We thank Dr. Kristen Verhey (University of Michigan, Ann Arbor) and Dr. Gregory Pazour (University of Massachusetts Medical School) for providing some plasmids. The project was supported by grants from the National Institutes of Health (NIH GM097037, GM116204 and GM122552 to W.Y.).

1. NIH-3T3 cell preparation for SPEED microscopy from stock
<ol>
	<li>1.5 weeks in advance of the experiment, recover a fresh culture of NIH-3T3 cells from a frozen stock by thawing at 37 &#176;C and transferring the cells to a 25 cm2 cell culture flask with 3 mL of Dulbecco&#39;s Modified Eagle&#39;s Medium (DMEM) supplemented with 110 mg/mL sodium pyruvate, 2 mM glutamine, 10% fetal bovine serum, and 1% penicillin/streptomycin.</li>
	<li>Incubate the cells at 37 &#176;C in a 5% CO2 incubator.</li>
	<li>Split cells at 80% confluency, about every two days, at least three times before experimental day to ensure homogeneity of cell cycle. Trypsinize cells with 0.25% trypsin for 2 min at 37 &#176;C, aspirate trypsin and replace it with 2 mL of medium. Pipette the medium repeatedly to break up cell clusters, remove the desired number of cells and bring total volume of media back up to 3 mL. 
	NOTE: NIH-3T3 were previously genetically engineered to express NPHP-4, a protein that localizes to the TZ26, fused at the C terminus to mCherry. mCherry is a fluorophore which can be excited with 561 nm illumination to quantitatively localize the TZ selectivity barrier and orient the primary cilia.</li>
	<li>Two days before the experiment, plate the cells into a 35 mm glass bottom dish at 60-70% confluency with 1.5 mL of the same medium as step 1.1 and return the cells to the incubator.</li>
	<li>One day before the experiment, chemically transfect the cells with the desired plasmid. Mix 500 - 1000 ng of the desired plasmid (see the Note below) in a 1:2.5 ratio with transfection reagent in 0.25 mL of reduced serum media without antibiotics for 30 min. Aspirate media from the 35 mm glass bottom dish and replace it with the 0.25 mL plasmid/transfection reagent mix plus an extra 1.25 mL of reduced serum media without antibiotics. Reduced serum media serves the purpose facilitating a successful transfection, inducing primary cilia growth, as well as keeping the cells alive long enough to perform experiment. Return cells to the incubator for the experiment on the following day. 
	NOTE: When performing single-molecule tracking of IFT20, a plasmid containing a genetically-modified IFT20 fused at its C terminus to GFP is used25. When performing single-molecule tracking of SSTR3, a plasmid containing a genetically-modified SSTR3 fused at its N terminus to an acceptor peptide (AP) domain and C terminus to GFP is used22. In addition to the SSTR3 construct, a plasmid containing the biotin ligase BirA must be co-expressed and the transfection media must be supplemented with 10 &#181;M biotin. BirA then attaches biotin to the AP domain of newly synthesized AP-SSTR3-GFP molecules at the level of the ER. Alexa647 conjugated to three of the four biotin-binding sites on streptavidin, on average, may then be supplemented to the media prior to imaging to fluorescently label the AP-SSTR3-GFP molecules on the external surface of the cell22,27. GFP and AlexaFluor647 are used in this method; however, other fluorescent probes can be used if they have similarly high photo-stability and quantum yield.</li>
	<li>If using the externally-labeled SSTR3 construct, remove the media from the glass bottom dish 1 h before experiment, wash the cell 5 times with 1 mL of phosphate-buffered saline (PBS), and add 1 mL of reduced serum media supplemented with 1 &#181;M Alexa647 conjugated streptavidin.</li>
	<li>No more than 15 min before the experiment, remove media from the glass bottom dish and wash the transfected and labeled cells 5 times with 1 mL of PBS.</li>
	<li>Place 1 mL of imaging buffer (20 mM HEPES, 110 mM KOAc, 5 mM NaOAc, 2 mM MgOAc, 1 mM EGTA, pH 7.3) in the glass bottom dish. 
	NOTE: In the imaging buffer, cells are viable for no longer than 3 h. Therefore, only 2 h of experiments are performed on each dish.</li>
</ol>
2. SPEED microscopy
Note: The SPEED microscopy setup includes an inverted fluorescence microscope equipped with a 1.4-NA 100&#215; oil-immersion apochromatic objective, a 35 mW 633 nm He-Ne laser, 50 mW solid state 488-nm and 561-nm lasers, an on-chip multiplication gain charge-coupled-device camera and a microscope software package for data acquisition and processing (Figure 1). For individual channel imaging, GFP, mCherry, and Alexa647 are excited by 488 nm, 561 nm, or 633 nm lasers, respectively. For single molecule tracking, single point illumination is used to track individual fluorescently-labeled molecules. For epifluorescence imaging, a concave lens is placed in the laser illumination path to expand the beam into a uniform field of illumination. The fluorescence emissions are collected by the same objective, filtered by a dichroic filter (405/488/561/635) and an emission filter (405/488/561/635), and imaged with the above CCD camera operating at 500 Hz for single molecule tracking or 2 Hz for epifluorescence imaging.
<ol>
	<li>Affix the glass bottom plate to the stage of the microscope and locate a cell that is properly expressing the desired constructs. Once a suitable cell has been found, align the NPHP4-mCherry spot at the base of the primary cilia with the location on the imaging plane that corresponds to the laser&#39;s single point illumination.</li>
	<li>Capture an epifluorescence image of NPHP4-mCherry and either IFT20-GFP or AP-SSTR3-GFP using the &#34;Snap&#34; function in the &#34;Camera&#34; tab of the &#34;Focus Controls&#34; window if using the digital microscopy software package (see Table of Materials). 
	NOTE: These images will act as a reference for the subsequent single molecule locations.</li>
	<li>Once the reference images are obtained, locally reduce the concentration of labeled single molecules. Photo-bleach the TZ with 1 mW laser illumination for 20 s or until the fluorescence intensity is close to that of background fluorescence. 
	NOTE: When the precise concentration can be controlled, 0.1-1 nM labeled single molecules are used.</li>
	<li>To prepare for single molecule tracking, reduce the laser illumination power to ~0.15 mW for single molecules labeled with GFP or ~0.5 mW for molecules labeled with Alexa647.</li>
	<li>As soon as the laser power and imaging parameters are set, maximum gain and intensification and 2 ms frame rate, for single molecule imaging, engage the appropriate illumination laser and record non-photobleached, labeled single molecules as they are transported through the photobleached region of the TZ by clicking the &#34;Stream&#34; button in the &#34;Camera&#34; tab of the &#34;Focus Controls&#34; window. 
	NOTE: No more than 2 min of video should be captured to minimize the effects of ciliary drift to a negligible level.</li>
	<li>After capturing the single molecule video, process the videos using a 2D Gaussian fitting algorithm, such as Glimpse by the Gelles Lab, which precisely localizes the centroid of each single molecule&#39;s excitation PSF in an encompassing area of interest (AOI).</li>
	<li>Select all single molecule locations with precision &#60;10 nm and correct the center of cilia based on distribution of single molecule locations fitted with a 2D Gaussian function. 
	NOTE: By using 2D to 3D transformation algorithm, the 3D transport routes of IFT20-GFP and AP-SSTR3 routes are clearly shown on ciliary axonemal or ciliary membrane, respectively.</li>
</ol>
3. 2D to 3D Transformation
<ol>
	<li>Once several thousand localizations for transiting molecules (signal to noise ratio &#62; 11) in the cilium are collected, select the long axis of the cilium as the X-dimension. Make a Y dimension histogram of the locations and obtain the bin sums in 10 nm increments. 
	NOTE: The 2D to 3D transformation may be evaluated by hand or any software or programming language. The authors have successfully implemented the transformation in both Matlab and Python 2.7.</li>
</ol>

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The authors declare no conflicts of interest.

This protocol describes the application of SPEED microscopy to the primary cilium, a cellular signaling organelle that is highly reliant on efficient protein transport. SPEED microscopy can provide high resolution (&#60; 10 nm) locations for fluorescently-labeled molecules as they pass through the single point illumination centered on the TZ. Previously it has been applied to study the protein trafficking through the NPC6,7,8. However, it may be extended to study trafficking through any sub-diffraction cellular cavity. This technique has an advantage over other high-resolution imaging techniques in that it is able to capture the dynamics of single protein transport in live cell systems at a spatial and temporal resolution of &#60;10 nm and &#60;2 ms, respectively. Another advantage is that one must only fluorescently-label a protein of interest and express the construct via transfection, a common molecular biology approach to studying protein localization, to begin performing SPEED microscopy. One must keep in mind that the single point illumination with a high-power laser is the feature that provides the well-separated, high resolution single molecule locations. However, this also limits the illumination area and makes the technique not suitable for wide field localization. Fortunately, there are many wide field super resolution techniques for investigators to choose from if desired. Another point to consider is that SPEED microscopy only captures locations of moving single molecules. Therefore, combination with FRAP, which can determine the fraction of total molecules that are mobile, may be a useful addition to analysis.
The conversion process between the Cartesian and the cylindrical coordination system is to generate a virtual 3D probability density map rather than a 3D view based on 3D single-molecule tracking. In detail, electron microscopy data have revealed that the primary cilia have a rotationally symmetrical structure which could generate a uniform molecule distribution along certain radius. This uniform distribution leads that the spatial distribution along &#952;&#160;dimension in the cylindrical system is constant. Then the 3D coordinates (R, X, &#952;) can be simplified to be the 2D coordinates (R, X, constant). Actually, our transform process between the Cartesian and the cylindrical systems is from 2D (X, Y) to 2D (R, X, constant). The constant &#952;, refers to the spatial density p, is calculated by using the equation A<img alt="Equation 3" src="/files/ftp_upload/56475/56475eq3.jpg" />(Figure 2). Use a matrix calculator to calculate the vector <img alt="Equation 4" src="/files/ftp_upload/56475/56475eq4.jpg" />, which corresponds with the relative areas between neighboring concentric rings in the cross-section view6,7&#160;Aj s the total interaction sites at the position j measured directly from experiments; Thus, pi, the spatial probability density of the interaction sites at ri, can be calculated by solving the matrix equation of A<img alt="Equation 3" src="/files/ftp_upload/56475/56475eq3.jpg" />.
A critical step in the protocol is to minimize any perturbation of the microscopy setup between the reference image acquisition and single molecule video. If any movement occurs, it is not possible to confidently map the single molecule locations to the ultrastructure of the primary cilia that was obtained via epifluorescence imaging. Another essential step is to photobleach the area of illumination enough to locally reduce the concentration of fluorescently labeled single molecules but not so much that the population is completely photobleached. In the case of SSTR3, the diffusion coefficient is slow compared to soluble molecules and the movement of a population of un-photobleached SSTR3 molecules back into the photobleached area will be longer than two minutes, the time beyond which the ciliary drift is a non-negligible factor. If the proper level of photobleaching is difficult to attain and a high level of background fluorescence is still present, an angled illumination laser may be used to reduce the amount of fluorescently labeled single molecules that are excited above and below the imaging plane. This will reduce the background and improve the signal-to-noise ratio for each captured single molecule.
In summary, SPEED microscopy is able to be readily implemented on conventional microscopy setups by the addition of a laser illumination source, associated mirrors, and high speed CCD camera. The main advancement over other similar techniques are the inclined laser which greatly reduces the background fluorescence and the 2D-to-3D transformation algorithm which reconstructs the 3D spatial probability density map from the 2D single-molecule locations. From a biological standpoint, no extra labeling besides a fluorophore-conjugated protein of interest is needed. These features allow SPEED microscopy to track single molecules with high spatiotemporal (5-10 nm, 0.4-2 ms) resolution as they traffic through a sub-micrometer bio-cavity or bio-channel with rotationally symmetric structures. Coupled with a 3D probability density transformation algorithm, the 3D transport routes of the tagged proteins may be determined through the cavity or channel. The application of this technique has been previously used to distinguish 3D transport routes of proteins and RNAs through the NPC and, here, we show that the 3D transport of a transmembrane protein, SSTR3, and a cytosolic protein, IFT20, may be achieved in the TZ of primary cilia. Other super resolution techniques, such as 3D-STORM, have obtained x,y,z positions for single molecules by placing a cylindrical lens in the optical path which creates an asymmetrical distortion of the single-molecule PSF depending on its position above or below the focal plane28. Another advance may attempt to implement virtual pinhole technology to reduce the width of the emission PSF and thus increase resolution even further. The above techniques could be implemented on SPEED microscopy quite readily. Overall, SPEED microscopy provides a unique approach in simultaneously determining transport kinetics at single-molecule level and 3D map of transport pathways with super-high spatiotemporal resolution for inter- or intra-organelle molecular trafficking under certain circumstances in live cell systems.

Since stated by Ernst Abbe in 1873, the resolution of conventional light microscopy has been believed to be limited to approximately 200 nm due to light diffraction from the objective1,2. Currently, super-resolution light microscopy techniques break this limitation and allow the capture of dynamic images with sub-diffraction (&#60; 200 nm) resolution. The techniques generally fall into two broad categories: stimulated emission depletion (STED) microscopy based approaches, which generate sub-diffraction illumination volume due to nonlinear optical response of fluorophores in samples3; and photoactivated light microscopy (PALM) and stochastic optical reconstruction microscopy (STORM)-based super-resolution techniques, which utilize mathematical functions to localize the centroids of fluorophores and then reconstitute these centroids to form super-resolution images4,5. Currently, due to the relatively uncomplicated optical setup, PALM and STORM are extensively employed by only activating a small subset of fluorophores in each frame of a long video of a biological preparation. This allows for the more accurate localization by 2D Gaussian fitting of the fluorescent spot, termed the point spread function (PSF), of fluorescently-labeled proteins in each frame of the video. The 2D location of each fluorescently-labeled molecule can then be superimposed on a single imaging plane to produce a super-resolution image of the biological preparation1,2. While these single-molecule localization, super-resolution approaches to microscopy certainly revolutionized how imaging of biological samples was performed, there are still challenges to be overcome. For example, STORM and PALM can achieve their best spatial resolutions after fixation of biological samples and thus present a static representation of the fluorescently-labeled proteins, which is a similar limitation of electron microscopy. Additionally, to achieve high spatial resolution for each fluorescently-labeled protein in live cells, samples must be imaged at very long framerates which are unable to capture protein dynamics. Therefore, it is necessary to overcome these main technical hurdles.
To obtain a high spatiotemporal resolution that is well-suited for detecting fast-moving proteins or RNAs in live cells, we have developed super-resolution SPEED microscopy in our laboratory (Figure 1)6,7,8. Several major technical advances in SPEED microscopy have previously enabled us to successfully track nucleocytoplasmic transport of small molecules, proteins, mRNA and virus through native nuclear pore complexes (NPCs)6,7,8. Briefly, the following features of SPEED microscopy will be used to track fast-moving macromolecules through sub-micrometer rotationally symmetrical structures in live cells, such as NPCs and primary cilia: (1) An inclined or a vertical illumination PSF enables the excitation of single molecules within a small diffraction-limit volume in the focal plane (Figure 1); (2) The inclined PSF can greatly avoid out-of-focus fluorescence and thus improve the signal-to-noise ratio. (3) The optical density of 100-500 kW/ cm2 in the illumination PSF allows thousands of photons to be collected from single fluorophores with fast detection speeds (&#62; 500 Hz). (4) The fast detection speed also greatly reduces the single-molecule spatial localization error (&#60; 10 nm) in determining the spatial trajectories of moving fluorescent molecules in live cells, because molecular diffusion is one of major factors causing imperfections of single-molecule localization for moving molecules. (5) Well-established 2D to 3D transformation algorithms enable us to provide 3D spatial probability density maps of transport routes for molecules in the NPC or the primary cilium. It is noteworthy that our conversion process between the Cartesian and the cylindrical coordination system is used to generate a 3D spatial probability density map rather than 3D single-molecule tracking (Figure 2). Previously, electron microscopy data have revealed that the NPC9,10 and the primary cilium11 both have a rotationally symmetrical structure. In principle, randomly diffusing molecules moving through the NPC or primary cilium should also have rotationally symmetrical distributions. As shown in Figure 2, a high number of randomly diffusing molecules inside the cylinder would generate rotationally symmetrical distributions at the cross-section view as that in the NPC, further resulting in an approximately uniform spatial distribution within each very small sub-region between two neighboring rings (Figure 2E). This uniform distribution leads that the spatial distribution along &#952; dimension in the cylindrical system is constant. Then the 3D coordinates (R, X, &#952;) can be simplified to be the 2D coordinates (R, X, constant). Actually, our conversion process between the Cartesian and the cylindrical systems is from 2D (X, Y) to 2D (R, X, constant). The constant &#952;, refers to the spatial density p in Figure 2E, is calculated by using the equation A<img alt="Equation 1" src="/files/ftp_upload/56475/56475eq1.jpg" />.
Ultimately, single-molecule tracking has broad application in biological research, thus, it is natural that a plethora of techniques will be developed to fill specific biological niches12,13,14. Such is the case with SPEED microscopy. Previously, when coupled with a 3D transformation algorithm, this technique was developed to resolve 3D transport routes of transiting molecules through the NPCs, a sub-diffraction-sized and rotationally symmetric biological structure6. In this paper, primary cilia are shown to be excellent model organelles as well. Primary cilia are cylindrical, antenna-like organelles (~125 nm radius) that project from the surface of most mammalian cells15,16,17. They are responsible for receiving external signals and transmitting an intracellular response typically associated with growth and metabolism15,16. Therefore, flux of structural proteins, recycling of transmembrane receptors, and transmission of intracellular messengers are vital responsibilities of primary cilia. At the juncture between the primary cilia and the cell body is a critical selectivity barrier, called the transition zone or TZ, through which all this protein transport must occur11,18,19,20. In addition to the gating function of the TZ, at least two transport processes, intraflagellar transport and passive diffusion, are thought to be responsible for the movement of protein through this region16,21,22. From a human health standpoint, the loss of primary cilia and subsequent deregulation of downstream signaling is characteristic of many cancers. In addition, many genetic diseases, such as Bardet-Biedl syndrome and polycystic kidney disease, are associated with defective protein transport23. Both the sub-diffraction limit size and the complex process of selective protein transport through the TZ make the primary cilia a prime target for this technique. In this methods paper, we will demonstrate the tracking of a ciliary transmembrane protein, somatostatin receptor 3 (SSTR3)24, labeled externally with Alexa Fluor 647 and a component of IFT, IFT2025, labeled with a fused GFP molecule.

<table><tbody><tr><td>25 cm&lt;sup&gt;2&lt;/sup&gt; tissue culture dish</td><td>Corning</td><td>VV-01936-00</td><td></td></tr><tr><td>Penicillin/streptomycin</td><td>ThermoFisher</td><td>15140122</td><td></td></tr><tr><td>Fetal bovine serum</td><td>ThermoFisher</td><td>10438018</td><td></td></tr><tr><td>DMEM</td><td>ThermoFisher</td><td>10566-016</td><td></td></tr><tr><td>OPTIMEM</td><td>ThermoFisher</td><td>31985062</td><td></td></tr><tr><td>Trypsin</td><td>ThermoFisher</td><td>25300054</td><td></td></tr><tr><td>Phosphate buffered saline</td><td>Sigma-Aldrich</td><td>P3813-1PAK</td><td></td></tr><tr><td>Transit LT1</td><td>Mirus</td><td>MIR 2300</td><td></td></tr><tr><td>35 mm glass bottom dish</td><td>MatTek</td><td>P35GCOL-0-14-C</td><td></td></tr><tr><td>AlexaFluor 647-conjugated streptavidin</td><td>ThermoFisher</td><td>S21374</td><td></td></tr><tr><td>Biotin</td><td>Sigma-Aldrich</td><td>B4501-100MG</td><td></td></tr><tr><td>633 nm He-Ne laser</td><td>Melles Griot</td><td>25-LHP-928-249</td><td></td></tr><tr><td>561 nm solid state laser</td><td>Coherent</td><td>OBIS 561-50 LS</td><td></td></tr><tr><td>488 nm solid state laser</td><td>Coherent</td><td>1185053</td><td></td></tr><tr><td>Inverted fluorescence microscope</td><td>Olympus</td><td>IX81</td><td></td></tr><tr><td>1.4-NA 100&amp;times; oil-immersion apochromatic objective</td><td>Olympus</td><td>UPLSAPO 100&amp;times;</td><td></td></tr><tr><td>On-chip multiplication gain charge-coupled-device camera</td><td>Roper Scientific</td><td>Cascade 128+</td><td></td></tr><tr><td>Dichroic filter</td><td>Semrock</td><td>Di01- R405/488/561/635-25x36</td><td></td></tr><tr><td>Emission filter</td><td>Semrock</td><td>NF01-405/488/561/635-25X5.0</td><td></td></tr><tr><td>Slidebook 6.0</td><td>Intelligent Imaging Innovations</td><td></td><td>digital microscopy software</td></tr></tbody></table>

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.

This section demonstrates the data obtained from performing SPEED microscopy at the TZ of primary cilia to study the transport route of SSTR3 connected by a ~15 nm external linker to Alexa647 (Figure 3A). It serves the dual purpose of verifying the 3D transformation algorithm. Alexa647 should only label the external surface of the primary cilium and therefore, the 3D transport route should reveal a high-density transport route at that location. NIH-3T3 stably expressing NPHP3-mCherry at the TZ must be transfected with AP-SSTR3-GFP and incubated according to the above protocol with streptavidin-conjugated Alexa647. Wide field imaging of the mCherry label is used to localize the TZ while the GFP is used to ensure that a cell has expressed the desired SSTR3 construct. Additionally, wide field imaging of the Alexa647 label using 633 nm illumination is necessary to confirm that the cell has properly expressed the BirA plasmid and has absorbed sufficient biotin from the media. Positive confirmation is characterized by a visible cilium distinct from the background fluorescence (Figure 3B). Efficient labeling of the AP-SSTR3-GFP construct with streptavidin-conjugated Alexa647 can only occur under these conditions. Once positive confirmation of selective labeling of the primary cilium by Alexa647 has been obtained, photobleaching and single point illumination of the TZ with a 633 nm laser will allow single molecules to be visualized. Before data analysis, superimposing the single molecule video on the wide field illumination is a useful validation step (Figure 3C). For example, if the laser is not aligned properly, no single molecules will appear to co-localize with the TZ. Another pre-data analysis validation step to ensure an adequate signal-to-noise ratio is to check the intensity at the TZ over the length of the single molecule video (Figure 3E). Such analysis can be performed quickly in ImageJ using the &#39;plot z axis profile&#39; function. The peaks in Figure 3F correspond to the appearance of a single molecule at the TZ with a signal-to-noise of ~5.0. This graph is also capable of ensuring that enough photobleaching has occurred as evidenced by the well-separated frequency of single molecules. Figure 3G demonstrates a low signal-to-noise ratio of &#60; 0.5. One explanation would be that the laser is not aligned directly on the TZ. Another reason may be that the laser illumination power is too low. Both explanations result in low excitation and, therefore, low emission of the fluorophores at the TZ. After these checks have been made, 2D Gaussian fitting of the single molecules will produce their trajectories in the TZ (Figure 3H). Superimposing many trajectories will produce the transport route of the labeled protein of interest in the TZ (Figure 3I). Since the 2D Gaussian fitting corresponds to the pixel values on the detector, the x, y single molecule data may be merged with the wide field image of the TZ and primary cilia to further illustrate the protein of interest&#39;s localization (Figure 3J).
IFT20 is a component of the IFT complex, which carries cargo along microtubules inside primary cilia25. Arl13b-mCherry, a widely used ciliary marker protein was used to label primary cilia19. Wide-field epi-fluorescence microscopy was used to image the primary cilium expressing both Arl13b-mCherry and IFT20-GFP and then switched to SPEED microscopy to track individual IFT20-GFP proteins in primary cilia after a pre-photobleaching of GFP fluorescence down to single-molecule level in the illumination area of SPEED microscopy (Figure 4A-4C).
Eventually, hundreds of single-molecule IFT20-GFP locations with a systematical localization precision of &#60; 16 nm can be obtained from the primary cilium of a live cell (Figure 4D). According to our simulation, 250 single molecule locations with a radius of 95 nm was enough to generate a reliable 3D transport route (Figure 5). A final determination of IFT20-GFP transport route is based on thousands of single-molecule IFT20-GFP locations collected from ten cilia (Figure 4E). Since primary cilia possess a structure with rotational symmetry, the 2D to 3D transformation algorithm was applied to the 2D projected data. Interestingly, the 3D spatial probability density maps of IFT20 indicated that only a single high-density region with a ~60-nm width peaked at ~95 nm along the radius of primary cilia (Figure 4E). This high-density region likely co-localizes with the axonemal microtubules, in agreement with the known location of the IFT route (Figure 4F).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56475/56475fig1.jpg" /> 
Figure 1. Simplified schematic of SPEED Microscopy. A vertical (path 1) or an inclined (path 2) point spread function is used to illuminate single fluorophore molecules in the focal plane. &#34;d&#34; refers to the distance between vertical illumination laser beam which passes through the center of the objective and angled illumination which is achieved by focusing the illumination laser on the edge of the objective. Two or more lasers regulated by an optical chopper to have an on-off excitation mode are used to track the single molecules transiting through the NPC or the primary cilium. The off time is at least ten-fold longer than the photobleaching time of the fluorophore label on the targeted molecules. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/56475/56475fig1large.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/56475/56475fig2.jpg" /> 
Figure 2. Graphical demonstration of 2D to 3D conversion process. Schematics demonstrate the 2D to 3D transformation algorithm for molecules, for example, if they diffuse within the central lumen (A-F) or a peripheral region (G-L) of a tube. (A) Ideal 3D spatial locations of randomly diffusing molecules inside a tube can be coordinated in a cylindrical coordination system (R, X, &#952;). In SPEED, only the 3D density map is ultimately calculated from the 2D spatial locations. However, this is an idealized case to illustrate that the 3D density map is equivalent whether the 2D or 3D spatial locations are known. (B) The 3D molecular locations in A are projected onto a 2D plane in a Cartesian coordination system (X, Y, Z) by microscopy imaging. (C) A very thin slice (&#916;x) cut from the cylinder in A along x dimension. (D) The 3D spatial locations in the slice shown in C can be projected within a narrow 2D region. (E) Cross-section view of all the locations in the thin slice shown in C. These locations can be grouped into the sub-regions between concentric rings. Given the high-number randomly distributed molecules in the cylinder and the cut very thin slice, the spatial density of locations (pi) in each sub-region (Si) between two neighboring rings will be rotationally symmetrical and uniform. These locations can be further projected into 1D along the Y dimension. If the locations along Y dimension are clustered in a histogram with j columns. The total number of locations in each column (Aj) is equal to <img alt="Equation 2" src="/files/ftp_upload/56475/56475eq2.jpg" />, which can be experimentally measured as shown in (F). (G-L) Similar as the above, the transformation process is presented for molecules diffusing within a peripheral region of a tube. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/56475/56475fig2large.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/56475/56475fig3.jpg" /> 
Figure 3. 3D spatial location of the transport route for SSTR3 on live primary cilia mapped by SPEED microscopy. (A) Graphical representation of SPEED microscopy being applied to tracking of GFP-tagged SSTR3 through the TZ marked by NPHP4-mCherry. (B) Epifluorescence image of NPHP4-mCherry (red), AP-SSTR3-GFP (green), Alexa647 used to externally label AP-SSTR3-GFP (white), and the final merged image. Scale bar: 2 &#181;m. (C) Tracking of a single Alexa 647-labeled AP-SSTR3-GFP (white) that passes through the point illumination field for four frames (2 ms/frame). Scale bar: 2 &#181;m. (D) Single molecule trajectory (white) from (C) superimposed on the epifluorescence image of NPHP4-mCherry (red) and AP-SSTR3-GFP (green). Scale bar: 2 &#181;m. (E) White dashed square highlights the area of single point illumination centered on the TZ. Scale bar: 2 &#181;m. (F) Photon count vs Frame number graph for a single molecule video with a high signal to noise ratio determined by dividing the maximum fluorescence of the single molecule divided by the background fluorescence. (G) Photon count vs Frame number graph for a single molecule video with a low signal to noise ratio. (H) Trajectory from (E) and (D) localized in each frame by 2D Gaussian fitting and superimposed on an accurate graphical representation of the TZ, also localized by 2D Gaussian fitting. The 2D Gaussian fitting process fits a Gaussian function to the X and Y dimensions of the intensity profile of an area of interest (AOI) encompassing the single molecule&#39;s PSF.(I) 2D super-resolution spatial distribution of 220 individual Alexa Fluor 647-labeled AP-SSTR3-GFP locations collected from a single primary cilium. (J) Super-resolution single molecule locations from (I) superimposed on the epifluorescence image of NPHP4-mCherry (red) and AP-SSTR3-GFP (green). Scale bar: 2 &#181;m. (K) By a 2D to 3D transformation algorithm, the spatial probability density distribution of Alexa Fluor 647-labeled AP-SSTR3-GFP on primary cilia along the R dimension was obtained. Based on Gaussian function fitting, Alexa Fluor 647-labeled AP-SSTR3-GFP primarily located on the ciliary membrane at a radius of 131&#177;3 nm with a full width at half maximum (FWHM) of 24&#177;1 nm. (L) Cross-section view of the spatial probability density distribution (green cloud) of Alexa Fluor 647-labeled AP-SSTR3-GFP on primary cilia. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/56475/56475fig3large.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/56475/56475fig4.jpg" /> 
Figure 4. 3D spatial location of the transport route for IFT20 inside live primary cilia mapped by SPEED microscopy. (A-C) Representative image of Arl13b-mCherry (A) and IFT20-GFP (B) co-expressed in an NIH3T3 cell and the merged (C). The circle (cyan) indicates the location of single-point illumination of SPEED microscopy on a primary cilium grew on the side of a cell (white dashed lines) whose edges are determined by IFT20-GFP fluorescence in the cell body and bright field illumination (not shown). Scale bar: 10 &#181;m. (D) 2D super-resolution spatial distribution of 286 individual IFT20-GFP locations collected from a single primary cilium. (E) By a 2D to 3D transformation algorithm, the spatial probability density distribution of IFT20-GFP inside primary cilia along the R dimension is obtained. Based on Gaussian function fitting, IFT20-GFP primarily locate at a radius of 95&#177;1 nm with a full width at half maximum (FWHM) of 56&#177;5 nm. (F) Cross-section view of the spatial probability density distribution (green clouds) of IFT20-GFP in primary cilia, overlaid with the schematic in H. Scale bar: 100 nm. <a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/56475/56475fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/56475/56475fig5.jpg" /> 
Figure 5. Simulation was used to estimate the minimum number of 2D single-molecule locations to generate a reliable 3D spatial probability density map for the 2D to 3D transformation algorithms. (A) 100 computationally-generated single-molecule locations randomly sample from a radial density normal distribution centered at 95 nm (corresponding to the primary transport route of IFT20 determined in our experiments). Localization precision of 16 nm is simulated for each single-molecule location by sampling from a normal distribution with &#963; = 16 nm. As shown in Figure 2C, a very thin slice (&#916;x) in the X dimension is used for transformation algorithms and therefore the X dimension is not shown in the computationally-generated 2D single-molecule locations. (B) Transformation between Y-dimensional project data and 3D R-dimensional density generate histograms of 3D R-dimensional densities for five different simulated data sets (each with 100 points). The number above is the peak position &#177; fitting error. (C-D) Simulation results based on 250 single-molecule locations. (E-F) Simulation results based on 500 single-molecule locations. (G-I) Average 3D density histograms from 100-, 250- and 500-points simulations respectively. Error bars represent variability in the histogram bin heights while the number above the peak is the average peak position &#177; standard deviation.&#160;<a href="https://cloudflare-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/56475/56475fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Application of High-speed Super-resolution SPEED Microscopy in Live Primary Cilium at JoVE.com

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

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 ...

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8.2K 조회수.  Temple University. 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.

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

2D and 3D Human Induced Pluripotent Stem Cell-Based Models to Dissect Primary Cilium Involvement during Neocortical Development

Primary cilia (PC) are non-motile dynamic microtubule-based organelles that protrude from the surface of most mammalian cells. They emerge from the older centriole during the G1/G0 phase of the cell cycle, while they disassemble as the cells re-enter the cell cycle at the G2/M phase boundary. They function as signal hubs, by detecting and transducing extracellular signals crucial for many cell processes. Similar to most cell types, all neocortical neural stem and progenitor cells (NSPCs) have been shown harboring a PC allowing them to sense and transduce specific signals required for the normal cerebral cortical development. Here, we provide detailed protocols to generate and characterize two-dimensional (2D) and three-dimensional (3D) cell-based models from human induced pluripotent stem cells (hIPSCs) to further dissect the involvement of PC during neocortical development. In particular, we present protocols to study the PC biogenesis and function in 2D neural rosette-derived NSPCs including the transduction of the Sonic Hedgehog (SHH) pathway. To take advantage of the three-dimensional (3D) organization of cerebral organoids, we describe a simple method for 3D imaging of in toto immunostained cerebral organoids. After optical clearing, rapid acquisition of entire organoids allows detection of both centrosomes and PC on neocortical progenitors and neurons of the whole organoid. Finally, we detail the procedure for immunostaining and clearing of thick free-floating organoid sections preserving a significant degree of 3D spatial information and allowing for the high-resolution acquisition required for the detailed qualitative and quantitative analysis of PC biogenesis and function.

We present detailed protocols for the generation and characterization of 2D and 3D human induced pluripotent stem cell (hIPSC)-based models of neocortical development as well as complementary methodologies enabling qualitative and quantitative analysis of primary cilium (PC) biogenesis and function.

2D 및 3D 인간 유도 만능 줄기 세포 기반 모델은 신코르티컬 개발 중 1 차용성 개입을 해부합니다.

Primary cilia (PC) are non-motile dynamic microtubule-based organelles that protrude from the surface of most mammalian cells. They emerge from the older ...

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발생학

Observation of the Ciliary Movement of Choroid Plexus Epithelial Cells Ex Vivo

The choroid plexus is located in the ventricular wall of the brain, the main function of which is believed to be production of cerebrospinal fluid. Choroid plexus epithelial cells (CPECs) covering the surface of choroid plexus tissue harbor multiple unique cilia, but most of the functions of these cilia remain to be investigated. To uncover the function of CPEC cilia with particular reference to their motility, an ex vivo observation system was developed to monitor ciliary motility during embryonic, perinatal and postnatal periods. The choroid plexus was dissected out of the brain ventricle and observed under a video-enhanced contrast microscope equipped with differential interference contrast optics. Under this condition, a simple and quantitative method was developed to analyze the motile profiles of CPEC cilia for several hours ex vivo. Next, the morphological changes of cilia during development were observed by scanning electron microscopy to elucidate the relationship between the morphological maturity of cilia and motility. Interestingly, this method could delineate changes in the number and length of cilia, which peaked at postnatal day (P) 2, while the beating frequency reached a maximum at P10, followed by abrupt cessation at P14. These techniques will enable elucidation of the functions of cilia in various tissues. While related techniques have been published in a previous report1, the current study focuses on detailed techniques to observe the motility and morphology of CPEC cilia ex vivo.

Observation of the Ciliary Movement of Choroid Plexus Epithelial Cells Ex Vivo

In this study, a detailed light microscopic technique was optimized for real-time observation and analysis of the motion of CPEC cilia ex vivo together with an electron microscopic method for ultrastructural analysis.

맥락총 상피 세포의 섬모 운동의 관측<em&gt; 전의 VIVO</em

The choroid plexus is located in the ventricular wall of the brain, the main function of which is believed to be production of cerebrospinal fluid. ...

신경과학

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment and maintenance of epithelial tissue morphology and organ morphogenesis. Moreover, the BM is essential for tissue modeling, serving as a signaling platform, and providing external forces to shape tissues and organs. Despite the many important roles that the BM plays during normal development and pathological conditions, the biological pathways controlling the intracellular trafficking of BM-containing vesicles and how basal secretion leads to the polarized deposition of BM proteins are poorly understood. The follicular epithelium of the Drosophila ovary is an excellent model system to study the basal deposition of BM membrane proteins, as it produces and secretes all major components of the BM. Confocal and super-resolution imaging combined with image processing in fixed tissues allows for the identification and characterization of cellular factors specifically involved in the intracellular trafficking and deposition of BM proteins. This article presents a detailed protocol for staining and imaging BM-containing vesicles and deposited BM using endogenously tagged proteins in the follicular epithelium of the Drosophila ovary. This protocol can be applied to address both qualitative and quantitative questions and it was developed to accommodate high-throughput screening, allowing for the rapid and efficient identification of factors involved in the polarized intracellular trafficking and secretion of vesicles during epithelial tissue development.

Confocal and Super-Resolution Imaging of Polarized Intracellular Trafficking and Secretion of Basement Membrane Proteins During Drosophila Oogenesis

The basement membrane is essential for tissue and organ morphogenesis during development. To better understand the mechanisms leading to proper placement of this structure, the protocol presented describes methods to visualize and characterize the intracellular trafficking and secretion of basement membrane proteins in epithelial cells using confocal and super-resolution microscopy.

초 파리 Oogenesis 동안 양극화 된 세포 내 밀매 및 기저막 단백질의 분비의 공초점 및 수퍼 해상도 이미징

The basement membrane (BM) - a specialized sheet of extracellular matrix present at the basal side of epithelial cells - is critical for the establishment ...

Ex vivo Method for High Resolution Imaging of Cilia Motility in Rodent Airway Epithelia

An ex vivo technique for imaging mouse airway epithelia for quantitative analysis of motile cilia function important for insight into mucociliary clearance function has been established. Freshly harvested mouse trachea is cut longitudinally through the trachealis muscle and mounted in a shallow walled chamber on a glass-bottomed dish. The trachea sample is positioned along its long axis to take advantage of the trachealis muscle to curl longitudinally. This allows imaging of ciliary motion in the profile view along the entire tracheal length. Videos at 200 frames/sec are obtained using differential interference contrast microscopy and a high speed digital camera to allow quantitative analysis of cilia beat frequency and ciliary waveform. With the addition of fluorescent beads during imaging, cilia generated fluid flow also can be determined. The protocol time spans approximately 30 min, with 5 min for chamber preparation, 5-10 min for sample mounting, and 10-15 min for videomicroscopy.

Ex vivo Method for High Resolution Imaging of Cilia Motility in Rodent Airway Epithelia

An easy and reliable technique for visualizing and quantifying airway cilia motility and cilia generated flow using mouse trachea is described. This technique can be modified to determine how a wide range of factors influence cilia motility, including pharmacological agents, genetic factors, environmental exposures, and/or mechanical factors such as mucus load.

설치류기도 상피의 섬모 운동성의 높은 해상도 이미지를위한 생체 방법

An ex vivo technique for imaging mouse airway epithelia for quantitative  analysis of motile cilia function important for insight into mucociliary  ...

생물학

Super-Resolution Imaging to Study Co-Localization of Proteins and Synaptic Markers in Primary Neurons

Synapses are the functional elements of neurons and their defects or losses are at the basis of several neurodegenerative and neurological disorders. Imaging studies are widely used to investigate their function and plasticity in physiological and pathological conditions. Because of their size and structure, localization studies of proteins require high-resolution imaging techniques. In this protocol, we describe a procedure to study in primary neurons the co-localization of target proteins with synaptic markers at a super-resolution level using structured illumination microscopy (SIM). SIM is a patterned-light illumination technique that doubles the spatial resolution of wide-field microscopy, reaching a detail of around 100 nm. The protocol indicates the required controls and settings for robust co-localization studies and an overview of the statistical methods to analyze the imaging data properly.

This protocol shows how to employ super-resolution microscopy to study protein co-localization in primary neuronal cultures.

1차 뉴런에서 단백질과 시냅스 마커의 공동 국소화를 연구하는 초해상도 이미징

Synapses are the functional elements of neurons and their defects or losses are at the basis of several neurodegenerative and neurological disorders. ...

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.

고속 3D 구조 조명 현미경을 사용하여 라이브 박테리아의 Cytokinetic Z 반지의 슈퍼 해상도 이미징 (F3D-SIM)

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

Efficient and Cost Effective Electroporation Method to Study Primary Cilium-Dependent Signaling Pathways in the Granule Cell Precursor

The primary cilium is a critical signaling organelle found on nearly every cell that transduces Hedgehog (Hh) signaling stimuli from the cell surface. In the granule cell precursor (GCP), the primary cilium acts as a pivotal signaling center that orchestrates precursor cell proliferation by modulating the Hh signaling pathway. The investigation of primary cilium-dependent Hh signaling machinery is facilitated by in vitro genetic manipulation of the pathway components to visualize their dynamic localization to the primary cilium. However, transfection of transgenes in the primary cultures of GCPs using the currently known electroporation methods is generally costly and often results in low cell viability and undesirable transfection efficiency. This paper introduces an efficient, cost-effective, and simple electroporation protocol that demonstrates a high transfection efficiency of ~80-90% and optimal cell viability. This is a simple, reproducible, and efficient genetic modification method that is applicable to the study of the primary cilium-dependent Hedgehog signaling pathway in primary GCP cultures.

Here, we present a reproducible in vitro electroporation protocol for genetic manipulation of primary cerebellar granule cell precursors (GCPs) that is cost-effective, efficient, and viable. Moreover, this protocol also demonstrates a straightforward method for the molecular study of primary cilium-dependent Hedgehog signaling pathways in primary GCP cells.

과립 세포 전구체에서 1 차적인 실륨 의존적 신호 경로를 연구하는 효율적이고 비용 효율적인 전기 포기법

The primary cilium is a critical signaling organelle found on nearly every cell that transduces Hedgehog (Hh) signaling stimuli from the cell surface. In ...

Simple Detection of Primary Cilia by Immunofluorescence

Primary cilia are dynamically regulated during cell cycle progression, specifically during the G0/G1 phases of the cell cycle, being resorbed prior to mitosis. Primary cilia can be visualized with highly sophisticated methods, including transmission electron microscopy, 3D imaging, or using software for the automatic detection of primary cilia. However, immunofluorescent staining of primary cilia is needed to perform these methods. This publication describes a protocol for the easy detection of primary cilia in vitro by staining acetylated alpha tubulin (axoneme) and gamma tubulin (basal body). This immunofluorescent staining protocol is relatively simple and results in high-quality images. The present protocol describes how four cell lines (C2C12, MEF, NHLF, and skin fibroblasts) expressing primary cilia were fixed, immunostained, and imaged with a fluorescent or confocal microscope.

Primary cilia are extracellular structures associated with the centriole. Primary cilia detection by immunofluorescent staining is a relatively simple procedure that results in extremely high-quality images. In this protocol, fibroblasts expressing primary cilia were fixed, immunostained, and imaged in a fluorescent or confocal microscope.

면역 형광에 의한 1 차 적인 섬모의 간단한 검출

Primary cilia are dynamically regulated during cell cycle progression, specifically during the G0/G1 phases of the cell cycle, being resorbed prior to ...

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.

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.

세포 전 형 구조의 슈퍼 해상도 라이브 세포 이미징

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 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.

Caenorhabditis elegans 생식계열 내 시냅톤 복합체의 초고해상도 현미경

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