders.

Sodium, Potassium, Chloride: Elevations are often caused by laboratory error.

Protein Electrophoresis: Separates plasma proteins into 5 main protein fractions: albumen, alpha-1, alpha-2, beta, and gamma globulins. Alpha globulins increase with acute-phase inflammation. Beta globulins mainly are IgM, IgA, and fibrinogen, also predominantly acute-phase proteins. Gamma globulins are immunoglobulins, IgA, IgM, IgE, and IgG). Abnormalities can indicate inflammation (elevated globulins), and patterns can be used to suggest or support certain diseases; decreased albumen can also be helpful in the assessment of hepatic or renal disease. The electrophoretogram can also be useful for monitoring response to disease.


Urinalysis

Indicated when renal disease is suspected. All samples will have bacterial contamination from the cloaca (sterile samples can collected by catheterization of the ureter). Normal specific gravity is 1.005-1.020. Normal pH is 6.0-8.0. Trace protein is normal in birds. The urinary threshold for glucosuria is 600mg/dL in most species. The most valuable assay in urinalysis is evaluation of the sediment. RBC’s and WBC’s should be 0-3/HPF in normal urine, and no epithelial cells should be present. The presence of casts is abnormal.

Miscellaneous:
Blood lead level: measured from unseparated blood or from red blood cells. Do not use EDTA.

Blood zinc level: measured from plasma. Contamination from rubber stoppers can cause false elevations.

Fecal Analysis:
Direct analysis: Perform direct fecal analysis to evaluate for parasites, including giardia. Megabacteria can often be seen using this technique with a darkened field.
Gram’s stain: evaluate presence of bacteria and differentiate Gram negative from Gram positive. Quantitate amounts of each cell population. Evaluate for the presence of yeast. Megabacteria can often be seen, and are much larger than the normal bacteria, with small basophilic granules. Occasionally, giardia can be identified on Gram stain in severe cases.
Trichrome or IFA stains can be used to confirm the presence of giardia. Iodine can be used in-house to identify cysts, but will interfere with flagellate motility.

Clinical Techniques

Fluid Administration /catheterization
· Maintenance fluid rate: 50 ml/kg/24 hrs; deficits = % dehydration x body weight + ongoing losses.
· LRS, Plasma-Lyte, Normal saline, D5W; Half-strength saline. Can be given as 3-4 IV boluses through a 24-hour period. Stable, alert patients often will not tolerate IV catheters; subcutaneous fluids may be preferable in these situations if other parameters do not mandate IV fluid replacement.
· Intravenous
· Jugular vein – right side only; left usually vestigial
· Note: pigeons have a venous plexus; jugular catheterization will not be as feasible in these birds.
· Cutaneous ulnar vein (Brachial, basilic, wing vein) – ventral aspect of the wing, running lateral to the humerus. Best visualization at the point where this vein crosses the radius and ulna at the elbow. Place catheter slightly proximal to this point to avoid the bones. Vein is very superficial. Note: can cause excessive bruising.
· Medial metatarsal vein – medial tibiotarsus extending across the medial aspect of the tarsometatarsal bone. Best to access near the junction of scaled and smooth skin of the foot. May bruise.
· Intraosseous
· Administer fluids slowly, as there is greater pressure and increased patient discomfort with intraosseous catheter use in comparison to intravenous fluid administration.
· If a spinal needle of appropriate size is not available, use a standard hypodermic needle (22-25 ga). It is common for a bone plug to occlude the needle. If this occurs, withdraw the original needle, and replace it with a new sterile needle. The bone plug is removed with the original needle, and it is usually quite simple to locate the site of entry of the original needle.
· Proximal tibiotarsus:
· With the stifle flexed, palpate the cranial tibial (cnemial) crest distal to the femoral condyles. Prep the area with a surgical prep. Use a spinal needle of appropriate size (generally 20-22 ga), and introduce it along the tibial crest into the marrow cavity. Remove the stylet and aspirate; if blood appears in the hub, flush and tape into place.
· Flush; sometimes the medial metatarsal vein will blanch as the fluid is flushed.
· Distal ulna:
· The dorsal condyle of the distal ulna is located on the dorsal aspect of the carpus. Flex the carpus and palpate dorsally just proximal to the angle of the carpus. The ulna is the larger, more ventral bone. Prep the area surgically and insert a spinal or hypodermic needle of appropriate size under the dorsal condyle into the marrow space while holding the ulna stable.
· Flush; the cutaneous ulnar vein will blanch with flushing if the needle is correctly positioned.
· Subcutaneous
· Lateral flank – Extend the leg caudally; identify the skin fold in the craniolateral aspect of the flank. Insert needle just underneath the skin; visualize the fluid “bubble” as you administer fluids. Beneath the skin is a thin muscle layer; deep to that is air sac. Use caution to avoid advancing the needle into the air sac, as fluids administered in this fashion may cause “drowning”.
· Medial thigh – Extend the leg laterally; administer fluids in the skin fold under the thigh. There is less potential space in this area; may cause patient discomfort until fluids are absorbed.
· Dorsal scapular – Between the shoulder blades, similar to mammals. Observe the fluid bubble underneath the skin as you administer. Beneath the skin lie the cervical and sometimes the cranial thoracic air sacs; use caution to avoid administering fluids into the air sac.
· Patagium (wing web) ventral wing web near axilla; use caution to avoid the cutaneous ulnar (wing) vein. May cause patient discomfort.

Gavage feeding
· Volume to administer: 1cc per 30g patient weight (adult birds; juvenile crops are much more expansive.)
· Using your right hand, facing the patient, advance the feeding tube from the bird’s left oral cavity into the esophagus on the right. The crop is located just above the keel; advance the tube into the crop.
· Palpate the feeding tube and the trachea to assure that they are 2 separate, distinct structures. Once this is evident, the food can be bolused, but always observe the back of the mouth for regurgitation. Keep the bird upright for feeding.
· This should be performed at the end of treatments; once gavage fed, place the bird immediately back in its cage to minimize the risk of regurgitation.
· If patient regurgitates, DO NOT turn upside down! Place patient down immediately, on a table or floor or in cage, and let the bird clear the regurgitated food without intervention.
· Feeding tubes: ball-tipped metal feeding tubes (do not force, can cause trauma); appropriately sized red rubber tubes (large; use mouth speculum).

Intubation
Tracheal – open the beak (use gauze strips if necessary in psittacines). The glottis is easily visualized at the base of the tongue. If it is not clearly accessible, placing a thumb under the mandible to elevate the tongue can assist in visualization. Do not inflate the cuff; as birds have complete tracheal rings.


Vascular Access and Nutritional Suggestions for Birds
Venipuncture sites: Jugular (right only), Cutaneous ulnar and Medial metatarsal
Intravenous Catheter Placement: Jugular (right only), Cutaneous ulnar and Medial metatarsal
Intraosseous catheter placement: Distal ulna, Proximal tibiotarsus
Fluid Volume/choice: LRS, PlasmaLyte, Normosol, 0.9% NaCl, Hetastarch, Oxyglobin, Whole blood (same species). 50ml/kg/day maintenance, Supplement with 5% dextrose if necessary.
Nutritional Suggestions: Avian Critical Care (Lafeber), Emeraid (Lafeber), Ensure, Deliver, Hand-feeding formula.
Comments: Gavage feeding preferred: 1cc per 30g body weight.





References

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Natalie Antinoff, D.V.M., Diplomate ABVP (Avian)

Natalie Antinoff obtained her DVM from the University of Wisconsin School of Veterinary Medicine. She then completed a 1-year postgraduate internship in small animal medicine and surgery at the Animal Medical Center in New York. After practicing in a high-volume small animal and exotics practice in New York, Dr. Antinoff returned to the Animal Medical Center in New York for a residency in Avian and Exotic Pet Medicine and Surgery, one of only 3 in the country at that time. She is currently board certified by the American Board of Veterinary Practitioners in the Avian Practice Specialty. In January 1997, Natalie established Gulf Coast Avian & Exotics as a part of Gulf Coast Veterinary Specialists. This is the first specialty exotics practice operating as part of a referral center in the country.

Dr. Antinoff has authored chapters and articles in several veterinary texts and refereed veterinary journals. She also has spoken locally, nationally, and internationally to pet owners and veterinary associations while serving as a mentor for others preparing for Avian Board Certification. Dr. Antinoff was named the Texas Veterinary Medical Association Non-traditional Species Practitioner of the Year in 2003 and has been a T.J. Lafeber Avian Practitioner Award Finalist in both 2003 and 2004. She is currently authoring a text for practicing veterinarians for a British Publishing Company.
Dr Antinoff is a member of the American Veterinary Medical Association, the American Board of Veterinary Practitioners, the Association of Avian Veterinarians, the Association of Reptile and Amphibian Veterinarians, the Texas Veterinary Medical Association and the Harris County Veterinary Medical Association.