The fidelity of signal transduction requires spatiotemporal control of the production of signaling agents. Phosphatidic acid (PA) is a pleiotropic lipid second messenger whose modes of action differ based on upstream stimulus, biosynthetic source, and site of production. How cells regulate the local production of PA to effect diverse signaling outcomes remains elusive. Unlike other second messengers, sites of PA biosynthesis cannot be accurately visualized with subcellular precision. Here, we describe a rapid, chemoenzymatic approach for imaging physiological PA production by phospholipase D (PLD) enzymes. Our method capitalizes on the remarkable discovery that bulky, hydrophilic trans-cyclooctene–containing primary alcohols can supplant water as the nucleophile in the PLD active site in a transphosphatidylation reaction of PLD’s lipid substrate, phosphatidylcholine. The resultant trans-cyclooctene–containing lipids are tagged with a fluorogenic tetrazine reagent via a no-rinse, inverse electron-demand Diels–Alder (IEDDA) reaction, enabling their immediate visualization by confocal microscopy in real time. Strikingly, the fluorescent reporter lipids initially produced at the plasma membrane (PM) induced by phorbol ester stimulation of PLD were rapidly internalized via apparent nonvesicular pathways rather than endocytosis, suggesting applications of this activity-based imaging toolset for probing mechanisms of intracellular phospholipid transport. By instead focusing on the initial 10 s of the IEDDA reaction, we precisely pinpointed the subcellular locations of endogenous PLD activity as elicited by physiological agonists of G protein-coupled receptor and receptor tyrosine kinase signaling. These tools hold promise to shed light on both lipid trafficking pathways and physiological and pathological effects of localized PLD signaling.

Strategies to visualize cellular membranes with light microscopy are restricted by the diffraction limit of light, which far exceeds the dimensions of lipid bilayers. Here, we describe a method for super-resolution imaging of metabolically labeled phospholipids within cellular membranes. Guided by the principles of expansion microscopy, we develop an all-small molecule approach that enables direct chemical anchoring of bioorthogonally labeled phospholipids into a hydrogel network and is capable of super-resolution imaging of cellular membranes. We apply this method, termed lipid expansion microscopy (LExM), to visualize organelle membranes with precision, including a unique class of membrane-bound structures known as nuclear invaginations. Compatible with standard confocal microscopes, LExM will be widely applicable for super-resolution imaging of phospholipids and cellular membranes in numerous physiological contexts.

A cognitively intensive companion service course has been introduced to the main fall general chemistry class at Cornell University. For years 2015 and 2016, priority students (those from groups under-represented and economically disadvantaged) show respectively improvement of +0.67 and +0.51 standard deviations in final course grade compared to priority students not in the program. Non-priority students show respectively a +0.66 and +0.62 standard deviation improvement. Progressive improvement (as measured by higher than expected Final Exam scores than what would have been expected solely from a given student’s earlier Exam 1 score) demonstrates conclusively the service course’s role in the enhanced outcomes. Progressive retention (as measured by the following year fall semester’s organic chemistry exam scores compared to what would have been expected based on a given student’s general chemistry final exam score) demonstrates that, on the average, the earlier observed progressive improvement is significantly retained in a chemistry course one year later. Preliminary retention statistics suggest a significant increase in first year to second year retention. A meta analysis of results from previously reported chemistry service courses indicate that such performance gains are difficult to achieve and hence common elements of the few effective programs may be of high value to the STEM education community.

Strategies to visualize cellular membranes with light microscopy are restricted by the diffraction limit of light, which far exceeds the dimensions of lipid bilayers. Here, we describe a method for super-resolution imaging of metabolically labeled phospholipids within cellular membranes. Guided by the principles of expansion microscopy, we develop an approach featuring cell-permeable reagents that enables direct chemical anchoring of bioorthogonally labeled phospholipids into a hydrogel network and is capable of tunable, isotropic expansion, thus facilitating super-resolution imaging of cellular membranes. We apply this method, termed lipid expansion microscopy, to visualize organelle membranes with precision, including a unique class of membrane-bound structures known as nuclear invaginations. As it is compatible with standard confocal microscopes, lipid expansion microscopy will be widely applicable for super-resolution imaging of phospholipids and cellular membranes in numerous physiological contexts.

Membrane architectures whose dimensions and features are smaller than the diffraction limit of light orchestrate diverse cellular events such as lipid transport, vesicle formation, and calcium signaling. These structures, which include membrane invaginations, organelle contact sites, and membrane microdomains, are primarily composed of phospholipids, making methods to visualize these biomolecules vital to our understanding of cellular function. Techniques to accurately image phospholipid‐containing structures with fluorescence microscopy are challenged by the diffusion of lipids within the bilayer, even in fixed samples. Expansion microscopy (ExM) utilizes hydrogel formation to fix biomolecules in place and swell samples to produce high‐resolution images of protein‐ and nucleic acid‐containing cellular structures 30‐70 nm in size. Using chemical reporter metabolites and a novel multifunctional fluorophore probe, here we present Lipid Expansion Microscopy (LExM), which enables the high‐resolution imaging of phospholipids with a tunable expansion factor using the principles of ExM. We will present key synthetic and technological advances critical to the development of LExM as well as its application to visualize nanoscale membrane structures, phospholipid‐organelle colocalization, and the spatial component of flux through specific lipid biosynthetic pathways within intact cells.