Best practices for sample processing and storage prior to microbiome DNA analysis freeze? buffer? process?

So – we have been having a running discussion with people in my lab about one key issue in microbiome studies – how does one store samples prior to doing DNA extractions and does it matter?  As background for those who do not do this kind of work – the general principle behind DNA based analysis of microbes and microbial communities is that you can go to a sample (soil, water, air, tissue, etc) and extract DNA from that sample and then study the microbes in the sample by looking at the DNA.

This can be represented in the following cartoon:


But one of the challenges is, it is not always possible or ideal to directly isolate DNA from ones sample.  And how samples are processed can affect what DNA comes out the other end and thus can affect results.

From a simple examination of a set of projects, it seems to me there are something like five main classes of approaches used in going from sample to DNA which I represent in this cartoon.  Some details are in the text below.


  1. Freeze. Collect samples, whatever they may be (soil, swabs, tissue, etc) and freeze them, preferably at a very cold temperature.  Then, when it is time to do DNA extractions, thaw them out and extract DNA.
  2. Buffer. Mix with some buffer or DNA stabilization / extraction solution and then let sit, possibly at room temperature, for an extended period of time.  Then extract DNA.
  3. Process. Process fresh samples immediately (or as rapidly as possible) and extract DNA and then store the DNA for later use.
  4. Dry. Collect samples and dry them and then store them dried.  This is done usually in cases where the main goals do not involve DNA analysis.  But sometimes people would like to go to the samples afterwards and try to study microbes in and on the samples.
  5. Fix. Collect samples and then mix them with some sort of fixative (e.g., formalin, alcohol).  This is done with all sorts of samples where the main goal is to do something other than DNA analysis.  But then, after the fact, many people would like to get DNA out of these samples.

Of course, this is a bit of an oversimplification, but to me these seem to be the main categories of what is commonly done with samples.  There are also other “minor” categories of what is done with samples but these five seem to cover most cases.

I am writing this post for three main reasons.

First, I would like to generate a community discussion around what people do with their samples and why.  For example, I was told recently that there is some literature indicating the freezing fecal samples directly (rather than putting in some buffer) is less than ideal because the microbiomes retrieved from such samples change over time.

Second, I would like this to serve in a way as a place for people to ask questions about recommendations for what to do for their samples.  For example, I got asked yesterday about how to collect plant leaf samples in the field for microbiome studies if one does not have access to a freezer.

Third, I would like to use this as a launching pad for starting to collect together formal protocols that people use in microbiome and microbiology of the built environment studies.  We may use Protocols for this, but am still experimenting on systems.

So please -anyone who works on microbiome studies – we would welcome comments about methods and best practices and references and any other detail you can provide.

46 thoughts on “Best practices for sample processing and storage prior to microbiome DNA analysis freeze? buffer? process?

  1. For our cave samples our current protocol is this:
    Backpack cooler load with dry ice and space blankets
    Gather the samples in sterilized (inside and out) falcon tubes
    Wrap in dry ice and space blankets then place inside the backpack cooler
    On return to basecamp the tubes go into a better cooler with lots of dry ice
    Keep frozen until processing

  2. We’ve always processed human faeces as soon as it’s collected for DNA extraction. Though my colleagues and I acknowledge we’re fortunate to have access to a clinical unit and good anaerobic facilites to allow transport of samples to the lab within 1-2 hours if produced at a subject’s home. The reason we process fresh faeces for DNA extraction is because when we did a lot of FISH work, we found that a month of freezing caused big losses in Gram-negatives, especially Bacteroidales. We wanted our DNA- and FISH-based work to be as comparable as possible. We found storing aliquots of homogenate in PBS/glycerol (50:50, v/v) was better than storing 1-2 g portions of faeces for recovery of DNA and diversity. By far the biggest factor that we’ve found to affect diversity studies is the extraction method used. Kits don’t even come close to the efficiency of old-school phenol/chloroform. Consequently, for samples collected from in vitro work, we’ll still go with the old-school method. For human studies, in which we tend to have large numbers of samples, we use FastDNA kits. Gave up with Qiagen kits long ago.

    1. thanks – this is really helpful – I am a bit worried about all the people saying they are creating “fecal sample banks” for later analysis when what they do is just freeze the samples

    1. I think you have unintentionally misrepresented the paper.
      “Samples frozen with and without glycerol as cryoprotectant indicated a major loss of Bacteroidetes in unprotected samples, resulting in higher proportions of Firmicutes.”

  3. Thanks for starting the important discussion on Best practices for sample processing and storage prior to microbiome DNA analysis freeze? buffer? process? Lesley has made an important observation that a month of freezing of faeces caused big losses in Gram-negatives, especially Bacteroidales. Is this data published.
    One of the major problems faced by us using human faeces is that the extracted DNA using the Qiagen kit yielde highly fragmented DNA of 1Kb size or less. Any suggestions? Phenol/chloroform method yielded higher concentrations but yet fragmented.

    1. We have not published our assessment work, and have no intention of doing so. Colleagues at the Rowett Institute recently published on loss of Bacteroidetes in unprotected samples (not faeces). Regarding DNA extraction, we’ve never had problems with fragmentation with kits or phenol/chloroform methods. Where possible we have avoid vortexing, preparing to mix samples by pipetting.

    2. Qiagen stool mini kit uses chemical way to lyze. The resulted DNA should not be highly fragmented. I have used a protocol from JGI to produce long DNA:

      But it is a one-day-long protocol for a small number of samples.

      Qiagen stool kit should be a good and easy way to get long DNAs, but it will bias greatly towards Gram negative bacteria, because it is difficult to lyze Gram+ with chemicals alone.

      Bead bashing has less bias, but cannot generate very long DNA. It depends on how long DNA you need.

  4. I am working on a disposable sterile device designed to produce a homogenized sample within an encapsulated anaerobic environment. I’d be interested in collaborating with interested parties.

  5. Very helpful discussions. I want to carry out gut microbiome analysis and was wondering on the best way to preserve my stool samples

  6. Just saw this new paper which is of relevance to this post:

    Effect of preservation method on spider monkey (Ateles geoffroyi) fecal microbiota over 8 weeks.

    Summary from the paper below:

    • We examine differences in microbial communities due to fecal preservation method.
    • Fresh, frozen, and ethanol-preserved samples have most similar microbial communities.
    • 8 weeks of preservation had little effect on microbial composition and diversity.
    • DNA amount/purity did not correlate to microbe amplification, diversity, and composition.
    • Preservation methods exhibit bias toward/against some bacterial phyla.

    Studies of the gut microbiome have become increasingly common with recent technological advances. Gut microbes play an important role in human and animal health, and gut microbiome analysis holds great potential for evaluating health in wildlife, as microbiota can be assessed from non-invasively collected fecal samples. However, many common fecal preservation protocols (e.g. freezing at − 80 °C) are not suitable for field conditions, or have not been tested for long-term (greater than 2 weeks) storage. In this study, we collected fresh fecal samples from captive spider monkeys (Ateles geoffroyi) at the Columbian Park Zoo (Lafayette, IN, USA). The samples were pooled, homogenized, and preserved for up to 8 weeks prior to DNA extraction and sequencing. Preservation methods included: freezing at − 20 °C, freezing at − 80 °C, immersion in 100% ethanol, application to FTA cards, and immersion in RNAlater. At 0 (fresh), 1, 2, 4, and 8 weeks from fecal collection, DNA was extracted and microbial DNA was amplified and sequenced. DNA concentration, purity, microbial diversity, and microbial composition were compared across all methods and time points. DNA concentration and purity did not correlate with microbial diversity or composition. Microbial composition of frozen and ethanol samples were most similar to fresh samples. FTA card and RNAlater-preserved samples had the least similar microbial composition and abundance compared to fresh samples. Microbial composition and diversity were relatively stable over time within each preservation method. Based on these results, if freezers are not available, we recommend preserving fecal samples in ethanol (for up to 8 weeks) prior to microbial extraction and analysis.

  7. I think it is very important to consider what are the natural conditions of the sample. Feces arise from warm, anaerobic situations and thus may be difficult to stabilize.

    In contrast, our samples were dry sand from coastal sand dunes. We considered storing dry at room temperature but instead opted for storing on ice in a cooler during the day in the field and that evening popping them directly into the -80C freezer until DNA extraction by bead beating three weeks later, using standard MolBio protocol a recommended by Noah Fierer.

    If the sand had been damp, we probably would have stored on ice in cooler in the field and then prioritized immediate extraction, so as to skip the freezing. Soils with more organic matter (lower bulk density) might require a similar treatment, although I might try to mimic natural soil temperatures during interim storage in the field.

  8. I tested sample preservation (buffers (SLB and GIT) and freezing in liquid nitrogen in the field) and DNA extraction methods for a wide variety of hot spring samples (pH range ~2-9). The outcome metric was detected richness (measured by DGGE, this was back in the dark days). SLB preserved samples stored at ambient temperature for up to 2 weeks then -80, extracted with a hot CTAB/SDS method was the winner, though SLB and Mobio kit was a close second.
    Sorry it’s not open access, contact me and I’m happy to send on a reprint.

  9. Just saw a new paper on this topic

    Methods for Improving Human Gut Microbiome Data by Reducing Variability through Sample Processing and Storage of Stool.

    Gut microbiome community analysis is used to understand many diseases like inflammatory bowel disease, obesity, and diabetes. Sampling methods are an important consideration for human microbiome research, yet are not emphasized in many studies. In this study, we demonstrate that the preparation, handling, and storage of human faeces are critical processes that alter the outcomes of downstream DNA-based bacterial community analyses via qPCR. We found that stool subsampling resulted in large variability of gut microbiome data due to different microenvironments harbouring various taxa within an individual stool. However, we reduced intra-sample variability by homogenizing the entire stool sample in liquid nitrogen and subsampling from the resulting crushed powder prior to DNA extraction. We experimentally determined that the bacterial taxa varied with room temperature storage beyond 15 minutes and beyond three days storage in a domestic frost-free freezer. While freeze thawing only had an effect on bacterial taxa abundance beyond four cycles, the use of samples stored in RNAlater should be avoided as overall DNA yields were reduced as well as the detection of bacterial taxa. Overall we provide solutions for processing and storing human stool samples that reduce variability of microbiome data. We recommend that stool is frozen within 15 minutes of being defecated, stored in a domestic frost-free freezer for less than three days, and homogenized prior to DNA extraction. Adoption of these simple protocols will have a significant and positive impact on future human microbiome research.

  10. Hello,

    Our company is looking for the DNA sample kit to obtain the sample and send it to a storage facility for 2.5 years. Can your company provide such service? I understand there are some restrictions/requirements/standards to take the sample.

    Although our company is located in Honduras, it has access to the USPS to receive the kit and send the sample to USA. Our company also has a Plant Physician to take the sample.

    Thanks in advance

  11. hello, I do enjoy reading this blog and it has been very helpful.
    However, is there any methods in preserving any perishable food items like milk products for any prolong future meta-genomic analysis?

  12. Hello, I enjoy reading this blog and it has been very helpful.
    However, is there any preservation methods that can be used for perishable items like milk products for later meta-genomic analysis?

  13. Thank you for the informative post. I am looking to perform FISH on dental plaque samples to be transported from the field. Any suggestions will be helpful!

  14. Hi everyone ! I had some raw materials, Curculionidae, using for dna extraction. However, after storing in alcohol 70% for 2 weeks they gave such a bad result, not much DNA. Is there any solution to solve this problem ?

    1. Can you clarify a few things?

      1. What do you mean by “not much DNA”?
      2. Do you get more DNA from fresh samples?
      3. Probably better to use 100% ethanol
      4. How did you do the DNA extraction?

      1. Hi Jonathan Eisen !
        First of all, I want to say thank you for your reply !
        1. Actually, I saw some weak bands.
        2. Yes, I do. After having that bad result, I made another test by using fresh samples and the result was much better, no more weak bands.
        3. Maybe I will try 100% ethanol :)
        4. I used CTAB method.

  15. Hello!
    Thanks for providing space for discussion and I have somewhat relevant input for this subject.

    My research focuses on describing gut fungal communities by combining ITS1 sequencing and phenotying cultured fungal isolates. The preliminary results from comparing fresh vs frozen (-80 C) stool sample indicates loss in fungal viability, which was not a big surprise. Because of this reason, I’m leaning towards using fresh samples for culturing efforts then freezing (-80 C) the remaining samples for DNA extraction in batches. I presume that the approach suggested by Gorzelak and colleagues (whole stool homogenization after flash freezing) would lead to significant loss of viable bacterial and fungal cells and would not be a practical method for me. Further, it would be interesting for such studies to include changes in fungal community detection in different sample storage conditions (I haven’t found any yet!).


  16. Hi there! Nice blog

    I am thinking in starting a project on skin microbiome and I have a very basic question. How do you sterilize the solutions and equipment for NOT having microbe DNA in/on them???? Filters, UV, autoclave, combinations or others?

    Thank you!

  17. Hello all – I stumbled upon this page via Google. What a fantastic page – truly. My group’s question is more proteomic. It seems that the preservatives discussion tends toward either DNA quality or diagnostic parasitology domain. We will be performing an assay to determine the level of calprotectin in stool (human). An expert that I pinged stated that, “Fcal is biologically functional as a hetoerodimer and heterotetrimer (and this is what the assay measures), I assume the ethanol would denature the 3* and 4* structure and cause a problem.” Given that consideration, does anyone have any thoughts about the best preservative in this case?

    1. David,

      Just saw your comment and I am actually interested in validating our stool preservative for the purpose of stabilizing calprotectin. If would be open to testing this, please let me know !


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Jonathan Eisen

I am an evolutionary biologist and a Professor at U. C. Davis. My lab is in the UC Davis Genome Center and I hold appointments in the Department of Medical Microbiology and Immunology in the School of Medicine and the Department of Evolution and Ecology in the College of Biological Sciences. My research focuses on the origin of novelty (how new processes and functions originate). To study this I focus on sequencing and analyzing genomes of organisms, especially microbes and using phylogenomic analysis (see my lab site here which has more information on lab activities).  In addition to research, I am heavily involved in the Open Access publishing and Open Science movements.