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...

Injection buffer (IB) 
 50 mM potassium glutamate 
 0.5 M MgCl2 (APPENDIX 2A) 
 Store up to 2 years at &#8722;20&deg; C
&nbsp;
G-buffer 
 2 mM Tris&#8901;Cl, pH 8.0 (APPENDIX 2A) 
 0.2 mM CaCl2 (APPENDIX 2A) 
 Add just before use: 
 0.2 mM ATP (see recipe for 100 mM) 
 0.5 mM 2-mercaptoethanol
Store up to 1 day at 4&deg; C

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

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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

Actin Co-Sedimentation Assay; for the Analysis of Protein Binding to F-Actin

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates tension and helps maintains cellular shape.  Although the actin cytoskeleton is a rigid structure, it is a dynamic structure that is constantly remodeling.  A number of proteins can bind to the actin cytoskeleton.  The binding of a particular protein to F-actin is often desired to support cell biological observations or to further understand dynamic processes due to remodeling of the actin cytoskeleton. The actin co-sedimentation assay is an in vitro assay routinely used to analyze the binding of specific proteins or protein domains with F-actin.  The basic principles of the assay involve an incubation of the protein of interest (full length or domain of) with F-actin, ultracentrifugation step to pellet F-actin and analysis of the protein co-sedimenting with F-actin.  Actin co-sedimentation assays can be designed accordingly to measure actin binding affinities and in competition assays.

Proteins bind to filamentous actin (F-actin) through distinct actin binding modules. In this video we demonstrate the procedure of actin co-sedimentation, which is an in vitro assay routinely used to analyze proteins or specific domains that bind F-actin.

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates ...

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Assembly, Loading, and Alignment of an Analytical Ultracentrifuge Sample Cell

The analytical ultracentrifuge (AUC) is a powerful biophysical tool that allows us to record macromolecular sedimentation profiles during high speed centrifugation. When properly planned and executed, an AUC sedimentation velocity or sedimentation equilibrium experiment can reveal a great deal about a protein in regards to size and shape, sample purity, sedimentation coefficient, oligomerization states and protein-protein interactions.
This technique, however, requires a rigorous level of technical attention. Sample cells hold a sectored center piece sandwiched between two window assemblies. They are sealed with a torque pressure of around 120-140 in/lbs. Reference buffer and sample are loaded into the centerpiece sectors and then after sealing, the cells are precisely aligned into a titanium rotor so that the optical detection systems scan both sample and reference buffer in the same radial path midline through each centerpiece sector while rotating at speeds of up to 60, 000 rpm and under very high vacuum
Not only is proper sample cell assembly critical, sample cell components are very expensive and must be properly cared for to ensure they are in optimum working condition in order to avoid leaks and breakage during experiments. Handle windows carefully, for even the slightest crack or scratch can lead to breakage in the centrifuge. The contact between centerpiece and windows must be as tight as possible; i.e. no Newton s rings should be visible after torque pressure is applied. Dust, lint, scratches and oils on either the windows or the centerpiece all compromise this contact and can very easily lead to leaking of solutions from one sector to another or leaking out of the centerpiece all together. Not only are precious samples lost, leaking of solutions during an experiment will cause an imbalance of pressure in the cell that often leads to broken windows and centerpieces. In addition, plug gaskets and housing plugs must be securely in place to avoid solutions being pulled out of the centerpiece sector through the loading holes by the high vacuum in the centrifuge chamber. Window liners and gaskets must be free of breaks and cracks that could cause movement resulting in broken windows.
This video will demonstrate our procedures of sample cell assembly, torque, loading and rotor alignment to help minimize component damage, solution leaking and breakage during the perfect AUC experiment.

The analytical ultracentrifuge (AUC) sample cell holds sample and reference buffer and during experiments and is exposed to high vacuum and rotor speeds up to 60,000 rpm. This video will demonstrate the rigorous attention to detail necessary for assembly, loading and alignment of this very important component of an AUC experiment.

The analytical ultracentrifuge (AUC) is a powerful biophysical tool that allows us to record macromolecular sedimentation profiles during high speed ...

Fluorescent Labeling of COS-7 Expressing SNAP-tag Fusion Proteins for Live Cell Imaging

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. These systems offer a broad selection of fluorescent substrates optimized for a range of imaging instrumentation. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again. There are two steps to using this system: cloning and expression of the protein of interest as a SNAP-tag fusion, and labeling of the fusion with the SNAP-tag substrate of choice. The SNAP-tag is a small protein based on human O6-alkylguanine-DNA-alkyltransferase (hAGT), a DNA repair protein. SNAP-tag labels are dyes conjugated to guanine or chloropyrimidine leaving groups via a benzyl linker. In the labeling reaction, the substituted benzyl group of the substrate is covalently attached to the SNAP-tag. CLIP-tag is a modified version of SNAP-tag, engineered to react with benzylcytosine rather than benzylguanine derivatives. When used in conjunction with SNAP-tag, CLIP-tag enables the orthogonal and complementary labeling of two proteins simultaneously in the same cells.

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest in live cells. Once cloned and expressed, the tagged protein can be used with a variety of substrates for numerous downstream applications without having to clone again.

SNAP-tag and CLIP-tag protein labeling systems enable the specific, covalent attachment of molecules, including fluorescent dyes, to a protein of interest ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

Intravital Microscopy for Imaging Subcellular Structures in Live Mice Expressing Fluorescent Proteins

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, we use the salivary glands of live mice since they provide several advantages. First, they can be easily exposed to enable access to the optics, and stabilized to facilitate the reduction of the motion artifacts due to heartbeat and respiration. This significantly facilitates imaging and tracking small subcellular structures. Second, most of the cell populations of the salivary glands are accessible from the surface of the organ. This permits the use of confocal microscopy that has a higher spatial resolution than other techniques that have been used for in vivo imaging, such as two-photon microscopy. Finally, salivary glands can be easily manipulated pharmacologically and genetically, thus providing a robust system to investigate biological processes at a molecular level.
In this study we focus on a protocol designed to follow the kinetics of the exocytosis of secretory granules in acinar cells and the dynamics of the apical plasma membrane where the secretory granules fuse upon stimulation of the beta-adrenergic receptors. Specifically, we used a transgenic mouse that co-expresses cytosolic GFP and a membrane-targeted peptide fused with the fluorescent protein tandem-Tomato. However, the procedures that we used to stabilize and image the salivary glands can be extended to other mouse models and coupled to other approaches to label in vivo cellular components, enabling the visualization of various subcellular structures, such as endosomes, lysosomes, mitochondria, and the actin cytoskeleton.

Intravital microscopy is a powerful tool that enables imaging various biological processes in live animals. In this article, we present a detailed method for imaging the dynamics of subcellular structures, such as the secretory granules, in the salivary glands of live mice.

Here we describe a procedure to image subcellular structures in live rodents that is based on the use of confocal intravital microscopy. As a model organ, ...

Real-time Imaging of Plant Cell Surface Dynamics with Variable-angle Epifluorescence Microscopy

A plant&#8217;s cell surface is its interface for perceiving environmental cues; it responds with cell biological changes such as membrane trafficking and cytoskeletal rearrangement. Real-time and high-resolution image analysis of such intracellular events will increase the understanding of plant cell biology at the molecular level. Variable angle epifluorescence microscopy (VAEM) is an emerging technique that provides high-quality, time-lapse images of fluorescently-labeled proteins on the plant cell surface. In this article, practical procedures are described for VAEM specimen preparation, adjustment of the VAEM optical system, movie capturing and image analysis. As an example of VAEM observation, representative results are presented on the dynamics of PATROL1. This is a protein essential for stomatal movement, thought to be involved in proton pump delivery to plasma membranes in the stomatal complex of Arabidopsis thaliana. VAEM real-time observation of guard cells and subsidiary cells in A. thaliana cotyledons showed that fluorescently-tagged PATROL1 appeared as dot-like structures on plasma membranes for several seconds and then disappeared. Kymograph analysis of VAEM movie data determined the time distribution of the presence (termed &#8216;residence time&#8217;) of the dot-like structures. The use of VAEM is discussed in the context of this example.

The goal of this protocol is to demonstrate how to monitor fluorescently-tagged protein dynamics on plant cell surfaces with variable-angle epifluorescence microscopy, showing blinking dots of GFP-tagged PATROL1, a membrane trafficking protein, in the cell cortex of the stomatal complex in Arabidopsis thaliana.

A plant&#8217;s cell surface is its interface for perceiving environmental cues; it responds with cell biological changes such as membrane trafficking and ...

Three-dimensional Super Resolution Microscopy of F-actin Filaments by Interferometric PhotoActivated Localization Microscopy (iPALM)

Fluorescence microscopy enables direct visualization of specific biomolecules within cells. However, for conventional fluorescence microscopy, the spatial resolution is restricted by diffraction to ~ 200 nm within the image plane and &#62; 500 nm along the optical axis. As a result, fluorescence microscopy has long been severely limited in the observation of ultrastructural features within cells. The recent development of super resolution microscopy methods has overcome this limitation. In particular, the advent of photoswitchable fluorophores enables localization-based super resolution microscopy, which provides resolving power approaching the molecular-length scale. Here, we describe the application of a three-dimensional super resolution microscopy method based on single-molecule localization microscopy and multiphase interferometry, called interferometric PhotoActivated Localization Microscopy (iPALM). This method provides nearly isotropic resolution on the order of 20 nm in all three dimensions. Protocols for visualizing the filamentous actin cytoskeleton, including specimen preparation and operation of the iPALM instrument, are described here. These protocols are also readily adaptable and instructive for the study of other ultrastructural features in cells.

Three-dimensional Super Resolution Microscopy of F-actin Filaments by Interferometric PhotoActivated Localization Microscopy iPALM

We present a protocol for the application of interferometric PhotoActivated Localization Microscopy (iPALM), a 3-dimensional single-molecule localization super resolution microscopy method, to the imaging of the actin cytoskeleton in adherent mammalian cells. This approach allows light-based visualization of nanoscale structural features that would otherwise remain unresolved by conventional diffraction-limited optical microscopy.

Fluorescence microscopy enables direct visualization of specific biomolecules within cells. However, for conventional fluorescence microscopy, the spatial ...

Exploiting Live Imaging to Track Nuclei During Myoblast Differentiation and Fusion

Nuclear positioning within cells is important for multiple cellular processes in development and regeneration. The most intriguing example of nuclear positioning occurs during skeletal muscle differentiation. Muscle fibers (myofibers) are multinucleated cells formed by the fusion of muscle precursor cells (myoblasts) derived from muscle stem cells (satellite cells) that undergo proliferation and differentiation. Correct nuclear positioning within myofibers is required for the proper muscle regeneration and function. The common procedure to assess myoblast differentiation and myofiber formation relies on fixed cells analyzed by immunofluorescence, which impedes the study of nuclear movement and cell behavior over time. Here, we describe a method for the analysis of myoblast differentiation and myofiber formation by live cell imaging. We provide a software for automated nuclear tracking to obtain a high-throughput quantitative characterization of nuclear dynamics and myoblast behavior (i.e., the trajectory) during differentiation and fusion.

Skeletal muscle differentiation is a highly dynamic process, which particularly relies on nuclear positioning. Here, we describe a method to track nuclei movements by live cell imaging during myoblast differentiation and myotube formation and to perform a quantitative characterization of nuclei dynamics by extracting information from automatic tracking.

Nuclear positioning within cells is important for multiple cellular processes in development and regeneration. The most intriguing example of nuclear ...

Examination of Mitotic and Meiotic Fission Yeast Nuclear Dynamics by Fluorescence Live-cell Microscopy

Live-cell imaging is a microscopy technique used to examine cell and protein dynamics in living cells. This imaging method is not toxic, generally does not interfere with cell physiology, and requires minimal experimental handling. The low levels of technical interference enable researchers to study cells across multiple cycles of mitosis and to observe meiosis from beginning to end. Using fluorescent tags such as Green Fluorescent Protein (GFP) and Red Fluorescent Protein (RFP), researchers can analyze different factors whose functions are important for processes like transcription, DNA replication, cohesion, and segregation. Coupled with data analysis using Fiji (a free, optimized ImageJ version), live-cell imaging offers various ways of assessing protein movement, localization, stability, and timing, as well as nuclear dynamics and chromosome segregation. However, as is the case with other microscopy methods, live-cell imaging is limited by the intrinsic properties of light, which put a limit to the resolution power at high magnifications, and is also sensitive to photobleaching or phototoxicity at high wavelength frequencies. However, with some care, investigators can bypass these physical limitations by carefully choosing the right conditions, strains, and fluorescent markers to allow for the appropriate visualization of mitotic and meiotic events.

Here, we present live-cell imaging which is a non-toxic microscopy method that allows researchers to study protein behavior and nuclear dynamics in living fission yeast cells during mitosis and meiosis.

Live-cell imaging is a microscopy technique used to examine cell and protein dynamics in living cells. This imaging method is not toxic, generally does ...