From the Field to the Lab: Sequencing Wild Microbiomes
Research on the animal microbiome—the amazing communities of bacteria, archaea, protists, fungi, and viruses present throughout an organism—is a rapidly growing field. This is because these communities of tiny organisms serve important functions in our body, like helping us digest food or regulating immune system functions . While the microbiome is partially passed down from mothers to their offspring , the environment is also a major factor influencing the types of microbes living in an organism . Environmental changes can alter the composition and diversity of our microbe communities, which in turn can affect the host organism’s health !
Much of our understanding of microbial communities comes from sequencing the DNA of the whole microbial ecosystem. This allows researchers to identify the variety of different microbes present in the sample. For ecologists and evolutionary biologists who want to understand the relationship between an animal and their microbes under natural environmental conditions, this means that they somehow have to collect microbiome samples from animals in the wild! So, how do we do this? I was thinking about this question as I set off to conduct my pilot study on wild lemur microbiomes this past summer at Beza Mahafaly Special Reserve in southwest Madagascar.
Fig. 1 - This is what prepping my sample collection tubes looks like! These tubes are all filled with a solution that will keep the microbial composition of my samples stable for several weeks at room temperature—perfect for collecting samples in remote field sites.
A majority of studies on the microbiome of wild animals solely focus on the gut microbiome, even though there are unique microbial communities throughout different regions of the body [3, 4]. The focus on gut microbiota makes sense though: out of all the different microbial communities in an animal, this one is the easiest to study noninvasively. Researchers often collect scat—or fecal samples—from wild animals and measure the microbial diversity within that fecal sample as a proxy for directly sampling the gut microbiome. These types of samples are relatively easy to collect, since you can follow the animal or groups of animals to collect scat, and you don’t actually have to handle the animal to get the samples. But here I had a problem: not only was I interested in the lemur gut microbiome, I was also interested in understanding the oral, genital, and hair microbiomes of lemurs. Different body regions actually have unique microbiomes, with different types and proportions of microbes that have functions specific to the part of the body in which they’re found ! Therefore, by only studying gut microbiota, researchers are missing out on a ton of information about the other types of microbes present in an organism. These types of microbiome samples require direct handling and sampling of the lemurs, and I also had to be careful to not contaminate the microbiome samples as I collected them.
Luckily, I collaborated with an existing research team that conducts annual capture and release of one species of lemur, the Verreaux’s sifaka. This team puts identifying collars on juveniles while also taking body measurements and small DNA samples from them, all for the goal of developing effective conservation strategies for these critically endangered primates. I was also able to follow ring-tailed lemur groups to opportunistically collect fecal samples from them. Normally during the capture and release of sifaka, the researchers and ecological monitoring team I worked with search the reserve for groups with juveniles, catch sedated individuals in a blanket and gently carry them to camp to take measurements and give them collars. However, I needed to collect hair microbiome samples from the lemurs without any contamination from people (or a blanket that has touched hundreds of other lemurs). So, I brought along individually wrapped, sterile blankets to use for each individual we caught! I would collect my hair samples right after catching them in a blanket to try and minimize contamination of their hair before taking them to camp. There I used sterile cotton swabs to gently swab their mouths and ano-genital regions. By the time I left the field site, I had 102 microbiome samples ready to analyze!
Fig. 2 - Me collecting a hair sample from a sedated Verreaux’s sifaka (not pictured) in the field with my sterile blanket in the lower right corner; two Verreaux’s sifaka individuals resting in the afternoon heat.
Back in the lab, I began the first process necessary for microbiome genomic DNA sequencing: DNA extraction from the samples. This step should be done in a sterile environment with little risk of contamination, so I conduct all of my DNA extractions in a biosafety cabinet. This hooded workspace is helpful for maintaining a sterile environment
because it has a constant outward airflow, preventing potentially contaminated air from coming into the workspace. By breaking open the bacterial cells and then removing all the proteins and extra cell debris, we can extract pure DNA from the sample. Then I will make a lot of copies of the 16S rRNA gene found in the extracted DNA. 16S is a gene found in most bacteria that changes very little between different groups of bacteria. These small changes between lineages allow us to identify the types of bacteria within the sample . The next step will be utilizing an amazing resource on campus—the Genomics Resource Laboratory—to sequence these samples and compare the bacterial diversity between and within all of these microbiome samples. This entire process of getting microbiome samples from wild lemurs took me across the world and back, and illustrates the range of work that can go into conducting biological research. I’ll be busy sequencing and analyzing my samples for the next few months, but I hope to be back in Madagascar this summer, collecting another set of samples for this project!
Fig. 3 – Amanda Fuchs and me extracting microbial DNA from field samples.
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