Wonderful World of Microbes

Sep 08, 2014

Enid Gonzalez-OrtaHello!  My name is Enid Gonzalez-Orta and I am an Associate Professor of Biological Sciences at California State University, Sacramento, where I teach undergraduate students about the wonderful world of microbes.  In addition, I embed an authentic research experience into my microbial diversity class each spring where we study soil samples collected from CSU Sacramento Arboretum or from local vernal pool sites.  We study the bacterial community of these samples using traditional methods, like culturing samples onto laboratory media, sequencing of the 16s rRNA gene through the Sanger method, and building phylogenetic trees.  However, I began to think about how much the field of microbial ecology has changed and is changing.  I thought about how culture-independent methods allow us to peek into environments seldom studied in the laboratory and how these methods reveal members of bacterial communities that were previously not known to inhabit these niches.  I also thought about how next generation sequencing (NGS) methods are becoming becoming the “tradition” in this field.  And, I thought about how important it is for undergraduate students to have hands-on experience with bioinformatics and computing in order to interpret the volume of sequencing data that is produced through NGS.  But, how I could I do this when I myself had little-to-no experience with processing 16s rRNA NGS data?  Then came EDAMAME to the rescue!


When the opportunity to attend EDAMAME presented itself, I just couldn’t pass it up.  I haven’t been a student for a little while, so I was excited to be in a classroom sitting in a desk and not in the front of the room for a change.  Not only would this course help me teach undergraduate students in my courses how to process NGS data, but it would help to advance my research program CSU Sacramento as well.   My research focuses on the bacterial diversity of the California Vernal Pool Ecosystem in the Sacramento Valley.   This project is done in collaboration with my fellow colleague Dr. Jamie Kneitel and  has been worked on by many undergraduate students.   One of them, Dana Carper, joined my lab as a graduate student focused her Master’s work on this topic .


My experience with EDAMAME so far has been awesome.  We are officially half-way through the course and I can say that I have learned a lot from Ashley, Tracy, and Josh.  And, on top of some great instruction, we have amazing guest lectures!  I can’t wait to take what I’ve learned and teach it to my undergraduates.

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Seagrass: The Whale of the Plant World

Sep 02, 2014

My name is Cassie Ettinger and I am a 2nd year PhD student in Jonathan Eisen’s laboratory at the University of California, Davis. I am working with Hannah Holland-Moritz, a Junior Specialist, and Jenna Morgan-Lang, the Post-doc in charge of the project, to try to characterize the Seagrass Microbiome, the entire microbial community associated with seagrass.


Why study the seagrass microbiome? Seagrass has an interesting evolutionary history.  Starting out as marine algae, an ancestor of seagrass made the transition from marine to terrestrial with its descendants eventually becoming what we know today as plants.  Some of these plants later returned to the marine home world from whence their ancestors had originated (just like whales evolved from a land-based ancestor that returned to the marine environment!). Seagrasses are the only known flowering plants to have made this return to the sea. This is because surviving in a marine environment poses significant physiological and morphological challenges. We are interested in the role microbes might have played in the adaptation of seagrass to the marine environment and if any co-evolution has occurred between seagrass and its microbiome.

Additionally, seagrass plays a vital role in coastal marine ecosystems by cycling nutrients, protecting the coastline from erosion and serving as a food source and hatchery for many marine macrofauna. Microbes have been found to have a significant effect on terrestrial plant fitness, so it only makes sense to try to determine what role microbes might have on marine plant fitness as well.

I am currently investigating the nature of “edge effects” in terms of microbial diversity by analyzing a dataset we have from microbial communities obtained from the inside, on the edge and outside seagrass beds. Hannah is working on determining the intraplant microbial biogeography (how the microbial community composition differs across a single plant) and Jenna is coordinating our collaboration with the ZEN (Zostera Experimental Network), a group of 40+ labs that work on Zostera marina, an abundant seagrass species. They have all agreed to send us leaf, root, sediment and water samples from their respective study sites around the globe! (And of course we are using MoBio PowerSoil kits for all our DNA extractions!)

My goal for EDAMAME is that I will leave the workshop with a better understanding of the data analysis algorithms I am using; I hope that what I learn at the workshop will help inform my choice of parameters and programs to use for my respective datasets. I am super excited to learn more about how to analyze metagenomic data sets so that I can use those skills to investigate the functional roles of different microbes in the seagrass microbiome. I am also enthusiastic to learn how to use R to interpret data obtained from QIIME and mothur and to make beautiful publication ready graphs. I also plan on sharing everything I learn with my fellow lab members back home!

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Dandelions, Microbes, and 300 Middle Schoolers..Oh My!

Aug 25, 2014

My email dings soon after lunch on Monday, I open the window, and there it is in all its glory. The MiSeq data has arrived! I promptly download the file giddy with excitement…and wait. And wait. The file is huge, and incomprehensible, and somehow contains nearly 15 million sequences.


Luckily, I had signed up to attend the Explorations in Data Analysis for Metagenomic Advances in Microbial Ecology (EDAMAME) workshop hosted by Michigan State University. So here I am at Kellogg Biological Station on Gull Lake taking baby step-by-baby step through the shell, QIIME, Mothur, and R. Many of these I have used in the past, believing that I had taught myself how to run them. But here’s the difference: what I had done was copy and paste some commands, hope it worked, and then compared my results to results from other programs. Very efficient! Now: we are really learning the meat of these programs down to the last –i and / and the knowledge of these details allows the understanding of what happens in each step. My goals while here at EDAMAME 2014 are not to merely analyze this dataset, but to be able to take this knowledge and power and translate it to any dataset.


Dandelion photo credit: Lea Shell of yourwildlife.org


But, I have gotten ahead of myself. Where did these data come from you ask? The short answer: middle school teachers. The long answer: a bit more complex. I am a postdoctoral research associate at the North Carolina Museum of Natural Sciences (NCMNS) in the Genomics and Microbiology Lab, and, together with the Rob Dunn Group at North Carolina State University, we are embarking on a journey to bring real science into middle school classrooms. Students Discover (http://education.yourwildlife.org) is a program at the intersection of research, education, and citizen science. This summer we had 12 middle school teachers working in the labs at NCMNS, and specifically 3 teachers working with me in the G&M lab on the dandelion rhizosphere. In 3 weeks, these teachers went from learning how to use a pipette, to running an experiment, and finally setting up a multiplexed Illumina MiSeq run. Their excitement and hunger to learn was infectious.

Photo credit:  Lea Shell of yourwildlife.org

Why dandelions? Well first, they are really cool. But importantly: they are everywhere, everyone can recognize them, and they can withstand a lot of disturbance. Perfect for citizen science! So this summer the three North Carolina middle school teachers, Laura Cochrane, Arthina Blanchard, and Amy Lawson, and I transplanted dandelions into nine different soil types to determine what happens to the rhizosphere microbes. This served as a pilot study that will be repeated in their classrooms throughout the school year. Roughly 300 middle school students will be driving research based on the results of this preliminary dataset. By maintaining this connection between NCMNS, NC State, and the classrooms, the students will have an active, hands-on role in current science.


For now, building confidence in proper management, handling, and analysis of these data will help foster confidence to move forward in this scientific adventure with some dandelions, their microbes, and a whole lot of middle schoolers.

-Julia L. Stevens

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EDAMAME, onward!


EDAMAMEGreetings, Culture Dish followers!


My name is Ashley Shade and I am an assistant professor at Michigan State University in the Department of Microbiology and Molecular Genetics. I am excited to share our first days’ adventures in our new workshop, Explorations in Data Analyses for Metagenomic Advances in Microbial Ecology (EDAMAME). EDAMAME is a week-long intensive training in bioinformatics tools and ecological analyses of microbial metagenomics data, held at Michigan State’s Kellogg Biological Station.


I know what you are thinking: Silly new professor! Why dedicate preparation time and a week of your life to a non-credited summer course that won’t count towards your teaching requirements?


The answer is simple. EDAMAME is happening because there is a hunger for this training. I am leading EDAMAME because I am passionate about informed analyses of metagenomic data. I want to empower other microbiologists to own their analyses, from raw sequencing reads to ecological interpretation.


EDAMAME was designed to advance computational skills in students with strong backgrounds in biology. My co-instructors, Joshua Herr and Tracy Teal, both of MSU, and I have worked hard to prepare lectures and hands-on tutorials, and to invite interesting guest speakers. Today is Day 2, and it is amazing what we have already accomplished. Yesterday we dug right into a tutorial on shell. Then, we spent the afternoon yesterday and full day today having the “QIIME” of our life. Titus Brown and Jack Gilbert gave guest research lectures today, and Pat Schloss is arriving tomorrow to work through a day of “mothur”-ing .


Did I mention, it is only Day 2?


Last evening, the students gave 5-minute lightning chalk talks to introduce themselves and their research. I was impressed by their diverse expertise; their research questions addressed topics ranging from animal nutrition to soil processes. We have students, post-docs, and professors learning side by side. They are tenacious! I like them. I hope that you will be hearing from them with some guests posts in the very near future. (They were happy to have the new MO BIO t-shirts. Photo op to come!)


If you’d like to join in on the conversation, we are tweeting! #edamame2014. We also have all of our course materials online at edamame-course.org.


EDAMAME, onward!

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Where in the World: Salt Marsh Microbes

Want to play in the mud?  Like the ocean?  If your answer to both questions is “yes,” you may have a tinge of jealousy when it comes to this Where in the World project.  François Thomas, PhD, works as a postdoctoral scientist with Stefan Sievert, PhD, at Woods Hole Oceanographic Institution on Cape Cod in Massachusetts.  His collaborative project* funded by the National Sciences Foundation studies microbes in New England salt marshes. By sampling sediments and roots of the salt marsh grass known as Spartina, he uses both cultivation and molecular techniques to understand the microbial communities in each environmental niche. In particular, he is studying sulfur-oxidizing microbes, which can profit from the release of oxygen from the roots and might play a critical role in linking the carbon, nitrogen and sulfur cycles. To get back to your childhood memories, check out this video from the field.

*Anne Giblin, PhD, and Zoe Cardon, PhD, of Marine Biological Laboratory in Woods Hole.

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Troubleshooting Low DNA yields

Jun 24, 2014
Michelle Tetreault Carlson

One of the top five technical concerns we get at MO BIO relates to low yields of DNA.   Whether it be sediment, swabs, waste water or sludge, at some point we’ve probably heard that you didn’t get enough DNA out of it.  After all, there is no such a thing as getting too much DNA.   Nope, haven’t heard that grievance even once!

mb110dWhen someone tells us they didn’t get enough DNA, we rule out the obvious offenders. It could be that your lab-mate left your samples out all night at room temperature and didn’t tell you. It might also be that you’ve made an error in the way you’ve interpreted or conducted a protocol or perhaps you’re using an inappropriate kit. The list goes on and on.  However, the answer could be a lot simpler.  Low DNA yields (defined here as those that can’t be easily measured with an absorbance spectrophotometer, for example, a Nanodrop) may not indicate that something went wrong at all.   It might just be that there wasn’t a lot of DNA to begin with.  This is very common with low biomass samples like swabs, filters or sediment.

Whenever you’re working with a sample that has never been tested, you may not know the taxa or number of cells present, so you may not know how much DNA to expect.   This is especially true of environmental samples.  For example, every type of soil is different, with varying pH, humics, carbon content, etc.   But fortunately you don’t need to march blindly into the darkness.   Instead you can do something much better;  run a control.

Soil in bead tubeFor soil, a good control is to “spike” your sample with a known quantity of bacteria or fungus and run it through the kit protocol.   You can start by growing an overnight culture of your favorite bug.   It’s best to pick something that is representative of what you might be looking for in your sample.  Then, take 1-2 ml of the culture and centrifuge to pellet the cells.  A good rule of thumb for a bacteria like E. coli is that 1 ml of an overnight culture contains 10^9 cells or approximately 5 micrograms of DNA.  Fungus have roughly 10 times the  concentration of DNA per cell as bacteria.   Find out  the genome size of your test bug’s DNA, measure the cell density in your culture and calculate the expected yield.  Now, you can spin down the cells from the culture into a pellet and then resuspend  in a small amount of neutral buffer like 100 uL saline.  Add this to your soil sample and let incubate for at least 10 minutes before starting your protocol.  Extract this alongside your original samples and voila;  you’ll be able to see how much DNA is present in your sample because it will be the difference  in concentration between your spiked and unspiked sample.  If not, then there might be something else going on.   Perhaps there is something in the soil like metals that is interfering with your prep or your homogenization protocol isn’t adequate for cell lysis.  If you can’t figure it out, contact MO BIO Technical support.

Similar “spiking” experiments can be done with other samples as well.   Just treat the cultured sample as you would your working sample.    For example, if you are filtering pond water then you can spike in your microbial culture and filter just the same.    In this way you can be confident that your DNA Isolation method is working properly, determine the concentration of  your sample and eliminate one more scientific unknown.   In fact, it’s always a good idea to run this type of control, even if you’re comfortable with your current extraction method.   It’s a little extra time up front but will likely save you loads of trouble down the road.   Here is to basic science 101!


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A Birthday Salute to a Special Sub

Jun 09, 2014

Last week marked the deep sea submersible Alvin’s 50th Birthday.  This translates to 50 years of unprecedented ocean exploration and discovery!

MO BIO Laboratories, Inc. would like to thank Alvin and the very dedicated folks that make Alvin’s story so historic in scientific discovery!

Thanks to Chris Linder, Alvin and MO BIO’s cultivating curiosity were united on R/V Atlantis during the first scientific expedition with the newly outfitted Alvin, awesome sauce!  I can’t help but share my own special moments with Alvin, thanks for checking in!





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96 Well Plate Shaker: Your High-Thoughput Companion

May 27, 2014
Michelle Tetreault Carlson

Considering a move to high throughput DNA/RNA isolation? Many of our customers, who’ve been using MO BIO single prep kits for some time, now want to scale up.  We’ve been getting a lot of questions as to what equipment is required to make the switch.  Scaling up usually means moving sample preps over from 2 ml tubes to standard 96 well microplates. While many customers have experience with multichannel pipetting for these plates, many have never done any bead beating in 96 well format.

Most MO BIO kits take advantage of bead beating to assist in cell lysis because it is an exceptionally good method for isolating gram+/- and spore and oocyst DNA & RNA.  But bead beating, while easily done on a vortex in 2 ml bead tubes, requires much beefier equipment for 96 well plates.

For high through-put disruption of biological samples, we recommend the MO BIO 96 Well Plate Shaker used along with our Plate Adapter Set. When used with our Plate Adapter Set, the 96 Well Plate Shaker can shake two 96 well 1.0 or 2.0 ml bead beating plates for a total of 192 simultaneous samples.    Cell types of all types (gram +/-, spores, oocysts) can be lysed including those in tough materials: bone, conifers, grains, hair, minerals, plants, seeds, soil, feces, tissue samples and waste material.
All of the MO BIO high through-put MO BIO DNA/RNA Isolation kits have been optimized and come with instructions for use on the MO BIO 96 Well Plate Shaker. The Adapter Plates are self-centering so that they simplify the process of flipping the bead beating plates halfway through the homogenization protocol, making it easy to apply equal energy to all samples during homogenization. The instruction manual that comes with each kit includes ideal settings for each particular sample type making success more likely.  We’ve put together a handy VIDEO showing the 96 Well Plate Shaker in action.

New Option: The MO BIO 96 Well Plate Shaker just got even more versatile!  We now have the 2 ml Tube Holder Set which can be used for high through-put bead beating for 2 ml bead tubes!   This is especially helpful for kits like the PowerBiofilm DNA Isolation kit which do not come with a 96 well format bead plate option.

NOTE: Use of other manufacturer's 2 ml tubes has not been tested.

The 2 ml Tube Holder Set consists of 2 identical plastic blocks capable of holding up to 48 MO BIO 2 ml bead tubes in each block.  This allows for homogenization of 96 samples on a single run with the 96 Well Plate Shaker. In contrast, a vortex adapter only shakes up to 24, 2 ml bead tubes.  The Tube Holder fits inside the same Plate Adapter Set that is used to hold 96 well plates.

The following is a list of 96 well kits which are compatible with and contain instructions for bead beating on the MO BIO 96 Well Plate Shaker:

PowerSoil® -htp 96 Well Soil DNA Isolation Kit

UltraClean® -htp 96 Well Tissue DNA Isolation Kit

UltraClean® -htp 96 Well Swab DNA Kit

PowerPlant® Pro -htp 96 Well DNA Isolation Kit

PowerMag Soil DNA Isolation kit (for epMotion® & KingFisher®)

PowerMag Microbial DNA Isolation kit (for epMotion® & KingFisher®)

PowerMag® Seed DNA Isolation Kit (for KingFisher®)

PowerMag™ Microbiome RNA/DNA Isolation Kit for epMotion® & KingFisher®

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*Freeze Dried*

Apr 16, 2014

Craig Cary, a friend of the MO BIO Laboratories’ family came by to say hello and also updated us on his work.

Craig works on soils from Antarctica in a region called the Dry Valleys in Victoria Land.   Talk about extreme!  Imagine glaciers, extremely cold temperatures, very dry air, rocky salty soils, lack of vegetation, “Kadabatic” winds derived from the mountains, 24 hours of sunlight during “summer”, and to top it off mummified seals!  This place has been described as one of the most extreme environments on the planet, and is used as a model ecosystem to better understand the conditions on Mars, a planet akin to this “freeze dried environment”.

In 1989, an important treaty called the Montreal Protocol was put into effect banning the use of CFC’s in any commercial products due to its deleterious effects on our ozone layer.  It turns out it this treaty has made a profound effect on positively regenerating the ozone layer which is concentrated in the upper atmosphere of the poles.  This comes with concern about how this will effect the environment in Antarctica, ozone traps heat including the climate changing CO2 we are pumping in to it.  Will Antarctica continue to warm?  We know that the Antarctic Peninsula, a fingerlike projection sticking up towards South America, is one of the fastest warming regions on the planet.  What will happen with the rest of the continent, will it warm too?  This is what Craig wants to know.  He states that 70% of the fresh water on planet is stored in the glaciers.  I think to myself, maybe I shouldn’t be so eager to live on the coast, stick to the hills!

Prior to the 1970s, the thought was there was no life at all in the Dry Valleys.  Here is where Craig and his audacious team take the stage.  They are studying changes using the biology of this environment as their proxy.  The paradigms so far that dispute prior knowledge about this “thought to be lifeless environment” are that the diversity of the microbes is way higher than previously thought, similar to your backyard as a matter of fact!  Not only that, but each valley has it’s own biological clock if you will.  There are microniche environments on small geographic scales where turnover rates can be as short as 2 years.  The main drive behind Cary’s group is to figure out what determines and can predict distribution of biota in the Dry Valleys and beyond.

Their partners in crime for this study are mosses, lichens, endoliths (bacteria living in rocks), springtails, mites, nematodes, rotifers, and of course     microorganisms.  Each specialized research group will monitor changes in these communities and generate a predictive model that will help better understand not only this environment but other environments as well.  Springtails, for example, have two genetic morphs, one adapted to cold and one adapted to heat.  By setting up “springtail traps” you can count the numbers of each morph to monitor temperature changes in the field.  Sounds like Flintstone material to me “Hey Freddie, can you please set out the springtail thermometer for me, I need to know if we need more clothing for Pebbles!”

Generating this kind of data does not go without struggle and is not cheap.  To get to their base camp near Spaulding Pond in the Taylor Valley, the team of twenty Kiwis and Americans start off on a C130 military aircraft from New Zealand to McMurdo Station and are then airlifted via helicopter where they set up camp including a bucket as their bathroom.  The locations they sampled from day to day over 4 weeks at times required 30-40 kms of hiking in sub zero temperatures.  Their goal of collecting samples was organized according to a method where they overlaid tiles determined using satellite images on to a map and determined which spots are the best to sample.  What’s even more interesting, is that they have some really high tech equipment to help facilitate.  In one study to understand lichen primary production, they have a fiber optic cable that will send a pulse of light to the edge of a lichen and wait to receive a signal back that informs them about the physiological state of the lichen.  This information along with temperature, humidity, and barometric pressure are sent via a satellite phone real time!  They are also working with drones that will “autopilot” aerial surveys and provide low level mapping of the area, totally teched out!  When the field season was over, they left nothing behind literally returning rocks to their original location.  What they brought home is some hefty “no shower for 4 weeks” stench but also a plethora of data and samples that could help us change the world we live in today!

Thanks Craig for the invigorating talk!  Need any volunteers?

Craig Cary uses MO BIO’s nucleic acid extraction kits to extract his samples.

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Swabs Revisited

Mar 24, 2014
Michelle Tetreault Carlson

Three years ago we published a tech article in which we went into detail on using swabs with MO BIO DNA Isolation kits.   Recommendations depend both on levels of PCR inhibitors and whether one wants microbial or eukaryotic DNA.  The article included details on how to transfer swab samples to bead tubes and collection tubes for use in each kit.  Swabs continue to be one of the fastest and easiest methods for sample collection and remain a popular subject for tech support questions.

Since the previous swab article, we’ve developed two new kits that we recommend for swabs, one for RNA Isolation and the other, for high through-put rapid mechanical lysis of microbial cells on swabs from low biomass/low inhibitor samples.

RNA from Swabs

The PowerMicrobiome RNA Isolation Kit is designed for fast and easy purification of total RNA from samples high in PCR inhibitors. In order to use a swab with this kit place the swab into bead tube containing 650 ul of Solution PM1/bME.  If the swab was previously frozen be sure not to allow the swab to thaw before placing it into the bead tube because any freeze thaw cycle will pop open cells and release tons of RNases into your sample.  Just go directly from freezer to buffer.  Rotate the swab in the buffer and let it soak for a several minutes to release the cells into the solution.  If the swab has a head that can be snapped off, go ahead and do so, leaving the swab in the tube. Otherwise remove the swab while gently squeezing it against the wall of the tube to remove as much of the solution as possible and proceed with the rest of the protocol.

High Through-put Swabs

The UltraClean –htp 96 Well Swab DNA Kit can be used for rapid mechanical lysis of microbial cells from swabs in high through-put format.  It is not a DNA purification kit but rather a kit for direct PCR.   Swabs are placed into the wells of a 96 well block containing 0.1 mm glass beads and SW1 lysis buffer.  The swab handles are snapped off just below the height of the plate and the plate undergoes bead beating.  The lysate can then be directly removed and used for PCR.  This kit will only work for low biomass samples with low levels of inhibitors; for example skin swabs.

The following table combines kit recommendations from the previous swab article along with those for our most recent swab-compatible kits.  We hope this will assist you to quickly find the right kit for your needs.

What is the best kit for your swab application?
Host DNA
Microbial DNA
Host or Microbial RNA
Low reading
Low Inhibitors PowerLyzer UltraClean® Tissue & Cells RNA Isolation Kit
High Inhibitors
(no bead beating)
Low Inhibitors
High Inhibitors

One more thing…Swab Storage

Many of our more recent questions involve the storage of swabs rather than which kit to use them in. Customers often ask us what sort of storage buffer they should put their swabs into until they can get them back to a lab.  Surprisingly, it turns out that the best answer may be “nothing.”

A study published in the Proceedings of the National Academy of Sciences (PNAS) by Rob Knight’s lab at the University of Colorado (Ref 1) looked at the effect of storage conditions on skin-associated bacterial communities collected on cotton swabs.    Skin surfaces were swabbed with pre-moistened sterile swabs and were either immediately frozen at −20 °C or −80 °C or left to dry in 15-mL conical tubes and left out on a laboratory bench.  Bacterial DNA was extracted from swabs after either 3 days or 14 days. Bacterial community analysis showed little difference between swabs stored at room temperature, -20 °C and -80 °C during this time period.  This is good news for those of you who might not be close to a freezer when samples are taken and might make you a bit more relaxed about isolating swab DNA.

A Swab is a Useful Thing

As you have seen, even if a MO BIO kit doesn’t have “swab” in the name,  it might be perfect for your swab samples.  If you are unsure which one to use think about PCR inhibitors and cell type.   And you don’t necessarily need any fancy buffers to store the swabs.   If there is more than one type of kit that will work with your swabs we’d be happy to send you a sample of each to try.  Always feel free to contact Technical Support if you have any comments or questions.

(1)Forensic identification using skin bacterial communities
Noah Fierer, Christian L. Lauber, Nick Zhou, Daniel McDonald, Elizabeth K. Costello, and Rob Knight
PNAS, Apr 2010; 107: 6477 – 6481.

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