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 s...

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 border="0" cellpadding="0" cellspacing="0" style="width:752px;" width="752">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">Adhesive tape</td>
			<td style="width:171px;">Leica</td>
			<td style="width:119px;">#39475214</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:296px;">Alexa fluor 488-goat anti-Rabbit secondary antibody</td>
			<td style="width:171px;">Invitrogen</td>
			<td style="width:119px;">A-11034</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">Antifade Mountant with DAPI</td>
			<td style="width:171px;">Invitrogen</td>
			<td style="width:119px;">P36931</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">Bovine serum albumin</td>
			<td style="width:171px;">Sigma</td>
			<td style="width:119px;">A2153</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">Coverslips</td>
			<td style="width:171px;">Fisher Brand</td>
			<td style="width:119px;">12-545-E</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">Cryostat</td>
			<td style="width:171px;">Leica</td>
			<td style="width:119px;">CM1850</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">EDTA</td>
			<td style="width:171px;">Sigma</td>
			<td style="width:119px;">E6758</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">Fluorescence microscope</td>
			<td style="width:171px;">Olympus</td>
			<td style="width:119px;">BX51</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">Gelatin</td>
			<td style="width:171px;">Sigma</td>
			<td style="width:119px;">G1890</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">In Situ&#160;Cell Death Detection Kit</td>
			<td style="width:171px;">Millipore</td>
			<td style="width:119px;">S7165</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">Microscope slides</td>
			<td style="width:171px;">Fisher Brand</td>
			<td style="width:119px;">12-550-15</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">OCT Compound</td>
			<td style="width:171px;">Fisher Healthcare</td>
			<td style="width:119px;">23-730-571</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">Paraformaldehyde (PFA)</td>
			<td style="width:171px;">Sigma</td>
			<td style="width:119px;">P6148&#160;</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">Phosphate buffered saline (PBS)</td>
			<td style="width:171px;">Sigma</td>
			<td style="width:119px;">P4417</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">Polyethylene glycol&#160;tert-octylphenyl ether</td>
			<td style="width:171px;">Sigma</td>
			<td style="width:119px;">T9284</td>
			<td style="width:167px;">Triton X-100</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">Proteinase K</td>
			<td style="width:171px;">Invitrogen</td>
			<td style="width:119px;">AM2542</td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:296px;">Rabbit anti-Ki67 antibody</td>
			<td style="width:171px;">Cell Signaling Technology</td>
			<td style="width:119px;">9129</td>
			<td>Lot#:3; RRID:AB_2687446</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:296px;">Rabbit anti-pSmad1/5/9 antibody</td>
			<td style="width:171px;">Cell Signaling Technology</td>
			<td style="width:119px;">13820</td>
			<td>Lot#:3; RRID:AB_2493181</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">Sodium citrate</td>
			<td style="width:171px;">Sigma</td>
			<td style="width:119px;">1613859</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">Sucrose</td>
			<td style="width:171px;">Sigma</td>
			<td style="width:119px;">S9378</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:296px;">Tris</td>
			<td style="width:171px;">Sigma</td>
			<td style="width:119px;">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 pro...

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

Research

JoVE Journal

Developmental Biology

12.3K Views.  Wuhan University. 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.

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

Automated Quantification of Hematopoietic Cell &#8211; Stromal Cell Interactions in Histological Images of Undecalcified Bone

Confocal microscopy is the method of choice for the analysis of localization of multiple cell types within complex tissues such as the bone marrow. However, the analysis and quantification of cellular localization is difficult, as in many cases it relies on manual counting, thus bearing the risk of introducing a rater-dependent bias and reducing interrater reliability. Moreover, it is often difficult to judge whether the co-localization between two cells results from random positioning, especially when cell types differ strongly in the frequency of their occurrence. Here, a method for unbiased quantification of cellular co-localization in the bone marrow is introduced. The protocol describes the sample preparation used to obtain histological sections of whole murine long bones including the bone marrow, as well as the staining protocol and the acquisition of high-resolution images. An analysis workflow spanning from the recognition of hematopoietic and non-hematopoietic cell types in 2-dimensional (2D) bone marrow images to the quantification of the direct contacts between those cells is presented. This also includes a neighborhood analysis, to obtain information about the cellular microenvironment surrounding a certain cell type. In order to evaluate whether co-localization of two cell types is the mere result of random cell positioning or reflects preferential associations between the cells, a simulation tool which is suitable for testing this hypothesis in the case of hematopoietic as well as stromal cells, is used. This approach is not limited to the bone marrow, and can be extended to other tissues to permit reproducible, quantitative analysis of histological data.

A strategy to quantitatively analyze histological data in the bone marrow is presented. Confocal microscopy of fluorescently labeled cells in tissue sections results in 2-dimensional images, which are automatically analyzed. Co-localization analyses of different cell types are compared to data from simulated images, giving quantitative information about cellular interactions.

Confocal microscopy is the method of choice for the analysis of localization of multiple cell types within complex tissues such as the bone marrow. ...

Ex Utero Electroporation and Organotypic Slice Culture of Mouse Hippocampal Tissue

Mouse genetics offers a powerful tool determining the role of specific genes during development. Analyzing the resulting phenotypes by immunohistochemical and molecular methods provides information of potential target genes and signaling pathways. To further elucidate specific regulatory mechanisms requires a system allowing the manipulation of only a small number of cells of a specific tissue by either overexpression, ablation or re-introduction of specific genes and follow their fate during development. To achieve this ex utero electroporation of hippocampal structures, especially the dentate gyrus, followed by organotypic slice culture provides such a tool. Using this system to generate mosaic deletions allows determining whether the gene of interest regulates cell-autonomously developmental processes like progenitor cell proliferation or neuronal differentiation. Furthermore it facilitates the rescue of phenotypes by re-introducing the deleted gene or its target genes. In contrast to in utero electroporation the ex utero approach improves the rate of successfully targeting deeper layers of the brain like the dentate gyrus. Overall ex utero electroporation and organotypic slice culture provide a potent tool to study regulatory mechanisms in a semi-native environment mirroring endogenous conditions.

Ex Utero Electroporation and Organotypic Slice Culture of Mouse Hippocampal Tissue

Here we present a protocol providing a tool to examine regulatory mechanisms of specific genes during hippocampal development. Employing ex utero electroporation and organotypic slice culture allows the up- and down-regulation of the expression of genes of interest in single cells and follow their fate during development.

Mouse genetics offers a powerful tool determining the role of specific genes during development. Analyzing the resulting phenotypes by immunohistochemical ...

Generating Primary Fibroblast Cultures from Mouse Ear and Tail Tissues

Primary cells are derived directly from tissue and are thought to be more representative of the physiological state of cells in vivo than established cell lines. However, primary cell cultures usually have a finite life span and need to be frequently re-established. Fibroblasts are an easily accessible source of primary cells. Here, we discuss a simple and quick experimental procedure to establish primary fibroblast cultures from ears and tails of mice. The protocol can be used to establish primary fibroblast cultures from ears stored at RT for up to 10 days. When the protocol is carefully followed, contaminations are unlikely to occur despite the use of non-sterile tissue stored for extended time in some cases. Fibroblasts proliferate rapidly in culture and can be expanded to substantial numbers before undergoing replicative senescence.

We describe a simple and quick experimental procedure for generating primary fibroblasts from the ears and tails of mice. The procedure does not require special animal training and can be used for the generation of fibroblast cultures from ears stored at RT for up to 10 days.

Primary cells are derived directly from tissue and are thought to be more representative of the physiological state of cells in vivo than established cell ...

Identification of Critical Conditions for Immunostaining in the Pea Aphid Embryos: Increasing Tissue Permeability and Decreasing Background Staining

The pea aphid Acyrthosiphon pisum, with a sequenced genome and abundant phenotypic plasticity, has become an emerging model for genomic and developmental studies. Like other aphids, A. pisum propagate rapidly via parthenogenetic viviparous reproduction, where the embryos develop within egg chambers in an assembly-line fashion in the ovariole. Previously we have established a robust platform of whole-mount in situ hybridization allowing detection of mRNA expression in the aphid embryos. For analyzing the expression of protein, though, established protocols for immunostaining the ovarioles of asexual viviparous aphids did not produce satisfactory results. Here we report conditions optimized for increasing tissue permeability and decreasing background staining, both of which were problems when applying established approaches. Optimizations include: (1) incubation of proteinase K (1 &#181;g/ml, 10 min), which was found essential for antibody penetration in mid- and late-stage aphid embryos; (2) replacement of normal goat serum/bovine serum albumin with a blocking reagent supplied by a Digoxigenin (DIG)-based buffer set and (3) application of methanol rather hydrogen peroxide (H2O2) for bleaching endogenous peroxidase; which significantly reduced the background staining in the aphid tissues. These critical conditions optimized for immunostaining will allow effective detection of gene products in the embryos of A. pisum and other aphids.

A protocol of whole-mount immunostaining on aphid embryos is presented, in which critical conditions for decreasing background staining and increasing tissue permeability are addressed. These conditions are specifically developed for effective detection of protein expression in the embryonic tissues of aphids.

The pea aphid Acyrthosiphon pisum, with a sequenced genome and abundant phenotypic plasticity, has become an emerging model for genomic and developmental ...

Bioengineering of Humanized Bone Marrow Microenvironments in Mouse and Their Visualization by Live Imaging

Human hematopoietic stem cells (HSCs) reside in the bone marrow (BM) niche, an intricate, multifactorial network of components producing cytokines, growth factors, and extracellular matrix. The ability of HSCs to remain quiescent, self-renew or differentiate, and acquire mutations and become malignant depends upon the complex interactions they establish with different stromal components. To observe the crosstalk between human HSCs and the human BM niche in physiological and pathological conditions, we designed a protocol to ectopically model and image a humanized BM niche in immunodeficient mice. We show that the use of different cellular components allows for the formation of humanized structures and the opportunity to sustain long-term human hematopoietic engraftment. Using two-photon microscopy, we can live-image these structures in situ at the single-cell resolution, providing a powerful new tool for the functional characterization of the human BM microenvironment and its role in regulating normal and malignant hematopoiesis.

A method to create and live-image different humanized bone-marrow niches in mice is presented. Based on the supportive niche created by human mesenchymal cells, the addition of human endothelial cells induces the formation of human vessels, while the addition of rhBMP-2 induces the formation of human-mouse chimeric mature bone tissue.

Human hematopoietic stem cells (HSCs) reside in the bone marrow (BM) niche, an intricate, multifactorial network of components producing cytokines, growth ...

Live Imaging of Mouse Secondary Palate Fusion

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to cleft secondary palate, a common congenital anomaly in humans. Secondary palate fusion has been extensively studied leading to several proposed cellular mechanisms that may mediate this process. However, these studies have been mostly performed on fixed embryonic tissues at progressive timepoints during development or in fixed explant cultures analyzed at static timepoints. Static analysis is limited for the analysis of dynamic morphogenetic processes such a palate fusion and what types of dynamic cellular behaviors mediate palatal fusion is incompletely understood. Here we describe a protocol for live imaging of ex vivo secondary palate fusion in mouse embryos. To examine cellular behaviors of palate fusion, epithelial-specific Keratin14-cre was used to label palate epithelial cells in ROSA26-mTmGflox reporter embryos. To visualize filamentous actin, Lifeact-mRFPruby reporter mice were used. Live imaging of secondary palate fusion was performed by dissecting recently-adhered secondary palatal shelves of embryonic day (E) 14.5 stage embryos and culturing in agarose-containing media on a glass bottom dish to enable imaging with an inverted confocal microscope. Using this method, we have detected a variety of novel cellular behaviors during secondary palate fusion. An appreciation of how distinct cell behaviors are coordinated in space and time greatly contributes to our understanding of this dynamic morphogenetic process. This protocol can be applied to mutant mouse lines, or cultures treated with pharmacological inhibitors to further advance understanding of how secondary palate fusion is controlled.

Here, we present a protocol for live imaging of mouse secondary palate fusion using confocal microscopy. This protocol can be used in combination with a variety of fluorescent reporter mouse lines, and with pathway inhibitors for mechanistic insight. This protocol can be adapted for live imaging in other developmental systems.

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to ...

Dissection and Coronal Slice Preparation of Developing Mouse Pituitary Gland

The pituitary gland or hypophysis is an important endocrine organ secreting hormones essential for homeostasis. It consists of two glands with separate embryonic origins and functions &#8212; the neurohypophysis and the adenohypophysis. The developing mouse pituitary gland is tiny and delicate with an elongated oval shape. A coronal section is preferred to display both the adenohypophysis and neurohypophysis in a single slice of the mouse pituitary.
The goal of this protocol is to achieve proper pituitary coronal sections with well-preserved tissue architectures from developing mice. In this protocol, we describe in detail how to dissect and process pituitary glands properly from developing mice. First, mice are fixed by transcardial perfusion of formaldehyde prior to dissection. Then three different dissecting techniques are applied to obtain intact pituitary glands depending on the age of mice. For fetal mice aged embryonic days (E) 17.5 - 18.5 and neonates up to 4 days, the entire sella regions including the sphenoid bone, gland, and trigeminal nerves are dissected. For pups aged postnatal days (P) 5 - 14, the pituitary glands connected with trigeminal nerves are dissected as a whole. For mice over 3 weeks old, the pituitary glands are carefully dissected free from the surrounding tissues. We also display how to embed the pituitary glands in a proper orientation by using the surrounding tissues as landmarks to obtain satisfying coronal sections. These methods are useful in analyzing histological and developmental features of pituitary glands in developing mice.

We present a protocol to dissect pituitary glands and prepare pituitary coronal sections from developing mice.

The pituitary gland or hypophysis is an important endocrine organ secreting hormones essential for homeostasis. It consists of two glands with separate ...

A Simplified and Efficient Method to Isolate Primary Human Keratinocytes from Adult Skin Tissue

Primary human keratinocytes isolated from fresh skin tissues and their expansion in vitro have been widely used for laboratory research and for clinical applications. The conventional isolation method of human keratinocytes involves a two-step sequential enzymatic digestion procedure, which has been proven to be inefficient in generating primary cells from adult tissues due to the low cell recovery rate and reduced cell viability. We recently reported an advanced method to isolate human primary epidermal progenitor cells from skin tissues that utilizes the Rho kinase inhibitor Y-27632 in the medium. Compared with the traditional protocol, this new method is simpler, easier, and less time-consuming, and increases epithelial stem cell yield and enhances their stem cell characteristics. Moreover, the new methodology does not require the separation of the epidermis from the dermis, and, therefore, is suitable for isolating cells from different types of adult tissues. This new isolation method overcomes the major shortcomings of conventional methods and is more suitable for producing large numbers of epidermal cells with high potency both for laboratory and for clinical applications. Here, we describe the new method in detail.

Here we present a protocol to efficiently isolate primary human keratinocytes from adult skin tissues. This method simplifies the conventional procedure by using the ROCK Inhibitor Y-27632 in the inoculation medium to spontaneously separate epidermal cells from dermal cells.

Primary human keratinocytes isolated from fresh skin tissues and their expansion in vitro have been widely used for laboratory research and for clinical ...

Dissection, Culture and Analysis of Primary Cranial Neural Crest Cells from Mouse for the Study of Neural Crest Cell Delamination and Migration

Over the past several decades there has been an increased availability of genetically modified mouse models used to mimic human pathologies. However, the ability to study cell movements and differentiation in vivo is still very difficult. Neurocristopathies, or disorders of the neural crest lineage, are particularly challenging to study due to a lack of accessibility of key embryonic stages and the difficulties in separating out the neural crest mesenchyme from adjacent mesodermal mesenchyme. Here, we set out to establish a well-defined, routine protocol for the culture of primary cranial neural crest cells. In our approach we dissect out the mouse neural plate border during the initial neural crest induction stage. The neural plate border region is explanted and cultured. The neural crest cells form in an epithelial sheet surrounding the neural plate border, and by 24 h after explant, begin to delaminate, undergoing an epithelial-mesenchymal transition (EMT) to become fully motile neural crest cells. Due to our two-dimensional culturing approach, the distinct tissue populations (neural plate versus premigratory and migratory neural crest) can be readily distinguished. Using live imaging approaches, we can then identify changes in neural crest induction, EMT and migratory behaviors. The combination of this technique with genetic mutants will be a very powerful approach for understanding normal and pathological neural crest cell biology.

This protocol describes the dissection and culture of cranial neural crest cells from mouse models, primarily for the study of cell migration. We describe the live imaging techniques used and the analysis of speed and cell shape changes.

Over the past several decades there has been an increased availability of genetically modified mouse models used to mimic human pathologies. However, the ...

Clonal Genetic Tracing using the Confetti Mouse to Study Mineralized Tissues

Labeling an individual cell in the body to monitor which cell types it can give rise to and track its migration through the organism or determine its longevity can be a powerful way to reveal mechanisms of tissue development and maintenance. One of the most important tools currently available to monitor cells in vivo is the Confetti mouse model. The Confetti model can be used to genetically label individual cells in living mice with various fluorescent proteins in a cell type-specific manner and monitor their fate, as well as the fate of their progeny over time, in a process called clonal genetic tracing or clonal lineage tracing. This model was generated almost a decade ago and has contributed to an improved understanding of many biological processes, particularly related to stem cell biology, development, and renewal of adult tissues. However, preserving the fluorescent signal until image collection and simultaneous capturing of various fluorescent signals is technically challenging, particularly for mineralized tissue. This publication describes a step-by-step protocol for using the Confetti model to analyze growth plate cartilage that can be applied to any mineralized or nonmineralized tissue.

This method describes the use of the R26R-Confetti (Confetti) mouse model to study mineralized tissues, covering all steps from the breeding strategy to the image acquirements. Included is a general protocol that can be applied to all soft tissues and a modified protocol that can be applied to mineralized tissues.

Labeling an individual cell in the body to monitor which cell types it can give rise to and track its migration through the organism or determine its ...