Validating a high-throughput in vitro model for culturing antibiotic-altered microbiome communities

The gut microbiome plays a central role in host health, and its disruption is associated with metabolic, infectious, and immune conditions. Stool sampling is frequently used to characterize snapshots of the gut microbiota. To screen microbiota communities in vitro, multi-stage bioreactors have been developed that model distinct physiological environments along the gastrointestinal tract. However, such bioreactors are often operationally complex and provide limited throughput. Stool-derived in vitro communities offer several advantages to study microbiota due to their ease of use, high-throughput nature, and have previously been shown to remain stable following repeat passaging. Yet, prior characterization of stool-derived cultures has relied on 16S rRNA sequencing of unaltered gut microbiota, leaving several key questions unanswered: 1) how closely does in vitro culture recapitulate the functional potential and resistome composition of its inoculum and 2) can these cultures model antibiotic disrupted microbial communities? In this work, we generated stool-derived in vitro cultures from male and female mice with both unaltered and antibiotic altered gut microbiomes and applied shotgun metagenomic sequencing to characterize taxonomic composition, functional pathway capacity, and antimicrobial resistance gene (ARG) content. Antibiotic-altered microbiota communities, dominated by Enterobacteriaceae, were faithfully recapitulated in culture with preservation of both taxonomy and ARGs. By contrast, unaltered communities underwent substantial restructuring in culture, with Bifidobacterium and Enterococcus blooming and driving sex-divergent shifts in functional capacity and a substantial amplification of the resistome. Together, these findings validate a high-throughput stool-derived in vitro culture as a tractable proxy for an antibiotic altered gut microbiome to screen interventions.