We are grateful to the German Research Foundation (DFG) for financial support (grant Nr. RO2414/5-1 to KR).

<ol>
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The authors have nothing to disclose.

Here we discuss critical steps in the techniques described in this article, and how they can be optimized for application in different experimental conditions.
Microinjection is a method that can be applied to monitor in cells the instant effects from introducing exogenous proteins, inhibitors, or drugs. It can be particularly advantageous for determining the functions of proteins in difficult to transfect cell types or in situations when long-term expression is not desired. It must be noted that survival of certain cell types varies depending on the extracellular matrix they are seeded on. Most endothelial, epithelial, or fibroblast-like cell types, even small ones like fish keratocytes (see Dang et al.21 and Anderson and Cross22) can be successfully injected. However, there are exceptions, such as B16-F1 cells seeded on laminin, which constitute an excellent model system of cell migration, but are incompatible with injection on this type of substratum for unknown reason. For NIH3T3 fibroblast cells, we routinely perform injections on fibronectin substratum, and additional photomanipulation techniques such as FRAP (even with photoactivation; shown for B16-F1 cells here) can be equally well performed in these fibroblasts (see e.g., K&#246;stler et al.3). It must also be considered that different proteins, according to their functional properties and the experiment goals, may take different amounts of time to cause changes, varying from seconds to hours. An advantage of the technique is that the dosage/concentration of exogenous agent can be controlled more accurately at the single cell level than e.g., when using plasmid transfection. In addition, fluorescent tagging of a protein is not a necessity to guarantee its presence in the cell, which can increase flexibility if simultaneous multi-channel visualization of other fluorescently-tagged proteins is required. Microinjection can be particularly useful for analyzing instant effects of specific proteins or protein mixtures on dynamic changes of cell morphology or the cytoskeleton (e.g., Dang et al.21 for an example of instant effects on migration by the Arp2/3 complex inhibitor Arpin). A disadvantage of the technique is its invasiveness, which can cause cell damage or influence cell morphology. Therefore, an important consideration when performing microinjections is monitoring the cell viability. The method introduced here relies on manual manipulation. In conditions tested to be compatible with successful injections, such as fibroblasts growing on fibronectin substratum, the manual injection protocol described here allows a near 100% success rate; this is essential when combining this approach with sophisticated and time-consuming follow-up experiments including video microscopy or FRAP, as published previously3. This does not exclude that occasionally, individual cells might suffer from a microinjection event, which can be safely recognized by abrupt changes of contrast of both the nucleus and cytoplasm, followed by cell edge retraction. Such rare experimental cases are excluded and thus not considered for further analyses.
However, a half-automatic approach is also commonly used, for instance employing rapid (&#60;300 ms) machine-controlled needle lowering coincident with injection pressure increase, so that the needle only has to be positioned above each cell prior to respective injection. The success rate of half-automatic injections is by definition lower than the manual approach described above, simply because it is optimized for speed, followed by analysis of multiple cells that successfully survived this treatment; thus it does not rely on successful injection of an individual cell. Therefore, as opposed to single cell analysis, half-automatic injections are better suited for analyzing injection effects of several hundred cells, e.g., by video microscopy at low magnification or upon cell fixation and staining. Irrespective of the detailed approach employed, microinjection does not constitute an end-point assay, but can be combined with a variety of techniques, including FRAP or photoactivation3.
When determining the protein turnover rate by FRAP, the intensity of the laser must be optimized, depending on the microscope setup and imaging conditions (magnification, objectives, etc., as well as the cell type, structure, and fluorescent protein for photobleaching). Note that at optimal laser power, efficient bleaching is combined with the least possible photodamage, to avoid shrinkage or complete retraction of the structure under analysis (e.g., lamellipodia or filopodia) or even damage at the cellular level. Ideally, at least 70&#8211;80% of bleaching efficiency should be achieved, although complete bleaching may be hampered by extremely rapid turnover of the protein, in which case, anything above 50% might also be acceptable. Optimal bleaching power for a given structure and fluorescent dye should be experimentally tested, starting from a low laser power followed by its gradual increase. Of course, any fluorescent dye can by definition be bleached with laser light close to its peak of excitation (488 nm for frequently used green dyes such as FITC or EGFP). However, lasers with shorter wavelengths, such as near-UV lasers, deliver higher powers and can thus also be used for efficient bleaching of commonly used dyes. We routinely employ a 405 nm diode laser (120 mW) for bleaching of both EGFP and red fluorescent dyes (such as mCherry), albeit with slightly lower efficiency in case of the latter (data not shown). As the 405 nm-diode can also be used for photoactivation of PA-GFP (see below), it endows this system with maximal flexibility.
For the B16-F1 cell structures and fluorescent proteins photobleached here, 405 nm-laser powers between 65&#8211;100 mW were applied. When analyzing a photobleached region, it is important to consider whether the given structure is preserved in its original shape over the analysis time period. For instance, when analyzing turnover of proteins at lamellipodia tips, care should be taken whether the curvature of lamellipodia is significantly altered over time, as changes in curvature might lead to inaccurate results if the region/contour analyzed does not fully encompass the entirety of the structure in each measured frame. In addition, it should be noted that bundles embedded into lamellipodia, such as microspikes, might cause deviations in fluorescence intensity. As illustrated in Figure 2b (white arrow in 9 s time frame), a microspike-like structure is situated next to the measured photobleached region, but remains outside of it throughout the duration of measurement, and thus does not cause any inaccuracy. For analysis of protein turnover, important considerations when selecting location and size of analyzed regions are that their fluorescence over time should not be significantly influenced by changes in cell morphology or factors other than hard to avoid acquisition photobleaching. For instance, structures providing significant quantitative contribution to the analyzed structure should not move out of the measured region during analysis; in addition, unrelated, fluorescent entities such as vesicular structures that attract the protein should not enter the field of interest during analysis. For determining the rate of lamellipodial actin polymerization, care should be taken that no retracting or ruffling (i.e., upwards folding) lamellipodia are analyzed, as this will strongly influence the accuracy of the results. In addition, retraction of lamellipodial regions might appear as rapid rearward translocation, potentially leading to overestimation of rates of lamellipodial actin polymerization. An additional consideration is the distance of intracellular normalization regions (taken as reference positions for the correction of&#160;acquisition photobleaching) from the actual position of photobleaching, which should be large enough to avoid direct influence by the photobleached area.
When setting up optimal conditions for photoactivation of PA-GFP-tagged constructs, care should be taken to avoid instant bleaching during photoactivation. In our work, the best results were obtained with laser powers 5-10 times lower than normally employed for bleaching of EGFP. For image acquisition of photoactivated molecules, exposure time and time interval between frames should be optimized by considering the size of regions and structures to be photoactivated and analyzed, as well as the potential mobility of photoactivated proteins to other subcellular locations. As for all types of fluorescence imaging, maintenance of cell viability is crucial for obtaining physiologically relevant results.
In principle, green-to-red photoconversion of fluorescent proteins such as mEos or Dronpa variants12 constitutes an equally powerful method of following dynamics and turnover of subcellular structures such as the lamellipodium (see e.g., Burnette et al.23). The advantage of the latter method as opposed to PA-GFP would be the possibility to follow protein dynamics before and after conversion with two distinct colors, without the need to co-express an additional red fluorescent protein. However, in our preliminary experiments, the extent of contrast change and intensity of fluorescent signal achieved upon photoactivation of PA-GFP was larger as compared to photoconverted probes, perhaps due to the superior spectral features of green versus red fluorescent probes (data not shown). In any case, detailed studies on actin filament turnover in cell-edge protrusions such as lamellipodia or Vaccinia virus-induced actin tails have so far only been published using PA-GFP derivatives5,6,24.
When considering which cell region to analyze following photoactivation, several factors should be taken into account, which are discussed using the specific example shown here (incorporation of actin monomers at the cell edge upon activation in the cytosol), but can certainly be extrapolated to various analogous scientific problems. First, when measuring the rate of lamellipodial incorporation of cytosolically photoactivated proteins, for instance, in distinct experimental conditions (as shown in Dimchev et al.6), sizes of cytosolic regions and their distances to lamellipodial edges should be comparable between experimental groups. It is also important to consider that when photoactivating cytosolic regions, the cell thickness is greater in positions closer to the nucleus. Activating thicker cellular regions might result in higher amounts of activated proteins, given that the distribution of the protein to be activated is homogenously distributed in the cytosol. Lastly, expression levels of the protein to be activated can certainly be highly variable in individual cells. Due to all these considerations of variability, it is crucial to compare incorporation levels of cytosolically activated proteins elsewhere in the cell relative to the total fluorescence obtained upon activation in the specific regions.
We have described how microinjection can be used as a tool for investigating the effects of proteins on cell morphology and have exemplified this by demonstrating the potent induction of lamellipodial structures in NIH3T3 fibroblast cells microinjected with the small GTPase Rac1. We have previously applied this technique to interfere with Arp2/3 function in cells microinjected with the C-terminal WCA domain of Scar/WAVE3. Various parameters in microinjected cells can be analyzed by other assays, such as FRAP or photoactivation. We have described how FRAP and photoactivation can be employed for investigating the subcellular dynamics and mobility of actin monomers. FRAP has been used by our group previously5 to investigate the turnover of proteins localizing to lamellipodia, such as VASP, Abi, cortactin, cofilin, and capping protein, or for elucidating the turnover of components in focal adhesions in the presence and absence of Rac signaling4. Moreover, measuring actin polymerization rates can be accomplished by photobleaching EGFP-tagged &#946;-actin5, but alternative methods exist. Tracking fluorescent inhomogeneities as seen by live cell imaging-compatible probes labeling cellular actin filaments, such as Lifeact25, can also be employed6,26. The advantage here is that the overexpression of &#946;-actin can be avoided, which is capable of increasing cell edge protrusion and migration, and thus potentially interferes&#160;with the specific assay or experimental question (see e.g., Kage et al.26; Peckham et al.27). However, a distinct disadvantage of the Lifeact probe constitutes its rapid on/off kinetics of binding to actin filaments, so that bleaching of actin filament structures labeled by Lifeact in cells&#160;provides information only on the probe turnover, but not the turnover of the actin filaments, to which it binds25. The tracking of fluorescence inhomogeneities employed previously6,26 does provide a practical compromise, much similar to the widely used tracking of fluorescence speckles incorporated into filamentous cytoskeletal structures (see e.g., Salmon and Waterman28), but may not be as straight forward to use and as precise as FRAP of EGFP-tagged F-actin structures. Photoactivation has been applied by us for estimating the rates of monomeric actin incorporation into protruding lamellipodia, as well as its mobility throughout the cytosol, in the context of experimentally tuned cytosolic F-actin levels6. The technique is useful when examining mobility and distribution of proteins derived from relatively large areas, such as cytosolic regions. However, examining the distribution of proteins derived from relatively small photoactivated structures; e.g., growth cones might be challenging due to the low numbers of fluorescent molecules activated, weak signals, and thus lack of sensitivity. Potential alternative techniques to photoactivation or photoconversion of fluorescence (see above) may include inverse FRAP, which relies on photobleaching the entire cell except the ROI, followed by tracking the mobility of fluorescent molecules away from this region. The technique does not require overexpressing photoactivatable versions of proteins, but will always involve exposure to an unusually high dose of laser power, potentially causing undesired side effects such as photodamage.
Clearly, photoactivation and FRAP cannot distinguish whether proteins are moving as monomers, dimers, or even small oligomers, and whether they move in combination with additional binding partners. Information of that kind can be obtained instead from fluorescence correlation spectroscopy techniques29 or, alternatively, FLIM-FRET30. Nonetheless, FRAP and photoactivation constitute straightforward approaches to directly assess local and global protein dynamics in cells, irrespective of the protein of interest, subcellular location, or cell type studied.

Monitoring the spatiotemporal dynamics of proteins and other molecules in living cells has become an essential tool in many fields of cell and molecular biology. Advanced fluorescence microscopy techniques including fluorescence resonance energy transfer (FRET) and FRET-fluorescence lifetime imaging (FRET-FLIM), or FRAP, fluorescence loss in photobleaching (FLIP) and photoactivation as well as many others allow for the temporal and spatial tracking of protein-protein interactions, conformational changes, as well as determining the kinetics of diffusion and localization of different proteins in the cell1,2. FRAP and photoactivation techniques, in particular, are widely applicable for examining the regulators of the actin cytoskeleton and cell migration. These techniques can be applied alone or in combination with additional micromanipulation techniques such as microinjection3, and involve the expression of fluorescently-labeled proteins. They allow the estimation of the kinetics of protein association to actin-rich structures involved in cell migration, such as filopodia or lamellipodia, the turnover of proteins in focal adhesions4, or branched actin networks5. They also enable the determination of lamellipodial actin polymerization rates, the assessment of the dispersion of monomeric actin within the cytosol, the rate of subcellular actin monomer translocation to polymerizing actin filaments in protruding lamellipodia6, and other parameters.
FRAP is a method for visualizing and quantifying the mobility of proteins within a living cell, originally developed in the 1970s by Axelrod7. A region of interest (ROI) within a cell, populated with fluorescently-labeled proteins, is transiently exposed to a laser of high intensity, sufficient to cause bleaching of the fluorophore molecules present in this region during a given short period of time. The unbleached, fluorescently labeled proteins located outside the ROI during bleaching, will diffuse and infiltrate the bleached region depending on their spatiotemporal dynamics, causing the displacement of photobleached molecules over time. The rate of fluorescence recovery in bleached regions is dependent on various factors, including the size and the rate of diffusion of a given molecule, and of course its turnover rate within the putative associated bleached structure. Thus, soluble proteins will mediate the recovery of fluorescence within the bleached ROI rapidly through diffusion, while proteins tightly associated with structures, such as focal adhesions, will have longer turnover times, as their fluorescence recovery will depend both on the diffusion of the soluble fraction of the protein and dissociation-association kinetics of the structure-associated fraction. Fluorescence recovery is usually acquired and quantified until the initial level of pre-bleach intensity of fluorescence is reached. However, this does not occur if a part of the initial fluorescence intensity belongs to the so-called immobile fraction, which is unable to be replenished by diffusion or is replenishing at very slow rates as compared to the majority of molecules comprising the mobile fraction. To determine the rate of protein turnover, FRAP curves are generated, representing the extent of fluorescence recovery over time. From these recovery curves, average half-times of protein recovery can be calculated. By creating curve fits of the average FRAP data, and hence mathematical analyses, it is also possible to deduce whether the average turnover rate of the mobile fraction constitutes a composite of one homogeneous population of molecules, or whether it is composed of two or more subpopulations of molecules turning over at differential rates. In addition to estimating protein turnover rates by quantitative approaches, tracking the recovery of photobleached regions in lamellipodia can also allow for accurate quantification of lamellipodial motility parameters such as retrograde flow, protrusion, and actin polymerization rates. Thus, FRAP constitutes a versatile tool to be applied for assessing various parameters within structures of living cells.
Photoactivation is a method used to track the diffusion and mobility of proteins or molecules originating from a designated cellular location. The technique employs, for instance, a variant of wild-type green fluorescent protein (GFP), initially developed by Patterson and Lippincott-Schwartz8, which is mutated in a manner that allows its fluorescence to be highly increased upon exposure to ultraviolet (UV) light (around 400 nm; here, 405 nm). As described by Patterson et al., wild-type GFP chromophores exist as a mixed population of neutral phenols and anionic phenolates, which produce a major absorbance peak at approximately 397 nm and a minor one at 475 nm, respectively. Upon irradiation of the protein with UV light, the population undergoes photoconversion, shifting towards the anionic form. When excited by 488 nm, the photoconverted/photoactivated protein exhibits a 3-fold increase in fluorescence, insufficient in practice for distinguishing between activated and non-activated GFP due to the high intrinsic background fluorescence. However, a decrease in background intensity has been achieved by introducing a single amino acid mutation into the GFP sequence (histidine substitution at position 203). The resulting T203H mutant, also known as photoactivatable-GFP (PA-GFP) is characterized by a significant reduction in absorbance of the minor peak, which upon irradiation with UV light is increased nearly 100-fold when subsequently excited by 488 nm light. Hence, overexpression of PA-GFP-tagged proteins is a widely used approach, which allows the determination of diffusion and motility of molecules within cells. We have previously applied PA-GFP-tagged actin to determine the rate of dispersion of actin monomers away from cytosolic regions, allowing not only exploration of their mobility within the cytosol, but also their incorporation rate into the protruding lamellipodial actin network6. More recent literature also describes novel, photo-convertible proteins that can in principle be used in an analogous fashion, but harboring the potential advantage to be visible already before photo-conversion. Examples for this group of fluorescent proteins include Dendra2 and mEos29,10,11,12.
In this article, we explain the methodology of microinjecting cells with proteins. We further explain how this technique can be combined with FRAP, by photobleaching proteins involved in actin cytoskeleton regulation and motility, and how FRAP curves and half-time of recovery of mobile fractions can be derived. In addition, we provide an example of how the FRAP technique can be used to determine actin polymerization rates of lamellipodial networks. We also provide instructions and tips on how to perform photoactivation experiments, which can be used to determine cytosolic mobility of monomeric actin and rates of actin incorporation into lamellipodia. These techniques, of course, are not only limited to tracking actin cytoskeleton components, but upon potentially required moderate adaption or optimization, can be widely applied to other cell types or to investigate different proteins, structures, and parameters.

<table><tbody><tr><td>B16-F1 mouse skin melanoma cells&amp;nbsp;</td><td>American Type Culture Collection, Manassas, VA</td><td>CRL-6323</td><td></td></tr><tr><td>NIH-3T3 cells</td><td>American Type Culture Collection, Manassas, VA</td><td>CRL-1658</td><td></td></tr><tr><td>DMEM 4.5g/L glucose&amp;nbsp;</td><td>Life Technologies, Thermno Fisher Scientific, Germany</td><td>41965-039</td><td></td></tr><tr><td>Ham&amp;rsquo;s F-12 medium</td><td>Sigma-Aldrich</td><td>N8641</td><td></td></tr><tr><td>Fetal calf serum&amp;nbsp; (FCS)</td><td>PAA Laboratories, Linz, Austria</td><td>A15-102</td><td></td></tr><tr><td>Fetal bovine serum (FBS)</td><td>Sigma-Aldrich, Germany</td><td>F7524</td><td>Lot054M3396</td></tr><tr><td>MEM Non essential amino acids&amp;nbsp;</td><td>Gibco, ThermoFisher Scientific, Germany</td><td>11140035</td><td></td></tr><tr><td>L-Glumatine 200mM (100x)</td><td>Life Technolgies</td><td>25030-024</td><td></td></tr><tr><td>Pen-Strep 5000 U/mL</td><td>Life technologies</td><td>15070063</td><td></td></tr><tr><td>Sodium Pyruvate (100 mM)</td><td>Gibco, ThermoFisher Scientific, Germany</td><td>11360-039</td><td></td></tr><tr><td>Laminin</td><td>Sigma-Aldrich</td><td>L-2020</td><td></td></tr><tr><td>Laminin coating buffer&amp;nbsp;</td><td></td><td></td><td>Self-made: 50mM Tris ph7.4, 150mM NaCl</td></tr><tr><td>Fibronectin from human plasma&amp;nbsp;</td><td>Roche Diagnostics, Mannheim, Germany</td><td>11 051 407 001</td><td></td></tr><tr><td>Jetpei</td><td>Polyplus Transfection, Illkirch, France</td><td>101-10N</td><td></td></tr><tr><td>JetPei buffer</td><td>Polyplus Transfection, Illkirch, France</td><td>702-50</td><td>150mM NaCl</td></tr><tr><td>PA-GFP-actin plasmid DNA&amp;nbsp;</td><td></td><td></td><td>described in Koestler et al.2008</td></tr><tr><td>pEGFP-actin plasmid DNA</td><td>Clontech, Mountain View, CA, USA</td><td></td><td></td></tr><tr><td>Rac1 protein for microinjection</td><td></td><td></td><td>Purified as GST-tagged version, and cleaved from GST prior to injection</td></tr><tr><td>Microinjection buffer</td><td></td><td></td><td>Self-made: 100mM NaCl, 50mM Tris-HCl ph7.5, 5mM MgCl&lt;sub&gt;2&lt;/sub&gt;, 1mM DTT</td></tr><tr><td>Dextran, Texas Red, 70,000 MW, Lysine Fixable</td><td>Molecular Probes, Thermno Fisher Scientific, Germany</td><td>D1864</td><td></td></tr><tr><td>Microscope circular cover glasses 15mm, No.1&amp;nbsp;</td><td>Karl Hecht, Aisstent, Sondheim, Germany</td><td>1001/15</td><td></td></tr><tr><td>Eppendorf Femtotips Microloader Tips</td><td>Eppendorf, Hamburg, Germany</td><td>5242 956 003</td><td></td></tr><tr><td>Eppendorf Femtotip Microinjection Capillary Tips</td><td>Eppendorf, Hamburg, Germany</td><td>930000035</td><td></td></tr><tr><td>Silicone Grease&amp;nbsp;</td><td>ACC Silicones, Bridgewater, England</td><td>SGM494</td><td></td></tr><tr><td>Aluminium Open Diamond Bath Imaging Chamber</td><td>Warner instruments</td><td>RC-26</td><td></td></tr><tr><td>Automatic temperature controller</td><td>Warner Instruments</td><td>TC-324B</td><td></td></tr><tr><td>Microscope: Axio Observer&amp;nbsp;</td><td>Carl Zeiss, Jena, Germany</td><td></td><td></td></tr><tr><td>CoolSnap-HQ2 camera&amp;nbsp;</td><td>Photometrics, Tucson, AZ</td><td></td><td></td></tr><tr><td>Lambda DG4 light source&amp;nbsp;</td><td>Sutter Instrucment, Novato, CA</td><td></td><td></td></tr><tr><td>Laser source&amp;nbsp;</td><td>Visitron Systems</td><td></td><td></td></tr><tr><td>Eppendorf FemtoJet microinjector&amp;nbsp;</td><td>Eppendorf, Hamburg, Germany</td><td></td><td>With built-in compressor for pressure supply</td></tr><tr><td>Nikon Narishige Micromanipulator system</td><td>Nikon Instruments, Japan</td><td></td><td></td></tr><tr><td>Visiview software v2.1.4</td><td>Visitron Systems, Puchheim, Germany</td><td></td><td></td></tr><tr><td>Metamorph software v7.8.10</td><td>Molecular Devices, Sunnyvale, CA</td><td></td><td></td></tr><tr><td>Sigma Plot v.12</td><td>Systat Software Inc.</td><td></td><td></td></tr></tbody></table>

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.

Figure 1g, h show&#160;phase contrast images of an NIH3T3 fibroblast cell prior and 10 min post-microinjection of Rac1, which is a small Rho-family GTPase capable of inducing lamellipodia formation through its interaction with the WAVE complex. The cell is first visualized before the microinjection (Figure 1g), to confirm its viability and morphology, e.g., lack of lamellipodia. At 10 min post-microinjection, the cell has clearly changed its morphology, which is expected from this treatment, and&#160;indicates a successful injection (Figure 1h).
For simplicity and clarity, we next provide exemplary results for FRAP and photoactivation analysis in cells, which have not been additionally microinjected.
Analysis of the turnover of EGFP-tagged VASP at the lamellipodium tip is shown in Figure 2a-f. Note that VASP in addition targets to nascent and focal adhesions, small and elongated dots in the cell interior18,19. The fluorescence intensity of a lamellipodial region with a clear VASP accumulation at the tip was bleached and measured for each time frame, by following the contour of the ROI before, during, and after bleaching as the lamellipodium protrudes forwards. As bleached EGFP-VASP proteins are being recycled by non-bleached molecules at these sites, gradual recovery of fluorescence is observed (Figure 2b). The FRAP recovery curve obtained in this fashion and normalized to the pre-bleach intensity (expressed as 1) can be seen in Figure 2c. Photobleaching efficiency can vary and was approximately 20% of the value before bleaching in this example, as determined from the value at t0 (the first frame after photobleaching). The increase of fluorescence reaches a plateau in the example shown at roughly 80% of the fluorescence before bleaching. In a static structure during the time course of the experiment, such as a focal adhesion, the difference between the pre-bleach intensity and the plateau fluorescence reached after recovery is defined as the immobile fraction (IF, red arrow in Figure 2c, e), whereas the amount of fluorescence recovered between the time of bleaching and full recovery is defined as the mobile fraction (green double-headed arrow in Figure 2c, e). Note that in a dynamically changing structure such as the lamellipodium tip analyzed here, the extent of the IF might not only represent immobile molecules, but also derive from a reduction of protrusion speed, as EGFP-VASP intensity is known to depend on this parameter18. To calculate the half-time of recovery, a fit curve was created on Sigma plot (Figure 2d). In this case, the value of the &#34;b&#34; parameter extracted from solving Equation 2 is equal to 0.0754, which when applied to the logarithmic function (Equation 4) results in an estimated half-time of recovery&#160;of 9.19 s (Figure 2d, far right panel), which is relatively fast in this particular cell as compared to the average published previously5. It must be noted that recovery half-times&#160;may sometimes vary significantly from cell to cell within the same population. Therefore, for obtaining representative results, we recommend determining this parameter as an average from at least 15-20 cells. To illustrate the degree of variance, arithmetic means of EGFP-VASP recovery averaged from 15 cells for each time-point were generated (Figure 2e), and average curve fits created and displayed in an analogous fashion (Figure 2f).
The polymerization rate of the lamellipodial actin network comprises the sum of forward network protrusion and retrograde flow. FRAP can be applied for measuring the actin polymerization rate by transfecting cells (in this case B16-F1) with EGFP-tagged &#946;-actin and photobleaching a protruding lamellipodial region (Figure 2g). For analysis of lamellipodial actin network polymerization, the fluorescence recovery upon bleaching of EGFP-tagged &#946;-actin is assessed over time. As the polymerization of actin monomers progresses at the barbed ends of lamellipodial actin filaments (which all point towards the front20), the network is constantly translocated rearwards and progressing forwards, the rates of which can be easily obtained through polarized recovery of fluorescence upon photobleaching. Fluorescence recovery of the lamellipodium is complete as soon as the bleached zone has reached the transition zone between the rear part of the lamellipodium and the lamella, which is characterized by a lower density of more horizontally-arranged filament bundles turning over much more slowly than what is observed in the lamellipodium. As illustrated in Figure 2g, fluorescence recovery can be visualized as a line horizontal to the edge and flowing backwards towards the lamella, which allows measuring the distances of protrusion and retrograde flow (individually represented in the far right panel of Figure 2g as orange and red double-headed arrows, respectively).
We have also applied photoactivation in B16-F1 cells transfected with PA-GFP-actin to track the mobility of actin monomers within the cytosol and the rate of their incorporation within protruding lamellipodia. As illustrated in Figure 3a, b, a cytosolic region was photoactivated by exposure to a 405 nm laser, while images were acquired on the GFP channel every 1.5 s for visualizing the distribution of GFP-tagged, photoactivated actin. Photoactivated GFP-actin can be seen diffusing out of the cytosolic region in Figure 3b. The rate of fluorescence intensity decrease in the photoactivated cytosolic region is represented as the percentage of the initial intensity at t0 (first frame after photoactivation; Figure 3c). Photoactivated actin also integrates at the tips of lamellipodia, where new actin monomers are added to the growing barbed ends of elongating actin filaments during protrusion. To estimate the rate of lamellipodial incorporation, we measured the fluorescence intensity over time of a two-dimensional contour/region of approximately 5 &#181;m in width and 1 &#181;m in height; the region was constantly re-positioned at the tip of the lamellipodium as it protruded. Actin incorporation was represented as the percentage of fluorescence intensity of the photoactivated cytosolic region at t0 (Figure 3d). As elongation of actin filaments progressed, new actin monomers were incorporated at the lamellipodial front. A fraction of these actin monomers was stochastically derived from the cytosolic pool where monomers were photoactivated. This results in the rapid increase of fluorescence in lamellipodia in the first 20 s after photoactivation. As new monomers are being added to the lamellipodial front, previously incorporated actin monomers flow with filaments towards the lamella by retrograde flow. Over time, the ROI is completely filled with fluorescent monomers and a plateau in fluorescence is reached (Figure 3d). A gradual drop in fluorescence is then observed when, following diffusion of photoactivated monomers throughout the cell, non-photoactivated actin monomers are increasingly being re-added to the lamellipodial front. This decrease in fluorescence will find a new plateau, which will be reached as soon as a balance in the entire cell between photoactivated and non-photoactivated monomers is reached (data not shown).
The mobility of actin monomers throughout the cytosol was derived by measuring fluorescence intensities in regions of equal size positioned distally from the photoactivated region (exemplified on Figure 3a by color coded regions labeled R1-R5). As illustrated in Figure 3e, fluorescence intensity in each of these regions is gradually decreasing away from the cytosolically photoactivated region, as the fraction of photoactivated actin monomers becomes increasingly diluted with non-activated (i.e., non-fluorescent) monomers. Furthermore, the peak of fluorescence is reached later: the more distant the measured region is located from the photoactivated region, the longer the time that is required for actin monomers to diffuse into these regions. A representative value for the degree of actin monomer infiltration into each region can be derived by quantifying the half time of reaching the fluorescence plateau. The more distant the&#160;region,&#160;the longer it takes for the photoactivated actin to diffuse&#160;into it, and thus more time is required for the fluorescent plateau to be reached, ultimately leading to a higher t1/2 value (Figure 3e).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/57643/57643fig1.jpg" /> 
Figure 1: Imaging chamber assembly and microinjection procedure. (a) Imaging chamber components. (b) Silicone grease is carefully smeared around the opening of a plastic sealer. (c) The coverslip is positioned with the cell-side facing up into the center of the imaging chamber opening. (d) A secure seal is established by positioning the plastic sealer on top of the coverslip and by tightening the side clamps. (e) Microscopy medium is pipetted into the chamber slot. (f) The imaging chamber is positioned on the microscope stage, heat detector and electrodes are linked to a heating unit pre-set to 37 &#176;C, and cells are allowed to adapt for at least 30 min before microscopy is initiated. In this example, the microscope stage is also equipped with a micromanipulator for performing microinjections, and the microinjection needle is dipped into the medium covering the cell layer in the imaging chamber. (g) An NIH3T3 fibroblast cell is visualized before microinjection by phase-contrast microscopy. The red cross in the perinuclear compartment indicates the location of the future microinjection, which corresponds to a high cytoplasmic region due to the close proximity to the bulky nucleus. (h) 10 min following microinjection with Rac1, the cell reacts by prominent formation of lamellipodia around the entire cell periphery (indicated by green arrows). <a href="/files/ftp_upload/57643/57643fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/57643/57643fig2.jpg" /> 
Figure 2: FRAP allows determining rates of protein turnover or lamellipodial actin polymerization. (a) Representative example of B16-F1 cell expressing EGFP-VASP before photobleaching of a lamellipodial region as indicated. Differently colored contours/shapes are labeled to indicate which regions were considered for fluorescence intensity measurements over time. Note the red contour marked with an exclamation mark, which labels a cytosolic region positioned in an area containing multiple vesicles and cell surface ruffles. Dynamic areas like this should be avoided for selecting regions of fluorescence reference, as they are characterized by strong short-term fluctuations of fluorescence, potentially causing inaccurate results. (b) Lamellipodial region of the EGFP-VASP expressing cell before and after photobleaching. The recovery of fluorescent signal after photobleaching within the region marked in purple is visualized over time. Arrow indicates the tip of a microspike, enriched for VASP likely due to the high density of actin filaments polymerizing there19. (c) An example of a FRAP recovery curve as derived from quantifying the fluorescent intensity of the photobleached lamellipodium (purple contour) in b. Red and green lines on the right indicate, respectively, immobile and mobile fractions. (d) A fit of the FRAP recovery curve in c (left panel) and an example of the calculation method used to derive the recovery half time&#160;(right panel). (e) An example of a FRAP recovery curve derived from averaging the fluorescence recovery curves of 15 cells, with SEM bars indicating the degree of variability within the sample population. (f) A curve fit derived from averaging the FRAP recovery curve fits of 15 cells (left panel) and an example of the calculation method used to derive the recovery half time&#160;(right panel). (g) Time-lapse panels of protruding lamellipodium of a B16-F1 cell expressing EGFP-tagged &#946;-actin before and after bleaching of a lamellipodial region as indicated, followed by fluorescence recovery in the lamellipodium over time. On the far right panel, values measured for protrusion and retrograde distances are provided (in orange and red, respectively). Calculations under the image panels reveal how the sum of protrusion and retrograde distances are used to derive polymerization rate of the lamellipodial actin network. <a href="/files/ftp_upload/57643/57643fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/57643/57643fig3.jpg" /> 
Figure 3: Photoactivation of PA-GFP-actin for monomer tracking throughout the cell. (a) A representative example of a B16-F1 cell expressing PA-GFP-actin before triggering photoactivation in a cytosolic region as indicated by the red circle (PA). Differently colored contours are labeled to indicate which regions were considered for fluorescence intensity measurements over time. (b) An illustration of the temporal distribution of PA-GFP-actin following photoactivation. Note the gradual reduction of fluorescence in the photoactivated, cytosolic region (red circle), as the photoactivated actin diffuses away from it. Due to their diffusion to the front and assembly into the network, photoactivated actin monomers are gradually enhanced in&#160;lamellipodia (cyan region) and throughout the cytosol (different color-coded regions) in a distance- and time-dependent fashion. (c) Representative, temporal decline of fluorescence within the photoactivated cytosolic region (red contour in b). (d) Temporal changes in fluorescence intensity in the lamellipodial region (cyan contour in b). (e) Curves representative of the temporal changes in fluorescence intensity of cytosolic regions (color-coded in b) due to positioning in variable distances from the area of photoactivation. Note how half-times of&#160;reaching the fluorescence plateau (indicated in legend on the right) increase with the distance of given region to the area of photoactivation, likely correlating with the increased times needed for diffusion of actin monomers into the respective region. <a href="/files/ftp_upload/57643/57643fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Micromanipulation Techniques Allowing Analysis of Morphogenetic Dynamics and Turnover of Cytoskeletal Regulators at JoVE.com

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

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.

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how ...

micromanipulation-techniques-allowing-analysis-morphogenetic-dynamics

Nanyang Technological University

Research

JoVE Journal

Biology

10.0K Views.  Technische Universität Braunschweig. 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.

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

Spatio-Temporal Manipulation of Small GTPase Activity at Subcellular Level and on Timescale of Seconds in Living Cells

Dynamic regulation of the Rho family of small guanosine triphosphatases (GTPases) with great spatiotemporal precision is essential for various cellular functions and events1, 2. Their spatiotemporally dynamic nature has been revealed by visualization of their activity and localization in real time3. In order to gain deeper understanding of their roles in diverse cellular functions at the molecular level, the next step should be perturbation of protein activities at a precise subcellular location and timing. 
To achieve this goal, we have developed a method for light-induced, spatio-temporally controlled activation of small GTPases by combining two techniques: (1) rapamycin-induced FKBP-FRB heterodimerization and (2) a photo-caging method of rapamycin. With the use of rapamycin-mediated FKBP-FRB heterodimerization, we have developed a method for rapidly inducible activation or inactivation of small GTPases including Rac4, Cdc424, RhoA4 and Ras5, in which rapamycin induces translocation of FKBP-fused GTPases, or their activators, to the plasma membrane where FRB is anchored. For coupling with this heterodimerization system, we have also developed a photo-caging system of rapamycin analogs. A photo-caged compound is a small molecule whose activity is suppressed with a photocleavable protecting group known as a caging group. To suppress heterodimerization activity completely, we designed a caged rapamycin that is tethered to a macromolecule such that the resulting large complex cannot cross the plasma membrane, leading to virtually no background activity as a chemical dimerizer inside cells6. Figure 1 illustrates a scheme of our system. With the combination of these two systems, we locally recruited a Rac activator to the plasma membrane on a timescale of seconds and achieved light-induced Rac activation at the subcellular level6.

A method for spatio-temporal control of small GTPase activity by light is described. This method is based on rapamycin-induced FKBP-FRB heterodimerization and photo-caging systems. Optimization of light-irradiation enables the spatio-temporally controlled activation of small GTPases at the subcellular level.

Dynamic regulation of the Rho family of small guanosine triphosphatases (GTPases) with great spatiotemporal precision is essential for various cellular ...

Bioengineering

An Optogenetic Method to Control and Analyze Gene Expression Patterns in Cell-to-cell Interactions

Cells should respond properly to temporally changing environments, which are influenced by various factors from surrounding cells. The Notch signaling pathway is one of such essential molecular machinery for cell-to-cell communications, which plays key roles in normal development of embryos. This pathway involves a cell-to-cell transfer of oscillatory information with ultradian rhythms, but despite the progress in molecular biology techniques, it has been challenging to elucidate the impact of multicellular interactions on oscillatory gene dynamics. Here, we present a protocol that permits optogenetic control and live monitoring of gene expression patterns in a precise temporal manner. This method successfully revealed that intracellular and intercellular periodic inputs of Notch signaling entrain intrinsic oscillations by frequency tuning and phase shifting at the single-cell resolution. This approach is applicable to the analysis of the dynamic features of various signaling pathways, providing a unique platform to test a functional significance of dynamic gene expression programs in multicellular systems.

Here, we present a protocol to analyze cell-to-cell transfer of oscillatory information by optogenetic control and live monitoring of gene expression. This approach provides a unique platform to test a functional significance of dynamic gene expression programs in multicellular systems.

Cells should respond properly to temporally changing environments, which are influenced by various factors from surrounding cells. The Notch signaling ...

Genetics

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

Drosophila immature eggs are called egg chambers, and their structure resembles primitive organs that undergo morphological changes from a round to an ellipsoid shape during development. This developmental process is called oogenesis and is crucial to generating functional mature eggs to secure the next fly generation. For these reasons, egg chambers have served as an ideal and relevant model to understand animal organ development.
Several in vitro culturing protocols have been developed, but there are several disadvantages to these protocols. One involves the application of various covers that exert an artificial pressure on the imaged egg chambers in order to immobilize them and to increase the imaged acquisition plane of the circumferential surface of the analyzed egg chambers. Such an approach may negatively influence the behavior of the thin actomyosin machinery that generates the power to rotate egg chambers around their longer axis.
Thus, to overcome this limitation, we culture Drosophila egg chambers freely in the media in order to reliably analyze actomyosin machinery along the circumference of egg chambers. In the first part of the protocol, we provide a manual detailing how to analyze the actomyosin machinery in a limited acquisition plane at the local cellular scale (up to 15 cells). In the second part of the protocol, we provide users with a new Fiji-based plugin that allows the simple extraction of a defined thin layer of the egg chambers&#8217; circumferential surface. The following protocol then describes how to analyze actomyosin signals at the tissue scale (&#62;50 cells). Finally, we pinpoint the limitations of these approaches at both the local cellular and tissue scales and discuss its potential future development and possible applications.

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

This protocol provides a Fiji-based, user-friendly methodology along with straightforward instructions explaining how to reliably analyze actomyosin behavior in individual cells and curved epithelial tissues. No programming skills are required to follow the tutorial; all steps are performed in a semi-interactive manner using the graphical user interface of Fiji and associated plugins.

Drosophila immature eggs are called egg chambers, and their structure resembles primitive organs that undergo morphological changes from a round to an ...

Developmental Biology

A Microfluidics Approach for the Functional Investigation of Signaling Oscillations Governing Somitogenesis

Periodic segmentation of the presomitic mesoderm of a developing mouse embryo is controlled by a network of signaling pathways. Signaling oscillations and gradients are thought to control the timing and spacing of segment formation, respectively. While the involved signaling pathways have been studied extensively over the last decades, direct evidence for the function of signaling oscillations in controlling somitogenesis has been lacking. To enable the functional investigation of signaling dynamics, microfluidics is a previously established tool for the subtle modulation of these dynamics. With this microfluidics-based entrainment approach endogenous signaling oscillations are synchronized by pulses of pathway modulators. This enables modulation of, for instance, the oscillation period or the phase-relationship between two oscillating pathways. Furthermore, spatial gradients of pathway modulators can be established along the tissue to study how specific changes in the signaling gradients affect somitogenesis.
The present protocol is meant to help establish microfluidic approaches for the first-time users of microfluidics. The basic principles and equipment needed to set up a microfluidic system are described, and a chip design is provided, with which a mold for chip generation can conveniently be prepared using a 3D printer. Finally, how to culture primary mouse tissue on a microfluidic chip and how to entrain signaling oscillations to external pulses of pathway modulators are discussed.
This microfluidic system can also be adapted to harbor other in vivo and in vitro model systems such as gastruloids and organoids for functional investigation of signaling dynamics and morphogen gradients in other contexts.

A protocol for the culture and manipulation of mouse embryonic tissue on a microfluidic chip is provided. By applying pulses of pathway modulators, this system can be used to externally control signaling oscillations for the functional investigation of mouse somitogenesis.

Periodic segmentation of the presomitic mesoderm of a developing mouse embryo is controlled by a network of signaling pathways. Signaling oscillations and ...

Biophysical Assays to Probe the Mechanical Properties of the Interphase Cell Nucleus: Substrate Strain Application and Microneedle Manipulation

In most eukaryotic cells, the nucleus is the largest organelle and is typically 2 to 10 times stiffer than the surrounding cytoskeleton; consequently, the physical properties of the nucleus contribute significantly to the overall biomechanical behavior of cells under physiological and pathological conditions. For example, in migrating neutrophils and invading cancer cells, nuclear stiffness can pose a major obstacle during extravasation or passage through narrow spaces within tissues.1 On the other hand, the nucleus of cells in mechanically active tissue such as muscle requires sufficient structural support to withstand repetitive mechanical stress. Importantly, the nucleus is tightly integrated into the cellular architecture; it is physically connected to the surrounding cytoskeleton, which is a critical requirement for the intracellular movement and positioning of the nucleus, for example, in polarized cells, synaptic nuclei at neuromuscular junctions, or in migrating cells.2 Not surprisingly, mutations in nuclear envelope proteins such as lamins and nesprins, which play a critical role in determining nuclear stiffness and nucleo-cytoskeletal coupling, have been shown recently to result in a number of human diseases, including Emery-Dreifuss muscular dystrophy, limb-girdle muscular dystrophy, and dilated cardiomyopathy.3 To investigate the biophysical function of diverse nuclear envelope proteins and the effect of specific mutations, we have developed experimental methods to study the physical properties of the nucleus in single, living cells subjected to global or localized mechanical perturbation. Measuring induced nuclear deformations in response to precisely applied substrate strain application yields important information on the deformability of the nucleus and allows quantitative comparison between different mutations or cell lines deficient for specific nuclear envelope proteins. Localized cytoskeletal strain application with a microneedle is used to complement this assay and can yield additional information on intracellular force transmission between the nucleus and the cytoskeleton. Studying nuclear mechanics in intact living cells preserves the normal intracellular architecture and avoids potential artifacts that can arise when working with isolated nuclei. Furthermore, substrate strain application presents a good model for the physiological stress experienced by cells in muscle or other tissues (e.g., vascular smooth muscle cells exposed to vessel strain). Lastly, while these tools have been developed primarily to study nuclear mechanics, they can also be applied to investigate the function of cytoskeletal proteins and mechanotransduction signaling.

We present two independent, microscope-based tools to measure the induced nuclear and cytoskeletal deformations in single, living adherent cells in response to global or localized strain application. These techniques are used to determine nuclear stiffness (i.e., deformability) and to probe intracellular force transmission between the nucleus and the cytoskeleton.

In most eukaryotic cells, the nucleus is the largest organelle and is typically 2 to 10 times stiffer than the surrounding cytoskeleton; consequently, the ...

Quantifying Cytoskeleton Dynamics Using Differential Dynamic Microscopy

Cells can crawl, self-heal, and tune their stiffness due to their remarkably dynamic cytoskeleton. As such, reconstituting networks of cytoskeletal biopolymers may lead to a host of active and adaptable materials. However, engineering such materials with precisely tuned properties requires measuring how the dynamics depend on the network composition and synthesis methods. Quantifying such dynamics is challenged by variations across the time, space, and formulation space of composite networks. The protocol here describes how the Fourier analysis technique, differential dynamic microscopy (DDM), can quantify the dynamics of biopolymer networks and is particularly well suited for studies of cytoskeleton networks. DDM works on time sequences of images acquired using a range of microscopy modalities, including laser-scanning confocal, widefield fluorescence, and brightfield imaging. From such image sequences, one can extract characteristic decorrelation times of density fluctuations across a span of wave vectors. A user-friendly, open-source Python package to perform DDM analysis is also developed. With this package, one can measure the dynamics of labeled cytoskeleton components or of embedded tracer particles, as demonstrated here with data of intermediate filament (vimentin) networks and active actin-microtubule networks. Users with no prior programming or image processing experience will be able to perform DDM using this software package and associated documentation.

Differential dynamic microscopy (DDM) combines features of dynamic light scattering and microscopy. Here, the process of using DDM to characterize reconstituted cytoskeleton networks by quantifying the subdiffusive and caged dynamics of particles in vimentin networks and the ballistic motion of active myosin-driven actin-microtubule composites is presented.

Cells can crawl, self-heal, and tune their stiffness due to their remarkably dynamic cytoskeleton. As such, reconstituting networks of cytoskeletal ...

3D Particle Tracking for Noninvasive In Vivo Analysis of Synaptic Microtubule Dynamics in Dendrites and Neuromuscular Junctions of Drosophila

Microtubules (MTs) play critical roles in neuronal development, but many questions remain about the molecular mechanisms of their regulation and function. Furthermore, despite progress in understanding postsynaptic MTs, much less is known about the contributions of presynaptic MTs to neuronal morphogenesis. In particular, studies of in vivo MT dynamics in Drosophila sensory dendrites yielded significant insights into polymer-level behavior. However, the technical and analytical challenges associated with live imaging of the fly neuromuscular junction (NMJ) have limited comparable studies of presynaptic MT dynamics. Moreover, while there are many highly effective software strategies for automated analysis of MT dynamics in vitro and ex vivo, in vivo data often necessitate significant operator input or entirely manual analysis due to inherently inferior signal-to-noise ratio in images and complex cellular morphology. &#160;To address this, this study optimized a new software platform for automated and unbiased in vivo particle detection. Multiparametric analysis of live time-lapse confocal images of EB1-GFP labeled MTs was performed in both dendrites and the NMJ of Drosophila larvae and found striking differences in MT behaviors. MT dynamics were furthermore analyzed following knockdown of the MT-associated protein (MAP) dTACC, a key regulator of Drosophila synapse development, and identified statistically significant changes in MT dynamics compared to wild type. These results demonstrate that this novel strategy for the automated multiparametric analysis of both pre- and postsynaptic MT dynamics at the polymer-level significantly reduces human-in-the-loop criteria. The study furthermore shows the utility of this method in detecting distinct MT behaviors upon dTACC-knockdown, indicating a possible future application for functional screens of factors that regulate MT dynamics in vivo. Future applications of this method may also focus on elucidating cell type and/or compartment-specific MT behaviors, and multicolor correlative imaging of EB1-GFP with other cellular and subcellular markers of interest.&#160;

This study presents a noninvasive intravital neuronal imaging strategy combined with a new software strategy to achieve automated, unbiased tracking and analysis of in vivo microtubule (MT) plus-end dynamics in the sensory dendrites and the neuromuscular junctions of Drosophila.

Microtubules (MTs) play critical roles in neuronal development, but many questions remain about the molecular mechanisms of their regulation and function. ...

Neuroscience

Tracking and Quantifying Developmental Processes in C. elegans Using Open-source Tools

Quantitatively capturing developmental processes is crucial to derive mechanistic models and key to identify and describe mutant phenotypes. Here protocols are presented for preparing embryos and adult C. elegans animals for short- and long-term time-lapse microscopy and methods for tracking and quantification of developmental processes. The methods presented are all based on C. elegans strains available from the Caenorhabditis Genetics Center and on open-source software that can be easily implemented in any laboratory independently of the microscopy system used. A reconstruction of a 3D cell-shape model using the modelling software IMOD, manual tracking of fluorescently-labeled subcellular structures using the multi-purpose image analysis program Endrov, and an analysis of cortical contractile flow using PIVlab (Time-Resolved Digital Particle Image Velocimetry Tool for MATLAB) are shown. It is discussed how these methods can also be deployed to quantitatively capture other developmental processes in different models, e.g., cell tracking and lineage tracing, tracking of vesicle flow.

Tracking and Quantifying Developmental Processes in C. elegans Using Open-source Tools

Here it is shown how to track and quantify developmental processes in C. elegans. The methods presented are based on open-source tools that can be easily implemented. It is demonstrated how to reconstruct 3D cell-shape models, how to manually track subcellular structures, and how to analyze cortical contractile flow.

Quantitatively capturing developmental processes is crucial to derive mechanistic models and key to identify and describe mutant phenotypes. Here ...

High-resolution Imaging and Analysis of Individual Astral Microtubule Dynamics in Budding Yeast

Dynamic microtubules are fundamental to many cellular processes, and accurate measurements of microtubule dynamics can provide insight into how cells regulate these processes and how genetic mutations impact regulation. The quantification of microtubule dynamics in metazoan models has a number of associated challenges, including a high microtubule density and limitations on genetic manipulations. In contrast, the budding yeast model offers advantages that overcome these challenges. This protocol describes a method to measure the dynamics of single microtubules in living yeast cells. Cells expressing fluorescently tagged tubulin are adhered to assembled slide chambers, allowing for stable time-lapse image acquisition. A detailed guide for high-speed, four-dimensional image acquisition is also provided, as well as a protocol for quantifying the properties of dynamic microtubules in confocal image stacks. This method, combined with conventional yeast genetics, provides an approach that is uniquely suited for quantitatively assessing the effects of microtubule regulators or mutations that alter the activity of tubulin subunits.

Budding yeast is an advantageous model for studying microtubule dynamics in vivo due to its powerful genetics and the simplicity of its microtubule cytoskeleton. The following protocol describes how to transform and culture yeast cells, acquire confocal microscopy images, and quantitatively analyze microtubule dynamics in living yeast cells.

Dynamic microtubules are fundamental to many cellular processes, and accurate measurements of microtubule dynamics can provide insight into how cells ...

Measurement of Microtubule Dynamics by Spinning Disk Microscopy in Monopolar Mitotic Spindles

We describe a modification of an established method to determine microtubule dynamics in living cells. The protocol is based on the expression of a genetically encoded marker for the positive ends of microtubules (EB3 labelled with tdTomato fluorescent protein) and high-speed, high-resolution, live-cell imaging using spinning disk confocal microscopy. Cell cycle synchronization and increased density of microtubules are achieved by inhibiting centrosomal separation in mitotic cells, and analysis of growth is performed using open-source U-Track software. The use of a bright and red-shifted fluorescent protein, in combination with the lower laser power and reduced exposure time required for spinning disk microscopy reduce phototoxicity and the probability of light-induced artifacts. This allows for imaging a larger number of cells in the same preparation while maintaining the cells in a growth medium under standard culture conditions. Because the analysis is performed in a supervised automatic fashion, the results are statistically robust and reproducible.

Here we present a robust and detailed method of microtubule dynamics analysis in cells synchronized in prometaphase using live-cell spinning disk confocal microscopy and MATLAB-based image processing.

We describe a modification of an established method to determine microtubule dynamics in living cells. The protocol is based on the expression of a ...

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

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Spatio-Temporal Manipulation of Small GTPase Activity at Subcellular Level and on Timescale of Seconds in Living Cells

An Optogenetic Method to Control and Analyze Gene Expression Patterns in Cell-to-cell Interactions

Analysis of Actomyosin Dynamics at Local Cellular and Tissue Scales Using Time-lapse Movies of Cultured Drosophila Egg Chambers

A Microfluidics Approach for the Functional Investigation of Signaling Oscillations Governing Somitogenesis

Biophysical Assays to Probe the Mechanical Properties of the Interphase Cell Nucleus: Substrate Strain Application and Microneedle Manipulation

Quantifying Cytoskeleton Dynamics Using Differential Dynamic Microscopy

3D Particle Tracking for Noninvasive In Vivo Analysis of Synaptic Microtubule Dynamics in Dendrites and Neuromuscular Junctions of Drosophila

Tracking and Quantifying Developmental Processes in C. elegans Using Open-source Tools

High-resolution Imaging and Analysis of Individual Astral Microtubule Dynamics in Budding Yeast

Measurement of Microtubule Dynamics by Spinning Disk Microscopy in Monopolar Mitotic Spindles