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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#215;18 No.1)</td>
			<td>MATSUNAMI</td>
			<td>C018181</td>
			<td></td>
		</tr>
		<tr>
			<td>Cover glass (24&#215;24 No.1)</td>
			<td>MATSUNAMI</td>
			<td>C024241</td>
			<td></td>
		</tr>
		<tr>
			<td>Cover glass (25&#215;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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5.0K 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

Proper Care and Cleaning of the Microscope

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a microscope causes reduction in contrast and resolution. DIC is especially sensitive to contamination and scratches on the lens surfaces. This protocol details the procedure for keeping the microscope clean.

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a microscope causes reduction in contrast and resolution. DIC is especially sensitive to contamination and scratches on the lens surfaces. This protocol details the procedure for keeping the microscope clean.

Keeping the microscope optics clean is important for high-quality imaging. Dust, fingerprints, excess immersion oil, or mounting medium on or in a ...

Major Components of the Light Microscope

The light microscope is a basic tool for the cell biologist, who should have a thorough understanding of how it works, how it should be aligned for different applications, and how it should be maintained as required to obtain maximum image-forming capacity and resolution. The components of the microscope are described in detail here.

The light microscope is a basic tool for the cell biologist, who should have a thorough understanding of how it works, how it should be aligned for different applications, and how it should be maintained as required to obtain maximum image-forming capacity and resolution. The components of the microscope are described in detail here.

The light microscope is a basic tool for the cell biologist, who should have a thorough understanding of how it works, how it should be aligned for ...

Phase Contrast and Differential Interference Contrast (DIC) Microscopy

Phase-contrast microscopy is often used to produce contrast for transparent, non light-absorbing, biological specimens. The technique was discovered by Zernike, in 1942, who received the Nobel prize for his achievement. DIC microscopy, introduced in the late 1960s, has been popular in biomedical research because it highlights edges of specimen structural detail, provides high-resolution optical sections of thick specimens including tissue cells, eggs, and embryos and does not suffer from the  phase halos  typical of phase-contrast images. This protocol highlights the principles and practical applications of these microscopy techniques.

Phase Contrast and Differential Interference Contrast DIC Microscopy

This protocol highlights the principles and practical applications of Phase and Differential Interference Contrast (DIC) Microscopy

Phase-contrast microscopy is often used to produce contrast for transparent, non light-absorbing, biological specimens. The technique was discovered by ...

Born Normalization for Fluorescence Optical Projection Tomography for Whole Heart Imaging

Optical projection tomography is a three-dimensional imaging technique that has been recently introduced as an imaging tool primarily in developmental biology and gene expression studies. The technique renders biological sample optically transparent by first dehydrating them and then placing in a mixture of benzyl alcohol and benzyl benzoate in a 2:1 ratio (BABB or Murray s Clear solution). The technique renders biological samples optically transparent by first dehydrating them in graded ethanol solutions then placing them in a mixture of benzyl alcohol and benzyl benzoate in a 2:1 ratio (BABB or Murray s Clear solution) to clear. After the clearing process the scattering contribution in the sample can be greatly reduced and made almost negligible while the absorption contribution cannot be eliminated completely. When trying to reconstruct the fluorescence distribution within the sample under investigation, this contribution affects the reconstructions and leads, inevitably, to image artifacts and quantification errors.. While absorption could be reduced further with a permanence of weeks or months in the clearing media, this will lead to progressive loss of fluorescence and to an unrealistically long sample processing time. This is true when reconstructing both exogenous contrast agents (molecular contrast agents) as well as endogenous contrast (e.g. reconstructions of genetically expressed fluorescent proteins).

We suggest a Born normalized approach for Optical Projection Tomography (BnOPT) that accounts for the absorption properties of imaged samples to obtain accurate and quantitative fluorescence tomographic reconstructions. We use the proposed algorithm to reconstruct the fluorescence molecular probe distribution within small animal organs.

Optical projection tomography is a three-dimensional imaging technique that has been recently introduced as an imaging tool primarily in developmental ...

Fluorescence Activated Cell Sorting of Plant Protoplasts

High-resolution, cell type-specific analysis of gene expression greatly enhances understanding of developmental regulation and responses to environmental stimuli in any multicellular organism. In situ hybridization and reporter gene visualization can to a limited extent be used to this end but for high resolution quantitative RT-PCR or high-throughput transcriptome-wide analysis the isolation of RNA from particular cell types is requisite. Cellular dissociation of tissue expressing a fluorescent protein marker in a specific cell type and subsequent Fluorescence Activated Cell Sorting (FACS) makes it possible to collect sufficient amounts of material for RNA extraction, cDNA synthesis/amplification and microarray analysis.
An extensive set of cell type-specific fluorescent reporter lines is available to the plant research community. In this case, two marker lines of the Arabidopsis thaliana root are used: PSCR::GFP (endodermis and quiescent center) and PWOX5::GFP (quiescent center). Large numbers (thousands) of seedlings are grown hydroponically or on agar plates and harvested to obtain enough root material for further analysis. Cellular dissociation of plant material is achieved by enzymatic digestion of the cell wall. This procedure makes use of high osmolarity-induced plasmolysis and commercially available cellulases, pectinases and hemicellulases to release protoplasts into solution.
FACS of GFP-positive cells makes use of the visualization of the green versus the red emission spectra of protoplasts excited by a 488 nm laser. GFP-positive protoplasts can be distinguished by their increased ratio of green to red emission. Protoplasts are typically sorted directly into RNA extraction buffer and stored for further processing at a later time.
This technique is revealed to be straightforward and practicable. Furthermore, it is shown that it can be used without difficulty to isolate sufficient numbers of cells for transcriptome analysis, even for very scarce cell types (e.g. quiescent center cells). Lastly, a growth setup for Arabidopsis seedlings is demonstrated that enables uncomplicated treatment of the plants prior to cell sorting (e.g. for the cell type-specific analysis of biotic or abiotic stress responses). Potential supplementary uses for FACS of plant protoplasts are discussed.

A method for isolating specific cell types from plant material is demonstrated. This technique employs transgenic marker lines expressing fluorescent proteins in particular cell types, cellular dissociation and Fluorescence Activated Cell Sorting. Additionally, a growth setup is established here that facilitates treatment of Arabidopsis thaliana seedlings prior to cell sorting.

High-resolution, cell type-specific analysis of gene expression greatly enhances understanding of developmental regulation and responses to environmental ...

Robotics and Dynamic Image Analysis for Studies of Gene Expression in Plant Tissues

Gene expression in plant tissues is typically studied by destructive extraction of compounds from plant tissues for in vitro analyses. The methods presented here utilize the green fluorescent protein (gfp) gene for continual monitoring of gene expression in the same pieces of tissues, over time. The gfp gene was placed under regulatory control of different promoters and introduced into lima bean cotyledonary tissues via particle bombardment. Cotyledons were then placed on a robotic image collection system, which consisted of a fluorescence dissecting microscope with a digital camera and a 2-dimensional robotics platform custom-designed to allow secure attachment of culture dishes. Images were collected from cotyledonary tissues every hour for 100 hours to generate expression profiles for each promoter. Each collected series of 100 images was first subjected to manual image alignment using ImageReady to make certain that GFP-expressing foci were consistently retained within selected fields of analysis. Specific regions of the series measuring 300 x 400 pixels, were then selected for further analysis to provide GFP Intensity measurements using ImageJ software. Batch images were separated into the red, green and blue channels and GFP-expressing areas were identified using the threshold feature of ImageJ. After subtracting the background fluorescence (subtraction of gray values of non-expressing pixels from every pixel) in the respective red and green channels, GFP intensity was calculated by multiplying the mean grayscale value per pixel by the total number of GFP-expressing pixels in each channel, and then adding those values for both the red and green channels. GFP Intensity values were collected for all 100 time points to yield expression profiles. Variations in GFP expression profiles resulted from differences in factors such as promoter strength, presence of a silencing suppressor, or nature of the promoter. In addition to quantification of GFP intensity, the image series were also used to generate time-lapse animations using ImageReady. Time-lapse animations revealed that the clear majority of cells displayed a relatively rapid increase in GFP expression, followed by a slow decline. Some cells occasionally displayed a sudden loss of fluorescence, which may be associated with rapid cell death. Apparent transport of GFP across the membrane and cell wall to adjacent cells was also observed. Time lapse animations provided additional information that could not otherwise be obtained using GFP Intensity profiles or single time point image collections.

We report a method for introduction, tracking and quantitative analysis of GFP expression in plant cells. This method utilizes a custom-designed robotics system for semi-continuous image collection from large numbers of samples, over time. We also demonstrate the use of ImageJ and ImageReady for analysis of image series.

Gene expression in plant tissues is typically studied by destructive extraction of compounds from plant tissues for in vitro analyses. The methods ...

Time-lapse Imaging of Mitosis After siRNA Transfection

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and cellular content. Hallmark events of mitosis, such as chromosome movement, can be readily tracked on an individual cell basis using time-lapse fluorescence microscopy of mammalian cell lines expressing specific GFP-tagged proteins. In combination with RNAi-based depletion, this can be a powerful method for pinpointing the stage(s) of mitosis where defects occur after levels of a particular protein have been lowered. In this protocol, we present a basic method for assessing the effect of depleting a potential mitotic regulatory protein on the timing of mitosis. Cells are transfected with siRNA, placed in a stage-top incubation chamber, and imaged using an automated fluorescence microscope. We describe how to use software to set up a time-lapse experiment, how to process the image sequences to make either still-image montages or movies, and how to quantify and analyze the timing of mitotic stages using a cell-line expressing mCherry-tagged histone H2B. Finally, we discuss important considerations for designing a time-lapse experiment. This strategy is complementary to other approaches and offers the advantages of 1) sensitivity to changes in kinetics that might not be observed when looking at cells as a population and 2) analysis of mitosis without the need to synchronize the cell cycle using drug treatments. The visual information from such imaging experiments not only allows the sub-stages of mitosis to be assessed, but can also provide unexpected insight that would not be apparent from cell cycle analysis by FACS.

Here we describe a basic protocol to image and quantify the mitotic timing of live mammalian tissue culture cells after siRNA transfection.

Changes in cellular organization and chromosome dynamics that occur during mitosis are tightly coordinated to ensure accurate inheritance of genomic and ...

Tandem High-pressure Freezing and Quick Freeze Substitution of Plant Tissues for Transmission Electron Microscopy

Since the 1940s transmission electron microscopy (TEM) has been providing biologists with ultra-high resolution images of biological materials. Yet, because of laborious and time-consuming protocols that also demand experience in preparation of artifact-free samples, TEM is not considered a user-friendly technique. Traditional sample preparation for TEM used chemical fixatives to preserve cellular structures. High-pressure freezing is the cryofixation of biological samples under high pressures to produce very fast cooling rates, thereby restricting ice formation, which is detrimental to the integrity of cellular ultrastructure. High-pressure freezing and freeze substitution are currently the methods of choice for producing the highest quality morphology in resin sections for TEM. These methods minimize the artifacts normally associated with conventional processing for TEM of thin sections. After cryofixation the frozen water in the sample is replaced with liquid organic solvent at low temperatures, a process called freeze substitution. Freeze substitution is typically carried out over several days in dedicated, costly equipment. A recent innovation allows the process to be completed in three hours, instead of the usual two days. This is typically followed by several more days of sample preparation that includes infiltration and embedding in epoxy resins before sectioning. Here we present a protocol combining high-pressure freezing and quick freeze substitution that enables plant sample fixation to be accomplished within hours. The protocol can readily be adapted for working with other tissues or organisms. Plant tissues are of special concern because of the presence of aerated spaces and water-filled vacuoles that impede ice-free freezing of water. In addition, the process of chemical fixation is especially long in plants due to cell walls impeding the penetration of the chemicals to deep within the tissues. Plant tissues are therefore particularly challenging, but this protocol is reliable and produces samples of the highest quality.

Obtaining high-quality transmission electron microscopy images is challenging, especially in the case of plant cells, which have abundant large water-filled vacuoles and aerated spaces. Tandem high-pressure freezing and quick freeze substitution greatly reduce preparation time of plant samples for TEM while producing samples with excellent ultrastructural preservation.

Since the 1940s transmission electron microscopy (TEM) has been providing biologists with ultra-high resolution images of biological materials. Yet, ...

Visualizing Stromule Frequency with Fluorescence Microscopy

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but whose functions remain mysterious. Alongside growing attention on the role of chloroplasts in coordinating plant responses to stress, interest in stromules and their relationship to chloroplast signaling dynamics has increased in recent years, aided by advances in fluorescence microscopy and protein fluorophores that allow for rapid, accurate visualization of stromule dynamics. Here, we provide detailed protocols to assay stromule frequency in the epidermal chloroplasts of Nicotiana benthamiana, an excellent model system for investigating chloroplast stromule biology. We also provide methods for visualizing chloroplast stromules in vitro by extracting chloroplasts from leaves. Finally, we outline sampling strategies and statistical approaches to analyze differences in stromule frequencies in response to stimuli, such as environmental stress, chemical treatments, or gene silencing. Researchers can use these protocols as a starting point to develop new methods for innovative experiments to explore how and why chloroplasts make stromules.

Protocols to investigate the dynamics of chloroplast stromules, the stroma-filled tubules that extend from the surface of chloroplasts, are described.

Stromules, or "stroma-filled tubules", are narrow, tubular extensions from the surface of the chloroplast that are universally observed in plant cells but ...

ACT-PRESTO: Biological Tissue Clearing and Immunolabeling Methods for Volume Imaging

The identification and exploration of the detailed organization of organs or of the whole body at the cellular level are fundamental challenges in biology. Transitional methods require a substantial amount of time and effort to obtain a 3D image and including sectioning the intact tissue, immunolabeling, and imaging serially-sectioned tissue, which produces a loss of information at each step of the process. In recently developed approaches for high-resolution imaging within intact tissue, molecular characterization has been restricted to the labeling of proteins. However, currently available protocols for organ clearing require a considerably long process time, making it difficult to implement tissue clearing techniques in the lab. We recently established a rapid and highly-reproducible protocol termed ACT-PRESTO (active clarity technique&#8211;pressure related efficient and stable transfer of macromolecules into organs), which allows for tissue clearance within several hours. Moreover, ACT-PRESTO enables rapid immunolabeling with conventional methods and accelerates antibody penetration into the deep layer of densely-formed, thick specimens by applying pressure or convection flow. We describe how to prepare tissues, how to clear by lipid removal using electrophoresis, and how to immuno-stain by a pressure-assisted delivery. The rapidity and consistency of the protocol will expedite the performance of 3D histological research and volume-based diagnoses.

ACT-PRESTO (active clarity technique&#8211;pressure related efficient and stable transfer of macromolecules into organs) enables rapid, efficient, but inexpensive tissue clearing and rapid antibody penetration into cleared tissues by passive diffusion or pressure-assisted delivery. Using this method, a wide range of tissues can be cleared, immunolabeled by multiple antibodies, and volume-imaged.