For pancreatic β-cells to secrete insulin in response to elevated blood glucose, insulin granules retained within the subplasmalemmal space must be transported to sites of secretion on the plasma membrane. Using a combination of super-resolution STORM imaging and live cell TIRF microscopy we investigate how the organization and dynamics of the actin and microtubule cytoskeletons in INS-1 β-cells contribute to this process. GFP-labeled insulin granules display 3 different modes of motion (stationary, diffusive-like, and directed). Diffusive-like motion dominates in basal, low glucose conditions. Upon glucose stimulation no gross rearrangement of the actin cytoskeleton is observed but there are increases in the 1) rate of microtubule polymerization; 2) rate of diffusive-like motion; and 3) proportion of granules undergoing microtubule-based directed motion. By pharmacologically perturbing the actin and microtubule cytoskeletons, we determine that microtubule-dependent granule transport occurs within the subplasmalemmal space and that the actin cytoskeleton limits this transport in basal conditions, when insulin secretion needs to be inhibited.

Intracellular cargo transport relies on myosin Va molecular motor ensembles to travel along the cell's three-dimensional (3D) highway of actin filaments. At actin filament intersections, the intersecting filament is a structural barrier to and an alternate track for directed cargo transport. Here we use 3D super-resolution fluorescence imaging to determine the directional outcome (that is, continues straight, turns or terminates) for an ∼10 motor ensemble transporting a 350 nm lipid-bound cargo that encounters a suspended 3D actin filament intersection in vitro. Motor–cargo complexes that interact with the intersecting filament go straight through the intersection 62% of the time, nearly twice that for turning. To explain this, we develop an in silico model, supported by optical trapping data, suggesting that the motors' diffusive movements on the vesicle surface and the extent of their engagement with the two intersecting actin tracks biases the motor–cargo complex on average to go straight through the intersection.

We measured myosin crossbridge detachment rate and the rates of MgADP release and MgATP binding in mouse and rat myocardial strips bearing one of the two cardiac myosin heavy chain (MyHC) isoforms. Mice and rats were fed an iodine-deficient, propylthiouracil diet resulting in ~100% expression of β-MyHC in the ventricles. Ventricles of control animals expressed ~100% α-MyHC. Chemically-skinned myocardial strips prepared from papillary muscle were subjected to sinusoidal length perturbation analysis at maximum calcium activation pCa 4.8 and 17°C. Frequency characteristics of myocardial viscoelasticity were used to calculate crossbridge detachment rate over 0.01 to 5 mM [MgATP]. The rate of MgADP release, equivalent to the asymptotic value of crossbridge detachment rate at high MgATP, was highest in mouse α-MyHC (111.4±6.2 s−1) followed by rat α-MyHC (65.0±7.3 s−1), mouse β-MyHC (24.3±1.8 s−1) and rat β-MyHC (15.5±0.8 s−1). The rate of MgATP binding was highest in mouse α-MyHC (325±32 mM−1.s−1) then mouse β-MyHC (152±23 mM−1.s−1), rat α-MyHC (108±10 mM−1.s−1) and rat β-MyHC (55±6 mM−1.s−1). Because the events of MgADP release and MgATP binding occur in a post power-stroke state of the myosin crossbridge, we infer that MgATP release and MgATP binding must be regulated by isoform- and species-specific structural differences located outside the nucleotide binding pocket, which is identical in sequence for these four myosins. We postulate that differences in the stiffness profile of the entire myosin molecule, including the thick filament and the myosin-actin interface, are primarily responsible for determining the strain on the nucleotide binding pocket and the subsequent differences in the rates of nucleotide release and binding observed among the four myosins examined here.

A Thomson parabola ion spectrometer has been designed for use at the Multiterawatt (MTW) laser facility at the Laboratory for Laser Energetics (LLE) at the University of Rochester. This device uses parallel electric and magnetic fields to deflect particles of a given mass-to-charge ratio onto parabolic curves on the detector plane. Once calibrated, the position of the ions on the detector plane can be used to determine the particle energy. The position dispersion of both the electric and magnetic fields of the Thomson parabola was measured using monoenergetic proton and alpha particle beams from the SUNY Geneseo 1.7 MV tandem Pelletron accelerator. The sensitivity of Fujifilm BAS-TR imaging plates, used as a detector in the Thomson parabola, was also measured as a function of the incident particle energy over the range from 0.6 MeV to 3.4 MeV for protons and deuterons and from 0.9 MeV to 5.4 MeV for alpha particles. The device was used to measure the energy spectrum of laser-produced protons at MTW.

Activated ezrin-radixin-moesin (ERM) proteins link the plasma membrane to the actin cytoskeleton to generate apical structures, including microvilli. Among many kinases implicated in ERM activation are the homologues LOK and SLK. CRISPR/Cas9 was used to knock out all ERM proteins or LOK/SLK in human cells. LOK/SLK knockout eliminates all ERM-activating phosphorylation. The apical domains of cells lacking LOK/SLK or ERMs are strikingly similar and selectively altered, with loss of microvilli and with junctional actin replaced by ectopic myosin-II–containing apical contractile structures. Constitutively active ezrin can reverse the phenotypes of either ERM or LOK/SLK knockouts, indicating that a central function of LOK/SLK is to activate ERMs. Both knockout lines have elevated active RhoA with concomitant enhanced myosin light chain phosphorylation, revealing that active ERMs are negative regulators of RhoA. As RhoA-GTP activates LOK/SLK to activate ERM proteins, the ability of active ERMs to negatively regulate RhoA-GTP represents a novel local feedback loop necessary for the proper apical morphology of epithelial cells.