The development of qFSM is funded by the NIH grant U01 GM06230.

Section 1: Obtaining your fluorescently labeled actin for FSM
Materials required: purified actin, fluorophore (Alexa, X-rhodamine recommended). G-buffer, ultracentrifuge
<ol>
<li>A key component to acquiring good FSM movies requires the proper labeling of actin (or other cytoskeletal proteins) with the fluorophore of your choice. We recommend using fluorophores of Alexa (488, 568 wavelengths) and X-rhodamine succinimidyl ester derivatives that target exposed lysine residues on the surface of actin. </li>
<li>When choosing your fluorophore and its appropriate wavelength, ensure that your microscope setup has the appropriate filters (excitation and emission) and illuminators (mercury arc lamp and/or lasers) to accommodate your fluorescent tag. We recommend selecting wavelengths in the orange to red wavelengths (560-630 nm) to minimize the effects of auto-fluorescence and photo-damage and to ensure its compatibility with GFP-conjugates. </li>
<li>Pure actin can be extracted from muscle tissue belonging to Rat or Chicken, but requires a lengthy extraction procedure from muscle acetone powder. Muscle acetone powder can be purchased from Invitrogen. The labeling procedure is not discussed in this protocol, but can be found here[1] </li>
<li>Alternatively, you can purchase labeled actin proteins from several vendors including Cytoskeleton Inc., and Invitrogen. Custom labeling services are also available through Invitrogen. Purified, non-labeled actin (both muscle and nonmuscle actin) can be purchased from Cytoskeleton Inc.</li>
<li>Whether you decide to purchase or prepare your own labeled actin, you want to ensure that the labeling ratio is somewhere around 0.3 to 0.7 dyes per monomer of actin. Too high a labeling ratio may effect the turnover kinetics (and binding to associated proteins) of the actin filament, while too low a labeling ratio will lead to poor looking speckles (low signal to noise) and may cause microinjection problems (see below). This is a critical factor for getting good speckle images.</li>
<li>Prior to long term storage, the labeled actin solution should be clarified by spinning at 75,000 - 80,000xg for 20 minutes at 4 degrees Celcius. Labeled actin should be stored in G-buffer solution and kept -80 degrees Celcius. Preparing 2 uL aliquots is highly recommended to reduce the detrimental effects of repeated freezing-thawing (leads to insoluble aggregates). Refrain from exposing the labeled actin solution to direct light (will photobleaching of fluorophores). We recommend using opaque microcentrifuge tubes for long term storage.</li>
</ol>
Section 2: Preparation of Cells and Microinjection of labeled actin
Materials required: microinjection needles, microsyringe, microloaders, prewarmed cell media, injection buffer, bucket of ice, microscope (phase ring, phase contrast objective 40X, transjector, needle puller, micromanipulator)
<ol>
<li>Many cell types are amenable to FSM, but getting good speckle images is directly dependent on the success of the microinjection procedure. When selecting cells for FSM, cells should be large ( &gt; 50 um in diameter), should spread when plated, and be naturally adherent to substrates (or plastic tissue culture dishes). It also helps if the cell&rsquo;s cytosol occupies an area greater than its nucleus. The aforementioned criteria ensure that cells are amenable to microinjection. Successful candidate cell lines include (but not limited to) PtK1[2,3], newt lung cells[4], and Aplysia neurons[3]. </li>
<li>Cells should be plated on square glass coverslips (22 x 22 mm; no. 1-1/2) that have been acid-washed. Standard substrate coating can be applied to the coverslip and will not affect microinjection.</li>
<li>The correct selection of microinjection needles is dependent on the size of the tip opening. Typically, hole sizes can very in size from 0.5 um to as large as 3 um. However, we recommend that injection of actin requires an optimal opening size between 0.5 to 1 um. If clogging problems are encountered consistently, larger hole sizes can accommodate for the delivery of monomeric as well as unwanted polymerized actin filaments. Microinjection needles can be purchased through several vendors including Eppendorf systems which make femtotip microinjection needles. Alternatively if a commercial needle puller is available, glass microinjection tubes can purchased from Sutter, and pulled to specification. For beginners, we recommend having 10-20 pulled needles on hand before starting the injection procedure.</li>
<li>Ideal &lsquo;injection concentration&rsquo; (final concentration of actin loaded in your needle) of actin is between 0.5 to 1.0 ug/ul based on a 40% labeling ratio of dye to actin monomer. Lower labeling ratio of your actin can be compensated by injecting at higher concentrations (1-3 ug/ul). However, the clogging frequency of your needle will be increased. This can be partially alleviated by using needles with larger openings. Stock concentration of actin can be diluted by adding ice-cold injection buffer. Ice-cold ddH2O can also be used to dilute for quick injections. At this point all working solutions should be kept on ice. </li>
<li>Load 0.5 to 1.0 uL of your actin solution using a microsyringe (narrow-gauge &ndash; Hamilton), or alternatively by using a microloader pipette tip. </li>
<li>Carefully dispense the actin solution, by gently releasing the solution. Bubbles should be avoided, but can be easily removed by taking up excess liquid surrounding the bubble. </li>
<li>Ensure that the actin solution is resting completely at the needle tip. Excess solution anywhere along the capillary of the needle can disrupt the injection flow. </li>
<li>Now, carefully attach the needle, avoiding contact with the needle tip, to the injector holder. Position the holder so that the needle makes a 35-50 degree angle to the base of the microscope stage. Lower angles are preferred.</li>
<li>Lower the tip of the needle until it barely breaks the surface of the media bathing your cells. </li>
<li>Now position the needle tip (without lowering) so that it rests directly above the center of the objective. If the needle tip is aligned with the objective, you should see a bright halo through the eyepiece. This is the end of the needle tip. Slowly moving the needle side to side (using the manipulator) may help better locate the halo. </li>
<li>Adjust the z-focus on the microscope until your cells are in clear view. Now, slowly lower the tip until the halo gradually constricts until finally converging into the needle tip &ndash; you will need to control the focus (z-position) simultaneously. An additional feature to look for is the shaft of the needle, which will cast linear shadows. Lowering the needle will gradually define the needle shaft. When done correctly the end of the needle and your cells should both be in clear view. Slight adjustment of the phase ring may be required to improve contrast. </li>
<li>When navigating, ensure the needle tip is well above your cells to prevent breaking the needle tip. </li>
<li>When selecting cells to inject, cells with free edges should be facing the needle tip. This will ensure declining slope of the cell (from thickest point, where the nucleus resides, to the thinnest point the leading edge) will meet the needle tip, rather than running parallel to it. </li>
<li>The needle pressure should be set at 0.3 to 0.8 psi. The constant pressure applied will result in a steady flow of the actin solution. </li>
<li>After deciding on a cell to inject, position your needle so the tip is next to the nucleus (perinuclear region, thickest part of the cell where the nucleus is not directly below). </li>
<li>Slowly lower, and readjust the position of the needle until the tip is depressing the membrane, now incrementally lower until the needle tip has pierced the membrane. If the needle has fully pierced the membrane, the cell will appear brighter, as soon as you see this difference raise the needle until it is no longer touching the cell. The actual piercing and injecting process will take 0.5 to 2 seconds. </li>
<li>When done correctly, the cell shape will appear unchanged, whereas too much injection will result in quick retraction of the cell edge and in some cases you will actually see the cell explode. </li>
<li>If you do not see changes in the cell when microinjecting (does not get brighter), the needle tip may be closed. You can test this by pressing the &lsquo;clean&rsquo; button if available on your transjector. If open, this will significantly increase the constant pressure flow which can be seen through the eyepieces &ndash; it will appear as ripples and nearby debris can be seen dispersing. </li>
<li>If the needle tip is closed, you can choose to insert a new microinjection needle or attempt to break the tip of the needle by gently lowering the needle until the tip of the needle presses against the glass coverslip, thus causing the very end of the tip to break. The breakage area should be no more than a 1/5th of the area of the needle point (the portion of the needle that starts to converge to a point). </li>
<li>Continue with the injection, spending no more than 20 &ndash; 45 minutes for each microinjection session (will depend on cell type). As a general rule, shorter sessions are recommended to minimize stress on cells.</li>
<li>After finishing the microinjection, change the cell media in the dish by adding fresh prewarmed media. Then allow cells to recover for 30 minutes in an incubator before imaging. This will also allow for the actin to integrate into the existing cytoskeleton network. </li>
</ol>
Section 3: Assembling the Imaging Chamber
Materials Required: coverglass, double sided tape, q-tips, forceps, valap, kim wipe, oxyrase, imaging media, razor blade, p200 pipette, pure ethanol
<ol>
<li>Begin by prewarming the imaging media and thawing a 15 uL aliquot of Oxyrase. Make sure to protect the photosensitive oxyrase from light. The addition of oxyrase will greatly reduce the effects of photobleaching. </li>
<li>Prewarm the valap, by setting the hot plate temperature to low. Do not overheat/burn the valap for it will lose its efficacy as a sealant. </li>
<li>Wipe down a coverglass with a kimwipe doused with pure ethanol to clean and remove small debris. </li>
<li>Stack two equal sized lengths (2.5 cm long) of double-sided tape on top of each other, using a clean flat surface as a platform. Press out any trapped air bubbles to create a completely flat plane.</li>
<li>Using a razor blade, cut two straight strips along the long axis, so that each strip measures 0.5 cm in width. </li>
<li>Using forceps, place each strip against the long edge of your coverglass so that the two strips of tape creates a gap in the center of your coverglass. Press gently on the tape to make a nice seal with the glass. The strips of tape will act as spacers between the coverglass and the coverslip.</li>
<li>Make sure the valap has completely liquefied before you begin the next step. </li>
<li>Retrieve your prewarmed imaging media, adding 15 ul of oxyrase to every 500 uL of media.</li>
<li>Fold kimwipes into triangles to create 3 hard edges. This will be used to create a dry surface for the tape to stick on. </li>
<li>Retrieve your cells from the incubator. Using forceps, grab a corner of the coverslip. Quickly place the coverslip on a kimwipe (cell side is facing up, not touching the tissue), and use the folded kimwipe to wipe two opposing sides of the coverslip to create a semi-dry surface (on the surface where your cells rest). </li>
<li>Using forceps, grab a corner the coverslip and place it on your coverglass so that the dry surfaces are aligned to the two strips of tape. Notice now you have two closed sides and two open slits. Slowly pipette 200 uL of your imaging-media-oxyrase solution close to one opening. Importantly, no air bubbles should be introduced. The solution should be sucked up into the newly assembled chamber by capillary forces, thus fully immersing your cells in solution. </li>
<li>Using a Q-tip dab the four corners of your coverslip with melted valap to quickly affix and stabilize the coverslip. You&rsquo;ll notice that the valap dries instantly (within seconds at room temperature). </li>
<li>Now beginning with the open sides (non-tape side), lay a thin strip of valap by using a Q-tip and mimicking a brushing stroke. Repeat for the closed sides. Slowly build up the coat of valap, thus reinforcing the seal by repeating the coating steps. The valap should be confined to the edges, leaving the center of the coverslip clear. </li>
<li>Gently clean the center portion of the coverslip by using a Q-tip with ddH2O to remove any residual media, being mindful not to crush the cells. Repeat cleaning step with pure ethanol. </li>
<li>You will now have a fully enclosed imaging chamber optimized for speckle imaging. Having a fully sealed enclosure will prevent your fluorophores from quickly photobleaching.</li>
</ol>
Section 4: Imaging cells
Materials required: inverted widefield microscope, mecury lamp, appropriate filters, cooled CCD camera with 6.7 micron pixels, High numerical aperture, oil-immersion plan-Apochromatic objectives (60X to 100X, phase or DIC). Vibration table. Imaging software.
<ol>
<li>Prewarm the microscope stage to 37 degrees. Place your newly completed imaging chamber, and wait 10-15 minutes for the temperature to stabilize before imaging. </li>
<li>Start by focusing your cells in Bright-field, and then find your injected cells by toggling the correct fluorescence settings. Try to minimize the time of fluorescence exposure to reduce photobleaching.</li>
<li>Look for injected cells that have healthy curved leading edges. Over-injected cells (exploded cells) will be characterized by retracted and jagged edges, resembling numerous filopodia &ndash; avoid imaging these cells. </li>
<li>Acquisition settings will depend on the cell type, the quality of the labeled actin, and microscope components. However, exposure settings should not be greater than 2 seconds, and time interval between frames should be set between 5 and 10 seconds. Typical lengths for a time series (per imaged cell) can vary anywhere between 10 to 30 minutes, and is limited by photobleaching effects. Gain settings on the camera can be turned up to increase the signal to noise.</li>
<li>Ideal speckle characteristics should have each speckle spaced by ~2 speckle diameters to the next neighboring speckle. This will warrant optimal spatial and temporal sampling of the actin structure. For epithelial cells, the speckled actin network will appear homogeneous, with speckles evenly spread apart. Cell lines that feature dense stress fibers or lamellipodia will be distinguishable, but will have a dotted/speckled appearance. Exposure settings should be adjusted to get the best speckled images. </li>
<li>Over-microinjected cells will have densely packed speckles that do not appear to be discrete. Stress fibers and the lamellipodia will lose its dotted appearance.</li>
<li>Under-microinjected cells will appear very dim through the eyepiece, containing speckles that are far apart (&gt;10 speckles apart) and appearing randomly scattered. Stress fibers and the lamellipodia in these cells will be indistinguishable.</li>
<li>Key to obtaining &lsquo;good&rsquo; speckle movies is maintaining focus. This is particularly important for post-analysis involving speckle flow and turnover measurements. Thus the quality of the quantitative analysis will depend on locking in the focus for the duration of the time-lapse. This can be particularly challenging if the imaging plane is quite thin. Other factors will depend on the stability of the microscope system, particularly the stage housing and the imaging chamber. Contrast-based focusing can be used if image interval lengths are sufficient to accommodate this feature. Other options include purchasing a near-infrared based focusing system that provides reliable real-time focus adjustments. You can find these offerings from Nikon, Olympus and ASI. </li>
</ol>
Section 5: qFSM analysis
This section is not discussed in detail, but the information on the analysis software can be found here [5]. Software download requests can be made on our website (lccb.scripps.edu). Topics Covered:
<ul>
<li>Noise calibration</li>
<li>Speckle detection</li>
<li>Tracking using a hybrid approach of image correlation-based tracking of the coarse motion of speckle areas and single particle tracking of the detailed motion of individual speckles.</li>
<li>Computation of the polymer assembly and disassembly rates from the intensity fluctuations during speckle appearance and disappearance events. </li>
</ul>

<ol>
	<li>Waterman-Storer, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fluorescent+speckle+microscopy+(FSM)+of+microtubules+and+actin+in+living+cells.">Fluorescent speckle microscopy (FSM) of microtubules and actin in living cells.</a> Curr Protoc Cell Biol. Chapter 4, Unit 4.10-Unit 4.10 (2002).</li><li>Ponti, A., Machacek, M., Gupton, S. L., Waterman-Storer, C. M., Danuser, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Two+distinct+actin+networks+drive+the+protrusion+of+migrating+cells.">Two distinct actin networks drive the protrusion of migrating cells.</a> Science. 305, 1782-1786 (2004).</li><li>Zhang, X. F., Schaefer, A. W., Burnette, D. T., Schoonderwoert, V. T., Forscher, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Rho-dependent+contractile+responses+in+the+neuronal+growth+cone+are+independent+of+classical+peripheral+retrograde+actin+flow.">Rho-dependent contractile responses in the neuronal growth cone are independent of classical peripheral retrograde actin flow.</a> Neuron. 40, 931-944 (2003).</li><li>Salmon, W. C., Adams, M. C., Waterman-Storer, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dual-wavelength+fluorescent+speckle+microscopy+reveals+coupling+of+microtubule+and+actin+movements+in+migrating+cells.">Dual-wavelength fluorescent speckle microscopy reveals coupling of microtubule and actin movements in migrating cells.</a> J Cell Biol. 158, 31-37 (2002).</li><li>Danuser, G., Waterman-Storer, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Quantitative+fluorescent+speckle+microscopy+of+cytoskeleton+dynamics.">Quantitative fluorescent speckle microscopy of cytoskeleton dynamics.</a> Annu Rev Biophys Biomol Struct. 35, 361-387 (2006).</li></ol>

The authors have nothing to disclose.

Several key factors are crucial for successfully obtaining speckle images, however good speckles begin and end with the quality of your fluorescently labeled actin. A high labeling ratio of dye to actin monomer, set around 0.4 to 0.7 will ensure that speckles will appear bright and discrete, while the labeled protein itself should be soluble and free of aggregates. Equally important is the microinjection procedure. To ensure normal cell homeostasis (and survival), cells need to be injected with a low flow pressure, int...

<table><tbody><tr><td colspan="4">&lt;p&gt;&lt;strong&gt;&lt;em&gt;Injection buffer (IB)&lt;/em&gt;&lt;/strong&gt;&lt;br/&gt; 50 mM potassium glutamate&lt;br/&gt; 0.5 M MgCl2 (&lt;em&gt;APPENDIX 2A&lt;/em&gt;)&lt;br/&gt; Store up to 2 years at &amp;#8722;20&amp;#x00B0; C&lt;strong&gt;&lt;/strong&gt;&lt;/p&gt;&lt;p&gt;&lt;strong&gt;&lt;em&gt;&lt;/em&gt;&lt;/strong&gt;&lt;/p&gt;&lt;p&gt;&lt;strong&gt;&lt;em&gt;G-buffer&lt;/em&gt;&lt;/strong&gt;&lt;br/&gt; 2 mM Tris&amp;#8901;Cl, pH 8.0 (&lt;em&gt;APPENDIX 2A&lt;/em&gt;)&lt;br/&gt; 0.2 mM CaCl2 (&lt;em&gt;APPENDIX 2A&lt;/em&gt;)&lt;br/&gt; Add just before use:&lt;br/&gt; 0.2 mM ATP (see recipe for 100 mM)&lt;br/&gt; 0.5 mM 2-mercapt&amp;#8211;thanol&lt;/p&gt;Store up to 1 day at 4&amp;#x00B0; C</td></tr></tbody></table>

live cell imaging of f-actin dynamics via fluorescent speckle microscopy (fsm)

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique advantage of FSM is its ability to capture the movement and turnover kinetics (assembly/disassembly) of the F-actin network within living cells. This technique is particularly useful in deriving quantitative measurements of F-actin dynamics when paired with computer vision software (qFSM).  We describe  the selection, microinjection and visualization of fluorescent actin probes in living cells. Importantly, similar procedures are applicable to visualizing other macomolecular assemblies. FSM has been demonstrated for microtubules, intermediate filaments, and adhesion complexes.  

Watch this Scientific Journal Video about Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy FSM at JoVE.com

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy FSM

Selection, microinjection, and imaging of fluorescently-labeled F-actin via fluorescent speckle microscopy (FSM).

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy (FSM)

In this protocol we describe the use of Fluorescent Speckle Microscopy (FSM) to capture high-resolution images of actin dynamics in PtK1 cells. A unique ...

live-cell-imaging-f-actin-dynamics-via-fluorescent-speckle-microscopy

Universidad de Cartagena UDEC

Research

JoVE Journal

Biology

15.9K Views.  Scripps Institute. Selection, microinjection, and imaging of fluorescently-labeled F-actin via fluorescent speckle microscopy (FSM). 

Video: Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy FSM

Live-cell Imaging of Migrating Cells Expressing Fluorescently-tagged Proteins in a Three-dimensional Matrix

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound healing, tissue regeneration, and cancer metastasis, cells must navigate through complex, three-dimensional extracellular tissue. To better understand the mechanisms behind these biological processes, it is important to examine the roles of the proteins responsible for driving cell migration. Here, we outline a protocol to study the mechanisms of cell migration using the epithelial cell line (MDCK), and a three-dimensional, fibrous, self-polymerizing matrix as a model system. This optically clear extracellular matrix is easily amenable to live-cell imaging studies and better mimics the physiological, soft tissue environment. This report demonstrates a technique for directly visualizing protein localization and dynamics, and deformation of the surrounding three-dimensional matrix. 
Examination of protein localization and dynamics during cellular processes provides key insight into protein functions. Genetically encoded fluorescent tags provide a unique method for observing protein localization and dynamics. Using this technique, we can analyze the subcellular accumulation of key, force-generating cytoskeletal components in real-time as the cell maneuvers through the matrix. In addition, using multiple fluorescent tags with different wavelengths, we can examine the localization of multiple proteins simultaneously, thus allowing us to test, for example, whether different proteins have similar or divergent roles. Furthermore, the dynamics of fluorescently tagged proteins can be quantified using Fluorescent Recovery After Photobleaching (FRAP) analysis. This measurement assays the protein mobility and how stably bound the proteins are to the cytoskeletal network.
By combining live-cell imaging with the treatment of protein function inhibitors, we can examine in real-time the changes in the distribution of proteins and morphology of migrating cells. Furthermore, we also combine live-cell imaging with the use of fluorescent tracer particles embedded within the matrix to visualize the matrix deformation during cell migration. Thus, we can visualize how a migrating cell distributes force-generating proteins, and where the traction forces are exerted to the surrounding matrix. Through these techniques, we can gain valuable insight into the roles of specific proteins and their contributions to the mechanisms of cell migration.

Cellular processes such as cell migration have traditionally been studied on two-dimensional, stiff plastic surfaces. This report describes a technique for directly visualizing protein localization and analyzing protein dynamics in cells migrating in a more physiologically relevant, three-dimensional matrix.

Traditionally, cell migration has been studied on two-dimensional, stiff plastic surfaces. However, during important biological processes such as wound ...

Bioengineering

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration have been thoroughly analyzed in vitro. However, in vivo cell migration somehow differs from in vitro migration, and has proven more difficult to analyze, being less accessible to direct observation and manipulation. This protocol uses the migration of the prospective prechordal plate in the early zebrafish embryo as a model system to study the function of candidate genes in cell migration. Prechordal plate progenitors form a group of cells which, during gastrulation, undergoes a directed migration from the embryonic organizer to the animal pole of the embryo. The proposed protocol uses cell transplantation to create mosaic embryos. This offers the combined advantages of labeling isolated cells, which is key to good imaging, and of limiting gain/loss of function effects to the observed cells, hence ensuring cell-autonomous effects. We describe here how we assessed the function of the TORC2 component Sin1 in cell migration, but the protocol can be used to analyze the function of any candidate gene in controlling cell migration in vivo.

Analyzing In Vivo Cell Migration using Cell Transplantations and Time-lapse Imaging in Zebrafish Embryos

Combining cell transplantation, cytoskeletal labeling and loss/gain of function approaches, this protocol describes how the migrating zebrafish prospective prechordal plate can be used to analyze the function of a candidate gene in in vivo cell migration.

Cell migration is key to many physiological and pathological conditions, including cancer metastasis. The cellular and molecular bases of cell migration ...

Developmental Biology

From Fast Fluorescence Imaging to Molecular Diffusion Law on Live Cell Membranes in a Commercial Microscope

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the regulation of many cellular functions. However, due to the fast dynamics and the tiny structures involved, a very high spatio-temporal resolution is required to catch the real behavior of molecules. Here we present the experimental protocol for studying the dynamics of fluorescently-labeled plasma-membrane proteins and lipids in live cells with high spatiotemporal resolution. Notably, this approach doesn&#8217;t need to track each molecule, but it calculates population behavior using all molecules in a given region of the membrane. The starting point is a fast imaging of a given region on the membrane. Afterwards, a complete spatio-temporal autocorrelation function is calculated correlating acquired images at increasing time delays, for example each 2, 3, n repetitions. It is possible to demonstrate that the width of the peak of the spatial autocorrelation function increases at increasing time delay as a function of particle movement due to diffusion. Therefore, fitting of the series of autocorrelation functions enables to extract the actual protein mean square displacement from imaging (iMSD), here presented in the form of apparent diffusivity vs average displacement. This yields a quantitative view of the average dynamics of single molecules with nanometer accuracy. By using a GFP-tagged variant of the Transferrin Receptor (TfR) and an ATTO488 labeled 1-palmitoyl-2-hydroxy-sn-glycero-3-phosphoethanolamine (PPE) it is possible to observe the spatiotemporal regulation of protein and lipid diffusion on &#181;m-sized membrane regions in the micro-to-milli-second time range.

Spatial distribution and temporal dynamics of plasma membrane proteins and lipids is a hot topic in biology. Here this issue is addressed by a spatio-temporal image fluctuation analysis that provides conceptually the same physical quantities of single particle tracking, but it uses small molecular labels and standard microscopy setups.

It has become increasingly evident that the spatial distribution and the motion of membrane components like lipids and proteins are key factors in the ...

Cortical Actin Flow in T Cells Quantified by Spatio-temporal Image Correlation Spectroscopy of Structured Illumination Microscopy Data

Filamentous-actin plays a crucial role in a majority of cell processes including motility and, in immune cells, the formation of a key cell-cell interaction known as the immunological synapse. F-actin is also speculated to play a role in regulating molecular distributions at the membrane of cells including sub-membranous vesicle dynamics and protein clustering. While standard light microscope techniques allow generalized and diffraction-limited observations to be made, many cellular and molecular events including clustering and molecular flow occur in populations at length-scales far below the resolving power of standard light microscopy. By combining total internal reflection fluorescence with the super resolution imaging method structured illumination microscopy, the two-dimensional molecular flow of F-actin at the immune synapse of T cells was recorded. Spatio-temporal image correlation spectroscopy (STICS) was then applied, which generates quantifiable results in the form of velocity histograms and vector maps representing flow directionality and magnitude. This protocol describes the combination of super-resolution imaging and STICS techniques to generate flow vectors at sub-diffraction levels of detail. This technique was used to confirm an actin flow that is symmetrically retrograde and centripetal throughout the periphery of T cells upon synapse formation.

To investigate flow velocities and directionality of filamentous-actin at the T cell immunological synapse, live-cell super-resolution imaging is combined with total internal reflection fluorescence and quantified with spatio-temporal image correlation spectroscopy.

Filamentous-actin plays a crucial role in a majority of cell processes including motility and, in immune cells, the formation of a key cell-cell ...

Immunology and Infection

Using Microfluidics and Fluorescence Microscopy to Study the Assembly Dynamics of Single Actin Filaments and Bundles

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor individual reactions live in well-controlled conditions. To do so, live single-filament experiments have emerged over the past 20 years, mostly using total internal reflection fluorescence (TIRF) microscopy, and have provided a trove of key results. In 2011, in order to further expand the possibilities of these experiments and to avoid recurring problematic artifacts, we introduced simple microfluidics in these assays. This study details our basic protocol, where individual actin filaments are anchored by one end to the passivated coverslip surface, align with the flow, and can be successively exposed to different protein solutions. We also present the protocols for specific applications and explain how controlled mechanical forces can be applied, thanks to the viscous drag of the flowing solution. We highlight the technical caveats of these experiments and briefly present possible developments based on this technique. These protocols and explanations, along with today&#39;s availability of easy-to-use microfluidics equipment, should allow non-specialists to implement this assay in their labs.

We present protocols for simple actin filament microfluidic assays, in combination with fluorescence microscopy, that allow one to accurately monitor individual actin filaments in real-time while sequentially exposing them to different protein solutions.

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor ...

Biochemistry

Visualizing Intracellular SNARE Trafficking by Fluorescence Lifetime Imaging Microscopy

Soluble N-ethylmaleimide sensitive fusion protein (NSF) attachment protein receptor (SNARE) proteins are key for membrane trafficking, as they catalyze membrane fusion within eukaryotic cells. The SNARE protein family consists of about 36 different members. Specific intracellular transport routes are catalyzed by specific sets of 3 or 4 SNARE proteins that thereby contribute to the specificity and fidelity of membrane trafficking. However, studying the precise function of SNARE proteins is technically challenging, because SNAREs are highly abundant and functionally redundant, with most SNAREs having multiple and overlapping functions. In this protocol, a new method for the visualization of SNARE complex formation in live cells is described. This method is based on expressing SNARE proteins C-terminally fused to fluorescent proteins and measuring their interaction by F&#246;rster resonance energy transfer (FRET) employing fluorescence lifetime imaging microscopy (FLIM). By fitting the fluorescence lifetime histograms with a multicomponent decay model, FRET-FLIM allows (semi-)quantitative estimation of the fraction of the SNARE complex formation at different vesicles. This protocol has been successfully applied to visualize SNARE complex formation at the plasma membrane and at endosomal compartments in mammalian cell lines and primary immune cells, and can be readily extended to study SNARE functions at other organelles in animal, plant, and fungal cells.

This protocol describes a new method allowing for the quantitative visualization of complex formation of SNARE proteins, based on F&#246;rster resonance energy transfer, and fluorescence lifetime imaging microscopy.

Soluble N-ethylmaleimide sensitive fusion protein (NSF) attachment protein receptor (SNARE) proteins are key for membrane trafficking, as they catalyze ...

Study of the Actin Cytoskeleton in Live Endothelial Cells Expressing GFP-Actin

The microvascular endothelium plays an important role as a selectively permeable barrier to fluids and solutes. The adhesive junctions between endothelial cells regulate permeability of the endothelium, and many studies have indicated the important contribution of the actin cytoskeleton to determining junctional integrity1-5. A cortical actin belt is thought to be important for the maintenance of stable junctions1, 2, 4, 5. In contrast, actin stress fibers are thought to generate centripetal tension within endothelial cells that weakens junctions2-5. Much of this theory has been based on studies in which endothelial cells are treated with inflammatory mediators known to increase endothelial permeability, and then fixing the cells and labeling F-actin for microscopic observation. However, these studies provide a very limited understanding of the role of the actin cytoskeleton because images of fixed cells provide only snapshots in time with no information about the dynamics of actin structures5. 
Live-cell imaging allows incorporation of the dynamic nature of the actin cytoskeleton into the studies of the mechanisms determining endothelial barrier integrity. A major advantage of this method is that the impact of various inflammatory stimuli on actin structures in endothelial cells can be assessed in the same set of living cells before and after treatment, removing potential bias that may occur when observing fixed specimens. Human umbilical vein endothelial cells (HUVEC) are transfected with a GFP-&beta;-actin plasmid and grown to confluence on glass coverslips. Time-lapse images of GFP-actin in confluent HUVEC are captured before and after the addition of inflammatory mediators that elicit time-dependent changes in endothelial barrier integrity. These studies enable visual observation of the fluid sequence of changes in the actin cytoskeleton that contribute to endothelial barrier disruption and restoration. 
Our results consistently show local, actin-rich lamellipodia formation and turnover in endothelial cells. The formation and movement of actin stress fibers can also be observed. An analysis of the frequency of formation and turnover of the local lamellipodia, before and after treatment with inflammatory stimuli can be documented by kymograph analyses. These studies provide important information on the dynamic nature of the actin cytoskeleton in endothelial cells that can used to discover previously unidentified molecular mechanisms important for the maintenance of endothelial barrier integrity.

Microscopic imaging of live endothelial cells expressing GFP-actin allows characterization of dynamic changes in cytoskeletal structures. Unlike techniques that use fixed specimens, this method provides a detailed assessment of temporal changes in the actin cytoskeleton in the same cells before, during, and after various physical, pharmacological, or inflammatory stimuli.

The microvascular endothelium plays an important role as a selectively permeable barrier to fluids and solutes. The adhesive junctions between endothelial ...

Fluorescence Imaging with One-nanometer Accuracy (FIONA)

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in the x-y plane. Here a summary of the FIONA technique is reported and examples of research that have been performed using FIONA are briefly described. First, how to set up the required equipment for FIONA experiments, i.e., a total internal reflection fluorescence microscopy (TIRFM), with details on aligning the optics, is described. Then how to carry out a simple FIONA experiment on localizing immobilized Cy3-DNA single molecules using appropriate protocols, followed by the use of FIONA to measure the 36 nm step size of a single truncated myosin Va motor labeled with a quantum dot, is illustrated. Lastly, recent effort to extend the application of FIONA to thick samples is reported. It is shown that, using a water immersion objective and quantum dots soaked deep in sol-gels and rabbit eye corneas (&#62;200 &#181;m), localization precision of 2-3 nm can be achieved.

Fluorescence Imaging with One-nanometer Accuracy FIONA

Single fluorophores can be localized with nanometer precision using FIONA. Here a summary of the FIONA technique is reported, and how to carry out FIONA experiments is described.

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in ...

Analyses of Actin Dynamics, Clutch Coupling and Traction Force for Growth Cone Advance

To establish functional networks, neurons must migrate to their appropriate destinations and then extend axons toward their target cells. These processes depend on the advances of growth cones that located at the tips of neurites. Axonal growth cones generate driving forces by sensing their local microenvironment and modulating cytoskeletal dynamics and actin-adhesion coupling (clutch coupling). Decades of research have led to the identification of guidance molecules, their receptors, and downstream signaling cascades for regulating neuronal migration and axonal guidance; however, the molecular machineries required for generating forces to drive growth cone advance and navigation are just beginning to be elucidated. At the leading edge of neuronal growth cones, actin filaments undergo retrograde flow, which is powered by actin polymerization and actomyosin contraction. A clutch coupling between F-actin retrograde flow and adhesive substrate generates traction forces for growth cone advance. The present study describes a detailed protocol for monitoring F-actin retrograde flow by single speckle imaging. Importantly, when combined with an F-actin marker Lifeact, this technique can quantify 1) the F-actin polymerization rate and 2) the clutch coupling efficiency between F-actin retrograde flow and the adhesive substrate. Both are critical variables for generating forces for growth cone advance and navigation. In addition, the present study describes a detailed protocol of traction force microscopy, which can quantify 3) traction force generated by growth cones. Thus, by coupling the analyses of&#160;single speckle imaging and traction force microscopy, investigators can monitor the molecular mechanics underlying growth cone advance and navigation.

To advance, growth cones must exert traction forces against the external environment. The generation of traction forces is dependent on actin dynamics and clutch coupling. The present study describes methods for analyzing actin dynamics, clutch coupling and traction forces for growth cone advance.

To establish functional networks, neurons must migrate to their appropriate destinations and then extend axons toward their target cells. These processes ...

Neuroscience

Micromanipulation Techniques Allowing Analysis of Morphogenetic Dynamics and Turnover of Cytoskeletal Regulators

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how fluorescent recovery after photobleaching (FRAP) and photoactivation techniques can be used to study the spatiotemporal dynamics of proteins in subcellular locations. We also show how these techniques enable straightforward determination of various parameters linked to actin cytoskeletal regulation and cell motility. Moreover, the microinjection of cells is additionally described as an alternative treatment (potentially preceding or complementing the aforementioned photomanipulation techniques) to trigger instantaneous effects of translocated proteins on cell morphology and function. Micromanipulation such as protein injection or local application of plasma membrane-permeable drugs or cytoskeletal inhibitors can serve as powerful tool to record immediate consequences of a given treatment on cell behavior at the single cell and subcellular level. This is exemplified here by immediate induction of lamellipodial cell edge protrusion by the injection of recombinant Rac1 protein, as established a quarter-century ago. In addition, we provide a protocol for determining the turnover of enhanced green fluorescent protein (EGFP)-VASP, an actin filament polymerase prominently accumulating at lamellipodial tips of B16-F1 cells, employing FRAP and including associated data analysis and curve fitting. We also present guidelines for estimating the rates of lamellipodial actin network polymerization, as exemplified by cells expressing EGFP-tagged &#946;-actin. Finally, instructions are given for how to investigate the rates of actin monomer mobility within the cell cytoplasm, followed by actin incorporation at sites of rapid filament assembly, such as the tips of protruding lamellipodia, using photoactivation approaches. None of these protocols is&#160;restricted to components or regulators of the actin cytoskeleton, but can easily be extended to explore in analogous fashion the spatiotemporal dynamics and function of proteins in various different subcellular structures or functional contexts.

We describe how micro- and photomanipulation techniques such as FRAP and photoactivation enable the determination of motility parameters and the spatiotemporal dynamics of proteins within migrating cells. Experimental readouts include subcellular dynamics and turnover of motility regulators or of the underlying actin cytoskeleton.

Live Cell Imaging of F-actin Dynamics via Fluorescent Speckle Microscopy (FSM)

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Study of the Actin Cytoskeleton in Live Endothelial Cells Expressing GFP-Actin

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Analyses of Actin Dynamics, Clutch Coupling and Traction Force for Growth Cone Advance

Micromanipulation Techniques Allowing Analysis of Morphogenetic Dynamics and Turnover of Cytoskeletal Regulators