Aug 13, 2024
Bone and tooth collagen extraction for stable isotope analysis and radiocarbon dating
- Prudence Robert1,2,3,
- Mathieu Boudin4,
- Samuel Bodé1
- 1ISOFYS - Isotope Bioscience Laboratory- Faculty of Bioscience Engineering, Ghent University;
- 2ArcheOs – Research Laboratory for Biological Anthropology, Department of Archaeology, Ghent University;
- 3A&MS- Atomic and Mass spectrometry research group, Ghent University;
- 4Royal Institute for Cultural Heritage, Brussels
Protocol Citation: Prudence Robert, Mathieu Boudin, Samuel Bodé 2024. Bone and tooth collagen extraction for stable isotope analysis and radiocarbon dating. protocols.io https://dx.doi.org/10.17504/protocols.io.36wgqnr75gk5/v1
Manuscript citation:
License: This is an open access protocol distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited
Protocol status: Working
We use this protocol and it's working
Created: March 18, 2024
Last Modified: August 13, 2024
Protocol Integer ID: 96866
Keywords: archaeology, collagen extraction, bioarchaeology, stable isotope analysis, biological anthropology, tooth collagen extraction for stable isotope analysis, dating several collagen extraction protocol, collagen extraction protocol, several collagen extraction protocol, tooth collagen extraction, stable isotope analysis, filtration of the collagen, collagen of appropriate quality, collagen, extraction, extraction protocol, radiocarbon, preserved sample
Abstract
Several collagen extraction protocols are described in the literature, but most lack detailed descriptions of the laboratory manipulations and the specific material used.
This collagen extraction protocol described here largely follows Longin's (1971) with a few changes. Shortly; The full bone is demineralised using diluted HCl (0.2 to 0.5M). This is followed by removing humic acids using a short NaOH rinse, and by solubilisation and filtration of the collagen.
This protocol aims to provide collagen of appropriate quality for both stable isotope analysis and 14C dating, at a low cost and with a decent duration. This protocol was found to also be suitable, with minor adaptations, to badly preserved samples.
Image Attribution
Prudence Robert
Guidelines
We suggest reading carefully the guidelines and every step of the protocol before starting because some steps and equipment may have to be adapted to your samples.
Numerous bone and dentine collagen extraction protocols are readily available, but they often lack detailed instructions regarding the process and the material needed. This protocol is tailored for researchers willing to implement stable isotope analysis on skeletal material but, for whom the significant amount of collagen extraction protocols published and tested make their pre-treatment choice uneasy. It can be adapted to fragile archaeological remains (e.g. poorly preserved skeletal material or fish bones).
The protocol has been designed to consider multiple factors simultaneously :
- Ensuring a reliable and satisfactory collagen purity suitable for both 14C dating and stable isotope analysis. Discussions with experts such as Mathieu Boudin from The Royal Institute of Belgium (KIK-IRPA) have influenced this aspect, drawing inspiration from Wojcieszak et al. (2020).
- Providing a minimally destructive and flexible pre-treatment approach, considering the condition, mass and age of the samples (e.g. Reducing the HCl concentration and temperature used during the demineralisation of highly degraded samples or lightweight samples).
- Emphasising affordability by utilising inexpensive materials and equipment, making the protocol accessible to researchers with limited budgets.
- Assuring a relatively short duration for the extraction process.
Materials
Equipment
- Drill tool (e.g. Dremel 3000; Proxxon)
- Heater (e.g. Oven; heating bath; -Ideally with a program function).
- Ultrasonic bath
- Freeze-dryer
- Fume hood
- Freezer
- Analytical balance
- Timer
Consumables
- Aluminium foil
- Mini-saw (Dremel; Proxxon): For most mammifère bones, we advise a maximum diameter of 22mm to avoid leaving traces in unexpected places. The width should not be over 0.7 mm to avoid the risk of losing too much sample in the form of a powder.
- Diamond-tipped drill bits (e.g. Dremel- 7103 2mm diamond)
- PP tubes with screw caps - Tubes resistant to heat (holding 13-15 ml and with a diameter of 16 mm; e.g. Böttger® - round-bottom tubes 100*16mm with screw caps ). Ezee filters fit in these sample tubes.
- Disposable pasteur pipettes.
- Ezee filters (Elkay laboratory products® - 9ml Ezee Filter separator (127-3193-000))
- pH paper
- Glass vials with well-sealing screw caps (e.g. Wheaton® - liquid scintillation vial with attached cap (DWK986541))
- Parafilm
Chemicals
- 70% ethanol solution
- 'DI-water'. It refers to deionised and/or ultrapure water (e.g. MQ water) because ultrapure water is also deionised.
- Diluted HCl solution in ultrapure water. For most samples, 0.5M is used. For more degraded samples a lower concentration is advisable (see protocol).
- 0.25M NaOH.
- "pH3 solution" made using ultrapure water and diluted HCl. pH paper is sufficiently precise for its use in this protocol.
Troubleshooting
Safety warnings
This protocol involves harmful machines and chemicals, please wear the PPE equipment (a lab coat, gloves, goggles and a mask when sampling bones especially).
Before start
400 mg of dry bone sample or half a tooth root (app. 200 mg) is usually sufficient to obtain enough collagen for 14C dating and carbon, nitrogen and sulphur isotope analysis. However, the quantity and quality of the collagen obtained at the end of this extraction protocol remain largely sample-dependent.
Sample acquisition and cleaning
Place the skeletal material to sample on a clean sheet of aluminium foil in a well-ventilated space.
With a clean mini diamond saw mounted on a rotary multifunction tool (e.g. Dremel 3000 or Proxxon), carefully cut a piece of the skeletal material aiming for the appropriate mass. The bone powder created during this step can be kept in the foil for further analysis of the material.
With a diamond drill bit, mechanically clean the bone/tooth surface by removing approximately 2mm of its outer surface.
Note
Thoroughly clean the surfaces with ethanol between each sample and replace the mini saw and the drill bit with clean ones.
The saws and drill bits can be cleaned by being soaked in an ultrasonic bath in DI-water, then in ethanol, then in water again (Repeat until the water is clear), 5 minutes each time.
Place the samples in a small container. Label the container with the sample ID on several surfaces, and then add DI-water. Sonicate the samples in the container for approximately 3 minutes, avoiding temperature increase. The water may become murky.
Note
To prevent warming the samples, avoid sonication for extended periods.
Remove the water from the container, rinse the samples 1 to 3 times and add fresh DI-water. Repeat until the water in the container remains clear. In general, two sonification steps suffice.
Let the samples dry Overnight in the container placed in a fume hood.
Weight the sample once it is dry.
Demineralisation
Insert the bone or dentin sample in a reaction tube with a screw cap*.
In a fume hood, add approximately 10 mL of the diluted HCl solution to the reaction tube, ensuring the sample is fully submerged. For well-preserved samples, a 0.5M HCl solution at room temperature can be used.
If the sample seems fragile or highly degraded, consider working at low temperatures (e.g. 4 °C ) and/or with a more dilute acid solution (e.g. 0.2M HCl solution).
Do not close the tubes tightly as CO2 forms during the demineralisation.
Note
The demineralisation might take longer when using a lower temperature and/or a lower HCl concentration.
Every two to three days, replace the HCl solution in the tube with fresh HCl. If no bubbling is observed within 30 minutes after the addition of fresh HCl at room temperature (01:00:00 if HCl is stored at 4 °C ), the sample can be considered as demineralised.
Note
Often, the bone or dentin sample, once demineralised, is flexible, transparent and/or floating in the solution.
However, it is not mandatory to wait until the sample is fully flexible or slushy, the lack of reaction between the HCl solution and the material is sufficient to consider the sample adequately demineralised.
Once the skeletal material is demineralised (i.e. no bubbling), rinse it in the tube three times with DI-water. Rinse also the edges of the tube. If the sample is fragmented, use an Ezee filter.
Note
*To use appropriately an Ezee filter, the sample tube should have a diameter of 16 mm (See 'Material' section).
If the NaOH step is not directly following, the rinsed demineralised sample can be preserved in the fridge in DI- water. If the sample is crumbly after the demineralisation, it is also possible to remove the DI-water and let the sample sit in the freezer.
Note
Different samples within the same batch are likely to demineralise at different rates. Therefore, you may want to store the samples that are ready first in the fridge or freezer while the others are still demineralising.
Removal of lipids and humic acids
Remove any DI-water from the tubes with the samples. If the sample was frozen, wait for it to thaw and remove the excess of DI-water.
Add 0.25 M NaOH solution making sure to cover the sample.
Let the sample in contact with the NaOH solution for 00:15:00 . Discard the NaOH solution from the tube using a Paster pipette or an Ezee filter if the sample consists of small fragments.
Note
The correct timing of the contact time with the NaOH is important. For this, it is advisable to space the addition of the NaOH enough (e.g. 2 minutes) between each sample and to keep track of the NaOH addition time.
Note
*To appropriately use an Ezee filter, the sample tube should have a diameter of 16 mm (See 'Material' section).
15m
Rinse the sample and the edges of the tube 3 times with DI-water.
Neutralising the NaOH
Add the diluted solution of HCl (same concentration as for the demineralisation step). Make sure the solution covers the sample.
Note
This step is important because the NaOH solution may absorb atmospheric CO2 and thus, bias the radiocarbon dates.
Let the sample and the solution react for 00:05:00 .
5m
Rinse the sample three times with DI-water.
Solubilisation of the collagen
Add the pH 3 HCl solution to the tubes. Make sure it largely covers the samples.
Note
Adding a generous amount of pH 3 HCl solution helps to prevent significant collagen loss during
.
Transfer the tubes to a heat-resistant rack.
Heat the sample in pH 3 HCL solution at 70 °C for 48:00:00 .
Note
For that step, using a programmable oven is handy.
2d
While waiting for the samples to solubilise, label the glass vials for each sample and weigh them. Also, prepare small pieces of parafilm to cover each glass vial.
Filtration
Once the samples are solubilised, filter the liquid samples by slowly pressing the Ezee filters into the sample tubes. Then transfer the filtered liquid to the pre-weighed glass vials designated for each sample.
Note
*To appropriately use an Ezee filter, the sample tube should have a diameter of 16 mm (See 'Material' section).
After the initial filtration, some of the collagen solution (pH 3 HCl solution) may remain at the bottom of the tube. To maximise recovery, add more of the pH 3 HCl solution to the tube. Then, proceed to 25 to filter the solution again.
Note
This will ensure that as much collagen as possible is collected.
Add a piece of parafilm on top of each glass vial and pierce it several times with a sharp tool (e.g. a clean toothpick for each sample).
Freezing
Put a cap above the parafilm and place the glass vials with the filtered samples in a freezer overnight.
Lyophilisation
Remove the caps from the vials and place the frozen samples in a freeze-dryer for at least 24 hours.
Note
When transferring samples between the freezer and the freeze-drier, do so quickly to prevent the samples from thawing.
Weighing and storing
Once you obtained fully dry collagen, remove the samples from the freeze-drier.
Expected result
A well-preserved collagen should look fluffy and white/beige.
Note
Ice crystals can resemble dry collagen.
It is important not to thaw a partially dried collagen sample as this might complicate the following processing.
If unsure whether the collagen is dry after some time in the freeze-drier, let the sample in for a longer time.
If unsure whether the sample is fully dehydrated after removing it from the freeze-drier, quickly weigh the collagen and place it back in the freeze-drier for an additional time, e.g., 12 hours. Then, weigh it again. If the sample was not fully dehydrated, the mass must have decreased. Repeat until the mass of the collagen is constant.
After removing the parafilm, place the caps on the glass vials to prevent the collagen from taking up humidity.
Note
Freeze-dried collagen is highly hygroscopic.
Weight the dry collagen in the glass vials and calculate the collagen yield.
Note
A collagen yield of ≥ 1 % is generally considered the threshold for determining well-preserved collagen in skeletal remains (van Klinken, 1999).
Citation
LINK
Storage
The collagen can be stored in a constantly dry environment (e.g. a freezer).
Protocol references
Brown TA, Nelson DE, Vogel JS, Southon JR (1988) Improved Collagen Extraction by Modified Longin Method. Radiocarbon 30:171–177. https://doi.org/10.1017/S0033822200044118
Buckley M, Kansa SW (2011) Collagen fingerprinting of archaeological bone and teeth remains from Domuztepe, South Eastern Turkey. Archaeol Anthropol Sci 3:271–280. https://doi.org/10.1007/s12520-011-0066-z
Collins MJ, Nielsen–Marsh CM, Hiller J, et al (2002) The survival of organic matter in bone: a review. Archaeometry 44:383–394. https://doi.org/10.1111/1475-4754.t01-1-00071
Fülöp R-H, Heinze S, John S, Rethemeyer J (2013) Ultrafiltration of Bone Samples is Neither the Problem nor the Solution. Radiocarbon 55:491–500. https://doi.org/10.1017/S0033822200057623
Jorkov MLS, Heinemeier J, Lynnerup N (2007) Evaluating bone collagen extraction methods for stable isotope analysis in dietary studies. Journal of Archaeological Science 34:1824–1829. https://doi.org/10.1016/j.jas.2006.12.020
Longin R (1971) New method of collagen extraction for radiocarbon dating. Nature 230: 241–242.
Pestle WJ (2010) Chemical, elemental, and isotopic effects of acid concentration and treatment duration on ancient bone collagen: an exploratory study. Journal of Archaeological Science 37:3124–3128. https://doi.org/10.1016/j.jas.2010.07.013
Schoeninger MJ, Moore K (1992) Bone stable isotope studies in archaeology. J World Prehist 6:247–296. https://doi.org/10.1007/BF00975551
Sealy J, Johnson M, Richards M, Nehlich O (2014) Comparison of two methods of extracting bone collagen for stable carbon and nitrogen isotope analysis: comparing whole bone demineralization with gelatinization and ultrafiltration. Journal of Archaeological Science 47:64–69. https://doi.org/10.1016/j.jas.2014.04.011
Talamo S, Fewlass H, Maria R, Jaouen K (2021) “Here we go again”: the inspection of collagen extraction protocols for 14 C dating and palaeodietary analysis. STAR: Science & Technology of Archaeological Research 7:62–77. https://doi.org/10.1080/20548923.2021.1944479
Wojcieszak M, Van den Brande T, Ligovich G, Boudin M (2020) Pretreatment Protocols Performed At The Royal Institute For Cultural Heritage (Rich) Prior To Ams 14 C Measurements. Radiocarbon 62:e14–e24. https://doi.org/10.1017/RDC.2020.641
Citations
Step 32
G.J. van Klinken. Bone Collagen Quality Indicators for Palaeodietary and Radiocarbon Measurements
https://doi.org/10.1006/jasc.1998.0385