TITLE: Preparation of mollusc larval shells for individual geochemical analysis
AUTHORS: Vincent Mouchi, Thomas Broquet, Thierry Comtet

[DESCRIPTION]
This protocol describes the digestion process of mollusc larvae and is modified from the procedure found in Becker et al. (2005). Here, we indicate the entire process to obtain the digestion solution, as well as cleaning glassware, actual larval digestion and extraction of shells until storage for subsequent geochemical analysis or observation. This protocol is applicable to shelly larvae of any species, once they are collected and brought to the laboratory.

The goal of this preparation is to isolate each larval shell to perform individual geochemical analyses, instead of dissolving several specimens for bulk analysis. To do this, great care needs to be taken to the handling of the shells using ultra-pure solutions and acid-washed glassware to avoid chemical contamination of all kind.

Reference:
Becker, B.J., Fodrie, F.J., McMillan, P.A., Levin, L.A., 2005. Spatial and temporal variation in trace elemental fingerprints of mytilid mussel shells: a precursor to invertebrate larval tracking. Limnology and Oceanography, 50, 48-61. doi: 10.4319/lo.2005.50.1.0048.

[GUIDELINES]
The protocol that follows was used to study the elemental fingerprints of Shinkailepas tollmanni larval shells. This is a small hydrothermal vent limpet that is particularly abundant on the shells of larger provannid gastropods. We collected nearly ready-to-hatch veliger larvae (approximately 100 μm) by opening egg capsules deposited by females in ridges at the surface of Ifremeria nautileii shells. Larvae were collected from Ifremeria shells aboard the research vessel L'Atalante during an oceanographic cruise in the southwest Pacific in spring 2019 (CHUBACARC, Hourdez & Jollivet, 2019). Upon collection, larvae were stored dry at -20°C in groups of 10 to 100 in 2mL microtubes until processed according to the protocol described here.

Reference:
Hourdez, S. & Jollivet, D. 2019. CHUBACARC cruise, L’Atalante R/V. doi: 10.17600/18001111. Sismer.

[STEPS]
SECTION: Preparation of the digestion solution (for 20 mL final volume of approximately 15% H2O2)
1. In a glass beaker
Add 10 mL  Suprapur® hydrogen peroxide (H2O2) 30%



SECTION: Preparation of the digestion solution (for 20 mL final volume of approximately 15% H2O2)
2. 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.

SECTION: Preparation of the digestion solution (for 20 mL final volume of approximately 15% H2O2)
3. 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.
SECTION: Preparation of the digestion solution (for 20 mL final volume of approximately 15% H2O2)
2.2. Using a 1 mL cone with pipette, slowly introduce sodium hydroxide (NaOH) 0.15 M in the beaker until pH reaches 8.5. 
 

SECTION: Preparation of the digestion solution (for 20 mL final volume of approximately 15% H2O2)
2.1. Activate shaker and place pH meter electrode in the H2O2 solution.
SECTION: Preparation of the digestion solution (for 20 mL final volume of approximately 15% H2O2)
2.3. Remove pH meter electrode and deactivate shaker.
SECTION: Cleaning tools and containers
4. Put 500 mL of Primar PlusTM – Trace analysis grade nitric acid (HNO3) (>68%) in each of two 2-L PTFE beakers (subsequently referred to as beakers A and B, respectively).
SECTION: Cleaning tools and containers
5. Complete each beaker with 1.5 L  ultra-pure water (resistivity 18.2 MΩcm).
SECTION: Cleaning tools and containers
6. 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.
SECTION: Cleaning tools and containers
7. Cover with a PTFE lid both beakers and leave   under a fume hood.
SECTION: Cleaning tools and containers
8. 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 10 min .

SECTION: Cleaning tools and containers
9. Empty and discard the HNO3 solution from the PTFE beaker A in a dedicated chemical waste container.
SECTION: Cleaning tools and containers
10. Rinse the PTFE beaker A (now empty).
SECTION: Cleaning tools and containers
10.1. In the PTFE beaker A: add 1 L ultrapure water and discard in the sink.

SECTION: Cleaning tools and containers
10.2. Repeat step 10.1 twice.
SECTION: Cleaning tools and containers
10.3. In the PTFE beaker A: add 2 L ultrapure water.

SECTION: Cleaning tools and containers
11. Transfer all materials from the PTFE beaker B (containing HNO3) to the beaker A (now containing ultrapure water).
SECTION: Cleaning tools and containers
12. 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.
SECTION: Cleaning tools and containers
13. Rinse the PTFE beaker B (now empty) following Step 10.1 and Step 10.2.
SECTION: Cleaning tools and containers
14. Place both PTFE beakers (A: containing ultrapure water and all materials; B: empty)   under the laminar flow cabinet with PTFE lids.

SECTION: Cleaning tools and containers
15. On the following morning, add 2 L ultrapure water to the PTFE beaker B.

SECTION: Cleaning tools and containers
16. 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.
SECTION: Cleaning tools and containers
18. Once dried, place the tools in the cleaned plastic bags and zip. Leave those under the laminar flow cabinet until use.
SECTION: Cleaning tools and containers
17. Discard used ultrapure water from both PTFE beakers in the sink.
SECTION: Cleaning laboratory room
19. At the beginning of each sample preparation day, activate the laminar flow cabinet and wait for at least 60 min prior starting with the sample preparation. Wear lab coat and nitrile gloves at all times in the room.

SECTION: Cleaning laboratory room
20. In the meantime, clean all surfaces of the laboratory with low lint wipes and absolute ethanol, including the inside of the laminar flow cabinet.
SECTION: Cleaning laboratory room
21. Sweep the room.
SECTION: Digestion of larvae
22. 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.
SECTION: Digestion of larvae
23. Label the glass salt dip with sample name using a permanent marker.
SECTION: Digestion of larvae
24. Let thaw the samples stored in microtubes (approximately 15 minutes).
SECTION: Digestion of larvae
25. 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.
SECTION: Digestion of larvae
26. 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.
SECTION: Digestion of larvae
27. Slowly add 2 mL  H2O2 to fill the glass salt dip with pipette and a low-retention tip. Cover with a lid and leave   .
SECTION: Digestion of larvae
28. 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).
SECTION: Digestion of larvae
29. 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.
SECTION: Extract cleaned larval shells
30. When digestion is over, remove most of the solution using pipette and low-retention tip without extracting larval shells and discard tip.
SECTION: Extract cleaned larval shells
31. Fill the glass salt dip with ultrapure water with pipette and low-retention tip. Discard tip.
SECTION: Extract cleaned larval shells
32. Leave 60 min  with Petri dish as cover.
SECTION: Extract cleaned larval shells
33. 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.
SECTION: Extract cleaned larval shells
34. Clean a sable brush with absolute ethanol. Dip the brush in ultrapure water for re-hydration and avoiding hydrophobic interactions.
SECTION: Extract cleaned larval shells
35. 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).
SECTION: Extract cleaned larval shells
36. 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.
SECTION: Extract cleaned larval shells
37. Clean the brush with absolute ethanol and re-hydrate with ultrapure water.
SECTION: Extract cleaned larval shells
38. Repeat steps 35 to 37 as necessary to place all specimens on the tab.
SECTION: Extract cleaned larval shells
39. Let the surface of the tab dry under the laminar-flow cabinet.
SECTION: Extract cleaned larval shells
40. Surround the group of deposited larval shells on the tab using a permanent marker to facilitate their location during geochemical acquisition.
SECTION: Extract cleaned larval shells
41. Clean the glass salt dip with ultrapure water and absolute ethanol, check cleanliness under the stereomicroscope.
SECTION: Extract cleaned larval shells
42. Let the glass salt dip dry under the laminar-flow cabinet.
SECTION: Extract cleaned larval shells
43. Store the slide in a clean slide box and cover the box with parafilm for storage until geochemical analyses.
SECTION: Extract cleaned larval shells
44. At the end of the day, wipe all surfaces in the room with low lint wipes and absolute ethanol.
SECTION: Extract cleaned larval shells
45. Switch off the laminar flow cabinet before leaving the room.