This work was supported by the Japan Society for the Promotion of Science (Grant-in-Aid for Scientific Research on Innovative Areas (JP16H06464, JP16H06280 to T.H.), Grant-in-Aid for Scientific Research (B, JP17H03697 to D.K.), Grant-in-Aid for Challenging Exploratory Research (JP18K19331 to D.K.), Grant-in-Aid for Scientific Research on Innovative Areas (JP20H05358 for D.K.)) and the Japan Science and Technology Agency (PRESTO program (JPMJPR15QC to Y.M., JPMJPR18K4 to D.K.)). The authors are thankful to the Live Imaging Center at the Institute of Transformative Bio-Molecules (WPI-ITbM) of Nagoya University for supporting the microscopic studies and Editage (www.editage.com) for English language editing.

1. Preparation of clearing solutions
<ol>
	<li>To prepare CS solution, dissolve 10% (w/v) xylitol, 15% (w/v) sodium deoxycholate, and 25% (w/v) urea in distilled water on a magnetic stirrer. 
	NOTE: Sodium deoxycholate powder should be weighed in a draft chamber as it easily floats in the air. CS can be stored at room temperature in dark for more than 1 year.</li>
	<li>To prepare CSA solution, add sodium sulfite (50 mM final concentration) to the CS solution obtained above. 
	&#8203;NOTE: Add sodium sulfite to the CS solution right before use as the reducing agent gets deactivated easily.</li>
</ol>
2. Preparation of the fixative solution
<ol>
	<li>Transfer 40 mL of sterilized water into a conical tube and add 2 g of paraformaldehyde. Add 200 &#181;L of 2 N NaOH to increase the pH of the solution. After closing and sealing the tube with parafilm, incubate at 60 &#176;C with occasional inversions until everything is dissolved.</li>
	<li>After cooling the solution to room temperature, add 5 mL of 10x phosphate-buffered saline (PBS) to adjust the pH to 7.4. Add sterilized water to make up the volume to 50 mL. 
	&#8203;NOTE: A freshly prepared fixative solution is preferred. The solution can also be stored at -30 &#176;C for several months.</li>
</ol>
3. Fixation of samples
<ol>
	<li>Immerse plant samples in the fixative solution in a microtube (Figure 1A), ensuring that the volume of the fixative solution is more than five times the sample volume.</li>
	<li>Seal the microtube with a parafilm and make holes using a needle. Do not leave the tube open owing to a risk of sample spillage during vacuum decompression.</li>
	<li>Place the microtube in a desiccator and slowly adjust the degree of vacuum (~690 mmHg) so that bubbles appear gradually from the samples (Figure 1B-C). Turn off the vacuum pump after evacuating the desiccator. Leave the microtube undisturbed for 30 min at room temperature.</li>
	<li>Vent the desiccator carefully to prevent disturbing the samples. Turn on the vacuum pump again and turn it off after evacuating the desiccator. Leave the microtube undisturbed for 30 min at room temperature. 
	NOTE: Care should be taken to prevent damage to the samples while venting the desiccator. The penetration of the fixative solution into the samples is enhanced by two vacuum treatments. Further vacuum treatment helps to penetrate the fixative solution into thicker samples.</li>
	<li>Open the desiccator carefully without bumping the fixative solution in the microtube. Using a micropipette, remove the fixative solution and add 1x PBS. After storing for 1 min, replace the old PBS with new 1x PBS (Figure 1D).</li>
</ol>
4. Clearing
<ol>
	<li>After removing PBS, add five times the sample volume of the clearing solution.</li>
	<li>Seal the microtube with parafilm and make holes using a needle. Place the samples in the desiccator, evacuate as in step 3.3, and turn off the vacuum pump. Leave the microtube undisturbed for 60 min at room temperature.</li>
	<li>Open the desiccator gently. Close the microtube with parafilm and store it at room temperature in the dark to avoid photobleaching of fluorescent proteins. Invert the microtube every 1-2 days to accelerate the clearing process.</li>
	<li>When the clearing solution turns green, replace it with new clearing solutions until the solution remains colorless (Figure 1E-H).</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/63428/63428fig01.jpg" /> 
Figure 1: Procedure for CS treatment. (A) Arabidopsis seedling in 4% PFA (Paraformaldehyde) solution. (B) The sample is placed into a desiccator. (C) The seedling is fixed under a vacuum. (D) The seedling is soaked in the PFA solution after vacuum treatment. (E) Resulting 3-day clearing solution-treated seedling. Note the green color of the clearing solution. (F) The clearing solution is replaced 3 days after treatment. (G) Resulting 5-day clearing solution-treated seedling. (H) The clearing solution is replaced 5 days after treatment. Scale bars: 1 cm (A,C-H) and 5 cm (B). <a href="https://www.jove.com/files/ftp_upload/63428/63428fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
5. Chemical dye staining
<ol>
	<li>To the microtube, add Hoechst 33342 (final concentration of 10 &#181;g/mL) for nuclear staining or Calcofluor White (final concentration of 1 mg/mL) for cell wall staining and wait for 1 h. After removing the dye solution, wash the sample with a fresh clearing solution for 1 h. 
	&#8203;NOTE: Overnight staining and washing can improve fluorescent dye penetration into tissues and reduce background fluorescence. Various fluorescent dyes are compatible with the CS solution such as Basic Fuchsin9 (lignin), Auramine O9 (lignin, suberin, and cutin), Nile Red9 (suberin), Direct Yellow 969, Direct Red 239, and SR220010,11 (cell walls).</li>
</ol>
6. Observation
<ol>
	<li>Cut silicone rubber sheet with a razor blade to prepare a frame for the spacer (Figure 2A). 
	NOTE: Adjust the thickness of the silicon rubber sheet according to the thickness of the sample. As samples treated with clearing solution are soft, they will be damaged if they are covered directly with the cover glass.</li>
	<li>Place the silicone frame on cover glass (e.g., 25 x 60 mm) (Figure 2B). Place the treated samples within the frame and add ~100 &#181;L of clearing solution to remove any bubbles in the frame. Cover with another cover glass (18 x 18 mm or 24 x 24 mm) to prevent evaporation of the clearing solution (Figure 2C).</li>
	<li>Observe the samples under a fluorescent microscope. After observation, return the samples to the clearing solution taken in a microtube and store at room temperature in the dark.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/63428/63428fig02.jpg" /> 
Figure 2: Sample preparation for microscopic observation. (A) Cut a 0.2 mm thick silicone sheet into a frame. (B) Put the silicone sheet frame onto the cover glass. (C) Place the sample treated with clearing solution (marked by dotted border) within the frame and cover it with a cover glass. Scale bars: 5 mm. <a href="https://www.jove.com/files/ftp_upload/63428/63428fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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ClearSee is commercialized (Fujifilm Wako Pure Chemical Corporation, Japan). Nagoya University holds a patent (JP6601842) on the clearing reagent ClearSee. D. Kurihara is the patent inventor. The authors declare no competing financial interests.

This method consists of fixation, washing, and cleaning. Fixation is a critical step in this protocol. If the fluorescent protein is not observed after PFA fixation, it will not be observed after treatment with clearing solution. The penetration of PFA solution into tissues is critical, but high vacuum treatment is not recommended because it can destroy the cell structure. Vacuum conditions and fixation periods should be optimized for each type of tissue and species. It is recommended to check fluorescent proteins even after fixation. Although samples were usually fixed for 30-60 min at room temperature, they can be fixed at 4 &#176;C for a longer time (overnight or more).
As shown in Figure 6A, some sodium deoxycholates had a pale-yellow color when dissolved. Such sodium deoxycholate solutions showed strong autofluorescence in the 400-600 nm region after excitation at 380 nm (Figure 6B). This autofluorescence prevents optical clearing and fluorescence imaging. Users should check the color of sodium deoxycholate solution as the quality of the reagent might differ owing to purity, lot-to-lot variation, or other reasons.
The clearing solutions used here have high concentrations of sodium deoxycholate, which could destroy the membrane structure. The plasma membrane marker (RPS5Apro::tdTomato-LTI6b) was observed even after CS treatment7. However, it might be better to reduce the concentration of sodium deoxycholate, depending on the structure and tissue of interest. Indeed, images with improved clarity were obtained for Arabidopsis pistils with modified CS, in which the concentration of sodium deoxycholate is reduced by half; however, reduced concentrations of sodium deoxycholate required prolonged treatment times (e.g., 1 month for Arabidopsis pistils).
CS can reduce red autofluorescence (&#62;610 nm) to remove chlorophyll in treated samples. However, 500-600 nm range autofluorescence (yellow to orange) remained even in CS-treated samples7. This autofluorescence is thought to be derived from the cell wall and other cellular components, such as lignin12,13. Therefore, it is difficult to make tissues, such as stems with developed secondary walls, completely transparent by CS treatment.
Several clearing reagents besides the ones used here have been developed to observe fluorescent proteins in plants using fluorescent microscopy14,15,16,17. Compared with these methods, CS and CSA remove chlorophyll and reduce autofluorescence, making the plant tissues more transparent. Recently, Sakamoto et al. developed an improved method, iTOMEI, for fixation, detergent clearing, and mounting to adjust the refractive index mismatch18. In Arabidopsis seedlings, iTOMEI cleared the tissue within 26 h.
CS is applicable to a wide range of plant species, such as Arabidopsis thaliana, Physcomitrium patens7, Chrysanthemum morifolium, Cucumis sativus, Nicotiana benthamiana, Nicotiana tabacum, Torenia fournieri8, Allium ochotense19, Astragalus sinicus20, avocado21, barley22, Brassica rapa23, Callitriche24, Eucalyptus25, maize26, Marchantia polymorpha27, Monophyllaea glabra28, Orobanche minor29, petunia30, rice31, Solanum lycopersicum32, soybean33, strawberry34, wheat35, and Wolffiella hyalina36. For thicker tissues, CS can also make the vibratome sections transparent37,38. This method allowed studies of the cellular structure and gene expression patterns in plants37,38. Moreover, nematode infections20,39, fungal infections, and symbiosis19,40,41 were also observed deep inside the CS-treated tissues. Thus, this method is useful for whole tissue imaging from micro- to macro-scales and could help to discover novel interactions among various cells, tissues, organs, and organisms.

Visualization of cellular structures and localization of proteins in living organisms is important for clarifying their functions in vivo. However, since the living body is not transparent, it is challenging to observe the internal structure of living organisms without dissection. Especially, in the case of plant tissues, which are multi-layered with cells of different shapes, the index mismatch caused by their structure and the presence of light-absorbing pigments is problematic. For example, plant leaves have a complex structure that allows them to efficiently utilize the light that enters their bodies for photosynthesis1, whereas the structure also causes refractive index mismatch, making them difficult to observe. However, leaves have many light-absorbing pigments, such as chlorophyll, which emit strong red fluorescence and brownish pigments are produced by oxidation2,3. These pigments also hinder whole-mount fluorescence microscopy observations in plants. Therefore, for observing the internal structure of plants, decolorization and fixation by alcohol and clearing using chloral hydrate have been used for a long time to eliminate the refractive index mismatch and autofluorescence4,5. These conventional methods have been adopted for many years, but they have the drawback of eliminating the fluorescence of fluorescent proteins at the same time6,7. This is problematic since fluorescent proteins have become indispensable in current fluorescent imaging.
Therefore, ClearSee (CS) and ClearSeeAlpha (CSA) have been developed as optical clearing reagents for plant tissues. Both reagents reduce chlorophyll autofluorescence while maintaining the stability of fluorescent proteins7,8. CSA is particularly useful when brown pigments are produced owing to tissue oxidation. Using these clearing reagents, it is possible to observe the cellular structure and protein localization inside the plant body without physical sectioning.

<table><tbody><tr><td>Calcofluor White</td><td>Sigma-Aldrich</td><td>F3543</td><td>Fluorescent Brightener 28; 100 mg/mL in H2O</td></tr><tr><td>ClearSee</td><td></td><td></td><td>10% (w/v) xylitol, 15% (w/v) sodium deoxycholate, 25% (w/v) urea</td></tr><tr><td>Cover glass (18&amp;times;18 No.1)</td><td>MATSUNAMI</td><td>C018181</td><td></td></tr><tr><td>Cover glass (24&amp;times;24 No.1)</td><td>MATSUNAMI</td><td>C024241</td><td></td></tr><tr><td>Cover glass (25&amp;times;60 No.1)</td><td>MATSUNAMI</td><td>C025601</td><td></td></tr><tr><td>Desiccator</td><td>AS One</td><td>1-5801-11</td><td></td></tr><tr><td>Hoechst 33342</td><td>DOJINDO</td><td>346-07951</td><td>1 mg/mL in H2O</td></tr><tr><td>Needle</td><td>TERUMO</td><td>NN-2238S</td><td></td></tr><tr><td>Parafilm</td><td>Bemis</td><td>PM-996</td><td></td></tr><tr><td>Paraformaldehyde</td><td>Nacalai Tesque</td><td>26126-25</td><td></td></tr><tr><td>Phosphate buffered saline, pH 7.4</td><td></td><td></td><td></td></tr><tr><td>Silicone rubber sheet</td><td>AS One</td><td>6-9085-12</td><td></td></tr><tr><td>Sodium deoxycholate</td><td>Tokyo Chemical Industry</td><td>C0316</td><td>Figure 6_1; Lot PSGYK-QB</td></tr><tr><td>Sodium deoxycholate</td><td>Kishida Chemical</td><td>260-71412</td><td>Figure 6_2; Lot C05543H</td></tr><tr><td>Sodium deoxycholate</td><td>Sigma-Aldrich</td><td>D6750</td><td>Figure 6_3; Lot SLBS7362</td></tr><tr><td>Sodium deoxycholate</td><td>Sigma-Aldrich</td><td>30970</td><td>Figure 6_4; Lot BCBW0612</td></tr><tr><td>Sodium deoxycholate</td><td>Nacalai Tesque</td><td>10712-96</td><td>Figure 6_5; Lot M5R3403</td></tr><tr><td>Sodium deoxycholate</td><td>FUJIFILM Wako Pure Chemical</td><td>194-08311</td><td>Figure 6_6; Lot LKL0648</td></tr><tr><td>Sodium hydroxide</td><td>Nacalai Tesque</td><td>31511-05</td><td></td></tr><tr><td>Sodium sulphite</td><td>FUJIFILM Wako Pure Chemical</td><td>190-03411</td><td></td></tr><tr><td>Urea</td><td>FUJIFILM Wako Pure Chemical</td><td>211-01213</td><td></td></tr><tr><td>Vacuum pump</td><td>BUCHI</td><td>V-700</td><td></td></tr><tr><td>Xylitol</td><td>FUJIFILM Wako Pure Chemical</td><td>248-00545</td><td></td></tr></tbody></table>

optical clearing of plant tissues for fluorescence imaging

It is challenging to directly observe the internal structure of multi-layered and opaque plant specimens, without dissection, under a microscope. In addition, autofluorescence attributed to chlorophyll hampers the observation of fluorescent proteins in plants. For a long time, various clearing reagents have been used to make plants transparent. However, conventional clearing reagents diminish fluorescent signals; therefore, it has not been possible to observe the cellular and intracellular structures with fluorescent proteins. Reagents were developed that can clear plant tissues by removing chlorophyll while maintaining fluorescent protein stability. A detailed protocol is provided here for the optical clearing of plant tissues using clearing reagents, ClearSee (CS) or ClearSeeAlpha (CSA). The preparation of cleared plant tissues involves three steps: fixation, washing, and clearing. Fixation is a crucial step in maintaining the cellular structures and intracellular stability of fluorescent proteins. The incubation time for clearing depends on the tissue type and species. In Arabidopsis thaliana, the time required for clearing with CS was 4 days for leaves and roots, 7 days for seedlings, and 1 month for pistils. CS also required a relatively short time of 4 days to make the gametophytic leaves of Physcomitrium patens transparent. In contrast, pistils in tobacco and torenia produced brown pigment due to oxidation during CS treatment. CSA reduced the brown pigment by preventing oxidation and could make tobacco and torenia pistils transparent, although it took a relatively long time (1 or 2 months). CS and CSA were also compatible with staining using chemical dyes, such as DAPI (4',6-diamidino-2-phenylindole) and Hoechst 33342 for DNA and Calcofluor White, SR2200, and Direct Red 23 for the cell wall. This method can be useful for whole-plant imaging to reveal intact morphology, developmental processes, plant-microbe interactions, and nematode infections.

CS can clear leaves of various species (Figure 3A-H). It is difficult for the CS solution to penetrate a rice leaf because the leaf surface is covered by cuticular wax in this plant. However, after extracting the cuticular wax by dipping in chloroform for 10 s, CS could clear the rice leaves (Figure 3H). CS could not, however, penetrate Chamaecyparis obtusa leaves which are less permeable to CS (Figure 3I,J). In Chrysanthemum leaves, brown pigmentation induced by polyphenol oxidation was observed in the CS-treated leaves (Figure 3K,L). Similarly, Tobacco and torenia pistils showed brown pigmentation during the CS treatment (Figure 3M,O). As the sodium sulfite component in CSA prevents polyphenol oxidation owing to the reducing effect, CSA could clear tobacco and torenia pistils without any brown pigmentation (Figure 3N,P).
Figures 4B shows that CS treatment reduced the pale green color of the Arabidopsis H2B-mClover leaf (bright field) and enhanced the fluorescence intensity of H2B-mClover compared with PBS incubation (Figure 4A). In addition to flowering plants, CS is also applicable to moss plants (Figure 4C); after 4 days of CS treatment, the fluorescence of H2B-mRFP was clearly detected for the entire gametophore with reduced chlorophyll autofluorescence. Figures 4D,E show 3D reconstruction images of the H2B-mClover pistil in Nicotiana benthamiana cleared using CSA. The sample depth was 440 &#181;m. As the depth-coded maximum intensity projection image shows, CSA allows for deep imaging of challenging tissues, such as tobacco pistils.
CS and CSA were also compatible with fluorescent dye staining. Figure 5 shows that CS could simultaneously be used to observe the fluorescent protein (H2B-mClover) and organic fluorescent dye staining (Calcofluor White). After 3D reconstruction from the z-stack images, any section could be observed.
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/63428/63428fig03.jpg" /> 
Figure 3: Optical clearing of leaves and pistils using clearing solutions. (A-L) Fixed leaves of various species were incubated in PBS (A,C,E,G,I,K) or CS (B,D,F,H,J,L) for 8 days and CS (M,O) or CSA (N,P) for 2 days. (A,B,O,P) Torenia fournieri, (C,D) Nicotiana tabacum, (E,F) Cucumis sativus, (G,H) Oryza sativa, (I,J) Chamaecyparis obtusa, (K,L) Chrysanthemum morifolium, (M,N) Nicotiana benthamiana. Scale bars: 1 cm. <a href="https://www.jove.com/files/ftp_upload/63428/63428fig03large.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/63428/63428fig04.jpg" /> 
Figure 4: Fluorescence imaging of tissues treated with clearing solutions. (A,B) UBQ10pro::H2B-mClover leaves of Arabidopsis thaliana were treated with PBS (A) or CS (B) for 3 days. (C) H2B-mRFP leafy gametophores of Physcomitrium patens were treated with CS for 4 days. The nuclei were labeled with H2B-mRFP (green). The CS treatment reduced chlorophyll autofluorescence (magenta). The H2B-mRFP signal in the apical region was clearly observed for both merged images of H2B-mRFP and autofluorescence or bright field. (D,E) The UBQ10pro::H2B-mClover stigma of Nicotiana benthamiana was treated in CSA for 1 month. Maximum-intensity projection (D) and depth-coded maximum-intensity projection (E) were generated from 88 z-stack images at 5 &#181;m intervals. Scale bars: 100 &#181;m. Images were taken using wide-field (A,B), confocal (C), and two-photon excitation (D,E) microscopy. <a href="https://www.jove.com/files/ftp_upload/63428/63428fig04large.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/63428/63428fig05.jpg" /> 
Figure 5: Fluorescent dye staining is compatible with CS. (A) CS-treated leaves observed by two-photon excitation microscopy with 950 nm excitation. Cell wall is stained with Calcofluor White (cyan). Nuclei are labeled with UBQ10pro::H2B-mClover (yellow). The yz (B) and xz (C) images are cross-sections at the position indicated by the white dashed lines in (A). Scale bar: 100 &#181;m. <a href="https://www.jove.com/files/ftp_upload/63428/63428fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/63428/63428fig06.jpg" /> 
Figure 6: Transparency of sodium deoxycholate. (A) Colors of various 15% sodium deoxycholates (listed in Table of Materials). (B) Fluorescence spectrum of each 15% sodium deoxycholate with 380 nm excitation. <a href="https://www.jove.com/files/ftp_upload/63428/63428fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Optical Clearing of Plant Tissues for Fluorescence Imaging at JoVE.com

Optical Clearing of Plant Tissues for Fluorescence Imaging

Here, a method is described for making plant tissues transparent while maintaining the stability of fluorescent proteins. This technique facilitates deep imaging of cleared plant tissues without physical sectioning.

It is challenging to directly observe the internal structure of multi-layered and opaque plant specimens, without dissection, under a microscope. In ...

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Plant Cell Structure and Organization

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5.1K Views.  Nagoya University. Here, a method is described for making plant tissues transparent while maintaining the stability of fluorescent proteins. This technique facilitates deep imaging of cleared plant tissues without physical sectioning.

Video: Optical Clearing of Plant Tissues for Fluorescence Imaging

Multiphoton Microscopy of Cleared Mouse Brain Expressing YFP

Multiphoton microscopy of intrinsic fluorescence and second harmonic generation (SHG) of whole mouse organs is made possible by optically clearing the organ before imaging.1,2 However, for organs that contain fluorescent proteins such as GFP and YFP, optical clearing protocols that use methanol dehydration and clear using benzyl alcohol:benzyl benzoate (BABB) while unprotected from light3 do not preserve the fluorescent signal. The protocol presented here is a novel way in which to perform whole organ optical clearing on mouse brain while preserving the fluorescence signal of YFP expressed in neurons. Altering the optical clearing protocol such that the organ is dehydrated using an ethanol graded series has been found to reduce the damage to the fluorescent proteins and preserve their fluorescent signal for multiphoton imaging.4 Using an optimized method of optical clearing with ethanol-based dehydration and clearing by BABB while shielded from light, we show high-resolution multiphoton images of yellow fluorescent protein (YFP) expression in the neurons of a mouse brain more than 2 mm beneath the tissue surface.

Multiphoton microscopy of whole mouse organs is possible by optically clearing the organ before imaging, but not all protocols preserve the fluorescent signal of fluorescent proteins. Using an optical clearing method with ethanol-based dehydration and benzyl alcohol:benzyl benzoate clearing, we show high-resolution multiphoton images of whole mouse brain expressing YFP.

Multiphoton microscopy of intrinsic fluorescence and second harmonic generation (SHG) of whole mouse organs is made possible by optically clearing the ...

Neuroscience

Whole-mount Clearing and Staining of Arabidopsis Flower Organs and Siliques

Due to its formidable tools for molecular genetic studies, Arabidopsis thaliana is one of the most prominent model species in plant biology and, especially, in plant reproductive biology. However, plant morphological, anatomical, and ultrastructural analyses traditionally involve time-consuming embedding and sectioning procedures for bright field, scanning, and electron microscopy. Recent progress in confocal fluorescence microscopy, state-of-the-art 3-D computer-aided microscopic analyses, and the continuous refinement of molecular techniques to be used on minimally processed whole-mount specimens, has led to an increased demand for developing efficient and minimal sample processing techniques. In this protocol, we describe techniques for properly dissecting Arabidopsis flowers and siliques, basic clearing techniques, and some staining procedures for whole-mount observations of reproductive structures.

Whole-mount Clearing and Staining of Arabidopsis Flower Organs and Siliques

In this protocol, we describe techniques for the proper dissection of Arabidopsis flowers and siliques, some basic clearing techniques, and selected staining procedures for whole-mount observations of reproductive structures.

Due to its formidable tools for molecular genetic studies, Arabidopsis thaliana is one of the most prominent model species in plant biology and, ...

Developmental Biology

Rapid Optical Clearing for Semi-High-Throughput Analysis of Tumor Spheroids

Tumor spheroids are fast becoming commonplace in basic cancer research and drug development. Obtaining data regarding protein expression within the spheroid at the cellular level is important for analysis, yet existing techniques are often expensive, laborious, use non-standard equipment, cause significant size distortion, or are limited to relatively small spheroids. This protocol presents a new method of mounting and clearing spheroids that address these issues while allowing for confocal analysis of the inner structure of spheroids. In contrast to existing approaches, this protocol provides for rapid mounting and clearing of a large number of spheroids using standard equipment and laboratory supplies. Mounting spheroids in a pH-neutral agarose-PBS gel solution before introducing a refractive-index-matched clearing solution minimizes size distortion common to other similar techniques. This allows for detailed quantitative and statistical analysis where the accuracy of size measurements is paramount. Furthermore, compared to liquid clearing solutions, the agarose gel technique keeps spheroids fixed in place, allowing for the collection of three-dimensional (3D) confocal images. The present article elaborates how the method yields high-quality two- and 3D images that provide information about inter-cell variability and inner spheroid structure.

Tumor spheroids are becoming increasingly utilized to assess tumor cell-microenvironment interactions and therapy response. The present protocol describes a robust but simple method for semi-high-throughput imaging of 3D tumor spheroids using rapid optical clearing.

Tumor spheroids are fast becoming commonplace in basic cancer research and drug development. Obtaining data regarding protein expression within the ...

Cancer Research

Imaging Cleared Intact Biological Systems at a Cellular Level by 3DISCO

Tissue clearing and subsequent imaging of transparent organs is a powerful method to analyze fluorescently labeled cells and molecules in 3D, in intact organs. Unlike traditional histological methods, where the tissue of interest is sectioned for fluorescent imaging, 3D imaging of cleared tissue allows examination of labeled cells and molecules in the entire specimen. To this end, optically opaque tissues should be rendered transparent by matching the refractory indices throughout the tissue. Subsequently, the tissue can be imaged at once using laser-scanning microscopes to obtain a complete high-resolution 3D image of the specimen. A growing list of tissue clearing protocols including 3DISCO, CLARITY, Sca/e, ClearT2, and SeeDB provide new ways for researchers to image their tissue of interest as a whole. Among them, 3DISCO is a highly reproducible and straightforward method, which can clear different types of tissues and can be utilized with various microscopy techniques. This protocol describes this straightforward procedure and presents its various applications. It also discusses the limitations and possible difficulties and how to overcome them.

To obtain high-resolution images of fluorescently labeled cells within large tissues, ideally, the biological samples should be imaged without sectioning. 3DISCO is a straightforward tissue clearing procedure based on sequential incubation with organic solvents. Upon clearing, the organs become transparent allowing an end-to-end laser scan of the specimen.

Tissue clearing and subsequent imaging of transparent organs is a powerful method to analyze fluorescently labeled cells and molecules in 3D, in intact ...

Optical Clearing of the Mouse Central Nervous System Using Passive CLARITY

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique non-linear deformations, dramatically hampering one's ability to evaluate cellular morphology, distribution and connectivity in the central nervous system (CNS). However, optical clearing techniques are changing the way tissues are visualized. These approaches permit one to probe deeply into intact organ preparations, providing tremendous insight into the structural organization of tissues in health and disease. Techniques such as Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) achieve this goal by providing a matrix that binds important biomolecules while permitting light-scattering lipids to freely diffuse out. Lipid removal, followed by refractive index matching, renders the tissue transparent and readily imaged in 3 dimensions (3D). Nevertheless, the electrophoretic tissue clearing (ETC) used in the original CLARITY protocol can be challenging to implement successfully and the use of a proprietary refraction index matching solution makes it expensive to use the technique routinely. This report demonstrates the implementation of a simple and inexpensive optical clearing protocol that combines passive CLARITY for improved tissue integrity and 2,2&#8242;-thiodiethanol (TDE), a previously described refractive index matching solution.

Optical clearing techniques are revolutionizing the way tissues are visualized. In this report we describe modifications of the original Clear Lipid-exchanged Acrylamide-hybridized Rigid Imaging-compatible Tissue-hYdrogel (CLARITY) protocol that yields more consistent and less expensive results.

Traditionally, tissue visualization has required that the tissue of interest be serially sectioned and imaged, subjecting each tissue section to unique ...

Microsurgical Dissection and Tissue Clearing for High Resolution Intact Whole Retina and Vitreous Imaging

Neuronal and vascular structures of the retina in physiologic and pathologic conditions can be better visualized and characterized by using intact whole retina imaging techniques compared to conventional retinal flat mount preparations and sections. However, immunofluorescent imaging of intact whole retina is hindered by the opaque coatings of the eyeball, i.e., sclera, choroid, and retinal pigment epithelium (RPE) and the light scattering properties of retinal layers that prevent full thickness high resolution optical imaging. Chemical bleaching of the pigmented layers and tissue clearing protocols have been described to address these obstacles; however, currently described methods are not suitable for imaging endogenous fluorescent molecules such as green fluorescent protein (GFP) in intact whole retina. Other approaches bypassed this limitation by surgical removal of pigmented layers and the anterior segment of the eyeball allowing intact eye imaging, though the peripheral retina and hyaloid structures were disrupted. Presented here is an intact whole retina and vitreous immunofluorescent imaging protocol that combines surgical dissection of the sclera/choroid/retina pigment epithelium (RPE) layers with a modified tissue clearing method and light sheet fluorescent microscopy (LSFM). The new approach offers an unprecedented view of&#160;unperturbed vascular and neuronal elements of the retina as well as the vitreous and hyaloid vascular system in pathologic conditions.

Presented here is a protocol for intact whole retina imaging in which the outer opaque/pigmented layers of the eyeball are surgically removed, and optical clearing is applied to render retina transparent enabling the visualization of the peripheral retina and hyaloid vasculature in intact retina using light sheet fluorescent microscopy.

Neuronal and vascular structures of the retina in physiologic and pathologic conditions can be better visualized and characterized by using intact whole ...

Optical Clearing and Imaging of Immunolabeled Kidney Tissue

Optical clearing techniques render tissue transparent by equilibrating the refractive index throughout a sample for subsequent three-dimensional (3-D) imaging. They have received great attention in all research areas for the potential to analyze microscopic multicellular structures that extend over macroscopic distances. Given that kidney tubules, vasculature, nerves, and glomeruli extend in many directions, which have been only partially captured by traditional two-dimensional techniques so far, tissue clearing also opened up many new areas of kidney research. The list of optical clearing methods is rapidly growing, but it remains difficult for beginners in this field to choose the best method for a given research question. Provided here is a simple method that combines antibody labeling of thick mouse kidney slices; optical clearing with cheap, non-toxic and ready-to-use chemical ethyl cinnamate; and confocal imaging. This protocol describes how to perfuse kidneys and use an antigen-retrieval step to increase antibody- binding without requiring specialized equipment. Its application is presented in imaging different multicellular structures within the kidney, and how to troubleshoot poor antibody penetration into tissue is addressed. We also discuss the potential difficulties of imaging endogenous fluorophores and acquiring very large samples and how to overcome them. This simple protocol provides an easy-to-setup and comprehensive tool to study tissue in three dimensions.

The combination of antibody labeling, optical clearing, and advanced light microscopy allows three-dimensional analysis of complete structures or organs. Described here is a simple method to combine immunolabeling of thick kidney slices, optical clearing with ethyl cinnamate, and confocal imaging that enables visualization and quantification of three-dimensional kidney structures.

Optical clearing techniques render tissue transparent by equilibrating the refractive index throughout a sample for subsequent three-dimensional (3-D) ...

Medicine

High-Resolution 3D Reconstruction of Human Brain with SHORT and LSFM

Optical Clearing and Labeling for Light-sheet Fluorescence Microscopy in Large-scale Human Brain Imaging

Despite the numerous clearing techniques that emerged in the last decade, processing postmortem human brains remains a challenging task due to its dimensions and complexity, which make imaging with micrometer resolution particularly difficult. This paper presents a protocol to perform the reconstruction of volumetric portions of the human brain by simultaneously processing tens of sections with the SHORT (SWITCH - H2O2 - Antigen Retrieval - 2,2'-thiodiethanol [TDE]) tissue transformation protocol, which enables clearing, labeling, and sequential imaging of the samples with light-sheet fluorescence microscopy (LSFM). SHORT provides rapid tissue clearing and homogeneous multi-labeling of thick slices with several neuronal markers, enabling the identification of different neuronal subpopulations in both white and grey matter. After clearing, the slices are imaged via LSFM with micrometer resolution and in multiple channels simultaneously for a rapid 3D reconstruction. By combining SHORT with LSFM analysis within a routinely high-throughput protocol, it is possible to obtain the 3D cytoarchitecture reconstruction of large volumetric areas at high resolution in a short time, thus enabling comprehensive structural characterization of the human brain.

Author Spotlight: Advancing 3D Cytoarchitecture Analysis - Rapid Volumetric Reconstruction of the Human Brain 

The present protocol provides a step-by-step procedure for rapid and simultaneous optical clearing, muti-round labeling, and 3D volumetric reconstruction of tens of postmortem human brain sections by combining the (SWITCH - H2O2 - Antigen Retrieval - 2,2'-thiodiethanol [TDE]) SHORT tissue transformation technique with light-sheet fluorescence microscopy imaging in a routinely high-throughput protocol.

Despite the numerous clearing techniques that emerged in the last decade, processing postmortem human brains remains a challenging task due to its ...

Deep Fluorescence Observation in Rice Shoots via Clearing Technology

The recently developed clearing technology that eliminates refractive index mismatches and diminishes auto-fluorescent material has made it possible to observe plant tissues in three dimensions (3D) while preserving their internal structures. In rice (Oryza sativa L.), a monocot model plant and a globally important crop, clearing technology has been reported in organs that are relatively easy to observe, such as the roots and leaves. Applications of clearing technology in shoot apical meristem (SAM) and stems have also been reported, but only to a limited degree because of the poor penetration of the clearing solution (CS) in these tissues. The limited efficiency of the clearing solutions in these tissues has been attributed to auto-fluorescence, thickening, and hardening of the tissues in the stem as the vascular bundles and epidermis develop and layering of the SAM with water-repellent leaves. The present protocol reports the optimization of a clearing approach for continuous and 3D observation of gene expression from the SAM/young panicle to the base of the shoots during development. Fixed tissue samples expressing a fluorescent protein reporter were trimmed into sections using a vibrating micro-slicer. When an appropriate thickness was achieved, the CS was applied. By specifically targeting the central tissue, the penetration rate and uniformity of the CS increased, and the time required to make the tissue transparent decreased. Additionally, clearing of the trimmed sections enabled the observation of the internal structure of the whole shoot from a macro perspective. This method has potential applications in deep imaging of tissues of other plant species that are difficult to clear.

Deep Fluorescence Observation in Rice Shoots via Clearing Technology

The present protocol describes a clearing technique for rice shoots, which are difficult to prepare for internal structural observations owing to the hard, thick, or layered nature of the tissues. This method facilitates continuous and deep fluorescence observations, even in adult rice plants.

The recently developed clearing technology that eliminates refractive index mismatches and diminishes auto-fluorescent material has made it possible to ...

Combining Clearing and Fluorescence Microscopy for Visualising Changes in Gene Expression and Physiological Responses to Plasmodiophora brassicae

Infection of Brassica crops by the soilborne protist Plasmodiophora brassicae leads to gall formation on the underground organs. The formation of galls requires cellular reprogramming and changes in the metabolism of the infected plant. This is necessary to establish a pathogen-oriented physiological sink toward which the host nutrients are redirected. For a complete understanding of this particular plant-pathogen interaction and the mechanisms by which host growth and development are subverted and repatterned, it is essential to track and observe the internal changes accompanying gall formation with cellular resolution. Methods combining fluorescent stains and fluorescent proteins are often employed to study anatomical and physiological responses in plants. Unfortunately, the large size of galls and their low transparency act as major hurdles in performing whole-mount observations under the microscope. Moreover, low transparency limits the employment of fluorescence microscopy to study clubroot disease progression and gall formation. This article presents an optimized method for fixing and clearing galls to facilitate epifluorescence and confocal microscopy for inspecting P. brassicae-infected galls. A tissue-clearing protocol for rapid optical clearing was used followed by vibratome sectioning to detect anatomical changes and localize gene expression with promoter fusions and reporter lines tagged with fluorescent proteins. This method will prove useful for studying cellular and physiological responses in other pathogen-triggered structures in plants, such as nematode-induced syncytia and root knots, as well as leaf galls and deformations caused by insects.

Combining Clearing and Fluorescence Microscopy for Visualising Changes in Gene Expression and Physiological Responses to Plasmodiophora brassicae

The present protocol describes an optimized method for the histological observation of galls induced by Plasmodiophora brassicae. Vibratome sections of hypocotyls are cleared before fluorescence imaging to study the involvement of transcription factors and phytohormones during disease progression. This protocol overcomes resin embedding limitations, enabling in planta visualization of fluorescent proteins.

Infection of Brassica crops by the soilborne protist Plasmodiophora brassicae leads to gall formation on the underground organs. The formation of galls ...

Optical Clearing of Plant Tissues for Fluorescence Imaging

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Combining Clearing and Fluorescence Microscopy for Visualising Changes in Gene Expression and Physiological Responses to Plasmodiophora brassicae