A schematic sampling protocol for contaminant monitoring in raptors

Birds of prey, owls and falcons are widely used as sentinel species in raptor biomonitoring programmes. A major current challenge is to facilitate large-scale biomonitoring by coordinating contaminant monitoring activities and by building capacity across countries. This requires sharing, dissemination and adoption of best practices addressed by the Networking Programme Research and Monitoring for and with Raptors in Europe (EURAPMON) and now being advanced by the ongoing international COST Action European Raptor Biomonitoring Facility. The present perspective introduces a schematic sampling protocol for contaminant monitoring in raptors. We provide guidance on sample collection with a view to increasing sampling capacity across countries, ensuring appropriate quality of samples and facilitating harmonization of procedures to maximize the reliability, comparability and interoperability of data. The here presented protocol can be used by professionals and volunteers as a standard guide to ensure harmonised sampling methods for contaminant monitoring in raptors. Electronic supplementary material The online version of this article (10.1007/s13280-020-01341-9) contains supplementary material, which is available to authorized users.


TABLE OF CONTENTS
Schematic protocol -Main menu 3 Important general guidelines 4 Schematic protocol for blood 5 Schematic protocol for plasma / serum 6 Schematic protocol for deserted and addled eggs 7 Schematic protocol for feathers 8 Schematic protocol for preen oil 9 Schematic protocol for regurgitate pellets / prey remains 10 Schematic protocol for internal tissues / gastric content 11 Schematic protocol for blood / plasma / serum / red blood cells: additional information 16 Schematic protocol for deserted and addled eggs: additional information 18 Schematic protocol for feathers: additional information 20 Schematic protocol for preen oil: additional information 21 Schematic protocol for regurgitate pellets / prey remains: additional information 21 Schematic protocol for internal tissues / gastric content: additional information 22

References 27
Click on the name of the matrix to see the schematic protocol for each sample type.
Click here to see important general guidelines related to permits and health and safety issues when sampling.

Table 1. Volume/Mass of sample, type of container and transport conditions required for contaminant monitoring in different matrices
a Please note that these are general guidelines. Take advice from the laboratory undertaking the chemical analysis. b Volume criteria: A general rule is that the collection weight should not exceed 2% of the body weight of the animal in any 14-day period, or 1% at any one time. Values provided in the table are volume/mass ranges generally needed in the Toxicology lab for analysis, but it will depend on the technique used. c From live birds, plucked contour body feathers (e.g. back/breast feathers) are preferred. Moulted feathers, chick down feathers and feathers from museum specimens are also useful. Consideration should be given to possible external contamination of museum feathers, e.g. due to conservation treatments. d This protocol does only deal with non-destructive sampling, thus it only refers to deserted or addled eggs. e Internal tissues collected during necropsies. 1 For feathers when they are wet or have tissue/blood attached to them, they need to be cleaned/dried or they need to be stored in a freezer and not room temperature (as then this may lead to further decay) 2 Pharmaceuticals is a broad group, and plastic containers may contain some compounds (e.g. UV filters), this should be considered or part of the plastic container analysed. Take advice from the laboratory undertaking the analysis. 3 −80°C recommended for some drugs and for long storage periods (> 3 months), take advice from the laboratory undertaking the analysis. 4 Agrochemicals and pharmaceuticals are broad groups. Some are not easy to break down (e.g. PCBs and most chlorinated pesticides has been found to be stable for at least one year at −20°C) but others may be rapidly degraded over time (

Schematic protocol for blood/plasma/serum/red blood cells: additional information
Prepare the correct needle and syringe: Take blood samples using a hypodermic needle and a syringe. Change needles between birds. Use the smallest needle possible: for birds < 500 g body weight: 30 to 25-gauge hypodermic needle and a 1-2 ml syringe / > 500 g body weight: 23-gauge hypodermic needle and a 5-10 ml syringe. Volume should be sufficient to ensure suitable analytical limits of detection (see Table 1 and Figure 2a). However, in no circumstances should the collection volume exceed 1% of the body weight.
Use anticoagulants for whole blood/plasma (e.g. heparin: 2-3 drops in a 1.5 ml-tube or heparinized tubes). See Figure 2a. EDTA may be problematic for biochemistry and metals, while heparin may interfere with PCR analysis (more info: Espín et al., 2014). Figure 2b): Stimulate the local blood circulation, e.g. by allowing wing flapping before puncturing (for brachial vein collection), use antiseptic at the phlebotomy site and take blood samples puncturing the vein. Press the puncture site with sterile dry cloth or non-woven gauze before pulling the needle from the vein, and keep pressure on the cloth at the puncture site for some minutes to avoid bleeding and haematomas (more info: Espín et al., 2014).

Transfer blood to proper tube:
Remove needle before placing the sample in tubes (see Figure 2a). Tubes containing anticoagulants should be adequately filled in order to provide a proper blood-to-anticoagulant ratio.
Transport samples at 4-10 ºC. Avoid direct contact with cold blocks/ice bags and temperatures <4 ºC to avoid haemolysis (see Table 1).
For serum/plasma collection: Use anticoagulants in the tube to obtain plasma, otherwise you will obtain serum. Centrifuge tube as soon as possible (10 minutes, 1600-3000 g), ideally within 6 hours (max. 24 h) after collection; the longer the elapsed time, the higher the risk of clotting and rupture of red blood cells. Plasma/serum/red cells separation is possible on fresh blood only and cannot be done on samples that have been frozen. Use different pipette tips for each sample during plasma/serum separation to avoid cross contamination. Keep all separated fractions (red cells, plasma/serum) in different labelled tubes.

Storage:
Keep frozen at -20°C /-80°C /liquid N2 (depending on the analyte or the studied biomarker; see Table 1). Take advice from the laboratory undertaking the chemical/biochemical analysis for further information about temperature and duration. Specific protocols for biomarkers may exist.

Schematic protocol for deserted and addled eggs: additional information
Collect only deserted eggs or addled eggs from the nest. Be careful about the timing of egg collection to avoid nest abandonment.
Transport eggs in suitable containers (e.g. polypropylene jars, chicken eggs boxes) to avoid breaking (see Figure 3a). Keep cool and process egg as quickly as possible. Use a graphite pencil to write information on both the eggshell and the container. Collect pieces of the eggshells found in the nest and keep them in sealed plastic bags (see Figure 3a), they may be useful for some contaminant analysis.
Take measurements and examine eggs before freezing: Measure length and width, and weigh the egg. Open at the equator of the egg and empty its contents into flasks, weigh and homogenise the content (using clean tools), and keep frozen until analysis (see Table 1). Examine eggs for putrefaction, embryo development (see Figure 3b) and deformities. If an embryo is present, keep frozen for future analyses. Rinse eggshell with tap water to remove all remains of egg contents from the inner surface. Dry eggshells at room temperature to a constant weight, and record the constant eggshell weight. Measure eggshell thickness at equator after drying at room temperature using a calliper (digital if it is possible) to take at least five measurements by the same investigator from the dry shell (see Figure 3a). A micrometer rather than a calliper may provide more comparable measurements.
Transfer egg content and eggshells to proper containers (see Table 1) and store samples (homogenized content at -20°C and eggshells at room temperature, see Table 1).

Schematic protocol for feathers: additional information
Collect feathers: Plucked (or cut at the skin) contour body feathers are preferred (see Figure 4). In adult birds, plucking tail or flight feathers should not be collected as it can impair the flight ability of the bird. From dead birds, all feather can be collected. Freshly moulted feathers found in the nest or field can also be collected.
Transport feathers in sealed plastic bags or envelopes (see Figure 4) at ambient temperature or using cold blocks. Before they are stored, feathers that have been plucked from living birds or collected from carcasses should be cleaned of all fresh tissue (blood, muscle) and they should be dried if they are wet. Otherwise, if they are stored at ambient temperature in sealed bags, rotting will occur. Alternatively, freeze the uncleaned feathers in sealed plastic bags.
Regarding the sample amount, see Table 1.
Identify and measure feathers: Identify type and number of the feather (left or right). In case of contour feathers, indicate the location on the body. Use the conventional numbering system for primary flight feathers from the inside out. (more info: Espín et al., 2014) Store feathers: Feathers can be kept at room temperature if stored properly and if any soft tissue or blood residue is removed. Store feathers in plastic sealed bags or envelopes, in darkness, and in a dry place (or use silica) if stored at room temperature. Alternatively, you can freeze the feathers in sealed plastic bags (see Table 1). Container materials should be checked to be free of contamination.
Click here to see video / Click here to see Figure 1. What can we measure in each sample type? (a. Active monitoring / b. Passive monitoring)

Schematic protocol for internal tissues/gastric content: additional information
Collect carcasses in sealed plastic bags to avoid dessication and label the bag (with waterproof marker). (More info: Espín et al., 2014) Transport the carcasses to the lab under cold conditions using cold blocks.
Perform necropsy: Necropsies should be carried out using protocols that avoid both potential exposure of the researcher to zoonotic diseases and chemical contamination of the sample. A proper professional necropsy requires a trained veterinarian (pathologist); however, when the aim is to collect a tissue for contaminant monitoring purposes, trained personnel may collect the samples. Necropsies should be done on fresh carcasses where possible or the carcass should be kept frozen (−20°C) until necropsy. If the carcass is frozen, thaw it overnight. External examination of the carcass is necessary to find possible signs of trauma or evidence of clinical symptoms previous to the death (e.g. haemorrhages, diarrhoea, salivation, etc). The cause of death should be determined if possible with the help of an experienced pathologist. Body/nutritive condition can be estimated as a relative score using the criteria in Figure 5a. During necropsy, record organ weight, lesions/alterations, sex and status of the gonads (developmental stage) (Figure 5b). Take pictures if possible. Take advice from the laboratory undertaking the chemical analysis as to selection of tissues (see Table 1 and Figure 1b). Use suitable dissection material ( Figure 5c) and disposable gloves, disinfect instruments and surfaces and clean the material between the organ sampling and between individuals. Regarding sampling of liver, kidney and other internal organs, the whole organ must be taken if possible. In case of muscle sampling, the pectoral muscle is the preferred choice. If sampling tissues are dispersed through parts of the body, such as fat or bone, it is recommended that the tissue is sampled consistently from the same part of the body. Collect all the gastric content. A standardised necropsy protocol should be followed. We provide a necropsy form ( Figure 5d) and figures of anatomy of birds ( Figure 5e) to facilitate the sampling. (more info: Espín et al., 2014) Transfer organs/samples to separate containers/sealed plastic bags (to avoid freezer burn) (see Table 1 and Figure 5c) and label the containers.
Store the samples at -20°C or -80°C (depending on the analyte or the studied biomarker; see Table 1). Take advice from the laboratory undertaking the chemical analysis for further information.  (2004) Note: The shape of the breast muscle generally shows active flight behavior. Nestlings often have a bilaterally concave shape, but the birds have a good nutritive condition. Therefore, this method should be used in adult individuals.

Nutritive condition
A method to assess the nutritive condition of a raptor is to measure the subcutaneous fat between the skin and the belly muscles caudal of the sternum, to measure the body fat between the belly muscles and the gizzard/gut at the caudal margin of the sternum and measure the width of the coronary fat.
Measuring body fat tissue between the belly muscles and the gizzard. The sternum has already been removed ( Figure: O. Krone).