Elemental and isotopic assays
Otolith Decontamination for Elemental or Isotopic Analysis
Otolith Decontamination for Elemental or Isotopic Analysis
The basic handling of otoliths intended for elemental or isotopic assay is to clean, then protect from potential contaminants. A simplified protocol is as follows:
- Remove sagittal otolith pair from fish immediately after capture; alternatively, freeze fish, but do not store in liquid preservative.
- Upon otolith removal, immediately remove all adhering tissue. Handling with metal forceps or fingers at this stage is acceptable.
- Decontaminate otolith by sonifying in a series of distilled, deionized, reverse osmosis water baths (Super Q or Milli Q water) in acid-washed polyethylene vials. Brushing with an acid-washed nylon toothbrush under a flow of Super Q water can be used to remove any adherent tissue before the first sonification. All handling at this and subsequent stages must be with non-metallic, acid-washed tools (except for radiocarbon assays).
- Dry decontaminated otoliths in a positive flow laminar flow fume hood (Class 100); store in dry, acid-washed polyethylene vials. Storage in aluminum foil is appropriate for samples to be assayed for radiocarbon. However, vials are easier to use, and less likely to result in loss of small pieces when opened.
A detailed protocol for each of the decontamination steps is shown below:
Preparation of Plastic Equipment
All new plastic equipment (vials, caps, forceps, etc.) is rinsed in 95% ethanol then in SuperQ water, air-dried in a positive pressure chamber (Clean Cell) and repackaged in plastic bags prior to being acid washed.
Acid washing operations are performed in a fume hood and all equipment is handled by an operator wearing acid-resistant rubber gloves and using Teflon tongs. 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 immersed for at least 8 hours in 6N HCl trace metal grade, rinsed 3 times in SuperQ, immersed for 4 hours (rest-rinse) in SuperQ, triple-rinsed in SuperQ and stored in sealed plastic bags. After this operation, the acid washed equipment is taken to a Clean Cell to be air-dried then repackaged in sealed plastic bags prior to being used.
Warning: Hydrochloric acid 12N is diluted 1:1 with SuperQ to obtain the required 6N. The acid is added to the water. This operation is performed in a fume hood.
Otolith Decontamination and Handling
During decontamination, otoliths are handled by an operator wearing plastic gloves. Otoliths are in contact with ultra-clean chemicals and equipment/instruments made of Teflon, polyethylene and polypropylene only. Each otolith is placed in a labeled 50 ml acid-washed conical vial, covered with SuperQ water, and sonified for 5 minutes in an ultrasonic cleaner. Vials are placed in groups of 5 attached with an elastic band and sonified at the same time. After the first sonification, each otolith is scrubbed using an acid-washed toothbrush under running SuperQ (1 minute) to remove particles and/or membranes from its surface. Each otolith is then triple-rinsed with SuperQ and placed back in its acid-washed vial with fresh SuperQ. Each otolith is sonified for 3 minutes (vials in groups of 5 again) then triple-rinsed with SuperQ. Finally, the otoliths are air-dried in their vial cap inside the Clean Cell for 18-24 hours and later stored inside their respective vials tightly closed. Note that the 50 ml conical vials can be re-used for decontaminating the next set of otoliths (max 5 times), if they are not going to be used for digestion.
Vials containing the otoliths are taken to the balance for weighing one at a time. Minimizing time outside the vial and exposure to ambient air, each otolith is quickly transferred to an acid-washed vial cap kept inside the balance (tared before weighing). During this operation, the vial is kept closed. As always, the otoliths are handled with plastic forceps/plastic gloves.
If otoliths are to be digested in their 50-ml conical vial, they can be stored dry in those vials until then. Otherwise, they can each be stored in a labeled 2.5 ml acid-washed plastic vial (5 ml for bigger otoliths), tightly closed. All vials are placed in peg trays covered with a sheet of plastic inside the Clean Cell, or sealed within an acid-washed plastic bag.
In terms of vials which can be used for both otolith decontamination and acid digestion, we use 50-ml polypropylene conical vials from Fisher Scientific (part number 05-539-6). For acid washing, we buy trace metal grade 12N HCl and dilute to 50% in Super Q water. The initial ethanol soak of new vials (before acid washing) is done in 95% non-denatured ethanol. Gloves for handling otoliths and vials are disposable polyethylene from Fisher (part 11-394-100A). You do NOT need to acid wash these gloves before using. Surgical gloves often have a powder in them to make them more comfortable, but this powder can contaminate your assays, so should not be used.
The Clean-Cell is a vertical laminar flow work station, rated to Class 10 or 100. You'll probably want to buy a roll of 3 mil polyethylene sheeting or bags to line the Clean Cell floor with, which can be replaced periodically. Of course, the sheeting should be wiped down with ethanol after being put in the Clean Cell and before use.
Decontamination in preparation for bomb radiocarbon assay is similar, but somewhat less stringent, than that used for elemental assay. The cleaning and sonification steps are the same as described above. However, acid-washed plastics are not required (although they must be clean and free of all organic debris). Also, metallic instruments do not contaminate radiocarbon.
Elemental Analysis using ICPMS
Elemental Analysis using ICPMS
After the otoliths have been decontaminated, they can be analyzed for elemental composition as described under Elemental Tags. Here we describe the details of preparation subsequent to decontamination.
Digestion is carried out in pre-cleaned (acid washed) polypropylene vials (eg- 30-ml skirted polypropylene centrifuge tubes available from Sarstedt). Other vials can be used, but they should have a non-pigmented screw cap without a cap liner. Assuming the otoliths are between 100-500 mg in weight, they are digested for several hours in 2 ml of 1:1 SuperQ water (water which has been distilled, deionized, and run through reverse osmosis) and high purity nitric acid (Seastar) in the original sample containers and then warmed for 15 mins at 50-60 degrees C to complete digestion. For the larger otoliths, an additional 0.5-1.0 ml of acid is added before dilution to complete digestion. Solutions are then diluted to a final volume of 30 mL with SuperQ water. Solutions are then diluted again 1:10 in SuperQ to leave them in the final concentation 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.
A full method blank is run which includes all materials and reagents involved in the analysis, including sample vials, volumetric tubes, pipette tips, plastic film (may be used to cover open samples), water, acids and even the toothbrush used for cleaning the specimens.
Simple dilute acid standard solutions are used for the initial instrument calibrations, but high calcium matrix standards are used to monitor (and correct for) ionization interferences and instrument drift.
The samples are prepared and analyzed in separate "batches". Two or three reagent blanks are processed with each group of specimens.
A "pool" of approximately twenty otoliths is prepared and portions of this bulk solution are analyzed within each analytical batch. A similar "pool" of otolith material from previous work is also analyzed with each batch. These are not standard materials, but were chosen to provide a means of monitoring within-run and between-run precision.
The samples are generally supplied in groups, based upon sample collection location. The samples are randomized within each group and each group is divided among the preparation batches. The samples are further randomized within each preparation batch. This is done to ensure that sequence effects do not have a pronounced effect on one particular group of samples.
Microsampling techniques are utilized to mechanically extract a portion of the otolith for subsequent analysis. Portions can be extracted as either powders, milled from discrete depths (e.g. annual growth zones) or as cores (e.g. 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.
1) Microsampling Device (automated):
We have prepared a 15-minute VIDEO DEMONSTRATION of the use of the Micromill for isolating otolith material, which provides an easy and complete introduction to the process. The information below also describes micromilling, but provides more detail on some of the materials and suppliers than does the video.
A photograph of our Micromill microsampling device is depicted below. This system includes the following:
- Pentium-based PC installed with MicroMill software, Coreco Bandit video board and drivers.
- 17" monitor, maximum resolution of 1280x1024.
- MicroMill sampler (New Wave Research) consists of a Leica stereo microscope fitted with a high resolution colour video camera, fine resolution (0.25 micron) motorized XYZ stages, high torque milling chuck with adjustable speed and carbide tipped or hardened cobalt steel milling bits.
Using on screen digitization of the sample area and precise depth control this system permits automated and accurate microsampling.
In our usual mode of operation the drill bit is used to carve out areas of the otolith, leaving discrete chunks which can be decontaminated prior to analysis.
2) Hand-held Drill:
A photograph of our hand-held sampling setup is depicted to the right. This system includes the following:
- Gesswein Power Hand 2X controller and rotary tool (Z-55X) with maximum speed of 55,000 r.p.m.
- a selection of drilling bits including diamond-tipped orb and circular cutting tools
- Wild stereo microscope
- High-intensity fiber optic illuminator (Nikon) with dual gooseneck cables
- Acrylic containment device, to prevent loss of sample
- HEPA filter vacuuming unit to remove fine dust from the work area
Using a combination of drilling and sculpturing techniques this hand-held setup offers an affordable alternative to the automated MicroMill system. Although accessible, this system is labour intensive to operate and its affordability is outweighed by its lack of precision.
Examples of papers where various microsampling procedures were used include Campana (1997), Campana and Jones (1998) and Schwarcz et al (1998).
With the realization that elemental fingerprints can be used very effectively to separate mixtures of fish coming from different sources, there is increasing demand for statistical software to separate the group mixtures. Discriminant analysis is not a good option here, since the 'priors' parameter is unknown. Here, we've released a working copy of two different methods for use in separating stock mixtures based on elemental fingerprints or other continuous or categorical variables.
A Bayesian stock mixture analysis (mix.Fish) which allows for the simultaneous analysis of both continuous (elemental) and categorical data (genetic, meristic) was described in Smith and Campana (2010). The mix.Fish R package (which includes a sample dataset) is available for free download and was written for R 3.0.1. Mix.Fish for Unix is also available. Installation instructions are provided.
The Integrated Stock Mixture Analysis (ISMA) program (written for the S-Plus environment) was first described in Campana et al. (1999). The ISMA function is a maximum likelihood-based method to analyze an unknown mixture given some known (reference) groups. Below you will find the ISMA function and a companion file which provides operating instructions. To read the function into your Splus session, just enter the following: source("d:\\assess\\scaling\\ISMA.ssc") ... using your appropriate paths, of course.
Note that no matter which method is used for the stock mixture analysis, mixture analyses don't tell you if there are differences among your reference groups to start with. For this, a MANOVA or discriminant analysis is required. 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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