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Community-shared SOPs


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Page last modified on: 2026-October-14

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This page lists experimental SOPs from external sources and in-house protocols as shared by NFDI4Microbiota members. More community-shared protocols can be found on NFDI4Microbiota’s protocols.io workspace

The protocols from the International Human Microbiome Consortium IHMS and the Earth Microbiome Project EMP are well-established and experimentally validated for their given fields, human and earth microbiome research respectively.

Protocols from other sources, especially if provided by NFDI4Microbiota community members, have been successfully implemented and shared by individuals working in their given fields.

List of SOPs

Category   Title      Original Provider       Source Links
DNA Extraction DNA Extraction: Shortened Godon protocol with optional enzymatic lysis University Hospital of RWTH Aachen CRC1832 Gut-liver-axis SOP, NFDI4Microbiota SOP
DNA Extraction DNA Extraction Protocol Earth Microbiome DNA Extraction Protocol
DNA Extraction Standard operating procedure for fecal samples DNA Extraction, protocol H International Human Microbiome Standards (IHMS) SOP 07
DNA Extraction Standard operating procedure for fecal samples DNA Extraction, protocol Q International Human Microbiome Standards (IHMS) SOP 06
DNA Sequencing 16S Illumina Amplicon Protocol Earth Microbiome 16S Illumina Amplicon Protocol
DNA Sequencing 18S Illumina Amplicon Protocol Earth Microbiome 16S Illumina Amplicon Protocol
DNA Sequencing ITS Illumina Amplicon Protocol Earth Microbiome ITS Illumina Amplicon Protocol
DNA Sequencing 16S rRNA gene amplicon sequencing of synovial fluid CoSOP (Community Collection of Validation Experiments informing Preanalytic SOPs), Klaus Neuhaus /CoSOP/Synovial
Lipid Extraction Lipid and fatty acid extraction protocol from biological samples CLOSED ACCESS Bird et al (2011).Analytical Chemistry, 83(17), 6648–6657. ,  NFDI4Microbiota SOP
Medium Preparation Media for freezing homogenised faecal samples for re-cultivation University Hospital of RWTH Aachen CRC1832 Gut-liver-axis SOP ,  NFDI4Microbiota SOP
Metabolite Extraction Metabolite extraction from adherent mammalian cells CLOSED ACCESS Yuan et al. (2021). Nat Protoc 7, 872–881 (2012) ,  NFDI4Microbiota SOP
Metabolite Extraction Metabolite extraction from plant tissue CLOSED ACCESS Novák et al. (2016). Current Protocols in Plant Biology, 1(2), 419–430. ,  NFDI4Microbiota SOP
Metabolite Extraction Comparison of fecal samples before and after metabolite extraction for 16S rRNA gene amplicon sequencing CoSOP (Community Collection of Validation Experiments informing Preanalytic SOPs), Klaus Neuhaus /CoSOP/AfterMetabEx
Protein Purification Proteomics TMT labelled SP3 method EMBL Protein purification. (2022). In EMBL. ,  NFDI4Microbiota SOP
Sample Collection Skin swab: Skin samples were collected using Catch-All-Swabs (Epicentre Technologies, Wisconsin, USA) shortly after birth but before the skin-to-skin contact with the infant, by swabbing the upper area of the maternal breast (intermammary cleft). After pre-moistening with 2 ml SCF-1 buffer (50 mM Tris buffer, pH 7.6, 1mM EDTA, pH 8.0, and 0.5% Tween-20) (Human Microbiome Project Consortium, 2012) contained in a 15 ml sterile screw top collection tube (Sarstedt, Nümbrecht, Germany), the swab head was rubbed back and forth for approximately 30 seconds over the area (repeating twice) before the swab was returned to the buffered solution. The lower part of the swab was broken to ensure closure of the tube (see below). After sampling, deterge the sampled area with a clean pad of cotton and water.   (in-house protocol provided by Pamela Ferretti)
Sample Collection Tongue dorsum swabs: Skin samples were collected using Catch-All-Swabs (Epicentre Technologies, Wisconsin, USA). Samples were collected by rubbing a swab on the central area of the back of the tongue for approximately 5 seconds. The swab head was then placed in a 15 ml collection tube containing 2 ml SCF-1 buffer (see photos above). Samples were collected while wearing protective disposable gloves to avoid skin contamination.   (in-house protocol provided by Pamela Ferretti)
Sample Collection Vaginal swabs: Skin samples were collected using Catch-All-Swabs (Epicentre Technologies, Wisconsin, USA). The swab was rubbed 5 times, with a circular motion, in the vaginal introitus and then the swab head was placed in a 15 ml sterile screw top collection tube containing 2 ml SCF-1 buffer.   (in-house protocol provided by Pamela Ferretti)
Sample Collection Faeces: Standard operating procedure for fecal samples frozen preserved self-Collection, laboratory analysis handled within 24 hours to 7 days International Human Microbiome Standards (IHMS) SOP 04
Sample Collection Faeces: Standard operating procedure for fecal samples identification International Human Microbiome Standards (IHMS) SOP 01
Sample Collection Faeces: Standard operating procedure for fecal samples preserved in stabilizing solution self-Collection, laboratory analysis handled within 24 hours to 7 days International Human Microbiome Standards (IHMS) SOP 05
Sample Collection Faeces: Standard operating procedure for fecal samples self-Collection, laboratory analysis handled within 4 hours International Human Microbiome Standards (IHMS) SOP 02
Sample Collection Faeces: Standard operating procedure for fecal samples self-Collection, laboratory analysis handled within 4 to 24 hours International Human Microbiome Standards (IHMS) SOP 03
Sample Collection & Storage Activated sludge from wastewater: collected samples stored at 4°C for transport, within 6 h centrifuged at 6 500 g for 15 min at 4°C, supernatant removed and pellets stored at -20°C   (Morin et al., 2020)
Sample Collection & Storage Antarctic ticks: collected from rocks, placed in RNALater and stored at -80°C within 4-8 h of collection   (Wille et al., 2020)
Sample Collection & Storage Atlantic salmon: dissection samples placed in RNAlater and stored at -80°C   (Karlsen et al., 2022)
Sample Collection & Storage Faeces/gut content : flash freeze in liquid nitrogen and store at -80°C (dissolve in Stool DNA Stabilizer (INVITEK Molecular) while thawing; or if you have more time dissolve in Stool DNA Stabilizer right after collection and normally freeze and store at -80°C)   (in-house protocol provided by Johannes Masson)
Sample Collection & Storage Frog swabbing : rinse caught frog with 50 ml sterilised water to remove transient bacteria, swab entire skin surface with a sterile Dacron swab for 30 sec, place swab in microcentrifuge tube containing PBS and store on ice, then transfer to -80°C storage   (Ellison et al., 2021)
Sample Collection & Storage Frog toes: stored in sterile 1.5 ml tubes at -80°C   (Boyle et al., 2004)
Sample Collection & Storage Fruit fly larvae: sterilise larvae with a solution of 0.5 % Tween 80, 0.5 % sodium hypochlorite and 80 % ethanol for 30 sec, rinse them 3 times in PBS for 30 sec, use sterile pestle to crush larvae and store in BHI containing 20 % glycerol at -80°C   (Majumder et al., 2020)
Sample Collection & Storage Hooded seal brain: whole samples are placed in 4°C glucose artificial cerebrospinal fluid (aCSF) saturated with 95 % O2 - 5 % CO2. After experiment (induced normoxia and hypoxia), tissues were stored in RNAlater stabilisation fluid (Life Technologies, USA) and kept at 4°C.   (in-house protocol provided by Justine Vandedorpe)
Sample Collection & Storage Invertebrates: whole samples are stored in 80 % ethanol, in fridge for short-term storage, and in -20°C freezer for long-term storage   (in-house protocol provided by Justine Vandedorpe)
Sample Collection & Storage Leaves: surface sterilise: rinse for 5 sec 95% ethanol, 30 sec in 70% ethanol, 30 sec in 10% bleach, followed by three 2-minute washes in sterile water, air-dry for 5 min, then add to 2 ml tube containing 350 µL 10 mM MgSO4 and a sterile 5 mm steel ball and homogenise using a TissueLyser (QIAGEN) run at max speed (50 Hz) for up to 60 sec, then flash freeze in liquid nitrogen and store at -80°C   (Humphrey & Whiteman, 2020)
Sample Collection & Storage Maternal Milk (with glycerol): Minimum of 2ml of milk was self-collected by the mothers in sterile falcon tubes with added 20% glycerol while wearing disposable gloves. Samples without were stored at -20C for cultivation experiments. Note that for colostrum and other milk samples within the first week of life the 2ml min quantity might not be possible. In that case, the mother collected whatever quantity was possible in 10 min timespan.   (in-house protocol provided by Pamela Ferretti)
Sample Collection & Storage Maternal Milk (without glycerol) : Minimum of 2ml of milk was self-collected by the mothers in sterile falcon tubes while wearing disposable gloves. No buffer was added. Samples were stored at -4C right after collection and moved to -80C within a week. Note that for colostrum and other milk samples within the first week of life the 2ml min quantity might not be possible. In that case, the mother collected whatever quantity was possible in 10 min timespan.   (in-house protocol provided by Pamela Ferretti)
Sample Collection & Storage Mosquitos: human bait adult catching method, pool 3-5 mosquitos, store at -80°C   (Ali et al., 2021)
Sample Collection & Storage Oral swabs: after collection swab is immersed in 10:1 Tris-EDTA buffer immediately and stored at −80 °C   (Gao et al., 2020)
Sample Collection & Storage Penguin cloacal swaps: placed in tubes containing viral transport medium (brain heart infusion [BHI] broth-based medium [Oxoid] with 0.3 mg/ml penicillin, 5 mg/ml streptomycin, 0.1 mg/ml gentamicin, and 2.5 μg/ml amphotericin B), kept on ice for 4 hours, then stored at -80°C   (Wille et al., 2020)
Sample Collection & Storage Pooled tear collection: centrifuged at 270 g for 3 min, supernatant stored in 1.5 ml microcentrifuge tubes at -80°C   (Willis et al., 2020)
Sample Collection & Storage Prokaryotes in sea water: water sampled without filtration, 2% final concentration glutaraldehyde added, stored at 4°C for 15 min in the dark, flash-frozen in liquid nitrogen and stored at -80°C   (Baltar et al., 2018)
Sample Collection & Storage Saliva: passive drooling procedure in sterile 50 ml screw top tubes (approximately 3 ml of saliva per sample). About 30 minutes before collection, all volunteers performed a mouth rinse with drinking water. Samples were vortexed before 2 ml of each was centrifuged at 10,000 × g for 10 minutes to collected the bacterial pellets. Total DNA was extracted using the PowerSoil DNA Isolation Kit (MoBio), according to manufacturer’s instructions. DNA was stored at −20 °C until used.   (in-house protocol provided by Pamela Ferretti)
Sample Collection & Storage Seawater: filtered through a 0.2 µm filter, filter stored in 2 ml tubes containing lysis buffer (EDTA 40 mmol l−1 pH 8.0, Tris-HCl 50 mmol l−1 pH 8.0, sucrose 0.75 mol l−1) flash frozen in liquid nitrogen, filtered & unfiltered water stored in 15 ml falcons at -20°C   (Michoud et al., 2021)
Sample Collection & Storage Seawater: filtered through a 200 µm, then a 20 µm mesh,  then vacuum filtered through a 0.8 µm filter, then a 0.2 µm filter, flash frozen in liquid nitrogen & stored at -80°C (all done with ~4h; water at 4°C during processing)   (Acinas et al., 2021)
Sample Collection & Storage Sponges: washed three times with filter-sterilised natural seawater, one fragment placed in 70% ethanol, one fragment placed in RNALater, stored at -20°C   (Ruocco et al., 2021)
Sample Collection & Storage Stool samples: samples were self-collected using collection tubes specific for faecal material (Sarstedt, Nümbrecht, Germany). Toilet paper was placed at the bottom of the WC, to prevent stool samples from sinking and getting contaminated. The collection was performed at the upper part of the feces, the one not in contact with the toilet paper, WC walls or other material. Subjects were instructed, whenever possible, to urinate before evacuating the stools. Samples were collected in variable quantity, depending on availability (meconium is usually only one spoon). Once collected the tube was placed at -4C as soon as possible, and then transferred to -80C within a week.   (in-house protocol provided by Pamela Ferretti)
Sample Collection & Storage Swedish cervical biobank: stored in liquid at -25°C, using liquid-based cytology cells suspended and fixed in Thinprep (TP) containing 20 ml PreservCyt (ThinPrep Hologic, Boxborough, MA). 4 ml of liquid-based cytology sample from the bottom of the patient tube is transferred into a conical tube and allowed to sediment for 30 min. 300 µl of the sediment are transferred to a cryotube and stored at -25°C   (Perskvist et al., 2013)
Sample Collection & Storage Turkey Trachea swabs: placed in 1 ml PBS, then stored at -20°C   (Kursa et al., 2021)
Sample Collection & Storage Yoghurt: after purchase, samples were placed immediately at 4°C for transport and within 12 h stared at -80°C   (Islam et al., 2021)
Sample Collection & Storage Urine: Effect of storage time and urine conditioning buffer on urinary microbiota composition CoSOP (Community Collection of Validation Experiments informing Preanalytic SOPs), Lena M. Biehl /CoSOP/Effect of Storage time on Urine samples
Sample Processing Biobank of human stool samples – sample processing University Hospital of RWTH Aachen CRC1832 Gut-liver-axis SOP ,  NFDI4Microbiota SOP
Sample Processing, DNA Extraction & Sequencing Comparison of non-enzymatic bead beating devices on human stool samples for DNA isolation and subsequent 16S rRNA gene sequencing CoSOP (Community Collection of Validation Experiments informing Preanalytic SOPs), Klaus Neuhaus /CoSOP/BeadBeater
Sample Purification Viral purification from bacterial culture EMBL protocols.io ,  NFDI4Microbiota SOP
Workflow CoSOP: Community Collection of Validation Experiments informing Preanalytic SOPs for wetlab researchers and data analysis NFDI4Microbiota Flex Funds Project, Lena M. Biehl, Klaus Neuhaus /CoSOP

List of SOPs with host-body site and environmental ontology tags

When depositing data to public repositories, such as sequencing data to the ENA, you will notice that some metadata fields request or require ontology terms when providing host-body-site specific data or the sample provenance and environmental information.

In this list we tried to restructure the above table to give more context. Here is a preview, the full table (last update: 2026-04) can be found as csv.

Category Title & Description Provider Primary Source Link Ontology Tags *Biological sample or Environmental medium/ env_medium Host/ host_body_site
DNA Extraction DNA Extraction Protocol Earth Microbiome DNA Extraction Protocol ENVO:environmental biome; Sample:soil/water; Host:environmental microbiome soil ENVO:00001998| water ENVO:00002006 | sediment ENVO:00002007 biome ENVO:00000428
DNA Extraction DNA extraction: Shortened Godon protocol with optional enzymatic lysis NFDI4Microbiota community member CRC1832 Gut-liver-axis SOP ENVO:fecal material; Host:Homo sapiens; Sample:stool; OBI:enzymatic lysis Fecal material ENVO:00002003 human; Homo sapiens ncbitaxon:9606
DNA Extraction Standard operating procedure for fecal samples DNA Extraction, protocol H International Human Microbiome Standards (IHMS) SOP 07 ENVO:fecal material; Host:Homo sapiens; Sample:stool Fecal material ENVO:00002003 human; Homo sapiens ncbitaxon:9606

References

  1. Morin, L., Goubet, A., Madigou, C., Pernelle, J.-J., Palmier, K., Labadie, K., Lemainque, A., Michot, O., Astoul, L., Barbier, P., & et al. (2020). Colonization Kinetics and implantation follow-up of the sewage microbiome in an urban wastewater treatment plant. Scientific Reports, 10(1). https://doi.org/10.1038/s41598-020-68496-z
  2. Wille, M., Harvey, E., Shi, M., Gonzalez-Acuña, D., Holmes, E. C., & Hurt, A. C. (2020). Sustained RNA virome diversity in Antarctic penguins and their ticks. The ISME Journal, 14(7), 1768–1782. https://doi.org/10.1038/s41396-020-0643-1
  3. Karlsen, C., Tzimorotas, D., Robertsen, E. M., Kirste, K. H., Bogevik, A. S., & Rud, I. (2022). Feed microbiome: Confounding factor affecting Fish Gut Microbiome Studies. ISME Communications, 2(1). https://doi.org/10.1038/s43705-022-00096-6
  4. Ellison, S., Knapp, R., & Vredenburg, V. (2021). Longitudinal patterns in the skin microbiome of wild, individually marked frogs from the Sierra Nevada, California. ISME Communications, 1(1). https://doi.org/10.1038/s43705-021-00047-7
  5. Boyle, D. G., Boyle, D. B., Olsen, V., Morgan, J. A. T., & Hyatt, A. D. (2004). Rapid quantitative detection of chytridiomycosis (batrachochytrium dendrobatidis) in amphibian samples using real-time Taqman PCR assay. Diseases of Aquatic Organisms, 60, 141–148. https://doi.org/10.3354/dao060141
  6. Majumder, R., Sutcliffe, B., Taylor, P. W., & Chapman, T. A. (2020). Fruit host-dependent fungal communities in the microbiome of wild Queensland fruit fly larvae. Scientific Reports, 10(1). https://doi.org/10.1038/s41598-020-73649-1
  7. Humphrey, P. T., & Whiteman, N. K. (2020). Insect herbivory reshapes a native leaf microbiome. Nature Ecology &Amp; Evolution, 4(2), 221–229. https://doi.org/10.1038/s41559-019-1085-x
  8. Ali, R., Jayaraj, J., Mohammed, A., Chinnaraja, C., Carrington, C. V., Severson, D. W., & Ramsubhag, A. (2021). Characterization of the virome associated with Haemagogus Mosquitoes in Trinidad, West Indies. Scientific Reports, 11(1). https://doi.org/10.1038/s41598-021-95842-6
  9. Gao, L., Kang, M., Zhang, M. J., Reza Sailani, M., Kuraji, R., Martinez, A., Ye, C., Kamarajan, P., Le, C., Zhan, L., & et al. (2020). Polymicrobial periodontal disease triggers a wide radius of effect and unique virome. Npj Biofilms and Microbiomes, 6(1). https://doi.org/10.1038/s41522-020-0120-7
  10. Willis, K. A., Postnikoff, C. K., Freeman, A., Rezonzew, G., Nichols, K., Gaggar, A., & Lal, C. V. (2020). The closed eye harbours a unique microbiome in dry eye disease. Scientific Reports, 10(1). https://doi.org/10.1038/s41598-020-68952-w
  11. Baltar, F., Gutiérrez-Rodríguez, A., Meyer, M., Skudelny, I., Sander, S., Thomson, B., Nodder, S., Middag, R., & Morales, S. E. (2018). Specific effect of trace metals on marine heterotrophic microbial activity and Diversity: Key role of iron and zinc and hydrocarbon-degrading bacteria. Frontiers in Microbiology, 9. https://doi.org/10.3389/fmicb.2018.03190
  12. Michoud, G., Ngugi, D. K., Barozzi, A., Merlino, G., Calleja, M. L., Delgado-Huertas, A., Morán, X. A., & Daffonchio, D. (2021). Fine-scale metabolic discontinuity in a stratified prokaryote microbiome of a red sea deep halocline. The ISME Journal, 15(8), 2351–2365. https://doi.org/10.1038/s41396-021-00931-z
  13. Acinas, S. G., Sánchez, P., Salazar, G., Cornejo-Castillo, F. M., Sebastián, M., Logares, R., Royo-Llonch, M., Paoli, L., Sunagawa, S., Hingamp, P., & et al. (2021). Deep Ocean metagenomes provide insight into the metabolic architecture of bathypelagic microbial communities. Communications Biology, 4(1). https://doi.org/10.1038/s42003-021-02112-2
  14. Ruocco, N., Esposito, R., Zagami, G., Bertolino, M., De Matteo, S., Sonnessa, M., Andreani, F., Crispi, S., Zupo, V., & Costantini, M. (2021). Microbial diversity in Mediterranean sponges as revealed by metataxonomic analysis. Scientific Reports, 11(1). https://doi.org/10.1038/s41598-021-00713-9
  15. Perskvist, N., Norman, I., Eklund, C., Litton, J.-E., & Dillner, J. (2013). The Swedish cervical cytology Biobank: Sample handling and storage process. Biopreservation and Biobanking, 11(1), 19–24. https://doi.org/10.1089/bio.2012.0036
  16. Kursa, O., Tomczyk, G., Sawicka-Durkalec, A., Giza, A., & Słomiany-Szwarc, M. (2021). Bacterial communities of the upper respiratory tract of turkeys. Scientific Reports, 11(1). https://doi.org/10.1038/s41598-021-81984-0
  17. Islam, S. M., Tanzina, A. Y., Foysal, M. J., Hoque, M. N., Rumi, M. H., Siddiki, A. M., Tay, A. C.-Y., Hossain, M. J., Bakar, M. A., Mostafa, M., & et al. (2021). Insights into the nutritional properties and microbiome diversity in sweet and sour yogurt manufactured in Bangladesh. Scientific Reports, 11(1). https://doi.org/10.1038/s41598-021-01852-9

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