November 26, 2014

 

Patterns are soothing for left-brained folks like me, with the exception being those terrible patterned holiday sweaters that will come out of mothball-ridden closets soon (unsettling for everyone, really). Today’s images are from a paper describing a new micropatterning technique to look at plasma membrane proteins. 

The plasma membrane of a cell is riddled with many multi-protein complexes that facilitate communication and transport. These complexes provide a challenge to biologists due to their ubiquitous localization around the cell, their large, complex size, and their transient interactions with proteins. A recent paper describes a technique to study signaling complexes at the plasma membrane, using micropatterns within the plasma membrane. Löchte and colleagues expressed a protein bait in a micropattern in a living cell’s plasma membrane. Dynamics of the interactions between the protein bait and ligand target can then be quantified using live microscopy and on a single-molecule level. Löchte and colleagues used the IFN interferon signaling complex, using one of the receptor units (IFNAR2) as bait. In the top images above, the micropatterned receptor bait (IFNAR2) was able to recruit the other IFN receptor subunit (IFNAR1, green) after the addition of the ligand (red; time represents the addition of the ligand). Bottom image shows a closer view of the high-affinity, micropatterned binding that requires simultaneous interaction of the ligand (red) with both receptor subunits.


Lochte, S., Waichman, S., Beutel, O., You, C., & Piehler, J. (2014). Live cell micropatterning reveals the dynamics of signaling complexes at the plasma membrane originally published in the Journal of Cell Biology, 207 (3), 407-418 DOI: 10.1083/jcb.201406032

October 17, 2014

For years, the prettiest cells to image were flat cells in a dish. Thanks to the tireless work of many, beautiful high-resolution images can now come from tissue within a living organism. Today’s image is from a paper showing improved techniques for imaging fine cellular processes within large volumes, from the lab of recent Nobel prize winner, Eric Betzig. 

A material’s refractive index refers to how light travels through it; the simplest example being how light bends when passed through water. The refractive index heterogeneities stemming from the many cell types, morphologies, and subdomains within a living organism are a challenge to microscopists. As described in a paper from earlier this year, Wang and colleagues improved on previous techniques for imaging within large volumes. Wang and colleagues use adaptive optics (AO), which corrects for the microscope’s aberrations that limit image resolution, in a mode fast enough to correct for the various aberrations within a large sample, without inducing photodamage or photobleaching. The image above shows a 3D rendering from deep within a living zebrafish brain, with oligodendrocytes (magenta) and neuronal nuclei (green) visible.

Wang, K., Milkie, D., Saxena, A., Engerer, P., Misgeld, T., Bronner, M., Mumm, J., & Betzig, E. (2014). Rapid adaptive optical recovery of optimal resolution over large volumes Nature Methods, 11 (6), 625-628 DOI: 10.1038/nmeth.2925
Adapted by permission from Macmillan Publishers Ltd, copyright ©2014

September 17, 2014

All good things must end—even the focal adhesions that are so key to cell migration. Today’s notable image is the first live cell visualization of ECM degradation at focal adhesions, in a recent paper that reports the link between CLASPs, exocytosis, and focal adhesion turnover. 

Cell migration depends on the precisely-timed formation of focal adhesions (FAs) that link the crawling cell to the extracellular matrix (ECM). FAs serve as anchor points for the crawling cell, yet must later disassemble in order to allow continued movement of the cell. A recent paper describes how CLASP proteins link FA-associated microtubules, exocytosis, and FA turnover. CLASP proteins are +TIP proteins, which means that they are found on the growing ends of microtubules. Stehbens and colleagues found that the clustering of CLASPs around FAs correlates with the timing of FA disassembly, and that CLASPs are required for ECM degradation. Stehbens and colleagues also found that the tethering of microtubules to FAs, via CLASPs, serve as a transport pathway for exocytic vesicles at FAs. The images above are the first live cell images of ECM degradation (visualized as dark regions, top panel) at FAs (magenta).

BONUS! For more information on the scanning angle interference microscopy used in this paper, check out Matthew Paszek’s Nature Methods paper here.

Stehbens, S., Paszek, M., Pemble, H., Ettinger, A., Gierke, S., & Wittmann, T. (2014). CLASPs link focal-adhesion-associated microtubule capture to localized exocytosis and adhesion site turnover Nature Cell Biology, 16 (6), 561-573 DOI: 10.1038/ncb2975
Adapted by permission from Macmillan Publishers Ltd, copyright ©2014

September 11, 2014

As your therapist likely tells you, understanding where you came from is key to accepting where you are now. Take that therapist’s task and multiply it by several million—you now understand the tough job ahead of developmental biologists trying to track cell lineages in complex organisms. Today’s colorful image is from a paper describing a new computational framework for reconstructing cell lineages. 

The successful tracking of cell position, division, and movement in a developing organism has been a goal for countless developmental biologists. Reconstructing cell lineages in organisms like fruit flies and mice, however, is difficult due to the complexity of cell organization and behavior, poor image quality of thick embryos, the enormous size of the data sets, and an uncompromising need for accuracy. A recent paper by Amat and colleagues describes the development and use of a new open-source framework that reconstructs cell lineages with high accuracy and speed. Their system uses four dimensional and terabyte-sized image data sets of nuclei-tracked embryos, imaged using three different types of fluorescence microscopy. The images above show the first reconstruction of early fruit fly nervous system development (S1 neuroblasts), with precursor cell tracks color-coded for time (purple to yellow).

Amat, F., Lemon, W., Mossing, D., McDole, K., Wan, Y., Branson, K., Myers, E., & Keller, P. (2014). Fast, accurate reconstruction of cell lineages from large-scale fluorescence microscopy data Nature Methods, 11 (9), 951-958 DOI: 10.1038/nmeth.3036
Adapted by permission from Macmillan Publishers Ltd, copyright ©2014

August 26, 2014

If you have little ones in your house, you might assume that the phrase “randomly fluctuating forces” is referring to your home. This phrase actually refers to the background force in a cell coming from active and motor-driven cell processes. Today’s image is from a study that developed a way to measure these forces. 

Actin- and microtubule-based motors move many types of material around a cell to drive critical cellular events. These motor-driven movements and other active processes in the cell contribute to a background of fluctuating forces in a cell. These stochastic forces collectively drive random motion of organelles and proteins within a cell, in turn affecting the dynamics and metabolic state of a cell. To measure these forces, Guo and colleagues developed force spectrum microscopy (FSM) to directly quantify the fluctuating forces in a cell’s cytoplasm, specifically by measuring the movements of individual injected particles. Guo and colleagues found that these forces are strong enough to move both large and small components, and that malignant cells have a higher level of fluctuating forces compared to benign cells. Image above shows a cell with injected particles (green), with the 2-minute trajectories (black) superimposed.

Guo, M., Ehrlicher, A., Jensen, M., Renz, M., Moore, J., Goldman, R., Lippincott-Schwartz, J., Mackintosh, F., & Weitz, D. (2014). Probing the Stochastic, Motor-Driven Properties of the Cytoplasm Using Force Spectrum Microscopy Cell, 158 (4), 822-832 DOI: 10.1016/j.cell.2014.06.051
Copyright ©2014 Elsevier Ltd. All rights reserved.

August 7, 2014

No matter how many brilliant discoveries are made by countless brilliant scientists, there will always be a lot of unknowns out there. These unknowns are what keep our mental wheels turning, our imaginations running, and our labs busy. Today’s image is from a paper that describes a newly-discovered process of vascular development called “canalogenesis.”

Schlemm’s canal (SC) is a flattened tube that encircles the anterior portion of the eye and drains fluid from the area. Abnormal drainage contributes to glaucoma, a disease that causes vision loss, yet a description of SC development and SC endothelial cells (SECs) is incomplete. In a recent study, Kizhatil and colleagues developed a new whole-mount procedure and used high-resolution confocal microscopy to study large regions of the SC during development. Kizhatil and colleagues found that the phenotype of SECs is a blend of blood and lymphatic endothelial cells, and that the SC develops through by a newly-discovered process called “canalogenesis.” Canalogenesis has features that are similar to, yet different from, the three well-studied vascular development programs—vasculogenesis, angiogenesis, and lymphangiogenesis. The image above was acquired using the new whole-mount procedure that protects the delicate ocular drainage structures. The SC (blue) is visualized in 3D relative to nearby blood vessels (magenta).

Kizhatil, K., Ryan, M., Marchant, J., Henrich, S., & John, S. (2014). Schlemm's Canal Is a Unique Vessel with a Combination of Blood Vascular and Lymphatic Phenotypes that Forms by a Novel Developmental Process PLoS Biology, 12 (7) DOI: 10.1371/journal.pbio.1001912

July 10, 2014

Do your thoughts and feelings have colors? Do you feel red with rage during traffic, or green with envy when your lady swoons over Ryan Gosling? A recent methods paper introduces a very cool technique that allows the visualization and measurement of voltage within an excited neuron.

Biologists build tools that are ideally accurate, fast, and non-damaging to the cells and organisms on which they are used. In a recent paper in Nature Methods, Hochbaum and colleagues describe the improved technique for simultaneous imaging of neuron stimulation and the resulting action potentials. Hochbaum and colleagues engineered a vector, called Optopatch, that uses their actuator (CheRiff) to induce action potentials and their voltage indicators (QuasAr1 and QuasAr2) to visualize and measure membrane voltage. Optopatch allows the measurement of action potentials on a microsecond timescale, without the need for electrodes. In the images above, a neuron expressing Optopatch shows action potential propagation (left to right, arrow is site of action potential initiation).

Hochbaum, D., Zhao, Y., Farhi, S., Klapoetke, N., Werley, C., Kapoor, V., Zou, P., Kralj, J., Maclaurin, D., Smedemark-Margulies, N., Saulnier, J., Boulting, G., Straub, C., Cho, Y., Melkonian, M., Wong, G., Harrison, D., Murthy, V., Sabatini, B., Boyden, E., Campbell, R., & Cohen, A. (2014). All-optical electrophysiology in mammalian neurons using engineered microbial rhodopsins Nature Methods DOI: 10.1038/nmeth.3000
Adapted by permission from Macmillan Publishers Ltd, copyright ©2014

February 19, 2014

It will take cell biologists an eternity to understand how cells function in a dish. It will take developmental biologists even longer to understand how cells function within a developing organism. Today’s image is from a paper describing the use of liquid droplets as cell biological crash test dummies to determine cell-generated forces within living tissue.

The development of an organism and the generation of its organs depend on mechanical forces that can move cells and groups of cells. There are many techniques that have been helpful in understanding these forces, but these techniques have not been applicable for measuring forces in three dimensional living tissue (or, in the case of laser ablation, have only provided relative force measurements). Campàs and colleagues have just published a paper describing the clever use of oil microdroplets to measure force within living tissues and organs. These spherical microdroplets are a similar size as cells, are fluorescently labeled, and display cell surface adhesion receptors. After these microdroplets are injected into tissue, their deformation by surrounding cells exerting mechanical stress can be measured and quantified. The images above show a microdroplet (white arrow, top) embedded in an incisor tooth bud of a mouse embryo (E13.5). Higher magnification views of the microdroplet (bottom row) show the pixel-resolution contour of the droplet (middle). The higher curvature regions (arrows, right image) correlate with cell-cell junctions of adjacent cells. 

Campàs O, Mammoto T, Hasso S, Sperling RA, O'Connell D, Bischof AG, Maas R, Weitz DA, Mahadevan L, & Ingber DE (2014). Quantifying cell-generated mechanical forces within living embryonic tissues. Nature methods, 11 (2), 183-9 PMID: 24317254
Adapted by permission from Macmillan Publishers Ltd, copyright ©2014

May 24, 2013

There isn’t a cell biologist out there who doesn’t fantasize about reaching her hands into a cell and physically manipulating whatever protein or structure that she obsesses over.  While we can’t do that with our own hands, optical tweezers can…and the information we learn is invaluable.  Today’s image is from a paper that uses optical tweezers to measure the forces within a mitotic spindle.

The mechanics within a mitotic spindle are complicated, and cannot be fully understood until all of the forces that exist within it are determined.  A recent paper measures the forces of chromosome and spindle pole movements using optical tweezers to trap either kinetochores or spindle poles.  Optical tweezers use refracted light to trap small objects within a cell, and are able to stop the movement of the object as the laser power of the trap approaches (or exceeds) the cell’s own force on the object.  With this technology, Ferraro-Gideon and colleagues measured the forces in mitotic spindles in several samples—a flatworm, a crane fly, and a mammalian spindle.  The force used to stop chromosome movement was closer to the theoretical values of chromosome force, much less than the force values calculated by experiments performed by Bruce Nicklas in the 1980s using needles to manipulate chromosome movement.  In the DIC (left) and fluorescence images (right) above, a trap (square) is applied to kinetochores in mammalian cells.  The trap did not damage the microtubules of the spindle (right image).

Ferraro-Gideon, J., Sheykhani, R., Zhu, Q., Duquette, M., Berns, M., & Forer, A. (2013). Measurements of forces produced by the mitotic spindle using optical tweezers Molecular Biology of the Cell, 24 (9), 1375-1386 DOI: 10.1091/mbc.E12-12-0901

Check out an extended report on these experiments, and others from this lab:
Sheykhani, R., Baker, N., Gomez-Godinez, V., Liaw, L., Shah, J., Berns, M., & Forer, A. (2013). The role of actin and myosin in PtK2 spindle length changes induced by laser microbeam irradiations across the spindle Cytoskeleton, 70 (5), 241-259 DOI: 10.1002/cm.21104

May 21, 2013

“LET THERE BE LIGHT!” said the microscopist.  Light plays a crucial role in microscopy and cell biology, and a recent paper describes the use of light to understand protein secretion.

Light is used in microscopy in countless ways—to illuminate a sample, excite a fluorophore, and signal the localization or dynamics of a protein.  Light can also be used to manipulate cellular events through the use of “caged” compounds that become active after illumination by certain wavelengths of light.  This technology gives biologists the ability to spatially and temporally control cellular events in order to understand them better.  Recent advances in this technology use illumination of plant photoreceptors to control protein-protein interactions, but some cellular processes such as protein secretion have been difficult to manipulate.  A recent paper describes the use of the plant photoreceptor UVR8 in the first light-triggered protein secretion system developed.  Chen and colleagues have shown that the recently described UVR8 can conditionally sequester proteins bound for secretion in the ER, and then upon illumination with UV light releases these proteins to the plasma membrane.  In the images above, a neuron before (left) and after (right) UV illumination with this UVR8 system shows the movement of proteins known to be secreted from the soma and dendritic processes (arrowheads), where the ER is distributed, and into the Golgi (arrow), a necessary step in protein secretion.

Chen, D., Gibson, E., & Kennedy, M. (2013). A light-triggered protein secretion system originally published in the Journal of Cell Biology, 201 (4), 631-640 DOI: 10.1083/jcb.201210119

November 9, 2012

Size really does matter, folks! Ask any scientist about the potential uses for nanoparticles, and you’ll quickly agree that these tiny little suckers deserve a spot at your dinner table, complete with cute nano-plates and nano-forks. Today’s image is from a paper describing a clever use of nanoparticles to systematically monitor how cells respond to force.

A cell encounters force from its entire environment. A cell responds to mechanical force by regulating signals, development, and migration, among other things. Biologists have been trying to understand how cells respond to force for years, and clever techniques have included manipulation by optical tweezers, micropipettes, and atomic force microscopy. These techniques, however, have been limited by their ability to monitor one cell at a time. A recent paper describes a technique in which many cells respond to uniform forces on the cortex. In this study, Tseng and colleagues plated cells on micropatterned magnetic substrates. Magnetic nanoparticles within the cells were then manipulated by a magnetic field in order to apply a uniform mechanical force on the cell cortex. By monitoring large numbers of individual cells under these forces, Tseng and colleagues developed a way for biologists to have higher control and accuracy in understanding how force affects cells. The images above show representative images of cells under varying amounts of force. The dose of nanoparticles (blue) in each cell (increasing from bottom to top) and strength of the magnetic field (increasing from left to right) both affect the response from cells. A higher nanoparticle dose and magnetic field gradient caused cells to produce more filopodial membrane extensions from cells (actin is in green) from the region where the force was applied.

Tseng, P., Judy, J., & Di Carlo, D. (2012). Magnetic nanoparticle–mediated massively parallel mechanical modulation of single-cell behavior Nature Methods, 9 (11), 1113-1119 DOI: 10.1038/nmeth.2210
Adapted by permission from Macmillan Publishers Ltd, copyright ©2012

August 10, 2012

Biologists are always looking for ways to push the limits of what our microscopes are capable of doing. These Microscopy Olympians (which should be a real thing, seriously) understand the physics and biology behind imaging cells in order to build, tweak, and test fancy-shmancy new technologies. And, just like the real Olympics, I watch from my couch with a bag of pretzels in front of me.

To accurately capture high resolution images of cells, biologists are always improving ways to capture three dimensional structures over time in a living cell. Combining nanoscale resolution with live imaging has proven a challenge, recently helped by a paper that describes the tweaks made to earlier microscopy setups. Paszek and colleagues presented their technique called scanning angle interference microscopy, in which dynamic cellular events and structures are imaged with nanoscale resolution in three dimensions. In the image above, microtubules imaged using this technique show their location in three dimensions. The “height” along each microtubule is color-coded (in nm) to show that microtubules bend down towards the cell cortex.

Paszek MJ, Dufort CC, Rubashkin MG, Davidson MW, Thorn KS, Liphardt JT, & Weaver VM (2012). Scanning angle interference microscopy reveals cell dynamics at the nanoscale. Nature methods, 9 (8), 825-7 PMID: 22751201

August 2, 2012

 Sometimes you watch a movie, and then you watch it again. And again. And again. Maybe it was Goonies, back when you were a kid (or adult)…or it was a pair of otters holding hands or a baby panda sneezing. Or maaaaybe it was watching stem cells and their progeny divide in real time, within the actual organism. Thanks to the technical gymnastics of a group of stem cell researchers, we can now see how stem cells and their progeny within a hair follicle divide and regenerate hair.

Tissue regeneration begins with stem cells and their immediate progeny. Knowledge of this process has been incomplete due to the difficulty of tracking all of the cell behaviors in living tissue. A group recently used high resolution imaging to track hair regeneration within hair follicles of mice over a long period of time. By tracking cell behaviors, Rompolas and colleagues found that the immediate progeny of stem cells actively divide to regenerate the tissue, while stem cells divide more slowly to contribute to hair regeneration over time. The stem cell progeny cell divisions are mostly aligned with the long axis of the hair. Finally, Rompolas and colleagues used cell ablation to show that a mesenchymal group of cells called the dermal papilla is required for hair regeneration. In the images above, stem cell progeny divisions can be seen in a live hair follicle undergoing growth. Three different cells are colored to show their divisions (right columns).

BONUS! Check out a movie of stem cell progeny divisions here and here, and a movie of stem cell divisions here. For all movies from this paper, click here.

Panteleimon Rompolas, Elizabeth R. Deschene, Giovanni Zito, David G. Gonzalez, Ichiko Saotome, Ann M. Haberman, & Valentina Greco (2012). Live imaging of stem cell and progeny behaviour in physiological hair-follicle regeneration Nature, 487 (7408), 496-499 DOI: 10.1038/nature11218
Adapted by permission from Macmillan Publishers Ltd, copyright ©2012

 

May 31, 2012

I love it when things really throw me for a loop. Like when I heard that Kim Kardashian earned a perfect SAT score and was accepted into Harvard. Okay, that’s not really true…but just imagine our collective “whaaaa?!” In science there are always things that flip your lid, and today’s image is from a paper that does just that. Clathrin-coated pit closure just got more interesting.**

Clathrin-mediated endocytosis is a process during which material is brought into the cell through pits coated with a clathrin lattice, which provides structure to the developing vesicle. Electron microscopy images of clathrin-coated pits show great detail about their structure, but can only provide snapshots of what occurs inside of the cell. A recent paper describes clathrin-coated pit closure using a technique that combines the imaging of structural detail seen in electron microscopy (in this case, called scanning ion conductance microscopy) with live confocal microscopy used to track fluorescently-tagged proteins. During conventional clathrin pit closure, the pit is closed and cleaved from flat membrane sheets. Shevchuk and colleagues, however, found that 70% of pits close using an alternative mechanism. In this mechanism, a membrane protrusion grew from one side of the clathrin pit and covered the pit to close it, as seen in the images above.

**Halvsies if this tagline prompts next summer’s blockbuster movie. 

Shevchuk, A., Novak, P., Taylor, M., Diakonov, I., Ziyadeh-Isleem, A., Bitoun, M., Guicheney, P., Lab, M., Gorelik, J., Merrifield, C., Klenerman, D., & Korchev, Y. (2012). An alternative mechanism of clathrin-coated pit closure revealed by ion conductance microscopy originally published in the Journal of Cell Biology, 197 (4), 499-508 DOI: 10.1083/jcb.201109130

May 3, 2012

Where did you come from? Understanding the answer to this question helps you get a grasp on who you are currently, and who you are becoming. Whoa, that was deep…enough of that crap! A recent paper puts the psychotherapy treatment on cells in the developing heart and adds to our understanding of where they are from.

The change that takes place in a developing heart is astounding—a simple tube structure with a single layer of cardiac muscle cells called cardiomyocytes must develop into a complex adult structure. How the early cardiomyocytes divide and move around to form the adult heart is difficult to map out due to their dynamic behavior and their location in a tissue that is difficult to image. A recent paper describes the use of a recently-developed technology called Brainbow, in which different cells can be labeled with about 90 different colors. Using Brainbow techniques, Gupta and Poss were able to track the division and movement of individual cardiomyocytes in the developing zebrafish heart, and found that the cells expand the cardiac tissue in patches of various sizes and shapes. By using clonal dominance as a mechanism for tissue expansion, the process is reminiscent of stem cells. The image above shows the surface of the ventricular side of a developing heart, each color representing a different clonal patch that arose from a single cardiomyocyte.

Gupta, V.; Poss, K. (2012). Clonally dominant cardiomyocytes direct heart morphogenesis Nature, 484 (7395), 479-484 DOI: 10.1038/nature11045

Adapted by permission from Macmillan Publishers Ltd, copyright ©2012

 

February 23, 2012

My bucket list has a few unrealistic things on it, such as “establish a close friendship with an elephant.” When I see how fast imaging technology improves, my goal to “float in a cell with swimmies on my arms” may not be as far off base as finding my BFF pachyderm. Although I may never be able to dive into a cell, I’m willing to bet I can eventually find myself standing in some amazing 3D images of a cell. Today’s image is from a paper describing improved imaging methods that produce super-resolution images that’ll knock your socks off.

Image resolution refers to the ability to distinguish two closely positioned lines or objects. Super-resolution imaging techniques have allowed biologists to see details in small structures, with resolutions down to about 20nm. A group of biologists recently pushed this resolution limit even further, and by doing so, was able to capture images of individual actin filaments with amazing detail. Xu and colleagues improved the 3D STORM (stochastic optical reconstruction microscopy) imaging methods by using two objective lenses to double the amount of fluorescent signal collected off of a cell sample. By imaging individual actin filaments in cells using this dual-objective STORM method, the 3D ultrastructure of the cytoskeleton revealed two layers of actin networks in a sheet-like membrane protrusion. The images above show the distinct organization of each of these two layers, each with a thickness of about 30-40nm. Color coding of the filaments (violet to red) provides a representation of the three-dimensional space (violet actin is closest to the glass coverslip, while red is furthest from the coverslip).

Xu, K., Babcock, H., & Zhuang, X. (2012). Dual-objective STORM reveals three-dimensional filament organization in the actin cytoskeleton Nature Methods, 9 (2), 185-188 DOI: 10.1038/nmeth.1841

January 23, 2012

The mammalian brain has billions of synaptic connections, which is more than a little difficult to sort and map out. Today’s image is from a group of biologists who used their own brilliant synaptic connections and developed a technique for mapping out these connections in mammals.

One of the most commonly used fluorescent markers in cell biology is GFP, which stands for Green Fluorescent Protein. Scientists have been very creative in using this powerful tool to understand more about cell biology. For example, GRASP (GFP reconstitution across synaptic partners) is a technique developed a few years back in order to map out synaptic connections in worms and flies. In GRASP, one half of the GFP protein is expressed in one type of neuron, while the other half of GFP is expressed in a different type of neuron. Alone, each of these GFP fragments cannot fluoresce, but when the two neurons are close together in a synaptic connection, the GFP will fluoresce green light that pinpoints the location of a connection. Recently, a group of biologists made major modifications in this technique in order for it to work in mammals, whose synaptic architecture can vary a lot from flies and worms. In the images above, dendrites (red) in a mouse brain intersect with axons (blue), as seen as the green GRASP signal.

Kim, J., Zhao, T., Petralia, R., Yu, Y., Peng, H., Myers, E., & Magee, J. (2011). mGRASP enables mapping mammalian synaptic connectivity with light microscopy Nature Methods, 9 (1), 96-102 DOI: 10.1038/nmeth.1784
Adapted by permission from Macmillan Publishers Ltd, copyright ©2011

December 5, 2011

When I read papers like the one that gave us today’s image, I think that one day my wish of jumping into a cell to float in the cytoplasm wearing goggles and swimmies may actually come true. Physical manipulation of proteins gets me so excited about how far our tools and technology have come. In this paper, biologists physically yanked on actin filaments to show how tension affects the presence and function of an actin-modulating protein.

Actin has many regulatory proteins that do a variety of things, such as promoting filament nucleation, branching, and severing. Cofilin is a ubiquitous protein that functions in actin filament severing and reorganization. Cofilin binds to the actin filament itself and induces a slight twist in the actin, which makes it easily severed. A recent paper describes the use of optical tweezers and manipulations to show that the binding of cofilin to actin, and in turn its severing of actin, is regulated by tension in the filaments. Hayakawa and colleagues bound one end of an actin filament to a glass coverslip and manipulated the other end using optical tweezers. When the filament was put under tension, the actin filament was not severed (or it took longer, in some cases). In another manipulation, a fine glass pipette was used to pull bundles of actin. Tension applied to the actin filaments caused a drop in the binding rate of cofilin to actin filaments, as seen in the images above. Top row shows actin (left) and cofilin (right, fat arrows) in a tension-relieved actin bundle, while bottom row shows actin and reduced cofilin binding in an actin bundle that was stretched by 20%.

Hayakawa, K., Tatsumi, H., & Sokabe, M. (2011). Actin filaments function as a tension sensor by tension-dependent binding of cofilin to the filament originally published in The Journal of Cell Biology, 195 (5), 721-727 DOI: 10.1083/jcb.201102039

October 20, 2011

One man’s trash is another man’s treasure. Photobleaching is an unavoidable side effect of imaging that leads to weakened fluorescent signals. Most of us pooh-pooh photobleaching, but some clever cell biologists use photobleaching as a fantastic tool instead. Today’s image is my new favorite use of photobleaching.

Glial cells function in the nervous system to support neurons and their signal transmission. Schwann cells are glial cells found at the neuromuscular junction (where neurons signal to muscles), and monitor the neurotransmission exchanged. Schwann cells can even function to form and regenerate the neuromuscular junction. A recent paper describes how Schwann cells establish their arrangement around the neuromuscular junction. In this paper, Brill and colleagues labeled individual Schwann cells and used live imaging to monitor their positioning. Schwann cells are dynamic during development, finding their appropriate position by competing for space with other Schwann cells. Schwann cells in adult animals, however, are much more static. In the images above, individual Schwann cells (pseudo-colored yellow, blue, white, and purple) were labeled by sequentially photobleaching one cell at a time, leading to distinct levels of fluorescence in each cell. The axon of the neuromuscular junction is in green. The Schwann cells are very dynamic in young mice (left) compared with adult mice (right), seen as the frequent formation and retraction of cell protrusions (arrowheads in the images of the boxed regions).

Brill, M., Lichtman, J., Thompson, W., Zuo, Y., & Misgeld, T. (2011). Spatial constraints dictate glial territories at murine neuromuscular junctions originally published in The Journal of Cell Biology, 195 (2), 293-305 DOI: 10.1083/jcb.201108005

July 14, 2011

Do you spend your photon budget wisely? This is a question that a recent paper asks, and answers back with a new technique that produces images that will blow your socks off. Once you put your socks back on, check out today’s image.

Cellular imaging is a constantly evolving field made of biologists on a never-ending quest for higher resolution of structures and faster image acquisition of a living cell. There are several challenges to these demands. For example, cells are not pancake-thin. Current techniques use illumination that leads to background noise in an image due to excited out-of-focus light. In addition, these techniques can cause phototoxic effects on cells, and can photobleach the fluorescent tags used to mark structures. Biologists have addressed these problems by using plane-illumination microscopy, which uses a separate excitation lens positioned orthogonally to the detection objective lens, leading to a more confined excitation of the focal plane. Planchon and colleagues recently improved this technique by using thinner sheets of light to illuminate the sample. The images produced using this Bessel beam plane illumination are remarkable, and allow for very fast 3D imaging of living cells. Images above show filopodia on a HeLa cell (left column), and the membrane ruffles on a kidney cell (right group of images). Purple arrowheads point to vacuole formation by macropinocytosis.

BONUS!! For a movie of the filopodia in the image above, click here. For a movie of the membrane ruffles and vacuole formation, click here. For many other knock-your-socks-off movies, click here.

Planchon, T., Gao, L., Milkie, D., Davidson, M., Galbraith, J., Galbraith, C., & Betzig, E. (2011). Rapid three-dimensional isotropic imaging of living cells using Bessel beam plane illumination Nature Methods, 8 (5), 417-423 DOI: 10.1038/nmeth.1586
Adapted by permission from Macmillan Publishers Ltd, copyright 2011