National Wildlife Health Center

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Collection of Blood Samples From Adult Amphibians


Standard Operating Procedure
ARMI SOP No. 101
Revised 12 Feb 2001

  1. PURPOSE: Provide guidelines for safe collection of blood samples from live adult amphibians.

  2. SCOPE: These guidelines apply to adult or post-metamorphic frogs, toads and salamanders, and neotenes. The described techniques may be used in the field or in a laboratory. These techniques are not recommended for amphibian larvae.

  3. EQUIPMENT and SUPPLIES.
    1. Anesthetized amphibian
    2. Syringes, 1 cc with 26, 27 or 28 gauge needle
    3. Glass slides (microscope slides, 25 X 75 mm), frosted end
    4. Microscope slide box
    5. Capillary tubes, plain (NOT heparinized)
    6. Sealing wax (eg, CritosealŽ)
    7. Blood tubes to hold blood-filled capillary tubes
    8. Pencil (to label glass slides) and felt-tip pen to label blood tubes
    9. Sharps safety disposal (for used syringes and broken glass slides and capillary tubes)
    10. Scales: electronic or spring

  4. PREPARATION:
    1. Calculation of amount of blood that can be removed safely.
      In most vertebrates, blood is 10% of the body mass. In ectotherms, about 50% of the blood can be removed at one time. This is roughly 5% of the body mass. Hence, if a toad weighs 30 grams, its blood volume is 3 gm [1 gram - 1 ml] (10% of 30g = 3 gm or 3 ml); half this blood volume is 1.5 ml. For the ARMI Health Surveys, a minimum of 0.2 ml and a maximum of 0.5 ml of whole blood are requested. The smallest amphibian from which 0.2 ml whole blood can be removed is 4 grams; the smallest amphibian from which 0.5 ml whole blood can be removed weighs 10 grams.
    2. Arrangement of Blood Collection Supplies. While the amphibian is being anesthetized in benzocaine or MS222, the blood collection and blood storage supplies should be arranged for immediate use. These supplies should all be laid out and easily accessable, because blood in the syringe will clot within 30-90 seconds, and clotted blood cannot be used for making blood smear slides and cannot be put into capillary tubes.
      1. Syringe and needle
      2. Slides, 2 frosted-end clean slides
      3. Capillary tubes: 2-5 tubes
      4. Sealing wax for capillary tubes
      5. Blood tubes to hold (store and mail) capillary tubes
      6. Tape and indelible ink pen to label slides and blood tubes
      7. Alternative: for large amphibians if >0.4 ml of blood are expected, a plain blood tube may substitute for capillary tubes.

  5. METHOD
    1. Anurans : Collection of blood from the heart
      1. Weigh amphibian and calculate maximum volume of blood that may be removed (see paragraph IV-A above).
      2. Positioning the amphibian. Place sedated or anesthetized amphibian on its back on a flat surface (see NWHC ACUC Protocol 2001-006, Anesthesia of Amphibians in the Field).
      3. Locate xiphoid. In the ventral chest area, locate the xiphoid cartilage which is the last portion of the sternum adjacent to the abdomen; this cartilage is in the ventral midline and with careful examination, the beating heart can be seen moving the skin on one or both sides of the xiphoid. The heart is located under the center of the sternum in the midline, but the needle will be inserted through the skin and muscles at the edge of the xiphoid. The needle is NOT inserted through the sternum nor through the xiphoid.
      4. Cleaning the skin. The skin should be gently rinsed with a stream of clean water from a squirt-bottle. Optionally, the site may then be sprayed with a short burst of BactineŽ (disinfectant). Do not use any other disinfectant (eg, alcohol, other sprays, iodine-based disinfectants, bleach, etc) on the skin of amphibians. For more explanation of disinfectants see also NWHC ACUC Protocol 2001-004, Toe-Clipping of Frogs and Toads.
      5. Syringe. Remove the syringe from its paper wrapping. Move the plunger back and forth about 2-4 cm in the syringe case, then push in the plunger completely.
      6. Remove the safety sheath from the needle and position the needle on the frog's skin in the midline at the tip (free edge) of the xiphoid. The syringe is held at a very acute angle, such that the needle and syringe are almost laying on the belly skin; the needle should be pointing towards the frog's snout at an imaginary point between the eyes.
      7. Insertion of needle into heart. The needle and syringe are pushed through the skin gently, slowly and steadily. When the needle has advanced about 2 mm through the skin, the plunger is gently pulled back about 3-5 mm and then the needle and syringe are advanced further towards the heart.
      8. Drawing blood. While keeping a steady but gentle vacuum ("suction") in the syringe by gently pulling back on the plunger, the needle and syringe are slowly advanced towards the heart. When the needle enters the heart, a small amount of blood will appear in the syringe at the hub. Immediately stop advancing the syringe and needle and allow blood to continue to flow into the syringe. It is not necessary to pull the plunger more than 5 mm in the syringe case to create sufficient vacuum to draw blood. If blood stops flowing, the syringe may be moved 0.5 to 1 mm back and forth to resume blood flow into the syringe. Sometimes it also is helpful to simply spin the syringe a quarter or half turn to reposition the bevel at the tip of the needle. NOTE: blood flow into the syringe will be very slow because of the small diameter of the needle and because of lower blood pressure in amphibians (compared to mammals and birds).
      9. Withdrawing needle. When 0.2 to 0.5 ml of blood are obtained, the plunger is released before pulling out the needle from the frog's body (this avoids aspirating air into the syringe which may greatly accelerate clotting of the blood).
      10. Slides and capillary tubes. Two blood smear slides are immediately prepared with blood in the syringe (see NWHC ACUC Protocol 2001-003). Remaining blood in the syringe is then very gently injected into capillary tubes. Each capillary tube is filled only two-thirds (~65%) and then sealed at both ends with sealing wax. Do not completely fill capillary tubes.
      11. Wound treatment. Treatment of the needle hole in the skin is optional. A brief spray of BactineŽ onto the hole in the skin is recommended.
      12. Tip on holding the syringe: The most efficient method of holding the syringe is to grasp it between the first two fingers and the thumb. The small finger can then be used to move the plunger. The other hand is then free to steady the frog or hold the forelimbs out of the way.
      13. Failures to obtain blood. If blood initially enters the syringe and then stops, gently rotate the syringe 90-180 degrees. If no blood is obtained on the first insertion of the needle, almost to the point that the needle leaves the skin, then gently pushed the needle towards the heart a second time. If the second insertion of the needle into the area of the heart fails, then cease all further attempts. NOTE: do not repeatedly plunge the needle and syringe back and forth into the chest; the limit on insertions of the needle into the area of the heart is twice.
      14. Large amphibians. If blood is to be collected from the heart of large amphibians (>100 g) such as adult bullfrogs, pig frogs, or cane toads, then a longer needle probably will be needed, and a larger diameter needle (eg., 23 gauge) may be used.

    2. Caudates (salamanders): Collection of blood from the tail.
      1. Weigh animal. For those salamanders that weight >10 g, blood can be obtained from the ventral tail vein. For salamanders weighing 2-10 g, blood may be collected from the heart.
      2. Calculate maximum volume of blood that may be removed (see paragraph IV-A above).
      3. Positioning the amphibian. Place anesthetized amphibian on its back on a flat surface (see NWHC ACUC Protocol 2001-006, Anesthesia of Amphibians in the Field).
      4. Location of ventral tail vein. The ventral tail vein is located in the ventral midline of the tail immediately under (ventral to) the vertebrae. This vein cannot be seen in live salamanders.
      5. Insertion site. The needle and syringe will be inserted into the vein just posterior (caudal) to the vent opening. If the vent opening is roughly 5 mm long, then the site for insertion of the needle should be 5 to 10 mm from the caudal edge of the vent opening; if the vent opening is 12 mm long, then the site of insertion should be 12 to 24 mm from the edge of the vent opening.
      6. Cleaning the skin. The skin should be gently rinsed with a stream of clean water from a squirt-bottle. Optionally, the site may then be sprayed with a short burst of BactineŽ (disinfectant). Do not use any other disinfectant (eg, alcohol, other cans of aerosol sprays, iodine-based disinfectants, bleach, etc) on the skin of amphibians. For more explanation of disinfectants see also NWHC ACUC Protocol 2001-004, Toe-Clipping of Frogs and Toads.
      7. Syringe. Remove the syringe from its paper wrapping. Move the plunger back and forth about 2-4 cm in the syringe case, then completely push in the plunger.
      8. Insertion of needle into vein. Remove the safety sheath from the needle and position the needle on the salamander's tail in the midline caudal to vent. The syringe is held at approximatedly 30 to 45E angle to the body with the needle pointed towards the head. The needle is advanced through the skin at 30 to 45E angle towards the vertebra, keeping the needle in the midline. The needle is advanced until it contacts the bone of the tail vertebra (coccygeal vertebra) and then the plunger of the syringe is pulled. If no blood appears in the syringe, then pull the syringe back very slightly from the bone while keeping a vacuum in the syringe (by gently pulling back on the plunger about 5 mm). If no blood appears in the syringe, gently rotate the needle and syringe 90-180 degrees and pull the plunger.
      9. Second attempt. If blood is not obtained from the ventral tail vein on the first attempt, and after the syringe has been moved back and forth slightly a maximum of 2 times to locate the vein, then removed the syringe, select a fresh syringe, and attempt to collect blood about 5 mm posterior to the first site (closer to the tail tip, NOT closer to the vent opening).
      10. Withdrawal of needle from tail. When 0.2 to 0.5 ml (or more) of blood are obtained, the plunger is released before pulling out the needle from the tail (this avoids aspirating air into the syringe which may greatly accelerate clotting of the blood).
      11. Blood smear slides and Capillary Tubes. Immediately place a small drop of blood on two slides and make the blood smears. Next, insert the needle into the end of each capillary tube and inject blood into each tube until the tube is about two-thirds full. Seal both ends of each capillary tube with sealing wax and put the capillary tubes into a standard blood tube; label the blood tube.
      12. Wound treatment. Treatment of the needle hole in the skin is optional. A brief spray of BactineŽ onto the hole in the skin is recommended.

    3. Caudates: Collection of blood from the heart of small salamanders
      1. Weigh animal. For those salamanders that weigh 4-10 g, blood may be collected from the heart
      2. Calculate maximum volume of blood that may be removed (see paragraph IV-A above).
      3. Position. The sedated or anesthetized salamander is placed on its back on a flat surface.
      4. Location of heart The heart of salamanders is located in the ventral midline of the anterior chest but in a position much closer to the head than in frogs and toads; the heart of salamanders may appear to be located in the neck.
      5. Supplies. Arrange syringes, slides, capillary tubes, sealing wax and labeling supplies for easy and rapid accession once blood has been taken.
      6. Cleaning the skin. The skin of the ventral neck and chest should be gently rinsed with a stream of clean water from a squirt-bottle. Optionally, the site may then be sprayed with a short burst of BactineŽ (disinfectant). Do not use any other disinfectant (eg, alcohol, other sprays, iodine-based disinfectants, bleach, etc) on the skin of amphibians. For more explanation of disinfectants see also NWHC ACUC Protocol 2001-004, Toe-Clipping of Frogs and Toads.
      7. Syringe. Remove the syringe from its paper wrapping. Move the plunger back and forth about 2-4 cm in the syringe case, then push in the plunger completely. Remove the safety sheath from the needle and position the needle in the ventral midline of the chest about in the middle of the sternum with the needle pointed towards the head. The needle and syringe should be at a very acute angle (15-30 degrees) with the syringe almost parallel to the body.
      8. Insertion of needle into heart. The needle and syringe are pushed through the skin gently, slowly and steadily. When the needle has advanced 1- 2 mm through the skin, the plunger is gently pulled back about 5 mm. The needle should just barely slide off the anterior (neck area) edge of the sternum. Advance the needle passed the sternum into the heart.
      9. Drawing blood. Keep steady but gentle vacuum ("suction") in the syringe. When the needle enters the heart, a small amount of blood will appear in the syringe at the hub. Immediately stop advancing the syringe and needle and allow blood to continue to flow into the syringe. It is not necessary to pull the plunger more than 5 mm in the syringe case to create sufficient vacuum to draw blood. If blood stops flowing, the syringe may be moved 0.5 to 1 mm back and forth to resume blood flow into the syringe. Sometimes it also is helpful to simply rotate the syringe 90 to 180 degrees in order to reposition the bevel at the tip of the needle. NOTE: blood flow into the syringe will be very slow because of the small diameter of the needle and because of lower blood pressure in amphibians (compared to mammals and birds).
      10. Withdrawing needle. When 0.2 to 0.5 ml of blood are obtained, the plunger is released before pulling out the needle from the salamander's body (this avoids aspirating air into the syringe which may greatly accelerate clotting of the blood).
      11. Slides and capillary tubes. Two blood smear slides are immediately prepared with blood in the syringe (see NWHC ACUC Protocol 2001-003). Remaining blood in the syringe is then very gently injected into capillary tubes. Each capillary tube is filled only two-thirds (~65%) full and then sealed at both ends with sealing wax. Do not completely fill capillary tubes.
      12. Wound treatment. Treatment of the needle hole in the skin is optional. A brief spray of BactineŽ onto the hole in the skin is recommended.
      13. Failures to obtain blood. If blood initially enters the syringe and then stops, gently rotate the syringe 90-180 degrees. If no blood is obtained on the first insertion of the needle, withdraw the needle almost to the point that the needle leaves the skin, then gently pushed the needle towards the heart a second time. If the second insertion of the needle into the area of the heart fails, then cease all further attempts. NOTE: do not repeatedly plunge the needle and syringe back and forth into the chest; the limit on insertions of the needle into the area of the heart is twice.

  6. Possible Complications.
    1. Bleeding from needle hole in skin. This problem rarely occurs because the skin is elastic and quickly seals the hole when needle is withdrawn. If more than two drops of blood appear at the needle hole, gentle pressure on the hole with a cloth or piece of cotton will stem blood flow and promote clotting.
    2. Slow anesthetic recovery. Occasionally, amphibians may take longer than 1 hour to recover from anesthesia and blood collection. If an animal appears to be recovering slowly, rinse its entire body thoroughly in fresh (non-chlorinated) water to remove any residual anesthetic on the skin. About 2% of anesthetized amphibians may recover slowly from anesthesia; these animals may have to be held for several hours or overnight. During this holding period, keep the amphibian out of direct sunlight, and in a cool shaded location.
    3. Death. Deaths during or shortly after collection of blood from the heart are rare (less than 1%). Occasionally, some amphibians may appear dead because of limpness, flaccidity of limbs, and lack of response to touch to the eyes. Nevertheless, such amphibians may still have a beating heart, so they should be held overnight in a protected, cool, shaded location. If an amphibian remains completely limp and flaccid after being held overnight (or it becomes stiff from rigor mortis), it is recommended that the carcass be preserved immediately in 75% ethanol or 10% buffered neutral formalin. The carcass should be slit open in the ventral midline to allow rapid fixation of internal organs before immersion into the fixative. The carcass can then be submitted for necropsy (dissection) and other diagnostic tests.
    4. Disease. Occasionally, blood may be collected from amphibians at a site where a die-off is occurring or occurred in the last 2-3 weeks. All amphibians from a recent or ongoing casualty site should be presumed to be infected with a contagious agent. To avoid spreading disease from one amphibian to another, never reuse needles and syringes. It is recommended that the person(s) anesthetizing amphibians and collecting blood wash their hands between each animal, (or change gloves between each animal), and that the table or other surface be washed between amphibians. All surfaces and instruments should be washed and then thoroughly rinsed in fresh water to avoid having soap, disinfectants or other chemicals coming into contact with amphibian skin.

  7. References.
    1. Literature
      1. Baranowski-Smith, L.L., and C.J.V. Smith. 1983. A simple method for obtaining blood samples from mature frogs. Laboratory Animal Science 1983:386-387.
      2. Wright, K. 1995. Blood collection and hematological techniques in amphibians. Bulletin of the Association of Reptilian and Amphibian Veterinarians 5:8-10.
    2. SOPs (Standard Operating Procedures):
      1. ACUC Protocol 1997-004. Collection, Preservation and Mailing of Amphibians...
      2. ACUC Protocol 1997-005. Short Term Storage of Amphibians
      3. ACUC Protocol 1997-006. Anesthesia and Euthanasia [of amphibians]
      4. ACUC Protocol 2001-002. Storage and Processing of Amphibian Blood Samples
      5. ACUC Protocol 2001-006. Anesthesia of Amphibians in the Field
      6. ACUC Protocol 2001-003. Making Amphibian Blood Smear Slides
 

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