Preparation of mollusc larval shells for individual geochemical analysis

In a glass beaker

Add 10mL Suprapur® hydrogen peroxide (H₂O₂) 30%

Place the beaker with hydrogen peroxide on a magnetic stirrer and control pH. This step is made to prevent the calcified shells from dissolving in the H₂O₂ solution.

Activate shaker and place pH meter electrode in the H₂O₂ solution.

Using a 1 mL cone with pipette, slowly introduce sodium hydroxide (NaOH) 0.15 M in the beaker until pH reaches 8.5.

Remove pH meter electrode and deactivate shaker.

Cover the beaker with parafilm and leave it at least 24h, for the solution to release gas (may cause the shells to “explode” when filled with gas otherwise). This solution is active for approximately a week. For multiple series of successive sample digestions ( i.e. , over more than a week), prepare this solution every Monday to use it from Tuesday to Friday.

Put 500mL of Primar Plus^TM – Trace analysis grade nitric acid (HNO₃) (>68%) in each of two 2-L PTFE beakers (subsequently referred to as beakers A and B, respectively).

Complete each beaker with 1.5L ultra-pure water (resistivity 18.2 MΩcm).

Place in the PTFE beaker A: Pasteur pipettes (at least one per sample to prepare, plus backup), Petri dishes, zip plastic bags for storage and transport of all materials.

Cover with a PTFE lid both beakers and leave under a fume hood.

On the next morning, transfer all materials from the PTFE beaker A to the PTFE beaker B (the one that contains the clean HNO₃ solution) for rinsing for 0h 10m 0s .

Empty and discard the HNO₃ solution from the PTFE beaker A in a dedicated chemical waste container.

Rinse the PTFE beaker A (now empty).

In the PTFE beaker A: add 1L ultrapure water and discard in the sink.

Repeat step 10.1 twice.

In the PTFE beaker A: add 2L ultrapure water.

Transfer all materials from the PTFE beaker B (containing HNO₃) to the beaker A (now containing ultrapure water).

Transfer the HNO₃ solution from the PTFE beaker B to another beaker for storage. This “used” HNO₃ solution can be used as a first bath (equivalent of beaker A at Step 6) for additional tools to clean.

Rinse the PTFE beaker B (now empty) following Step 10.1 and Step 10.2.

Place both PTFE beakers (A: containing ultrapure water and all materials; B: empty) under the laminar flow cabinet with PTFE lids.

On the following morning, add 2L ultrapure water to the PTFE beaker B.

Have each material from the PTFE beaker A rinsed a few seconds in the ultrapure water from the PTFE beaker B, then placed to dry under the laminar flow cabinet. Pasteur pipettes can hold some water at their tips even under the laminar flow. Try and dry those by shaking the pipettes by hand with a few strong strokes.

Discard used ultrapure water from both PTFE beakers in the sink.

Once dried, place the tools in the cleaned plastic bags and zip. Leave those under the laminar flow cabinet until use.

At the beginning of each sample preparation day, activate the laminar flow cabinet and wait for at least 1h 0m 0s prior starting with the sample preparation. Wear lab coat and nitrile gloves at all times in the room.

In the meantime, clean all surfaces of the laboratory with low lint wipes and absolute ethanol, including the inside of the laminar flow cabinet.

Sweep the room.

Clean glass salt dips with absolute ethanol using a squeeze bottle (no wipes to avoid lint) over sink. Leave to dry under the laminar flow cabinet.

All subsequent steps must be performed under the laminar flow cabinet.

Label the glass salt dip with sample name using a permanent marker.

Let thaw the samples stored in microtubes (approximately 15 minutes).

Collect some (approximately 50) larvae from their storage container ( e.g. , microtubes) using an acid-washed Pasteur pipette. Place those in the glass salt dip under the stereomicroscope, drop by drop, spacing the drops on the glass salt dip surface to avoid previously released larvae to stick on the pipette. Check that no remaining larvae are stuck inside the pipette by capillarity. If so, attempt extracting them with ultrapure water. Discard the Pasteur pipette.

Remove most of the water (that was collected with the larvae) from the glass salt dip using a pipette with a low-retention tip. Avoid contact between the tip and the shells as those can attach on the tip by capillarity. Discard tip.

Slowly add 2mL H₂O₂ to fill the glass salt dip with pipette and a low-retention tip. Cover with a lid and leave .

Repeat steps 24 to 27 for each sample to a maximum of 200 specimens total. Expect to process approximately 200 larval shells on the next day. Switch off the laminar flow cabinet before leaving the room (leaving the cabinet on overnight has proven to induce substantial evaporation of H₂O₂ where samples were being digested in, even if the containers were covered with a lid).

On the following morning, check that the digestion is over. Shells should be translucent under natural light under the stereomicroscope and should exhibit Maltese cross pattern under polarised light. If digestion is still ongoing, remove most of the solution using pipette and low-retention tip without extracting larval shells, fill the glass salt dip (2 mL) with clean H₂O₂ solution with pipette and clean low-retention tip and leave until digestion is complete. Discard tip. If digestion is not complete by mid-day, the H₂O₂ solution may not be sufficiently active anymore. Our experience is that at this point, larval shells tend to start exhibiting abnormal flexibility and dissolution.

When digestion is over, remove most of the solution using pipette and low-retention tip without extracting larval shells and discard tip.

Fill the glass salt dip with ultrapure water with pipette and low-retention tip. Discard tip.

Leave 1h 0m 0s with Petri dish as cover.

Clean a microscope slide with a low-lint wipe with absolute ethanol. Place an Extra Pure adhesive tab on the slide and write the name of the sample on the slide.

Clean a sable brush with absolute ethanol. Dip the brush in ultrapure water for re-hydration and avoiding hydrophobic interactions.

Carefully collect larval shells with the brush (1-5 specimens at a time) by capillarity under the stereomicroscope (use polarized light for easy visualization of the larval shells).

Carefully place the larval shells on the Extra Pure adhesive tab on the slide (under the stereomicroscope in natural light using the cold light source directed on the tab’s surface) by slowly sweeping the brush on the tab’s surface.

Clean the brush with absolute ethanol and re-hydrate with ultrapure water.

Repeat steps 35 to 37 as necessary to place all specimens on the tab.

Let the surface of the tab dry under the laminar-flow cabinet.

Surround the group of deposited larval shells on the tab using a permanent marker to facilitate their location during geochemical acquisition.

Clean the glass salt dip with ultrapure water and absolute ethanol, check cleanliness under the stereomicroscope.

Let the glass salt dip dry under the laminar-flow cabinet.

Store the slide in a clean slide box and cover the box with parafilm for storage until geochemical analyses.

At the end of the day, wipe all surfaces in the room with low lint wipes and absolute ethanol.

Switch off the laminar flow cabinet before leaving the room.