Name: the cutting and floating method for paraffin-embedded tissue for sectioning
Uploaded: 2018-09-05T15:30:00.000+00:00
Duration: 5 min
Description: Watch this Scientific Journal Video about The Cutting and Floating Method for Paraffin-embedded Tissue for Sectioning at JoVE.com

This work was supported by the National Natural Science Foundation of China (Grant No. 31400936, 31460260) and the Natural Science Foundation of Jiangxi Province of China (20171BAB215020). We also thank the joint program between Nanchang University and Queen Mary University of London for supporting this work.

All methods described here have been approved by the Animal Care and Use Committee of Nanchang University.
1. Assemble the Equipment and Connect the Microtome
<ol>
	<li>Design the parameter per the requirements (Supplementary Figure 1).</li>
	<li>Submit the parameter to a local factory to manufacture the acrylic boards.</li>
	<li>Assemble all parts in sequence: Use chloroform to combine 7 commercial acrylic boards into a tank with a section channel and water bath (Figure 1). 
	CAUTION: Chloroform produces toxic substances when it meets light and oxygen. Assemble the apparatus in a fume hood.</li>
	<li>Install the tubular electric heating element through the hole in acrylic board #1 (Figure 2A, arrowhead).</li>
	<li>To install the temperature controller, create a small hole on the #2 acrylic board using an electric drill, and install the temperature detector (Figure 2A, arrow). Install a digital thermostat on the tank (Figure 2A). 
	NOTE: The temperature detector should be within 1 cm below the water surface.</li>
	<li>To provide power access, adjust the voltage below 24 V before turning on the equipment.</li>
	<li>Remove the section waste tray from the microtome.</li>
	<li>Link the sections channel with the microtome blade holder with neutral silicone sealant (Figure 2B). 
	NOTE: Do not smear any neutral silicone sealant above the top of the blade so that the old blade is easily replaced.</li>
	<li>Dry overnight to allow the chloroform and neutral silicone sealant time to solidify.</li>
	<li>Install the blade to the blade holder. Pour water into the water bath (Figure 2C). The water surface should just touch the top of the blade (Figure 2D, arrow head).</li>
</ol>
2. Paraffin Sectioning
NOTE: Debris often accumulate on the upper or lower edges of the wax block and the water surface. This debris should be cleared regularly.
<ol>
	<li>Perform Conventional paraffin sectioning10.
	<ol>
		<li>Clamp the paraffin-embedded specimen on a rotary microtome and place the blade in a blade holder.</li>
		<li>Turn the handwheel and section the specimen.</li>
		<li>Transfer the paraffin sectioned ribbon to the 38.5 &#176;C warm water bath.</li>
		<li>Pick up the sections onto the microscope slide from the water surface after unfolding.</li>
		<li>Dry the slides in an oven at 42 &#176;C overnight.</li>
	</ol>
	</li>
	<li>Improved paraffin sectioning (Figure 3)
	<ol>
		<li>Turn on the equipment.</li>
		<li>Open the heater with temperature detection switch and set the targeted temperature between 38.0 &#176;C to 40.0 &#176;C for 1 h in advance to warm the water.</li>
		<li>Modify the paraffin block acquired by paraffin embedding; then place the paraffin block on the microtome&#39;s cassette clamp and lock it. 
		NOTE: The superior and inferior borders of the paraffin block should be modified horizontally to prevent the section ribbon from forming an arc and becoming unable to flow through the narrow sections channel.</li>
		<li>Quickly approach the sample by cutting thicker sections.</li>
		<li>Adjust to a suitable section thickness (10 &#181;m for adult mouse brains, adult mouse kidneys, and embryo mouse brains, 4 &#181;m for zebrafish eyes). Turn the handwheel and the sections will automatically enter the sections channel and bath.</li>
		<li>Section the specimen in a slow and uniform cutting stroke (Figure 3A). 
		NOTE: Thethickness of the first section will be altered after the handwheel has been stopped for a few minutes. The disposable blades should be replaced when the sections have consecutive scratches.</li>
		<li>Guide the sections into the bath chamber to float in a line (Figure 3B, 3C). 
		NOTE: If the head section adheres to the chamber wall, use a brush guide to keep it floating.</li>
		<li>Adhere sections onto microscope slides. After the sections fully unfold in the water bath, separate several sections with tweezers. Then pick them up from the water bath onto the microscope slides (Figure 3D). 
		NOTE: Sections should be picked up from the posterior of the water bath, as those in the sections channel and the anterior of the water bath will not yet be fully unfolded.</li>
		<li>Dry the slides by placing them in a 42 &#176;C oven overnight. The slides can then be stored at room temperature indefinitely.</li>
	</ol>
	</li>
	<li>Use Hematoxylin-Eosin (HE) staining for the sake of evaluation of the improved method8.</li>
</ol>

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The authors have a patent on this device and declare no competing interests.

To improve paraffin section morphology and solve the problem of wasted time during conventional paraffin sectioning, we created an improved paraffin sectioning method that combines cutting and unfolding. This improved method relies on simple equipment that comprises a section channel, a water bath and a heater with a temperature detection switch. The section ribbon enters the water bath through the section channel and unfolds automatically while cutting. Therefore, this method improves paraffin sectioning quality and efficiency, which was verified by paraffin sectioning of the adult mouse hippocampus, adult mouse kidneys, E15.5 mouse brains, and adult zebrafish eyes.
To avoid section curling during conventional paraffin sectioning, many adjustments have been made, such as moistening the tissue surface, cooling the wax block, increasing the humidity, and combining cryosectioning6,7,8; however, these adjustments make paraffin sectioning more tedious. A commercial section transfer system that relies on fluid water has been more successful. In this system, the section contacts the water surface immediately after cutting, and the electric motor-driven water stream transfers the sections from the blade holder to the lower warm water bath9. Due to its mechanism, static electricity does not inhibit the sectioning, resulting in improved section quality. In addition, as this system avoids manually transferring sections from the blade holder to the water bath, easily damaging fragile sections, it also helps in acquiring high-quality thinner sections.
With our improved method, the simple equipment draws on the experience of the commercial section transfer system with some advantages. First, the equipment is cheaper, which could popularize this method in laboratories in developing regions and may complement the commercial section transfer system. Second, the water in the equipment is static and never disturbs the sections, making it possible to acquire thinner sections. We consecutively sectioned zebrafish eyes of 2 &#181;m thickness using the improved method, and the acquired sections were of higher quality. Third, a larger water bath tank is used in the equipment, which permits at least 40 sections to unfold and float simultaneously, allowing two operators to cooperate. One person can cut the tissue specimen continuously and the other can directly fish up the sections. This reduces the time waiting for sections to unfold. Successive cutting with a uniform cutting stroke also improves section quality.
The structure and appearance of our simple equipment requires improvement. For example, the components, such as the power, a heater, and a temperature controller, can be integrated to make this equipment more integral and portable. Additionally, commercial rotary microtomes have different sizes. To connect to other microtomes, the angle and height of this equipment should be adjusted. Paraffin sectioning combined with other molecular techniques is widely used in neurobiology, tissue morphology and other research fields11,12,13. Potentially, this method can improve clinical inspection efficiency14,15 and other biological specimens beyond the three tissue types used here.

Morphological study is important in the biological research. Although new technology has allowed researchers to observe their targets directly from the whole tissue or organisms1,2,3, cutting the specimen into thin sections, followed by staining, remains the primary method fornot only tissue morphology but also protein targeting directly in the tissue. Light microscopy uses three section types: paraffin, frozen, and semithin. Although cryosectioning is common for protecting tissue antigenicity, and the specimen preparation is simple, the retained tissue morphology is poor and unsuitable for thin sectioning4,5. Paraffin sectioning is the most frequently used method for exhibiting well preserved morphology. As the specimens are dehydrated completely and embedded in wax, the paraffin blocks can be stored indefinitely. In addition, paraffin sectioning produces thin sections that improve biological probe access in further experiments and reduce cell layer overlay in the Z direction.
However, conventional paraffin sectioning is tedious and demands operator skill. Paraffin sections undergo fixation, dehydration, embedding, cutting, and floating. Importantly, transferring section ribbons from the knife holder to the water bath is necessary but difficult for junior operators. Especially in dry air, the section ribbons will twist due to static electricity and are difficult to unfold on the warm water surface. To improve section quality, moistening the exposed tissue surface between microtome blade passes, cooling the wax blocks by immersing them in ice water, or raising the humidity with a humidifier near the microtome are recommended6,7. Newer methods for improving paraffin sectioning include hybrid paraffin embedding, cryosectioning8, and commercial section transfer system assistance9. Although these methods partially improve paraffin sectioning speed and quality, they make sectioning much more cumbersome, and commercial section transfer systems are expensive.
In this protocol, we demonstrate how to create simple, cheap and flexible equipment step by step, which can be connected to the blade holder of a rotary microtome. This equipment is comprised of a section channel, a water bath, and a heater with a temperature detection switch. After cutting, dozens of sections flow into the section channel and enter the water bath directly, thus unfolding automatically. This improves the efficiency of paraffin sectioning and makes this technology more convenient. Using this method, more adult mouse hippocampal sections, adult mouse kidney sections, embryonic 15.5 day-old (E15.5) mouse brain sections, and adult zebrafish eye sections were harvested in less time and remained more intact morphologically. This method can also be used for other tissue samples that require accelerated paraffin sectioning while avoiding loss of section distinction.

<table><tbody><tr><td>Incubator</td><td>Boekel Scientific</td><td>133000-2</td><td></td></tr><tr><td>Ethanol&amp;nbsp;</td><td>Sinopharm Chemical Reagent Co.,Lid</td><td>64-17-5</td><td></td></tr><tr><td>Xylene&amp;nbsp;</td><td>Sinopharm Chemical Reagent Co.,Lid</td><td>1330-20-7</td><td></td></tr><tr><td>Paraplast</td><td>Leica</td><td>39601006</td><td></td></tr><tr><td>Heated Paraffin Embedding Module</td><td>Leica</td><td></td><td></td></tr><tr><td>Commercial acrylic board</td><td></td><td></td><td></td></tr><tr><td>Trichloromethane</td><td>Sinopharm Chemical Reagent Co.,Lid</td><td>67-66-3</td><td></td></tr><tr><td>Tubular electric heating element(12 V 200 W)</td><td></td><td></td><td></td></tr><tr><td>Temperature controller(12 V 120 W)</td><td>Mingsuo</td><td>XH-W3002</td><td></td></tr><tr><td>Rotary microtome&amp;nbsp;</td><td>Leica</td><td></td><td></td></tr><tr><td>Neutral silicone sealant</td><td></td><td></td><td>Link the water channel with the microtome knife holder</td></tr><tr><td>Voltage transformer</td><td>Dearll</td><td>S-250-12</td><td></td></tr><tr><td>Disposable blade</td><td>Accu-Edge</td><td>4689</td><td></td></tr><tr><td>Hematoxylin</td><td>Baso Diagnostics Inc.</td><td>BA-4025</td><td></td></tr><tr><td>Eosin&amp;nbsp;</td><td>Baso Diagnostics Inc.</td><td>BA-4025</td><td></td></tr><tr><td>Microslide</td><td>Sail Brand</td><td>7105</td><td></td></tr><tr><td>Neutral balsam</td><td>Sinopharm Chemical Reagent Co.,Lid</td><td>10004160</td><td></td></tr><tr><td>Coverslip&amp;nbsp;</td><td>Citoglas</td><td>10212424C</td><td></td></tr><tr><td>Microscope</td><td>Carl Zeiss</td><td></td><td></td></tr><tr><td>Hydrochloric acid</td><td>Xilong Chemical</td><td>7647-01-0</td><td></td></tr><tr><td>Water bath for paraffin sections</td><td>Leica</td><td></td><td></td></tr><tr><td>HistoCore Arcadia C&amp;nbsp;- Cold Plate</td><td>Leica</td><td></td><td></td></tr><tr><td>paraffin repellent spray&amp;nbsp;</td><td>Thermo Scientific</td><td>9990420</td><td></td></tr></tbody></table>

the cutting and floating method for paraffin-embedded tissue for sectioning

Sectioning of the paraffin-embedded tissue is widely used in histology and pathology. However, it is tedious. To improve this method, several commercial companies have devised complex section transfer systems using fluid water. To simplify this technology, we created a simple method using homemade equipment that combines cutting and floating within a simple thermostatic chamber; therefore, the sections automatically enter the water bath on the water surface. The hippocampus from adult mouse brains, adult mouse kidneys, embryonic mouse brains, and adult zebrafish eyes were cut using both conventional paraffin sectioning and the presented method for comparison. Statistical analysis shows that our improved method saved time and produced higher quality sections. In addition, paraffin sectioning of a whole specimen in a short time is easy for junior operators.

The improved method increased the number of intact paraffin sections. We tested this new method on adult mouse hippocampal tissue, adult mouse kidneys, embryonic mouse brains, and zebrafish eyes. Water was added to the tank, and the water temperature was maintained between 38.0 &#176;C to 40.0 &#176;C. After a serially preparing the tissue samples, they were sectioned and compared to conventional sectioning. The new method avoided section loss and increased the proportion of intact sections in the adult mouse hippocampus, adult mouse kidneys, E15.5 mouse brains, and adult zebrafish eyes (Figure 4A, 4C, 4E, 4G). In conclusion, our method improved the sectioning by preventing section loss and harvesting more intact sections per specimen.
The improved method decreased the time spent per specimen. In comparing the time taken between the conventional method and our improved method, we performed whole-series sectioning (from the first section to the last section of an organ) on the adult mouse hippocampus, adult mouse kidneys, E15.5 mouse brains, and adult zebrafish eyes. The results suggest that the improved method accelerated the paraffin sectioning speed in all tested samples (Figure 4B, 4D, 4F, 4H). In general, our method allowed for faster high-quality paraffin sectioning.
Series sections of the zebrafish optic retina was observed by H and E staining. For series sectioning of the optic disc segment of the zebrafish retina,the sections harvested from the improved method were better has better integrity than those from the conventional paraffin sectioning method (Figure 5A, 5B). There was no significant difference between these two methods regarding the quality of most harvested sections (Figure 5C, 5D). Occasionally, the quality of sections harvested from the conventional method were relatively low (Figure 5E, 5F).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58288/58288fig1.jpg" /> 
Figure 1: The process for assembling the 7 commercial acrylic boards into a tank. (A) Overall view of the 7 acrylic boards. (B-G) Stepwise assembly of the tank from board # 1 to board #7. (H) Primary tank after assembly. (I) General tank size; the protective membrane on the tank surface was removed for further installation. L = 400 mm, L&#39; = 220 mm, H = 122 mm, W = 200 mm. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58288/58288fig1large.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/58288/58288fig2.jpg" /> 
Figure 2: The appropriately assembled equipment ready for use. (A) The intact equipment after being connected to the rotary microtome. Arrow indication, the temperature detector is installed through the small hole by electric drill. Arrow head, the heater (tubular electric heating element) is installed through the hole in board #1. (B) The amplified dashed area in A, the feature of the connection between the microtome blade holder and the sections channel. Arrow head, the blade holder and sections channel are connected by neutral silicone sealant. (C) Same as B, with the water poured and blade installed. (D) Side view of the dashed area in C. Arrow head, the water surface is close to the blade edge. VT: voltage transformer; WB: water bath; DT: digital thermostat; B: blade; BH: blade holder; SC: sections channel. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58288/58288fig2large.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/58288/58288fig3.jpg" /> 
Figure 3: The improved paraffin sectioning process. The figures located at the lower left corner are the amplified dashed areas in A-D respectively. Arrow indications, the first section. (A, A&#39;) The section enters the sections channel immediately and directly after cutting. (B, B&#39;) The sections ribbon crosses the border between the sections channel and water bath. (C, C&#39;) The sections float to the posterior of the water bath and are fully unfolded. (D, D&#39;) The sections are picked up by microscope slide when they reach the posterior of the water bath and are fully unfolded. SC: sections channel; WB, water bath. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58288/58288fig3large.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/58288/58288fig4.jpg" /> 
Figure 4: The improved sectioning method improved the efficiency and yield rate significantly for whole-series sections. The yield rate equaled the proportion of intact sections in all cut sections. Data are shown as the mean &#177; SEM and acquired by the same operator. (A, C, E, G) The improved method prevented section loss of the adult mouse hippocampus (91.04 &#177; 1.429% N = 3, vs 97.63 &#177; 0.4864% N = 3, p = 0.0120), adult mouse kidneys (92.30 &#177; 0.5689% N = 3, vs 95.99 &#177; 0.7055% N = 3, p = 0.0153), E15.5 mouse brain (90.67 &#177; 1.549%, N = 3 vs 97.01 &#177; 0.2816%, N = 3, p = 0.0158), and adult zebrafish eyes (91.27 &#177; 0.9734% N = 4 vs 96.32 &#177; 0.7217% N = 4, p = 0.0059). (B, D, F, H) The improved method saved time compared with the conventional method for the adult mouse hippocampus (126.0 &#177; 3.606 min vs 79.67 &#177; 1.856 min, N = 3, p = 0.0003), adult mouse kidneys (66.67 &#177; 1.764 min vs 41.33 &#177; 2.186 min, N = 3, p = 0.0008), E15.5 mouse brain (99.67 &#177; 3.480 min vs 52.33 &#177; 4.333 min, N = 3, p = 0.0010), and adult zebrafish eyes (48.00 &#177; 2.160 min vs 30.75 &#177; 1.797 min, N = 4, p = 0.0009). * P &#60; 0.05, ** P &#60; 0.01, *** P &#60; 0.005. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58288/58288fig4large.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/58288/58288fig5.jpg" /> 
Figure 5: H and E staining of the whole-series sections of the optic disc segment of the zebrafish eyes acquired from the conventional methods and improved paraffin sectioning by the same operator are compared. (A) Conventional paraffin sectioning (21 sections). (B) Improved paraffin sectioning (24 sections). (C) Dashed area of A6. (D) Dashed area of B7. (E) Dashed area of A17. (F) Dashed area of B19. Asterisks, the sections with relatively poor quality. Arrow heads, optic disc; Arrows, severe tear of retinal tissues. Scale bar = 200 &#181;m (A and B); Scale bar =&#160;50 &#181;m&#160;(C, D, E, F). <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58288/58288fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplementary Figure 1: The dimension parameters of the 7 commercial acrylic boards. The number represents the length of corresponding board margin. Units: mm. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58288/supplementary_figure.AI">Please click here to download this file.</a>

Watch this Scientific Journal Video about The Cutting and Floating Method for Paraffin-embedded Tissue for Sectioning at JoVE.com

The Cutting and Floating Method for Paraffin-embedded Tissue for Sectioning

Here, we present a protocol to improve paraffin sectioning. This method combines cutting and floating using a simple thermostatic chamber to avoid the transfer process required by the conventional method. As a result, the efficiency and the number of intact paraffin sections were greatly improved.

Sectioning of the paraffin-embedded tissue is widely used in histology and pathology. However, it is tedious. To improve this method, several commercial ...

the-cutting-floating-method-for-paraffin-embedded-tissue-for

Universidad San Sebastian

Research

JoVE Journal

Neuroscience

18.2K Views.  Nanchang University. Here, we present a protocol to improve paraffin sectioning. This method combines cutting and floating using a simple thermostatic chamber to avoid the transfer process required by the conventional method. As a result, the efficiency and the number of intact paraffin sections were greatly improved.

Video: The Cutting and Floating Method for Paraffin-embedded Tissue for Sectioning

Thin Sectioning of Slice Preparations for Immunohistochemistry

Many investigations in neuroscience, as well as other disciplines, involve studying small, yet macroscopic pieces or sections of tissue that have been preserved, freshly removed, or excised but kept viable, as in slice preparations of brain tissue.  Subsequent microscopic studies of this material can be challenging, as the tissue samples may be difficult to handle.  Demonstrated here is a method for obtaining thin cryostat sections of tissue with a thickness that may range from 0.2-5.0 mm.  We routinely cut 400 micron thick Vibratome brain slices serially into 5-10 micron coronal cryostat sections.  The slices are typically first used for electrophysiology experiments and then require microscopic analysis of the cytoarchitecture of the region from which the recordings were observed.  We have constructed a simple device that allows controlled and reproducible preparation and positioning of the tissue slice.  This device consists of a cylinder 5 cm in length with a diameter of 1.2 cm, which serves as a freezing stage for the slice.  A ring snugly slides over the cylinder providing walls around the slice allowing the tissue to be immersed in freezing compound (e.g., OCT).  This is then quickly frozen with crushed dry ice and the resulting  wafer  can be position easily for cryostat sectioning.  Thin sections can be thaw-mounted onto coated slides to allow further studies to be performed, such as various staining methods, in situ hybridization, or immunohistochemistry, as demonstrated here.

The present method allows reproducible cryostat sectioning of small, difficult-to-manage, tissue pieces, such as biopsies and brain slices. We utilize a simple aluminum freezing stage to facilitate handling of tissue and a standard cryostat to routinely produce 5-10 micron serial sections from 400 micron thick brain slices.

Many investigations in neuroscience, as well as other disciplines, involve studying small, yet macroscopic pieces or sections of tissue that have been ...

Biology

Sectioning Mammary Gland Whole Mounts for Lesion Identification

Normal mammary gland development may be altered by exposure to environmental toxicants and pharmaceutical products, excessive exposure to hormones, and genetic alterations. Mammary gland whole mounts are an inexpensive method to capture the progression of morphological changes that may arise after exposure. However, in later life, when abnormalities are more prone to develop, sole reliance on this one method may not always provide enough information to make a proper diagnosis of the abnormality. Historically, in chemical test guideline studies, a single mammary gland is removed at necropsy and prepared as a hematoxylin and eosin (H&#38;E)-stained section. The incorporation of contralateral mammary whole-mount collection and analysis decreases the likelihood of a false-negative assessment. Evaluation of the whole mount is limited by the presence of one or two entire mammary glands on a slide, and in some cases, the abnormalities observed in the whole mount are not uniformly represented in the H&#38;E section.&#160; The goal of this study was to develop a protocol for converting coverslipped mammary whole mounts to H&#38;E-stained sections so that lesions that would otherwise have been missed or that are difficult to diagnose can be identified. Here, we detail a method to produce a high-quality, paraffin-embedded H&#38;E section from a mammary gland that was initially prepared as a whole mount. In comparison to a tissue that was intentionally prepared for H&#38;E sectioning, the whole mount requires additional preparation for tissue removal and processing. However, this method is considered inexpensive, as it requires common lab reagents and little additional time. As a result, this method can provide invaluable information on how chemical and environmental exposures alter normal mammary development, as well as display changes that occur because of genetic modifications.

We developed a method to successfully remove, process, section, and stain, for histopathological evaluation, mammary tissue that had originally been fixed on slides as whole mounts. This method may promote the collection and evaluation of mammary gland whole mounts in reproductive and developmental test guideline studies.

Normal mammary gland development may be altered by exposure to environmental toxicants and pharmaceutical products, excessive exposure to hormones, and ...

Cancer Research

Paraffin Embedding and Thin Sectioning of Microbial Colony Biofilms for Microscopic Analysis

Sectioning via paraffin embedding is a broadly established technique in eukaryotic systems. Here we provide a method for the fixation, embedding, and sectioning of intact microbial colony biofilms using perfused paraffin wax. To adapt this method for use on colony biofilms, we developed techniques for maintaining each sample on its growth substrate and laminating it with an agar overlayer, and added lysine to the fixative solution. These optimizations improve sample retention and preservation of micromorphological features. Samples prepared in this manner are amenable to thin sectioning and imaging by light, fluorescence, and transmission electron microscopy. We have applied this technique to colony biofilms of Pseudomonas aeruginosa, Pseudomonas synxantha, Bacillus subtilis, and Vibrio cholerae. The high level of detail visible in samples generated by this method, combined with reporter strain engineering or the use of specific dyes, can provide exciting insights into the physiology and development of microbial communities.

We describe fixation, paraffin embedding, and thin sectioning techniques for microbial colony biofilms. In prepared samples, biofilm substructure and reporter expression patterns can be visualized by microscopy.

Sectioning via paraffin embedding is a broadly established technique in eukaryotic systems. Here we provide a method for the fixation, embedding, and ...

Immunology and Infection

Preparation of Non-human Primate Brain Tissue for Pre-embedding Immunohistochemistry and Electron Microscopy

Despite all the technological advances at the light microscopy&#160;level, electron microscopy remains the only tool in neuroscience to examine and characterize ultrastructural and morphological details of neurons, such as synaptic contacts. Good preservation of brain tissue for electron microscopy can be obtained by rigorous cryo-fixation methods, but these techniques are rather costly and limit the use of immunolabeling, which is crucial to understand the connectivity of identified neuronal systems. Freeze-substitution methods have been developed to allow the combination of cryo-fixation with immunolabeling. However, the reproducibility of these methodological approaches usually relies on costly freezing devices. Moreover, achieving reliable results with this technique is very time-consuming and skill-challenging. Hence, the traditional chemically fixed brain, particularly with acrolein fixative, remains a time-efficient and low-cost method to combine electron microscopy with immunohistochemistry. Here, we provide a reliable experimental protocol using chemical acrolein fixation that leads to the preservation of primate brain tissue and is compatible with pre-embedding immunohistochemistry and transmission electron microscopic examination.

Here, we provide an easy, low-cost, and time-efficient protocol to chemically fix primate brain tissue with acrolein fixative, allowing for long-term preservation that is compatible with pre-embedding immunohistochemistry for transmission electron microscopy.

Despite all the technological advances at the light microscopy&#160;level, electron microscopy remains the only tool in neuroscience to examine and ...

Establishment and Analysis of Tumor Slice Explants As a Prerequisite for Diagnostic Testing

Organotypic primary tissue explant cultures, which include precision-cut slices, represent the three-dimensional (3-D) tissue architecture as well as the multicellular interactions of native tissue. Tissue&#160;slices immediately cut from freshly resected tumors preserve spatial aspects of intratumor heterogeneity (ITH), thus making them useful surrogates of in vivo biology. Careful optimization of tissue slice preparation and cultivation conditions is fundamental for the predictive diagnostic potential of tumor slice explants. In this regard, murine models are valuable, as these provide a consistent flow of tumor material to perform replicate and reproducible experiments. This protocol describes the culturing of murine lung tumor-derived tissue slices using a rotating incubation unit, a system that enables intermittent exposure of tissues to oxygen and nutrients. Our previous work showed that the use of rotating incubation units improves the viability of tissue compared to other culture methods, particularly floating slices and stagnant filter supports. Here, we further show that slice thickness influences the viability of cultured slices, suggesting that optimization of slice thickness should be done for different tissue types. Pronounced ITH in relevant oncogenic functions, such as signaling activities, stromal cell infiltration or expression of differentiation markers, necessitates evaluation of adjacent tissue slices for the expression of markers altered by drug treatment or cultivation itself. In summary, this protocol describes the generation of murine lung tumor slices and their culture on a rotating incubation unit and demonstrates how slices should be systematically analyzed for the expression of heterogeneous tissue markers, as a prerequisite prior to drug response studies.

We provide a method for the generation, cultivation and systematic analysis of organotypic slices derived from murine lung tumors. We also describe how to optimize for slice thickness, and how to select drug concentrations to treat tumor slices.

Organotypic primary tissue explant cultures, which include precision-cut slices, represent the three-dimensional (3-D) tissue architecture as well as the ...

Hybrid-Cut: An Improved Sectioning Method for Recalcitrant Plant Tissue Samples

Maintaining plant section integrity is essential for studying detailed anatomical structures at the cellular, tissue, or even organ level. However, some plant cells have rigid cell walls, tough fibers and crystals (calcium oxalate, silica, etc.), and high water content that often disrupt tissue integrity during plant tissue sectioning.
This study establishes a simple Hybrid-Cut tissue sectioning method. This protocol modifies a paraffin-based sectioning technique and improves the integrity of tissue sections from different plants. Plant tissues were embedded in paraffin before sectioning in a cryostat at -16 &#176;C. Sectioning under low temperature hardened the paraffin blocks, reduced tearing and scratching, and improved tissue integrity significantly. This protocol was successfully applied to calcium oxalate-rich Phalaenopsis orchid tissues as well as recalcitrant tissues such as reproductive organs and leaves of rice, maize, and wheat. In addition, the high quality of tissue sections from Hybrid-Cut could be used in combination with in situ hybridization (ISH) to provide spatial expression patterns of genes of interest. In conclusion, this protocol is particularly useful for recalcitrant plant tissue containing high crystal or silica content. Good quality tissue sections enable morphological and other biological studies.

This protocol describes a simple Hybrid-Cut tissue sectioning method that is useful for recalcitrant plant tissues. Good quality tissue sections enable anatomical studies and other biological studies including in situ hybridization (ISH).

Maintaining plant section integrity is essential for studying detailed anatomical structures at the cellular, tissue, or even organ level. However, some ...

Pancreatic Islet Embedding for Paraffin Sections

Experiments using isolated pancreatic islets are important for diabetes research, but islets are expensive and of limited abundance. Islets contain a mixed cell population in a structured architecture that impacts function, and human islets are widely variable in cell type composition. Current frequently used methods to study cultured islets include molecular studies performed on whole islets, lumping disparate islet cell types together, or microscopy or molecular studies on dispersed islet cells, disrupting islet architecture. For in vivo islet studies, paraffin-embedded pancreas sectioning is a powerful technique to assess cell-specific outcomes in the native pancreatic environment. Studying post-culture islets by paraffin sectioning would offer several advantages: detection of multiple outcomes on the same islets (potentially even the exact-same islets, using serial sections), cell-type-specific measurements, and maintaining native islet cell-cell and cell-substratum interactions both during experimental exposure and for analysis. However, existing techniques for embedding isolated islets post-culture are inefficient, time consuming, prone to loss of material, and generally produce sections with inadequate islet numbers to be useful for quantifying outcomes. Clinical pathology laboratory cell block preparation facilities are inaccessible and impractical for basic research laboratories. We have developed an improved, simplified bench-top method that generates sections with robust yield and distribution of islets. Fixed islets are resuspended in warm histological agarose gel and pipetted into a flat disc on a standard glass slide, such that the islets are distributed in a plane. After standard dehydration and embedding, multiple (10+) 4 - 5 &#181;m sections can be cut from the same islet block. Using this method, histological&#160;and immunofluorescent analyses can be performed on mouse, rat, and human islets. This is an effective, inexpensive, time-saving approach to assess cell-type-specific, intact-architecture outcomes from cultured islets.

Ex vivo pancreatic islet studies are important for diabetes research. Existing techniques to study cultured islets in their native 3-dimensional architecture are time consuming, inefficient, and infrequently used. This work describes a new, simple, and efficient method for generating high-quality paraffin sections of whole cultured islets.

Experiments using isolated pancreatic islets are important for diabetes research, but islets are expensive and of limited abundance. Islets contain a ...

Microscopy Techniques for Interpreting Fungal Colonization in Mycoheterotrophic Plants Tissues and Symbiotic Germination of Seeds

Structural botany is an indispensable perspective to fully understand the ecology, physiology, development, and evolution of plants. When researching mycoheterotrophic plants (i.e., plants that obtain carbon from fungi), remarkable aspects of their structural adaptations, the patterns of tissue colonization by fungi, and the morphoanatomy of subterranean organs can enlighten their developmental strategies and their relationships with hyphae, the source of nutrients. Another important role of symbiotic fungi is related to the germination of orchid seeds; all Orchidaceae species are mycoheterotrophic during germination and seedling stage (initial mycoheterotrophy), even the ones that photosynthesize in adult stages. Due to the lack of nutritional reserves in orchid seeds, fungal symbionts are essential to provide substrates and enable germination. Analyzing germination stages by structural perspectives can also answer important questions regarding the fungi interaction with the seeds. Different imaging techniques can be applied to unveil fungi endophytes in plant tissues, as are proposed in this article. Freehand and thin sections of plant organs can be stained and then observed using light microscopy. A fluorochrome conjugated to wheat germ agglutinin can be applied to the fungi and co-incubated with Calcofluor White to highlight plant cell walls in confocal microscopy. In addition, the methodologies of scanning and transmission electron microscopy are detailed for mycoheterotrophic orchids, and the possibilities of applying such protocols in related plants is explored. Symbiotic germination of orchid seeds (i.e., in the presence of mycorrhizal fungi) is described in the protocol in detail, along with possibilities of preparing the structures obtained from different stages of germination for analyses with light, confocal, and electron microscopy.

This protocol aims to provide detailed procedures for collecting, fixing, and maintaining mycoheterotrophic plant samples, applying different microscopy techniques such as scanning and transmission electron microscopy, light, confocal, and fluorescence microscopy to study fungal colonization in plants tissues and seeds germinated with mycorrhizal fungi.

Structural botany is an indispensable perspective to fully understand the ecology, physiology, development, and evolution of plants. When researching ...

Optimized Thin Endometrium Mouse Model with Low Mortality and Minimal Adhesions

Establishing a Mouse Model of Thin Endometrium

Thin endometrium (TE) has been widely recognized as a critical cause of infertility. However, the pathogenesis of TE remains unclear, and satisfactory treatment options are still urgently needed. Several animal models of TE have been developed, but the mouse model involving abdominal surgery and injection of 95% ethanol presents a formidable challenge due to the high mortality rate and risk of intrauterine adhesions if not performed correctly. Here, we describe a detailed protocol that produces reliable and reproducible TE with a very low mortality rate and minimal intrauterine adhesions by injecting 95% ethanol into the mouse uterus with varying infusion times. The results showed that all of the mice successfully developed TE with infusion times ranging from 1-3 min, characterized by a typical reduction in endometrial thickness and the number of glands, as well as excessive endometrial fibrosis. These findings suggest that this mouse model is suitable for studying thin endometrium and can serve as a platform for developing future TE treatments.

This article describes a detailed protocol for producing a reliable and reproducible thin endometrium with a very low mortality rate and minimal intrauterine adhesions by injecting 95% ethanol into the mouse uterus within 1-3 min.

Thin endometrium (TE) has been widely recognized as a critical cause of infertility. However, the pathogenesis of TE remains unclear, and satisfactory ...

Formalin-Fixed Paraffin Embedded (FFPE) Tissue Preservation: A Method for Studying Tissue Morphology

Source: Nagaraj A.S., et al., <a href="https://www-jove-com.bdigitaluss.remotexs.co/v/58569/establishment-analysis-tumor-slice-explants-as-prerequisite-for">Establishment and Analysis of Tumor Slice Explants As a Prerequisite for Diagnostic Testing.</a> J. Vis. Exp. (2018)
The following video describes how to preserve morphology of a tissue with formalin fixation and paraffin embedding.