Microbiome research has experienced a surge of interest in recent years due to the advances and reduced cost of next-generation sequencing technology. The production of high quality and comparable data is dependent on proper sample collection and storage and should be standardized as far as possible. However, this becomes challenging when samples are collected in the field, especially in resource-limited settings. We investigated the impact of different stool storage methods common to the TB-CHAMP clinical trial on the microbial communities in stool. Ten stool samples were subjected to DNA extraction after 48-hour storage at -80°C, room temperature and in a cooler-box, as well as immediate DNA extraction. Three stool DNA extraction kits were evaluated based on DNA yield and quality. Quantitative PCR was performed to determine the relative abundance of the two major gut phyla Bacteroidetes and Firmicutes, and other representative microbial groups. The bacterial populations in the frozen group closely resembled the immediate extraction group, supporting previous findings that storage at -80°C is equivalent to the gold standard of immediate DNA extraction. More variation was seen in the room temperature and cooler-box groups, which may be due to the growth temperature preferences of certain bacterial populations. However, for most bacterial populations, no significant differences were found between the storage groups. As seen in other microbiome studies, the variation between participant samples was greater than that related to differences in storage. We determined that the risk of introducing bias to microbial community profiling through differences in storage will likely be minimal in our setting.

We performed qPCR to determine the abundance of various microbial subpopulations, namely Bacteroidetes, Firmicutes, Enterobacteriaceae, Bifidobacterium spp. and Lactobacillus spp., relative to Eubacterial 16S rRNA gene amplification. We also amplified fungal species by targeting the ITS1 region. Amplification was performed on the Rotor-Gene Q thermocycler (Qiagen) as singleplex reactions using 1X KAPA 2G SYBR Fast Uni Kit (KAPA Biosystems), 0.2 µM of each primer (see below) and nuclease free water (Qiagen) in 20 µL reactions. DNA input was standardized to 30 ng per reaction and all reactions were performed in triplicate. The adapted cycling conditions for all bacterial populations were as follows: denaturation at 95°C for 3 minutes, followed by 40 cycles of denaturation (95°C for 5 seconds) and annealing/extension (60°C for 30 seconds). The cycling conditions for fungal amplification were denaturation at 95°C for 3 minutes, followed by denaturation (95°C for 5 seconds) and annealing/extension at 64, 62 and 60°C for 30 seconds for 10, 10 and 20 cycles respectively. Fluorescence was acquired to the green channel during the annealing step. The Rotor-Gene software was used to calculate the efficiency and detection threshold for each primer set using individual standard curves. The efficiencies ranged between 0.91 and 1.04 with R² values > 0.99.

Dataset details:

Quality and Purity results:

This dataset contains A260/A280 and A230/A280 purity and quality metrics of extracted DNA, and DNA yield as determined by Qubit.

Cts:

These data are comma separated values representing the Cycle threshold (Ct) values of 7 microbial qPCRs tested for 4 different storage conditions. 10 samples were tested in triplicate for each storage condition.

Missing values:

Cts: Sample 3, Fungal amplification - all three replicates were lower than the detection limit of the qPCR.