Team:NYU Abu Dhabi/Documentation/DOCS 20ee279bfcdc46b09c4fb108851b2757/Biology 93d1eff7b0cd4d6ca8529879e773d615/Sample Collection d07098acd0a747ecb5df04fe0b20d0a8

Sample Collection

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Sample Collection

@Marko Susic

  • Swab Method on Adult Amphibians
    • Notes
      • Optimum conditions for Bd proliferation: 22°C, neutral pH, 100% moisture saturation
    • Protocol
      1. Sample collection approach (swab):
        1. Swabbing common infection sites (hands, feet, thighs, and venter) minimizes false-negatives
        1. For samples that will be stored for long periods of time or exposed to heat and high levels of humidity ethanol (70%) should be used to preserve DNA in the sample; use of swabs with wooden shaft is advised
        1. For real-time analysis, preferred to place swabs directly into the sample tube with no preservative; use of swabs with plastic shaft is advised
      1. Sample collection personnel (swab):
        1. Person 1 is to capture and handle the amphibian, collect the fungal sample, and take desired measurements. Gloves must be changed between every animal to prevent cross-sample contamination.
        1. Pearson 2 is to keep sterile items (vials, swabs) protected from contamination and record all data. Gloves should still be worn to prevent accidental contamination.
      1. Sample collection procedure (swab):
        1. Only person 1 should attempt to capture amphibians.
        1. Minimize contact with any substrate; if the animal is dirty after capture it must NOT be washed. Rinsing zoospores off skin reduces chances of detecting light infections; it may introduce large amounts of pathogen into the aquatic environment and facilitate transmission; washing animal in contaminated water might result in false positives.
        1. In case of frogs: grip the animal by both legs anterior to the hind legs, with the ventral side up to allow access to the ventral side, feet and hands. These areas commonly have a high concentration of Bd sporangia in infected animals and will maximize the sensitivity of the assay. Position and grip on a small individual may need to be altered to swab feet first, then pelvic patch and hands.
        1. In case of anurans: swab each: rear foot, ventral surface of thigh, and ventral abdominal surface, in that order, using a sweeping motion along the length of each of the five areas to be swabbed. Five sweeps of each area, for a total of 25 sweeps per individual, will be sufficient to sample the animal.
        1. In case of salamanders there are three main methods depending on the behavior of the species:
          1. If the salamander cooperates, the swab can be placed under the ventral surface, behind the front legs; and then dragged under the animal. This process is repeated up to ten times.
          1. The salamander can simply be picked up with the swab and allowed to hang on while you turn the swab. More contact and longer contact time with the swab should increase the probability of detection, but no relationship has been established.
          1. Procedures i) and ii) work well for Bolitoglossine and other arboreal salamanders. For salamanders that will not cooperate with the first two sampling methods, place the salamander into a clean collection bag, let salamander move about in the bag, and subsequently swab the inside of the bag. This technique is best for fossorial salamanders such as ambystomatids and Oedipina spp.
      1. Preservation of sample (swab):
        1. Person 2 holds an open, alcohol filled, pre-labeled vial as Person 1 carefully inserts the used swab into the vial so that the cotton tip is suspended just above the bottom of the vial.
        1. Person 1 then carefully breaks the end of the applicator off into the vial (wooden shaft swabs only; for plastic shafts use scissors to cut shaft appropriately).
        1. Finally, Person 2 secures the cap to the vial, assures that all information has been recorded, and stores the sample in a closeable bag labeled with the date, time, and location.
      1. Disposal and decontamination of equipment:
        1. Use fresh gloves for all amphibians to be sampled
        1. Dispose of the remaining portion of the swab into waste bag/container
        1. Carefully remove gloves by pinching around the opening, and remove glove by inverting it as you remove it. You now have both contaminated gloves in one hand. With your clean ungloved hand, once again pinch the ring around the opening of the glove remaining on your hand, and remove glove by inverting it as you remove it.
        1. At the end of the survey all gear and materials in the waste bag must be decontaminated. Use a bleach solution (1:9 bleach:water) for decontamination of equipment, but be careful not to get any bleach on vials, gloves or swabs. Bleach will quickly degrade the DNA in a sample, and invariably produce a negative result, even if the animal swabbed is positive.
  • Swab Method on Tadpoles
    • Notes
      • Tadpoles normally killed and their mouthparts excised to test for pathogen
      • Swabbing was consistently less sensitive than lethal sampling, but still detected Bd
      • Experimental Bd prevalence was 41.1% when estimated by destructively sampling mouthparts and 4.7 to 36.6% (mean = 21.4%) when estimated with swabs
      • Tadpoles are often easier to detect and capture because they are confined to aquatic habitats and are generally less ephemeral and cryptic than metamorphosed individuals.
      • Tadpoles, however, generally become infected with Bd only on their mouthparts
    • Protocol
      1. mouthparts were swabbed using rounded, wooden toothpicks with sharply pointed ends. A new, clean toothpick was used for each tadpole session.
      1. The points of the toothpicks were scraped and rolled over and between the tooth rows and the keratinized beak of the tadpoles. Broken pieces of labial teeth were usually visible on the toothpick after swabbing
    • Results
      • Bd was detected on swabs in every 2-wk session. First session (2W) levels were lowest (6.7%), while levels in the final session (8W) were highest (44.0%)
      • Multiple swabs of infected individuals increased the likelihood of detecting Bd. Infection was detected on only 14% of tadpoles swabbed once. For tadpoles swabbed 2 or 3 times, 20 and 30% of individuals, respectively, were found to be infected; 6.7% of tadpoles in each of these 2 categories had positive swabs every time. Infection was detected on 53% of tadpoles swabbed 4 times, but only 8.8% of tadpoles had positive swabs all 4 times.
      • Bd detection rates varied among samplers from 11.7 to 90.0% (mean = 54.2%). Given the mean detection rate, there was approximately a 1-in-2 chance of the average swabber detecting Bd when sampling an infected tadpole. Samplers who participated in >1 session became more likely to detect Bd, perhaps due to improving their technique or an increase in the tadpoles’ infection levels, or both.
  • Biopsy Punch Method on Adult Amphibians
    • Notes
      • Non-lethal Bd isolation using biopsy punches from toe webbing from live amphibians in the field
      • Bd disproportionately colonizes skin of the abdomen, pelvic patch and feet
      • This method does not decrease survival after 1 year
      • Used sterile biopsy punches (Integra Miltex) to collect either 1.5 mm (juveniles and adults) or 3.0 mm (adults only) diameter skin tissue samples
      • 1.5mm samples collected from juvenile frogs were more likely to show Bd growth than adults
      • 3mm biopsy punch size was more likely to detect Bd growth than the 1.5mm in adult frogs
      • One of the main factors that limited ability to purify isolates was contamination by other fungi.
      • Only 1 tissue sample was taken per individual; more could improve detection rates
      • The method described prioritizes isolation success at the spatiotemporal level instead of the level of the individual animal
      • 79% (31/39) of the frogs tested positive for Bd
      • Observed Bd growth via microscopy in 87.5% (35/40) of the samples collected
    • Materials

      1) TGhL agar with antibiotics

      2) Sterile needles

      3) Hand lens or magnifying glass (optional)

      4) Sterile forceps

      5) 70% ethanol

      6) Lighter for flaming tools

      7) Ziplock bags (2-gallon)

      8) Parafilm

      9) 1.5 or 3mm diameter biopsy punches and rigid, wipeable surfaces for performing punches

    • Protocol

      1) Place the frog on the rigid work surface and hold it securely while you spread the webbing of one hind foot

      a. Choose the punch size that is appropriate for the size of the individual animal

      b. Select a punch location that is in the middle of the webbing.

      2) Position the punch on the webbing and press down firmly, then twist slightly to ensure a complete puncture

      a. Remove the punch and pick up the skin piece using a sterile needle or forceps (make sure these are completely cool if flame was used for sterilization)

    • Image

References

(Author, year)LinkDescription
Forrest Brem, 2007https://www.researchgate.net/profile/Karen_Lips/publication/267778285_FIELD-SAMPLING_PROTOCOL_FOR_BATRACHOCHYTRIUM_DENDROBATIDIS_FROM_LIVING_AMPHIBIANS_USING_ALCOHOL_PRESERVED_SWABS/links/54f8f0f00cf28d6deca2c895/FIELD-SAMPLING-PROTOCOL-FOR-BATRACHOCHYTRIUM-DENDROBATIDIS-FROM-LIVING-AMPHIBIANS-USING-ALCOHOL-PRESERVED-SWABS.pdfClassical swabbing method on adult amphibians
Richard W. R. Retallick, 2006https://www.int-res.com/abstracts/dao/v72/n1/p77-85/Swabbing method on tadpoles
Kimberly J. Cook, 2018https://www.int-res.com/abstracts/dao/v129/n2/p159-164/Biopsy punching method on adult amphibians