Gas Chromatography-Mass Spectrometry-Based Targeted Metabolomics of Hard Coral Samples

Summary

Here, we present the extraction and preparation of polar and semi-polar metabolites from a coral holobiont, as well as separated coral host tissue and Symbiodiniaceae cell fractions, for gas chromatography-mass spectrometry analysis.

Abstract

Gas chromatography-mass spectrometry (GC-MS)-based approaches have proven to be powerful for elucidating the metabolic basis of the cnidarian-dinoflagellate symbiosis and how coral responds to stress (i.e., during temperature-induced bleaching). Steady-state metabolite profiling of the coral holobiont, which comprises the cnidarian host and its associated microbes (Symbiodiniaceae and other protists, bacteria, archaea, fungi, and viruses), has been successfully applied under ambient and stress conditions to characterize the holistic metabolic status of the coral.

However, to answer questions surrounding the symbiotic interactions, it is necessary to analyze the metabolite profiles of the coral host and its algal symbionts independently, which can only be achieved by physical separation and isolation of the tissues, followed by independent extraction and analysis. While the application of metabolomics is relatively new to the coral field, the sustained efforts of research groups have resulted in the development of robust methods for analyzing metabolites in corals, including the separation of the coral host tissue and algal symbionts.

This paper presents a step-by-step guide for holobiont separation and the extraction of metabolites for GC-MS analysis, including key optimization steps for consideration. We demonstrate how, once analyzed independently, the combined metabolite profile of the two fractions (coral and Symbiodiniaceae) is similar to the profile of the whole (holobiont), but by separating the tissues, we can also obtain key information about the metabolism of and interactions between the two partners that cannot be obtained from the whole alone.

Introduction

Metabolites represent the end products of cellular processes, and metabolomics - the study of the suite of metabolites produced by a given organism or ecosystem - can provide a direct measure of organismal functioning¹. This is particularly critical for exploring ecosystems, symbiotic interactions, and restoration tools, as the goal of most management strategies is to preserve (or restore) specific ecosystem service functions². Coral reefs are one aquatic ecosystem that demonstrates the potential value of metabolomics for elucidating symbiotic interactions and linking coral physiological responses to community-level and ecosystem-level impacts³. The application of high-throughput gas chromatography-mass spectrometry (GC-MS) is especially valued due to its capacity to rapidly analyze a broad range of metabolite classes simultaneously with high selectivity and sensitivity, provide rapid compound identification when spectral libraries are available, and provide a high level of reproducibility and accuracy, with a relatively low cost per sample.

Corals are holobionts consisting of the coral animal, photosynthetic dinoflagellate endosymbionts (family: Symbiodiniaceae⁴), and a complex microbiome^5,6. Overall, the fitness of the holobiont is maintained primarily through the exchange of small molecules and elements to support the metabolic functioning of each member^7,8,9,10. Metabolomic approaches have proven especially powerful for elucidating the metabolic basis of symbiosis specificity^9,11, the bleaching response to thermal stress^7,8,12,13, disease responses¹⁴, pollution exposure responses¹⁵, photoacclimation¹⁶, and chemical signalling¹⁷ in corals, as well as aiding in biomarker discovery^18,19. Additionally, metabolomics can provide valuable confirmation of the conclusions inferred from DNA- and RNA-based techniques^9,20. There is, therefore, considerable potential for the use of metabolomics for assessing reef health and developing tools for reef conservation³, such as through the detection of metabolic biomarkers of stress^18,19 and for examining the potential of active management strategies such as nutritional subsidies²¹.

Separating the host and symbiont cells and analyzing their metabolite profiles independently, rather than together as the holobiont, can yield more information about the partner interactions, independent physiological and metabolic statuses, and potential molecular mechanisms for adaptation^{11,12,22,23,24}. Without separating the coral and Symbiodiniaceae, it is almost impossible to elucidate the contribution and metabolism of coral and/or Symbiodiniaceae independently, except for with complex genome reconstruction and metabolic modeling²⁵, but this has yet to be applied to the coral-dinoflagellate symbiosis. Furthermore, attempting to extract information about the individual metabolism of the host or algal symbiont from the metabolite profile of the holobiont can lead to misinterpretation.

For example, until recently, the presence of C18:3n-6, C18:4n-3, and C16 polyunsaturated fatty acids in extracts from coral and holobiont tissues was thought to be derived from the algal symbiont, as corals were assumed to not possess the ωx desaturases essential for the production of omega-3 (ω3) fatty acids; however, recent genomic evidence suggests that multiple cnidarians have the ability to produce ω3 PUFA de novo and further biosynthesize ω3 long-chain PUFA²⁶. Combining GC-MS with stable isotopic labeling (e.g., ¹³C-bicarbonate, NaH¹³CO₃) can be used to track the fate of photosynthetically fixed carbon through coral holobiont metabolic networks under both control conditions and in response to external stressors^27,28. However, a critical step in the tracking of ¹³C fate is the separation of the coral tissue from the algal cells-only then can the presence of a ¹³C-labeled compound in the coral host fraction be unequivocally assigned as a Symbiodiniaceae-derived metabolite translocated to the coral or a downstream product of a translocated labeled compound. This technique has demonstrated its power by challenging the long-held assumption that glycerol is the primary form in which photosynthate is translocated from symbiont to host²⁹, as well as elucidating how inter-partner nutritional flux changes during bleaching^27,28 and in response to incompatible Symbiodiniaceae species¹¹.

While the decision to separate tissues is primarily driven by the research question, the practicality, reliability, and potential metabolic impacts of this approach are important to consider. Here, we provide detailed, demonstrated methods for the extraction of metabolites from the holobiont, as well as the separate host and symbiont fractions. We compare the metabolite profiles of the host and symbiont independently and how these profiles compare to the holobiont metabolite profile.

Protocol

NOTE: The experimental design, sample collection and storage have been described in detail elsewhere^2,30,31. Permit approval for the collection of wild corals must be obtained prior to collection and experimentation. The samples here were collected from colonies of Montipora mollis (green colour-morph) imported from Batavia Coral Farms (Geraldton, WA), originally collected from a reef off the Abrohlos Islands (Western Australia; 28°52'43.3"S 114°00'17.0"E) at a depth of 1 m under the Aquaculture License AQ1643. Prior to sampling, the colonies were held in an 800 L aquarium at 35 PSU, under blue and white light at 150 µmol photons·m⁻²·s⁻¹, and at 25 °C ± 0.5 °C for 3 months. The coral fragments (~5 cm², N = 6) were snap-frozen in liquid nitrogen and stored at −80 °C until processing.

1. Preparation of the extraction solutions and equipment

1. At least 1 day prior to coral tissue removal, prepare the extraction solutions and equipment.
2. Prechill ultrapure water in clean, detergent-free glassware at 4 °C.
3. Mix 100% LC-grade methanol with a 10 µg·mL⁻¹ final concentration of appropriate internal standard(s) (e.g., ¹³C₆ sorbitol).
4. Create a 50% methanol extraction solution using half 100% LC-grade methanol and half ultrapure water. Store both methanol solutions at −20 ˚C.
   NOTE: To help prevent degradation of the metabolites, it is recommended to perform the sample processing steps in batches of five coral fragments at a time, with one additional biological (water only) blank (total samples N = 6). Once each coral sample has been separated into the two fractions (coral host tissue, henceforth "Host", and microalgal cells, henceforth "Symbiont"), the total number of samples in one processing batch will be 12.

2. Coral metabolism quenching

NOTE: The experimental design, sample collection and storage have been described in detail elsewhere^2,30,31. However, it should be noted that the time taken to quench metabolism (i.e., the time between coral sample collection and preservation) is critical to capture the original response³⁰. Preserve the sample as quickly as possible after collection to prevent changes in the metabolite composition due to sample degradation or non-target physiological responses³².

1. Place coral fragment in a sterile sample collection bag, and drain any excess seawater as much as possible. Submerge the sample in liquid nitrogen for a minimum of 30 s. Move the samples as soon as possible to a −80 ˚C freezer for storage.
   NOTE: Samples can be frozen at −80°C in light-blocked containers until processing, avoiding freeze-thaw cycles.

3. Coral tissue removal from the skeleton

NOTE: The samples should be kept on ice (4 °C) at all times to ensure they are simultaneously in liquid form whilst preventing ongoing metabolism.

1. Place a clean, sterile sample collection bag on ice so that the bag is stable and open on top of the ice in a shallow well but is not submerged in the ice. Add 10 mL of cold (4 ˚C) ultrapure water to the bag.
   NOTE: This will help to avoid repeated freeze-thawing of the coral fragment due to the cold pressurized air and surrounding ice.
2. Select a coral fragment with sterilized tweezers, and rinse with cold (4 ˚C) ultrapure water using a sterile Pasteur pipette until no seawater residue remains. Submerge the rinsed coral fragment in the bag containing the 10 mL of ultrapure water.
   NOTE: This rinse is critical to remove any residual salts that would interfere with the downstream analysis. Avoid hand contact with the water or coral fragment through the bag to maintain the sample at 4 ˚C.
3. Tape a sterile 1 mL pipette tip over the end of an air gun with electrical tape, with ~5 mm cut off the end of the tip (Figure 1A).
4. Aim the air gun onto the coral fragment with the bag half-sealed and the air flow on low-medium to gently remove the tissue by encouraging a circular movement of the water over the coral fragment.
5. After ~3 min, or when all the tissue appears to have been removed from the skeleton, turn off the air, and remove the airbrush. Completely seal the bag.
6. Squeeze all the removed coral tissue to a bottom corner of the bag. Cut off the opposite corner and gently pour the contents of the bag into a 15 mL tube on ice.

4. Optional homogenization

NOTE: Some coral species are more viscous than others, meaning the air-brushing will remove the tissue in clumps instead of in a slurry. If clumps of tissue are visible in the air-brushed homogenate, a homogenization step at 4 °C can be added for all the samples.

1. Clean a mechanical saw-tooth homogenizer twice with 4 ˚C 70% methanol and finally with 4 °C ultrapure water.
2. Homogenize the coral sample in a 15 mL tube for ~1 min until the sample is fully homogenized and no clumps are visible.
3. Clean the homogenizer as in step 4.1 between each sample. Keep the homogenization time consistent across the samples.

5. Sample collection for normalization

1. Collect a 1,000 µL aliquot from the homogenized tissue for Symbiodiniaceae cell counts, coral host tissue protein content analysis, and chlorophyll a estimation. Store at −20 ˚C until ready to analyze (section 10).

6. Optional coral host tissue-Symbiodiniaceae cell separation

1. Centrifuge the coral homogenate at 2,500 × g for 5 min at 4 °C using a refrigerated centrifuge.
   NOTE: This speed is optimal to separate the heavier Symbiodiniaceae cells, while keeping their cell walls intact, from the host tissue, which is suspended in the supernatant.
2. Remove the supernatant containing the host material, and place in a new 15 mL tube.
   NOTE: The lipids from the host tissue typically form a narrow pink/white layer on top of the symbiont cells. This layer can be collected along with the soluble host supernatant via pipetting (Figure 1B).
3. Vigorously vortex the host for exactly 1 min. Keep the algal pellet sample and host supernatant sample on ice.
4. Add 2 mL of ultrapure water at 4 °C to the algal pellet. Vigorously vortex for exactly 2 min to resuspend the pellet.
   NOTE: If individual fragments of 1 cm were not collected from the coral colony for Symbiodiniaceae genotyping, a 200 µL aliquot of the Symbiodiniaceae cell suspension can be collected here, preserved in the preferred DNA buffer solution, and stored as described in Thurber et al.³⁰ for Symbiodiniaceae genotyping (e.g., as per González-Pech et al.¹²).
5. Repeat steps 6.1-6.4 once more.
   NOTE: The reliable separation of the host and symbiont depends on the coral biomass and species, as some species can be more viscous than others. A minimum of three wash steps is recommended, but this can be increased depending on the separation success. Repeat wash steps 4.7-4.9 until no Symbiodiniaceae cells can be seen in the bottom of the host fraction and until the Symbiodiniaceae fraction is visibly free of host material (e.g., no white layer on top) (Figure 1).
6. Remove the supernatant containing the host material, and place in a new 15 mL tube.
7. Retain the symbiont pellet in the 15 mL tube.

7. Sample drying

1. Freeze either the holobiont homogenate or both the separated host and Symbiodiniaceae fractions, at −80 °C for ~120 min. Lyophilize the samples overnight with a 0.01 mbar vacuum at −85 ˚C.
   NOTE: To avoid sample loss during lyophilization, it is recommended to use a lid cut from another sterile tube, or sterile parafilm, with a ~2 mm hole punched through carefully using a sterile 25 G needle.
2. When dry, using a laboratory balance, weigh one of the following: 1) 25 mg of the holobiont; 2) 15 mg of the symbiont fraction; or 3) 30 mg of the host tissue from each sample into separate 2 mL plasticizer-free microcentrifuge tubes.
   NOTE: Critical step: The optimization of the biomass for extraction is essential to ensure that the GC-MS is not overloaded while ensuring sufficient signal. The dried coral material is very static. To avoid sample loss, use anti-static devices to eliminate electrostatic charges from the samples and weighing vessels. A simple and cost-effective alternative is to place a laundry dryer sheet under the sample tube. The dried symbiont pellet can be cut to the desired weight using a sterile blade.

8. Intracellular metabolite extractions

1. Intracellular metabolite extraction from lyophilized holobiont:
   1. Add 400 µL of 100% cold (−20 ˚C) methanol with internal standard/s (IS; ¹³C₆ sorbitol and/or ¹³C₅-¹⁵N valine, at 10 µM) to each tube.
   2. Add a small number of 710-1,180 µm acid-washed glass beads (~10 mg) to each sample. Place in a bead mill at 50 Hz for 3 min in a prechilled (−20 ˚C) bead mill insert.
   3. Add an additional 600 µL of 100% cold (−20 ˚C) methanol with ISs (¹³C₆ sorbitol and/or ¹³C₅-¹⁵N valine, at 10 µM) to each tube.
   4. Vortex to mix for 1 min. Place on a rotisserie shaker at 4 ˚C for 30 min.
2. Intracellular metabolite extraction from separated lyophilized Symbiodiniaceae cells:
   1. Add 200 µL of 100% cold (−20 ˚C) methanol with ISs (¹³C₆ sorbitol and/or ¹³C₅-¹⁵N valine, at 10 µM) to the dried Symbiodiniaceae material.
   2. Add a small number of 710-1,180 µm acid-washed glass beads (~10 mg). Place in a bead mill at 50 Hz for 3 min in a prechilled (−20 ˚C) bead mill insert.
   3. Add a further 800 µL of 100% cold (−20 ˚C) methanol with ISs, and vortex for 30 s.
3. Intracellular metabolite extraction from separated lyophilized host tissue:
   1. Add 1 mL of 100% cold (−20 ˚C) LC-grade methanol containing ISs (¹³C₆ sorbitol and/or ¹³C₅-¹⁵N valine, at 10 µM) to the dried host material.
   2. Vortex to mix for 20 s. Place in a floating tube holder in a sonication bath set at 4 ˚C for 30 min.

9. Metabolite extract purification

1. Centrifuge the samples (holobiont/host/symbiont) at 3,000 × g for 30 min at 4 °C.
2. Transfer all the supernatant to a new 2 mL microcentrifuge tube, being careful not to disturb the cell debris pellet.
   NOTE: These are the semi-polar extracts. These can be kept on ice temporarily but stored long-term at −80 ˚C in the dark.
3. To the remaining cell debris, add 1,000 µL of 50% cold (−20 ˚C) methanol. Vigorously vortex for 1 min to resuspend.
4. Centrifuge the samples at 3,000 × g for 30 min at 4 °C.
5. Collect and pool the supernatant (polar extracts) with the semi-polar extracts from the same sample.
   NOTE: The cell debris can be stored at −80 ˚C and used for protein content normalization (section 11).
6. Centrifuge the pooled extracts at 16,100 g for 15 min to remove all the precipitates, and move the supernatant to a new plasticizer-free microcentrifuge tube (2 mL).
   NOTE: The sample extracts can be stored at −80 ˚C in the dark.
7. When ready to analyze, aliquot 50 µL of each extract into a glass insert. Concentrate for 30 min at 30 ˚C using a vacuum concentrator. Repeat four more times (for 250 µL of total dried extract).
   NOTE: The dried samples can be stored at room temperature under desiccant conditions until analysis.

10. Metabolite derivatization

NOTE : A two-step online derivatization process is used for the methoximation and trimethylsilylation of the polar metabolites.

1. Add 25 µL of methoxyamine hydrochloride (30 mg/mL in pyridine) to each sample.
2. Agitate at 37 °C on an orbital shaker set at 750 rpm for 2 h.
3. Add 25 µL of N,O-bis (trimethylsilyl)trifluoroacetamide + trimethylchlorosilane to each sample.
4. Agitate again at 37 °C and 750 rpm for 1 h.
5. Allow the samples to equilibrate at room temperature for 1 h before injecting 1 µL in a 1:10 split ratio onto the GC.

11. Gas chromatography-mass spectrometry analysis

NOTE: The mass spectrometer should be tuned according to the manufacturer's recommendations using tris-(perfluorobutyl)-amine (CF43).

1. Use ultra-high purity helium as the carrier gas at a constant column flow rate of 1 mL/min.
2. Use a 30 m DB-5 column with a 1 µm film thickness and a 0.25 mm internal diameter.
3. GC oven program
   1. Set the inlet temperature to 280 °C.
   2. Start at the injection with an oven temperature of 100 °C, and hold for 4 min.
   3. Increase the temperature by 10 °C/min to 320 °C, and then hold for 11 min.
4. Mass spectrometer parameters
   1. Set the MS transfer line to 280 °C, and adjust the ion source to 200 °C.
   2. Use argon as the collision cell gas to generate the multiple reaction monitoring (MRM) product ion.
   3. Achieve metabolite detection relative to a time-segmented MRM library containing MRM targets.

12. Symbiodiniaceae cell counts, coral host tissue protein content analysis, and chlorophyll a estimation

1. Symbiodiniaceae cell counts:
   1. Take an aliquot of the coral tissue homogenate.
   2. Centrifuge the samples at 2,000 × g to pellet the algae.
   3. Remove the ~200 µL supernatant from the algal pellet, and place in a new microcentrifuge tube.
      NOTE: This will be the protein sample which will be used to normalize the data; store it at −20 ˚C before analyzing, if necessary.
   4. Resuspend the algal pellet in 1 mL of filtered sea water by gently pipetting up and down. If necessary, dilute the algal suspension to facilitate the cell counting.
   5. Conduct a cell count using a hemocytometer under a light microscope by adding 10 µL to one of the chambers. Complete 8-10 counts per sample.
      NOTE: Alternative methods for counting the algal cells can also be applied where available (e.g., flow cytometry, high-throughput confocal microscopy).
   6. Calculate the concentration of the symbiont cells (mL⁻¹), taking into account any dilution factors used.
2. Assay for the protein content
   1. Quantify the sample protein content (e.g., via Bradford's colorimetric assay, as initially described by Bradford et al.³³, or the Lowry assay^34,35, the protocol for which has been described for cnidarians elsewhere³⁶).
3. Chlorophyll a extraction
   1. Use a cell pellet of ~200, 000 cells, frozen or fresh.
   2. Transfer each algal pellet into 2 mL of dimethylformamide (DMF) in a glass scintillation vial, and incubate in the dark at 4 ˚C for 48 h.
      NOTE: DMF is toxic and carcinogenic, so the sample preparation must be completed under a fume hood that is as dark as possible and on ice. If there are <200,000 cells, use less DMF.
   3. Centrifuge for 3 min at 16,000 × g.
   4. Transfer 200 mL into a UV-96 well plate for photometric measurements. Run each sample in triplicate with DMF as the blank.
   5. Measure the absorbance at wavelengths (E) 663.8 nm, 646 nm, and 750 nm. Subtract the absorbance at 750 nm from the absorbance at both of the other wavelengths.
      NOTE: Measuring at 750 nm corrects for any scattering or turbidity in the sample.
   6. Calculate the chlorophyll a concentration (µg/mL) using equation (1):
      Chl a concentration(µg/mL) = (12.00 × E^663.8) - (3.11 × E^646.8)    (1)

13. Quantification of the cell biomass following metabolite extractions for normalization

NOTE: There are two options for the quantification of cell biomass described below: the quantification of protein related to biomass using a modified Bradford colorimetric method and the measurement of the cell debris dry weight. Either method is appropriate to use, as both offer accurate quantification of the cell biomass.

1. Protein content of the cell debris
   1. Resuspend the frozen cell debris with 1 mL of 0.2 M NaOH, and incubate the samples at 98 ˚C for 20 min.
   2. Cool the samples on ice for ~10 min, and centrifuge at 3,000 × g for 5 min at an ambient temperature.
   3. Quantify the sample protein content (e.g., via Bradford's colorimetric assay, as initially described by Bradford et al.³³ and modified by Smart et al.³⁷).
2. Measurement of cell debris dry weight
   1. Resuspend the cell debris from the intracellular metabolite extraction in double-distilled water (~10 mL).
   2. Filter the solution under a vacuum using a preweighed membrane filter (0.22 µm pore, 47 mm).
   3. Wash the tubes containing biomass twice with ultrapure water to ensure the complete transfer of biomass to the membrane filter.
   4. Remove the membrane filter containing the biomass and dry it using a microwave oven (low power; ~250 W for 20 min).
   5. Store the filter paper in a desiccator overnight. Record the dry weight of the filter paper and calculate the dry weight of the biomass by subtracting the weight of the dry membrane filter (using a clean dry membrane filter dried alongside the sample filter) from the total weight.

14. Data analysis

1. Analyze metabolite targets using metabolite databases where each target is comprised of a quantifier and qualifier MRM.
2. Visually inspect the detected metabolite targets and manually integrate them as required.
3. Use a metabolite peak area to calculate relative abundance of each sample for each group. Values are blank corrected and normalized to sample internal standard peak area, and then to sample cell debris protein content as per Smart et al.³⁷.
4. Discard metabolites with a relative standard deviation greater than 35% in all the treatment groups (N = 23 metabolites).
5. Transform the data (e.g., cube root), and mean-center them; confirm a normal distribution and homogeneity of variance.
6. Perform the data analysis (ANOVA and heatmap construction; e.g., using https://www.metaboanalyst.ca)³⁸. Cluster the samples to examine within-treatment variability using the packages “cluster”, “factoextra,” and “klustR”. Calculate the gap statistic (a method to determine the optimal number of clusters³⁹) using the "clusGap" function in R and plots using the R package "tidyverse". Perform PERMANOVAs to examine the significance in the separation between the treatment metabolite profiles (e.g., in Primer).

Results

All the data produced during this work are available in the supplementary information.

Host-symbiont separation

Figure 1: Setup and validation of the separation of coral host tissues and Symbiodiniaceae cells. (A) The air gun setup for the removal of coral...

Discussion

The separation of the host and symbiont is easily and rapidly achievable via simple centrifugation, and the results here show that separating the fractions can provide valuable information indicative of specific holobiont member contributions, which can contribute toward the functional analysis of coral health. In adult corals, lipid synthesis is primarily performed by the resident algal symbiont⁴⁰, which supplies lipids (e.g., triacylglycerol and phospholipids)⁴¹ ...

Materials

Name	Company	Catalog Number	Comments
100% LC-grade methanol	Merck	439193	LC grade essential
2 mL microcentrifuge tubes, PP	Eppendorf	30121880	Polypropylene provides high resistance to chemicals, mechanical stress and temperature extremes
2030 Shimadzu gas chromatograph	Shimadzu	GC-2030
710-1180 µm acid-washed glass beads	Merck	G1152	This size is optimal for breaking the Symbiodiniaceae cells
AOC-6000 Plus Multifunctional autosampler	Shimadzu	AOC6000
Bradford reagent	Merck	B6916	Any protein colourimetric reagent is acceptable
Compressed air gun	Ozito	6270636	Similar design acceptable. Having a fitting to fit a 1 mL tip over is critical.
DB-5 column with 0.25 mm internal diameter column and 1 µm film thickness	Agilent	122-5013
DMF	Merck	RTC000098
D-Sorbitol-6-13C and/or 13C5–15N Valine	Merck	605514/ 600148	Either or both internal standards can be added to the methanol.
Flat bottom 96-well plate	Merck	CLS3614
Glass scintillation vials	Merck	V7130	20 mL, with non-plastic seal
Immunoglogin G	Merck	56834	if not availbe, Bovine Serum Albumin is acceptable
Primer		v4
R		v4.1.2
Shimadzu LabSolutions Insight software		v3.6
Sodium Hydroxide	Merck	S5881	Pellets to make 1 M solution
tidyverse		v1.3.1	R package
TissueLyser LT	Qiagen	85600	Or similar
TQ8050NX triple quadrupole mass spectrometer	Shimadzu	GCMS-TQ8050 NX
UV-96 well plate	Greiner	M3812
Whirl-Pak sample bag	Merck	WPB01018WA	Sample collection bag; Size: big enough to house a ~5 cm coral fragment, but not too big that the water is too spread