Several aquaporin (AQP) water channels are short-term regulated by the messenger cyclic adenosine monophosphate (cAMP), including AQP3. Bulk measurements show that cAMP can change diffusive properties of AQP3; however, it remains unknown how elevated cAMP affects AQP3 organization at the nanoscale. Here we analyzed AQP3 nano-organization following cAMP stimulation using photoactivated localization microscopy (PALM) of fixed cells combined with pair correlation analysis. Moreover, in live cells, we combined PALM acquisitions of single fluorophores with single-particle tracking (spt-PALM). These analyses revealed that AQP3 tends to cluster and that the diffusive mobility is confined to nanodomains with radii of ∼150 nm. This domain size increases by ∼30% upon elevation of cAMP, which, however, is not accompanied by a significant increase in the confined diffusion coefficient. This regulation of AQP3 organization at the nanoscale may be important for understanding the mechanisms of water AQP3-mediated water transport across plasma membranes.

Optical and electron microscopy have made tremendous inroads toward understanding the complexity of the brain. However, optical microscopy offers insufficient resolution to reveal subcellular details, and electron microscopy lacks the throughput and molecular contrast to visualize specific molecular constituents over millimeter-scale or larger dimensions. We combined expansion microscopy and lattice light-sheet microscopy to image the nanoscale spatial relationships between proteins across the thickness of the mouse cortex or the entire Drosophila brain. These included synaptic proteins at dendritic spines, myelination along axons, and presynaptic densities at dopaminergic neurons in every fly brain region. The technology should enable statistically rich, large-scale studies of neural development, sexual dimorphism, degree of stereotypy, and structural correlations to behavior or neural activity, all with molecular contrast.

Cell reprogramming is thought to be associated with a full metabolic switch from an oxidative- to a glycolytic-based metabolism. However, neither the dynamics nor the factors controlling this metabolic switch are fully understood. By using cellular, biochemical, protein array, metabolomic, and respirometry analyses, we found that c-MYC establishes a robust bivalent energetics program early in cell reprogramming. Cells prone to undergo reprogramming exhibit high mitochondrial membrane potential and display a hybrid metabolism. We conclude that MYC proteins orchestrate a rewiring of somatic cell metabolism early in cell reprogramming, whereby somatic cells acquire the phenotypic plasticity necessary for their transition to pluripotency in response to either intrinsic or external cues.

In eukaryotic cells, organelles and the cytoskeleton undergo highly dynamic yet organized interactions capable of orchestrating complex cellular functions. Visualizing these interactions requires noninvasive, long-duration imaging of the intracellular environment at high spatiotemporal resolution and low background. To achieve these normally opposing goals, we developed grazing incidence structured illumination microscopy (GI-SIM) that is capable of imaging dynamic events near the basal cell cortex at 97-nm resolution and 266 frames/s over thousands of time points. We employed multi-color GI-SIM to characterize the fast dynamic interactions of diverse organelles and the cytoskeleton, shedding new light on the complex behaviors of these structures. Precise measurements of microtubule growth or shrinkage events helped distinguish among models of microtubule dynamic instability. Analysis of endoplasmic reticulum (ER) interactions with other organelles or microtubules uncovered new ER remodeling mechanisms, such as hitchhiking of the ER on motile organelles. Finally, ER-mitochondria contact sites were found to promote both mitochondrial fission and fusion.

Type I collagen is the main component of bone matrix and other connective tissues. Rerouting of its procollagen precursor to a degradative pathway is crucial for osteoblast survival in pathologies involving excessive intracellular buildup of procollagen that is improperly folded and/or trafficked. What cellular mechanisms underlie this rerouting remains unclear. To study these mechanisms, we employed live-cell imaging and correlative light and electron microscopy (CLEM) to examine procollagen trafficking both in wild-type mouse osteoblasts and osteoblasts expressing a bone pathology-causing mutant procollagen. We found that although most procollagen molecules successfully trafficked through the secretory pathway in these cells, a subpopulation did not. The latter molecules appeared in numerous dispersed puncta colocalizing with COPII subunits, autophagy markers and ubiquitin machinery, with more puncta seen in mutant procollagen-expressing cells. Blocking endoplasmic reticulum exit site (ERES) formation suppressed the number of these puncta, suggesting they formed after procollagen entry into ERESs. The punctate structures containing procollagen, COPII, and autophagic markers did not move toward the Golgi but instead were relatively immobile. They appeared to be quickly engulfed by nearby lysosomes through a bafilomycin-insensitive pathway. CLEM and fluorescence recovery after photobleaching experiments suggested engulfment occurred through a noncanonical form of autophagy resembling microautophagy of ERESs. Overall, our findings reveal that a subset of procollagen molecules is directed toward lysosomal degradation through an autophagic pathway originating at ERESs, providing a mechanism to remove excess procollagen from cells.

During starvation the transcriptional activation of catabolic processes is induced by the nuclear translocation and consequent activation of transcription factor EB (TFEB), a master modulator of autophagy and lysosomal biogenesis. However, how TFEB is inactivated upon nutrient refeeding is currently unknown. Here we show that TFEB subcellular localization is dynamically controlled by its continuous shuttling between the cytosol and the nucleus, with the nuclear export representing a limiting step. TFEB nuclear export is mediated by CRM1 and is modulated by nutrient availability via mTOR-dependent hierarchical multisite phosphorylation of serines S142 and S138, which are localized in proximity of a nuclear export signal (NES). Our data on TFEB nucleo-cytoplasmic shuttling suggest an unpredicted role of mTOR in nuclear export.

The saga of fluorescence recovery after photobleaching (FRAP) illustrates how disparate technical developments impact science. Starting with the classic 1976 Axelrod et al. work in Biophysical Journal, FRAP (originally fluorescence photobleaching recovery) opened the door to extraction of quantitative information from photobleaching experiments, laying the experimental and theoretical groundwork for quantifying both the mobility and the mobile fraction of a labeled population of proteins. Over the ensuing years, FRAP's reach dramatically expanded, with new developments in GFP technology and turn-key confocal microscopy, which enabled measurement of protein diffusion and binding/dissociation rates in virtually every compartment within the cell. The FRAP technique and data catalyzed an exchange of ideas between biophysicists studying membrane dynamics, cell biologists focused on intracellular dynamics, and systems biologists modeling the dynamics of cell activity. The outcome transformed the field of cellular biology, leading to a fundamental rethinking of long-held theories of cellular dynamism. Here, we review the pivotal FRAP studies that made these developments and conceptual changes possible, which gave rise to current models of complex cell dynamics.

Different multicellular organisms undergo cell-cell fusion to form functional syncytia that support specialized functions necessary for proper development and survival. For years, monitoring the structural consequences of this process using live-cell imaging has been challenging due to the unpredictable timing of cell fusion events in tissue systems. Here we present a triggered vesicular stomatitis virus G-protein (VSV-G)-mediated cell-cell fusion assay that can be used to synchronize fusion between cells. This allows the study of cellular changes that occur during cell fusion. The process is induced using a fast wash of low pH isotonic buffer, promoting the fusion of plasma membranes of two or more adjacent cells within seconds. This approach is suitable for studying mixing of small cytoplasmic molecules between fusing cells as well as changes in organelle distribution and dynamics. © 2018 by John Wiley & Sons, Inc.

Eukaryotic cells are organized into membrane-bound organelles. These organelles communicate with one another through vesicular trafficking pathways and membrane contact sites (MCSs). MCSs are sites of close apposition between two or more organelles that play diverse roles in the exchange of metabolites, lipids and proteins. Organelle interactions at MCSs also are important for organelle division and biogenesis. For example, the division of several organelles, including mitochondria and endosomes, seem to be regulated by contacts with the endoplasmic reticulum (ER). Moreover, the biogenesis of autophagosomes and peroxisomes involves contributions from the ER and multiple other cellular compartments. Thus, organelle-organelle interactions allow cells to alter the shape and activities of their membrane-bound compartments, allowing them to cope with different developmental and environmental conditions.

This protocol describes how to apply appropriate pharmacological controls to induce mitochondrial fusion or fission in studies of mitochondria morphology for four different mammalian cell types, HepG2 human liver hepatocellular carcinoma cells, MCF7 human breast adenocarcinoma cells, HEK293 human embryonic kidney cells, and collagen sandwich culture of primary rat hepatocytes. The protocol provides methods of treating cells with these pharmacological controls, staining mitochondria with commercially available MitoTracker Green and TMRE dyes, and imaging the mitochondrial morphology in live cells using a confocal fluorescent microscope. It also describes the cell culture methods needed for this protocol. © 2018 by John Wiley & Sons, Inc.