This work was supported by the National Institutes of Health (R01DE020843 to Y.M.), the International FOP Association (Y.M.), and a grant-in-aid from the National Natural Science Foundation of China (31500788 to J.Y.).

All mouse experiments were carried out in accordance with University of Michigan guidelines covering the humane care and use of animals in research. All animal procedures used in this study were approved by the Institutional Animal Care and Use Committee (IACUC) at the University of Michigan (Protocol #PRO00007715).
1. Tissue Preparation
<ol>
	<li>Preparation of embryonic tissues 
	<ol>
		<li>Prepare one 10 cm dish and several 3.5 cm dishes containing phosphate buffered saline (PBS), and one 12-well culture plate containing 2 mL 4% paraformaldehyde (PFA) in PBS in each well for each pregnant mouse. Place all Petri dishes and the plate on ice. 
		NOTE: Handle 4% PFA in a fume hood.</li>
		<li>Dissect embryos from pregnant mice in ice-cold PBS with forceps and scissors as previously described14.
		<ol>
			<li>Briefly, euthanize a pregnant mouse with CO2, grab the skin below the center of the belly with forceps and cut through the skin only, then gently pull at the skin to separate it from the underlying the abdominal muscle wall.</li>
			<li>Next, cut into the abdominal cavity following the same line of the skin incision. Remove the uterus containing a string of embryos and remove the embryos by gently cutting away the uterine wall. The extraembryonic tissues such as the yolk sac and amnion will be removed.</li>
			<li>Cut and isolate head from each embryo.</li>
		</ol>
		</li>
		<li>Transfer each head into each well of a 12-well plate containing 4% PFA with a plastic transfer pipette or forceps. Fix samples in 4% PFA at 4 &#176;C for 4 h. Rinse samples in PBS at 4 &#176;C with gentle shaking for 12 h. 
		NOTE: For embryos younger than embryonic day 16.5 (E16.5), fix embryo heads with 4% PFA directly after isolation. For embryos at E16.5 or later, remove to discard skin and adipose tissue from the heads and rinse several times in ice cold PBS before fixation.</li>
		<li>Cryoprotect heads.
		<ol>
			<li>Transfer each head into a new 12-well plate containing 2 mL of 30% sucrose in PBS using a plastic transfer pipette or forceps. Agitate gently at 4 &#176;C until the head sinks to the bottom of the dish.</li>
		</ol>
		</li>
		<li>Embed heads.
		<ol>
			<li>Transfer the cryoprotected head into a mold containing Optimal Cutting Temperature (OCT) compound. Equilibrate samples in OCT for several minutes. Adjust the location and direction of samples with forceps.</li>
			<li>Place the mold on dry ice to freeze. Store resulting cryomolds in a plastic bag at -80 &#176;C until ready for cryosectioning. 
			NOTE: The trimmed side of the samples must face the bottom of the embedding mold.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Preparation of postnatal undecalcified hard tissues
	<ol>
		<li>Euthanize at 3 week or 3 month old mouse with CO2. Remove the skin and adipose tissues. Cut and isolate the head or long bones from the mouse.</li>
		<li>Fix and cryoprotect the head or long bone of mice as described in steps 1.1.3&#8211;1.1.4.</li>
		<li>Embed in 8% gelatin in a similar manner as step 1.1.5. Keep the cryomolds in a plastic bag at -80 &#176;C until cryosectioning. 
		NOTE: Decalcification is not necessary here. To prepare 8% gelatin, mix 8 g of gelatin with 100 mL of PBS and boil using a microwave. Be aware that the mixture boils over easily.</li>
	</ol>
	</li>
</ol>
2. Cryosectioning
<ol>
	<li>Set cryostat temperature to -18 &#176;C for soft tissues embedded in OCT or -25 &#176;C and lower for undecalcified hard tissues embedded in gelatin. Keep samples in the cryostat chamber for about 30 min to equilibrate to the cryostat temperature.</li>
	<li>Expel the block from the cryomold. Freeze the block onto the specimen chuck (tissue holder) via mounting with an OCT drop. Keep the trimmed side of the sample furthest from the chuck (facing the operator).</li>
	<li>Load the block-mounted chuck onto the cryostat object holder. Adjust the blade holder to make the angle of the blade 3&#176;&#8211;5&#176; relative to the sample.</li>
	<li>Collect 10 &#181;m sections onto coated microscope slides. Dry sections completely at RT, then store them at -80 &#176;C.</li>
</ol>
3. Histological Staining and Microscopic Imaging
<ol>
	<li>Immunofluorescence staining
	<ol>
		<li>Take out slides from -80 &#176;C. Keep slides at RT for 1 h to airdry sections. Rinse slides in 0.1% PBST (0.1% Polyethylene glycol tert-octylphenyl ether in PBS; see Table of Materials) three times for 5 min each to wash out OCT and permeabilize sections.</li>
		<li>Optionally, perform antigen retrieval (optional).
		<ol>
			<li>Preheat citrate buffer (10 mM sodium citrate pH 6) in the staining dish with steamer or water bath to 95&#8211;100 &#176;C. Immerse slides in the citrate buffer, incubate for 10 min.</li>
			<li>Take the staining dish out from steamer or water bath to RT. Cool the slides at RT for 20 min or longer15. 
			NOTE: As alternatives, use Tris-EDTA buffer (10 mM Tris base, 1mM EDTA, 0.05% Tween 20, pH 9.0) or EDTA buffer (1 mM EDTA, 0.05% Tween 20, pH 8.0) for heat-induced antigen retrieval. Use a pressure cooker, microwave, or water bath for heat-induced antigen retrieval, in addition to the hot steamer. An enzyme-induced antigen retrieval using trypsin or pepsin is another alternative. Optimize the concentration and treatment time of enzymatic retrieval to avoid damaging sections. Optimize the antigen retrieval method for each antibody/antigen combination.</li>
		</ol>
		</li>
		<li>Incubate each slide with 200 &#956;L of blocking solution (5% donkey serum diluted in 0.1% PBST) at RT for 30 min, then remove the blocking solution without rinse.</li>
		<li>Incubate each slide with 100 &#956;L of primary antibody or antibodies diluted in blocking solution for 1 h at RT or O/N at 4 &#176;C. Rinse slides with PBS three times for 10 min each at RT.</li>
		<li>Incubate each slide with 100 &#956;L of secondary antibody diluted in blocking solution for 1 h at RT. Rinse slides in PBS three times for 10 min each at RT. Protect slides from light.</li>
		<li>Mount slides.
		<ol>
			<li>Add two drops of anti-fade medium with DAPI (4&#39;, 6-diamidino-2-phenylindole) on the slide. Then cover with a coverslip.</li>
			<li>Store at 4 &#176;C in dark until ready to image. 
			NOTE: As an alternative, label the nuclei with DAPI or Hoechst 33324 dye diluted 1:2,000 in PBS at RT first, then mount with glycerol.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Terminal deoxynucleotidyl transferase dUTP nick end labeling (TUNEL) staining. 
	NOTE: Double-stranded DNA with 3&#8217;-hydroxyl termini (3&#39;OH DNA termini) will form during apoptosis in the cell. Here, we provide a protocol that label the free 3&#39;OH DNA termini in situ via labeling DNA fragments with the digoxigenin-nucleotide utilizing terminal deoxynucleotidyl transferase (TdT) by specific staining using a commercial kit (see Table of Materials).
	<ol>
		<li>Optionally, stain sections with primary and Alexa Fluor-488 labeled secondary antibodies prior to the TUNEL staining. Rinse the slides in PBS three times for 10 min each. 
		NOTE: This step is optional for a double staining of a protein and TUNEL in the same slide.</li>
		<li>Incubate each slide with 100 &#956;L Proteinase K (10 &#181;g/mL in 10 mM Tris pH 7.5 and 5 mM EDTA) for 5 min at RT. Rinse slides with PBS three times for 10 min each at RT. 
		NOTE: Adjust the incubation time and temperature of Proteinase K for each tissue type. For 10 &#181;m sections of embryo heads fixed in 4% PFA, incubate for 5 min at RT. In addition to the method using Proteinase K, use alternative treatments as needed, including (1) freshly prepared 0.1% Polyethylene glycol tert-octylphenyl ether, 0.1% sodium citrate, 10 min at 37 &#176;C; (2) 0.25%&#8211;0.5% Pepsin in HCl (pH 2) or 0.25% trypsin, 10 min at 37 &#176;C; and (3) microwave irradiation with 0.1 M citrate buffer (pH 6).</li>
		<li>Apply 200 &#956;L of blocking solution (5% donkey serum diluted in 0.1% PBST) to each slide, incubate at RT for 30 min, tap off the blocking solution without rinse.</li>
		<li>Apply 50 &#181;L of the equilibration buffer supplied by the kit to each slide at RT for at least 10 s. Tap off the buffer without rinse.</li>
		<li>Prepare reaction mixture (working strength TdT enzyme) by mixing TdT Enzyme with the reaction buffer supplied by the kit at the ratio of 3:7. Apply 50 &#181;L of the reaction mixture to each slide, and incubate at 37 &#176;C for 1 h. Tap off the buffer without rinsing.</li>
		<li>Apply 200 &#181;L of the stop buffer (1:30 diluted in ddH2O) supplied by the kit to each slide, then incubate at RT for 10 min. Rinse slides with PBS three times for 10 min each.</li>
		<li>Label with Rhodamine antibody.
		<ol>
			<li>Apply 50 &#181;L of pre-warmed (RT) anti-digoxigenin conjugate (rhodamine) (1:1 diluted in blocking solution) to each slide. Incubate at RT for 30 min in dark.</li>
			<li>Rinse slides with PBS three times for 10 min each. Mount slides as step 3.1.6.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
4. Imaging Acquisition
<ol>
	<li>Use positive controls (tissues positive for the target antigen) to check the signal labeling and negative controls (omit the primary antibody, isotype control, or tissues negative for the target antigen) to evaluate the background of images under the fluorescent microscope.</li>
	<li>Set the equipment and camera conditions (exposures and other general settings) for imaging based on the signal intensity of negative and positive controls. 
	NOTE: These conditions vary by (1) cameras and microscopes used for imaging, (2) antibodies, and (3) tissues for each experiment. Common conditions used for craniofacial tissues are ISO 200 with an exposure time ranging from 1/100 s to 1 s depend on the quality and specificity of antibodies. Appropriate magnifications vary depending on the size of the samples and purpose of experiments.</li>
	<li>Acquire images with conventional epifluorescence microscope or confocal microscope. Acquire images (including those of corresponding controls) in the same conditions for each color channel. Save images with the same format (tiff is best to preserve information).</li>
</ol>
5. Fluorescence Quantification
NOTE: Statistically comparing the staining between different groups will be more informative in many cases. With the immunofluorescence images, quantify the relative level of the protein by measuring signal density, counting positive cells, or calculating positive areas. For statistical analysis, the minimum number of biologically independent samples is 3. A typical method is to generate at least three sections from each sample and take images for at least three representative areas in each section.
<ol>
	<li>Quantification of fluorescence intensity using ImageJ
	<ol>
		<li>Open the software, and use Analyze &#62; Set Measurements to check that only Area and Integrated Density are selected. Use File &#62; Open to open images to be analyzed.</li>
		<li>Use Toolbar to select either the square or circle icon on the far left. Select the area to be analyzed on the image using the selection tool. Use Analyze &#62; Measure to get the readout of the selected area and integrated density in the Results window. Select a region next to a positive cell that has no fluorescence to read out the background.</li>
		<li>Repeat step 5.1.2 to analyze other images. Adjusted the area to be analyzed to match with that of the first image.</li>
		<li>Copy all the data in the Results window and paste into a spreadsheet when finished analyzing.</li>
		<li>Calculate the corrected fluorescence intensity (CTCF) as Integrated Density &#8212; (Area of selected cell x Mean fluorescence of background readings). Compare the difference of the corrected total cell fluorescence between samples and make a graph.</li>
	</ol>
	</li>
	<li>Quantification of the positive cell number of fluorescent images using ImageJ
	<ol>
		<li>Manual cell counting.
		<ol>
			<li>Use ImageJ &#62; Plugins &#62; Analysis to install the Cell Counter plugin.</li>
			<li>Use File &#62; Open to open images to be analyzed. Use Plugins &#62; Analysis &#62; Cell counter to open the Counter window and the Results window. 
			NOTE: Cell counter does not work on stacks. For counting stacks, plugin Plot Z Axis Profile, then use Image &#62; Stacks &#62; Plot Z Axis Profile to monitor the intensity of a moving ROI using a particle tracking tool. This tool can be either manual or automatic.</li>
			<li>Clicking one of buttons at the bottom of the Counter window to initiate counting. Click directly on a cell/object to be count until finishing.</li>
			<li>Click the Results button in the Count window. The total number of cells counted will be shown in the Results window. Save the result log as spreadsheet and analyze.</li>
		</ol>
		</li>
		<li>Automated cell counting.
		<ol>
			<li>Use File &#62; Open to open images to be analyzed. Convert the RGB image into a grey scale image before proceeding.</li>
			<li>Use Image &#62; Adjust &#62; Threshold to select all the areas that need to be counted.</li>
			<li>Use Analyze &#62; Analyze Particles to get the number of cells/particles. Set a range of the valid particle size (e.g., 100-Infinity) instead of the default of 0-Infinity to count cells/particles within a specific range. Save the result log as spreadsheet and analyze. 
			NOTE: To get other information from the image, besides area, go to Analyze &#62; Set Measurements and select the box next to the information needed.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Here we provide a detailed protocol for preparation of mouse head and undecalcified bone tissues, and cryosectioning for immunostaining of cell proliferation, cell death, and BMP signaling markers. We also detail the strategy for obtaining quantitative data from immunofluorescent images. Those methods can also be applicable to other tissues with appropriate modifications.
Conditions for tissue preparation vary by the size and type of tissues. The fixation and cryoprotection time usually need several hours to overnight. After fixation, the tissue can also be embedded in paraffin and sectioned with a microtome16. Though both paraffin and OCT work well for immunostaining, there are some differences between them. Paraffin blocks can be kept for multiple years at RT, while OCT blocks are for 1 year at -80 &#176;C. Paraffin preserves tissue morphology, while ice crystal formed during OCT embedding may negatively affect tissue structures. Paraffin sometimes masks epitopes of antigens, while OCT preserves enzyme activities and antigen epitopes. Therefore, there is no need of antigen retrieval for most of the antibodies if fixed in 4% PFA for only 4 h or less and embedded in OCT. It is, however, still possible to get better results by antigen retrieval, if positive controls did not show good staining results in cryosections.
Both Hoechst dye and DAPI can be used for nuclear counter-staining. They have similarities, as both (1) are UV-excited, minor groove-binding chemicals to emit signals proportional to total DNA content, and (2) are subjected to photo-bleaching after a long exposure. However, Hoechst dyes are typically used for staining DNA content in live cells due to their high permeability. DAPI is typically used for staining DNA in fixed cells due to its low membrane permeability. In addition, DAPI generates a stronger and more stable signal than Hoechst.
Proper controls are essential for IF. The specificity of every new antibody should be confirmed by Western blot analysis, if applicable. The optimal working concentration of a particular primary antibody should be determined through the use of serial dilutions. A positive control (tissue or cell which is proved to express the protein/antigen) should be included to check the IF process and specificity of antibody. A negative control should also be included, e.g., absence of the primary antibody, or the substitution of normal IgG from the same species for the primary antibody, or tissues negative for the target antigen. When taking pictures, the sample without secondary antibodies (background control) should be examined independently with each channel to set the limits of signal gain and offset to be adapted for the final imaging. For detection of multiple labels, background control and single-labeled controls need to be prepared to avoid spectral overlap artifacts. All channels that will be used to obtain an image of a multiple-label sample must be subjected to independent background correction, because the level of autofluorescence in each channel varies substantially.
We also provide the protocol for preparation and cryosectioning of undecalcified hard tissues embedded in gelatin. For the OCT embedded decalcified hard tissues, most of the hard tissue sections will be detached from slide glasses during immunostaining procedures, because of their low adhesion character on the slide. The adhesive tape designed to facilitate cryosectioning helps to generate good quality sections. But, those sections are easily damaged when the tape is peeling off. For gelatin embedded tissues, there is no need for a tape-transfer system to generate good quality sections. As an embedding medium, gelatin can infiltrate the sample well, although it has a lower viscosity compared with OCT. Gelatin has been used in other histological applications, such as brain tissue17,18 and ultrathin sections of cells for immunocytochemistry19. Here, gelatin was used to embed undecalcified bone, which generates blocks easier to cryosection than OCT. There are several small tips to get good sections of gelatin embedded undecalcified hard tissues. The critical step is to embed with gelatin instead of OCT. To get better penetration, keep samples in 30% sucrose one more day after samples sink to the bottom. It is equally important to set the temperature lower than usual at about -25 &#176;C. An ultra-sharp blade is not necessary. Although a lower cryo temperature (-25 &#176;C) makes some improvements for cryosectioning of OCT embedded undecalcified hard tissues, it is still difficult to get a good integrity of tissue structures. As shown in Figure 2, Figure 3, and Figure 4, good quality sections applicable for immunostaining were obtained from gelatin embedded hard tissues (e.g., trabecular bones, cortical bones, skull bones, nasal tissues, and incisor). Those results proved that gelatin embedding significantly improves the specimen integrity of hard tissue sections, but also enhances adhesion of the sections to slide glasses. In addition, gelatin preserves antigen functions, and exhibits compatibility with fluorescent signals and immunostaining. However, this technique only works well for up to 3 month-old samples. Potential improvements of this method are (1) to dissect the samples further to separate the target tissue from other parts to make the structure of the tissue simple (in the case of teeth, the mandible or maxilla should be dissected and fixed instead of the whole head) and (2) to use 10% EDTA to decalcify tissues for only 2&#8211;3 days before cryoprotection. This short time of decalcification will not compromise the immunostaining results. Another concern is that, as a non-aqueous embedding media, gelatin cannot be easily removed from slides, which may lead to higher background depending on staining methods (e.g., H&#38;E staining).
Immunostaining results are not easy to quantify, so they are usually used semi-quantitatively. Difficulties and limitations of the quantification of immunostaining of craniofacial tissues include but are not limited to the following: (1) it is difficult to define the area to be counted due to the complexity of the structure of craniofacial tissues; (2) it is difficult to define the labeled area or labeled cells due to the non-linear nature of immunostaining; (3) there is limited information for the dynamic range of the signal; (4) it is hard to compare the intensity of signals between images or groups due to the fading of fluorescent signal during image acquisition; and (5) the signal background may change significantly among antibodies, slides, and samples. To increase the reliability of the quantification results, the experiment should be carefully and strictly performed. All samples should be processed under the same conditions. During immunostaining, various controls are required to evaluate signal background and define the positive area or cells for the signal. Take convincing and representative images to clearly show the area to be counted and labeled cells with good contrast. In addition, during image acquisition, the camera setting and equipment setting must be kept consistent.
Taken together, we present a simple standard protocol for immunofluorescence on mouse craniofacial tissues, especially for undecalcified hard tissues. Immunostaining analysis of craniofacial tissues will not only help to understand the mechanism of morphogenesis during development, but also illustrate the changes during pathogenesis. In addition, immunostaining can also be used to study the expression pattern of other signaling pathway ligands, receptors, or other phenotypic markers, besides cell proliferation, cell death, and the BMP signaling pathway. However, the critical steps of an immunostaining experiment must be modified appropriately for each antigen/antibody or tissue to get specific staining and minimized non-specific background signals.

The face is a key part of human identity, and is composed of several different types of tissues, such as epithelium, muscle, bone, cartilage, tooth. Those tissues are derived from all three germ layers: ectoderm, endoderm, and mesoderm1,2. For proper patterning and development of craniofacial tissues, cell proliferation, death and differentiation need to be highly coordinated and regulated by specific signaling pathways, such as Wnt, Fgf, Hh and Bmp pathways3,4,5. Defects in proliferation, survival or differentiation of cells will lead to craniofacial malformations, which are among the most frequently occurring congenital birth defects. Transgenic mice are useful tools to study mechanisms of craniofacial morphogenesis and pathogenesis1,2,3,4,5. Understanding the changes in craniofacial structures during development and pathogenesis will help to clarify key developmental principles as well as the mechanisms of craniofacial malformations1,2,3,4,5.
The staining of whole mount or sectioned tissues with specific antibodies is an invaluable technique for determining spatial distribution of proteins of interest 6. Formally, tissue immunostaining can rely either upon immunohistochemistry (IHC) or immunofluorescence (IF). Compared with the opaque reaction product generated with a chromogenic substrate such as 3,3&#8217;-Diaminobenzidine (DAB) by IHC, IF involves the use of fluorescent conjugates visible by fluorescence microscopy. Therefore, IF may clearly differentiate positive cells from background noise, and allows images to be quantitatively analyzed and enhanced in a straightforward fashion by software such as ImageJ and Adobe Photoshop7,8. The whole mount staining approach works on small blocks of tissue (less than 5 mm thick), which can provide three-dimensional information about the location of proteins/antigens without the need for reconstruction from sections9,10. However, compared with tissue sections, whole mount immunostaining is time consuming and requires large volumes of antibody solutions. Not all antibodies are compatible with the basic whole mount approach. In addition, the incomplete penetration of antibodies will result in uneven staining or false negative staining. Here we will focus on the immunofluorescence detection of proteins/antigens on sectioned tissues. For hard tissues (eg, head, tooth, long bone), calcium deposition during development/pathogenesis makes the sample difficult to section and easily rinsed off during immunostaining treatment11,12. Most of the currently available protocols decalcify hard tissues before embedding to make sectioning easier, which is time consuming and can destroy morphology and antigens of samples if handled improperly11,12. To overcome the issues, we optimized an approach for cryosectioning of hard tissues without decalcification, leading to improved visualization of their morphology and distribution of signaling proteins.
The protocol described here is being used to determine morphometric and histological changes in the craniofacial tissues of BMP transgenic mice. Specifically, the protocol includes (1) harvesting and dissecting head tissues, (2) section and immunostaining of experimental markers (Ki67, pSmad1/5/9) along with TUNEL staining, (3) imaging the sections using fluorescence microscope, and finally (4) analyzing and quantifying the results. The protocol to prepare and cryosection hard tissues without decalcification is also described13. Those methods are optimized for craniofacial tissues. They are also applicable to other tissues from various ages of samples with appropriate modifications.

<table><tbody><tr><td>Adhesive tape</td><td>Leica</td><td>#39475214</td><td></td></tr><tr><td>Alexa fluor 488-goat anti-Rabbit secondary antibody</td><td>Invitrogen</td><td>A-11034</td><td></td></tr><tr><td>Antifade Mountant with DAPI</td><td>Invitrogen</td><td>P36931</td><td></td></tr><tr><td>Bovine serum albumin</td><td>Sigma</td><td>A2153</td><td></td></tr><tr><td>Coverslips</td><td>Fisher Brand</td><td>12-545-E</td><td></td></tr><tr><td>Cryostat</td><td>Leica</td><td>CM1850</td><td></td></tr><tr><td>EDTA</td><td>Sigma</td><td>E6758</td><td></td></tr><tr><td>Fluorescence microscope</td><td>Olympus</td><td>BX51</td><td></td></tr><tr><td>Gelatin</td><td>Sigma</td><td>G1890</td><td></td></tr><tr><td>In Situ&amp;nbsp;Cell Death Detection Kit</td><td>Millipore</td><td>S7165</td><td></td></tr><tr><td>Microscope slides</td><td>Fisher Brand</td><td>12-550-15</td><td></td></tr><tr><td>OCT Compound</td><td>Fisher Healthcare</td><td>23-730-571</td><td></td></tr><tr><td>Paraformaldehyde (PFA)</td><td>Sigma</td><td>P6148&amp;nbsp;</td><td></td></tr><tr><td>Phosphate buffered saline (PBS)</td><td>Sigma</td><td>P4417</td><td></td></tr><tr><td>Polyethylene glycol&amp;nbsp;tert-octylphenyl ether</td><td>Sigma</td><td>T9284</td><td>Triton X-100</td></tr><tr><td>Proteinase K</td><td>Invitrogen</td><td>AM2542</td><td></td></tr><tr><td>Rabbit anti-Ki67 antibody</td><td>Cell Signaling Technology</td><td>9129</td><td>Lot#:3; RRID:AB_2687446</td></tr><tr><td>Rabbit anti-pSmad1/5/9 antibody</td><td>Cell Signaling Technology</td><td>13820</td><td>Lot#:3; RRID:AB_2493181</td></tr><tr><td>Sodium citrate</td><td>Sigma</td><td>1613859</td><td></td></tr><tr><td>Sucrose</td><td>Sigma</td><td>S9378</td><td></td></tr><tr><td>Tris</td><td>Sigma</td><td>10708976001</td><td></td></tr></tbody></table>

tissue preparation and immunostaining of mouse craniofacial tissues and undecalcified bone

Tissue immunostaining provides highly specific and reliable detection of proteins of interest within a given tissue. Here we describe a complete and simple protocol to detect protein expression during craniofacial morphogenesis/pathogenesis using mouse craniofacial tissues as examples. The protocol consists of preparation and cryosectioning of tissues, indirect immunofluorescence, image acquisition, and quantification. In addition, a method for preparation and cryosectioning of undecalcified hard tissues for immunostaining is described, using craniofacial tissues and long bones as examples. Those methods are key to determine the protein expression and morphological/anatomical changes in various tissues during craniofacial morphogenesis/pathogenesis. They are also applicable to other tissues with appropriate modifications. Knowledge of the histology and high quality of sections are critical to draw scientific conclusions from experimental outcomes. Potential limitations of this methodology include but are not limited to specificity of antibodies and difficulties of quantification, which are also discussed here.

Embryonic craniofacial tissue sections 
Following the above steps, heads were dissected from control (P0-Cre) or mutant (constitutively activated Bmpr1a in neural crest cells, P0-Cre; caBmpr1a) embryos at embryonic day (E) 16.5 or 18.5. After fixing in 4% PFA for 4 h, samples were embedded in OCT and cryosectioned coronally. Resulted sections were immunostained with antibodies against pSmad1/5/9 (downstream BMP signaling factors) or Ki67 (a cell proliferation marker) without antigen retrieval according to the protocol. As shown, pSmad1/5/9 (Figure 1A) and Ki67 (Figure 1C) were positive in the frontal bones of control embryos. In mutant embryos, the levels of pSmad1/5/9 was increased (Figure 1B), while those of Ki67 was decreased (Figure 1D) in the frontal bones. Cell death in those samples were also checked according to the protocol. As shown, more apoptotic cells were observed in the frontal bones of mutant embryos than those of control embryos (Figure 1E,F).
Undecalcified craniofacial tissues or long bone sections 
Following the above steps for undecalcified hard tissues, heads from 3 week old mice (P0-Cre; mTmG (membrane-tomato and membrane GFP)) were fixed with 4% PFA and embedded in 8% gelatin. Coronal cryosections were washed with PBST and mounted with anti-fade medium with DAPI. Figure 2A,B demonstrate that gelatin does not interfere with fluorescent signals from sectioned tissues.
Heads and femora from 3 week-old or 3 month-old mice were employed to check whether gelatin embedded undecalcified tissues are good for IF. The whole heads and femora were processed and sectioned according to the protocol. The resulted sections were used for SOX9 immunostaining (Figure 3) or OSX and E11/Podoplanin double immunostaining (Figure 4). As shown, good quality sections were obtained from most of the 3 week hard tissues, including the trabecular and the cortical compartments of the femur (Figure 3A,B, Figure 4A&#8211;D), the frontal bones (Figure 4E,F), the incisor (Figure 3E,F, Figure 4I,J), nasal tissues (Figure 3C,D), and the skull including the nasal-premaxilla suture and surrounding bones (Figure 4G,H) of the head. While, with 3-month-old samples, good quality sections were only obtained in some of the hard tissues, including the trabecular compartments of the femur (Figure 3G,H, Figure 4K,L), nasal tissues (Figure 3I,J), and the skull including the nasal-premaxilla suture and surrounding bones (Figure 4M,N) of the head. As shown in Figure 3, SOX9 positive cells were detected specifically in the chondrocytes of the growth plate (Figure 3B) and the joint (Figure 3H) from the femur, and the nasal septum (Figure 3D,J). In the 3 week-old incisor, SOX9 was detected in the mesenchymal cells (Figure 3F). OSX and E11 double staining results showed that OSX was detected in osteoblasts, while E11 was detected in osteocytes of bones from the femur and the head (Figure 4B,D,H,L,N). In the 3 week incisor, OSX was positive in odontoblasts, while E11 was positive in follicle mesenchymal cells (Figure 4J). Those results indicate that undecalcified hard tissues embedded with gelatin well-preserve antigen functions.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/59113/59113fig1.jpg" /> 
Figure 1: Examples of IF results of pSmad1/5/9, Ki67 or TUNEL in control embryos and mutant embryos with enhanced BMP activity. Constitutively activated Bmpr1a (caBmpr1a) mice were crossed with P0-Cre mice to increase BMP signaling activity in neural crest cells (NCCs). Heads of control (P0-Cre; caBmpr1a+/+) and mutant (P0-Cre; caBmpr1afx/+) embryos were dissected at E16.5 or E18.5, fixed with 4% PFA for 4h, cryoprotected with 30% sucrose for 1 day, embedded in OCT, and cryosectioned at -18 &#176;C. Sections of the frontal bone (similar level with the eye) were used for immunodetection against pSmad1/5/9, Ki67, or TUNEL staining. (A, B) pSmad1/5/9 (green) staining patterns in the frontal bones of control (A) or mutant (B) embryos at E16.5. (C, D) Ki67 (green) staining patterns in the frontal bones of control (C) or mutant (D) embryos at E18.5. (E, F) TUNEL (red) staining patterns in the frontal bones of control (E) or mutant (F) embryos at E18.5. Nuclei were stained with DAPI (blue). FB = frontal bone, B = brain. Scale bars = 100 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59113/59113fig1large.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/59113/59113fig2.jpg" /> 
Figure 2: Examples of mTmG reporter signal results of undecalcified tissues in the head. Heads from 3 week old P0-Cre mice with membrane-tomato and membrane GFP (mTmG) reporter were dissected, fixed with 4% PFA for 4h, cryoprotected with 30% sucrose for 2 days, embedded in 8% gelatin, and cryosectioned at -25 &#176;C. Head sections clearly show GFP (green, Cre recombination positive) and Tomato (red, Cre recombination negative) signal in the nasal bone and nasal tissues (A, B). Nuclei were stained with DAPI (blue). NB = nasal bone, N = nasal tissues, NS = nasal septum. Scale bars = 250 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59113/59113fig2large.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/59113/59113fig3.jpg" /> 
Figure 3: Examples of SOX9 immunostaining results of undecalcified tissues in the head and the femora. Heads and femora were dissected from 3 week or 3 month old mice, fixed with 4% PFA for 4h, cryoprotected with 30% sucrose for 2 days, embedded in 8% gelatin, and cryosectioned at -25 &#176;C. Slides were used for immunodetection against SOX9 (red). Nuclei were stained with DAPI (blue) (B, D, F, H, J). Adjacent sections of those tissues were used for Hematoxylin &#38; Eosin (H&#38;E) staining (A, C, E, G, I). Arrow heads in A and B indicate growth plate and in G and H, articular cartilage. Arrows in A and G indicate trabecular bones and in C, D, I, and J, nasal septum. DM = dental mesenchyme, DE = dental epithelium, FM = follicle mesenchyme. Scale bars = 50 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59113/59113fig3large.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/59113/59113fig4.jpg" /> 
Figure 4: Examples of OSX and E11 double immunostaining results of undecalcified tissues in the head and the femora. Heads and femora were dissected from 3 week-old or 3 month-old mice, fixed with 4% PFA for 4h, cryoprotected with 30% sucrose for 2 days, embedded in 8% gelatin, and cryosectioned at -25 &#176;C. Sections were used for double immunostaining with antibodies against OSX (Red) and E11/Podoplanin (Green). Nuclei were stained with DAPI (blue) (B, D, F, H, J, L, N). Adjacent sections of those tissues were used for H&#38;E staining (A, C, E, G, I, K, M). Arrows in A, B, K, and L indicate trabecular compartments of the femur; C and D, cortical compartments of the femur; and in E and F, the frontal bones. Arrowheads in A and B indicate growth plate. BM = bone marrow, N = nasal tissues, DM = dental mesenchyme, DE = dental epithelium, FM = follicle mesenchyme, NPS = nasal premaxilla suture. The frontal bones (E, F) and the nasal-premaxilla suture and surrounding bones (G, H, M, N) are also shown. Scale bars = 50 &#956;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/59113/59113fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Tissue Preparation and Immunostaining of Mouse Craniofacial Tissues and Undecalcified Bone at JoVE.com

Tissue Preparation and Immunostaining of Mouse Craniofacial Tissues and Undecalcified Bone

Here, we present a detailed protocol to detect and quantify protein levels during craniofacial morphogenesis/pathogenesis by immunostaining using mouse craniofacial tissues as examples. In addition, we describe a method for preparation and cryosectioning of undecalcified hard tissues from young mice for immunostaining.

Tissue immunostaining provides highly specific and reliable detection of proteins of interest within a given tissue. Here we describe a complete and ...

tissue-preparation-immunostaining-mouse-craniofacial-tissues

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

12.4K Views.  Wuhan University. Hier presenteren we ons eenvoudige protocol voor immunofluorescentiedetectie op eiwitten op gedesnede materialen. Deze methode vereist geen antigeen ophalen. We zullen muizenkoppen gebruiken, inclusief onverkalkte harde weefsels.Het belangrijkste voordeel van deze techniek is het overslaan van antigeen ophalen en ontkalking van harde weefsels. Die leiden tot een goede kwaliteit van immunostaining resultaten. Deze methode bespaart ook zowel tijd als arbeid.Deze methode is ook van...

Video: Tissue Preparation and Immunostaining of Mouse Craniofacial Tissues and Undecalcified Bone

Tissue Preparation and Immunostaining of Mouse Sensory Nerve Fibers Innervating Skin and Limb Bones

Detection and primary processing of physical, chemical and thermal sensory stimuli by peripheral sensory nerve fibers is key to sensory perception in animals and humans. These peripheral sensory nerve fibers express a plethora of receptors and ion channel proteins which detect and initiate specific sensory stimuli. Methods are available to characterize the electrical properties of peripheral sensory nerve fibers innervating the skin, which can also be utilized to identify the functional expression of specific ion channel proteins in these fibers. However, similar electrophysiological methods are not available (and are also difficult to develop) for the detection of the functional expression of receptors and ion channel proteins in peripheral sensory nerve fibers innervating other visceral organs, including the most challenging tissues such as bone. Moreover, such electrophysiological methods cannot be utilized to determine the expression of non-excitable proteins in peripheral sensory nerve fibers. Therefore, immunostaining of peripheral/visceral tissue samples for sensory nerve fivers provides the best possible way to determine the expression of specific proteins of interest in these nerve fibers. So far, most of the protein expression studies in sensory neurons have utilized immunostaining procedures in sensory ganglia, where the information is limited to the expression of specific proteins in the cell body of specific types or subsets of sensory neurons. Here we report detailed methods/protocols for the preparation of peripheral/visceral tissue samples for immunostaining of peripheral sensory nerve fibers. We specifically detail methods for the preparation of skin or plantar punch biopsy and bone (femur) sections from mice for immunostaining of peripheral sensory nerve fibers. These methods are not only key to the qualitative determination of protein expression in peripheral sensory neurons, but also provide a quantitative assay method for determining changes in protein expression levels in specific types or subsets of sensory fibers, as well as for determining the morphological and/or anatomical changes in the number and density of sensory fibers during various pathological states. Further, these methods are not confined to the staining of only sensory nerve fibers, but can also be used for staining any types of nerve fibers in the skin, bones and other visceral tissue.

Immunocytochemical identification of peripheral sensory nerve fiber subtypes (and detection of protein expression therein) are key to the understanding of molecular mechanisms underlying peripheral sensation. Here we describe methods for preparation of peripheral/visceral tissue samples, such as skin and limb bones, for specific immunostaining of peripheral sensory nerve fibers.

Detection and primary processing of physical, chemical and thermal sensory stimuli by peripheral sensory nerve fibers is key to sensory perception in ...

Neuroscience

Undecalcified Bone Preparation for Histology, Histomorphometry and Fluorochrome Analysis

Undecalcified bone histology demonstrates the micro-architecture of bone. It shows both the mineralised and cellular components of bone, which provides vital information on bone turnover or bone formation and resorption. This has tremendous importance in a variety of clinical and research applications. It yields beautiful images1 and allows for techniques such as fluorochrome assessment and histomorphometry2. Fluorochrome analysis is a technique where fluorescent dyes that bind to calcium are injected at a particular time point, which allows for quantification of the amount of mineralisation at that given time. Histomorphometry is a process of bone quantification at the microscopic level. 
Performing undecalcified bone histology is technically challenging, particularly with large size specimens. It requires variations in technique from those used in standard paraffin embedded histology. This video illustrates the process of producing good quality sections and demonstrates the technical difficulties and methods with which to overcome them. Specimen preparation, fixation and processing are achieved with a manner similar to other soft tissues, however due to the density and lower permeability of bone considerably longer fixation and processing times are required, often taking several weeks. Embedding is achieved using a supporting medium with similar or equal hardness and density to the bone such as methacrylate- based resins, but unlike paraffin infiltration and embedding, this is an irreversible step. Sectioning can be achieved by grinding which produces a thicker section, which is optimal for studies such as fluorochrome analysis. This is best achieved using a diamond blade on a macrotome. Alternatively, thinner sections can be produced for light microscopy and this is achieved using a sledge microtome with a very sharp blade. The sledge microtome provides the additional strength and stability required for large, hard blocks. Resin embedded sections can be stained with a variety of stains, which are demonstrated.

Undecalcified bone histology provides important information for a variety of clinical and research applications. It is technically challenging, particularly with large size specimens. This video illustrates the process of producing good quality sections and demonstrates the technical difficulties and methods with which to overcome them.

Undecalcified bone histology demonstrates the micro-architecture of bone. It shows both the mineralised and cellular components of bone, which provides ...

Biology

Whole Mount Dissection and Immunofluorescence of the Adult Mouse Cochlea

The organ of Corti, housed in the cochlea of the inner ear, contains mechanosensory hair cells and surrounding supporting cells which are organized in a spiral shape and have a tonotopic gradient for sound detection. The mouse cochlea is approximately 6 mm long and often divided into three turns (apex, middle, and base) for analysis. To investigate cell loss, cell division, or mosaic gene expression, the whole mount or surface preparation of the cochlea is useful. This dissection method allows visualization of all cells within the organ of Corti when combined with immunostaining and confocal microscopy to image cells at different planes in the z-axis. Multiple optical cross-sections can also be obtained from these z-stack images. In addition, the whole mount dissection method can be used for scanning electron microscopy, although a different fixation method is needed. Here, we present a method to isolate the organ of Corti as three intact cochlear turns (apex, middle, and base). This method can be used for mice ranging from one week of age through adulthood and differs from the technique used for neonatal samples where calcification of the cochlea is incomplete. A slightly modified version can be used for dissection of the rat cochlea. We also demonstrate a procedure for immunostaining with fluorescently tagged antibodies.

We present a method to isolate the adult organ of Corti as three intact cochlear turns (apex, middle, and base). We also demonstrate a procedure for immunostaining with fluorescently tagged antibodies. Together these techniques allow the study of hair cells, supporting cells, and other cell types found in the cochlea.

The organ of Corti, housed in the cochlea of the inner ear, contains mechanosensory hair cells and surrounding supporting cells which are organized in a ...

Cryosectioning and Immunostaining Mouse Inner Ear Tissue: From Embryonic to Adult Stages

This protocol details general histology methods for preparing inner ear samples from embryonic, neonatal, and adult mice. The purpose of this protocol is to provide a straightforward and standardized method for inner ear tissue processing for researchers, possibly new to the field, interested in working with the mouse cochlea. Included here are protocols for dissection, fixation, embedding, cryosectioning, and immunostaining. Section immunostaining is one of the best methods for examining individual cells within the context of the entire cochlea.
Key steps include dissecting the inner ear away from the skull, fixing the tissue using 4% paraformaldehyde, embedding with Optimal Cutting Temperature compound, cryosectioning, and immunostaining with antibodies targeting specific proteins expressed by the cochlea.
Included in our methods are special considerations made for the cochlea given its 
shape and structure.&#160;Reasonably good focus and dissection skills are needed, as tissue damage can affect the quality of immunostaining and consistency among samples. However, with the procedures we outline, most users can be trained in a few weeks to make beautiful preparations. Overall, this protocol offers a valuable way to promote research to understand auditory development, function, and disease in mouse models.

This protocol, intended for novice users of mouse inner ear tissue, comprehensively details steps for processing mouse inner ears at different developmental stages for section immunostaining.

This protocol details general histology methods for preparing inner ear samples from embryonic, neonatal, and adult mice. The purpose of this protocol is ...

Isolating Central Nervous System Tissues and Associated Meninges for the Downstream Analysis of Immune cells

The central nervous system (CNS) is comprised of the brain and spinal cord and is enveloped by the meninges, membranous layers serving as a barrier between the periphery and the CNS. The CNS is an immunologically specialized site, and in steady state conditions, immune privilege is most evident in the CNS parenchyma. In contrast, the meninges harbor a diverse array of resident cells, including innate and adaptive immune cells. During inflammatory conditions triggered by CNS injury, autoimmunity, infection, or even neurodegeneration, peripherally derived immune cells may enter the parenchyma and take up residence within the meninges. These cells are thought to perform both beneficial and detrimental actions during CNS disease pathogenesis. Despite this knowledge, the meninges are often overlooked when analyzing the CNS compartment, because conventional CNS tissue extraction methods omit the meningeal layers. This protocol presents two distinct methods for the rapid isolation of murine CNS tissues (i.e., brain, spinal cord, and meninges) that are suitable for downstream analysis via single-cell techniques, immunohistochemistry, and in situ hybridization methods. The described methods provide a comprehensive analysis of CNS tissues, ideal for assessing the phenotype, function, and localization of cells occupying the CNS compartment under homeostatic conditions and during disease pathogenesis.

This paper presents two optimized protocols for examining resident and peripherally derived immune cells within the central nervous system, including the brain, spinal cord, and meninges. Each of these protocols helps to ascertain the function and composition of the cells occupying these compartments under steady state and inflammatory conditions.

The central nervous system (CNS) is comprised of the brain and spinal cord and is enveloped by the meninges, membranous layers serving as a barrier ...

Immunology and Infection

An Effective Manual Deboning Method To Prepare Intact Mouse Nasal Tissue With Preserved Anatomical Organization

The mammalian nose is a multi-functional organ with intricate internal structures. The nasal cavity is lined with various epithelia such as olfactory, respiratory, and squamous epithelia which differ markedly in anatomical locations, morphology, and functions. In adult mice, the nose is covered with various skull bones, limiting experimental access to internal structures, especially those in the posterior such as the main olfactory epithelium (MOE). Here we describe an effective method for obtaining almost the entire and intact nasal tissues with preserved anatomical organization. Using surgical tools under a dissecting microscope, we sequentially remove the skull bones surrounding the nasal tissue. This procedure can be performed on both paraformaldehyde-fixed and freshly dissected, skinned mouse heads. The entire deboning procedure takes about 20-30 min, which is significantly shorter than the experimental time required for conventional chemical-based decalcification. In addition, we present an easy method to remove air bubbles trapped between turbinates, which is critical for obtaining intact thin horizontal or coronal or sagittal sections from the nasal tissue preparation. Nasal tissue prepared using our method can be used for whole mount observation of the entire epithelia, as well as morphological, immunocytochemical, RNA in situ hybridization, and physiological studies, especially in studies where region-specific examination and comparison are of interest.

The mammalian nose is a multi-functional organ with intricate internal structures.  The nasal cavity is lined with various epithelia such as olfactory, ...

Cochlear Surface Preparation in the Adult Mouse

Auditory processing in the cochlea depends on the integrity of the mechanosensory hair cells. Over a lifetime, hearing loss can be acquired from numerous etiologies such as exposure to excessive noise, the use of ototoxic medications, bacterial or viral ear infections, head injuries, and the aging process. Loss of sensory hair cells is a common pathological feature of the varieties of acquired hearing loss. Additionally, the inner hair cell synapse can be damaged by mild insults. Therefore, surface preparations of cochlear epithelia, in combination with immunolabeling techniques and confocal imagery, are a very useful tool for the investigation of cochlear pathologies, including losses of ribbon synapses and sensory hair cells, changes in protein levels in hair cells and supporting cells, hair cell regeneration, and determination of report gene expression (i.e., GFP) for verification of successful transduction and identification of transduced cell types. The cochlea, a bony spiral-shaped structure in the inner ear, holds the auditory sensory end organ, the organ of Corti (OC). Sensory hair cells and surrounding supporting cells in the OC are contained in the cochlear duct and rest on the basilar membrane, organized in a tonotopic fashion with high-frequency detection occurring in the base and low-frequency in the apex. With the availability of molecular and genetic information and the ability to manipulate genes by knockout and knock-in techniques, mice have been widely used in biological research, including in hearing science. However, the adult mouse cochlea is miniscule, and the cochlear epithelium is encapsulated in a bony labyrinth, making microdissection difficult. Although dissection techniques have been developed and used in many laboratories, this modified microdissection method using cell and tissue adhesive is easier and more convenient. It can be used in all types of adult mouse cochleae following decalcification.

This article presents a modified cochlear surface preparation method that requires decalcification and use of a cell and tissue adhesive to adhere the pieces of cochlear epithelia to 10 mm round cover slips for immunohistochemistry in adult mouse cochleae.

Auditory processing in the cochlea depends on the integrity of the mechanosensory hair cells. Over a lifetime, hearing loss can be acquired from numerous ...

Combining Multiplex Fluorescence In Situ Hybridization with Fluorescent Immunohistochemistry on Fresh Frozen or Fixed Mouse Brain Sections

Fluorescent in situ hybridization (FISH) is a molecular technique that identifies the presence and spatial distribution of specific RNA transcripts within cells. Neurochemical phenotyping of functionally identified neurons usually requires concurrent labelling with multiple antibodies (targeting protein) using immunohistochemistry (IHC) and optimization of in situ hybridization (targeting RNA), in tandem. A &#34;neurochemical signature&#34; to characterize particular neurons may be achieved however complicating factors include the need to verify FISH and IHC targets before combining the methods, and the limited number of RNAs and proteins that may be targeted simultaneously within the same tissue section.
Here we describe a protocol, using both fresh frozen and fixed mouse brain preparations, which detects multiple mRNAs and proteins in the same brain section using RNAscope FISH followed by fluorescence immunostaining, respectively. We use the combined method to describe the expression pattern of low abundance mRNAs (e.g., galanin receptor 1) and high abundance mRNAs (e.g., glycine transporter 2), in immunohistochemically identified brainstem nuclei.
Key considerations for protein labelling downstream of the FISH assay extend beyond tissue preparation and optimization of FISH probe labelling. For example, we found that antibody binding and labelling specificity can be detrimentally affected by the protease step within the FISH probe assay. Proteases catalyze hydrolytic cleavage of peptide bonds, facilitating FISH probe entry into cells, however they may also digest the protein targeted by the subsequent IHC assay, producing off target binding. The subcellular location of the targeted protein is another factor contributing to IHC success following FISH probe assay. We observed IHC specificity to be retained when the targeted protein is membrane bound, whereas IHC targeting cytoplasmic protein required extensive troubleshooting. Finally, we found handling of slide-mounted fixed frozen tissue more challenging than fresh frozen tissue, however IHC quality was overall better with fixed frozen tissue, when combined with RNAscope.

Combining Multiplex Fluorescence In Situ Hybridization with Fluorescent Immunohistochemistry on Fresh Frozen or Fixed Mouse Brain Sections

This protocol describes a method for combining fluorescence in situ hybridization (FISH) and fluorescence immunohistochemistry (IHC) in both fresh frozen and fixed mouse brain sections, with the goal of achieving multilabel FISH and fluorescence IHC signal. IHC targeted cytoplasmic and membrane attached proteins.

Fluorescent in situ hybridization (FISH) is a molecular technique that identifies the presence and spatial distribution of specific RNA transcripts within ...

3D Visualization of Bone Remodeling Under Tensile Load in Suture Expansion Mice

A 3-D Visualization Technique for Bone Remodeling in a Suture Expansion Mouse Model

Craniofacial sutures play a crucial role beyond being fibrous joints connecting craniofacial bones; they also serve as the primary niche for calvarial and facial bone growth, housing mesenchymal stem cells and osteoprogenitors. As most craniofacial bones develop through intramembranous ossification, the sutures' marginal regions act as initiation points. Due to this significance, these sutures have become intriguing targets in orthopedic therapies like spring-assisted cranial vault expansion, rapid maxillary expansion, and maxillary protraction. Under orthopedic tracing force, suture stem cells are rapidly activated, becoming a dynamic source for bone remodeling during expansion. Despite their importance, the physiological changes during bone remodeling periods remain poorly understood. Traditional sectioning methods, primarily in the sagittal direction, do not capture the comprehensive changes occurring throughout the entire suture. This study established a standard mouse model for sagittal suture expansion. To fully visualize bone remodeling changes post-suture expansion, the PEGASOS tissue clearing method was combined with whole-mount EdU staining and calcium chelating double labeling. This allowed the visualization of highly proliferating cells and new bone formation across the entire calvarial bones following expansion. This protocol offers a standardized suture expansion mouse model and a 3-D visualization method, shedding light on the mechanobiological changes in sutures and bone remodeling under tensile force loading.

Author Spotlight: PEGASOS Tissue Clearing Technique to Visualize Bone Remodeling 

This protocol presents a standardized suture expansion mouse model and a 3-D visualization method to study the mechanobiological changes of the suture and bone remodeling under tensile force loading.

Craniofacial sutures play a crucial role beyond being fibrous joints connecting craniofacial bones; they also serve as the primary niche for calvarial and ...

Immunostaining of Mouse Craniofacial Tissues

Source: Yang, J., et al., <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/59113">Tissue Preparation and Immunostaining of Mouse Craniofacial Tissues and Undecalcified Bone.</a> J. Vis. Exp. (2019)
This video demonstrates the immunofluorescence staining of cryosections from mouse embryonic craniofacial tissues, using fluorescence microscopy to assess nuclear marker expression and determine cell proliferation and signaling activity.

Disclaimer text: All procedures involving animal samples have been reviewed and approved by the appropriate animal ethical review committee.&#160;&#160; ...

Tissue Preparation and Immunostaining of Mouse Craniofacial Tissues and Undecalcified Bone

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