SUMMARY Changes in neuronal activity create local and transient changes in energy demands at synapses. Here we discover a metabolic compartment that forms in vivo near synapses to meet local energy demands and support synaptic function in Caenorhabditis elegans neurons. Under conditions of energy stress, glycolytic enzymes redistribute from a diffuse localization in the cytoplasm to a punctate localization adjacent to synapses. Glycolytic enzymes colocalize, suggesting the ad hoc formation of a glycolysis compartment, or a ‘glycolytic metabolon’, that can maintain local levels of ATP. Local formation of the glycolytic metabolon is dependent on presynaptic scaffolding proteins, and disruption of the glycolytic metabolon blocks the synaptic vesicle cycle, impairs synaptic recovery, and affects locomotion. Our studies indicate that under energy stress conditions, energy demands in C. elegans synapses are met locally through the assembly of a glycolytic metabolon to sustain synaptic function and behavior.

SUMMARYChanges in neuronal activity create local and transient changes in energy demands at synapses. Here we discover a metabolic compartment that forms in vivo near synapses to meet local energy demands and support synaptic function in Caenorhabditis elegans neurons. Under conditions of energy stress, glycolytic enzymes redistribute from a diffuse localization in the cytoplasm to a punctate localization adjacent to synapses. Glycolytic enzymes colocalize, suggesting the ad hoc formation of a glycolysis compartment, or a 'glycolytic metabolon', that can maintain local levels of ATP. Local formation of the glycolytic metabolon is dependent on presynaptic scaffolding proteins, and disruption of the glycolytic metabolon blocks the synaptic vesicle cycle, impairs synaptic recovery, and affects locomotion. Our studies indicate that under energy stress conditions, energy demands in C. elegans synapses are met locally through the assembly of a glycolytic metabolon to sustain synaptic function and behavior. †

BackgroundThe sugar sensing and carbon catabolite repression in Baker’s yeast Saccharomyces cerevisiae is governed by three major signaling pathways that connect carbon source recognition with transcriptional regulation. Here we present a screening method based on a non-invasive in vivo reporter system for real-time, single-cell screening of the sugar signaling state in S. cerevisiae in response to changing carbon conditions, with a main focus on the response to glucose and xylose.ResultsThe artificial reporter system was constructed by coupling a green fluorescent protein gene (yEGFP3) downstream of endogenous yeast promoters from the Snf3p/Rgt2p, SNF1/Mig1p and cAMP/PKA signaling pathways: HXT1p/2p/4p; SUC2p, CAT8p; TPS1p/2p and TEF4p respectively. A panel of eight biosensors strains was generated by single copy chromosomal integration of the different constructs in a W303-derived strain. The signaling biosensors were validated for their functionality with flow cytometry by comparing the fluorescence intensity (FI) response in the presence of high or nearly depleted glucose to the known induction/repression conditions of the eight different promoters. The FI signal correlated with the known patterns of the selected promoters while maintaining a non-invasive property on the cellular phenotype, as was demonstrated in terms of growth, metabolites and enzyme activity.ConclusionsOnce verified, the sensors were used to evaluate the signaling response to varying conditions of extracellular glucose, glycerol and xylose by screening in 96-well microtiter plates. We show that these yeast strains, which do not harbor any recombinant pathways for xylose utilization, are lacking a signaling response for extracellular xylose. However, for the HXT2p/4p sensors, a shift in the flow cytometry population dynamics indicated that internalized xylose does affect the signaling. These results suggest that the previously observed effects of this pentose on the S. cerevisiae physiology and gene regulation can be attributed to xylose and not only to a lack of glucose.Electronic supplementary materialThe online version of this article (doi:10.1186/s12934-016-0580-x) contains supplementary material, which is available to authorized users.

Production of chemicals in microbes often employs potent biosynthetic enzymes, which can interact with the microbial native metabolism to affect cell fitness and product yield. However, production optimization largely relies on data collected from wild-type strains in the absence of metabolic perturbations, thus limiting their relevance to specific conditions. Here, we address this issue by coupling cell fitness to the production of thiamine diphosphate in Escherichia coli using a synthetic RNA biosensor. We use this strategy to interrogate a library of transposon mutants and elucidate the native gene network influencing both cell fitness and thiamine production. Ultimately, we identify effectors of the OxyR-Fur stress response that limit thiamine biosynthesis via alternative regulation of iron storage and Fe-S cluster inclusion in enzymes. This study presents a new approach for the reliable high-throughput identification of genetic targets of both known and unknown function that are directly relevant to a specific biosynthetic process.

Bile acids are essential molecules produced by vertebrates that are involved in several physiological roles, including the uptake of nutrients. Bacterial isolates capable of producing bile acids de novo have been identified and characterized. Such isolates may provide access to novel biochemical pathways suitable for the design of microbial cell factories. Here, we further characterized the ability of Maribacter dokdonensis, Dokdonia donghaensis, and Myroides pelagicus to produce bile acids. Contrary to previous reports, we did not observe de novo production of bile acids by these isolates. Instead, we found that these isolates deconjugated the amino acid moiety of bile acids present in the growth medium used in previous reports. Through genomic analysis, we identified putative bile salt hydrolases, which could be responsible for the different bile acid modifications observed. Our results challenge the hypothesis of de novo microbial bile acid production, while further demonstrating the diverse capacity of bacteria to modify bile acids.