Soils and sediments

Required materials:

• Gloves (powder-free, nitrile preferred, latex also possible)

• Sampling tool (e.g., a stainless steel spatula with one straight and one rounded edge)

• Sterilising reagent (e.g., acetone, isopropyl alcohol, dichloromethane)

• Unused, sterile glass tubes of various sizes

  • Alternative: clean aluminum foil and unused plastic bags

• Permanent marker and transluscent tape 

• Protective personal equipment (e.g., protective clothing, glasses, and mask - particularly important when cleaning with reagent)

• Optional: small balance to weigh sample in the field

Before sampling

Before sampling, prepare glass vials by labelling them with pertinent information (e.g., unit, stratigraphic and archaeological layer, feature, etc.) and placing translucent tape over the label to avoid smudging. Label caps as well. This can also be done immediately after sampling, as the location may slightly change if osbtacles (e.g., rocks) are encountered. 

Next, clean your sampling tools. As stated in the materials, a stainless steel spatula with one straight and one rounded edge is an ideal tool as this allows you to remove some surface soil/sediment with the square edge and collect the sample with the unused rounded edge. While a (non-rusty) trowel may also be used, it will be more difficult to transfer the soil/sediment to the glass vial. Clean aluminum foil may be used as a funnel to these ends.

Sampling tools should be cleaned using one of the above reagents (e.g., acetone), while wearing protective gear, in a well-ventilated area before and between each sample. Care must be taken to limit exposure to these reagents, including contact with exposed skin and inhalation. 

Sampling

If sampling a profile, first remove approximately 0.5 - 2 cm of the surface with the clean square edge of the spatula to avoid surface contamination (e.g., from excavators, organisms, carbonate formation in humid environments such as caves) in your sample. Using the rounded edge, collect the soil/sediment of interest and transfer to the glass vial. If an aluminum foil pouch (in an unused plastic bag) is being used, the sample should be placed in contact with the matte side of the foil, as the shiny side may have additional lipids or molecules from the manufacturing process. 

Physical handling of soils and sediments should be as limited as possible, and should not occur without gloves to avoid the addition of hand lipids to the sample. Be careful to not mix the soil/sediment of interest with other soils/sediments, and pay particular attention to contamination from material higher in the profile that comes loose with sampling. Any mixed soil should not be transfered to the vial (or aluminum foil pouch).

Be sure your sampling goals are clear, by asking questions such as the following: 

• Are you sampling at fixed depths/increments, or following the natural topography of the layer(s)? 

• How will your sampling strategy differ if sampling the profile vs. an exposed excavation surface? 

• How will you record these different types of spatial information?

• How will your sampling strategy be modified if you encounter an obstacle (e.g., rocks, archaeological material) in the targeted area? 

• Are you gathering enough material for all the analyses that may be applied to your samples?

• Are your controls truly representive of the sterile substrate? Have you sampled controls in multiple areas across the site?

• Are the sampled areas stratigraphically intact, or has there been some post-depositional reworking of the soils/sediments?

Sample Requirements

The amount of soil/sediment required for lipidomic and DNA analyses are as follows, although variation by laboratory, extraction protocol, preservation, and organic matter abundance can be expected.

Lipids (via GC-MS):

• Loose sediment:  2 – 5 grams (1)

• *Impregnated blocks/thin sections: ~ 200 mg  (2)

DNA:

• Loose sediment: ~ 40 – 200 mg 

  • • 38 – 160 mg (3)

    • 29 – 191 mg (4)

    • 41.5 – 80.5 (5)

• *Impregnated blocks/thin sections: ~ 50 mg (6)

Accounting for sample loss and/or low abundance or preservation, at least 5 grams of soil/sediment should be gathered for lipidomic analyses of loose sediment, while at least 200 mg should be gathered for genetic analysis. 

*Note that work on impregnated micromorphology blocks or thin sections is relatively new and still being developed, particularly for lipidomic analysis. These values are only indicative of the published literature.

Sources:

1. Jambrina-Enríquez et al., 2019 

2. Rodríguez de Vera et al., 2020 

3. Slon et al., 2017 

4. Zavala et al., 2021 

5. Slon et al., 2022 

6. Massilani et al., 2022 

Once gathered, soil and sediment samples should be stored in a cold environment, ideally at -20 °C, to halt microbial action. The precise temperature needed varies with analysis, such that a standard cold room (~4 °C) may be appropriate, or a colder freezer (e.g. -40 to -80 °C) may be needed. Samples will be stable for years at or below -20 °C.

Samples should be stored in glass vials (alternatively but less preferred: aluminum foil packets within unused plastic bags), and care should be taken to ensure the labels will remain legible after freezing. As freeze-thaw cycles should be avoided, it may be ideal to preform dry and homogenise samples (see below) prior to freezing. 

Required materials:

• Oven 

• Vessels for drying (e.g., clean glass trays, ceramic vessels, or aluminum foil tins)

  • Note: Plastic vessels can also be used, though the potential for contamination from plasticisers must be acknowledged

  • Note: Vessels can be cleaned with the reagents mentioned above, and glass vessels or ceramic vessels can be pre-combusted to limit contamination

• Mortar and pestle (agate or ceramic)

• 200 μm stainless steel sieve and receptacle

• Cleaning reagent (e.g., dichloromethane)

• Laboratory paper towel

• Wipes that do not leave fibers or residue (e.g., Kimtech wipes)

• Aluminum foil

• Optional: compressed air

Drying your samples

Drying is necessary to standardize residual moisture across samples and limit algal growth and microbial action. However, rushed drying at higher temperatures can damage organic matter in the sample and/or create false signals of heat-alteration. Oven drying is thus best achieved at 40 °C for 24 - 48 hours, though additional time may be required for samples with considerable residual moisture. Freeze-drying is preferred, according the manufacturer's instructions. To ensure homogenous drying, samples should be dried as flat as possible and soil aggregates should be broken up. 

Following drying, homogenise samples (see below) or transfer them back to their original (or a new) glass vial using clean aluminum foil or a glass funnel. 

Note: if residual moisture in the glass vials is a concern, samples can be dried for an additional hour in their glass vials. 

Homogenising your samples

Homogenising samples prior to any extraction or analysis is necessary to ensure the compounds and values obtained are representative of the sample as a whole. Samples may first be sieved to larger sizes (e.g., 2 mm,  500 μm) to remove larger rocks or archaeological material, particularly if working with bulk sediment samples. For smaller samples, a 200 μm sieve is ideal.

Homogenise soil/sediment samples by hand with a mortar and pestle, or using a ball-and-mill machine following the manufacturer's instructions*. Transfer the homogenised sample to the sieve and gently shake to homogenise. The fraction greater than 200 μm may be re-homogenised with the motar and pestle, several times, to minimise sample loss. Shake to homogenise, and collect the fraction smaller than 200 μm for organic residue analysis. 

The mortar and pestle, and sieve, must be thoroughly cleaned before and after each sample. First, use compressed air to remove particulate matter - this is particularly important for the sieve. Next, clean with the reagent of choice, following the manufacturer's safety guidelines (e.g., under a fume hood if using dichloromethane). Ensure no residue is left behind - of previous samples, reagents, or paper towel fibers. A Kimtech wipe can be used last to ensure no paper fibers remain. Avoid any contact of paper towel with the sieve as the fibers will become embedded in the mesh. 

Allow the sieve to dry completely (e.g., under a fume hood) before attempting another sample, as any moisture in the mesh will trap the sample and lead to sample loss. 

*It may be useful to sieve the material following homogenisation with a ball-and-mill machine to have an estimate of particle size. 

The identification of archaeological biomolecules – namely, DNA and lipids – in soils and sediments has enabled novel discoveries. 

For instance, the pioneering work of Slon et al., 2017 recovered Neandertal and Densiovan DNA from several Pleistocene archaeological sites, even in areas where no skeletal remains were found. DNA in sediment can also be employed to study faunal and hominin turnovers, as at Denisova Cave (Zavala et al., 2021). The recent work of Slon et al., 2022 found DNA in Levantine Palaeolithic sediments, in a context which surpasses the theoretical limit of DNA preservation in such a warm environment, suggesting that DNA may persist (given the appropriate conditions) longer than previously thought. Lastly, high-quality DNA may also be recovered from resin-impregnated micromorphology blocks, as in the work of Massilani et al., 2022, enabling the reconstruction of taxonomic composition and the linking of genetic information to archaeological and ecological records on a microstratigraphic scale. 

The study of lipids in soils and sediments has a deep research history. Like DNA, lipids are ubiquitous in the natural world and their presence in soils and sediments largely speaks to the environmental and ecological conditions at the time of pedogenesis (soil formation). Particular lipid compounds may also serve as biomarkers of human activity, such as cooking of animal meats or producing tars and resins, and for this reason their use in the study of hearths and burned organic matter continues to grow. The development of micro-contextual lipid analysis via the sampling of impregnated micromorphology blocks is also underway (Rodríguez de Vera et al., 2020), though the resin constituents pose a great challenge here both for lipid analysis and stable carbon isotopic analysis. Further limitations regarding the study of organic matter in soils and sediments are discussed below.

There are numerous methodological and interpretive challenges that arise when working with soils and sediments. 

Detailed information regarding the spatial origin of the sample and it's relationship to surrounding archaeological material is the first line of defense to counter some of these challenges. Variation undoubtedly occurs at a microstratigraphic scale, and such variation, as well as the sedimentation rates of the sampled area and possible post-depositional reworking of the soils/sediments, must be taken into consideration before proposing any interpretations. 

Basic characterisation of the sampled soil/sediment may also prove useful, such as minerology, granulometry, pH (to explore the impact of the chemistry of the depositional environment on organic matter preservation), carbon and nitrogen elemental analysis, and so forth. These analyses may aid in the interpretation of outlier data, and may also be employed as pre-screening methods to identify samples of priority interest. 

Given the great abundance of diverse organic matter in soils and sediments, the choice of methodology should also be catered to the specific research questions being pursued. If targeting anthropogenic biomarkers, the applied lipid extraction protocol will likely differ from a study primarily concerned with palaeoenvironmental reconstruction. 

Leaching and the mobility of organic compounds in soils and sediments is another significant concern that must be taken into consideration. Burned organic matter, such as charred residues and charcoal, are mobile. They can move both vertically and laterally in the profile under the influence of rain, flooding, percolating groundwater, freeze-thaw cycles, and weathering. Their molecular counterparts are similarly mobile. Therefore, how certain can you be that the lipids recovered from a hearth substrate are actually concurrent with the burning event(s)? Is it possible that they originate from somewhere else (e.g., higher) in the profile, and were deposited at this layer from leaching? Systematic sampling below features of interest (e.g., at 2, 4, 6, 8 and 10 cm depth) is one way to explore the effect of leaching on such deposits. 

Carter, M.R., Gregorich, E.G. (Eds.), 2007. Soil Sampling and Methods of Analysis, 2nd ed. CRC Press, Boca Raton. https://doi.org/10.1201/9781420005271

Diefendorf, A.F., Freimuth, E.J., 2017. Extracting the most from terrestrial plant-derived n-alkyl lipids and their carbon isotopes from the sedimentary record: A review. Org. Geochem. 103, 1–21. https://doi.org/10.1016/j.orggeochem.2016.10.016

Harrault, L., Milek, K., Huguet, A., Anquetil, C., Anderson, D.G., 2022. Preserved lipid signatures in palaeosols help to distinguish the impacts of palaeoclimate and indigenous peoples on palaeovegetation in northwest Siberia. Org. Geochem. 167, 104407. https://doi.org/10.1016/j.orggeochem.2022.104407

Jambrina-Enríquez, M., Herrera-Herrera, A.V., Rodríguez de Vera, C., Leierer, L., Connolly, R., Mallol, C., 2019. n-Alkyl nitriles and compound-specific carbon isotope analysis of lipid combustion residues from Neanderthal and experimental hearths: Identifying sources of organic compounds and combustion temperatures. Quat. Sci. Rev. 222, 105899. https://doi.org/10.1016/j.quascirev.2019.105899

Leierer, L., Jambrina-Enríquez, M., Herrera-Herrera, A.V., Connolly, R., Hernández, C.M., Galván, B., Mallol, C., 2019. Insights into the timing, intensity and natural setting of Neanderthal occupation from the geoarchaeological study of combustion structures: A micromorphological and biomarker investigation of El Salt, unit Xb, Alcoy, Spain. PLoS One 14, e0214955.

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Slon, V., Clark, J.L., Friesem, D.E., Orbach, M., Porat, N., Meyer, M., Kandel, A.W., Shimelmitz, R., 2022. Extended longevity of DNA preservation in Levantine Paleolithic sediments, Sefunim Cave, Israel. Sci Rep 12, 14528. https://doi.org/10.1038/s41598-022-17399-2

Slon, V., Hopfe, C., Weiß, C.L., Mafessoni, F., de la Rasilla, M., Lalueza-Fox, C., Rosas, A., Soressi, M., Knul, M.V., Miller, R., Stewart, J.R., Derevianko, A.P., Jacobs, Z., Li, B., Roberts, R.G., Shunkov, M.V., de Lumley, H., Perrenoud, C., Gušić, I., Kućan, Ž., Rudan, P., Aximu-Petri, A., Essel, E., Nagel, S., Nickel, B., Schmidt, A., Prüfer, K., Kelso, J., Burbano, H.A., Pääbo, S., Meyer, M., 2017. Neandertal and Denisovan DNA from Pleistocene sediments. Science 356, 605. https://doi.org/10.1126/science.aam9695

Thomas, C.L., Jansen, B., van Loon, E.E., Wiesenberg, G.L.B., 2021. Transformation of n-alkanes from plant to soil: a review. SOIL 7, 785–809. https://doi.org/10.5194/soil-7-785-2021

Zavala, E.I., Jacobs, Z., Vernot, B., Shunkov, M.V., Kozlikin, M.B., Derevianko, A.P., Essel, E., de Fillipo, C., Nagel, S., Richter, J., Romagné, F., Schmidt, A., Li, B., O’Gorman, K., Slon, V., Kelso, J., Pääbo, S., Roberts, R.G., Meyer, M., 2021. Pleistocene sediment DNA reveals hominin and faunal turnovers at Denisova Cave. Nature 595, 399–403. https://doi.org/10.1038/s41586-021-03675-0