We thank for the laboratory platform and assistance of Ear Institute, Shanghai Jiaotong University School of Medicine. This work was supported by Shanghai Pujiang Program (22PJ1409200); National Natural Science Foundation of China (No.11932012); Postdoctoral Scientific Research Foundation of Shanghai Ninth People&#39;s Hospital, Shanghai Jiao Tong University School of Medicine;Fundamental research program funding of Ninth People&#39;s Hospital affiliated to Shanghai Jiao Tong University School of Medicine (JYZZ154).

All experimental procedures described here were approved by the Animal Care Committee of Shanghai Ninth People&#39;s Hospital, Shanghai Jiao Tong University School of Medicine (SH9H-2023-A616-SB). 4-week-old C57BL/6 male mice were used in this study. All the instruments used were sterilized prior to the procedure.
1. Preparation of the suture expansion model
<ol>
	<li>Preparation of two retention holders.
	<ol>
		<li>Use 0.014&#39;&#39; Australian wire or stainless steel wire (see Table of Materials) to make a helical loop with light wire pliers. The diameter of the loop is 2 mm, and 1 mm tail is reserved on two sides (Figure 1A).</li>
		<li>Sterilize the wires and holders for the surgery in an autoclave, plasma sterilizer, or suitable cold sterilant (e.g., glutaraldehyde).</li>
	</ol>
	</li>
	<li>Preparation of the springs.
	<ol>
		<li>Prepare customized stainless-steel springs. 
		NOTE: The pressure spring with 0.2 mm wire diameter, 1.5 mm external diameter, 1 mm interval space and 7 mm length is used in this study (Figure 1B). Each 1 mm spring compression obtained a thrust of about 30 g.</li>
		<li>Cut the spring into an available length before setting. Confirm the force for the specialized spring before the experiment. 
		NOTE: The sizes of springs are varied as long as the force magnitude is the same in parallel experiments. The force is measured by a tabletop uniaxial testing instrument or a small electronic dynamometer (see Table of Materials)&#160;if the condition is limited.&#160;Length change is evaluated to the force magnitude (Figure 1C-E). Notably, it is necessary to change the spring force loading value based on the age, the bone condition of the mouse, and the research objects.</li>
		<li>Prepare one straight Australian wire or straight steel wire of 7 mm length, and make two pieces of paper with 2 mm diameters to use as the barriers (Figure 1F).</li>
	</ol>
	</li>
</ol>
2. Sagittal suture expansion surgery
<ol>
	<li>Anesthetization: Anesthetize the mice with tiletamine (25 mg/kg), zolazepam (25 mg/kg) and Xylazine (10 mg/kg). At the same time, apply sterile ophthalmic ointment to the eyes to prevent drying of the corneas. Judge the depth of anesthesia of the mice via a toe pinch. 
	NOTE: The following characteristics indicate that the mouse is anesthetized properly: slow and steady breathing, weakness of limbs, muscle relaxation, disappearance of skin acupuncture reflex and eyelid reflex, as well as weaker corneal reflex.</li>
	<li>Fur removal and disinfection: Carefully remove the fur on the top of the head with hair removal cream; avoid touching the eyes. Soon afterward, use alternating rounds of iodophor and 75% alcohol to disinfect the surgical site. Administer meloxicam (1 mg/kg) as analgesia to&#160;the mice by subcutaneous injection.</li>
	<li>Body position fixation: Put the mouse lying on the front and use surgical tapes to fix the limbs on the operating table.</li>
	<li>Open scalp flap: Use surgical scissors to perform an arcuate flap along the scalp near the neck of the mouse, fully exposing the sagittal suture and surrounding skull. After that, fix the scalp flap with a 6-0 suture on the operating table.</li>
	<li>Bond the retention holders after acid etching.
	<ol>
		<li>Dry the skull and etch it with 37% phosphoric acid for 20 s, and use normal saline to clean up the acid etching (Figure 2A). A chalky residue will be evident after drying the skull with the rubber pipette bulb.</li>
		<li>Cement the two holders (prepared in step 1.1) on both sides of the parietal bones 3 mm from the sagittal sutures with a light-cured adhesive (Figure 2B).</li>
	</ol>
	</li>
	<li>Reset the scalp flap.
	<ol>
		<li>Manually put back the scalp flap (Figure 2C). Label the holders&#39; position, cut two small holes in the scalp at the corresponding position of the bilateral retention holders, and then reset the flapped scalp.</li>
		<li>At the same time, pass the small loops through the holes to expose the skin&#39;s surface. Suture the curved incision with a 6-0 suture&#160;(Figure 2D). One to three days are allowed for the skin recovery.&#160;Administer meloxicam (1 mg/kg) to&#160;the mice by subcutaneous injection every 24 h&#160;for 1 to 3 days. 
		NOTE: After surgery, daily check the activity of the mouse scalp and whether the holders fall off. There are various types of skin in the matter of color and thickness; the healthy skin scalp will maintain the original color, and skin edges will gradually fuse together after suturing. If an infection of the scalp is found, or if the surgical device has fallen off, remove the mouse from the study and euthanize it.</li>
	</ol>
	</li>
	<li>Install the spring and guide wire.
	<ol>
		<li>Cut the spring 1 mm longer than the distance between the two holders.</li>
		<li>Compress the selected pressure spring and place it between the small coils on both sides.</li>
		<li>Pass the stainless-steel wire through the small coils and the spring, and release the spring to obtain a starting thrust of about 30 g.</li>
		<li>Check that the scalp under the spring area of the mouse is without any compression.</li>
		<li>After confirming that the bonding is firm without any looseness, bond two scraps of paper between the spring and the holders with a light-cured adhesive in order to set up barriers at both ends of the spring (Figure 2E,F).</li>
	</ol>
	</li>
</ol>
3. Double labeling of mineralized bones
<ol>
	<li>Preparation of the stock solution.
	<ol>
		<li>Prepare a dilution solution with distilled water that contains 0.9% NaCl&#160;and 2% NaHCO3.</li>
		<li>Use the diluent solution to prepare 20 mg/mL of Alizarin complex dihydrate and 10 mg/mL of Calcein (see Table of Materials).</li>
		<li>Adjust the pH to 7.4 using a pH measuring device. Flush the pH probe between measurements to ensure accurate readings.</li>
		<li>Put the dye into a sterile container and store it in a refrigerator at 4 &#176;C; foil is used to keep light out.</li>
	</ol>
	</li>
	<li>Prepare the working solution. Before injection, dilute the stock solution to 1 mg/mL for calcium and 2 mg/mL for Alizarin complex dihydrate using a dissolution solution.</li>
	<li>Intraperitoneally inject 5 mg/kg of Calcein green and 20 mg/kg of Alizarin red at two time points to label the mineralized bones and analyze the changes between two times. Generally, collect the samples 12-14 h after injection. 
	NOTE: Warm the dyes before injection, and ensure that the injection time is not less than 3 minutes. The injection interval depends on the time of expansion. In this study, Alizarin red and Calcein green were injected overnight (O/N) before expansion and O/N before collection, respectively.</li>
	<li>Collect whole calvarial bones prepared for tissue clearing procedure a day after the second injection.</li>
</ol>
4. EdU staining
<ol>
	<li>Intraperitoneal injection: Intraperitoneally inject EdU 2 h before euthanizing the mice (following institutionally approved protocols). The dose is 1 mg/10 g body weight.</li>
	<li>Labeling cocktail preparation: Mix Tris-buffered saline (100 mmol/L final, pH 7.6), CuSO4 (4 mmol/L final), Sulfo-Cyanine 3 Azide (2-5 &#181;mol/L final) and Sodium Ascorbate (100 mmol/L final, made fresh each use) (see Table of Materials).</li>
	<li>EdU whole-mount staining: After the tissue decolorization step in the PEGASOS tissue clearing method (step 7), put samples in the labeling cocktail for 1 day at room temperature (RT).</li>
</ol>
5. Micro-computed tomography imaging
<ol>
	<li>Tissue fixation and storage: Fix the calvarial bones in 4% paraformaldehyde (PFA) at 4 &#176;C O/N and store them in 0.5% PFA before scanning.</li>
	<li>Scanning and analysis: Scan the samples with a micro-computed tomography (&#181;CT) imaging system with high resolution and a voxel size of 7 &#181;m.</li>
</ol>
6. Preparation of work solution for PEGASOS tissue clearing
<ol>
	<li>4% polyformaldehyde (PFA): Dissolve 4 g of PFA powder in 1&#215; PBS to 100 mL.</li>
	<li>Cardiac perfusion solution: 0.02% Heparin, that is, 20 mg heparin powder dissolved in 1&#215; PBS to 100 mL; alternatively, use 0.05 mol/L EDTA, that is, 0.05 mol of ethylene diamine tetraacetic acid (EDTA) (see Table of Materials), dissolved in a deionized aqueous solution to a total volume of 1 L.</li>
	<li>Decalcification solution: 0.5 mol/L EDTA, that is, 0.5 mol of ethylene diamine tetraacetic acid (EDTA), dissolved in a deionized aqueous solution to a total volume of 1 L.</li>
	<li>Decolorization solution: 25% Quadrol, which is 250 mL of Quadrol (N, N, N&#39;, N&#39; - Tetrakis (2-Hydroxypropyl) ethylenediamine) (see Table of Materials) dissolved in deionized water to a total volume of 1 L. 
	NOTE: Due to the viscosity of Quadrol, it can be heated to 60 &#176;C to increase its fluidity before configuration.</li>
	<li>Degreasing solution: 30%, 50%, 70% tert-Butanol (tB) (see Table of Materials), prepared with water by volume fraction. 
	NOTE: Considering that pure tB is crystalline at room temperature, it must be heated to 60 &#176;C until it dissolves before it can be used. Subsequently, 3% to 5% (v/v) triethanolamine (TEA) is added to regulate the pH of the solution &#62;9.5.</li>
	<li>Dehydration solution: TB-PEG solution, 70% tB + 27% PEG-MMA + 3% Quadrol (v/v), wherein PEG-MMA is poly (ethylene glycol) methyl ether methacrylate with an average molecular weight of 500 (Poly (ethylene glycol) Methacrylate 500, PEG-MMA 500) (see Table of Materials).</li>
	<li>Transparent solution: BB-PEG solution, 75% BB + 22% PEG-MMA + 3% Quadrol (v/v), wherein BB is benzyl benzoate (BB).</li>
</ol>
7. Transparency of calvarial bones with the PEGASOS method
<ol>
	<li>Anesthetize the mouse&#160;by intraperitoneal injection of anesthetics (step 2.1), and judge the depth of anesthesia of the mouse&#160;through the pinch reaction to ensure that there is no resistance reaction before cardiac perfusion.</li>
	<li>Perform cardiac perfusion and fixation.
	<ol>
		<li>Hold the abdomen of the mouse upward, fix the limbs with adhesive tape, and carefully cut along the circumference of the thorax to fully expose it.</li>
		<li>Immediately after making a small cut in the right atrium with ophthalmic scissors, insert a perfusion 22 G needle at the apex of the left ventricle. 
		NOTE: Only the needle tip is inserted to avoid damaging the heart valve through excessive insertion. Otherwise, cardiac perfusion fluid will enter the pulmonary circulation, affecting the perfusion effect of the systemic circulation.</li>
		<li>Push the 30-50 mL cardiac perfusion fluid (step 6.2) in the syringe. It can be seen that the liver gradually turns pale from bright red.</li>
		<li>After the outflow fluid from the right atrium becomes completely clear, infuse 4% PFA solution in equal volume. Operation of PFA perfusion is conducted in a ventilated hood.</li>
		<li>Dissect and separate the tissues and organs, and fix them overnight with 4% PFA at 4 &#176;C.</li>
	</ol>
	</li>
	<li>Tissue decalcification: For samples processed by EdU staining, place the fixed hard tissue in 0.5 mol/L EDTA (pH 7.0) solution (10 mL) in a shaking table at 37 &#176;C for about 2 days, and change the fluid daily. 
	NOTE: Skip the decalcification process in the normalized PEGASOS method for calcium chelating agent labeled bones to fully preserve the signaling. Decalcification is required to treat the bones to visualize the endogenous fluorescence or whole-mount immunostained signals.</li>
	<li>Tissue decolorization: Place the fixed calvarial bones in 25% Quadrol solution (20 mL)&#160;in a shaking table at 37 &#176;C for 1 day, and change the fluid once. 
	NOTE: The decolorization time is related to the heme content in the tissue. Observe the color of the treatment liquid, as the depth of color represents the need to continue the fluid replacement treatment.</li>
	<li>EdU staining: Rinse the samples in 1&#215; PBS for 5 min (three times), and then immerse them in the labeling cocktail for 1 day. After that, rinse the samples in 1&#215; PBS for 1 h (three times).</li>
	<li>Tissue degreasing and dehydration
	<ol>
		<li>Place the tissues in 30% tB, 50% tB, and 70% tB solutions in sequence to perform gradient decreasing on a shaking table at 37 &#176;C. Place in each concentration for about 2 h.</li>
		<li>Subsequently, dehydrate the samples in TB-PEG solution for 6 h on a shaking table at 37 &#176;C. 
		NOTE: The above processing time can be increased or decreased depending on the number of the tissue.</li>
	</ol>
	</li>
	<li>Tissue transparency: Place the completely dehydrated tissue in BB-PEG solution on a shaking table at 37 &#176;C (2-4 h) until transparent. Currently, samples can be stored for up to 1-2 years. 
	&#8203;NOTE: After almost fully clearing, open the tube lid to expose the samples and medium to air with shaking will further improve the transparency. This method is suitable for improving the transparency of individual tissues and organs, as well as the entire head and body of young mice. However, for the transparency of the whole body of adult mice, the above steps should be performed using a full-cycle perfusion method.</li>
</ol>
8. Imaging
NOTE: Confocal microscopy was used for 3-D visualization of transparent tissues in this study. Light-sheet microscopy is also appropriate for this protocol. Several operating systems have been verified as available before. Here, a laser confocal microscope operating system is taken as an example (see Table of Materials).
<ol>
	<li>According to the user manual, follow the correct steps to open the laser confocal and LAS AF operation interface. Turn on the required laser in different shooting channels, and adjust the power. Set the optical path and the corresponding wavelength, 561 nm for Alizarin red and 488 nm for Calcein green, any of which is used for the EdU signal according to the labeling cocktail.</li>
	<li>Place the transparent samples in the concave glass Petri dish that can carry thick specimens, embedding boxes, or self-made placement devices such as waterproof glue to immerse the transparent samples in a transparent liquid.</li>
	<li>Select a low-power objective lens to quickly locate the sample. Make an overview of the whole calvarial tissue with the fast-scanning function, and find the interest area for imaging.</li>
	<li>Set output power, select the scanning mode, and first adjust the shooting coefficients (resolution, scanning speed, zoom factor, image quality, average background noise, etc.). 
	NOTE: Generally, Smart Gain ranges from 500 to 800 (both signal and noise change); Smart Offset is as close to 0 as possible while ensuring image quality (to reduce background noise). When the PMT gain is higher than 800, or HyD gain is greater than 100%, and the brightness is still insufficient, the laser intensity can be appropriately increased, but in principle, the lower, the better.</li>
	<li>Set the axial distance Z range and Z step, which is drilled down from the shallowest side of the transparent organization; that is, set the deepest and shallowest sides. 
	NOTE: Confirm the classification and the working distance for each lens. Carefully set the Z range, and do not operate the working distance exceeding the limit of the selected lens. The &#34;z-Galvo&#34; can be selected when fine regulation is needed.</li>
	<li>Set the shooting parameters again, and start shooting. After completion, merge and process the images through the multi-functional module. Finally, store and shut down the instrument in the order indicated.</li>
</ol>

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

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The authors have nothing to disclose.

We applied a standard suture expansion mouse model to observe the regular morphological changes that occur every week during the entire month-long remodeling cycle10. This model is useful for researching calvarial bone remodeling and regeneration by expanding calvarial sutures, as well as for studying various suture cells in vivo. To fully present the results of such research, three-dimensional visualization of stained tissues is needed. Therefore, PEGASOS technology, known for its effici...

Craniofacial sutures are fibrous tissues that connect craniofacial bones and play essential roles in the growth and remodeling of craniofacial bones. The structure of the suture resembles a river, providing a flow of cell resources to nourish and build the &#34;river bank&#34;, known as the osteogenic fronts, which contribute to the formation of craniofacial bones via intramembranous osteogenesis1.
Interest in craniofacial sutures has been driven by clinical needs to understand premature closure of cranial sutures and facial suture dysfunction, which may lead to craniofacial deformities and even life-threatening conditions in children. Open suturectomy is routinely used in clinical treatment, but long-term follow-up has shown incomplete re-ossification recurrence in some patients2. Minimally invasive craniotomy assisted by expansion springs or endoscopic stripe craniectomy may provide a safer approach to preserving the potential suture rather than discarding the tissues3. Similarly, orthopedic therapies such as facemasks and expansion appliances have been widely used to treat sagittal or horizontal maxillary hypoplasia, with some studies extending the age limitation to treat adult patients via miniscrew-assisted palatal expanders4,5,6. Additionally, cranial suture regeneration with mesenchymal stem cells (MSCs) combined with biodegradable materials is a potential therapy in the future, offering a novel direction for the treatment of related diseases7. However, the function process or regulatory mechanism of sutures remains elusive.
Bone remodeling mainly consists of a balance between bone formation conducted by osteoblasts and bone resorption conducted by osteoclasts, where osteogenic differentiation of stem cells stimulated by mechanical signals plays an important role. After decades of research, it has been found that craniofacial sutures are highly plastic mesenchymal stem cell niches8. Suture stem cells (SuSCs) are a heterogeneous group of stem cells, belonging to mesenchymal stem cells (MSCs) or bone stem cells (SSCs). SuSCs are labeled in vivo by four markers, including Gli1, Axin2, Prrx1, and Ctsk. Gli1+ SuSCs, in particular, have strictly verified the biological characteristics of stem cells, not only exhibiting high expression of typical MSC markers but also demonstrating excellent osteogenic and chondrogenic potential9. Previous research has shown that Gli1+ SuSCs actively contribute to new bone formation under tensile force, identifying them as the suture stem cell source supporting distraction osteogenesis10.
In the past, extensive mechanical characteristics of stem cells were studied in vitro via Flexcell, four-point bending, micro-magnet loading system, and others. Although mouse cranial suture-derived mesenchymal cells have been identified in vitro11, and human suture mesenchymal stem cells have also been isolated recently12, the biomechanical response of suture cells remains unclear in the in vitro system. To further investigate the bone remodeling process, a suture expansion model based on isolated calvaria organ culture has been established, paving the way for establishing a useful in vivo suture expansion model1,13. Rabbits14 and rats15 have been the most widely used animals in basic research for suture expansion. However, mice are preferred animal models for exploring human disease due to their highly homologous genome with humans, numerous gene modification lines, and strong reproductive hybridization ability. Existing mouse models of cranial suture expansion typically rely on stainless steel orthodontic spring wires to apply tensile force to the sagittal suture16,17. In these models, two holes are made in each side of the parietal bones to fix the expansion device, and the wires are embedded under the skin, which may affect the cell activation mode.
Regarding the visualization method, the two-dimensional observation of slices in the sagittal direction has been generally adopted for decades. However, considering that bone remodeling is a complex three-dimensional dynamic process, obtaining complete three-dimensional information has become an urgent need. The PEGASOS tissue transparency technique emerged to meet this requirement18,19. It offers unique advantages for the transparency of hard and soft tissues, enabling the complete bone remodeling process to be reproduced in three-dimensional space.
To gain a deeper and more comprehensive understanding of the physiological changes in the bone remodeling periods, a standard sagittal suture expansion mouse model with a spring setting between the handmade holders was established10. With a standardized acid etching and bonding procedure, the expansion device could be firmly bonded to the cranial bone, generating a tensile force perpendicular to the sagittal suture. Furthermore, the PEGASOS tissue clearing method was applied after double labeling of the mineralized bone post-expansion to fully visualize the bone modeling changes after suture expansion.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>37% Acid etching</td>
			<td>Xihubiom</td>
			<td>E10-02/1807011</td>
			<td></td>
		</tr>
		<tr>
			<td>Alizarin red</td>
			<td>Sigma-Aldrich</td>
			<td>A3882</td>
			<td></td>
		</tr>
		<tr>
			<td>AUSTRALIAN WIRE</td>
			<td>A.J.WILCOCK</td>
			<td>0.014&#39;&#39;</td>
			<td></td>
		</tr>
		<tr>
			<td>Benzyl benzoate</td>
			<td>Sigma-Aldrich</td>
			<td>B6630</td>
			<td></td>
		</tr>
		<tr>
			<td>Calcein green</td>
			<td>Sigma-Aldrich</td>
			<td>C0875</td>
			<td></td>
		</tr>
		<tr>
			<td>Copper(II) sulfate, anhydrous</td>
			<td>Sangon Biotech</td>
			<td>A603008</td>
			<td></td>
		</tr>
		<tr>
			<td>Dynamometer</td>
			<td>Sanliang</td>
			<td>SF-10N</td>
			<td></td>
		</tr>
		<tr>
			<td>EDTA</td>
			<td>Sigma-Aldrich</td>
			<td>E9884</td>
			<td></td>
		</tr>
		<tr>
			<td>EdU</td>
			<td>Invitrogen</td>
			<td>E104152</td>
			<td></td>
		</tr>
		<tr>
			<td>Laser Confocal Microscope</td>
			<td>Leica</td>
			<td>SP8</td>
			<td></td>
		</tr>
		<tr>
			<td>PBS</td>
			<td>Sangon Biotech</td>
			<td>E607008</td>
			<td></td>
		</tr>
		<tr>
			<td>PEG-MMA 500</td>
			<td>Sigma-Aldrich</td>
			<td>447943</td>
			<td></td>
		</tr>
		<tr>
			<td>PFA</td>
			<td>Sigma-Aldrich</td>
			<td>P6148&#160;</td>
			<td></td>
		</tr>
		<tr>
			<td>pH Meters</td>
			<td>Mettler Toledo</td>
			<td>S220</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>A4034</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium bicarbonate</td>
			<td>Sangon Biotech</td>
			<td>A500873</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium chloride</td>
			<td>Sangon Biotech</td>
			<td>A610476</td>
			<td></td>
		</tr>
		<tr>
			<td>Sodium hydroxide</td>
			<td>Sigma-Aldrich</td>
			<td>S5881</td>
			<td></td>
		</tr>
		<tr>
			<td>Spring</td>
			<td>TAOBAO</td>
			<td>0.2*1.5*1*7</td>
			<td></td>
		</tr>
		<tr>
			<td>Sulfo-Cyanine3 azide</td>
			<td>Lumiprobe</td>
			<td>A1330</td>
			<td></td>
		</tr>
		<tr>
			<td>tert-Butanol</td>
			<td>Sigma-Aldrich</td>
			<td>360538&#160;</td>
			<td>Protect from light. Do not freeze.</td>
		</tr>
		<tr>
			<td>Transbond MIP 
			Moisture Insensitive Primer</td>
			<td>3M Unitek</td>
			<td>712-025</td>
			<td></td>
		</tr>
		<tr>
			<td>Transbond XT 
			Light Cure Adhesive Paste</td>
			<td>3M Unitek</td>
			<td>712-035</td>
			<td></td>
		</tr>
		<tr>
			<td>Triethanolamine</td>
			<td>Sigma-Aldrich</td>
			<td>V900257</td>
			<td></td>
		</tr>
		<tr>
			<td>Tris-buffered saline</td>
			<td>Sangon Biotech</td>
			<td>A500027</td>
			<td></td>
		</tr>
	</tbody>
</table>

a 3-d visualization technique for bone remodeling in a suture expansion mouse model

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

Using this protocol, a mouse model for sagittal suture expansion has been established (Figure 1-2). For 3-D visualization of bone modeling changes after suture expansion, the PEGASOS tissue clearing method was applied to the entire calvarial bones following expansion. After perfusion, calvarial bones were separated (Figure 3A), and the appropriate PEGASOS process was continued (Table 1 and Table 2</stron...

Watch this Scientific Journal Video about PEGASOS Tissue Clearing Technique to Visualize Bone Remodeling at JoVE.com

Author Spotlight: PEGASOS Tissue Clearing Technique to Visualize Bone Remodeling 

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

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

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

a-3-d-visualization-technique-for-bone-remodeling-suture-expansion

Research

JoVE Journal

Biology

1.3K Views.  Shanghai Jiao Tong University School of Medicine. This protocol presents a standardized suture expansion mouse model and a 3-D visualization method to study the mechanobiological changes of the suture and bone remodeling under tensile force loading.

Video: Author Spotlight: PEGASOS Tissue Clearing Technique to Visualize Bone Remodeling 

A Rapid Technique for the Visualization of Live Immobilized Yeast Cells

We present here a simple, rapid, and extremely flexible technique for the immobilization and visualization of growing yeast cells by epifluorescence microscopy. The technique is equally suited for visualization of static yeast populations, or time courses experiments up to ten hours in length. My microscopy investigates epigenetic inheritance at the silent mating loci in S. cerevisiae. There are two silent mating loci, HML and HMR, which are normally not expressed as they are packaged in heterochromatin. In the sir1 mutant background silencing is weakened such that each locus can either be in the expressed or silenced epigenetic state, so in the population as a whole there is a mix of cells of different epigenetic states for both HML and HMR. My microscopy demonstrated that there is no relationship between the epigenetic state of HML and HMR in an individual cell. sir1 cells stochastically switch epigenetic states, establishing silencing at a previously expressed locus or expressing a previously silenced locus. My time course microscopy tracked individual sir1 cells and their offspring to score the frequency of each of the four possible epigenetic switches, and thus the stability of each of the epigenetic states in sir1 cells. See also  Xu et al., Mol. Cell 2006.

A rapid technique for the visualization of growing immobilized yeast cells, here applied to fluorescent reporters at the silent mating loci HML and HMR

Szybkie Technika Wizualizacja żywo unieruchomionych komórek drożdży

We present here a simple, rapid, and extremely flexible technique for the immobilization and visualization of growing yeast cells by epifluorescence ...

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Human cancer and response to therapy is better represented in orthotopic animal models. This paper describes the development of an orthotopic mouse model of ovarian cancer, treatment of cancer via oral delivery of drugs, and monitoring of tumor cell behavior in response to drug treatment in real time using in vivo imaging system. In this orthotopic model, ovarian tumor cells expressing luciferase are applied topically by injecting them directly into the mouse bursa where each ovary is enclosed. Upon injection of D-luciferin, a substrate of firefly luciferase, luciferase-expressing cells generate bioluminescence signals. This signal is detected by the in vivo imaging system and allows for a non-invasive means of monitoring tumor growth, distribution, and regression in individual animals. Drug administration via oral gavage allows for a maximum dosing volume of 10 mL/kg body weight to be delivered directly to the stomach and closely resembles delivery of drugs in clinical treatments. Therefore, techniques described here, development of an orthotopic mouse model of ovarian cancer, oral delivery of drugs, and in vivo imaging, are useful for better understanding of human ovarian cancer and treatment and will improve targeting this disease.

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Orthotopic animal models of ovarian cancer replicate better human disease and therefore enhance our understanding of cancer progression and tumor response to therapy. A mouse model receives an intrabursal injection of luciferase-expressing ovarian tumor cells. Treatment is administered via oral gavage. Tumor growth is monitored by in vivo imaging system.

Human cancer and response to therapy is better represented in orthotopic animal models.  This paper describes the development of an orthotopic mouse model ...

Visualization of Endoplasmic Reticulum Localized mRNAs in Mammalian Cells

In eukaryotes, most of the messenger RNAs (mRNAs) that encode secreted and membrane proteins are localized to the surface of the endoplasmic reticulum (ER). However, the visualization of these mRNAs can be challenging. This is especially true when only a fraction of the mRNA is ER-associated and their distribution to this organelle is obstructed by non-targeted (i.e. "free") transcripts. In order to monitor ER-associated mRNAs, we have developed a method in which cells are treated with a short exposure to a digitonin extraction solution that selectively permeabilizes the plasma membrane, and thus removes the cytoplasmic contents, while simultaneously maintaining the integrity of the ER. When this method is coupled with fluorescent in situ hybridization (FISH), one can clearly visualize ER-bound mRNAs by fluorescent microscopy. Using this protocol the degree of ER-association for either bulk poly(A) transcripts or specific mRNAs can be assessed and even quantified. In the process, one can use this assay to investigate the nature of mRNA-ER interactions.

Here we describe a method to visualize endoplasmic reticulum-associated mRNAs in mammalian tissue culture cells. This technique involves the selective permeabilization of the plasma membrane with digitonin to remove cytoplasmic contents followed by fluorescent in situ hybridization to detect either bulk poly(A) mRNA or specific transcripts.

In eukaryotes, most of the messenger RNAs (mRNAs) that encode secreted and membrane proteins are localized to the surface of the endoplasmic reticulum ...

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the function and regulation of protein folding was studied in vitro, in isolated tissue culture cells and in unicellular organisms. Recent studies have uncovered links between protein homeostasis (proteostasis), metabolism, development, aging, and temperature-sensing. These findings have led to the development of new tools for monitoring protein folding in the model metazoan organism Caenorhabditis elegans. In our laboratory, we combine behavioral assays, imaging and biochemical approaches using temperature-sensitive or naturally occurring metastable proteins as sensors of the folding environment to monitor protein misfolding. Behavioral assays that are associated with the misfolding of a specific protein provide a simple and powerful readout for protein folding, allowing for the fast screening of genes and conditions that modulate folding. Likewise, such misfolding can be associated with protein mislocalization in the cell. Monitoring protein localization can, therefore, highlight changes in cellular folding capacity occurring in different tissues, at various stages of development and in the face of changing conditions. Finally, using biochemical tools ex vivo, we can directly monitor protein stability and conformation. Thus, by combining behavioral assays, imaging and biochemical techniques, we are able to monitor protein misfolding at the resolution of the organism, the cell, and the protein, respectively.

Using Caenorhabditis elegans as a Model System to Study Protein Homeostasis in a Multicellular Organism

To study the relationship between protein homeostasis, stress and aging, we monitored changes in protein folding by following protein dysfunction, protein localization in the cell and protein stability at the organismal, cellular and protein levels, using the genetically tractable metazoan Caenorhabditis elegans as a model system.

The folding and assembly of proteins is essential for protein function, the long-term health of the cell, and longevity of the organism. Historically, the ...

Osteoclast Derivation from Mouse Bone Marrow

Osteoclasts are highly specialized cells that are derived from the monocyte/macrophage lineage of the bone marrow. Their unique ability to resorb both the organic and inorganic matrices of bone means that they play a key role in regulating skeletal remodeling. Together, osteoblasts and osteoclasts are responsible for the dynamic coupling process that involves both bone resorption and bone formation acting together to maintain the normal skeleton during health and disease.
As the principal bone-resorbing cell in the body, changes in osteoclast differentiation or function can result in profound effects in the body. Diseases associated with altered osteoclast function can range in severity from lethal neonatal disease due to failure to form a marrow space for hematopoiesis, to more commonly observed pathologies such as osteoporosis, in which excessive osteoclastic bone resorption predisposes to fracture formation.
An ability to isolate osteoclasts in high numbers in vitro has allowed for significant advances in the understanding of the bone remodeling cycle and has paved the way for the discovery of novel therapeutic strategies that combat these diseases. 
Here, we describe a protocol to isolate and cultivate osteoclasts from mouse bone marrow that will yield large numbers of osteoclasts.

Osteoclasts are the principal bone-resorbing cell in the body. An ability to isolate osteoclasts in large numbers has resulted in significant advances in the understanding of osteoclast biology. In this protocol, we describe a method for isolation, cultivating and quantifying osteoclast activity in vitro.

Osteoclasts are highly specialized cells that are derived from the monocyte/macrophage lineage of the bone marrow. Their unique ability to resorb both the ...

Validation of a Mouse Model to Disrupt LINC Complexes in a Cell-specific Manner

Nuclear migration and anchorage within developing and adult tissues relies heavily upon large macromolecular protein assemblies called LInkers of the Nucleoskeleton and Cytoskeleton (LINC complexes). These protein scaffolds span the nuclear envelope and connect the interior of the nucleus to components of the surrounding cytoplasmic cytoskeleton. LINC complexes consist of two evolutionary-conserved protein families, Sun proteins and Nesprins that harbor C-terminal molecular signature motifs called the SUN and KASH domains, respectively. Sun proteins are transmembrane proteins of the inner nuclear membrane whose N-terminal nucleoplasmic domain interacts with the nuclear lamina while their C-terminal SUN domains protrudes into the perinuclear space and interacts with the KASH domain of Nesprins. Canonical Nesprin isoforms have a variable sized N-terminus that projects into the cytoplasm and interacts with components of the cytoskeleton. This protocol describes the validation of a dominant-negative transgenic mouse strategy that disrupts endogenous SUN/KASH interactions in a cell-type specific manner. Our approach is based on the Cre/Lox system that bypasses many drawbacks such as perinatal lethality and cell nonautonomous phenotypes that are associated with germline models of LINC complex inactivation. For this reason, this model provides a useful tool to understand the role of LINC complexes during development and homeostasis in a wide array of tissues.

Nuclear envelope proteins play a central role in many basic biological processes and have been implicated in a variety of human diseases. This protocol describes a new Cre/Lox-based mouse model that allows for the spatiotemporal control of LINC complexes disruption.

Nuclear migration and anchorage within developing and adult tissues relies heavily upon large macromolecular protein assemblies called LInkers of the ...

Nephrotoxin Microinjection in Zebrafish to Model Acute Kidney Injury

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid decline in renal function and development of the clinical syndrome known as acute kidney injury (AKI). Pharmacological agents used to treat medical circumstances ranging from bacterial infection to cancer, when administered individually or in combination with other drugs, can initiate AKI. Zebrafish are a useful animal model to study the chemical effects on renal function in vivo, as they form an embryonic kidney comprised of nephron functional units that are conserved with higher vertebrates, including humans. Further, zebrafish can be utilized to perform genetic and chemical screens, which provide opportunities to elucidate the cellular and molecular facets of AKI and develop therapeutic strategies such as the identification of nephroprotective molecules. Here, we demonstrate how microinjection into the zebrafish embryo can be utilized as a paradigm for nephrotoxin studies.

Renal injuries incurred from nephrotoxins, which include drugs ranging from antibiotics to chemotherapeutics, can result in complex disorders whose pathogenesis remains incompletely understood. This protocol demonstrates how zebrafish can be used for disease modeling of these conditions, which can be applied to the identification of renoprotective measures.

The kidneys are susceptible to harm from exposure to chemicals they filter from the bloodstream. This can lead to organ injury associated with a rapid ...

An Ex Vivo Tissue Culture Model of Cartilage Remodeling in Bovine Knee Explants

Ex vivo culture systems cover a broad range of experiments dedicated to studying tissue and cellular function in a native setting. Cartilage is a unique tissue important for proper function of the synovial joint and is constituted by a dense extracellular matrix (ECM), rich in proteoglycan and type II collagen. Chondrocytes are the only cell type present within cartilage and are widespread and relatively low in number. Altered external stimuli and cellular signalling can lead to changes in ECM composition and deterioration, which are important pathological hallmarks in diseases such as osteoarthritis (OA) and rheumatoid arthritis.
Ex vivo cartilage models allow 1) profiling of chondrocyte mediated alterations of cartilage tissue turnover, 2) visualizing the cartilage ECM composition, and 3) chondrocyte rearrangement directly in the tissue. Profiling these alterations in response to stimuli or treatments are of high importance in various aspects of cartilage biology, and complement in vitro experiments in isolated chondrocytes, or more complex models in live animals where experimental conditions are more difficult to control.
Cartilage explants present a translational and easily accessible method for assessing tissue remodeling in the cartilage ECM in controllable settings. Here, we describe a protocol for isolating and culturing live bovine cartilage explants. The method uses tissue from the bovine knee, which is easily accessible from the local butchery. Both explants and conditioned culture medium can be analyzed to investigate tissue turnover, ECM composition, and chondrocyte function, thus profiling ECM modulation.

Here, we present a protocol describing isolation and culturing of cartilage explants from bovine knees. This method provides an easy and accessible tool to describe tissue changes in response to biological stimuli or novel therapeutics targeting the joint.

Ex vivo culture systems cover a broad range of experiments dedicated to studying tissue and cellular function in a native setting. Cartilage is a unique ...

Mass Spectrometry Analysis to Identify Ubiquitylation of EYFP-tagged CENP-A (EYFP-CENP-A)

Studying the structure and the dynamics of kinetochores and centromeres is important in understanding chromosomal instability (CIN) and cancer progression. How the chromosomal location and function of a centromere (i.e., centromere identity) are determined and participate in accurate chromosome segregation is a fundamental question. CENP-A is proposed to be the non-DNA indicator (epigenetic mark) of centromere identity, and CENP-A ubiquitylation is required for CENP-A deposition at the centromere, inherited through dimerization between cell division, and indispensable to cell viability.
Here we describe mass spectrometry analysis to identify ubiquitylation of EYFP-CENP-A K124R mutant suggesting that ubiquitylation at a different lysine is induced because of the EYFP tagging in the CENP-A K124R mutant protein. Lysine 306 (K306) ubiquitylation in EYFP-CENP-A K124R was successfully identified, which corresponds to lysine 56 (K56) in CENP-A through mass spectrometry analysis. A caveat is discussed in the use of GFP/EYFP or the tagging of high molecular weight protein as a tool to analyze the function of a protein. Current technical limit is also discussed for the detection of ubiquitylated bands, identification of site-specific ubiquitylation(s), and visualization of ubiquitylation in living cells or a specific single cell during the whole cell cycle.
The method of mass spectrometry analysis presented here can be applied to human CENP-A protein with different tags and other centromere-kinetochore proteins. These combinatory methods consisting of several assays/analyses could be recommended for researchers who are interested in identifying functional roles of ubiquitylation.

Mass Spectrometry Analysis to Identify Ubiquitylation of EYFP-tagged CENP-A EYFP-CENP-A

CENP-A ubiquitylation is an important requirement for CENP-A deposition at the centromere, inherited through dimerization between cell division, and indispensable to cell viability. Here we describe mass spectrometry analysis to identify ubiquitylation of EYFP-tagged CENP-A (EYFP-CENP-A) protein.

Studying the structure and the dynamics of kinetochores and centromeres is important in understanding chromosomal instability (CIN) and cancer ...

Ex Vivo Expansion and Genetic Manipulation of Mouse Hematopoietic Stem Cells in Polyvinyl Alcohol-Based Cultures

Self-renewing&#160;multipotent&#160;hematopoietic stem cells (HSCs) are an important cell type due to their abilities to support hematopoiesis throughout life and reconstitute the entire blood system following transplantation. HSCs are used clinically in stem cell transplantation therapies, which represent curative treatment for a range of blood diseases. There is substantial interest in both understanding the mechanisms that regulate HSC activity and hematopoiesis, and developing new HSC-based therapies. However, the stable culture and expansion of HSCs ex vivo has been a major barrier in studying these stem cells in a tractable ex vivo system. We recently developed a polyvinyl alcohol-based culture system that can support the long-term and large-scale expansion of transplantable mouse HSCs and methods to genetically edit them. This protocol describes methods to culture and genetically manipulate mouse HSCs via electroporation and lentiviral transduction. This protocol is expected to be useful to a wide range of experimental hematologists interested in HSC biology and hematopoiesis.

Ex Vivo Expansion and Genetic Manipulation of Mouse Hematopoietic Stem Cells in Polyvinyl Alcohol-Based Cultures

Presented here is a protocol to initiate, maintain, and analyze mouse hematopoietic stem cell cultures using ex vivo polyvinyl alcohol-based expansion, as well as methods to genetically manipulate them by lentiviral transduction and electroporation.

Self-renewing&#160;multipotent&#160;hematopoietic stem cells (HSCs) are an important cell type due to their abilities to support hematopoiesis throughout ...