Adsorption of Sub-Micron Amphiphilic Dumbbells to Fluid Interfaces

We investigate the adsorption of submicrometer bulk-synthesized polymer dumbbells to oil–water interfaces using freeze-fracture, shadow-casting (FreSCa) cryo-scanning electron microscopy. We find that the dumbbells are amphiphilic and adsorb to the interface with a preferred orientation. Most particles adsorb in a tilted configuration, with the polar and apolar lobes intersecting the interface and pointing toward the water and oil, respectively. Some particles adsorb with only one lobe attached to the interface. Moreover, we find that each lobe has a preferred angle of contact with the interface, identical in all observed configurations. A simple geometrical calculation using these contact angles accurately predicts the dominant configuration of particles at the interface. This calculation provides insight into how the shape and composition of dumbbells can be tuned to stand upright and pack efficiently on curved interfaces

Elastic Coupling of Nascent apCAM Adhesions to Flowing Actin Networks

<div><p>Adhesions are multi-molecular complexes that transmit forces generated by a cell’s acto-myosin networks to external substrates. While the physical properties of some of the individual components of adhesions have been carefully characterized, the mechanics of the coupling between the cytoskeleton and the adhesion site as a whole are just beginning to be revealed. We characterized the mechanics of nascent adhesions mediated by the immunoglobulin-family cell adhesion molecule apCAM, which is known to interact with actin filaments. Using simultaneous visualization of actin flow and quantification of forces transmitted to apCAM-coated beads restrained with an optical trap, we found that adhesions are dynamic structures capable of transmitting a wide range of forces. For forces in the picoNewton scale, the nascent adhesions’ mechanical properties are dominated by an elastic structure which can be reversibly deformed by up to 1 µm. Large reversible deformations rule out an interface between substrate and cytoskeleton that is dominated by a number of stiff molecular springs in parallel, and favor a compliant cross-linked network. Such a compliant structure may increase the lifetime of a nascent adhesion, facilitating signaling and reinforcement.</p></div

Membrane deforms but is too weak to explain observed traction forces.

<p>(A) Schematic representation of the setup: a 2 µm diameter apCAM-coated bead with two 100 nm fluorescent beads irreversibly attached on its surface is placed on the membrane of growth cone treated with 5 µM latrunculin B. (B) Displacement over time of the apCAM-coated bead (green) and the two fluorescent beads (red and blue) along the x-axis. (C) Positions in the x-y plane of the two fluorescent beads in the frame of reference of the center of mass of the apCAM-coated bead (green circle indicates the bead’s circumference). (D) Translational mean squared displacement of the apCAM-coated bead (green) and rotational mean squared displacements of the twisting angle (red) and rolling angle (blue). (E) Schematic representation of the setup: a 2 µm diameter apCAM-coated bead with a 100 nm diameter fluorescent bead irreversibly attached on its surface is placed on the membrane of a control growth cone. (F) Displacement over time of the apCAM-coated bead (green) and the small fluorescent bead (orange) along the actin flow. Green force scale bar applies only to the green trace. Rolling angle (blue) calculated from the small bead motion respect to the apCAM-coated bead motion. (G) Breakage force histograms calculated from centroids of the apCAM-coated bead (blue) and the fluorescent beads on its surface (red).</p

Schematic mechanical model of a nascent adhesion.

<p>The trapped bead alternates between frictional coupling with retrograde flow (left) and intermittent coupling to elastic intracellular structure (rignt).</p

Elastic properties of nascent adhesions.

<p>(<i>A</i>) Top panel: Bead (green) vs. trap (red) relative positions. Flow-coupled apCAM-coated bead (green) is successively clamped at different constant forces by changing the separation between the bead and the center of the optical trap (red). Middle panel: flow-coupled bead is first released from the optical trap (“unload” arrow) and then force-clamped (“reload” arrow) at 30, 20 and 10 pN successively. Bottom panel: bead velocity exhibits a transient for each change of force indicated by the black arrows (B) Steady bead velocity under constant force normalized to bead velocity under zero force. Same color symbols correspond to same bead clamped at different forces. For each particular experiment the laser power of the trap is first set and then the distance between trap and bead was adjusted. (C) Jump sizes of flow-coupled beads when released from (Δ<i>x</i>_{<i>unload</i>}) and reloaded by(Δ<i>x</i>_{<i>reload</i>}) optical traps. Same color symbols correspond to different events with the same bead and cell. (D-E) When a flow-coupled bead (green) was displaced by 0.7 µm from the trap center (red), the bead was released. Dashed lines in D and E depict the extrapolated position of actin features moving with retrograde flow. (F) In-plane force-displacement curve of nascent adhesion. Displacement is defined as the distance between the bead and the blue dashed line in D representing the trajectory of an unrestrained flow-coupled bead. (G) Force-velocity of the flow-coupled bead during relaxation calculated using bead trajectory in <i>F</i> and force-displacement curve in <i>H</i>.</p

Relaxation of bead-substrates released from optical tweezers.

<p>(<i>A</i>) <b>apCAM-coated bead (green) is restrained 10 s in a stationary trap (red)</b>. </p> <p>The trap is then turned off and the bead moves with actin retrograde flow. A closed-loop trapping system returns the bead to its original position after it has been displaced by about 3 µm. Shaded and white areas represent periods where the optical tweezers are on and off, respectively. An enlargement showing bead displacement kinetics before and after release (boxed region) is shown below. (<i>B</i>) Histogram of transient displacements defined as the distance traveled by the bead during the first 0.8 s after turning laser off. Inset: histogram of the steady state bead velocity after each transient event: 0.8 s after release, the steady-state velocity was measured over the next 8 to 12 s. (<i>C</i>) Transient displacements versus force applied to beads just before turning off the laser. Note that 100% of beads exhibited elastic relaxation when force was >10 pN. (<i>D</i>) Histogram of transient displacement for a growth cone treated with 50 µM blebbistatin. Inset: histogram of the steady state bead velocity after each transient. (<i>E</i>) Bar graph showing the percentage of beads exhibiting no jump (blue), and the percentage exhibiting a jump (red), in control or blebbistatin-treated experiments. The first and third plots include all events after laser turn-off, while the second and fourth plots include events where the pre-stress exceeded 10 pN.</p

Correlation of bead movements with actin flow.

<p>(A) Schematic depicting growth cone cytoplasmic domains and an apCAM coated bead restrained against retrograde flow on the surface of an <i>Aplysia</i> bag cell growth cone using an optical trap. (B) DIC image of trapped apCAM-coated bead. (C) F-actin labeled with fluorescent phalloidin imaged with confocal microscopy. (D) Kymograph of actin speckle displacement sampled along the yellow line in (C). Green line shows trapped bead trajectory. Magenta lines show F-actin retrograde flow speed. Red arrows denote breakage events. The yellow arrow indicates the point of maximum optical force where the bead escapes from the trap. (E) Time-lapse montage DIC (top) and phalloidin-tagged actin (bottom) of a bead forming an adhesion, coupling to retrograde flow, a breakage event followed by recoupling to actin flow and eventual bead escape. Bead positions are superimposed on the phalloidin montage (green circles). Red arrow: breakage event; white/black arrow, bead escape. Time and space for C-D indicated by respective scale bars.</p

Structural Diversity of Arthropod Biophotonic Nanostructures Spans Amphiphilic Phase-Space

Many organisms, especially arthropods, produce vivid interference colors using diverse mesoscopic (100–350 nm) integumentary biophotonic nanostructures that are increasingly being investigated for technological applications. Despite a century of interest, precise structural knowledge of many biophotonic nanostructures and the mechanisms controlling their development remain tentative, when such knowledge can open novel biomimetic routes to facilely self-assemble tunable, multifunctional materials. Here, we use synchrotron small-angle X-ray scattering and electron microscopy to characterize the photonic nanostructure of 140 integumentary scales and setae from ∼127 species of terrestrial arthropods in 85 genera from 5 orders. We report a rich nanostructural diversity, including triply periodic bicontinuous networks, close-packed spheres, inverse columnar, perforated lamellar, and disordered spongelike morphologies, commonly observed as stable phases of amphiphilic surfactants, block copolymer, and lyotropic lipid–water systems. Diverse arthropod lineages appear to have independently evolved to utilize the self-assembly of infolding lipid-bilayer membranes to develop biophotonic nanostructures that span the phase-space of amphiphilic morphologies, but at optical length scales

Surface Passivation Method for the Super-repellence of Aqueous Macromolecular Condensates

Solutions of macromolecules can undergo liquid–liquid phase separation to form droplets with ultralow surface tension. Droplets with such low surface tension wet and spread over common surfaces such as test tubes and microscope slides, complicating in vitro experiments. The development of a universal super-repellent surface for macromolecular droplets has remained elusive because their ultralow surface tension requires low surface energies. Furthermore, the nonwetting of droplets containing proteins poses additional challenges because the surface must remain inert to a wide range of chemistries presented by the various amino acid side chains at the droplet surface. Here, we present a method to coat microscope slides with a thin transparent hydrogel that exhibits complete dewetting (contact angles θ ≈ 180°) and minimal pinning of phase-separated droplets in aqueous solution. The hydrogel is based on a swollen matrix of chemically cross-linked polyethylene glycol diacrylate of molecular weight 12 kDa (PEGDA), and can be prepared with basic chemistry laboratory equipment. The PEGDA hydrogel is a powerful tool for in vitro studies of weak interactions, dynamics, and the internal organization of phase-separated droplets in aqueous solutions

Surface Passivation Method for the Super-repellence of Aqueous Macromolecular Condensates

Solutions of macromolecules can undergo liquid–liquid phase separation to form droplets with ultralow surface tension. Droplets with such low surface tension wet and spread over common surfaces such as test tubes and microscope slides, complicating in vitro experiments. The development of a universal super-repellent surface for macromolecular droplets has remained elusive because their ultralow surface tension requires low surface energies. Furthermore, the nonwetting of droplets containing proteins poses additional challenges because the surface must remain inert to a wide range of chemistries presented by the various amino acid side chains at the droplet surface. Here, we present a method to coat microscope slides with a thin transparent hydrogel that exhibits complete dewetting (contact angles θ ≈ 180°) and minimal pinning of phase-separated droplets in aqueous solution. The hydrogel is based on a swollen matrix of chemically cross-linked polyethylene glycol diacrylate of molecular weight 12 kDa (PEGDA), and can be prepared with basic chemistry laboratory equipment. The PEGDA hydrogel is a powerful tool for in vitro studies of weak interactions, dynamics, and the internal organization of phase-separated droplets in aqueous solutions