The authors would like to thank Integrated Imaging Core facility at Johns Hopkins University for microscopes and data analysis software. We thank J. Snedeker and Q. E. Yu for proofreading and suggestions, and X.C. lab members for helpful discussions and suggestions. Supported by NIGMS/NIH R35GM127075, the Howard Hughes Medical Institute, the David and Lucile Packard Foundation, and Johns Hopkins University startup funds (X.C.).

1. Preparation of live cell imaging cocktail (live cell media)
<ol>
	<li>Supplement Schneider&#8217;s Drosophila medium with 15% fetal bovine serum (FBS) and 0.6x penicillin/streptomycin. Adjust the pH to approximately 7.0.</li>
	<li>Just before using the medium, add insulin to a final concentration of 200 &#181;g/mL. 
	NOTE: This media is critical for maintaining normal cell division and development of the Drosophila testis during time-lapse imaging10,14.</li>
</ol>
2. Preparation of glass bottom cell culture dish
<ol>
	<li>Use a poly L-lysine coated glass-bottom cell culture dish with a diameter of 35 mm for the Drosophila testis. The inner well of the dish has a glass bottom with a diameter of 23 mm and a side with an elevation of 1 mm (Figure 1A-a). 
	NOTE: Choose a glass-bottom dish that allows for a shorter working distance, larger numerical aperture, and higher magnification objective; these features are crucial to obtain a super-resolution image.</li>
	<li>Use a dialysis membrane (MWCO: 12&#8211;14kD) that allows for the gaseous exchange. Cut the membrane into small pieces. 
	NOTE: The membrane pieces should be smaller than the glass area of the dish but large enough to cover the specimen.</li>
	<li>Soak the membrane with 100 &#181;L of live cell media for ~5 min. Do not use a dry membrane on the sample, as it may damage it. Membrane helps prevent hypoxic stress.</li>
	<li>Prepare 2&#8211;3 light-weight glass or plastic pieces to put onto the membrane so that the tissue will not float. To do this, first take the outer ring of the 50 mL centrifuge tube and cut it into small pieces. Then, sterilize the pieces with 70% ethanol before use. 
	NOTE: This step ensures that the sample remains on the surface during imaging and does not float, which is crucial for obtaining optimal results.</li>
	<li>Prepare a ring ~25 mm in diameter and place it at the elevated side of the dish. Put a coverslip on it to generate two chambers. The bottom chamber will contain the sample while the top chamber will serve as a humidity chamber to prevent the sample from drying.
	<ol>
		<li>To make this ring, cut off the outer ring from a 50 mL centrifuge tube, which will allow it to fit securely on the elevated side of the 35 mm dish. A similar ring can be made in other ways, for example, by using a rubber band or plastic twist tie to fit on the elevated side of the dish to properly hold the coverslip without interfering with the sample. 
		NOTE: This step is critical, especially for samples sensitive to stresses such as hypoxic conditions, and changes in temperature and humidity.</li>
	</ol>
	</li>
</ol>
3. Dissection of testes from male flies and mounting
<ol>
	<li>Take ~10 young male flies (2&#8211;3 day old) and dissect the testes in the live cell media to obtain ~10 pairs of Drosophila adult testes.
	<ol>
		<li>Dissect the flies under a dissection microscope in a dissection dish using fine forceps. 
		NOTE: The Drosophila melanogaster (fruit fly) strain carries UAS-&#945;-Tubulin-GFP transgene driven by an early germ cell driver nanos-Gal4.</li>
		<li>Generate the following knock-in Drosophila melanogaster strains using the CRISPR-Cas9 technology: Lamin-mCherry (C-terminal tag) and CENP-A-Dendra2-CENP-A [tag at the internal site (between 118th - 119th codon)]. 
		NOTE: While only one testis of excellent quality is needed for each experiment, mounting enough number of tissues (15&#8211;20) or cells will ensure that there will be at least one healthy sample with excellent fluorescence signals for time-lapse imaging.</li>
	</ol>
	</li>
	<li>Wash the testes twice in live cell media in the dissection dish.
	<ol>
		<li>Use a pipette to add media followed by removing the live cell media as a washing step.</li>
		<li>Remove excess tissue using forceps. Any extra tissue will interfere with imaging. 
		NOTE: Do not use phosphate-buffered saline (PBS) for washing, because it may affect the dynamics of certain cellular components.</li>
	</ol>
	</li>
	<li>Add 100&#8211;150 &#181;L of live cell media into the dish (prepared in step 2.1) and spread it on the glass surface using a pipette tip. Spreading the media on the surface will allow the tissue to properly stick to the dish.</li>
	<li>Transfer the testes to the dish using fine forceps and bring them to the center of the dish (Figure 1A-b). 
	NOTE: Avoid transferring debris because it will interfere with imaging and could reduce the quality of the image.</li>
	<li>Remove excess live cell media (leave approximately 10 &#181;L of media) to allow for the tissue to flatten and stick properly to the dish. Perform this step quickly to avoid drying the sample.</li>
	<li>Place the pre-wet membrane (prepared in step 2.2 and 2.3) on top of the testes (Figure 1A-c).</li>
	<li>Put 2&#8211;3 small plastic weights (prepared in Step 2.4) on the membrane so that the sample will not float and quickly add 100&#8211;150 &#181;L of live cell media (Figure 1A-d). 
	NOTE: Adding media quickly and gently ensures that the sample will not dry out or become displaced.</li>
	<li>Place the plastic ring (prepared in step 2.5) on the elevated side of the dish (Figure 1B-a).</li>
	<li>Place a 22 mm x 22 mm coverslip on top of the ring (Figure 1B-b). This generates two chambers.</li>
	<li>Take a piece of tissue paper, dampen it with water, then swirl it and put it on the coverslip, to create a humid chamber (Figure 1B-c). Close the lid of the dish (Figure 1B-d) and begin live cell imaging. 
	NOTE: If the sample is light sensitive, perform section 3 in the dark.</li>
</ol>
4. Live cell imaging of Drosophila male germline stem cells (GSC) in situ
<ol>
	<li>Place the dish under the super-resolution microscope and secure it with the stage clamps.
	<ol>
		<li>Open the imaging software, turn on the transmitted light, and use 63x objective to focus on the testis tissue with the focus knob. 
		NOTE: If there is a difficulty in locating a sample with the 63x objective, then first use the 40x or 20x objective before switching to the 63x.</li>
		<li>Turn on the lasers, click Live, and find the GSC with the optimal position and condition. Avoid testes that have a low fluorescence signal or have the hub (niche) away from the surface. 
		NOTE: In Drosophila testes, the GSCs are attached to the hub.</li>
	</ol>
	</li>
	<li>Adjust the focus for the GSCs and identify the right settings for the sample, including laser power, electron-multiplying (EM) gain, averaging, and zoom for the Airyscan mode. Use the following settings as an example for Drosophila testes expressing EGFP tagged &#945;-tubulin in early-stage germ cells.
	<ol>
		<li>Set the frame size between 512 x 512 pixels and 1024 x 1024 pixels by clicking Frame Size.</li>
		<li>Set the frame average to 1 or 2 by clicking Averaging. 
		NOTE: Increasing the frame size (&#62;1024 x 1024 pixels) and performing more averaging (i.e., more than 2) could result in photobleaching or phototoxicity.</li>
		<li>Set the laser power to 1%&#8211;2% by clicking Lasers. 
		NOTE: Boosting the laser power may also lead to photobleaching or phototoxicity.</li>
		<li>Set the EM gain below the recommended level (&#60; 800) by clicking Master Gain. 
		NOTE: A higher EM gain may generate artifacts.</li>
		<li>Zoom in to the region or cell of interest to reduce the image acquisition time and reduce photobleaching. This also allows the specimen to survive for a longer time.</li>
	</ol>
	</li>
	<li>Start the time-lapse image capture using the Airyscan mode by clicking Start Experiment.
	<ol>
		<li>Before clicking Start Experiment, make sure that the acquisition is optimally configured.</li>
		<li>If it displays a warning with the notice &#8220;Airyscan acquisition is not configured optimally&#8221;, then click Optimal in frame size section, click Optimal in z-stack section, and click Optimal for scan area section (a minimum of 1.3 zoom is required for super-resolution imaging).</li>
	</ol>
	</li>
	<li>Optimize the time interval, number of z-slices, and duration of the time-lapse imaging according to the experimental design and the type of specimen, so that the quality of the image is not compromised and the cells will not be arrested at particular stages of the cell cycle.
	<ol>
		<li>As an example, the parameters for the time-lapse imaging of Drosophila testes expressing EGFP-tagged &#945;-tubulin in early germline are shown in the following steps.</li>
		<li>For more than 5 h time-lapse imaging, image at a 10 min interval with 1% laser power and take up to 30 z-slices at each time point.</li>
		<li>For highly dynamic cellular processes, such as microtubule dynamics, set a 2 min interval between time points, perform 1&#8211;2 h time-lapse imaging at 1% laser power, and take up to 30 z-slices at each time point.</li>
		<li>For rare cellular events, such as anaphase to early telophase chromatin dynamics (2&#8211;3 min events), set live cell imaging with a 1 min interval between time points, perform 30 min time-lapse imaging at 1% laser power, and take up to 30 z-slices at each time point. 
		NOTE: These parameters may vary for different samples, so alteration will be needed for optimal use for the specific sample.</li>
	</ol>
	</li>
	<li>Perform either time-lapse imaging or live snapshot imaging, depending on the sensitivity of the sample and the experimental design. 
	NOTE: A short movie is 15&#8211;30 min, and a long movie is more than 5 h.</li>
	<li>If the sample is very sensitive and cannot be studied by time-lapse imaging with a super-resolution microscope, as is often the case with Drosophila male GSCs, perform a super-resolution live snapshot (SRLS) of the sample at different cell cycle stages (as shown in Figure 2, Figure 3, Figure 4).
	<ol>
		<li>If a rare cell biological event is being captured or the protein of interest has low expression level, then perform SRLS.</li>
		<li>For SRLS, find a cell at the specific cell cycle stage you are interested in. For example, if focusing on microtubules-kinetochore binding, then use a cell at prometaphase or metaphase. 
		NOTE: This will help ensure that you get a high-quality image of the cell of interest at the right time without risking phototoxicity to the sample or bleaching of the fluorophores, which may occur during a longer movie.</li>
		<li>If using SRLS, image different cell cycle stages of interest across multiple cells and arrange these images in chronological order. 
		NOTE: This can be used to construct a temporal order of events while maximizing signal and tissue health, to obtain excellent temporal resolution of the event for particularly sensitive samples or samples with low signal.</li>
	</ol>
	</li>
</ol>
5. Airyscan processing of the live cell images
<ol>
	<li>After image acquisition with the imaging software, select Processing, click Batch, and then select Airyscan Processing.
	<ol>
		<li>Select the images to be processed and then click Run/Process to obtain the super-resolution images.</li>
		<li>Perform the processing using the default setting. 
		NOTE: Airyscan processing will only work if the imaging settings are properly established for Airyscan imaging.</li>
	</ol>
	</li>
</ol>

<ol>
	<li>Breedijk, R. M. P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+live-cell+super-resolution+technique+demonstrated+by+imaging+germinosomes+in+wild-type+bacterial+spores.">A live-cell super-resolution technique demonstrated by imaging germinosomes in wild-type bacterial spores.</a> Scientific Reports. 10 (1), 5312 (2020).</li><li>Maddox, P. S., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+the+mitotic+spindle.">Imaging the mitotic spindle.</a> Methods in Enzymology. 505, 81-103 (2012).</li><li>Frigault, M. M., Lacoste, J., Swift, J. L., Brown, C. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live-cell+microscopy+-+tips+and+tools.">Live-cell microscopy - tips and tools.</a> Journal of Cell Science. 122 (6), 753-767 (2009).</li><li>Bates, M., Jones, S. A., Zhuang, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stochastic+optical+reconstruction+microscopy+(STORM):+A+method+for+super+resolution+fluorescence+imaging.">Stochastic optical reconstruction microscopy (STORM): A method for super resolution fluorescence imaging.</a> Cold Spring Harbor Protocols. 2013 (6), 075143 (2013).</li><li>Shroff, H., Galbraith, C. G., Galbraith, J. A., Betzig, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Live-cell+photoactivated+localization+microscopy+of+nanoscale+adhesion+dynamics.">Live-cell photoactivated localization microscopy of nanoscale adhesion dynamics.</a> Nature Methods. 5 (5), 417-423 (2008).</li><li>Vicidomini, G., Bianchini, P., Diaspro, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=STED+super-resolved+microscopy.">STED super-resolved microscopy.</a> Nature Methods. 15 (3), 173-182 (2018).</li><li>Huff, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Airyscan+detector+from+ZEISS:+confocal+imaging+with+improved+signal-to-noise+ratio+and+super-resolution.">The Airyscan detector from ZEISS: confocal imaging with improved signal-to-noise ratio and super-resolution.</a> Nature Methods. 12 (12), (2015).</li><li>Huff, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+new+2D+superresolution+mode+for+ZEISS+Airyscan.">The new 2D superresolution mode for ZEISS Airyscan.</a> Nature Methods. 14 (12), 1223 (2017).</li><li>Korobchevskaya, K., Lagerholm, B., Colin-York, H., Fritzsche, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Exploring+the+potential+of+Airyscan+microscopy+for+live+cell+imaging.">Exploring the potential of Airyscan microscopy for live cell imaging.</a> Photonics. 4 (4), 41 (2017).</li><li>Ranjan, R., Snedeker, J., Chen, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Asymmetric+centromeres+differentially+coordinate+with+mitotic+machinery+to+ensure+biased+sister+chromatid+segregation+in+germline+stem+cells.">Asymmetric centromeres differentially coordinate with mitotic machinery to ensure biased sister chromatid segregation in germline stem cells.</a> Cell Stem Cell. 25 (5), 666-681 (2019).</li><li>Kahney, E. W., Ranjan, R., Gleason, R. J., Chen, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Symmetry+from+asymmetry+or+asymmetry+from+symmetry.">Symmetry from asymmetry or asymmetry from symmetry.</a> Cold Spring Harbor Symposia on Quantitative Biology. 82, 305-318 (2017).</li><li>Wooten, M., Ranjan, R., Chen, X. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Asymmetric+histone+inheritance+in+asymmetrically+dividing+stem+cells.">Asymmetric histone inheritance in asymmetrically dividing stem cells.</a> Trends in Genetics. 36 (1), 30-43 (2020).</li><li>Wooten, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Asymmetric+histone+inheritance+via+strand-specific+incorporation+and+biased+replication+fork+movement.">Asymmetric histone inheritance via strand-specific incorporation and biased replication fork movement.</a> Nature Structural &amp; Molecular Biology. 26 (8), 732-743 (2019).</li><li>Prasad, M., Jang, A. C. C., Starz-Gaiano, M., Melani, M., Montell, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+protocol+for+culturing+Drosophila+melanogaster+stage+9+egg+chambers+for+live+imaging.">A protocol for culturing Drosophila melanogaster stage 9 egg chambers for live imaging.</a> Nature Protocols. 2 (10), 2467-2473 (2007).</li><li>Schermelleh, L., Heintzmann, R., Leonhardt, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+guide+to+super-resolution+fluorescence+microscopy.">A guide to super-resolution fluorescence microscopy.</a> Journal of Cell Biology. 190 (2), 165-175 (2010).</li><li>Sivaguru, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Comparative+performance+of+airyscan+and+structured+illumination+superresolution+microscopy+in+the+study+of+the+surface+texture+and+3D+shape+of+pollen.">Comparative performance of airyscan and structured illumination superresolution microscopy in the study of the surface texture and 3D shape of pollen.</a> Microscopy Research and Technique. 81 (2), 101-114 (2018).</li></ol>

The authors declare no conflict of interest.

Super-resolution microscopy methods provide spatial resolution as high as 10s of nanometers4,5,6. The STORM and PALM microscopy methods allow resolution up to 20 to 50 nm (XY-resolution), while STED microscopy offers resolution of 20 to 100 nm (XY-resolution). The spatial resolution of SIM microscopy is limited to 100 to 130 nm15. However, due to its high-photon density and lengthy acquisition time, it is extremely challenging to use these techniques for live cell imaging.
Imaging subcellular structures, such as the cytoskeleton, chromosomes, and organelles in live cells within tissues, requires the technique to be both ultrasensitive and noninvasive while still having a high spatial resolution. The Airyscan super-resolution technique presented here combines confocal imaging with a 0.2 Airy Unit pinhole, together with pixel reassignment and deconvolution. This technique can achieve a 1.7x higher resolution than that of conventional confocal microscopy7. The ~ 220-230 nm resolution of conventional confocal microscopy is restricted by the diffraction limit of light, but this imaging technique is able to achieve approximately 140 nm XY-resolution, which is comparable to other widely used super-resolution techniques, such as SIM (resolution at 110-120 nm)16.
The adaptability and versatility of Airyscan confocal microscopy will allow this protocol to be applicable to various other types of tissues and cells to study a diverse number of cell biology processes7,16. Using this technique, researchers can obtain time-lapse images at a super-resolution level to visualize cellular dynamics with unprecedented spatiotemporal detail.
The critical step in this technique is mounting the sample onto the dish so that it does not move or float during time-lapse imaging. Keeping the specimen close to or stuck to the imaging surface (i.e., the glass bottom of the dish) is critical for acquiring super-resolution images and maintaining the optimal microenvironment for the sample. This ensures that the sample is not stressed due to conditions such as hypoxia and changes in temperature or humidity.
Despite the usefulness of this protocol, there are a few caveats. Time-lapse imaging with the Airyscan mode is a relatively slow process compared to conventional confocal microscope imaging. Therefore, even the slightest physical shift of the sample could result in poor resolution. This situation can be prevented by using an immobilization method to adhere specimens to the imaging surface (e.g., poly L-lysine coating or other coating compatible with the sample). Also, placing a dialysis membrane on top of the tissue and adding a few small weights as described in this protocol helps prevent tissue movement or floating while also allowing for gaseous exchange. Time-lapse imaging cells that are located deep in the tissue with the super-resolution mode requires high illumination and can cause photobleaching. This can be minimized by adjusting the number of z-slices, illumination power, and averaging algorithm to achieve the optimal settings for the sample. If the cell or tissue is sensitive to laser toxicity, hypoxia, or any other stress during imaging, then cell cycle arrest may occur. Also, if a protein has an extremely low expression, short half-life, or low fluorescence signal, then photobleaching may occur which will result in an image with bad quality. This can be avoided by taking a SRLS, making a short movie, or shooting with long intervals between consecutive time points. Additionally, the microscope setting and the time-lapse imaging parameter (as described in this protocol) must be optimized to fit the sample.
Since it is essential that the sample does not move or degenerate during the process, prolonged time-lapse imaging lasting overnight or more than a day without compromising the image resolution may be challenging.
The existing methods mainly focus on optimizing the super-resolution microscopy and time-lapse imaging parameters separately1,9. However, these methods do not provide details regarding sample preparation or mounting parameters, which are critical to ensuring the viability of the sample and the adaptability of the microscope to achieve desired super-resolution during time-lapse imaging. In this method, we have optimized the imaging parameters, sample preparation and mounting parameters so that time-lapse imaging can be performed for an extended period of time to acquire super-resolution images without risking both the degeneration of the sample and the resolution.

Visualizing subcellular structures and protein dynamics in live cells with resolution beyond the diffraction limit of light is typically very challenging1-3. While multiple super-resolution techniques such as Stochastic-Optical-Reconstruction-Microscopy (STORM), Photo-Activated-Localization-Microscopy (PALM) and Stimulated-Emission-Depletion (STED)4,5,6 microscopy have been developed, complications in specimen preparation as well as the need to maintain viability and activity ex vivo, limit the use of conventional super-resolution microscopy for imaging live samples. Conventional confocal microscopy cannot reach spatial resolution beyond ~230 nm XY-resolution and is often insufficient to observe intricate cellular substructures5,6. However, a recent development in confocal microscopy, Airyscan super-resolution imaging, is able to achieve approximately 140 nm (XY-resolution)7,8 and has a relatively simple sample preparation that is compatible with live imaging. Since this imaging detection system requires a long acquisition time, its high spatial resolution does come at the cost of temporal resolution9. Therefore, a method is needed to ensure that live cell imaging is extended with high spatial resolution.
Here, we developed a method for observing live cells in intact tissue at its optimal resolution to decipher subcellular structures with detailed spatial information. This method is designed as such so that samples can be mounted stably for a long period of time (~ 10 h) without moving or degenerating. The live cell media used in this technique can support cellular function and avoid photobleaching for up to 10 hours under a super-resolution microscope. Finally, this protocol minimizes most stresses caused by the constant illumination of lasers over extended periods of time such as hypoxia, changes in humidity and temperature, as well as nutrient exhaustion.
Using this protocol to image Drosophila male germline stem cells (GSCs), we were able to observe how the asymmetric activity of microtubules allows for preferential interaction with epigenetically distinct sister chromatids10,11,12,13. These types of cellular events are highly dynamic and are very difficult to visualize in live cells using other super-resolution imaging methods such as STORM, PALM, or STED. We anticipate that this method will become highly useful for cell biologists as they aim to understand the dynamic subcellular structures of live cells residing in tissues. There are many areas in which this method can be applied to, such as studying the dynamics of proteins; understanding the movement of cells; and lineage tracing and cellular differentiation processes, among other possible applications.

<table><tbody><tr><td>Adobe Illustrator CS6 (figure making software)</td><td>Adobe</td><td>N/A</td><td></td></tr><tr><td>Dialysis membrane</td><td>Spectra/Por 1-4 Standard RC Dialysis Membrane</td><td>Cat No. 08-67121</td><td></td></tr><tr><td>FBS</td><td>Thermo Fisher Scientific</td><td>Cat. no. 26140079</td><td>15% (V/V)</td></tr><tr><td>Fiji (analysis software)</td><td>NIH</td><td>N/A</td><td>Image fluorescence intensity quantification</td></tr><tr><td>Glass bottom cell culture dishes (FluroDish)</td><td>World Precision Instrument, Inc.</td><td>FD35PDL-100</td><td></td></tr><tr><td>Imaris (image reconstruction software)</td><td>Bitplane</td><td>N/A</td><td>3D image reconstruction</td></tr><tr><td>Imerssion oil</td><td>Zeiss</td><td>Immersol 518F/30 &amp;deg;C</td><td></td></tr><tr><td>Insulin</td><td>Sigma</td><td>Cat. No. 15550</td><td>200 &amp;micro;g/ml</td></tr><tr><td>Penicillin/streptomycin</td><td>Invitrogen</td><td>Cat No. - 15140-122</td><td>0.6x</td></tr><tr><td>Schneider Drosophila media</td><td>Invitrogen</td><td>Cat No. - 11720-034</td><td></td></tr><tr><td>Spinning disc confocal microscope</td><td>Zeiss</td><td>N/A</td><td>equipped with an evolve camera (Photometrics), using a 63x Zeiss objective (1.4 NA).</td></tr><tr><td>Tissue paper</td><td>Kimwipe</td><td>N/A</td><td>Wet to form humid chamber</td></tr><tr><td>LSM 800 confocal microscope with AiryScan super-resolution module</td><td>Zeiss</td><td>N/A</td><td>equipped with highly sensitive GaAsP (Gallium Arsenide Phosphide) detectors using a 63x Zeiss objective (1.4 NA)</td></tr><tr><td>ZEN (imaging software)</td><td>Zeiss</td><td>N/A</td><td></td></tr></tbody></table>

super-resolution live cell imaging of subcellular structures

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally been restricted to be used only on fixed samples or live cells outside of tissue labeled with strong fluorescent signal. Current super-resolution live cell imaging techniques require the use of special fluorescence probes, high illumination, multiple image acquisitions with post-acquisition processing, or often a combination of these processes. These prerequisites significantly limit the biological samples and contexts that this technique can be applied to.
Here we describe a method to perform super-resolution (~140 nm XY-resolution) time-lapse fluorescence live cell imaging in situ. This technique is also compatible with low fluorescent intensity, for example, EGFP or mCherry endogenously tagged at lowly expressed genes. As a proof-of-principle, we have used this method to visualize multiple subcellular structures in the Drosophila testis. During tissue preparation, both the cellular structure and tissue morphology are maintained within the dissected testis. Here, we use this technique to image microtubule dynamics, the interactions between microtubules and the nuclear membrane, as well as the attachment of microtubules to centromeres.
This technique requires special procedures in sample preparation, sample mounting and immobilizing of specimens. Additionally, the specimens must be maintained for several hours after dissection without compromising cellular function and activity. While we have optimized the conditions for live super-resolution imaging specifically in Drosophila male germline stem cells (GSCs) and progenitor germ cells in dissected testis tissue, this technique is broadly applicable to a variety of different cell types. The ability to observe cells under their physiological conditions without sacrificing either spatial or temporal resolution will serve as an invaluable tool to researchers seeking to address crucial questions in cell biology.

Live cell imaging beyond the diffraction limit in Drosophila tissue, especially for GSCs, provides an opportunity to investigate the dynamics of subcellular events in the context of cell cycle progression. Recently, a study utilizing this protocol has shown that microtubule activities at the mother centrosome versus the daughter centrosome are temporally asymmetric in GSCs10. The mother centrosome emanates microtubules approximately 4 hours prior to the mitotic entry, whereas the daughter centrosome only emanates microtubules at the onset of mitosis. The highly active microtubules from the mother centrosome interact with the nuclear membrane and induce polarized Nuclear Envelope Break Down (NEBD). The local NEBD is proximal to the hub and allows microtubules from the mother centrosome to enter the nucleus and preferentially attach to the stronger sister kinetochore and sister centromere to ensure that they segregate to the future stem cell. These observations suggest that an asymmetric mitotic axis exists in which the microtubules, nuclear membrane, kinetochore, and centromeres interact in a spatiotemporally controlled manner, in order to ensure proper nonrandom chromosome segregation ultimately10,11,12.
Each of these results was supported using the SRLS approach outlined here. By performing live cell imaging of Drosophila testes expressing &#945;-tubulin-GFP in early-stage germ cells, we were able to identify temporal asymmetry in microtubule activity in GSCs (Figure 2). We were also able to see the asymmetric intensity of GFP signals at the two centrosomes as a brighter signal at the mother centrosome and a relatively weaker signal at the daughter centrosome side using a spinning disk confocal microscope (Figure 2A). The difference in brightness is likely reflected by the temporal asymmetry of microtubule nucleation, but the detailed morphology and quantity of microtubules could not be resolved using spinning disk confocal microscopy (Figure 2A). In contrast, live cell super-resolution imaging allowed us, to visualize the morphology and quantify the number of microtubules (Figure 2B). Its improved resolution revealed patterns of asymmetric microtubule nucleation, elongation, and increased interaction with the nuclear membrane (Figure 2B). As examples, the time-lapse movies of &#945;-tubulin-GFP-expressing GSCs have been published in a primary research paper (Videos S3 and Video S5)10.
Next, we performed live cell imaging on Drosophila testes co-expressing either &#945;-Tubulin-GFP with Lamin-mCherry or &#945;-Tubulin-mCherry with Lamin-GFP in early-stage germ cells. Our results show that asymmetric microtubule activity coincided with asymmetric NEBD in GSCs (Figure 3). Using the spinning-disk confocal microscope, we could visualize the asymmetric nuclear membrane invagination but not the individual microtubules that directly entered the nucleus (Figure 3A). In contrast, this live cell super-resolution technique allowed us to directly observe these events by imaging both the microtubules and the nuclear lamina simultaneously (Figure 3B).
Next, we performed live cell imaging on Drosophila testes expressing &#945;-Tubulin-mCherry and CENP-A-Dendra2 or CENP-A-GFP (Centromere Protein A) in early-stage germ cells. To better visualize the microtubule and centromere attachment, we live imaged them at a lower temperature (~18 &#730;C) to stabilize their attachment. If needed, the sample can be briefly chilled with ice or ice-cold live cell media for 4&#8211;5 minutes to stabilize the microtubule&#8211;centromere attachment and depolymerize any unattached microtubule. Our results show that the microtubules emanating from the early active mother centrosome preferentially attach to the stronger sister centromere (Figure 4). Using the spinning-disk confocal microscope, the &#945;-Tubulin-mCherry signals displayed an asymmetric brightness near the two centrosomes. We could also detect both sister centromeres as one signal, as shown in Figure 4A (metaphase), but were not able to visualize the microtubule&#8211;centromere attachment (Figure 4A, metaphase). In contrast, the super-resolution live imaging allowed us to visualize the microtubule-centromere attachment (Figure 4B). Our results show that mother centrosome-emanating microtubules attach to the centromere prior to the daughter centrosome-emanating microtubules (Figure 4B, early prophase).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61563/61563fig01.jpg" /> 
Figure 1: Scheme for the preparation and mounting of Drosophila testis.&#160;(A) Top view: the glass-bottom cell culture dish (A-a), transfer the Drosophila testis toward the center of the dish (A-b), place a dialysis membrane on the top of the testis (A-c), place three plastic weights on the dialysis membrane (A-d). (B) Lateral view: place the plastic ring at the elevated side of the dish (B-a), place a 22 mm &#215; 22 mm coverslip on the plastic ring (B-b), put a swirl and wet tissue paper on the coverslip (B-c), close the dish with the lid (B-d). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/61563/61563fig01large.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/61563/61563fig02.jpg" /> 
Figure 2: Super-resolution time-lapse imaging of microtubule dynamics in Drosophila tissue (testes) expressing &#945;-tubulin-GFP&#160;(A) Conventional confocal microscopy live cell image showing microtubule dynamics in mid G2 phase to metaphase. (B) Airyscan microscope live cell image showing asymmetric microtubule emanating from mother versus daughter centrosomes in mid G2 phase to metaphase, with detailed structural information. A cartoon depicts the Drosophila male germline stem cell. Scale bar: 5&#181;m; asterisk: hub (niche); green arrow: mother centrosome (stem cell side [M]); red arrow: daughter centrosome (differentiating daughter cell side [D]); yellow arrow: centrosome in spermatogonia cell (progenitor non-stem germ cell); magenta arrow: microtubule poking in nucleus. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/61563/61563fig02large.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/61563/61563fig03.jpg" /> 
Figure 3: Super-resolution time-lapse imaging of microtubule poking-in activity and asymmetric nuclear envelope breakdown in Drosophila tissue (testes) co-expressing Lamin-mCherry and &#945;-Tubulin-GFP.&#160;(A) Conventional confocal microscopy live cell image showing microtubule dynamics from G2-M phase to mitosis. Image showing higher &#945;-Tubulin-GFP intensity at the stem cell side and its interaction with the nuclear membrane. (B) Airyscan microscope live cell image showing microtubule poking-in and asymmetric NEBD at the stem cell side. Scale bar: 5&#181;m; asterisk: hub (niche); orange arrow: site of microtubule poking in nuclear membrane; magenta arrow: microtubules that poke in at stem cell side. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/61563/61563fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/61563/61563fig04.jpg" /> 
Figure 4: Super-resolution time-lapse imaging of microtubule and centromere attachment in Drosophila tissue (testes) expressing &#945;-Tubulin-mCherry and CENP-A-Dendra2.&#160;(A) Conventional confocal microscopy live cell image showing microtubule and centromere interaction from Early prophase to metaphase. Image showing higher &#945;-Tubulin-mCherry intensity at stem cell side and CENP-A-GFP signal. (B) Airyscan microscope live cell image showing microtubules emanating from the mother centrosome and attachment to the stronger centromeres in early prophase. At metaphase, centromeres are attached to opposite pole (bi-orientation). Scale bar: 5&#181;m; asterisk: hub (niche); magenta arrow: kinetochore fiber or K-fiber (microtubules attached to the centromere). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/61563/61563fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Super-Resolution Live Cell Imaging of Subcellular Structures at JoVE.com

Super-Resolution Live Cell Imaging of Subcellular Structures

Presented here is a protocol for super-resolution live-cell imaging in intact tissue. We have standardized the conditions for imaging a highly sensitive adult stem cell population in its native tissue environment. This technique involves balancing temporal and spatial resolution to allow for the direct observation of biological phenomena in live tissue.

There has long been a crucial tradeoff between spatial and temporal resolution in imaging. Imaging beyond the diffraction limit of light has traditionally ...

super-resolution-live-cell-imaging-of-subcellular-structures

Universidad San Sebastian

Research

JoVE Journal

Biology

4.7K Views.  The Johns Hopkins University. Presented here is a protocol for super-resolution live-cell imaging in intact tissue. We have standardized the conditions for imaging a highly sensitive adult stem cell population in its native tissue environment. This technique involves balancing temporal and spatial resolution to allow for the direct observation of biological phenomena in live tissue.

Video: Super-Resolution Live Cell Imaging of Subcellular Structures

Imaging Subcellular Structures in the Living Zebrafish Embryo

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such imaging experiments in higher vertebrates such as mammals generally requires surgical access to the system under study. The optical accessibility of embryonic and larval zebrafish allows such invasive procedures to be circumvented and permits imaging in the intact organism. Indeed the zebrafish is now a well-established model to visualize dynamic cellular behaviors using in vivo microscopy in a wide range of developmental contexts from proliferation to migration and differentiation. A more recent development is the increasing use of zebrafish to study subcellular events including mitochondrial trafficking and centrosome dynamics. The relative ease with which these subcellular structures can be genetically labeled by fluorescent proteins and the use of light microscopy techniques to image them is transforming the zebrafish into an in vivo model of cell biology. Here we describe methods to generate genetic constructs that fluorescently label organelles, highlighting mitochondria and centrosomes as specific examples. We use the bipartite Gal4-UAS system in multiple configurations to restrict expression to specific cell-types and provide protocols to generate transiently expressing and stable transgenic fish. Finally, we provide guidelines for choosing light microscopy methods that are most suitable for imaging subcellular dynamics.

Imaging the dynamic behavior of organelles and other subcellular structures in vivo can shed light on their function in physiological and disease conditions. Here, we present methods for genetically tagging two organelles, centrosomes and mitochondria, and imaging their dynamics in living zebrafish embryos using wide-field and confocal microscopy.

In vivo imaging provides unprecedented access to the dynamic behavior of cellular and subcellular structures in their natural context. Performing such ...

Developmental Biology

Super-resolution Imaging of the Natural Killer Cell Immunological Synapse on a Glass-supported Planar Lipid Bilayer

The glass-supported planar lipid bilayer system has been utilized in a variety of disciplines. One of the most useful applications of this technique has been in the study of immunological synapse formation, due to the ability of the glass-supported planar lipid bilayers to mimic the surface of a target cell while forming a horizontal interface. The recent advances in super-resolution imaging have further allowed scientists to better view the fine details of synapse structure. In this study, one of these advanced techniques, stimulated emission depletion (STED), is utilized to study the structure of natural killer (NK) cell synapses on the supported lipid bilayer. Provided herein is an easy-to-follow protocol detailing: how to prepare raw synthetic phospholipids for use in synthesizing glass-supported bilayers; how to determine how densely protein of a given concentration occupies the bilayer's attachment sites; how to construct a supported lipid bilayer containing antibodies against NK cell activating receptor CD16; and finally, how to image human NK cells on this bilayer using STED super-resolution microscopy, with a focus on distribution of perforin positive lytic granules and filamentous actin at NK synapses. Thus, combining the glass-supported planar lipid bilayer system with STED technique, we demonstrate the feasibility and application of this combined technique, as well as intracellular structures at NK immunological synapse with super-resolution.

We describe here a combination of the glass-supported lipid bilayer technique of forming immunological synapses with the super-resolution imaging technique of stimulated emission depletion (STED) microscopy. The goal of this protocol is to provide users with the instructions necessary to successfully carry out these two techniques.

The glass-supported planar lipid bilayer system has been utilized in a variety of disciplines. One of the most useful applications of this technique has ...

Immunology and Infection

A Guide to Structured Illumination TIRF Microscopy at High Speed with Multiple Colors

Optical super-resolution imaging with structured illumination microscopy (SIM) is a key technology for the visualization of processes at the molecular level in the chemical and biomedical sciences. Although commercial SIM systems are available, systems that are custom designed in the laboratory can outperform commercial systems, the latter typically designed for ease of use and general purpose applications, both in terms of imaging fidelity and speed. This article presents an in-depth guide to building a SIM system that uses total internal reflection (TIR) illumination and is capable of imaging at up to 10 Hz in three colors at a resolution reaching 100 nm. Due to the combination of SIM and TIRF, the system provides better image contrast than rival technologies. To achieve these specifications, several optical elements are used to enable automated control over the polarization state and spatial structure of the illumination light for all available excitation wavelengths. Full details on hardware implementation and control are given to achieve synchronization between excitation light pattern generation, wavelength, polarization state, and camera control with an emphasis on achieving maximum acquisition frame rate. A step-by-step protocol for system alignment and calibration is presented and the achievable resolution improvement is validated on ideal test samples. The capability for video-rate super-resolution imaging is demonstrated with living cells.

This article provides an in depth guide for the assembly and operation of a structured illumination microscope operating with total internal reflection fluorescence illumination (TIRF-SIM) to image dynamic biological processes with optical super-resolution in multiple colors.

Optical super-resolution imaging with structured illumination microscopy (SIM) is a key technology for the visualization of processes at the molecular ...

Bioengineering

Visualization of Endosome Dynamics in Living Nerve Terminals with Four-dimensional Fluorescence Imaging

Four-dimensional (4D) light imaging has been used to study behavior of small structures within motor nerve terminals of the thin transversus abdominis muscle of the garter snake. Raw data comprises time-lapse sequences of 3D z-stacks. Each stack contains 4-20 images acquired with epifluorescence optics at focal planes separated by 400-1,500 nm. Steps in the acquisition of image stacks, such as adjustment of focus, switching of excitation wavelengths, and operation of the digital camera, are automated as much as possible to maximize image rate and minimize tissue damage from light exposure. After acquisition, a set of image stacks is deconvolved to improve spatial resolution, converted to the desired 3D format, and used to create a 4D "movie" that is suitable for variety of computer-based analyses, depending upon the experimental data sought. One application is study of the dynamic behavior of two classes of endosomes found in nerve terminals-macroendosomes (MEs) and acidic endosomes (AEs)-whose sizes (200-800 nm for both types) are at or near the diffraction limit. Access to 3D information at each time point provides several advantages over conventional time-lapse imaging. In particular, size and velocity of movement of structures can be quantified over time without loss of sharp focus. Examples of data from 4D imaging reveal that MEs approach the plasma membrane and disappear, suggesting that they are exocytosed rather than simply moving vertically away from a single plane of focus. Also revealed is putative fusion of MEs and AEs, by visualization of overlap between the two dye-containing structures as viewed in each three orthogonal projections.

Four-dimensional (4D) imaging is utilized to study the behavior and interactions among two types of endosomes in living vertebrate nerve terminals. Movement of these small structures is characterized in three dimensions, permitting confirmation of events such as endosome fusion and exocytosis.

Four-dimensional (4D) light imaging has been used to study behavior of small structures within motor nerve terminals of the thin transversus abdominis ...

Neuroscience

Three-dimensional Imaging of Bacterial Cells for Accurate Cellular Representations and Precise Protein Localization

The shape of a bacterium is important for its physiology. Many aspects of cell physiology such as cell motility, predation, and biofilm production can be affected by cell shape. Bacterial cells are three-dimensional (3D) objects, although they are rarely treated as such. Most microscopy techniques result in two-dimensional (2D) images leading to the loss of data pertaining to the actual 3D cell shape and localization of proteins. Certain shape parameters, such as Gaussian curvature (the product of the two principal curvatures), can only be measured in 3D because 2D images do not measure both principal curvatures. Additionally, not all cells lie flat when mounting and 2D imaging of curved cells may not accurately represent the shapes of these cells. Accurately measuring protein localization in 3D can help determine the spatial regulation and function of proteins. A forward convolution technique has been developed that uses the blurring function of the microscope to reconstruct 3D cell shapes and to accurately localize proteins. Here, a protocol for preparing and mounting samples for live cell imaging of bacteria in 3D both to reconstruct an accurate cell shape and to localize proteins is described. The method is based on simple sample preparation, fluorescent image acquisition, and MATLAB-based image processing. Many high-quality fluorescent microscopes can be simply modified to take these measurements. These cell reconstructions are computationally intensive and access to high-throughput computational resources is recommended, although not necessary. This method has been successfully applied to multiple bacterial species and mutants, fluorescent imaging modalities, and microscope manufacturers.

This protocol explains how to prepare and mount bacterial samples for live three-dimensional imaging and how to reconstruct the three-dimensional shape of E. coli from those images.

The shape of a bacterium is important for its physiology. Many aspects of cell physiology such as cell motility, predation, and biofilm production can be ...

Super-resolution Imaging of the Bacterial Division Machinery

Bacterial cell division requires the coordinated assembly of more than ten essential proteins at midcell1,2. Central to this process is the formation of a ring-like suprastructure (Z-ring) by the FtsZ protein at the division plan3,4. The Z-ring consists of multiple single-stranded FtsZ protofilaments, and understanding the arrangement of the protofilaments inside the Z-ring will provide insight into the mechanism of Z-ring assembly and its function as a force generator5,6. This information has remained elusive due to current limitations in conventional fluorescence microscopy and electron microscopy. Conventional fluorescence microscopy is unable to provide a high-resolution image of the Z-ring due to the diffraction limit of light (~200 nm). Electron cryotomographic imaging has detected scattered FtsZ protofilaments in small C. crescentus cells7, but is difficult to apply to larger cells such as E. coli or B. subtilis. Here we describe the application of a super-resolution fluorescence microscopy method, Photoactivated Localization Microscopy (PALM), to quantitatively characterize the structural organization of the E. coli Z-ring8.
	PALM imaging offers both high spatial resolution (~35 nm) and specific labeling to enable unambiguous identification of target proteins. We labeled FtsZ with the photoactivatable fluorescent protein mEos2, which switches from green fluorescence (excitation = 488 nm) to red fluorescence (excitation = 561 nm) upon activation at 405 nm9. During a PALM experiment, single FtsZ-mEos2 molecules are stochastically activated and the corresponding centroid positions of the single molecules are determined with &lt;20 nm precision. A super-resolution image of the Z-ring is then reconstructed by superimposing the centroid positions of all detected FtsZ-mEos2 molecules.
	Using this method, we found that the Z-ring has a fixed width of ~100 nm and is composed of a loose bundle of FtsZ protofilaments that overlap with each other in three dimensions. These data provide a springboard for further investigations of the cell cycle dependent changes of the Z-ring10 and can be applied to other proteins of interest.

We describe a super-resolution imaging method to probe the structural organization of the bacterial FtsZ-ring, an essential apparatus for cell division. This method is based on quantitative analyses of photoactivated localization microscopy (PALM) images and can be applied to other bacterial cytoskeletal proteins.

Bacterial cell division requires the coordinated  assembly of more than ten essential proteins at midcell1,2. Central  to this process is the formation of ...

Super-resolution Imaging of the Cytokinetic Z Ring in Live Bacteria Using Fast 3D-Structured Illumination Microscopy (f3D-SIM)

Imaging of biological samples using fluorescence microscopy has advanced substantially with new technologies to overcome the resolution barrier of the diffraction of light allowing super-resolution of live samples. There are currently three main types of super-resolution techniques &#8211; stimulated emission depletion (STED), single-molecule localization microscopy (including techniques such as PALM, STORM, and GDSIM), and structured illumination microscopy (SIM). While STED and single-molecule localization techniques show the largest increases in resolution, they have been slower to offer increased speeds of image acquisition. Three-dimensional SIM (3D-SIM) is a wide-field fluorescence microscopy technique that offers a number of advantages over both single-molecule localization and STED. Resolution is improved, with typical lateral and axial resolutions of 110 and 280 nm, respectively and depth of sampling of up to 30 &#181;m from the coverslip, allowing for imaging of whole cells. Recent advancements (fast 3D-SIM) in the technology increasing the capture rate of raw images allows for fast capture of biological processes occurring in seconds, while significantly reducing photo-toxicity and photobleaching. Here we describe the use of one such method to image bacterial cells harboring the fluorescently-labelled cytokinetic FtsZ protein to show how cells are analyzed and the type of unique information that this technique can provide.

Super-resolution Imaging of the Cytokinetic Z Ring in Live Bacteria Using Fast 3D-Structured Illumination Microscopy f3D-SIM

Spatiotemporal information about dynamic proteins inside live cells is crucial for understanding biology. A type of super-resolution microscopy called fast 3D-structured illumination microscopy (f3D-SIM) reveals unique information about the cytokinetic Z ring in bacteria: both its bead-like appearance and the rapid dynamics of FtsZ within the ring.

Imaging of biological samples using fluorescence microscopy has advanced substantially with new technologies to overcome the resolution barrier of the ...

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

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

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

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

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

Application of High-speed Super-resolution SPEED Microscopy in Live Primary Cilium

The primary cilium is a microtubule-based protrusion on the surface of many eukaryotic cells and contains a unique complement of proteins that function critically in cell motility and signaling. Since cilia are incapable of synthesizing their own protein, nearly 200 unique ciliary proteins need to be trafficked between the cytosol and primary cilia. However, it is still a technical challenge to map three-dimensional (3D) locations of transport pathways for these proteins in live primary cilia due to the limitations of currently existing techniques. To conquer the challenge, recently we have developed and employed a high-speed virtual 3D super-resolution microscopy, termed single-point edge-excitation sub-diffraction (SPEED) microscopy, to determine the 3D spatial location of transport pathways for both cytosolic and membrane proteins in primary cilia of live cells. In this article, we will demonstrate the detailed setup of SPEED microscopy, the preparation of cells expressing fluorescence-protein-labeled ciliary proteins, the real-time single-molecule tracking of individual proteins in live cilium and the achievement of 3D spatial probability density maps of transport routes for ciliary proteins.

Recently we mapped the three-dimensional (3D) spatial locations of transport routes for various proteins translocating inside primary cilia in live cells. Here this paper details the experimental setup, the process of biological samples and the data analyses for the 3D super-resolution fluorescence imaging approach newly applied in live primary cilia.

The primary cilium is a microtubule-based protrusion on the surface of many eukaryotic cells and contains a unique complement of proteins that function ...

Super-Resolution Microscopy of the Synaptonemal Complex Within the Caenorhabditis elegans Germline

During meiosis, homologous chromosomes must recognize and adhere to one another to allow for their correct segregation. One of the key events that secures the interaction of homologous chromosomes is the assembly of the synaptonemal complex (SC) in meiotic prophase I. Even though there is little sequence homology between protein components within the SC among different species, the general structure of the SC has been highly conserved during evolution. In electron micrographs, the SC appears as a tripartite, ladder-like structure composed of lateral elements or axes, transverse filaments, and a central element. 
However, precisely identifying the localization of individual components within the complex by electron microscopy to determine the molecular structure of the SC remains challenging. By contrast, fluorescence microscopy allows for the identification of individual protein components within the complex. However, since the SC is only ~100 nm wide, its substructure cannot be resolved by diffraction-limited conventional fluorescence microscopy. Thus, determining the molecular architecture of the SC requires super-resolution light microscopy techniques such as structured illumination microscopy (SIM), stimulated-emission depletion (STED) microscopy, or single-molecule localization microscopy (SMLM). 
To maintain the structure and interactions of individual components within the SC, it is important to observe the complex in an environment that is close to its native environment in the germ cells. Therefore, we demonstrate an immunohistochemistry and imaging protocol that enables the study of the substructure of the SC in intact, extruded Caenorhabditis elegans germline tissue with SMLM and STED microscopy. Directly fixing the tissue to the coverslip reduces the movement of the samples during imaging and minimizes aberrations in the sample to achieve the high resolution necessary to visualize the substructure of the SC in its biological context.

Super-Resolution Microscopy of the Synaptonemal Complex Within the Caenorhabditis elegans Germline

Super-resolution microscopy can provide a detailed insight into the organization of components within the synaptonemal complex in meiosis. Here, we demonstrate a protocol to resolve individual proteins of the Caenorhabditis elegans synaptonemal complex.

During meiosis, homologous chromosomes must recognize and adhere to one another to allow for their correct segregation. One of the key events that secures ...

Super-Resolution Live Cell Imaging of Subcellular Structures

Summary

Explore More Videos

Chapters in this video

Imaging Subcellular Structures in the Living Zebrafish Embryo

Super-resolution Imaging of the Natural Killer Cell Immunological Synapse on a Glass-supported Planar Lipid Bilayer

A Guide to Structured Illumination TIRF Microscopy at High Speed with Multiple Colors

Visualization of Endosome Dynamics in Living Nerve Terminals with Four-dimensional Fluorescence Imaging

Three-dimensional Imaging of Bacterial Cells for Accurate Cellular Representations and Precise Protein Localization

Super-resolution Imaging of the Bacterial Division Machinery

Super-resolution Imaging of the Cytokinetic Z Ring in Live Bacteria Using Fast 3D-Structured Illumination Microscopy (f3D-SIM)

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

Application of High-speed Super-resolution SPEED Microscopy in Live Primary Cilium

Super-Resolution Microscopy of the Synaptonemal Complex Within the Caenorhabditis elegans Germline