|Diverse cells throughout the eukaryotic tree use actin to make protrusions for motility. Chytrid fungi, specifically, use branched actin networks to quickly build pseudopods and rapidly crawl.|
In the Mullins lab, we study actin and the cell cytoskeleton. I am the microscope specialist, helping researchers use the fluorescence microscopes to their highest potential. With Lil Fritz-Laylin, I studied pseudopod formation and rapid crawling in chytrid fungi cells.
|The supported lipid bilayer is an attractive tool for studying live cells because it is a model relatively simple to modify, image, and probe using modern lithography and microscopy techniques. This is a simplified cartoon of a living T cell interfaced with a supported lipid bilayer, including metal barriers that restrict motion of phospholipids and membrane proteins. In a living organism, antigen-presenting cells digest components of invading bodies and display those components on their surface. T cell receptors bind to such antigens and signal to the T cell that the antigens are either self or invasive. The interface between the T cell and the antigen-presenting cell is the immunological synapse. Support lipid bilayers make it easier to image and study the immunological synapse, because it eliminates the need for the antigen-presenting cell. Moreover, barriers to lateral diffusion (i.e. spatial mutations) can be introduced, which permit manipulating the location and density of the receptors, antigens, and adhesion complexes.|
In the lab of Professor Jay T. Groves at Berkeley, I used fluorescence microscopy and supported lipid bilayers as tools to study dynamics during cell-cell interactions. These studies revealed that mechanical properties of a cell-cell interface can influence endocytosis of receptor tyrosine kinases.
There is a persistent need for small-molecule fluorescent labels optimized for single-molecule imaging in the cellular environment. Application of these labels comes with a set of strict requirements: strong absorption, efficient and stable emission, water solubility and membrane permeability, low background emission, and red-shifted absorption to avoid cell autofluorescence. I have designed and characterized several fluorophores, termed DCDHF fluorophores, for use in live-cell imaging based on the push-pull design: an amine donor group and a 2-dicyanomethylene-3-cyano-2,5-dihydrofuran (DCDHF) acceptor group, separated by a π-rich conjugated network.1
The photophysics of DCDHF fluorophores can be synthetically tuned. For instance, increasing the size of the π-conjugated linker (e.g. from benzene, to naphthalene, to anthracene) redshifts the absorption and emission.
Also, DCDHF exhibit the special characteristic that they brighten upon rigidization. The vials show liquid and frozen solutions of the same concentrations; in the rigid ice, the fluorophore is significantly brighter.
In general, the DCDHF fluorophores are comparatively photostable, sensitive to local environment, and their chemistries and photophysics are tunable to optimize absorption wavelength, membrane affinity, and solubility. The DCDHFs provide a new class of bright photoactivatable small-molecule fluorophores, which are needed for imaging in living cells.2
I reengineered a red-emitting DCDHF push-pull fluorophore so that it is dark until photoactivated with a short burst of low-intensity violet light. Photoactivation of the dark fluorogen leads to conversion of an azide to an amine, which shifts the absorption to long wavelengths.3-4
|The azido DCDHF fluorophore (left) is dark until photoconverted to an amino DCDHF (right). The photoactivation was demonstrated to occur in solution and even inside living cells.|
After photoactivation, the fluorophore is bright and photostable enough to be imaged on the single-molecule level in living cells. This proof-of-principle demonstration provides a new class of bright photoactivatable fluorophores, as are needed for super-resolution imaging schemes that require active control of single molecule emission.
|The photochemical conversion of the azide to an electron-donating amine regenerates the push-pull character of the chromophore; and thus, the red-shifted charge-transfer peak grows in. The inset show that after photoactivation, the fluorescence increase significantly.|
Recently, a colleague in the Moerner lab used an azido DCDHF to perform 3D super-resolution measurements in a polymer film. Because the fluorophore emits millions of photons, it was possible to localize single molecules in three dimensions to less than 20 nm.5 We have also imaged protein structures below the diffraction limit in living bacteria cells using targeted DCDHF dyes.6
I demonstrated that cylindrical molecular brushes, each with a gradient of grafting density along the backbone, transition from rodlike to tadpole conformations. Using atomic force microscopy (AFM), I observed a coexistence of two conformational phases within individual molecules adsorbed on a mica substrate. I made these observations by compressing monolayers on the surface of water and then transferring a sample of the monolayer to a mica substrate for AFM studies.7
There is a competition between entropy (which wants the sidechains to desorb and allow the backbone to coil) and enthalpy (which encourages sidechains to adsorb to the surface and spread out). The density of the sidechain grafting as well as the pressure determines whether entropy or enthalpy wins.
In gradient brushes, therefore, there is a pressure at which one side of the brush collapses into a coil while the loosely grafted end stays adsorbed and the backbone remains extended.
Upon compression, the rod/globule transition occurs at the end where the brush is densely grafted, leaving a molecule with a globular "head" and an extended "tail" a so-called tadpole conformation. My research demonstrated the asymmetric changes in the molecular conformation, which is one of the prerequisites for directed motion.
|High-resolution imaging using AFM reveals not only the shape and height of the brushes, but also resolves individual polymer sidechains. The gradient in grafting density (and thus brush height) is evident in several molecules.|
I also studied other conformational effects on single polymer brushes, include the effect of humidity and super-critical CO2 pressure.
Using video fluorescence microscopy, I observed the dynamics of individual λ-phage DNA molecules in entangled solutions in extensional flow. I explored multiple methods to concentrate DNA solutions without inducing phase separation or clumping, including using a speed vac and centrifugal filters. Also, I examined steady and dynamic rheology of the solutions and used the plateau modulus to extract the number of entanglements per chain.
|I used a simple microfluidic device to observe stretching in individual DNA strands. Because the flow in the t-shaped cell is extentional, the strands are pulled open from their coiled conformation. Because there is a stagnation point in the flowfield, it is possible to trap single DNA strands and observe them for long periods.||Individual fluorescently labeled DNA chains in entangled 1.0 mg/mL solutions at equilibrium. The labeled-to-unlabeled ratio is approximately 1:105.|
Single-molecule experiments can reveal rich behavior regarding chain dynamics--information that is often unattainable from bulk measurements, such as molecular individualism. Previous studies had focused on the dynamics of individual DNA molecules in dilute and unentangled semidilute solutions, which were useful because they were able to test theoretical predictions of polymer scaling and non-equilibrium dynamics in unentangled solutions; however, theory predicts quantitative changes in scaling and dynamics when the chains become entangled at high concentrations. Therefore, the goal of my study was to experimentally investigate the non-equilibrium dynamics of entangled polymer chains using single-molecule techniques and compare these results to theoretical models.
Dynamic rheology of bulk DNA solutions reveal the properties such as the reptation time and extent of entanglement.
Here, a concentrated solution exhibits not only a clear plateau, but also the cross-over of the two moduli which scales with τrep.
I used rheology as a powerful tool for measuring complex fluids. An oscillatory (dynamic) frequency sweep of an entangled polymer solution or melt should generate a rubbery plateau of the storage modulus (G') at frequencies greater than the inverse reptation time (τrep). On time scales shorter than the characteristic relaxation time of the polymer, topological entanglements act like physical cross-links and the solution acts as a rubbery gel. From the value of G' at the minimum of the loss modulus (G"), I estimated the value of the plateau modulus (Ge) from which the number of entanglements per chain can be calculated. For the most concentrated solution I made, there were approximately 16 entanglements per chain.
After confirming entangled solutions using bulk rheology, I fluorescently labeled DNA with YOYO dyes, then used microfluidics and microscopy in order to visualize the motions and dynamics of single strands among many unlabeled strands.