Introduction to Conducting Stable Isotope Measurements for Animal Migration Studies

C H A P T E R

2 Introduction to Conducting Stable Isotope Measurements for Animal Migration Studies Leonard I. Wassenaar International Atomic Energy Agency, Vienna, Austria

2.1 INTRODUCTION

commercial, government, or university laboratories that measure samples on a fee-for-service basis. This leads to a discussion between the isotope analyst, who may have little knowledge about the project, and an ecologist who may have little knowledge about pertinent details of stable isotope measurements. The process usually begins with an enquiry from an ecologist interested in using stable isotopes to investigate animal movement, but who has little familiarity with what to do from an isotope analytical perspective. What follows is a dialogue that forms the framework of this chapter. What are stable isotopes? Which isotopes could be used to answer research questions for migratory species of interest? What animal tissues are best used for isotope analysis? How to prepare the samples? How much sample mass is needed? Are there caveats? How much will it cost? Why did duplicates give such different results?

Since publication of the first edition of Tracking Animal Migration with Stable Isotopes a decade ago, the application of stable isotopes (δ13C, δ2H, δ15N, δ18O, δ34S, 87Sr) in ecological sciences has blossomed into numerous and diverse studies of terrestrial and aquatic animal migrations (Vander Zanden, Soto, Bowen, & Hobson, 2016). Similarly, there have been significant advances in stable isotope methodologies along the way. This chapter briefly reviews the light stable isotope measurement terminology, methods and practices currently applicable for use in migration ecology. The analytical aspects of conducting stable isotope measurements for migration studies are particularly important to consider, especially since few biologists and ecologists operate their own stable isotope instrumentation laboratories. Most ecologists rely on

Tracking Animal Migration with Stable Isotopes DOI: https://doi.org/10.1016/B978-0-12-814723-8.00002-7

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2. INTRODUCTION TO CONDUCTING STABLE ISOTOPE MEASUREMENTS FOR ANIMAL MIGRATION STUDIES

These are relevant questions that should be considered at the beginning of a project to help ensure a successful outcome using stable isotopes. Addressing crucial analytical problems mid-way, after avoidable problems emerge, or forging ahead without a good understanding of what stable isotope analyses can or cannot do, or measuring tissues without understanding their dynamics, can lead to some painful discussions (MeierAugenstein, Hobson, & Wassenaar, 2013; Wunder et al., 2009). Because of the high potential for misunderstandings, this chapter attempts to bridge the gap by arming the ecologist with key information to ensure a basic understanding of the terminology and methods and issues around isotopic measurements. Thereby, the ecologist gains confidence that the isotopic assays selected are appropriate, conducted correctly, and that the results hopefully lead to meaningful and quantitative spatial information regarding migratory animal movement.

2.1.1 What are Light Stable Isotopes? Many of the individual elements of the periodic table have a range of stable isotopes, i.e., variants of one element having the same number of protons, but differing only in their number of neutrons (Criss, 1999; Hoefs, 2015; Sharp, 2017). However, there are very few elemental stable isotopes that are easily measurable or of practical use in animal migration studies. The most useful elements are the socalled “light isotopes” of carbon, hydrogen, nitrogen, oxygen, and sulfur (CHNOS), important elements which constitute the proteinaceous and building blocks of life: biosphere (plants, animals), hydrosphere (H2O) and atmosphere (N2, O2, and H2O). These five elements, in varying proportions, constitute almost 100% of the dry mass of animal and plant tissues, ranging from B50% for carbon, to around 6% for hydrogen in proteins (Table 2.1). Each of these elements have two or more stable isotopes whose ratios vary widely in nature. Other “heavier” elements (e.g., Hg,

TABLE 2.1 Approximate Elemental Abundances as (dry) Mass Fraction in wt. %, the Stable Isotope Ratios of Interest, and Expected Stable Isotopic Ranges for Bulk Tissues (e.g., α- or β-Keratins Like Hair or Feathers) Commonly Used in Migratory Research Element

Wt. %

Isotope Ratios

δ-Range

Mass Required

LIGHT ISOTOPES Carbona

30
40 wt. %

13

25 to 265m (PDB)

0.2
1.5 mg

27
40 wt. %

18

16

110 to 130m (VSMOW)

0.2
0.5 mg

12
19 wt. %

15

14

6
8 wt. %

2

b

Oxygen

a

Nitrogen

b

Hydrogen

Sulfur 5
20 wt. % HEAVY ISOTOPES Strontium ,100 2 x ppb

C/12C O/ O

22 to 125m (Air)

0.5
1.5 mg

1

2250 to 190m (VSMOW)

0.1
0.4 mg

32

220 to 130m (CDT)

1
2 mg

Absolute ratios

2
30 mgc

N/ N

H/ H

34

S/ S

87

Sr/86Sr

a

C and N isotopes are generally obtained simultaneously by CF-IRMS, but need sufficient sample obtain enough N. H and O can be obtained simultaneously by pyrolytic methods. Dependent on the Sr concentrations determined beforehand. Keratins commonly contain amino acids such as glycine, alanine, and cysteine. Mass of sample typically required for each isotopic assay also applies to other biological tissues like muscle, claw, and blood. The primary isotopic reference materials are PeeDee Belemnite (PDB), Vienna Standard Mean Ocean Water (VSMOW), atmospheric N2 (AIR), and Canyon Diablo Triolite (CDT). b c

TRACKING ANIMAL MIGRATION WITH STABLE ISOTOPES

2.1 INTRODUCTION

Pb, Fe, and Sr) also have stable isotopes that may be useful, but these elements typically occur at trace element (,ppm
ppb) mass fractions and do not form the core of tissue protein structure. The heavier isotopes are also difficult to extract, easily contaminated, and costly to measure, or otherwise exhibit very little useful isotope ratio differences in nature (Chapter 1: Animal Migration: A Context for Using New Techniques and Approaches). One particularly useful “heavy isotope” is 87Sr, a trace element that can be extracted from tissues and has controlled geological distributions that may be utilized (Table 2.1; Chapter 3: Isoscapes for Terrestrial Migration Research, Chapter 4: Application of Isotopic Methods to Tracking Animal Movements). The CHNOS elements have one highly abundant “light” isotope (e.g., 12C; 98.894%) and one or more “heavier” or rare isotopes (e.g., 13C; 1.1056%) (Criss, 1999; Hoefs, 2015). The ratios of the rare to common isotope vary minutely in natural materials due to a variety of biogeophysical processes, but these tiny variations in ratios can be used by scientists to great advantage (Brand & Coplen, 2012). It is exceedingly difficult to directly measure the absolute isotope ratios, or the concentrations of one stable isotope in a tissue. To overcome this, it was recognized over 70 years ago that it was far easier and convenient to measure the relative differences in rare-to-abundant isotope ratios of a sample converted to a pure gas (e.g., CO2, H2, SO2, and N2) compared to an identical reference gas with known ratios, by using isotope-ratio mass-spectrometers, or IRMS (McKinney, McCrea, Epstein, Allen, & Urey, 1950). This is a first point of confusion for newcomers; stable isotopes are assays comparing isotope ratios to isotope ratios (Fry, 2006). Generally, gas source IRMSs are used to measure the light isotopes in all environmental materials. This has important implications for the migration ecologist. The first implication is that their biological samples (feather, muscle,

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blood, and claws) cannot have stable isotope ratios measured on bulk material directly, instead milligram-sized subsamples must be sampled and quantitatively converted into ultrapure analyte gases like CO2, N2, or H2. In other words, stable isotope assays are sample destructive. The second implication is measured isotope ratios are obtained to 4
6 decimal places, numbers which are difficult to examine, and are not Syste`me International (SI) quantities. Instead, by convention, researchers receive their results tabulated as positive or negative “δ-values” (delta values). The δ-values are defined as the part-perthousand (m, or per mille) deviation of the ratios from a “zero-point” primary reference material (Fry, 2006; Hoefs, 2015). The idea of “deviations in ratio differences” is even more confusing to those accustomed to and who prefer SI concentration units like μg/kg or mg/L. Because mass spectrometers measure very small isotope ratio differences between two analyte gases, an examination of the standard δ-equation reveals what the δ-value really means:   Ratiosample δX in m 5
1 (2.1) Ratiostandard where X is the stable isotope of interest (δ2HVSMOW, δ13CPDB, δ15NAIR, etc.) expressed in m (Coplen, 2011). The right side of the equation is the isotopic ratio of the sample (gas) relative to a laboratory standard gas measured by the mass spectrometer for the isotopes of interest (13C/12C, 18O/16O, 2H/1H). The ratio of the laboratory standard gas must be determined by calibration to an appropriate primary reference material (usually defined to be “zero” m). Because isotope-ratio differences lie in the 4th
6th decimal place, the results from Eq. (2.1) are simply multiplied by 1000 to obtain numbers having 1
2 decimal places (e.g., δ2H 5 2150.1m). Because the primary reference materials and their ratios act as our

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2. INTRODUCTION TO CONDUCTING STABLE ISOTOPE MEASUREMENTS FOR ANIMAL MIGRATION STUDIES

zero point, the reported δ-values may be negative or positive values, only because they are relative to the ratios of the primary reference materials (Groning, 2004). Negative δ-values do not mean there are negative concentrations of the isotopes, and for simple 2- or 3-isotope systems one can easily convert negative δ-values into positive mass fractions like ppm (Fry, 2006; Speakman, 1997). The primary reference material anchors were established decades ago based on arbitrary natural materials that were limited in quantity or are now exhausted, hence laboratories need to measure samples against appropriate well-calibrated secondary reference materials (Paul, Skrzypek, & Forizs, 2007). Recently there was a proposal to replace the classical m (per mille) symbol with an SI compliant unit called the Urey (Ur) (Brand & Coplen, 2012). Adoption of the Ur remains inconsistent in the scientific literature, hence in this book we retain the conventional m symbol. The conversion is 1m 5 1 mUr.

2.1.2 Isotope Fractionation and Discrimination Mass dependent isotope fractionation (Criss, 1999; Hoefs, 2015; Sharp, 2017) occurs when a chemical, biological, or physical process drives a change in the isotope ratios of the source material and/or reactant due to slight chemical differences arising from the subtle differences in isotopic masses. There are two primary kinds of isotope fractionation processes. Unidirectional or irreversible isotope fractionation is referred to as kinetic isotope fractionation, whereas equilibrium fractionation is chemical reactions that are fully or partly reversible (Hoefs, 2015). In short, if there was no isotope fractionation in nature, all components of the hydrosphere, atmosphere, and biosphere would have identical isotopic ratios and stable isotopic assays would be pointless.

Fortunately, isotope fractionation of the light isotopes in nature is not only widespread, but highly diverse, and often characteristic in both magnitude and its direction. Many examples of isotope fractionation in environmental systems are found in the recommended textbooks listed below. For the migratory ecologist, isotope fractionation usually involves using measured δ-value differences between two or more related bulk substrates because samples are often of necessity “bulk” tissue materials with no clearly defined chemical stoichiometry that can be followed (e.g., comparing feather vs food). For example, in the global H-isotope system there are consistent and large δ-value “offsets” between water, plants, and bulk tissues of an organism. These isotope fractionations are here more appropriately referred to as net isotopic discriminations, since every hydrogen pool and chemical form of hydrogen in all three substrates cannot be fully known. Net isotope discrimination, while a needed oversimplification, allows us to exploit stable isotopic measurements in studying animal migration (see Chapters 3
6). As with all oversimplifications, the peril lies in the assumed details, and a researcher may be required to defend the assumptions with controlled experimentation (diet to tissue experiments), and maintain a critical eye toward what the measured isotopic data really represents.

2.1.3 Isotope Mass Spectrometry Instrumentation The type of IRMS instrumentation currently available to ecologists nowadays are predominantly continuous-flow isotope-ratio massspectrometers (CF-IRMS), which gained widespread adoption since the 1990s (Fry, Brand, Mersch, Tholke, & Garritt, 1992; Matthews & Hayes, 1978). Compared to their ultra-high precision dual-inlet counterparts (DI-IRMS)

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2.1 INTRODUCTION

preparative automation. Even δ34S, a historically difficult assay, can be conducted using CF-IRMS to fit-for-purpose analytical precision for migration studies (, 6 0.4m). For organic tissues used in animal migration studies, milligram subsamples are burned to convert samples to CO2, H2O, SO2, or N2 gas using EAs. Schematic illustrations of some contemporary CF-IRMS systems for C, N, H, and O isotope analyses are depicted in Fig. 2.1.

developed in the 1950s, there are key advantages to CF-IRMS-based assays. The pros are low cost analyses and rapid throughput, which are achieved by linking preparative modules like Elemental Analyzers (EAs) or HighTemperature Thermochemical (HTC) reactors for 13C 1 15N, or 2H, and 2H 1 18O isotopes. The trade-off used to be lower analytical precision, although CF-IRMS assays nowadays can exceed dual-inlet precision due to extensive Elemental Analyzer for δ13C and δ13N Auto sampler O2 injection

He Reference gas injectors

He flow in

Water trap Quartz wool Reduction 650ºC

Quartz tube Combustion 1050ºC Chromium oxide Quartz chips

Magnesium perchlorate GC column

Pneumatic needle valve Purge

50ºC 3m PoropakQ 50ºC

Copper

N2

Silvered cobaltous Cobaltic oxide Quartz wool

N2

CO2

CO2

Quartz wool

TCD NUPRO isolation valve

He diluter (Optional)

Stand-by valve

CF-IRMS

Elemental Analyzer for δ2H or δ18O Auto sampler Oxygen setup Hydrogen setup

He flow in

He Open split

Ceramic outer tube

Furnace >1100– 1300ºC Glassy carbon / Cr wire Quartz wool

CO

Diluter (Optional)

Glassy carbon inner tube H2 NiC Glassy carbon Chips Quartz wool

H2

Reference gas injectors

CO TCD

50ºC

GC column 1 m 5 A molecular sieve packed column

*90ºC for Hydrogen

Pneumatic needle valve Purge NUPRO isolation valve Stand-by valve

CF-IRMS

FIGURE 2.1 Typical Elemental Analyzer (EA, top right) preparative interface to a CF-IRMS system for δ13C, δ15N, or

δ34S (top left, S configuration not shown) and δ2H and δ18O (bottom left) in organic samples (courtesy of Elementar GmbH). The Uni-Prep device (bottom right) and autosampler for automated isothermal online “comparative equilibration” and sample drying mounted to an EA-IRMS (courtesy Eurovector SpA.)

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2. INTRODUCTION TO CONDUCTING STABLE ISOTOPE MEASUREMENTS FOR ANIMAL MIGRATION STUDIES

Much of the recent instrumental developments have focused on automation and multiple (13C, 15N, 34S) isotope assays, decreasing the sample size requirements, and increasing throughput while maintaining high precision and accuracy. Some mass spectrometer companies promote multielement stable isotopic 13 C 1 15N 1 34S measurements on single samples (Mambelli et al., 2016), although in practice, for the ecologist such assays are more difficult for samples with varying C:N:S elemental ratios. Radiogenic 87Sr isotopes, conversely, are conducted by trace-element solid source IRMS. Numerous papers and books describe the routine preparative and combustion-based conversions of environmental samples for stable isotope analyses by IRMS. Some summaries and historical perspectives are found in Volumes 1 and 2 of the Handbook of Stable Isotope Techniques (de Groot, 2004). The cost (USD) of stable isotope analyses in 2018 range from $7
20 for 13C 1 15N, to $15
50 for 18O/2H, $50
100 for 34S, to $100
200 or more for 87Sr. Costs, however, often vary widely and can be reduced through collaborative partnerships, or by the ecologist undertaking some of the costly sample preparation steps. Recent promising developments in IRMS analyses involve compound-specific or moleculespecific isotope analysis (CSIA-IRMS), where tissue samples are processed for C, N, or H isotope analysis of selected targeted compounds using either gas chromatography (GC) or high-pressure liquid chromatography separation interfaces to an IRMS (Fogel, Griffin, & Newsome, 2016). CSIA assays promise distinctive advantages over traditional bulk tissue analyses, but suffer from difficult or laborious sample preparation (i.e., derivatization), require highly specialized instrumentation, and accordingly low sample throughput due to long processing times required. Chapter 7, Amino Acid Isotope Analysis: A New Frontier in Studies of Animal Migration and Foraging Ecology, summarizes

the most recent and promising developments and the pros and cons of using CSIA for 13C, 15 N, and 2H analyses of selected amino acids for use in migratory studies. Since 2010, lower-cost laser-based isotope analyzers brought exciting potential for easy to operate instruments compared to IRMSs. Laser isotope analyzers have already largely displaced IRMS for water isotopes and some greenhouse gases (Lis, Wassenaar, & Hendry, 2008). Despite their potential, currently no automated sample preparative devices exist to convert biological bulk tissue samples to CO2 or H2O gas for laser-based isotope measurements. Perhaps someday the connection of automated EAs to lasers for δ2H or δ13C assays may be achievable (Koehler & Wassenaar, 2012). For further reading about stable isotoperatio mass spectrometry and broader applications of stable isotopes in environmental chemistry, the reader is referred to classic textbooks published over the past decades (Hoefs, 2015), and online textbooks like Principles of Stable Isotope Geochemistry (Sharp, 2017). For newcomers to environmental isotope measurements in ecology the following introductory books are highly recommended: Stable Isotope Ecology (Fry, 2006), Stable Isotopes in Ecology and Environmental Science (Lajtha & Michener, 2007), Isoscapes: Understanding Movement, Pattern, and Process on Earth through Isotope Mapping (West, Bowen, Dawson, & Tu, 2009), Stable Isotope Forensics: An Introduction to the Forensic Application of Stable Isotope Analysis (Meier-Augenstein, 2011), and Groundwater Geochemistry and Isotopes (Clark, 2015).

2.2 SAMPLE COLLECTION AND PREPARATIVE METHODS For animal migration studies, H (and possibly O) isotopes are typically of paramount interest. These two isotopes are considered

TRACKING ANIMAL MIGRATION WITH STABLE ISOTOPES

2.2 SAMPLE COLLECTION AND PREPARATIVE METHODS

global-spatial assays, because the patterns of H and O isotopes in terrestrial ecosystems are controlled by global-scale hydrologic processes that are seasonally and spatially predictable by latitude and elevation over multiyear timeframes, and over spatial scales ranging from kilometer to continental scales (Fig. 2.2; Chapter 3: Isoscapes for Terrestrial Migration Research). These patterns are revealed in the 50-year long-term Global Network for Isotopes in Precipitation (www.iaea.org/water) and its spatial water isotope maps (Terzer, Wassenaar, Aragua´s-Aragua´s, & Aggarwal, 2013). A prominent feature of isotopic spatial predictability is

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a high degree of confidence in making H and O isotopic predictions into areas where no observational data exist (Bowen, Wassenaar, & Hobson, 2005; Terzer et al., 2013). The oceans are global O and H reservoirs that are mostly isotopically homogenous and highly distinctive from terrestrial freshwater ecosystems, which allows for clear marine versus terrestrial distinctions to be made. The C, N, S, and Sr isotopes, conversely, are categorized as localspatial assays, because there is no a priori reason their isotope ratios vary predictably in time or continuously across landscapes to the same degree as H and O isotopes do, although

FIGURE 2.2 Predicted growing-season hydrogen isotope (δ2H) patterns in precipitation across the globe (Terzer et al., 2013). These distinctive H isotope patterns are mirrored in plants and upper-level trophic organisms and forms the foundational “isoscape” for tracking animal movement. Migratory movements are often North2South, spanning continental or regional isotopic gradients. Oxygen isotopes exhibit a similar pattern. Hydrogen and oxygen isoscape maps and data can be found in the Global Network of Isotopes in Precipitation www.iaea.org/water, with public access to GNIP data and maps at https://nucleus.iaea.org/wiser. Additional isoscape datasets for marine and aquatic systems are compiled at http://wateriso.utah.edu/waterisotopes.

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2. INTRODUCTION TO CONDUCTING STABLE ISOTOPE MEASUREMENTS FOR ANIMAL MIGRATION STUDIES

they could exhibit exploitable patterns at some relevant ecosystem scale (Chapter 3: Isoscapes for Terrestrial Migration Research).

2.2.1 Tissue Sample Considerations for Isotopic Assays The first issue facing scientists using CHNOS stable isotopes in migration research involves the type of samples to be collected for the species of interest for isotopic analysis. There are many possibilities: hair, claw, muscle, blood, wings, fins, thorax, tissue punch, carapace, etc., but two clear distinctions can be made from an isotopic perspective: fixed versus dynamic tissues (sometimes referred to as metabolically inactive vs metabolically active, respectively). A discussion of the pros and cons and potential of fixed versus active tissues are fully discussed in Chapter 4, Application of Isotopic Methods to Tracking Animal Movements. Further, from an isotopic measurement aspect, the researcher needs to be fully aware of issues of inter- and intrasample isotopic heterogeneity because of the large potential for obtaining confounding isotopic results. This issue applies to both bulk- and compound-specific isotope analyses. In short, what and where you sample for isotope measurements on a tissue is highly relevant and requires forethought. Intersample isotopic heterogeneity is defined as isotopic differences that occur among similar tissues (i.e., feathers) on the same animal due to inherent variance in isotope discriminations occurring during biochemical synthesis of tissues. We expect some isotopic intersample variance, e.g., among flight feathers grown by one individual at a location. Secondly, we expect intersample isotopic variance for similar feathers from different birds at the same location. Intersample isotopic heterogeneity of both types is always greater than instrumental error for all isotopes, but is ideally less than

large-scale isotopic patterns to make geospatial interpretations in migratory studies. Intrasample isotopic heterogeneity is defined as stable isotopic variance that occurs at the tiny mg to μg (or mm to cm) scale within or along the length of one sample used for isotopic analyses (Hobson et al., 2017). Intrasample isotopic heterogeneity is also greater than instrumental analytical uncertainty. For migrant animals, the problem may be further exacerbated in slowly growing tissues while the animal is moving across large spatial distances or feeding in different isotopic biomes, and can lead to distinctive isotopic trends within a single tissue (which may also be useful). One example of useful intrasample isotopic patterns is in fish otoliths which can be used to infer movement (Chapter 6: Isotopic Tracking of Marine Animal Movement). The isotope analyst’s uninformed answer to the problem of intrasample isotopic heterogeneity is often to “pulverize the sample” to produce a homogenous powder, a practice that could seriously complicate matters, as demonstrated below. Two contrasting examples of intrasample isotopic heterogeneity are illustrated in Fig. 2.3. One panel shows δ2H data along the length of a bald eagle flight feather. These data show distinctive longitudinal H isotopic patterns in the feather related to its southern migratory movement during feather growth. As the feather grew, it gained more positive δ2H values as the eagle moved southward from Canada into the United States, hence recorded a “net migratory track” inside a single “fixed” feather. By contrast, a single feather from a lesser scaup showed no significant intrasample tissue isotopic differences because it was fully grown at the molt site. The scaup feather represents an excellent candidate to assess molt origins regardless of the position on the feather where the subsample for isotope assays was taken. But, had the researcher been unaware the eagle feather was growing en route, and took one subsample for isotopic analysis to determine natal

TRACKING ANIMAL MIGRATION WITH STABLE ISOTOPES

2.2 SAMPLE COLLECTION AND PREPARATIVE METHODS

Scaup feather

Intrasample hydrogen (δ2H) isotopic heterogeneity in migratory bird feathers. The feather on the left is from a lesser scaup (Aythya affinis) that grew this feather at one location, and a bald eagle (Haliaeetus leucocephalus), right, that grew the feather during southward migration. Subsamples for δ2H were taken from the vane up and down each side, and from the top and bottom of the rachis (R).

FIGURE 2.3

Bald eagle feather –114 (R) –120

–128 (R)

–117 –120 –121

–112

–117

–114

–116

–110

–111 –123

–111

–111

–96 –94 –124

–124

26 cm

10 cm

–124

33

–95

–100

–95

–90 –85 –128 (R)

–82

–79

–93 (R)

origin, highly misleading results could occur. Movement during tissue growth is one reason why “duplicate” samples taken from different parts of the same feather or tissue (often unwittingly assumed by a researcher to be homogenous “repeats”) can give startlingly different isotopic results. In other words, what might be presumed to be a single fixed tissue sample representing only one location may in fact be an interesting spatial tracking “recorder.” Most investigators try to avoid issues of sample heterogeneity by always making measurements on the same tissue (e.g., a focal tail or wing feather) and measuring the same part of that tissue (e.g., just the vane section of the distal section) (Hutchinson & Trueman, 2006; Mazerolle, Hobson, & Wassenaar, 2005; Coplen and Qi, 2013). Yet another consideration is whether or not endogenous reserves are mobilized into tissue formation versus local diet only (Fox, Hobson, & Kahlert, 2009).

2.2.2 Expressing Sample Isotopic Uncertainty The sample heterogeneity issues described earlier will lead to varying degrees of sample isotopic variance, and is usually expressed in the standard deviation (SD) or standard error (SE) of n measurements of sample replication or using population data. The uncertainty of individual- or population-based measurements must be properly propagated into the assignment models to obtain realistic geospatial assignment and associated uncertainty estimates (Chapter 8: Design and Analysis for Isotope-Based Studies of Migratory Animals, Chapter 9: Isoscape Computation and Inference of Spatial Origins With Mixed Models Using the R package IsoriX). Correct expression of isotope analytical uncertainty is often overlooked, and unfortunately, there is very little consistency among

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2. INTRODUCTION TO CONDUCTING STABLE ISOTOPE MEASUREMENTS FOR ANIMAL MIGRATION STUDIES

stable isotope laboratories in determining and reporting measurement uncertainty (Wassenaar et al., 2018). It is possible, e.g., to repeat a feather sample 10 times for δ2H and calculate a SD that is lower than the uncertainty inherent in the primary reference material VSMOW. The uncertainty budget of an isotopic measurement contains several obtainable components: (1) the uncertainty of the primary reference anchor, (2) the uncertainty of the secondary calibration standard(s) used, and (3) optionally, the uncertainty obtained from replicates of the same sample (intrasample variance). The point is this: reported sample isotopic uncertainty (Uc) cannot be less than the summed uncertainty inherent in the reference and calibration materials used, and should be determined and conservatively reported using an error propagation method, like the squared root of the sum of the squares: Uc 5 O½ðUVSMOW Þ2 1 ðUCBS Þ2 1 ðUREPEATS Þ2 . . . To illustrate, a feather is measured five times for δ2H. The uncertainty of the primary reference material to which all results are reported (VSMOW) is 6 0.3m, the uncertainty of the laboratory keratin calibration standard (i.e., CBS keratin) is 6 0.9m, and the SD of five feather repeats is 6 0.2m (i.e., less than VSMOW, and indicating very low intrasample variance which is an important feature). The propagated uncertainty is correctly reported, to an appropriate number of significant digits, by: Uc 5O½ð0:3Þ2 1ð0:9Þ2 1ð0:2Þ5 61:0 m ðnot60:2 mÞ If the laboratory used for isotope analyses does not clearly reveal how their uncertainty is reported, it is worth asking how it was done, and armed with information about the calibration and control standards, can be determined with error propagation postprocessing.

2.2.3 Other Sample Collection Issues Museum or archeological tissue samples, often attractive for historical information regarding a migrating or extinct species, could have been chemically treated or preserved in formalin in the past. The researcher must be vigilant to potential isotope fractionation from sample degradation or by contamination from isotopic exchange with the chemicals used or artifacts from applied preservatives (Hobson, Gibbs, & Gloutney, 1997). Testing for treatment effects may be required. One study demonstrated that acetone preservation of archived dragonflies had no effect on δ2H or δ18O values of their chitin (Hobson, Soto, Paulson, Wassenaar, & Matthews, 2012). Ethical considerations include whether a tissue sample may be obtained nonlethally. For many species, several milligrams of hair, nail, fin clip, plucking a feather, or muscle biopsy could be safely taken without affecting the health of the individual (Hayden et al., 2015) or tissues are easily regrown. For small migratory species like insects (moths, dragonflies) the entire body may be required to obtain sufficient sample, hence the animal is euthanized. This may be a minor issue or a serious concern in the case of endangered species. Animal care and national or international regulations must be observed when tissue sampling. Finally, isotopic results obtained for fixed or dynamic tissues will need to be related to on-the-ground known-origin equivalent samples or predictively using water isotope spatial patterns (e.g., isotopic “base maps” or “isoscapes”) to facilitate quantification of geospatial movement. The assumptions and challenges in making robust geospatial connections and species-specific isoscape maps for migratory studies are fully discussed in Chapter 3, Isoscapes for Terrestrial Migration Research and Chapter 8, Design and Analysis for Isotope-Based Studies of Migratory Animals.

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2.2 SAMPLE COLLECTION AND PREPARATIVE METHODS

In summary, it is incumbent upon the researcher to carefully consider the sample type and isotopic assay most likely to satisfy the requirement of answering, wholly or in part, the migration research question. Data scrutiny requires knowledge of the biology of the species and possibly experimentation with the species or tissue under study, e.g., to better establish water-diet-tissue isotope discrimination, or to verify tissue isotopic heterogeneity. It is unwise to blindly extrapolate experimental findings regarding intrasample isotopic homogeneity from one to a different species, as illustrated in Fig. 2.3. However, for many passerines, waterfowl, insects, and fish the water-dietary-tissue isotope discriminations for δ2H and δ13C are remarkably uniform (Clark, Hobson, & Wassenaar, 2006; Hobson, 1999; Soto, Hobson, & Wassenaar, 2016). Notably, stable isotopic assays are never deceptive; it may be our understanding of the isotope biochemistry of the organism and its tissue that is lacking (Meier-Augenstein et al., 2013).

2.2.4 Sample Cleaning and Storing Once appropriate tissue samples have been collected there are questions concerning storage and subsequent laboratory preparation for isotope analysis. Field samples may be dirty, matted, bloody, or greasy, so some degree of sample cleaning may be required to remove extraneous contamination. While cleaning procedures for fixed tissues are often straightforward, there is debate concerning what to do with dynamic tissues (e.g., defatting, plasma separation, and acid treatment). In short, any sample treatment used should not alter the isotopic integrity and signal of the sample, whether it is a bulk tissue sample or targeted compound-specific analysis. For fixed tissues like feather, hair, nail, or claws which are tough and do not easily

35

decompose, the sample cleaning procedure is uncomplicated. If there is dirt or adherent material, samples can be washed with distilled water, and air, oven, or freeze-dried. Where there are natural oils (hair, feathers), samples must be cleansed using a solvent. Solvent cleaning is strongly recommended to remove surface oils for C and H isotopes because oil, grease, or waxes are markedly different (usually depleted in the heavy isotope) compared to bulk tissue, and so could impart uncontrolled negative isotopic biases (Hobson et al., 2017). Note that natural oils on feathers or hair typically do not usually contain S or N, so solvent cleaning is not needed for these isotopic assays. For tissue trace element isotopes (e.g., 87Sr) sample cleaning and contamination of tissues is a serious concern, since results are easily compromised by dust particles and by handling in the field or laboratory—extensive “clean-lab” procedures are usually needed for using trace element isotopes (Bortolotti, 2010; Font, Nowell, Pearson, Ottley, & Willis, 2007). For soft tissues like muscle, blood, or liver, the approach to cleaning and preservation procedures is not as clear. In the field, these samples are usually stored cold, frozen, or somehow preserved to avoid decomposition. In the laboratory, a longstanding debate revolves around the fact that solvent or acid treatments alter the isotopic composition of the bulk living tissue to some degree by selectively removing fatty acids, carbonate, or amino acids, and thereby altering the original bulk isotopic composition (CNS) of the sample (Pinnegar & Polunin, 1999; Post et al., 2007; Sotiropoulos, Tonn, & Wassenaar, 2004; Yurkowski, Hussey, Semeniuk, Ferguson, & Fisk, 2015). Opinions range from no cleaning, solvent cleaning, the use of C/N ratios to correct for fat content, or using empirical lipid correction models (Elliott, Davis, & Elliott, 2014; Skinner, Martin, & Moore, 2016). Unfortunately, there is unlikely to ever be a

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2. INTRODUCTION TO CONDUCTING STABLE ISOTOPE MEASUREMENTS FOR ANIMAL MIGRATION STUDIES

“one-size-fits-all” cleaning or lipid correction approach that applies universally to all tissues. Nevertheless, removal of lipids is generally agreed as an essential procedure to remove bias induced by 13C and 2H depleted lipids (Sessions, Burgoyne, Schimmelmann, & Hayes, 1999; Soto, Wassenaar, & Hobson, 2013). By removing lipids, we are better assured that the H and C isotopic analysis is conducted on bulk protein. In some countries, samples shipped to laboratories for isotopic analyses must be sterilized (e.g., Australia and New Zealand). Testing has shown that autoclaving (high pressure 120 C and steaming for 15 minutes) has no measurable impact on the C, N, or H isotopic composition of resilient keratins or chitin (T. Horton, Personal communication). However, depending on the resilience of the fixed tissue, less extreme sterilization methods may be preferable (alcohol storage, irradiation), the only criteria being that any sterilization process will not alter the original isotopic composition of the sample. It is not recommended to treat or wash samples with sodium hydroxide solutions, since NaOH decomposes organic samples. Both fixed and dynamic tissues for stable isotopic analyses should be properly stored before and after preparative procedures to prevent decomposition (and loss of isotope information or fractionation). Storage may include vials or envelopes at room temperature, or samples stored frozen. If properly prepared and preserved, the isotopic integrity with storage time will not be compromised. One known exception is formalin preservation which affects carbon and nitrogen isotope ratios (Edwards, Turner, & Sharp, 2002; Hobson et al., 1997; Barrow, Bjorndal, & Reich, 2008). When short-term storage is required for practical or field collection reasons, consider using 70% ethanol
water mixtures and plastic vials, then fully process the samples after returning from the field.

2.2.5 Sample Weighing All bulk tissue samples submitted for stable isotope measurements require careful analytical microbalance weighing into specialized cups or capsules prior to stable isotopic analysis. Many isotope laboratories perform weighing on a fee-for-service basis, and the client need not be overly concerned about the technical details. Sample weighing, including specialized capsules, cost $3
15 per sample. Accurate and precise microbalance weighing of samples is critical as the target mass for each isotope is instrument specific. This work cannot be automated and is painstakingly done by hand for each sample. However, given the precautions above regarding intrasample heterogeneity, clear subsampling instructions need to be given to the laboratory to avoid confounding results. Cost savings and improvement in sample turnaround time may be achieved if the researcher does the weighing and encapsulation. Only be aware microbalance weighing is tedious work and prone to external interferences from drafty labs, static electricity, and low relative humidity. Never begin any sample preparation work and microbalance weighing without clear laboratory guidance. Target sample weights are usually determined by the type of isotopic assay needed (C, H, S) and the class of mass spectrometer that will be used (ranges from 100 μg to .5 mg). Analytical microbalances therefore require a readability of 6 0.001 mg or better. For each isotope, the mass of each sample depends on the sensitivity of the IRMS instrument and the mass fraction (dry weight %) of the element (i.e., C, H) in the sample. Never use a “general-purpose” laboratory balance for isotopic weighing, and ensure readability and achievable precision are compatible with the isotope laboratory requirements. The types of sample capsules also depend on the type of isotope analysis—usually they are ultra-high purity tin (for C, N, and S) or silver capsules (for H and

TRACKING ANIMAL MIGRATION WITH STABLE ISOTOPES

2.2 SAMPLE COLLECTION AND PREPARATIVE METHODS

O). Specialty sample capsules cost up to $2 each and are available from scientific suppliers (see suppliers at www.isogeochem.com). It cannot be overstated—accurate and appropriately precise weighing is critical for technical reasons. The main reason is the IRMS instrument-specific dependency of the measured δ-value on the mass of the element being analyzed. This effect arises from differential gas pressures in the mass spectrometer ion source or its tuning sensitivity, as shown in Fig. 2.4. The δ-value dependency on the mass of sample used is often referred to as “ion source linearity.” Linearity is not always quantified in laboratories on a per-run basis, nor is the response always fully linear to allow for simple corrections. Usually, a laboratory targets a specific mass that the analyst has predetermined gives the lowest ion source δ-value variance. Thus the person preparing samples must closely adhere to the sample target weight and uncertainty guidelines by the laboratory. For C or N isotopes, these δ-value mass

dependencies are often a bit more forgiving (i.e., target weight 1000 6 100 μg), but always depend on the instrument used and its operational conditions. For δ2H, the δ-value mass dependency is highly sensitive for all types of IRMS instruments and is often nonlinear, easily to 10m per 100 μg of sample in a positive or negative direction. This means accuracy of target weighing to 6 10 μg is usually needed for H isotopes to reduce this significant source of isotopic variance. In short, inaccurate or high weighing variance will translate to high isotopic variance in the sample results. As noted, weighing pulverized or small pieces of sample tissue to an accuracy of 6 10 μg is very difficult work; especially for feathers, hair, or keratinaceous tissue powders that are strongly affected by static electricity (i.e., powder explodes off the microspatula). The use of antistatic matting, cotton clothing, using a room air humidifier, and deploying antistatic guns and wrist straps are all useful to reduce annoying static electricity effects.

FIGURE 2.4 The dependence of instrumental δ2H results on sample mass for two different isotope-ratio mass spectrometers using homogenized keratin powder. Weighing variance can result in δ2H changing by 10m per 100 μg of sample, well beyond acceptable uncertainty.

–105

–110 δD (o/oo) VSMOW

37

–115

–120

–125

–130 0

0.1

0.2

0.3

0.4

0.5

0.6

Keratin powder mass (mg)

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2. INTRODUCTION TO CONDUCTING STABLE ISOTOPE MEASUREMENTS FOR ANIMAL MIGRATION STUDIES

To reiterate, inaccurate or imprecise weighing will cause serious isotopic variance for replicated results within and among laboratories, and particularly for H isotopes. If the laboratory has not established δ-mass dependencies, this is a good question to ask beforehand. If contracting out weighing to a laboratory, ask for the sample weights recorded during the microbalance preparation and the elemental composition (mass fraction) automatically determined by the mass spectrometer (wt. %H, C) along with the isotope δ-value data. The mass fraction may not be routinely reported by laboratory unless explicitly requested. A cross plot of balance weight versus the % elemental (H or C) yield is a good proxy for assessing the quality of weighing and its impact on isotopic variance. An example of target masses typically used for stable isotope assays is summarized in Table 2.1. A suggested weighing procedure is shown in Table 2.2. One possible negative outcome of weighing errors or other sample handling mistakes is “isotopic outliers.” For example, samples from a population of local birds are isotopically clustered as expected, but there is an inexplicable outlier. Either the outlier is correct, or the data is faulty. The first thing the researcher should do is contact the isotope laboratory and double check if there was a weighing error or if samples were somehow mistakenly switched. If all is correct on the analytical side, repeating an outlier is warranted. If the outlier is correct, the researcher can ponder the ecological significance (e.g., recruitment of immigrants?).

2.3 GLOBAL-SPATIAL ISOTOPES 2.3.1 Hydrogen Isotopes Hydrogen (δ2H) isotopes are most frequently used in studies of long-distance animal migration due to proven and successful track records for many diverse and iconic

species ranging from birds to mammals to insects and over many spatial scales (Bowen et al., 2005; Cormie, Schwarcz, & Gray, 1994; Hobson & Wassenaar, 1997; Vander Zanden et al., 2016; Wassenaar & Hobson, 1998). Using δ2H measurements requires obtaining the H isotopic composition of the nonexchangeable H of tissue samples, which is carbonbound hydrogen that reflects the environmental H signal (water, diet) of the geographic location of tissue formation. However, hydrogen isotope analyses of organic samples suffer from two seemingly minor, interrelated, but critically important analytical concerns that do not occur with the isotopes of CNS. These concerns are the problem of (1) residual moisture in the sample plus, (2) uncontrolled H isotope exchange with exchangeable H continually interacting with ambient air moisture in the laboratory. Trying to disentangle both aspects to obtain the δ2H of nonexchangeable H is not trivial, and has long been a subject of analytical method development starting in the 1990s (Schimmelmann, Miller, & Leavitt, 1993; Soto, Koehler, Wassenaar, & Hobson, 2017; Wassenaar & Hobson, 2000). Residual moisture are water molecules from sample processing and from room air vapor that quickly adhere to organic tissue samples and are not fully removed due to insufficient drying (Qi & Coplen, 2011; Soto et al., 2017). Residual moisture H does not reflect the location of tissue formation. The definition of residual moisture also varies considerably by discipline, in the geotechnical literature it is the gravimetric difference between the sample after vacuum drying for 24 hours at 110 C. Exchangeable H is the fraction of nonremovable H atoms that form part of the organic sample tissue plus any residual moisture. Both pools readily and differentially exchange H atoms with the 6000
22,000 ppm H2O in ambient laboratory air (Schimmelmann, 1991; Sessions & Hayes, 2005; Wassenaar & Hobson, 2000). Both residual moisture and

TRACKING ANIMAL MIGRATION WITH STABLE ISOTOPES

2.3 GLOBAL-SPATIAL ISOTOPES

39

TABLE 2.2 Example Procedure for Feather Stable Isotopic Analysis Materials required: Analytical microbalance, tissue samples, clean culture tray(s), weighing utensils, methanol, Kimwipes, tray template, tape, marker, silver or tin EA capsules. 1. Obtain a clean 96 position plastic culture tray (Elisa Plate) and print out an Excel sample template. 2. Ensure feathers are solvent cleaned (2:1 v/v chloroform/methanol 24-h soak and 2 3 rinse) to remove surface oils. Air dry feathers in fume hood (24 h). 3. Clean weighing utensils using methanol and Kimwipes, allow to dry. 4. Cut off a small amount of feather material for analysis—cut samples from the same location on different feather samples if possible (e.g., near tip). Feather pieces are best cut using small stainless steel surgical scissors. 5. Make sure the microbalance is clean and calibrated. Ensure the doors are closed when taring and weighing. 6. Tare a silver EA capsule,a handling only with tweezers, remove, and set on a clean metal surface. Use the smallest available capsule that will contain the sample (e.g., 3.5 3 5.0 mm). 7. Using a spatula or tweezers, transfer a small amount of feather material into the capsule. 8. Reweigh, and continue adding or removing feather material until the target sample weight of 350 6 10 μg is obtained.b With practice this will take ,3
5 min per sample. Ensure the microbalance is accurate and stay within prescribed weight tolerance to avoid mass spectrometer linearity effects. Samples and references must be weighed to obtain comparable elemental mass yield as the samples. 9. To seal the capsule, crimp the top of the capsule using a pair of straight edge tweezers and fold tightly (as if folding down from the top of a paper bag). Then use the edge of the tweezers (use of two tweezers helps) to gently compact the tin or silver capsule into a small cube or ball. There should be no stray edges, loose sides or sample material poking out. Flattened samples (rather than cube/ball-shaped) or capsules with stray or loose edges can jam in an autosampler, cross-contaminate samples, and ruin an analysis. 10. Record final sample weight and sample name in a spreadsheet. Place the sample capsule in the 96-position tray and record the weight on the tray template. Clean all utensils with Kimwipes and methanol after each sample, air dry briefly. Secure the lid of the sample tray with rubber bands or masking tape and label the tray when done. Ensure samples cannot “jump” out of the cells when the Elisa lid is properly closed (some brands of trays allow this). 11. Record sample name and weights (in mg or μg) for each sample in the appropriate tray and its position (e.g., Tray 1, pos A5). When completed, transfer this information to appropriate isotope laboratory sample submission form. 12. Use 3.5 3 5.0 mm silver or tin capsules designed for elemental isotope analysis. Suggested suppliers are Costech (1-800-524-7219) and Elemental Microanalysis (1-800-659-9885). Silver capsules must be used for δ2H and δ18O analyses, tin capsules for δ13C, δ 15N, and δ34S. Consult isotope laboratory for target weights for each isotope. Example illustrated for feathers for H assays, but also applies similarly to claw, hair, and can be adapted for the other isotopes (CNS).

a

b

exchangeable-H together form the fraction of exchangeable H (fex) of a sample. Unfortunately, CF-IRMS instruments can only measure the total H of a sample and cannot distinguish between nonexchangeable H and fex H. The fex immediately undergoes uncontrolled isotope exchange whenever a sample is exposed to laboratory air.

Why does the net fraction of exchangeable H matter? Because if left uncontrolled or ignored, identical samples and organic calibration standards will give incomparable δ2H or δ18O results at different laboratories or at different times of the year depending on outside air temperature and moisture sources. Residual moisture heavily contributes to net fex of H.

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2. INTRODUCTION TO CONDUCTING STABLE ISOTOPE MEASUREMENTS FOR ANIMAL MIGRATION STUDIES

An apt illustrative example of this confounding effect is shown in a H-isotope intercomparison of keratins where the δ2H results (uncontrolled for fex) were unacceptably incomparable among different laboratories (Carter, Hill, Doyle, & Lock, 2009). It should be obvious that tissue samples having 20% residual moisture content will likely give a very different isotopic result for δ2H and δ18O and a higher fex than after being dried to ,1% residual moisture content. Ideally, tissue samples should have all the residual moisture removed before H isotope analysis, so that fex represents only the remaining nonremovable tissue exchangeable-H. Sample drying is therefore required, although drying practices range widely in both approaches and efficacy. Drying methods include air drying, N2 flushing, desiccator or vacuum desiccator drying (with various desiccant chemicals), oven or vacuum-oven drying (60
110 C), and freeze-drying. Reported drying times range from hours to days, sometimes using the samples already sealed preweighed in the capsules. While the drying methods used are rarely questioned, for H and O isotopes there is an additional problem as tissue samples are reexposed to air again following drying and during subsequent handling and weighing. Depending on the hygroscopic properties of the tissue (and whether in pieces or powdered), samples will immediately begin to reabsorb water moisture from the laboratory room air (Bowen, Chesson, Nielson, Cerling, & Ehleringer, 2005). Moisture reabsorption continues until air-sample H2O equilibrium is reached during the handling, weighing, or when encapsulated in trays for hours or weeks, or in the EA carousel. Drying and handling differences are one reason why fex values reported for keratins are so wildly inconsistent in the literature (ranging from 2% to 12%), likely from incomplete or incomparable moisture removal methods and different laboratory humidity conditions.

The preferred solution to remove moisture is by online heated vacuum drying (preferably .50 C), and using an approach where dried samples are never reexposed to air before their isotope analysis. At 0.1 bar vacuum, the boiling point of water is reduced to only 50 C, a temperature where many keratin and other organic tissues may be safely dried for 1
2 hours without any detrimental isotopic alteration. Residual moisture removal reduces fex and hence what is left reflects the tissue exchangeable H. Currently, the only commercial drying solution that meets all of these criteria is the Uni-Prep device for online use with HTC EAs and IRMS systems (Wassenaar, Hobson, & Sisti, 2015).

2.3.2 The “Comparative Equilibration” Approach to Organic δ 2H Analyses Another useful way to control net fex and to determine the δ2H of nonexchangeable H is an idea called comparative equilibration (Wassenaar & Hobson, 2003). Comparative “equilibration” leverages the fact that net H isotope exchange between ambient lab air moisture and fex H in samples is fast and reaches equilibrium in ,96 hours at room temperature, and much faster at hotter temperatures (Bowen et al., 2005; Soto et al., 2017; Wassenaar & Hobson, 2000). The “comparative” aspect means that each run of prepared unknown tissue samples includes at least 2
3 matrix-equivalent organic calibration standards whose nonexchangeable H δ2H values are known (see below) and are prepared along with unknown samples. During handling, as ambient laboratory moisture temporally changes its 2H content, both the samples and references equally reequilibrate their total exchangeable H in an identical fashion. These “comparatively equilibrated” standards and unknowns are then isolated from the atmosphere using a common zeroblank EA autosampler and analyzed together by IRMS in a single analysis session.

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41

2.3 GLOBAL-SPATIAL ISOTOPES

δD

The comparative equilibration method proved so successful it was commercialized into an online sample preparation device called the Uni-Prep (Wassenaar et al., 2015) (www. eurovector.at). The Uni-Prep allows exhaustive online vacuum moisture removal over a wide range of isothermal temperatures (thereby reducing fex), and subsequently facilitates controlled equilibration with injected H2O vapor under strictly isothermal conditions to “reset” the exchangeable H of all samples and standards to a uniform H isotopic composition. The advantage of the Uni-Prep approach is that samples and standards are never left uncontrollably exposed to ambient air moisture. Whether using the Uni-Prep or not, comparative equilibration at room temperatures with air exposed samples and standards can be used successfully to obtain comparable and robust δ2H

0 –20 –40 –60 –80 –100 –120 –140 –160

results for organic samples measured among laboratories worldwide, as depicted in Fig. 2.5 and Table 2.3 (Kelly, Bridge, Fudickar, & Wassenaar, 2009; Soto et al., 2017; Wassenaar & Hobson, 2003). In most laboratories H isotope measurements of tissue samples are performed on H2 derived from high-temperature EA pyrolysis and CF-IRMS (Fig. 2.1). Pure H2 is used as the sample analysis gas and the isotopic reference gas. A high-temperature EA and autosampler is used to pyrolyze samples to a pulse of H2 gas (and N2 and CO gas). The pyrolysis column consists of a ceramic or Ni-carbide tube partially filled with glassy carbon chips and/ or chromium powder held at 1100
1350 C, followed by a molecular sieve GC column at 80
100 C (Gehre et al., 2015; Schimmelmann et al., 2016; Soto et al., 2017). A GC column is

Lab 1 Lab 2 Lab 3

1

2

3

4

5

6

7

8

9 10 11 12 13 14 15 16 17 18 Sample #

FIGURE 2.5 Laboratory intercomparison of δ2H for 18 individual feathers using the “comparative equilibration” approach. No attempt was made to homogenize the feathers nor were they screened for intrasample heterogeneity. The average δ2H range per sample between laboratories was only 7m (with additional data from T. Jardine and R. Doucett).

TABLE 2.3 Laboratory Intercomparison of δ2H Results for Powdered Keratins Run Over Many Months Using the “Comparative Equilibration” Approach Lab 1 mean

Lab 1 SD

Moose hair

2 163.5

2.1

Vole hair

2 106.2

Human hair (IAEA-085) Keratin (Spectrum)

N

n

Lab 2 mean

Lab 2 SD

84

2 164.7

2.4

54

2.3

85

2 105.1

2.6

21

2 70

2.4

84

2 68.7

2.6

51

2 117.2

1.9

103

2 116.1

2.7

106

The results show excellent reproducibility and comparability of results among different laboratories Courtesy T. Jardine and R. Doucette, unpublished data.

TRACKING ANIMAL MIGRATION WITH STABLE ISOTOPES

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2. INTRODUCTION TO CONDUCTING STABLE ISOTOPE MEASUREMENTS FOR ANIMAL MIGRATION STUDIES

used to resolve sample H2 from N2 and CO. All δ2H results are reported in units of per mil (m) relative to the VSMOW-SLAP standard scale using new keratin reference materials (Table 2.4). Recently, the use of Cr-based, or C/Cr mixed bed reactors instead of pure C chips has been shown to improve the δ2H results for nitrogenous organics like keratins by reducing isotope fractionation that arises from formation of HCN (Coplen & Qi, 2016; Gehre et al., 2015). The sample throughput for δ2H is fast, about 2.5 minutes per sample, and at low cost. Analytical uncertainty is often better than 6 1.0m.

2.3.3 Organic Reference Materials for Nonexchangeable H The success of applying the “comparative equilibration” approach for δ2H assays fundamentally hinges on using matrix equivalent organic reference materials with similar exchangeable H and hygroscopic properties as the unknown samples, and having known δ2H values carefully calibrated to the VSMOWSLAP scale. Hydrogen isotope calibration of complex organics has long been problematic due to lack of appropriate reference materials.

As a result, laboratories previously used an array of in-house, or inappropriate standards (like minerals, nonequivalent organic standards), and generally inconsistent approaches (Meier-Augenstein et al., 2013), leading to more confusion and unacceptably irreproducible results among laboratories. Recently, a suite of four new keratinous organic keratin standards was made formally available by Environment Canada and the USGS. All are available through the USGS reference materials web portal (https://isotopes. usgs.gov/lab/referencematerials.html). These keratin standards (Table 2.4) span a wide range of δ2H values, and are fit-for-purpose for use with most proteinaceous tissues (keratin, muscle), although this assertion should be further tested by varied tissue type. These new keratin standards are USGS42 and USGS43 (Indian and Tibetan human hair), CBS (Caribou hoof), and KHS (Kudu horn). Their currently assigned δ2HVSMOW values are 272.9m, 244.4m, 2157.0m and 235.3m, respectively (Coplen & Qi, 2016; Soto et al., 2017). Their assigned values and fex are tabulated in Table 2.4. It is important to note that these four keratinous reference materials were released several years ago, and were initially incompatible with

TABLE 2.4 Recommended Organic Keratin Standards δ2HVSMOW Values for Nonexchangeable H and for δ18OVSMOW of Oxygen From 2017 Standard

δ 2HVSMOW

Wt. % H

fex Ha

δ 18OVSMOW

Wt. % O

Material

Type

Source

KHS

2 35.3 6 1.1m

6.6 6 0.1%

1.1 6 0.4

1 21.2 6 0.3

B22%

Kudu Horn

Powder

Env. Canadab

USGS43

2 44.2 6 1.0m

6.3 6 0.1%

1.3 6 0.3

1 14.11 6 0.1

22%

Human Hair

Powder

USGS

USGS42

2 72.2 6 0.9m

6.3 6 0.1%

1.0 6 0.3

1 8.56 6 0.1m

22%

Human Hair

Powder

USGS

2 157.0 6 0.9m

6.6 6 0.1%

1.1 6 0.3

1 2.4 6 0.3m

B22%

Caribou Hoof

Powder

Env. Canadab

CBS

Determined by online vacuum drying at 105 C and dual-water equilibrations using the Uni-Prep device. Prepared by L.I. Wassenaar and K.A. Hobson at Environment Canada, Saskatoon, Canada in 2010. These keratin H and O standards can be purchased in 0.5 g quantities (approx. 1 year supply) from the USGS at https://isotopes.usgs. gov/lab/referencematerials.html. Assigned δ2H values, fex, and dry mass fractions H are from Soto et al. (2017). Comparable δ2H and δ18O values and dry mass fractions of H and O are from Qi et al. (2016) and Coplen and Qi (2013). Conversion equations for previous (deprecated) δ2H values for all of the standards can be found in Soto et al. (2017). a

b

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2.3 GLOBAL-SPATIAL ISOTOPES

each other. They underwent testing and revisions, the currently accepted and unified δ2H results were published in 2017 (Soto et al., 2017). This means years of older publications using the initial USGS (USGS43/44) and Environment Canada (CBS/KHS) assigned δ2H values need their data corrected to maintain comparability with these unified standards going forward. The conversions for the old δ2H data are as follows: For older (pre-2017) publications using deprecated CBS and KHS δ2H values: Revised δ2 HVSMOW 5 10:774 1 0:852  δ2 Hold value (2.2) For older (pre-2017) publications using deprecated USGS 43 and USGS 44 δ2H values: Revised δ2 HVSMOW 5 5:743 1 0:993  δ2 Hold value (2.3) A good δ2H IRMS analysis template for comparative equilibration would be to use at least 2
4 CBS and KHS as standards in each run for conducting two-point normalization, and measure USGS42 and USG43 as unknown controls to ensure accurate results are obtained. Notably these keratinous standards need to be prepared along with the unknown samples and undergo identical preparation procedures. Despite the excellent δ-range of these new reference materials, in the future it would be useful to develop a fifth keratin standard with a δ2HVMSOW value in the 120m to 150m range since many avian species and other organisms exhibit such positive δ-values (De Ruyck, Hobson, Koper, Larson, & Wassenaar, 2013). In the case where migrant tissue samples are not closely matrix equivalent to these available keratin standards, the researcher will be forced to conduct vapor equilibration experiments with at least two isotopically distinctive waters and run all samples twice to be able to calculate the δ2H of the nonexchangeable H. The dual-vapor equilibration approach has

43

been fully described for wood reference materials (Qi, Coplen, & Jordan, 2016). The use of the Uni-Prep device will greatly facilitate and improve the speed and reliability of dualequilibration experiments.

2.3.4 Oxygen Isotopes While oxygen isotopes of tissues should essentially give the same spatial information as H isotopes, there are some nonanalytical disadvantages to using δ18O. First, the global range of δ18O values in animal tissues is relatively small, only around a 15m range. Along with an analytical uncertainty that is relatively high (B6 1.0m), oxygen isotopes have a lower signal-to-noise ratio than hydrogen isotopes in helping to resolve geospatial information. Second, the flow of oxygen in trophodynamics is more complicated than for hydrogen. There are more sources of isotopically variable oxygen (drinking water, air, food) and sinks (exhaled CO2 and O2, urine, sweat, excrement), which altogether cause more complicated net oxygen isotope fractionations between diet, drinking water, and tissue. Oxygen isotopes can be complicated to the point of being impossible for migratory animal tracking, partly depending on water use in the organism (Pietsch, Hobson, Wassenaar, & Tutken, 2011; Wolf, Bowen, & del Rio, 2011). On the other hand, for some species, O and H isotopes are reasonably well correlated (albeit noisily), which suggests there may be some value in using oxygen as a second isotope dimension for migratory assignments. Excursions from the expected linear H versus O relationship may also be instructive to better separate different migratory populations. The correlations of δ18O in tissues compared to water isoscapes for known origin samples are not nearly as good as hydrogen (Bowen et al., 2005; Hobson & Koehler, 2015; Hobson, deMent, Van Wilgenburg, & Wassenaar, 2009; Nielson &

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2. INTRODUCTION TO CONDUCTING STABLE ISOTOPE MEASUREMENTS FOR ANIMAL MIGRATION STUDIES

Bowen, 2010). Overall, there remains a paucity of δ18O data for migrant tissues, and the use of oxygen isotopes in conjunction with hydrogen and other isotopes remains largely problematic and unexplored. One clear benefit to oxygen isotopes is keratinous samples do not have any “exchangeable-O” to be concerned about, however, the strong precautions concerning residual moisture still apply. Exhaustive drying procedures to remove adsorbed water will improve δ18O results. The drying procedures described above for H still need to be followed, and are preferably done online using a device like the Uni-Prep so that dried samples never reabsorb water from air during handling. Oxygen isotope analyses of organic tissues are performed using high-temperature pyrolysis to CO and CF-IRMS (Fig. 2.1). Pure CO gas is used as the sample analysis gas and isotopic reference gas. A HTC analyzer and autosampler is used to automatically pyrolyze submilligram samples to a pulse of CO gas (and N2 and H2 gas as also produced). The pyrolysis column consists of a ceramic tube and glassy carbon tube insert, filled to the hot zone with glassy carbon chips held at .1200
1350 C, followed by a molecular sieve GC column at 40
60 C (Qi, Coplen, & Wassenaar, 2011). A 1 m GC column is used to resolve sample CO from the H2 and N2. All δ18O results are reported in units of m relative to the VSMOWSLAP standard scale using keratinous or other certified organic reference materials (i.e., IAEA Benzoic Acid Standards) or by using primary reference waters VSMOW2 and SLAP2. The sample throughput rate for δ18O is about 7
9 minutes per sample to allow for clean chromatographic separation of CO from interfering sample N2. The IAEA benzoic acid standards have δ18O values of 123.1m relative to VSMOW (IAEA601) and 171.4m VSMOW (IAEA-604). It should be noted that the δ18O of tissues generally fall between 1 10m and 120m, well

outside of the benzoic acid calibration range. Accordingly, it is much preferable to use the recently developed keratinous materials USGS42, USGS43, CBS, and KHS (Table 5). These new keratinous reference materials for δ18O are available for purchase from the USGS (https://isotopes.usgs.gov/lab/referencematerials.html). The CF-IRMS analytical methods for H2 and CO are almost identical, and because H2 gas is also produced in the same thermochemical reduction reaction as CO, some carbon-reactor IRMS systems allow for dual δ18O and δ2H assays on a single tissue sample (Hobson & Koehler, 2015). The only drawback is reduced sample throughput rate and a more complicated analytical setup involving multiple reference gases and IRMS peak jumping.

2.4 LOCAL-SPATIAL ISOTOPES 2.4.1 Stable Carbon and Nitrogen Isotopes Stable-carbon and stable-nitrogen isotope assays can be considered local-spatial analyses, but may be useful in delineating migratory populations or by providing additional information indicating type of habitat (see Chapter 1: Animal Migration: A Context for Using New Techniques and Approaches, Chapter 4: Application of Isotopic Methods to Tracking Animal Movements). There may also be larger scale spatial patterns (Chapter 3: Isoscapes for Terrestrial Migration Research) for δ13C and δ15N at regional or ecosystem scales related to plant and ecozone types (C3 vs C4 dominated landscapes). Several studies that used δ2H were able to leverage additional information from δ13C to improve the spatial resolution of migratory assignments in both terrestrial and marine systems (Cherel & Hobson, 2007; Garcı´a-Pe´rez & Hobson, 2014; Hobson, Wassenaar, & Taylor, 1999; Marra,

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Hobson, & Holmes, 1998). The analysis of carbon and nitrogen isotopes is routine in stable isotopes laboratories. The sample size requirements range from 0.5 to 1.5 mg, depending on the C:N ratio of the sample. The cost is low, often around $10 for both isotopes. There are very few complications other than the consideration about sample preparation (lipid removal is recommended for 13C), the general recommendation is to lipid extract samples for 13C but not for 15N, depending on the tissue type. The analyses of δ13C and δ15N are typically coupled isotope measurements done on the same organic tissue sample. Samples are flash combusted using an EA (Fig. 2.1). Purified CO2 and N2 are used as the sample analysis gas and the isotopic reference gases. A standard EA and autosampler holding up to 100 samples are used to quantitatively combust samples to a pulse of CO2 and N2 gas (combustion H2O is scrubbed out with a trap). The oxidation column consists of a quartz tube partially filled to the hot zone with chromium oxide held at 1050 C, followed by reduction column filled with copper wires (to reduce NOX to N2) held at 600
800 C, followed by a packed GC column held at 35
50 C. The GC often has a thermal conductivity detector to quantify and resolve CO2 from N2. Isotope peak jumping is used on the IRMS to switch between nitrogen and carbon isotopes. All δ13C results are reported in units of per mille (m) relative to the PDB standard. For organic samples and tissues, calibration is recommended by using three to four certified organic reference materials (L-glutamic acids or hair keratins). The glutamic acids standards (USGS41a, USGS40) have δ13C values of 136.55m and 226.39m relative to VPDB, and δ15N values of 147.55m and 24.52m relative to AIR, respectively. Two powdered hair C 1 N standards (USGS42, USGS43) have δ13C values of 221.09m and 221.28m VPDB, and δ15N values of 18.05 and 18.44m relative to AIR,

45

respectively. The hair C 1 N standards have similar C: N ratios as proteinaceous tissues, and combined with the glutamic acids, fully span the isotopic range encountered for migrant tissues in nature. These C 1 N reference materials are available for purchase from the USGS (https://isotopes.usgs.gov/lab/ referencematerials.html). Sample throughput is between 9 and 14 minutes per sample, with an analytical uncertainty around 6 0.2m.

2.4.2 Sulfur Isotopes For migratory organism tissues (hair, muscle, feather), sulfur is contained mainly in the form of amino acids (e.g., cysteine) often at low elemental concentrations (B, 4
5 wt. %). Conventionally, a 34S analysis on organic substrates requires a lengthy preparative process that involves quantitative highly oxidative 850 C or flash (Parr) bomb conversion of the total S in a tissue sample to an appropriate sulfate salt. The sulfate salt is converted to purified BaSO4 or to Ag2S. These purified matrices are in turn converted to SO2 or SF6 gas for analyses on a gas source DI-IRMS (Mayer & Krouse, 2004). The highest precision δ34S analyses are still done using conventional DI-IRMS, but very few isotope laboratories nowadays offer this capability. However, CF-IRMS methods evolved significantly over the past decades, with the advantage of lower cost and high sample throughput. Like C, tissue samples converted to BaSO4 or Ag2S can be combusted in an EA to produce SO2 gas. The SO2 is separated from CO2 and N2 using a GC column (Mayer & Krouse, 2004). Analysis can be made using BaSO4 or Ag2S, which requires initial preparative sample conversion steps. The precision for this method is about 6 0.4m, and requires about 15 minutes per sample (,100 μg as S). More recent advances forgo the initial sample conversion to an inorganic sulfur

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2. INTRODUCTION TO CONDUCTING STABLE ISOTOPE MEASUREMENTS FOR ANIMAL MIGRATION STUDIES

salt, and use direct EA combustion of tissue or keratin samples to SO2 without prior conversion to BaSO4 or Ag2S. This approach is promising, but is complicated by variable or high C: N:H:S ratios commonly found in biological samples. The high C:N:H:S ratios and larger sample mass required cogenerate large amounts of CO2, N2, and H2O and subsequent separation of these undesired gases by GC or trapping is essential. However, more recent innovations enabled concurrent C 1 N 1 S assays by using multiple chromatographic and chemical gas trapping methods (Fry, 2007; Mambelli et al., 2016). The prevailing consensus seems to be direct combustion and analysis is feasible when samples have .0.1 wt. % S (still requiring 2
5 mg of keratin sample), and the resulting δ34S precisions will be on the order of 6 0.5m, and this is sufficiently precise for many migration studies (marine vs terrestrial assignment). Analytical vigilance to the EA is required because the δ18O of the SO2 produced changes as the analyzer oxidant reagents deplete, requiring IRMS corrections to obtain correct δ34S values (Fry, Silva, Kendall, & Anderson, 2002). One further complication with direct tissue combustion is appropriate organic calibration standards having a wide range of δ34S values. These are nonexistent, and large discrepancies occur when mixing inorganic standards (Ag2S) and organic samples having high C:N:S ratios by EA-IRMS. Recently, a set of hair standards for δ34S was developed by the USGS. The USGS42 and USGS43 hair standards have δ34S (CDT) values of 17.84m and 110.46m, respectively, unfortunately a very small δ-range. Further, given the large sample sizes needed (2
5 mg), the USGS hair standards are costly and limited in quantity (0.5 g). The need to include many references within a CF-IRMS autorun means these USGS reference materials will be quickly used up, so it is likely that calibration and organic S reference material

development will remain in the realm of a few specialized laboratories. In short, the most uncomplicated and costeffective method for 34S remains offline combustion of organic tissue samples to pure BaSO4 salt and subsequent 34S analysis by EAIRMS using inorganic 34S calibration standards. The additional preparation costs and specialized IRMS assays result in higher costs per sample ( . $30
60).

2.4.3 Isotopes of Trace Elements—87Sr/86Sr Another type of local-spatial stable isotope analysis used in animal migration studies is the stable isotope ratios of the trace element strontium (87Sr/86Sr) (Chamberlain et al., 1997). One of the key advantages of the “heavy isotopes” over light stable isotopes is that there is little to no isotopic fractionation from geologic sources through the food web and into tissues (Blum, Taliaferro, & Holmes, 2001). Hence the Sr isotopes among all species in local food webs are expected to show fidelity to the 87Sr/86Sr ratios of the underlying bedrock or soil. Thus 87Sr/86Sr variations in and among landscapes and continents may be distinctive, but these can be highly variable at small scales in areas of complex geology and are not a priori suitable for continuous interpolation (Chapter 3: Isoscapes for Terrestrial Migration Research). As with δ34S, the 87Sr/86Sr differences between terrestrial and marine environments are distinctive (Kennedy, Chamberlain, Blum, Nislow, & Folt, 2005). Hence, similarly to the other localspatial isotopes, Sr isotopes are best used in conjunction with δ2H or other light isotopes in a multiisotope approach. One important difference between the light isotopes and Sr isotopes in fixed tissues like feathers is that Sr is a trace element at exceedingly low concentrations (e.g., ,20 μg Sr/g of

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feather) (Font et al., 2007). The only exception is calcium-bearing tissues that contain much higher Sr concentrations (e.g., bones, teeth, and otoliths) (Blum et al., 2001). For noncalciumbearing tissues this means the potential for extraneous contamination (e.g., entrained dust, handling, and background) is critically high and must be quantified and requires rigorous cleaning and QA/QC procedures (Font et al., 2007; Vautour, Poirier, & Widory, 2015). There is no standardized agreement on how this should be done. Further, trace Sr concentrations means that up to 25 mg of sample may be required. All samples require prescreening to determine the Sr content by inductively-coupled-plasma mass spectrometry (ICP-MS) before preparative procedures for isolating Sr for isotopic assays are begun. Further, extensive wet chemical or microwave digestions and selective ion chromatography are required to isolate Sr for isotopic analysis. Difficulties arise with organic samples in being able to fully extract all available Sr. Sr isotopic ratios are determined using thermal ionization mass spectrometry and using the NBS-987 standard. No organic or keratinous Sr isotope standards exist. The problem of intrasample isotopic variability has not been rigorously tested for Sr isotopes, although differences in Sr concentrations and isotope ratios between the rachis and vane of individual feathers have been reported (Font et al., 2007), although these differences were comparatively smaller than potential geospatial differences. From the preceding section this could be a major issue for slowly growing tissues of birds or animals that are moving among areas having variable 87Sr/86Sr, especially given the large sample requirements. For example, when we consider feathers of small birds weigh 10
20 mg, or hair strands 1 mg or less, sample tissue pooling may be required. The example of the eagle feather heterogeneity and the implications for confounding Sr isotope interpretations

47

regarding spatial interpretations is critical to consider. To date, the application of 87Sr isotopes in movement or migration studies are largely limited to proof of concept testing (Chau et al., 2017; Font et al., 2012), with no widespread application yet available. Finally, 87Sr/86Sr isotope analyses are very costly in comparison to the light isotopes ( . $200
300). These mass spectrometry costs do not include initial prescreening for trace Sr concentrations, nor cost of clean-lab chemical digestions. Only few laboratories worldwide have Sr isotope capabilities on offer for commercial assays. This and the large sample size requirements suggest that Sr isotopes will remain a specialized assay for projects where the value of the anticipated outcomes exceeds the high analytical cost considerations.

2.4.4 Biomineral C, O and Sr Isotopes As noted in Chapter 6, Isotopic Tracking of Marine Animal Movement, using marine isoscapes for assessing fish migratory movement often utilize isotope and trace elements in accretionary carbonate biominerals (fish otolith) or keratin (whale baleen). Baleen keratin and marine chitins are easily measured, as described earlier. Fish otoliths, however, require specialized sampling apparatus. To obtain calcite or aragonite from otolith annual rings, micromilling or laser ablation preparative device are needed, which uses a computerized microscope to target and remove accretionary carbonate bands or targets that represent a season or a time of growth (akin to sampling tree rings). The carbonate is analyzed for δ13C, δ18O, or 87Sr isotope analyses by IRMS or ICP-MS. The stable isotope analyses are conducted on tiny ,10
30 μg carbonate samples, and are painstaking and costly (Campana, Fowler, & Jones, 1994; McCulloch, Cappo, Aumend, & Mu¨ller, 2005; Wurster, Patterson, & Cheatham, 1999).

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2.5 CONCLUSIONS This chapter briefly reviews some practical aspects of measuring stable isotope analyses for use in animal migration research. The researcher is encouraged to be discriminating and critical in the application of stable isotopes. Carefully consider the isotopes to be used, and all aspects of sampling and sample preparation, as well as isotopic measurements. Only when utmost confidence in the stable isotope analyses are assured the researcher can proceed with the task of making spatial interpretations outlined in the subsequent chapters.

Acknowledgments Tim Jardine and Richard Doucett contributed data to Figs. 2.4 and 2.5 and Table 2.3.

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TRACKING ANIMAL MIGRATION WITH STABLE ISOTOPES