This work has received funding from the European Research Council (ERC) under the European Union&#39;s Horizon 2020 Research and Innovation Programme (grant agreement No 866127, project CAPTUR3D).

1. Sample preparation
<ol>
	<li>Before the microscopy experiment, subculture cells in dishes suited for microscopy applications.
	<ol>
		<li>Wash a 10 cm tissue-culture-treated dish of confluent HeLa or INS 1E (insulinoma &#946;-cell-like) cells twice with 0.01 M 1x PBS, add 1 mL of 0.05% trypsin-EDTA (1x), and place it in a 37 &#176;C, humidified, 5% CO2 incubator for 5 min.</li>
		<li>Resuspend the detached cells by adding 9 mL of complete DMEM (for HeLa cells) or RPMI 1640 (for INS-1E cells) medium, and collect the final 10 mL in centrifuge tubes.</li>
		<li>Seed approximately 2 &#215; 105 cells in each 35 mm x 10 mm dish in a final volume of 1 mL of the medium. Incubate the cells for 24 h at 37 &#176;C and 5% CO2.</li>
	</ol>
	</li>
	<li>To fluorescently label lysosomes, use LysoTracker Red DND-99.
	<ol>
		<li>Dilute the stock solution in 1 mL of prewarmed medium to a final dye concentration of 70 nM.</li>
		<li>Replace the medium from the dish with fresh LysoTracker-containing medium. Incubate the cells in LysoTracker-containing medium for 20 min at 37 &#176;C in a 5% CO2 atmosphere, and wash them twice with fresh medium before the experiment.</li>
		<li>To fluorescently label macropinosomes, use 70-kDa fluorescein isothiocyanate-dextran. Wash the subcultured cells three times with 0.01 M 1x phosphate-buffered saline (PBS), replace with dextran-containing medium (1 mg/mL), and incubate at 37 &#176;C for 30 min. Before proceeding with the microscopy experiment, wash the cells three times with fresh medium.</li>
		<li>To fluorescently label ISGs in INS-1E cells, transfect the cells using transfection reagent (see the Table of Materials) and C-peptide-enhanced green fluorescent protein (EGFP) plasmid13 according to the manufacturer&#39;s protocol, and incubate at 37 &#176;C for 24 h in a 5% CO2 atmosphere before the experiment.</li>
	</ol>
	</li>
</ol>
2. Data Acquisition
<ol>
	<li>To let the microscope equilibrate at the desired temperature and atmosphere, turn on the microscope incubator-controlling system at least 2 h before the experiment. 
	NOTE: Each acquisition is a time-lapse series.</li>
	<li>Acquire the images using an inverted confocal microscope equipped with a 60x, 1.2 Numerical Aperture (NA) water-immersion objective.</li>
	<li>Use a 488 nm Argon laser for excitation of EGFP (transfected cells) and fluorescein-labeled macropinosomes. Collect the fluorescence emission between 500 and 600 nm using a standard photomultiplier tube detector.</li>
	<li>Use a 543 nm HeNe laser to excite Lysotracker and collect its fluorescence emission between 555 and 655 nm.</li>
	<li>Set the diameter of the detection pinhole to the size of 1 Airy. For each acquisition, collect a series of 1000 sequential frames. Set the pixel-dwell time to 2 &#181;s/pixel for a frame time of 129 ms. 
	&#8203;NOTE: Each frame consisted of 256 x 256 pixels (16 bit/pixel) with a physical dimension of 69 nm/pixel, corresponding approximately to an area of 17 &#181;m x 17 &#181;m.</li>
</ol>
3. iMSD calculation
NOTE: To properly execute the calculation, use software capable of numeric calculation and script programming. The specific script (i.e., for the &#39;iMSD.m&#39; script file, see Supporting File 1) must be present in the same directory containing the image series to be processed. Each image of the series must be saved as a distinct &#39;.tif&#39; file.
<ol>
	<li>To properly initialize the instrumental parameters used for the acquisitions, open iMSD.m with the software text editor and modify its first section as follows.
	<ol>
		<li>Set N as the number of frames in the time series (e.g., 1000 in this protocol).</li>
		<li>Set px_size: pixel size, expressed in &#181;m (e.g., 0,069 in this protocol).</li>
		<li>Set f: temporal resolution of each frame, expressed in seconds (e.g., 0,129 in this protocol).</li>
		<li>Set Filter: binary input for background correction, set the value to &#39;0&#39; to process raw images, or set the value to &#39;1&#39; to perform a threshold-based background subtraction.</li>
		<li>Set av_toll: threshold for background correction; any pixel with intensity lower than this value will be set as 0 if Filter=1.</li>
		<li>Set bit as the integer number determining the intensity sampling (e.g., 8 bit, 16 bit).</li>
	</ol>
	</li>
	<li>Save and run the edited iMSD.m script file.</li>
	<li>Check the script execution. 
	NOTE: The calculation processing status can be checked on the command window; if any fatal problem occurs, the process will be interrupted, and a warning message will be displayed to show the error type and the related line code. Otherwise, follow steps 3.3.1-3.3.3.
	<ol>
		<li>Import the image stack and subtract the background (if required).</li>
		<li>Calculate the spatiotemporal correlation function G(&#958;,&#951;,&#964;) using the Fourier method.</li>
		<li>Fit the spatiotemporal correlation function with a 2D Gaussian function. 
		NOTE: At the end of the process, the output values of the &#963;2(&#964;) fitting procedures will be shown in the command window: averages, error, and corresponding goodness (R2) of fitting are reported.</li>
	</ol>
	</li>
	<li>Check the graphical output. 
	NOTE: The iMSD curve and the corresponding fitting curves are shown in three separate panels, each for a different type of fitting equation used: Brownian diffusion, anomalous diffusion, or confined diffusion. For each curve, the R2 value is reported in the graph legend.</li>
	<li>Check the text output. 
	NOTE: At the end of the process, an &#39;.xls&#39; file is created with the same name of&#160;the original time-lapse &#39;.tif&#39; file. The first sheet contains the values of &#958;, &#951;, &#964;, and &#963;2 calculated for each time-delay. The input parameters and principal calculated output values are reported in the second sheet, i.e., diffusion coefficient, anomalous coefficient, and y-axis &#963;20 intercept.</li>
</ol>

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The authors have no conflict of interest to declare.

The properties and advantages of iMSD are evident when compared to the techniques available to retrieve analogous information. For structural information, the preferred choice is transmission electron microscopy (TEM) analysis. By this method, ultrastructural details at molecular resolution and even beyond can be retrieved, even for subcellular nanostructures. Nevertheless, the peculiar spatial resolution of TEM is achieved at the expense of the information in the temporal dimension, which is of interest here. To compensate for this, the recent advances in live-cell imaging technologies are of particular interest. These include new fluorescent markers with increased performances (e.g., brightness and photostability), optimized labeling procedures, and more sensitive detectors. In addition, analytical tools are available to address both structural (e.g., &#39;size&#39; by phasor-based analysis of local image correlation spectroscopy, PLICS23, aggregation/oligomerization by Number&#38;Brightness analysis24) and dynamic (e.g., diffusion law by single-particle tracking, i.e.,&#160;(SPT)25,26,27,28) parameters on the subcellular scale. The SPT method affords direct access to the object trajectory and its MSD. However, the disadvantage is the need for a low density of the probe and very bright labels and many single-object trajectories to be measured to satisfy statistical criteria. With respect to the temporal resolution of the measurement, inorganic, photostable probes (e.g., quantum dots or metal nanoparticles) can increase SPT performance but at the expense of complex production and labeling procedures.
Compared to these standards, the iMSD method described here shows some key advantages. First, this approach can be used in conjunction with relatively dim fluorescent tags, such as genetically encoded fluorescent proteins (e.g., the application to ISGs). Thus, compared to SPT, a higher temporal resolution is achieved (using the same label) due to the lower amount of photons required8. Second, the iMSD method is limited only by the temporal resolution but not diffraction. In fact, despite the diffraction-limited optical setup used, average molecular displacements even below the diffraction limit can be measured, as already demonstrated for molecular flows by using STICS29. The actual resolution in the measurement of displacements depends on how accurately (in terms of signal to noise) the correlation function can be measured, thus explaining why it is not limited by diffraction. Thus, it appears clear that the minimum displacement that can be measured depends on the diffusivity of the object of interest and the temporal resolution of the imaging setup.
In this regard, it is important to consider that application to subcellular nanostructures, such as macropinosomes or insulin granules, with a laser scanning microscope is optimal: the scanning speed available is significantly higher than the dynamics of the object of interest. In such a case, the movement of the objects during the acquisition is negligible, and the correlation function can be approximated by a Gaussian function. Finally, the iMSD approach can be easily applied to a wide range of commercial optical microscopy setups based on raster scan or wide-field camera-based imaging, with no need for system calibration (required only if an accurate estimate of particle size needs to be achieved). An important parameter for the method to work is proper spatial sampling. As a general rule, to reach satisfactory convergence of the fitting algorithm, the minimum size of the region of interest for imaging should be at least 3 times larger than the maximum displacement of interest.
In conclusion, the iMSD method requires only a microscope equipped for fast acquisition. The structure of interest can be tagged to any genetically encoded or organic fluorophore, thus enabling multichannel imaging. It is envisioned that cross-iMSD analysis will be used in the near future to select subpopulations of subcellular nanostructures and reveal their interactions and co-diffusion within the cell, the latter being a hot topic in cellular biophysics. If any detail is lost by iMSD analysis, this is certainly related to the large amount of molecular information within dynamic subcellular nanostructures. Such information is inevitably averaged out during the measurement due to poor temporal resolution. Theoretically, however, there is no technical limit due to the possibility to retrieve molecular information, provided that sufficient acquisition speeds can be achieved8. Due to the continuous improvements in detector speed/sensitivity and imaging technologies, it is envisioned that information about the whole subcellular compartment and its molecular constituents will be extracted from a single dataset.

Subcellular nanostructures (e.g., endocytic/secretory vesicles, organelles) play a pivotal role in cell-signaling regulation1. Proper tuning of their structural&#160;(e.g., size) and/or dynamic (e.g., diffusivity) characteristics determines how the cell responds to internal or external stimuli2,3,4. Based on this evidence, it is not surprising that alterations of these characteristics are found in many pathological conditions. Examples encompass the role of misregulated endocytosis in cancer2,3, the structural and dynamic alterations found at the level of ISGs in &#946;-cells exposed to Type-2 Diabetes conditions5, the misregulation of lysosomal structural and transport properties in globoid-cell leukodystrophy or galactosylceramide lipidosis6, and dysfunctionalities in the endo-lysosomal pathway in neurodegenerative disorders (e.g., Alzheimer&#39;s disease)7.
In this context, researchers have recently proved that the performance of standard optical microscopy methods can be enhanced by properly tuning spatial and temporal sampling resolution8. This, in turn, may provide further insight into biological processes of relevance. In practice, this is made possible by an algorithm of spatiotemporal fluctuation analysis, which simultaneously extracts the average structural and dynamic properties of diffusing objects directly from the standard stack of optical microscopy images without any need for preliminary knowledge on the biological object of interest and extraction of single-object trajectories. All this information is enclosed in a single output of the method: an iMSD trace9 (details on the iMSD trace derivation and analysis are given in Supplemental File 1).
The resulting experimental protocol consists of a few steps. First, imaging of the region of interest is performed at high temporal resolution. Then, average spatial-temporal correlation functions are calculated from the stack of images. Finally, by Gaussian fitting of the series of correlation functions, the average &#39;diffusion law&#39; is obtained directly from imaging and analyzed to recognize the object diffusion mode. The potential of the method was already proved for a variety of biological objects, ranging from molecules to nanoparticles and even entire subcellular organelles/structures9,10,11,12,13,14,15.
This paper reports iMSD application to macropinosomes to highlight their intrinsic, irreversible time-evolving nature in terms of their average (i.e., at the whole population level) structural and dynamic properties. Further, these endocytic vesicles are compared to ISGs as a reference for subcellular structures in a &#39;stationary state&#39;, i.e., a state in which the average structural/dynamic properties of the whole population of granules remain constant at any time point. Macropinocytosis defines a series of events initiated by the extensive reorganization (or ruffling) of the plasma membrane to form an external macropinocytic structure that is then internalized16. The formed early-stage macropinosomes are very similar to phagosomes. At the same time, they can be distinguished from other forms of endocytic vesicles owing to their characteristic large size, morphological heterogeneity, and lack of protein-coat structures.
Biochemical assays revealed that, upon internalization, macropinosomes progressively get enriched with protein markers of other endocytic pathways, in turn, suggesting that their identities are continuously changing during trafficking17. Using antibodies against known markers of the endosomal pathway, it was demonstrated that macropinosomes progressively adopt classical endosomal features: they diminish in size, develop into late endocytic structures (e.g., lysosomes), or eventually lose their identity via membrane-mediated retrieval of specific molecular markers (e.g., sorting nexins)18,19. The overall scenario is that every single macropinosome within the cell irreversibly changes its structural and dynamic (as well as molecular) identity during trafficking from the plasma membrane to its final intracellular fate. As a result, the structural/dynamic/molecular properties of the whole population of macropinosomes are also changing along the same temporal path. Being intrinsically sensitive to the average properties of the whole population of observed objects, the iMSD method quantitatively depicts the &#39;evolving nature&#39; by the quantification of key average parameters, i.e., the local diffusivity and anomalous coefficient (dynamic properties) and the average size of the macropinosomes (structural property) at any stage of their intracellular trafficking.
For comparison, similar measurements were performed on a well-known intracellular membrane-enclosed structure, the ISG, in a model of &#946;-cells. Like macropinosomes, the regulation of the structural and dynamic properties of ISGs, from their genesis at the Trans Golgi Network (TGN) to their exocytosis at the plasma membrane, is pivotal to the proper execution of ISG function20. However, unlike macropinosomes, ISGs live in a &#39;stationary state&#39; in which, at any time, all the functional/structural/molecular stages of ISG lifespan are simultaneously present within the cell, and each is represented by a specific subpopulation of ISGs. This means that although every single granule irreversibly evolves from biogenesis to secretion, the average structural/dynamic properties of the whole population of granules is expected to remain constant at any time point (unless the stationary state conditions are changed, for instance, by external stimuli such as glucose, cholesterol, and cytokines13). This is confirmed by iMSD analysis.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>100x Penicillin-Streptomycin-Glutamine</td>
			<td>Gibco</td>
			<td>10378-016</td>
			<td>Cell medium supplement</td>
		</tr>
		<tr>
			<td>C-peptide-EGFP</td>
			<td></td>
			<td></td>
			<td>Plasmid</td>
		</tr>
		<tr>
			<td>DMEM High Glucose</td>
			<td>Gibco</td>
			<td>31053028</td>
			<td>Cell medium (HeLa)</td>
		</tr>
		<tr>
			<td>FBS</td>
			<td>Gibco</td>
			<td>10082147</td>
			<td>Cell medium supplement</td>
		</tr>
		<tr>
			<td>Fluorescein isothiocyanate-dextran 70 kDa</td>
			<td>Sigma Aldrich</td>
			<td>46945-100MG-F</td>
			<td>Reagent</td>
		</tr>
		<tr>
			<td>HeLa</td>
			<td>ATCC</td>
			<td>CCL-61</td>
			<td>Cell Line</td>
		</tr>
		<tr>
			<td>Lipofectamine 2000</td>
			<td>TermoFisher</td>
			<td>11668019</td>
			<td>Trasfection reagent</td>
		</tr>
		<tr>
			<td>Lysotracker Red DND-99</td>
			<td>Gibco</td>
			<td>L7528</td>
			<td>Reagent</td>
		</tr>
		<tr>
			<td>Matlab</td>
			<td>MathWork</td>
			<td></td>
			<td>Software</td>
		</tr>
		<tr>
			<td>Microscope-suitable cell dishes</td>
			<td>Willco</td>
			<td>GWSt-3522</td>
			<td>Petri dishes</td>
		</tr>
		<tr>
			<td>Olympus FV1000</td>
			<td>Olympus Japan</td>
			<td></td>
			<td>Confocal microscope</td>
		</tr>
		<tr>
			<td>RPMI 1640</td>
			<td>Gibco</td>
			<td>11835063</td>
			<td>Cell medium (INS-1E)</td>
		</tr>
	</tbody>
</table>

probing structural and dynamic properties of trafficking subcellular nanostructures by spatiotemporal fluctuation spectroscopy

Imaging-derived mean square displacement (iMSD) is used to address the structural and dynamic properties of subcellular nanostructures, such as vesicles involved in the endo/exocytotic trafficking of solutes and biomolecules. iMSD relies on standard time-lapse imaging, is compatible with any optical setup, and does not need to dwell on single objects to extract trajectories. From each iMSD trace, a unique triplet of average structural and dynamic parameters (i.e., size, local diffusivity, anomalous coefficient) is calculated and combined to build the "iMSD signature" of the nanostructure under study. 
The potency of this approach is proved here with the exemplary case of macropinosomes. These vesicles evolve in time, changing their average size, number, and dynamic properties passing from early to late stages of intracellular trafficking. As a control, insulin secretory granules (ISGs) are used as a reference for subcellular structures that live in a stationary state in which the average structural and dynamic properties of the whole population of objects are invariant in time. The iMSD analysis highlights these peculiar features quantitatively and paves the way to similar applications at the sub-cellular level, both in the physiological and pathological states.

The general workflow of the method is presented in Figure 1. It recapitulates the main steps presented in the protocol section, from sample preparation (Figure 1A) to time-lapse imaging of intracellular nanostructures (Figure 1B), fluctuation analysis for the calculation of the series of spatiotemporal correlation functions (Figure 1C), and fitting for the derivation of the average structural/dynamic properties of the object under study (Figure 1D).
A critical parameter is the time resolution adopted for imaging the subcellular object of interest. This experimental value will set the time threshold at which the minimum average displacement of the objects of interest will be measured. However, the preferred condition is setting a time resolution of imaging at which the object of interest appears &#39;immobile&#39; within the captured frame, i.e., it displays a characteristic size that, on average, is not deformed due to the imaging speed. This is technically possible if the object of interest is a membrane-enclosed subcellular structure or organelle (as in this case). Typically, subcellular structures exhibit local diffusion coefficients (D, &#181;m2/s, see Table 1) that are several orders of magnitude lower than those of isolated single molecules in the cytoplasm (e.g., GFP21). Validation can be performed by artificially immobilizing the organelle of interest (e.g., by chemical fixation). Indeed, this condition can serve as a reference to determine the actual organelle size under the experimental conditions used (e.g., excitation wavelength, pixel size, objective).
Here, although lysosomes were used as test organelles for this procedure, the results are valid independently from the target structure. Figure 2A shows an image of a fixed (i.e., immobile) lysosome along with an acquisition performed on live cells at the appropriate temporal resolution (i.e., typically below 100 ms/frame; e.g., 65 ms/frame in the example in Figure 2B) and an acquisition performed intentionally at very low temporal resolution (e.g., 10 s/frame in the example in Figure 2C). For each condition, the size of the diffusing object is extracted as follows: i) an intensity profile of the spot is derived by the line tool in ImageJ software; ii) the intensity profile is plotted and interpolated by a Gaussian function to calculate the full width at half maximum (FWHM) value that, in turn, is used as an estimate of the spot diameter (Figure 2D). As expected and shown in the plot of Figure 2E, the acquisition at very high temporal resolution (i.e., 65 ms/frame) yields an average size of the structure close to that obtained in the fixed sample, either by using the standard tool described above or by extracting the iMSD y-axis intercept. Instead, the acquisition at slow speed yields an increase in the apparent size of the structure, owing to the natural structure dynamics during imaging. Under suboptimal experimental conditions, the structural/dynamic information extracted does not faithfully reflect the intrinsic properties of the object under study.
Once the main experimental parameters are selected, datasets can be produced for the target intracellular structures. For macropinosomes, after 20 min of incubation of the cells with 70 kDa dextrans, time series of the labeled intracellular structures were acquired at different time points after treatment, from 30 min up to approximately 180 min. Interestingly, a gradual change in the structural and dynamic properties of macropinosomes is detected during trafficking (Figure 3; distributions of &#963;02, Dm, &#945;, and N for macropinosomes are reported in the plots on the left). While no obvious changes in the local diffusivity (Dm) of macropinosomes are detected during trafficking, both the characteristic size (&#963;02) and overall mode of motion (&#945;)&#160;evolve in time.
Of particular note, a decrease in the average size of the macropinosomes is observed during trafficking (Figure 3A, left), together with a concomitant increase in the subdiffusive nature of their motion (i.e., denoted as a decrease in &#945; values, Figure 3C, left panel). Additionally, the number of macropinosomes was extracted from each acquisition: the results, reported in Figure 3D, left panel, clearly reveal an increase in the number of macropinosomes in time. All these results are in good agreement with the expectations because dextran-labeled macropinosomes are supposed to originate as isolated, large membrane-enclosed vesicles at the plasma membrane (that are also competent for movement along cytoskeletal components) but are supposed to gradually communicate with the endo-lysosomal pathway made up of a large population of smaller and randomly diffusing structures.
As anticipated above, the results for macropinosomes are in contrast to similar measurements performed on ISGs (Figure 3, right column). Insulin granules do not show a time-evolving trend of the iMSD-derived structural/dynamic parameters (and their average number within the cell) in the same time window observed for macropinosomes. Moreover, the characteristic values of &#963;02, Dm, and &#945; are quite different from those of lysosomes, used again as a reference. This result confirms the idea, as anticipated above, that granules are probed in a &#39;stationary state&#39; in which, at any time, the average structural/dynamic properties of the whole population of ISGs are unchanged (i.e., they remain constant, unless the stationary-state conditions change, for instance, due to external stimuli).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/62790/62790fig01.jpg" /> 
Figure 1: Experimental workflow. (A) Cells were plated 24 h (48 h for transfection experiments) before confocal experiments onto cell dishes suited for microscopic applications. Cells were then suitably treated according to the labeling method (see protocol) to stain the cytoplasmic organelle of interest. (B) A typical confocal acquisition consists of a stack of images (time-lapse) of a cytoplasmic portion of a living cell, describing the time-evolution of labeled organelle dynamics. (C) Time-lapse movie is analyzed with a custom-made Matlab script, first calculating spatiotemporal image correlation function and Gaussian fitting to plot iMSD curves (D) and related extracted fitting parameters describing structural dynamics parameters of imaged organelles. Abbreviations: iMSD = imaging-derived mean square displacement; STICS = spatiotemporal image correlation spectroscopy; &#945; = anomalous diffusion coefficient; Dm = local diffusivity; &#963;2(&#964;) = variance. <a href="https://www.jove.com/files/ftp_upload/62790/62790fig01large.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/62790/62790fig02.jpg" /> 
Figure 2: Proper experimental parameters. (A) Exemplary image of stained lysosomes in a fixed sample. Scale bar = 2 &#181;m. (B) The first frame of a stack of images of stained lysosomes in a living cell, acquired with the appropriate parameters. Temporal resolution: 65 ms/frame. Scale bar = 2 &#181;m. (C) The first frame of a stack of images of stained lysosomes in a living cell, acquired at low speed: artifactual deformation of the apparent lysosome size due to organelle motion during imaging is visible. Temporal resolution: 10 s/frame. Scale bar = 2 &#181;m. (D) Example of size calculation for imaged lysosomes in a blue ROI of (A), (B), and (C). The intensity profile along the blue line was fitted with a Gaussian function to retrieve the FWHM, i.e., an estimate of spot size. FWHM values are reported for each fitting. (E) Graphical representation of size values obtained by image analysis described in panel (D) for all imaged lysosomes (black square, mean value, and standard deviation), for lysosomes enclosed within the blue ROI (blue triangle), and retrieved by iMSD analysis (red circle). This figure is from 22. Abbreviations: ROI = region of interest; FWHM = full-width at half-maximum; iMSD = imaging-derived mean square displacement. <a href="https://www.jove.com/files/ftp_upload/62790/62790fig02large.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/62790/62790fig03.jpg" /> 
Figure 3: Temporal evolution of labeled organelles. (A) Plots of iMSD-extracted size values vs. time progression of subsequent acquisitions represented as a mean of measured values in acquisitions performed on a 10-min time window. On the left, progressive reduction in the average size of macropinosomes (black circles) and on the right, the time-invariant size of insulin secretory granules (black squares) compared to lysosomes, represented as average size value (thicker red line) &#177; standard deviation (dashed red lines). (B) and (C) Time progression of Dm and &#945; coefficients for macropinosomes (left) and insulin granules (right) extracted by iMSD analysis. (D) Time-evolution of numbers of labeled macropinosomes and insulin granules measured in the first frame of each acquired time-lapse movie. Abbreviations: iMSD = imaging-derived mean square displacement; &#945; = anomalous diffusion coefficient; Dm = local diffusivity, N = number. <a href="https://www.jove.com/files/ftp_upload/62790/62790fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td colspan="2">Organelle</td>
			<td colspan="2">Labelling</td>
			<td>Cell Line</td>
			<td>Size (nm)</td>
			<td>Dm ( &#215; 10-3 &#956;m2/s)</td>
			<td>&#945;</td>
			<td>N</td>
			<td>Ref.</td>
		</tr>
		<tr>
			<td colspan="2">Early Endosome (EE)</td>
			<td colspan="2">CellLight Early Endosome GFP</td>
			<td>HeLa</td>
			<td>395&#177;74</td>
			<td>3.0&#177;2.4</td>
			<td>1.02&#177;0.20</td>
			<td>40</td>
			<td>10</td>
		</tr>
		<tr>
			<td colspan="2">Late Endosome (LE)</td>
			<td colspan="2">CellLight Late Endosome GFP</td>
			<td>HeLa</td>
			<td>693&#177;102</td>
			<td>15.4&#177;10.6</td>
			<td>0.57&#177;0.16</td>
			<td>58</td>
			<td>10</td>
		</tr>
		<tr>
			<td colspan="2">Lysosome (LY)</td>
			<td colspan="2">LysoTracker DND-99</td>
			<td>HeLa</td>
			<td>471&#177;76</td>
			<td>15.3&#177;9.0</td>
			<td>0.49&#177;0.13</td>
			<td>143</td>
			<td>10, 14</td>
		</tr>
		<tr>
			<td colspan="2">Caveola (CAV)</td>
			<td colspan="2">Caveolin-EGFP</td>
			<td>HeLa</td>
			<td>405&#177;49</td>
			<td>3.1&#177;1.8</td>
			<td>1.00&#177;0.22</td>
			<td>15</td>
			<td>10</td>
		</tr>
		<tr>
			<td colspan="2">Clathrin Coated Vescicle (CCV)</td>
			<td colspan="2">Transferrin-Alexa 488</td>
			<td>HeLa</td>
			<td>513&#177;62</td>
			<td>16.2&#177;9.9</td>
			<td>0.48&#177;0.17</td>
			<td>33</td>
			<td>10</td>
		</tr>
		<tr>
			<td colspan="2">Insulin Granule (IG)</td>
			<td colspan="2">C-peptide-EGFP</td>
			<td>INS-1E</td>
			<td>335&#177;56</td>
			<td>3.0&#177;1.7</td>
			<td>0.70&#177;0.14</td>
			<td>107</td>
			<td>11</td>
		</tr>
		<tr>
			<td colspan="2">Early Macropinosome (EMCR)</td>
			<td colspan="2">Fluorescein-Dextran 70 kDa</td>
			<td>HeLa</td>
			<td>979&#177;423</td>
			<td>8.3&#177;9</td>
			<td>0.79&#177;0.27</td>
			<td>36</td>
			<td>10</td>
		</tr>
		<tr>
			<td colspan="2">Intermediate Macropinosome (IMCR)</td>
			<td colspan="2">Fluorescein-Dextran 70 kDa</td>
			<td>HeLa</td>
			<td>702&#177;180</td>
			<td>13.7&#177;19.9</td>
			<td>0.60&#177;0.38</td>
			<td>29</td>
			<td>10</td>
		</tr>
		<tr>
			<td colspan="2">Late Macropinosome (LMCR)</td>
			<td colspan="2">Fluorescein-Dextran 70 kDa</td>
			<td>HeLa</td>
			<td>592&#177;127</td>
			<td>5.8&#177;4.7</td>
			<td>0.39&#177;0.21</td>
			<td>21</td>
			<td>10</td>
		</tr>
	</tbody>
</table>
Table 1: Structural and dynamic iMSD-extracted parameters. The table shows the values of size, Dm, and &#945; coefficient measured for different organelles, specifying labeling strategies, the cell line used, and the number of analyzed acquisitions. Values are reported as mean &#177; standard deviation. Abbreviations: iMSD = imaging-derived mean square displacement; &#945; = anomalous diffusion coefficient; Dm = local diffusivity, N = number; GFP = green fluorescent protein; EGFP = enhanced green fluorescent protein.
Supplemental File 1: Details on the iMSD trace derivation and analysis. <a href="https://www.jove.com/files/ftp_upload/62790/Supplementary File 1 - iMSD calculation.docx" target="_blank">Please click here to download this File.</a>
Supporting File 1: <a href="https://www.jove.com/files/ftp_upload/62790/Supplementary File 2 - MatLAB script.m" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Probing Structural and Dynamic Properties of Trafficking Subcellular Nanostructures by Spatiotemporal Fluctuation Spectroscopy at JoVE.com

Probing Structural and Dynamic Properties of Trafficking Subcellular Nanostructures by Spatiotemporal Fluctuation Spectroscopy

Imaging-derived mean square displacement (iMSD) analysis is applied to macropinosomes to highlight their intrinsic time-evolving nature in terms of structural and dynamic properties. Macropinosomes are then compared to insulin secretory granules (ISGs) as a reference for subcellular structures with time-invariant average structural/dynamic properties.

Imaging-derived mean square displacement (iMSD) is used to address the structural and dynamic properties of subcellular nanostructures, such as vesicles ...

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JoVE Journal

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1.7K Views.  Scuola Normale Superiore. Imaging-derived mean square displacement (iMSD) analysis is applied to macropinosomes to highlight their intrinsic time-evolving nature in terms of structural and dynamic properties. Macropinosomes are then compared to insulin secretory granules (ISGs) as a reference for subcellular structures with time-invariant average structural/dynamic properties.

Video: Probing Structural and Dynamic Properties of Trafficking Subcellular Nanostructures by Spatiotemporal Fluctuation Spectroscopy

Application of Light-cured Dental Adhesive Resin for Mounting Electrodes or Microdialysis Probes in Chronic Experiments

In chronic recording experiments, self-curing dental acrylic resins have been used as a mounting base of electrodes or microdialysis-probes. Since these acrylics do not bond to the bone, screws have been used as anchors. However, in small experimental animals like finches or mouse, their craniums are very fragile and can not successfully hold the anchors. In this report, we propose a new application of light-curing dental resins for mounting base of electrodes or microdialysis probes in chronic experiments. This material allows direct bonding to the cranium. Therefore, anchor screws are not required and surgical field can be reduced considerably. Past experiences show that the bonding effect maintains more than 2 months. Conventional resin's window of time when the materials are pliable and workable is a few minutes. However, the window of working time for these dental adhesives is significantly wider and adjustable.

In this report, we propose a new application of light-curing dental resins for mounting base of electrodes or microdialysis probes in chronic experiments. This material allows direct bonding to the cranium.

In chronic recording experiments, self-curing dental acrylic resins have been used as a mounting base of electrodes or microdialysis-probes. Since these ...

Analyzing Large Protein Complexes by Structural Mass Spectrometry

Living cells control and regulate their biological processes through the coordinated action of a large number of proteins that assemble themselves into an array of dynamic, multi-protein complexes1. To gain a mechanistic understanding of the various cellular processes, it is crucial to determine the structure of such protein complexes, and reveal how their structural organization dictates their function. Many aspects of multi-protein complexes are, however, difficult to characterize, due to their heterogeneous nature, asymmetric structure, and dynamics. Therefore, new approaches are required for the study of the tertiary levels of protein organization.
One of the emerging structural biology tools for analyzing macromolecular complexes is mass spectrometry (MS)2-5. This method yields information on the complex protein composition, subunit stoichiometry, and structural topology. The power of MS derives from its high sensitivity and, as a consequence, low sample requirement, which enables examination of protein complexes expressed at endogenous levels. Another advantage is the speed of analysis, which allows monitoring of reactions in real time. Moreover, the technique can simultaneously measure the characteristics of separate populations co-existing in a mixture. 
Here, we describe a detailed protocol for the application of structural MS to the analysis of large protein assemblies. The procedure begins with the preparation of gold-coated capillaries for nanoflow electrospray ionization (nESI). It then continues with sample preparation, emphasizing the buffer conditions which should be compatible with nESI on the one hand, and enable to maintain complexes intact on the other. We then explain, step-by-step, how to optimize the experimental conditions for high mass measurements and acquire MS and tandem MS spectra. Finally, we chart the data processing and analyses that follow. Rather than attempting to characterize every aspect of protein assemblies, this protocol introduces basic MS procedures, enabling the performance of MS and MS/MS experiments on non-covalent complexes. Overall, our goal is to provide researchers unacquainted with the field of structural MS, with knowledge of the principal experimental tools.

Mass spectrometry has proven to be a valuable tool for analyzing large protein complexes. This method enables insights into the composition, stoichiometry and overall architecture of multi-subunit assemblies. Here, we describe, step-by-step, how to perform a structural mass spectrometry analysis, and characterize macromolecular structures.

Living cells control and regulate their biological processes through the coordinated action of a large number of proteins that assemble themselves into an ...

Harvesting and Cryo-cooling Crystals of Membrane Proteins Grown in Lipidic Mesophases for Structure Determination by Macromolecular Crystallography

An important route to understanding how proteins function at a mechanistic level is to have the structure of the target protein available, ideally at atomic resolution. Presently, there is only one way to capture such information as applied to integral membrane proteins (Figure 1), and the complexes they form, and that method is macromolecular X-ray crystallography (MX). To do MX diffraction quality crystals are needed which, in the case of membrane proteins, do not form readily. A method for crystallizing membrane proteins that involves the use of lipidic mesophases, specifically the cubic and sponge phases1-5, has gained considerable attention of late due to the successes it has had in the G protein-coupled receptor field6-21 (<a href="http://www.mpdb.tcd.ie" target="_blank">www.mpdb.tcd.ie</a>). However, the method, henceforth referred to as the in meso or lipidic cubic phase method, comes with its own technical challenges. These arise, in part, due to the generally viscous and sticky nature of the lipidic mesophase in which the crystals, which are often micro-crystals, grow. Manipulating crystals becomes difficult as a result and particularly so during harvesting22,23. Problems arise too at the step that precedes harvesting which requires that the glass sandwich plates in which the crystals grow (Figure 2)24,25 are opened to expose the mesophase bolus, and the crystals therein, for harvesting, cryo-cooling and eventual X-ray diffraction data collection.
The cubic and sponge mesophase variants (Figure 3) from which crystals must be harvested have profoundly different rheologies4,26. The cubic phase is viscous and sticky akin to a thick toothpaste. By contrast, the sponge phase is more fluid with a distinct tendency to flow. Accordingly, different approaches for opening crystallization wells containing crystals growing in the cubic and the sponge phase are called for as indeed different methods are required for harvesting crystals from the two mesophase types. Protocols for doing just that have been refined and implemented in the Membrane Structural and Functional Biology (MS&amp;FB) Group, and are described in detail in this JoVE article (Figure 4). Examples are given of situations where crystals are successfully harvested and cryo-cooled. We also provide examples of cases where problems arise that lead to the irretrievable loss of crystals and describe how these problems can be avoided. In this article the Viewer is provided with step-by-step instructions for opening glass sandwich crystallization wells, for harvesting and for cryo-cooling crystals of membrane proteins growing in cubic and in sponge phases.

Herein is described procedures implemented in the Caffrey Membrane Structural and Functional Biology Group to harvest and cryo-cool membrane protein crystals grown in lipidic cubic and sponge phases for use in structure determination using macromolecular X-ray crystallography.

An important route to understanding how proteins function at a mechanistic level is to have the structure of the target protein available, ideally at ...

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, ...

Visualization of ATP Synthase Dimers in Mitochondria by Electron Cryo-tomography

Electron cryo-tomography is a powerful tool in structural biology, capable of visualizing the three-dimensional structure of biological samples, such as cells, organelles, membrane vesicles, or viruses at molecular detail. To achieve this, the aqueous sample is rapidly vitrified in liquid ethane, which preserves it in a close-to-native, frozen-hydrated state. In the electron microscope, tilt series are recorded at liquid nitrogen temperature, from which 3D tomograms are reconstructed. The signal-to-noise ratio of the tomographic volume is inherently low. Recognizable, recurring features are enhanced by subtomogram averaging, by which individual subvolumes are cut out, aligned and averaged to reduce noise. In this way, 3D maps with a resolution of 2 nm or better can be obtained. A fit of available high-resolution structures to the 3D volume then produces atomic models of protein complexes in their native environment. Here we show how we use electron cryo-tomography to study the in situ organization of large membrane protein complexes in mitochondria. We find that ATP synthases are organized in rows of dimers along highly curved apices of the inner membrane cristae, whereas complex I is randomly distributed in the membrane regions on either side of the rows. By subtomogram averaging we obtained a structure of the mitochondrial ATP synthase dimer within the cristae membrane.

We present a protocol of how to collect and process electron cryo-tomograms of whole mitochondria. The technique provides detailed insights into the structure, function, and organization of large membrane protein complexes in native biological membranes.

Electron cryo-tomography is a powerful tool in structural biology, capable of visualizing the three-dimensional structure of biological samples, such as ...

AFM-based Mapping of the Elastic Properties of Cell Walls: at Tissue, Cellular, and Subcellular Resolutions

We describe a recently developed method to measure mechanical properties of the surfaces of plant tissues using atomic force microscopy (AFM) micro/nano-indentations, for a JPK&#160;AFM. Specifically, in this protocol we measure the apparent Young&#8217;s modulus of cell walls at subcellular resolutions across regions of up to 100 &#181;m&#160;x 100 &#181;m in floral meristems, hypocotyls, and roots. This requires careful preparation of the sample, the correct selection of micro-indenters and indentation depths. To account for cell wall properties only, measurements are performed in highly concentrated solutions of mannitol in order to plasmolyze the cells and thus remove the contribution of cell turgor pressure.
In contrast to other extant techniques, by using different indenters and indentation depths, this method allows simultaneous multiscale measurements, i.e. at subcellular resolutions and across hundreds of cells comprising a tissue. This means that it is now possible&#160;to spatially-temporally characterize the changes that take place in the mechanical properties of cell walls during development, enabling these changes to be correlated with growth and differentiation. This represents a key step to understand how coordinated microscopic cellular changes bring about macroscopic morphogenetic events.
However, several limitations remain: the method can only be used on fairly small samples (around 100 &#181;m in diameter) and only on external tissues; the method is sensitive to tissue topography; it measures only certain aspects of the tissue&#8217;s complex mechanical properties. The technique is being developed rapidly and it is likely that most of these limitations will be resolved in the near future.

We describe a method to map mechanical properties of plant tissues using an atomic force microscope (AFM). We focus on how to record mechanical changes that take place in cell walls during plant development at wide-field mesoscale, enabling these changes to be correlated with growth and morphogenesis.

We describe a recently developed method to measure mechanical properties of the surfaces of plant tissues using atomic force microscopy (AFM) ...

Tracking Drug-induced Changes in Receptor Post-internalization Trafficking by Colocalizational Analysis

The intracellular trafficking of receptors is a collection of complex and highly controlled processes. Receptor trafficking modulates signaling and overall cell responsiveness to ligands and is, itself, influenced by intra- and extracellular conditions, including ligand-induced signaling. Optimized for use with monolayer-plated cultured cells, but extendable to free-floating tissue slices, this protocol uses immunolabelling and colocalizational analysis to track changes in intracellular receptor trafficking following both chronic/prolonged and acute interventions, including exogenous drug treatment. After drug treatment, cells are double-immunolabelled for the receptor and for markers for the intracellular compartments of interest. Sequential confocal microscopy is then used to capture two-channel photomicrographs of individual cells, which are subjected to computerized colocalizational analysis to yield quantitative colocalization scores. These scores are normalized to permit pooling of independent replicates prior to statistical analysis. Representative photomicrographs may also be processed to generate illustrative figures. Here, we describe a powerful and flexible technique for quantitatively assessing induced receptor trafficking.

Receptor trafficking modulates signaling and cell responsiveness to ligands and is, itself, responsive to cell conditions, including ligand-induced signaling. Here, we describe a powerful and flexible technique for quantitatively assessing drug-induced receptor trafficking using immunolabeling and colocalizational analysis.

The intracellular trafficking of receptors is a collection of complex and highly controlled processes. Receptor trafficking modulates signaling and ...

Analysis of SCAP N-glycosylation and Trafficking in Human Cells

Elevated lipogenesis is a common characteristic of cancer and metabolic diseases. Sterol regulatory element-binding proteins (SREBPs), a family of membrane-bound transcription factors controlling the expression of genes important for the synthesis of cholesterol, fatty acids and phospholipids, are frequently upregulated in these diseases. In the process of SREBP nuclear translocation, SREBP-cleavage activating protein (SCAP) plays a central role in the trafficking of SREBP from the endoplasmic reticulum (ER) to the Golgi and in subsequent proteolysis activation. Recently, we uncovered that glucose-mediated N-glycosylation of SCAP is a prerequisite condition for the exit of SCAP/SREBP from the ER and movement to the Golgi. N-glycosylation stabilizes SCAP and directs SCAP/SREBP trafficking. Here, we describe a protocol for the isolation of membrane fractions in human cells and for the preparation of the samples for the detection of SCAP N-glycosylation and total protein by using western blot. We further provide a method to monitor SCAP trafficking by using confocal microscopy. This protocol is appropriate for the investigation of SCAP N-glycosylation and trafficking in mammalian cells.

Analysis of SCAP N-glycosylation and Trafficking in Human Cells

We describe a modified method for membrane fraction isolation from human cells and sample preparation for the detection of SCAP N-glycosylation and total protein by using western blot. We further introduce a GFP-labeling method to monitor SCAP trafficking using confocal microscopy. This protocol can be used in regular biology laboratories.

Elevated lipogenesis is a common characteristic of cancer and metabolic diseases. Sterol regulatory element-binding proteins (SREBPs), a family of ...

Spatiotemporal Analysis of Cytokinetic Events in Fission Yeast

Cytokinesis, the final step in cell division is critical for maintaining genome integrity. Proper cytokinesis is important for cell differentiation and development. Cytokinesis involves a series of events that are well coordinated in time and space. Cytokinesis involves the formation of an actomyosin ring at the division site, followed by ring constriction, membrane furrow formation and extra cellular matrix remodeling. The fission yeast, Schizosaccharomyces pombe (S.&#160;pombe)&#160;is a well-studied model system that has revealed with substantial clarity the initial events in cytokinesis. However, we do not understand clearly how different cytokinetic events are coordinated spatiotemporally. To determine this, one needs to analyze the different cytokinetic events in great details in both time and in space. Here we describe a microscopy approach to examine different cytokinetic events in live cells. With this approach it is possible to time different cytokinetic events and determine the time of recruitment of different proteins during cytokinesis. In addition, we describe protocols to compare protein localization, and distribution at the site of cell division. This is a basic protocol to study cytokinesis in fission yeast and can also be used for other yeasts and fungal systems.

The fission yeast, Schizosaccharomyces pombe&#160;is an excellent model system to study cytokinesis, the final stage in cell division. Here we describe a microscopy approach to analyze different cytokinetic events in live fission yeast cells.

Cytokinesis, the final step in cell division is critical for maintaining genome integrity. Proper cytokinesis is important for cell differentiation and ...

Utilizing pHluorin-tagged Receptors to Monitor Subcellular Localization and Trafficking

Understanding membrane protein trafficking, assembly, and expression requires an approach that differentiates between those residing in intracellular organelles and those localized on the plasma membrane. Traditional fluorescence-based measurements lack the capability to distinguish membrane proteins residing in different organelles. Cutting edge methodologies transcend traditional methods by coupling pH-sensitive fluorophores with total internal reflection fluorescence microscopy (TIRFM). TIRF illumination excites the sample up to approximately 150 nm from the glass-sample interface, thus decreasing background, increasing the signal to noise ratio, and enhancing resolution. The excitation volume in TIRFM encompasses the plasma membrane and nearby organelles such as the peripheral ER. Superecliptic pHluorin (SEP) is a pH sensitive version of GFP. Genetically encoding SEP into the extracellular domain of a membrane protein of interest positions the fluorophore on the luminal side of the ER and in the extracellular region of the cell. SEP is fluorescent when the pH is greater than 6, but remains in an off state at lower pH values. Therefore, receptors tagged with SEP fluoresce when residing in the endoplasmic reticulum (ER) or upon insertion in the plasma membrane (PM) but not when confined to a trafficking vesicle or other organelles such as the Golgi. The extracellular pH can be adjusted to dictate the fluorescence of receptors on the plasma membrane. The difference in fluorescence between TIRF images at neutral and acidic extracellular pH for the same cell corresponds to a relative number of receptors on the plasma membrane. This allows a simultaneous measurement of intracellular and plasma membrane resident receptors. Single vesicle insertion events can also be measured when the extracellular pH is neutral, corresponding to a low pH trafficking vesicle fusing with the plasma membrane and transitioning into a fluorescent state. This versatile technique can be exploited to study localization, expression, and trafficking of membrane proteins.

Labeling the extracellular domain of a membrane protein with a pH sensitive fluorophore, superecliptic pHluorin (SEP), allows subcellular localization, expression, and trafficking to be determined. Imaging SEP-labeled proteins with total internal reflection fluorescence microscopy (TIRFM) enables the quantification of protein levels in the peripheral ER and plasma membrane.

Understanding membrane protein trafficking, assembly, and expression requires an approach that differentiates between those residing in intracellular ...