We thank Meghann K. Holt for editing the manuscript. This study was supported by NIH/NIDCR grants DE026461 and DE028345 and the startup funding from the Texas A&#38;M School of Dentistry to Dr. Xiaofang Wang.

All methods described here have been approved by the Institutional Animal Care and Use Committee (IACUC) for Texas A&#38;M University College of Dentistry.
1. Preparation of the EdU labeling cocktail
<ol>
	<li>Prepare the cocktail stock solutions as described in Table 1.</li>
	<li>Prepare the EdU labeling cocktail as described in Table 1.</li>
</ol>
2. Preparation of the mice and EdU solution
<ol>
	<li>Prepare the EdU solution. Dissolve EdU in DMSO at 20 mg/mL and store in a -20 &#176;C freezer. 
	CAUTION: EdU is a mutagen. Users should wear appropriate personal protective equipment (PPE) when handling this substance.</li>
	<li>Inject 5-day-old C57BL/6J mice with the prepared EdU solution intraperitoneally at 3 &#181;L/g body weight. Mice must receive the solution once a day for 7 consecutive days. Next, they undergo a chase period for 6 weeks before being harvested for analysis (Figure 1). 
	&#8203;CAUTION: Be careful to avoid needle pricking when injecting EdU into the mice. After injecting EdU for 7 consecutive days, place the mice in a new cage. After the mice are in their new home, dispose of the bedding in the old cage using appropriate biohazard rules.</li>
</ol>
3. Sample preparation by trans-cardiac perfusion (postnatal 53-day-old mice after the 6-week chase period)
NOTE: The mouse liver should turn pale after&#160;successful trans-cardiac perfusion.
<ol>
	<li>Anesthetize mice with an intraperitoneal injection. They need to be given a combination of xylazine and ketamine anesthetics (Xylazine 10-12.5 mg/kg; Ketamine, 80-100 mg/kg body weight).</li>
	<li>Wait for ~10 min until the mouse no longer responds to painful stimuli (such as paw pinch reflexes).</li>
	<li>Place the mice in a supine position stabilized on&#160;Styrofoam support with needles in each paw.</li>
	<li>Expose the chest cavity by opening up the overlying skin using tweezers and dissecting scissors.</li>
	<li>Cut open the diaphragm using sharp scissors. Take care not to pierce the heart.</li>
	<li>Cut through the ribcage, grabbing the base of the sternum with clamp scissors to expose the heart.</li>
	<li>Transfer the mouse to the perfusion stage near the circulation pump in a fume hood.</li>
	<li>Insert the 25 G needle (from the tubing filled with phosphate-buffered saline (PBS)/4% paraformaldehyde (PFA) solution) into the apex of the left ventricle. Take care to keep the tip of the needle in the lumen of the ventricle. 
	CAUTION: PFA is moderately toxic and a probable carcinogen. Use PPE to handle PFA and prepare the solution under a chemical fume hood. Waste PFA requires proper disposal per institution guidelines.</li>
	<li>Cut the right ventricle using a sharp scissor immediately after inserting the needle into the left ventricle. This step allows for blood to flow out of the mouse and drain into the collecting dish.</li>
	<li>Perfuse the samples with 50 mL of PBS solution at a flow rate of 7 mL/min.</li>
	<li>After PBS perfusion, switch the stopcock to allow for a flow of 20 mL of 4% PFA solution at the same flow rate of 5 mL/min.</li>
	<li>While the circulation pump is still on, remove the needle from the left ventricle. 
	&#8203;NOTE: The mouse is now fixed for tissue collection. To preserve the fluorescence, light exposure should be avoided whenever possible throughout the entire protocol. While the tissue is processing, the 50 mL conical tube with the tissue samples may be covered with aluminum foil.</li>
</ol>
4. Tissue clearing of the mandible using the PEGASOS technique
NOTE: A 15 mL or 50 mL conical tube as per the volume of tissues can be used to keep the samples ready for treatment in each step. Samples are processed at 37 &#176;C shakers (~100 rpm) from steps 4.2 to 4.7. Use polypropylene-based plastic containers that are resistant to organic solvents to avoid melting&#160;the plastic. Alternatively, glassware can be used.
CAUTION: The PEGASOS tissue clearing technique uses toxic solutions such as Quadrol, polyethylene glycol (PEG), benzyl benzoate (BB), MMA500, etc. Appropriate PPE is required to avoid potential exposures.
<ol>
	<li>Dissect mandibles and further fix samples in 4% PFA. Keep them at room temperature overnight.</li>
	<li>Remove the muscles from the mandibles and immerse them in 0.5 M EDTA solution (pH 8.0) to decalcify for 4 days. During this period, perform a daily medium change on a 37 &#176;C shaker. 
	NOTE: It is desirable to remove the muscles from the sample as much as possible. This precaution will simplify the imaging process and reduce the autofluorescence from the muscles.</li>
	<li>Decolorize the mandibles with a 25% Quadrol solution (v/v in H2O) on a 37 &#176;C shaker for one day to remove the remaining blood heme.</li>
	<li>Complete whole mount EdU staining.
	<ol>
		<li>Prepare a fresh EdU labeling cocktail according to Table 1. 
		NOTE: The EdU labeling cocktail needs to be made fresh each time with freshly prepared ascorbate solution added for obtaining optimum EdU labeling results.</li>
		<li>Rinse the mandibles with PBST for 30 min on a 37 &#176;C shaker.</li>
		<li>Rinse the mandibles with PBS three times for 3 min each time.</li>
		<li>Incubate the mandibles in a freshly made EdU labeling cocktail on a 37 &#176;C shaker overnight.</li>
		<li>Rinse the mandibles with PBS three times for 3 min each time.</li>
	</ol>
	</li>
	<li>Perform serial de-lipidation in each of the following solutions for 6 h on 37 &#176;C: 
	30%&#160;tert-butanol (tB) solution: 75% v/v H2O, 22% v/v tB, and 3% v/v Quadrol. 
	50%&#160;tB solution: 50% v/v H2O, 47% v/v tB, and 3% v/v Quadrol. 
	70%&#160;tB solution: 30% v/v H2O, 67% v/v tB, and 3% v/v Quadrol 
	NOTE: The de-lipidation step is critical. De-lipidation can be done between 4-6 h in 30% and 50% tB solution and for 1 day in 70% tB solution.</li>
	<li>Dehydrate the mandibles in tB-PEG solution for 2 days with the following daily medium change: 75% tB, 22% v/v polyethylene glycol methyl-ether methacrylate average MMA500, and 3% v/v Quadrol at 37 &#176;C.</li>
	<li>Clear the mandible in benzyl benzoate (BB)-PEG clearing medium to obtain refractive index matching: 75% v/v BB, 22% v/v PEG-MMA500, and 3% Quadrol until transparency is achieved. 
	NOTE: We have information on refractive index matching. The inorganic and organic components in tissues such as water, lipids, proteins, minerals, organelles, etc. have a different refractive index (RI) in the range of 1.33 to 1.55. This RI mismatch between the tissue components hinders the imaging process. Transparency can be achieved by eliminating RI mismatch within the tissue. Immersing the tissue-cleared samples in the clearing medium of BB-PEG is advised. This step will give the uniform internal RI of ~1.54 (called RI matching) to the whole specimen, easing the imaging process.</li>
	<li>Preserve the mandibles in the BB-PEG tissue clearing medium at room temperature for storage and imaging. 
	&#8203;NOTE: It is good to complete the imaging as soon as possible to avoid the gradual loss of fluorescent reporters. However, the PEGASOS method will keep fluorescence reporters well preserved even after several weeks of storage20,21. For long-term storage, the samples can also be stored at 4&#176;C.</li>
</ol>
5. Confocal imaging of the tissue-cleared mandible
<ol>
	<li>Mount the tissue cleared whole mandible on brand cavity slides in the BB-PEG clearing medium and cover with glass cover slides. Avoid any bubbles.</li>
	<li>Perform image acquisition with a suitable laser scanning confocal microscope. Select Acquire and set the image acquisition parameters at 512 x 512 pixel resolution and 400 Hz acquisition speed. These settings are found in the acquisition mode menu of the software that controls the microscope.</li>
	<li>In the panel for fluorescent excitation, activate the laser required (TRITC) to optimally excite fluorophores (the probe we used for LRCs visualization has fluorescent properties such as Cy3 with excitation at 548 nm and emission at 563 nm of laser wavelength).</li>
	<li>In the panel for fluorescent detection, move the slider to select the wavelengths that will be measured (for example, between 555 to 625 nm for a Cy3-like fluorophore).</li>
	<li>Place the mounted mandible on the microscope stage to visualize and find the sample using white light and a lower magnification objective (2-10x).</li>
	<li>Add a drop of immersion oil with a refractive index of 1.52 on top of the coverslip to match the refractive index of the glass coverslip and objective. 
	NOTE: Match the refractive index of objectives, imaging media, and tissue as closely as possible to minimize the introduction of optical distortions during image acquisition.</li>
	<li>Turn on the fluorescent lamps and choose an appropriate magnification objective to image regions of interest. We used a 20x objective lens with a&#160;numerical aperture of 0.9 with a working distance of 1.95 mm. A longer working distance lens may be required when imaging larger tissue samples in the confocal microscope. Alternatively, imaging can be done easily using a light-sheet microscope for larger tissue samples.</li>
	<li>On the confocal imaging software, press the Live button to start live-scan mode. To maximize the dynamic range of the images and avoid pixel saturation, adjust the gain and laser intensity for the channel selected.</li>
	<li>Identify the field of view to visualize the entire region of the mandible in the X and Y dimensions. Set the upper and lower Z-stage acquisition parameters that accurately cover the volume of the stem cell niche in the mice incisor. Use a Z-step size of 2 &#181;m or as per your requirements. The newer versions of confocal microscopes should be able to automatically acquire z-stacks of the various overlapping regions.</li>
	<li>Acquire and save Z-stack image files in .lif format. 
	&#8203;NOTE: The .lif flies can be converted into .tiff files and used for other 3D analysis software.</li>
</ol>
6. Image processing, 3D image reconstruction, and quantification of label-retaining cells by creating a surface, segmentation of the Region of Interest (ROI), masking the ROI, and creating spots
NOTE: We used Imaris (Bitplane 9.0.1) for image processing and 3D reconstruction, but similar image processing steps can be conducted using other software suites (e.g., ImageJ/Fiji, 3D slicer, Avizo, Arivis, Amira, etc.).
<ol>
	<li>In the imaging software, click the Arena button, then choose Image. Select the original .lif file and choose merged file, the images will be imported into the imaging software. 
	NOTE:&#160;Alternatively, use an imaging software file converter to convert Z stack image file to the native format file type for the software. For example: convert the .lif files into .ims files using the Imaris File Converter and open the .ims files in the imaging software. Converting data to the native file format of the software will allow rapid file conversion and minimize errors.</li>
	<li>Double click the imported file to open the image data set.</li>
	<li>Visit the Display Adjustment panel,click the name of the channel. It is usually labeled as Red in default mode.
	<ol>
		<li>In the Image Properties window, click on the Mapped Color tab. In the Color Table File option, choose Fire Flow, and then click on OK. The display color is normally chosen to distinguish the background fluorescence and the positive fluorescence from the sample.</li>
	</ol>
	</li>
	<li>On the upper left side of the screen in the Scene &#8211; Properties pane, click on the blue icon labeled Add New Surface. Unselect the Volume icon which will remove most of the background fluorescence and the positive fluorescence are distinctly visible (see Step 6.4.2).
	<ol>
		<li>Start the process of manual segmentation of region of interest (ROI) by clicking on the Create tab and selecting the Skip Automatic Creation, Edit Manually tab. In the manual surface creation wizard, click on the Contour tab to choose the orientation (XY, or YZ, or XZ) based on the samples to easily contour the ROI. Here, we choose XY orientation.</li>
		<li>Next, make sure the pointer menu in the right corner is in Select mode. Adjust the Slice Position to locate the ROI in the tissue. You will notice that most of the background fluorescence have been removed with distinct positive fluorescence as mentioned in Step 6.4. Click on the Draw button and draw a tentative region of interest. Select the Visibility option to none to avoid interference from other areas than ROI.</li>
		<li>Repeat step 6.4.2. slice by slice to draw the entire region of interest.</li>
		<li>After the ROI drawing is finished, click on Create Surface to get a 3D geometry of the ROI.</li>
		<li>Click on the Edit button in the manual surface creation wizard and select Mask All.</li>
		<li>In the popped-out Mask Channel window, select Channel 1 in the channel selection options. Afterward, select Duplicate Channel Before Applying the Mask. In the mask settings, select Constant Inside/Outside and set the voxel outside surface to 0.</li>
		<li>In the Scene-Properties pane, deselect the Surface Object and select the Volume Object. In the Display Adjustment, deselect the original &#34;Red&#8221; channel and select the Masked Red channel and adjust the color intensities to get the desired 3D-rendered and positively labeled fluorescence ROI.</li>
	</ol>
	</li>
	<li>Create a new spot object. Begin by clicking on the Add on&#160;Spots icon to quantify the 3D-rendered LRCs by creating 3D spots that comparably overlap with 3D-rendered LRCs. Use the bottom arrows to move between steps in the spot creation wizard.
	<ol>
		<li>In the spot creation wizard, there will be four sequential steps of instructions to create spots automatically with the software. Make sure the Segment Only a Region of Interest option is selected in the first step. Click on the Next icon.</li>
		<li>In the second step, there will be an &#34;XYZ 3D box&#34;. Adjust the XYZ box to cover the ROI. Click on the Next icon.</li>
		<li>In the third step, go to the Source Channel options. Choose the masked red channel; input the estimated XY spot diameter comparable to fit and overlap the sample fluorescence dots. Click on the Next icon.</li>
		<li>In the fourth step, add the &#34;Quality&#34; filter type and adjust the lower intensity threshold by moving the sliding window across the &#39;Quality&#39; histogram until the majority of identifiable LRCs are labeled. 
		NOTE: The segmentation and therefore statistical readouts (example: number of cells) very much depend on the intensity threshold values. Be careful to precisely set the appropriate threshold values.</li>
		<li>Click on the Finish button and select the Statistic button. Select the Overall tab to find the value for the total number of spots in the image.</li>
		<li>Export statistics by clicking on the Tab Display to File button and receive the results in an .xls file.</li>
		<li>Use a suitable diagram to present the quantification results.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Multiple doses of injections (BrdU, EdU) are usually used on growing neonatal mice to label proliferating cells as much as possible1,6,13. The chasing period is considered a critical step regarding the renewal rate of tissues6,13. The mouse incisor renews itself around every month. This trait allows researchers to set the chasing period to 4 weeks or longer<sup class="xr...

The continuously growing mouse incisor is an excellent model for studying adult stem cells1. The epithelial (labial and lingual cervical loop) and mesenchymal stem cells (between the labial and lingual cervical loop) that reside on the proximal side of the incisor differentiate into ameloblasts, odontoblasts, and dental pulp cells. This unique process provides a source of cells for compensation of tissue loss and turnover2. Though several stem cell markers such as Sox2, Gli1, Thy1/CD90, Bmi1, etc. have been identified in vivo for the subsets of adult stem cells in mouse incisors, they are inadequate in representing the stem cell populations when used alone1,3,4,5. Visualizing long-living quiescent cells by nucleotide analog DNA labeling and retention could provide unbiased detection for most subsets of adult stem cells6. Further, this approach is useful among many stem cell identification methods7 for understanding cell behavior and the homeostasis of dental stem cell populations3,8. While the dividing stem cells would lose their DNA labeling after a considerable chase, the putative quiescent stem cells retain their DNA label, deeming them label-retaining cells (LRCs)6. DNA labeling and retention by non-dividing stem cells will mark and locate the putative adult stem cells in their niches.
Over the past years, thymidine analog 5-bromo-2&#8242;-deoxyuridine (BrdU) labeling replaced the cumbersome, time-consuming, and high-resolution microscopy incompatible 3H-thymidine DNA labeling method for cell proliferation assays6,9. In recent years, the 5-ethynyl-2&#180;-deoxyuridine (EdU) labeling technique has been increasingly used over BrdU. This pattern emerged due to several reasons. First, the BrdU method is slow and labor-intensive. User conditions are variable, and it is unable to preserve the ultrastructure in specimens (due to DNA denaturation). Likewise, the BrdU method loses the antigenicity of cells, thus making it inefficient for downstream functional analyses and assays such as the co-localization experiments and in vivo stem cell transplantation3,7,9,10,11,12. BrdU is also a teratogen, which is not suitable for labeling LRCs in embryonic development6. Also, the BrdU method is inefficient when used in the whole-mount specimens. The disadvantages are low penetration of antibodies in the deep part of specimens or the requirement of a long antibody incubation period for deep penetration13. EdU labeling escapes the steps of denaturing specimens, thus preserving the ultrastructure. This feature is advantageous for downstream functional analyses such as co-localization experiments and stem cell transplantation11,12. Also, EdU labeling is highly sensitive and rapid; specimen penetration is high due to the use of rapidly absorbed and smaller-sized fluorescent azides for detecting EdU labels through a Cu(I)-catalyzed [3 + 2] cycloaddition reaction (&#34;click&#34; chemistry)14.
Another increasingly applied&#160;DNA labeling method is the use of engineered transgenic mice. These mice express histone 2B green fluorescent protein (H2B-GFP) controlled by a tetracycline-responsive regulator element5,14. After feeding mice with tetracycline chow/water for a 4-week to 4-month chasing period, the GFP fluorescence will diminish in cycling cells and only LRCs retain the fluorescence6. The advantage of this method is that the labeled LRCs can be isolated and remain viable for cell culture or downstream functional analyses6,7. Some studies reported inaccurate labeling of quiescent stem cells when chased for long-term use. This result was due to a leaky background expression from the H2B-GFP strain and not the appropriate tetracycline-regulated response15.
Moreover, most literature in the past used the LRCs detection mainly on sectioned slides, which are two dimensional and often erroneously biased in showing the accurate location and number of LRCs. The approach displayed incorrect angles for sections of complex tissue structures16. The other method was to obtain 3D images from serial sections and perform post-image reconstructions. These steps were inaccurate because of image distortion from variations in each serial section due to compressed or stretched sections, resulting in missing information16,17,18. The method was also laborious and time-consuming.
To facilitate the whole-mount imaging of LRCs, samples need to be made clear while the fluorescence be&#160;well maintained. Current tissue clearing techniques can be classified into three major categories: organic solvent-based tissue clearing techniques, aqueous reagent-based tissue clearing techniques, and hydrogel-based tissue clearing techniques17,19. The polyethylene glycol (PEG)-associated solvent system (PEGASOS) has been recently developed. This approach renders nearly all types of tissues transparent and preserves endogenous fluorescence, including hard tissues such as bone and teeth20. The PEGASOS method has advantages over other tissue clearing methods, especially in clearing tooth and bone materials. Most other methods could only partially clear hard tissues, have long processing times, or require costly reagents21. Also, the PEGASOS method can efficiently preserve endogenous fluorescence over other methods.
This literature led us to create a new method for cell study. We combined the LRCs detection advantages of EdU&#160;labeling with the most superior 3D whole-mount imaging of tissue-cleared specimens; samples were processed with&#160;advanced polyethylene glycol (PEG)-associated solvent system (PEGASOS) tissue clearing technique15. Hard tissue transparency enabled us to reconstruct the 3D signal of LRCs fluorescence in vivo without breaking the teeth or mandible, creating a more accurate way to visualize and quantify LRCs.
In this study, we provide an innovative guide to visualize LRCs in the mouse incisor. We made a 3D visual approach to determine the location and quantity of LRCs within the mouse incisor stem cell niche. This project used EdU labeling, PEGASOS tissue clearing techniques, and confocal microscopy. Our method of EdU labeling the LRCs on whole-mount tissue and the use of a cleared and transparent specimen overcomes both the limitations of traditional sectioned slides and other disadvantageous DNA labeling methods. Thus, our technique will be suitable for studies on stem cell homeostasis requiring LRCs detection, especially on hard tissues. The protocol can be equally advantageous to those focusing on stem cell homeostasis in other tissues and organs.

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	<tbody>
		<tr>
			<td>0.5 M EDTA</td>
			<td>Sigma Aldrcih</td>
			<td>E9884</td>
			<td></td>
		</tr>
		<tr>
			<td>20 &#215; Objective/NA 0.9</td>
			<td>Leica</td>
			<td>507702</td>
			<td></td>
		</tr>
		<tr>
			<td>50 mL Falcon Centrifuge Tubes</td>
			<td>Falcon</td>
			<td>352070</td>
			<td></td>
		</tr>
		<tr>
			<td>BD PrecisionGlide Needle</td>
			<td>BD</td>
			<td>REF 305111</td>
			<td></td>
		</tr>
		<tr>
			<td>Bezyl benzoate (BB)</td>
			<td>Sigma Aldrcih</td>
			<td>409529</td>
			<td></td>
		</tr>
		<tr>
			<td>Bitplane 9.0.1</td>
			<td>Imaris</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>BRAND cavity slides</td>
			<td>Millipore Sigma</td>
			<td>BR475505</td>
			<td></td>
		</tr>
		<tr>
			<td>C57BL/6J mice</td>
			<td>Jackson Laboratory</td>
			<td>Strain #:000664</td>
			<td></td>
		</tr>
		<tr>
			<td>Circulation Pump</td>
			<td>VWR</td>
			<td>23609-170</td>
			<td></td>
		</tr>
		<tr>
			<td>CuSO4</td>
			<td>Sigma Aldrcih</td>
			<td>451657</td>
			<td></td>
		</tr>
		<tr>
			<td>DMSO</td>
			<td>Sigma Aldrcih</td>
			<td>D8418</td>
			<td></td>
		</tr>
		<tr>
			<td>EdU</td>
			<td>Carboynth</td>
			<td>NE08701</td>
			<td></td>
		</tr>
		<tr>
			<td>Heparin</td>
			<td>Miiilipore Sigma</td>
			<td>H3149</td>
			<td></td>
		</tr>
		<tr>
			<td>Imaging System</td>
			<td>Olympus</td>
			<td>DP27</td>
			<td></td>
		</tr>
		<tr>
			<td>LAS X Software</td>
			<td>Leica</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Olympus Stereo Microscope</td>
			<td>Olympus</td>
			<td>SZX16</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraformaldehye</td>
			<td>Sigma Aldrich</td>
			<td>P6148</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>Sigma Aldrich</td>
			<td>P4417</td>
			<td></td>
		</tr>
		<tr>
			<td>PEGMMA500</td>
			<td>Sigma Aldrich</td>
			<td>447943</td>
			<td></td>
		</tr>
		<tr>
			<td>Quadrol</td>
			<td>Sigma Aldrich</td>
			<td>122262</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium Ascorbate</td>
			<td>Sigma Aldrich</td>
			<td>11140</td>
			<td></td>
		</tr>
		<tr>
			<td>Sulfa-Cyanine 3 Azide</td>
			<td>Lumiprobe</td>
			<td>D1330</td>
			<td></td>
		</tr>
		<tr>
			<td>TBS-10X</td>
			<td>Cell Signaling Technology</td>
			<td>12498</td>
			<td></td>
		</tr>
		<tr>
			<td>TCS SP8 Confocal Microscope</td>
			<td>Leica</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>tert-butanol (tB)</td>
			<td>Sigma Aldrich</td>
			<td>360538</td>
			<td></td>
		</tr>
		<tr>
			<td>Triton X-100</td>
			<td>Sigma Aldrich</td>
			<td>X100</td>
			<td></td>
		</tr>
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</table>

detection and quantitation of label-retaining cells in mouse incisors using a 3d reconstruction approach after tissue clearing

The murine incisor is an organ that grows continuously throughout the lifespan of the mouse. The epithelial and mesenchymal stem cells residing in the proximal tissues of incisors give rise to progeny that will differentiate into ameloblasts, odontoblasts, and pulp fibroblasts. These cells are crucial in supporting the sustained turnover of incisor tissues, making the murine incisor an excellent model for studying the homeostasis of adult stem cells. Stem cells are believed to contain long-living quiescent cells that can be labeled by nucleotide analogs such as 5-ethynyl-2&#180;-deoxyuridine (EdU). The cells retain this label over time and are accordingly named&#160;label-retaining cells (LRCs). Approaches for visualizing LRCs in vivo provide a robust tool for monitoring stem cell homeostasis. In this study, we described a method for visualizing and analyzing LRCs. Our innovative approach features LRCs in mouse incisors after tissue clearing and whole-mount EdU staining followed by confocal microscopy and a 3-dimensional (3D) reconstruction with the imaging software. This method enables 3D imaging acquisition and non-biased quantitation compared to traditional LRCs analysis on sectioned slides.

After EdU labeling and the PEGASOS tissue clearing process (Figure 2), the transparent mandible was obtained as shown in our image (Figure 3B). We compared the modified sample to a normal mandible without a tissue clearing process (Figure 3A). The transparent mandible (Figure 3B) with EdU labeling was subjected to confocal imaging. We focused on the incisor apex showing the stem cell niche as shown in (...

Watch this Scientific Journal Video about Detection and Quantitation of Label-Retaining Cells in Mouse Incisors using a 3D Reconstruction Approach after Tissue Clearing at JoVE.com

Detection and Quantitation of Label-Retaining Cells in Mouse Incisors using a 3D Reconstruction Approach after Tissue Clearing

The mouse incisor contains valuable label-retaining cells in its stem cell niche. We have a novel way to unbiasedly detect and quantify the label-retaining cells; our study used EdU labeling and a 3D reconstruction approach after PEGASOS tissue clearing of the mandible.

The murine incisor is an organ that grows continuously throughout the lifespan of the mouse. The epithelial and mesenchymal stem cells residing in the ...

detection-quantitation-label-retaining-cells-mouse-incisors-using-3d

Research

JoVE Journal

Developmental Biology

1.8K Views.  Texas A&M School of Dentistry. The mouse incisor contains valuable label-retaining cells in its stem cell niche. We have a novel way to unbiasedly detect and quantify the label-retaining cells; our study used EdU labeling and a 3D reconstruction approach after PEGASOS tissue clearing of the mandible.

Video: Detection and Quantitation of Label-Retaining Cells in Mouse Incisors using a 3D Reconstruction Approach after Tissue Clearing

Loss- and Gain-of-function Approach to Investigate Early Cell Fate Determinants in Preimplantation Mouse Embryos

Gene silencing and overexpression techniques are instrumental for the identification of genes involved in embryonic development. Direct target gene modification in preimplantation embryos provides a means to study the underlying mechanisms of genes implicated in, for instance, cellular differentiation into the trophectoderm (TE) and the inner cell mass (ICM). Here, we describe a protocol that examines the role of neogenin as an authentic receptor for initial cell fate determination in preimplantation mouse embryos. First, we discuss the experimental manipulations that were used to produce gain and loss of neogenin function by microinjecting neogenin cDNA and shRNA; the effectiveness of this approach was confirmed by a strong correlation between the pair-wise expression levels of either red fluorescent protein (RFP) or green fluorescent protein (GFP) and the immunocytochemical quantification of neogenin expression. Secondly, overexpression of neogenin in preimplantation mouse embryos leads to normal ICM development while neogenin knockdown causes the ICM to develop abnormally, implying that neogenin could be a receptor that relays extracellular cues to drive blastomeres to early cell fates. Given the success of this detailed protocol in investigating the function of a novel embryonic developmental stage-specific receptor, we propose that it has the potential to aid in exploration and identification of other stage-specific genes during embryogenesis.

The goal of this protocol is to describe a loss- and gain-of function method that is applicable to identify neogenin as a stage-specific receptor that leads to trophectoderm and inner cell mass differentiation in preimplantation mouse embryos.

 Gene silencing and overexpression techniques are instrumental for the identification of genes involved in embryonic development. Direct target gene ...

The Production of Pluripotent Stem Cells from Mouse Amniotic Fluid Cells Using a Transposon System

Induced pluripotent stem (iPS) cells are generated from mouse and human somatic cells by forced expression of defined transcription factors using different methods. Here, we produced iPS cells from mouse amniotic fluid cells, using a non-viral-based transposon system. All obtained iPS cell lines exhibited characteristics of pluripotent cells, including the ability to differentiate toward derivatives of all three germ layers in vitro and in vivo. This strategy opens up the possibility of using cells from diseased fetuses to develop new therapies for birth defects.

In this study, we generate induced pluripotent stem cells from mouse amniotic fluid cells, using a non-viral-based transposon system.

Induced pluripotent stem (iPS) cells are generated from mouse and human somatic cells by forced expression of defined transcription factors using ...

A Standardized Approach for Multispecies Purification of Mammalian Male Germ Cells by Mechanical Tissue Dissociation and Flow Cytometry

Fluorescence-activated cell sorting (FACS) has been one of the methods of choice to isolate enriched populations of mammalian testicular germ cells. Currently, it allows the discrimination of up to 9 murine germ cell populations with high yield and purity. This high-resolution in discrimination and purification is possible due to unique changes in chromatin structure and quantity throughout spermatogenesis. These patterns can be captured by flow cytometry of male germ cells stained with fluorescent DNA-binding dyes such as Hoechst-33342 (Hoechst). Herein is a detailed description of a recently developed protocol to isolate mammalian testicular germ cells. Briefly, single cell suspensions are generated from testicular tissue by mechanical dissociation, double stained with Hoechst and propidium iodide (PI) and processed by flow cytometry. A serial gating strategy, including the selection of live cells (PI negative) with different DNA content (Hoechst intensity), is used during FACS sorting to discriminate up to 5 germ cell types. These include, with corresponding average purities (determined by microscopy evaluation): spermatogonia (66%), primary (71%) and secondary (85%) spermatocytes, and spermatids (90%), further separated into round (93%) and elongating (87%) subpopulations. Execution of the entire workflow is straightforward, allows the isolation of 4 cell types simultaneously with the appropriate FACS machine, and can be performed in less than 2 h. As reduced processing time is crucial to preserve the physiology of ex vivo cells, this method is ideal for downstream high-throughput studies of male germ cell biology. Moreover, a standardized protocol for multispecies purification of mammalian germ cells eliminates methodological sources of variables and allows a single set of reagents to be used for different animal models.

This work describes the standardization of a method to obtain purified germ cell populations from testicular tissue of different mammalian species. It is a straightforward protocol that combines mechanical testis dissociation, staining with Hoechst-33342 and propidium iodide, and FACS sorting, with wide applications in comparative studies of male reproductive biology.

Fluorescence-activated cell sorting (FACS) has been one of the methods of choice to isolate enriched populations of mammalian testicular germ cells. ...

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 ...

Horizontal Whole Mount: A Novel Processing and Imaging Protocol for Thick, Three-dimensional Tissue Cross-sections of Skin

Processing a tissue of interest to generate a microscopic image that supports a scientific argument can be challenging. The acquisition of high-quality microscopic images is not entirely dependent upon the quality of the microscope, but also upon the methods of tissue processing, which often involve multiple critical actions or steps. Furthermore, mesenchymal cell types in the skin and other tissues represent a new challenge for tissue preparation and imaging. Here, we present a complete process, from tissue harvest to microscopy. Our technique, called &#34;horizontal whole mount,&#34; is one that novices can quickly become proficient in and that allows for antigen preservation and detection in 60-300 &#181;m-thick sections cut with a cryostat. Sections of this thickness provide enhanced visualization of tissue microarchitecture in a three-dimensional environment. In addition, the protocol preserves mesenchymal cells in a manner that enhances image quality when compared to standard cryostat or paraffin sections, thereby increasing the efficacy and reliability of immunostaining. We believe that this protocol will benefit all laboratories that visualize skin, and possibly other tissues and organs.

This work presents a novel processing and imaging protocol for thick, three-dimensional tissue cross-section analysis that enables the full exploitation of confocal imaging modalities. This protocol preserves antigenicity and represents a robust system to analyze skin histology and potentially other tissue types.

Processing a tissue of interest to generate a microscopic image that supports a scientific argument can be challenging. The acquisition of high-quality ...

Immuno-fluorescent Labeling of Microtubules and Centrosomal Proteins in Ex Vivo Intestinal Tissue and 3D In Vitro Intestinal Organoids

The advent of 3D in vitro organoids that mimic the in vivo tissue architecture and morphogenesis has greatly advanced the ability to study key biological questions in cell and developmental biology. In addition, organoids together with recent technical advances in gene editing and viral gene delivery promises to advance medical research and development of new drugs for treatment of diseases. Organoids grown in vitro in basement matrix provide powerful model systems for studying the behavior and function of various proteins and are well suited for live-imaging of fluorescent-tagged proteins. However, establishing the expression and localization of the endogenous proteins in ex vivo tissue and in in vitro organoids is important to verify the behavior of the tagged proteins. To this end we have developed and modified tissue isolation, fixation, and immuno-labeling protocols for localization of microtubules, centrosomal, and associated proteins in ex vivo intestinal tissue and in in vitro intestinal organoids. The aim was for the fixative to preserve the 3D architecture of the organoids/tissue while also preserving antibody antigenicity and enabling good penetration and clearance of fixative and antibodies. Exposure to cold depolymerizes all but stable microtubules and this was a key factor when modifying the various protocols. We found that increasing the ethylenediaminetetraacetic acid (EDTA) concentration from 3 mM to 30 mM gave efficient detachment of villi and crypts in the small intestine while 3 mM EDTA was sufficient for colonic crypts. The developed formaldehyde/methanol fixation protocol gave very good structural preservation while also preserving antigenicity for effective labeling of microtubules, actin, and the end-binding (EB) proteins. It also worked for the centrosomal protein ninein although the methanol protocol worked more consistently. We further established that fixation and immuno-labeling of microtubules and associated proteins could be achieved with organoids isolated from or remaining within the basement matrix.

Immuno-fluorescent Labeling of Microtubules and Centrosomal Proteins in Ex Vivo Intestinal Tissue and 3D In Vitro Intestinal Organoids

We present protocols for isolation of intestinal 3D structures from in vivo tissue and in vitro basement matrix embedded organoids, and detail different fixation and staining protocols optimized for immuno-labeling of microtubule, centrosomal, and junctional proteins as well as cell markers including the stem cell protein Lgr5.

The advent of 3D in vitro organoids that mimic the in vivo tissue architecture and morphogenesis has greatly advanced the ability to study key biological ...

Visualization and Analysis of Pharyngeal Arch Arteries using Whole-mount Immunohistochemistry and 3D Reconstruction

Improper formation or remodeling of the pharyngeal arch arteries (PAAs) 3, 4, and 6 contribute to some of the most severe forms of congenital heart disease. To study the formation of PAAs, we developed a protocol using whole-mount immunofluorescence coupled with benzyl alcohol/benzyl benzoate (BABB) tissue clearing, and confocal microscopy. This allows for the visualization of the pharyngeal arch endothelium at a fine cellular resolution as well as the 3D connectivity of the vasculature. Using software, we have established a protocol to quantify the number of endothelial cells (ECs) in PAAs, as well as the number of ECs within the vascular plexus surrounding the PAAs within pharyngeal arches 3, 4, and 6. When applied to the whole embryo, this methodology provides a comprehensive visualization and quantitative analysis of embryonic vasculature.

Here, we describe a protocol to visualize and analyze the pharyngeal arch arteries 3, 4, and 6 of mouse embryos using whole-mount immunofluorescence, tissue clearing, confocal microscopy, and 3D reconstruction.

Improper formation or remodeling of the pharyngeal arch arteries (PAAs) 3, 4, and 6 contribute to some of the most severe forms of congenital heart ...

Preparation of Small RNA Libraries for Sequencing from Early Mouse Embryos

MicroRNAs (miRNAs) are important for the complex regulation of cell fate decisions and developmental timing. In vivo studies of the contribution of miRNAs during early development are technically challenging due to the limiting cell number. Moreover, many approaches require a miRNA of interest to be defined in assays such as northern blotting, microarray, and qPCR. Therefore, the expression of many miRNAs and their isoforms have not been studied during early development. Here, we demonstrate a protocol for small RNA sequencing of sorted cells from early mouse embryos to enable relatively unbiased profiling of miRNAs in early populations of neural crest cells. We overcome the challenges of low cell input and size selection during library preparation using amplification and gel-based purification. We identify embryonic age as a variable accounting for variation between replicates and stage-matched mouse embryos must be used to accurately profile miRNAs in biological replicates. Our results suggest that this method can be broadly applied to profile the expression of miRNAs from other lineages of cells. In summary, this protocol can be used to study how miRNAs regulate developmental programs in different cell lineages of the early mouse embryo.

We describe a technique for profiling microRNAs in early mouse embryos. This protocol overcomes the challenge of low cell input and small RNA enrichment. This assay can be used to analyze changes in miRNA expression over time in different cell lineages of the early mouse embryo.

MicroRNAs (miRNAs) are important for the complex regulation of cell fate decisions and developmental timing. In vivo studies of the contribution of miRNAs ...

Accurate Follicle Enumeration in Adult Mouse Ovaries

Sexually reproducing female mammals are born with their entire lifetime supply of oocytes. Immature, quiescent oocytes are found within primordial follicles, the storage unit of the female germline. They are non-renewable, thus their number at birth and subsequent rate of loss largely dictates the female fertile lifespan. Accurate quantification of primordial follicle numbers in women and animals is essential for determining the impact of medicines and toxicants on the ovarian reserve. It is also necessary for evaluating the need for, and success of, existing and emerging fertility preservation techniques. Currently, no methods exist to accurately measure the number of primordial follicles comprising the ovarian reserve in women. Furthermore, obtaining ovarian tissue from large animals or endangered species for experimentation is often not feasible. Thus, mice have become an essential model for such studies, and the ability to evaluate primordial follicle numbers in whole mouse ovaries is a critical tool. However, reports of absolute follicle numbers in mouse ovaries in the literature are highly variable, making it difficult to compare and/or replicate data. This is due to a number of factors including strain, age, treatment groups, as well as technical differences in the methods of counting employed. In this article, we provide a step-by-step instructional guide for preparing histological sections and counting primordial follicles in mouse ovaries using two different methods: [1] stereology, which relies on the fractionator/optical dissector technique; and [2] the direct count technique. Some of the key advantages and drawbacks of each method will be highlighted so that reproducibility can be improved in the field and to enable researchers to select the most appropriate method for their studies.

Here, we describe, compare, and contrast two different techniques for accurate follicle counting of fixed mouse ovarian tissues.

Sexually reproducing female mammals are born with their entire lifetime supply of oocytes. Immature, quiescent oocytes are found within primordial ...

Whole Ovary Immunofluorescence, Clearing, and Multiphoton Microscopy for Quantitative 3D Analysis of the Developing Ovarian Reserve in Mouse

Female fertility and reproductive lifespan depend on the quality and quantity of the ovarian oocyte reserve. An estimated 80% of female germ cells entering meiotic prophase I are eliminated during Fetal Oocyte Attrition (FOA) and the first week of postnatal life. Three major mechanisms regulate the number of oocytes that survive during development and establish the ovarian reserve in females entering puberty. In the first wave of oocyte loss, 30-50% of the oocytes are eliminated during early FOA, a phenomenon that is attributed to high Long interspersed nuclear element-1 (LINE-1) expression. The second wave of oocyte loss is the elimination of oocytes with meiotic defects by a meiotic quality checkpoint. The third wave of oocyte loss occurs perinatally during primordial follicle formation when some oocytes fail to form follicles. It remains unclear what regulates each of these three waves of oocyte loss and how they shape the ovarian reserve in either mice or humans.
Immunofluorescence and 3D visualization have opened a new avenue to image and analyze oocyte development in the context of the whole ovary rather than in less informative 2D sections. This article provides a comprehensive protocol for whole ovary immunostaining and optical clearing, yielding preparations for imaging using multiphoton microscopy and 3D modeling using commercially available software. It shows how this method can be used to show the dynamics of oocyte attrition during ovarian development in C57BL/6J mice and quantify oocyte loss during the three waves of oocyte elimination. This protocol can be applied to prenatal and early postnatal ovaries for oocyte visualization and quantification, as well as other quantitative approaches. Importantly, the protocol was strategically developed to accommodate high-throughput, reliable, and repeatable processing that can meet the needs in toxicology, clinical diagnostics, and genomic assays of ovarian function.

Here, we present an optimized protocol for imaging entire ovaries for quantitative and qualitative analyses using whole-mount immunostaining, multiphoton microscopy, and 3D visualization and analysis. This protocol accommodates high-throughput, reliable, and repeatable processing that is applicable for toxicology, clinical diagnostics, and genomic assays of ovarian function.