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Elemental and isotopic assays for otoliths

Learn how to analyze otoliths through elemental and isotopic assay techniques.

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Decontamination

Reverse-flow fume hood used for keeping dust and contaminants off samples.

When handling otoliths intended for elemental or isotopic assay, you must clean and then protect them from potential contaminants. A simplified protocol includes:

  1. removing sagittal otolith pairs from fish immediately after capture
    • alternatively, freeze fish, but don’t store them in liquid preservative
  2. immediately removing all adhering tissue upon otolith removal
    • handling with metal forceps or fingers at this stage is acceptable
  3. decontaminating otolith by sonifying in a series of distilled, deionized, reverse osmosis water baths (Super-Q or Milli-Q water) in acid-washed polyethylene vials
    • you may brush them with an acid-washed nylon toothbrush under a flow of Super-Q water to remove any adherent tissue before the first sonification
  4. all handling at this and subsequent stages must be with non-metallic, acid-washed tools (except for radiocarbon assays)
  5. drying decontaminating otoliths in a positive flow laminar flow fume hood (Class 100)
  6. storing samples in dry, acid-washed polyethylene vials
    • vials are easier to use, and less likely to result in loss of small pieces when opened
    • storage in aluminum foil is appropriate for samples to be assayed for radiocarbon

Preparation of plastic equipment

To prepare all new plastic equipment, such as vials, caps and forceps:

  1. rinse in 95% ethanol then in SuperQ water
  2. air-dry in a positive pressure chamber
  3. repackage in plastic bags prior to being acid washed

Perform acid washing operations in a fume hood. Wear acid-resistant rubber gloves and use non-stick tongs while handling all equipment. The acid bath is made of cross-linked high-density polyethylene (XLPE) kept covered before and during immersion. Water baths are either made of XLPE or high density polyethylene (HDPE).

The plastic equipment to be decontaminated is:

  1. immersed for at least 8 hours in 6N HCl trace metal grade
  2. rinsed 3 times in Super-Q
  3. immersed for 4 hours (rest-rinse) in Super-Q
  4. triple-rinsed in Super-Q
  5. stored in sealed plastic bags

After this operation, the acid washed equipment is taken to a fume hood to be air-dried then repackaged in sealed plastic bags prior to being used.

Hydrochloric acid 12N is diluted 1:1 with Super-Q to obtain the required 6N. The acid is added to the water. Perform this operation in a fume hood.

Otolith decontamination and handling

During decontamination, the otolith handler should wear plastic gloves. Otoliths are only in contact with ultra-clean chemicals and equipment/instruments made of non-stick material, polyethylene and polypropylene.

Place each otolith in a labeled 50 mL acid-washed conical vial, covered with Super-Q water and sonified for 5 minutes in an ultrasonic cleaner. Group 5 vials together and attach with an elastic band. Sonify them together at the same time.

After the first sonification, scrub each otolith using an acid-washed toothbrush under running Super-Q for 1 minute to remove particles and/or membranes from its surface.

Then triple-rinse each otolith with Super-Q and place it back in its acid-washed vial with fresh Super-Q. Sonify each otolith for 3 minutes (with the vials in groups of 5 again) then triple-rinsed with Super-Q.

Finally, air-dry the otoliths in their vial cap inside the fume hood for 18 to 24 hours. Later, store them inside their respective vials, tightly closed. You may re-use the 50 mL conical vials for decontaminating the next set of otoliths for a maximum of 5 times if they’re not going to be used for digestion.

Balance and weighing

Take vials containing the otoliths to the balance for weighing one at a time.

Minimize the otolith’s time outside the vial and exposure to ambient air. Transfer each one quickly to an acid-washed vial cap kept inside the balance (tared before weighing). During this operation, keep the vial closed. As always, handle the otoliths with plastic forceps and plastic gloves.

Digesting

If otoliths are to be digested in their 50-mL conical vial, you may store them dry in those vials until then. Otherwise, they may each be stored in a labeled 2.5 mL acid-washed plastic vial (5 mL for bigger otoliths), tightly closed. Place all vials in peg trays covered with a sheet of plastic inside the fume hood, or sealed within an acid-washed plastic bag.

Vials for both decontamination and digestion

We use 50-mL polypropylene conical vials for both otolith decontamination and acid digestion.

For acid washing, we buy trace metal grade 12N HCl and dilute to 50% in Super-Q water. Do the initial ethanol soak of new vials (before acid washing) in 95% non-denatured ethanol.

Gloves

Use disposable polyethylene gloves for handling otoliths and vials. You don’t need to acid wash gloves before using. Do not use surgical gloves. They often have a powder in them to make them more comfortable, but this powder can contaminate your assays.

Elemental analysis

After the otoliths have been decontaminated, you may analyze them for elemental composition. To prepare for that analysis after decontamination, you must prepare the otoliths.

Digestion

Carry out digestion in pre-cleaned (acid washed) polypropylene vials, such as 30 mL skirted polypropylene centrifuge tubes. Other vials may be used, but they should have a non-pigmented screw cap without a cap liner.

Digest otoliths between 100 to 500 mg in weight for several hours in 2 mL of 1 part each:

Keep otoliths in the original sample containers and then warm them for 15 minutes at 50 to 60 degrees Celsius to complete digestion.

For the larger otoliths, add an additional 0.5 to 1.0 mL of acid before dilution to complete digestion.

Dilute the final solutions to a volume of 30 mL with Super-Q water. Dilute the solutions again 1:10 in Super-Q to leave them in the final concentration of about 0.1% w/v. Rhodium is sometimes added as an internal standard to monitor changes in nebulization efficiency. Since we assay most elements using isotope dilution, Cr-52 is added as the internal standard for Mn. Enriched isotopes for assay by isotope dilution ICPMS are added at this stage. All vials received the same amount of spike.

Method blank testing

Run a full method blank, which includes all materials and reagents involved in the analysis, including:

Use simple dilute acid standard solutions for the initial instrument calibrations, but high calcium matrix standards to monitor (and correct for) ionization interferences and instrument drift.

Prepare and analyze the samples in separate batches of 2 or 3 reagent blanks. Process each group of specimens.

Prepare a pool of approximately 20 otoliths and analyze portions of this bulk solution within each analytical batch. Also analyze a similar pool of otolith material from previous work with each batch. These aren’t standard materials, but they provide a means of monitoring within-run and between-run precision.

To ensure that sequence effects don’t affect one particular group of samples:

  1. supply samples in groups based upon sample collection location
  2. randomize samples within each group
  3. divide each group among the preparation batches
  4. further randomize the samples within each preparation batch

Microsampling techniques

Use microsampling techniques to mechanically extract a portion of the otolith for subsequent analysis. You may extract portions as either powders, milled from discrete depths (such as annual growth zones) or as cores (such as first year of growth).

Assays for bomb radiocarbon and stable isotope ratios are examples of applications where a particular age or date range in the otolith is required. For these applications, the best sampling procedures involve microsampling or coring techniques which physically remove a discrete portion of the otolith.

Automated microsampling device

Our micromill microsampling system includes a:

This system permits automated and accurate microsampling using on-screen digitization of the sample area and precise depth control.

Our usual mode of operation uses the drill bit to carve out areas of the otolith, leaving discrete chunks which can be decontaminated prior to analysis.

Hand-held drill

Our hand-held sampling system includes:

Using a combination of drilling and sculpturing techniques, this hand-held setup offers an affordable alternative to the automated MicroMill system. However, it’s labour intensive to operate and its affordability is outweighed by its lack of precision.

Micromill microsampling device.

Close up of the Micromill drill bit and an otolith cross section.

Hand-held microsampler.

Statistical analysis

The ability to use elemental fingerprints to separate mixtures of fish coming from different sources has increased the demand for statistical software to separate the group mixtures.

Discriminant analysis isn’t a good option here, since the 'priors' parameter is unknown. Here, we've released a working copy of 2 different methods for use in separating stock mixtures based on elemental fingerprints or other continuous or categorical variables.

Bayesian stock mixture analysis

The Bayesian stock mixture analysis (mix.Fish) allows for the simultaneous analysis of both continuous (elemental) and categorical data (genetic, meristic).

The mix.Fish R package (ZIP file, 46 KB) includes a sample dataset and was written for R 3.0.1. Mix.Fish for Unix (ZIP file, 22 KB) is also available. Installation instructions (DOC, 30 KB) are provided.

Integrated stock mixture analysis

The integrated stock mixture analysis (ISMA) program (written for the S-Plus environment) is a maximum likelihood-based method to analyze an unknown mixture given some known (reference) groups.

To read the function into your S-Plus session, use your appropriate paths and enter: source("d:\assess\scaling\ISMA.ssc").

Discriminant analysis

No matter which method you use for the stock mixture analysis, mixture analyses don't tell you if there’re differences among your reference groups to start with.

For this, you need a MANOVA or discriminant analysis. The more differentiated your reference groups are (based on the variables you use), the more accurate your classification of your unknown mixture will be.

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