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><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&#039;&#039;</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&amp;nbsp;</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&amp;nbsp;</td><td>Protect from light. Do not freeze.</td></tr><tr><td>Transbond MIP&lt;br/&gt; Moisture Insensitive Primer</td><td>3M Unitek</td><td>712-025</td><td></td></tr><tr><td>Transbond XT&lt;br/&gt; 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 ...

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Research

JoVE Journal

Biology

1.4K 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 

Sequential In vivo Imaging of Osteogenic Stem/Progenitor Cells During Fracture Repair

Bone turns over continuously and is highly regenerative following injury. Osteogenic stem/progenitor cells have long been hypothesized to exist, but in vivo demonstration of such cells has only recently been attained. Here, in vivo imaging techniques to investigate the role of endogenous osteogenic stem/progenitor cells (OSPCs) and their progeny in bone repair are provided. Using osteo-lineage cell tracing models and intravital imaging of induced microfractures in calvarial bone, OSPCs can be directly observed during the first few days after injury, in which critical events in the early repair process occur. Injury sites can be sequentially imaged revealing that OSPCs relocate to the injury, increase in number and differentiate into bone forming osteoblasts. These methods offer a means of investigating the role of stem cell-intrinsic and extrinsic molecular regulators for bone regeneration and repair.

Sequential In vivo Imaging of Osteogenic Stem/Progenitor Cells During Fracture Repair

Quantitative measurement of bone progenitor function in fracture healing requires high resolution serial imaging technology. Here, protocols are provided for using intravital microscopy and osteo-lineage tracking to sequentially image and quantify the migration, proliferation and differentiation of endogenous osteogenic stem/progenitor cells in the process of repairing bone fracture.

Bone turns over continuously and is highly regenerative following injury. Osteogenic stem/progenitor cells have long been hypothesized to exist, but in ...

Medicine

Computed Tomography and Optical Imaging of Osteogenesis-angiogenesis Coupling to Assess Integration of Cranial Bone Autografts and Allografts

A major parameter determining the success of a bone-grafting procedure is vascularization of the area surrounding the graft. We hypothesized that implantation of a bone autograft would induce greater bone regeneration by abundant blood vessel formation. To investigate the effect of the graft on neovascularization at the defect site, we developed a micro&#8211;computed tomography (&#181;CT) approach to characterize newly forming blood vessels, which involves systemic perfusion of the animal with a polymerizing contrast agent. This method enables detailed vascular analysis of an organ in its entirety. Additionally, blood perfusion was assessed using fluorescence imaging (FLI) of a blood-borne fluorescent agent. Bone formation was quantified by FLI using a hydroxyapatite-targeted probe and &#181;CT analysis. Stem cell recruitment was monitored by bioluminescence imaging (BLI) of transgenic mice that express luciferase under the control of the osteocalcin promoter. Here we describe and demonstrate preparation of the allograft, calvarial defect surgery, &#181;CT scanning protocols for the neovascularization study and bone formation analysis (including the in vivo perfusion of contrast agent), and the protocol for data analysis.
The 3D high-resolution analysis of vasculature demonstrated significantly greater angiogenesis in animals with implanted autografts, especially with respect to arteriole formation. Accordingly, blood perfusion was significantly higher in the autograft group by the 7th day after surgery. We observed superior bone mineralization and measured greater bone formation in animals that received autografts. Autograft implantation induced resident stem cell recruitment to the graft-host bone suture, where the cells differentiated into bone-forming cells between the 7th and 10th postoperative day. This finding means that enhanced bone formation may be attributed to the augmented vascular feeding that characterizes autograft implantation. The methods depicted may serve as an optimal tool to study bone regeneration in terms of tightly bounded bone formation and neovascularization.

Implantation of autologous and allogeneic bone grafts constitute accepted approaches to treat major craniofacial bone loss. Yet the effect of graft composition on the interplay among neovascularization, cell differentiation and bone formation is unclear. We present a multimodal imaging protocol aimed to elucidate the angiogenesis-osteogenesis interdependence at the graft proximity.

A major parameter determining the success of a bone-grafting procedure is vascularization of the area surrounding the graft. We hypothesized that ...

Bioengineering

Combined In vivo Optical and &#181;CT Imaging to Monitor Infection, Inflammation, and Bone Anatomy in an Orthopaedic Implant Infection in Mice

Multimodality imaging has emerged as a common technological approach used in both preclinical and clinical research. Advanced techniques that combine in vivo optical and &#956;CT imaging allow the visualization of biological phenomena in an anatomical context. These imaging modalities may be especially useful to study conditions that impact bone. In particular, orthopaedic implant infections are an important problem in clinical orthopaedic surgery. These infections are difficult to treat because bacterial biofilms form on the foreign surgically implanted materials, leading to persistent inflammation, osteomyelitis and eventual osteolysis of the bone surrounding the implant, which ultimately results in implant loosening and failure. Here, a mouse model of an infected orthopaedic prosthetic implant was used that involved the surgical placement of a Kirschner-wire implant into an intramedullary canal in the femur in such a way that the end of the implant extended into the knee joint. In this model, LysEGFP mice, a mouse strain that has EGFP-fluorescent neutrophils, were employed in conjunction with a bioluminescent Staphylococcus aureus strain, which naturally emits light. The bacteria were inoculated into the knee joints of the mice prior to closing the surgical site. In vivo bioluminescent and fluorescent imaging was used to quantify the bacterial burden and neutrophil inflammatory response, respectively. In addition, &#956;CT imaging was performed on the same mice so that the 3D location of the bioluminescent and fluorescent optical signals could be co-registered with the anatomical &#956;CT images. To quantify the changes in the bone over time, the outer bone volume of the distal femurs were measured at specific time points using a semi-automated contour based segmentation process. Taken together, the combination of in vivo bioluminescent/fluorescent imaging with &#956;CT imaging may be especially useful for the noninvasive monitoring of the infection, inflammatory response and anatomical changes in bone over time.

Combined In vivo Optical and  &#181;CT Imaging to Monitor Infection, Inflammation, and Bone Anatomy in an Orthopaedic Implant Infection in Mice

Combined optical and &#956;CT imaging in a mouse model of orthopaedic implant infection, utilizing a bioluminescent engineered strain of Staphylococcus aureus, provided the capability to noninvasively and longitudinally monitor the dynamics of the bacterial infection, as well as the corresponding inflammatory response and anatomical changes in the bone.

Multimodality imaging has emerged as a common technological approach used in both preclinical and clinical research. Advanced techniques that combine in ...

Adult Mouse Digit Amputation and Regeneration: A Simple Model to Investigate Mammalian Blastema Formation and Intramembranous Ossification

Here, we present a protocol of adult mouse distal terminal phalanx (P3) amputation, a procedurally simple and reproducible mammalian model of epimorphic regeneration, which involves blastema formation and intramembranous ossification analyzed by fluorescence immunohistochemistry and sequential in-vivo microcomputed tomography (&#956;CT). Mammalian regeneration is restricted to amputations transecting the distal region of the terminal phalanx (P3); digits amputated at more proximal levels fail to regenerate and undergo fibrotic healing and scar formation. The regeneration response is mediated by the formation of a proliferative blastema, followed by bone regeneration via intramembranous ossification to restore the amputated skeletal length. P3 amputation is a preclinical model to investigate epimorphic regeneration in mammals, and is a powerful tool for the design of therapeutic strategies to replace fibrotic healing with a successful regenerative response. Our protocol uses fluorescence immunohistochemistry to 1) identify early-and-late blastema cell populations, 2) study revascularization in the context of regeneration, and 3) investigate intramembranous ossification without the need for complex bone stabilization devices. We also demonstrate the use of sequential in vivo &#956;CT to create high resolution images to examine morphological changes after amputation, as well as quantify volume and length changes in the same digit over the course of regeneration. We believe this protocol offers tremendous utility to investigate both epimorphic and tissue regenerative responses in mammals.

Here, we present a protocol of adult mouse terminal phalanx amputation to investigate mammalian blastema formation and intramembranous ossification, analyzed by fluorescent immunohistochemistry and sequential in-vivo microcomputed tomography.

Here, we present a protocol of adult mouse distal terminal phalanx (P3) amputation, a procedurally simple and reproducible mammalian model of epimorphic ...

Developmental Biology

Three and Four-Dimensional Visualization and Analysis Approaches to Study Vertebrate Axial Elongation and Segmentation

Somitogenesis is a hallmark of vertebrate embryonic development. For years, researchers have been studying this process in a variety of organisms using a wide range of techniques encompassing ex vivo and in vitro approaches. However, most studies still rely on the analysis of two-dimensional (2D) imaging data, which limits proper evaluation of a developmental process like axial extension and somitogenesis involving highly dynamic interactions in a complex 3D space. Here we describe techniques that allow mouse live imaging acquisition, dataset processing, visualization and analysis in 3D and 4D to study the cells (e.g., neuromesodermal progenitors) involved in these developmental processes. We also provide a step-by-step protocol for optical projection tomography and whole-mount immunofluorescence microscopy in mouse embryos (from sample preparation to image acquisition) and show a pipeline that we developed to process and visualize 3D image data. We extend the use of some of these techniques and highlight specific features of different available software (e.g., Fiji/ImageJ, Drishti, Amira and Imaris) that can be used to improve our current understanding of axial extension and somite formation (e.g., 3D reconstructions). Altogether, the techniques here described emphasize the importance of 3D data visualization and analysis in developmental biology, and might help other researchers to better address 3D and 4D image data in the context of vertebrate axial extension and segmentation. Finally, the work also employs novel tools to facilitate teaching vertebrate embryonic development.

Here, we describe computational tools and methods that allow visualization and analysis of three and four-dimensional image data of mouse embryos in the context of axial elongation and segmentation, obtained by in toto optical projection tomography, and by live imaging and whole-mount immunofluorescence staining using multiphoton microscopy.

Somitogenesis is a hallmark of vertebrate embryonic development. For years, researchers have been studying this process in a variety of organisms using a ...

Establishment of a Segmental Femoral Critical-size Defect Model in Mice Stabilized by Plate Osteosynthesis

The use of tissue-engineered bone constructs is an appealing strategy to overcome drawbacks of autografts for the treatment of massive bone defects. As a model organism, the mouse has already been widely used in bone-related research. Large diaphyseal bone defect models in mice, however, are sparse and often use bone fixation which fills the bone marrow cavity and does not provide optimal mechanical stability. The objectives of the current study were to develop a critical-size, segmental, femoral defect in nude mice. A 3.5-mm mid-diaphyseal femoral ostectomy (approximately 25% of the femur length) was performed using a dedicated jig, and was stabilized with an anterior located locking plate and 4 locking screws. The bone defect was subsequently either left empty or filled with a bone substitute (syngenic bone graft or coralline scaffold). Bone healing was monitored noninvasively using radiography and in vivo micro-computed-tomography and was subsequently assessed by ex vivo micro-computed-tomography and undecalcified histology after animal sacrifice, 10 weeks postoperatively. The recovery of all mice was excellent, a full-weight-bearing was observed within one day following the surgical procedure. Furthermore, stable bone fixation and consistent fixation of the implanted materials were achieved in all animals tested throughout the study. When the bone defects were left empty, non-union was consistently obtained. In contrast, when the bone defects were filled with syngenic bone grafts, bone union was always observed. When the bone defects were filled with coralline scaffolds, newly-formed bone was observed in the interface between bone resection edges and the scaffold, as well as within a short distance within the scaffold. 
The present model describes a reproducible critical-size femoral defect stabilized by plate osteosynthesis with low morbidity in mice. The new load-bearing segmental bone defect model could be useful for studying the underlying mechanisms in bone regeneration pertinent to orthopaedic applications.

Although mouse models are invaluable tools for bone tissue engineering, models of long bone defects are sparse. This need motivated development of the present protocol which uses a locking plate with four screws and a dedicated jig to perform and stabilize a reproducible, femoral, critical-size defect with low morbidity.

The use of tissue-engineered bone constructs is an appealing strategy to overcome drawbacks of autografts for the treatment of massive bone defects. As a ...

Analysis of Craniomaxillofacial Malformations in Mice Using Three-dimensional Microcomputed Tomography

To model craniofacial malformations caused by vitamin A deficiency (VAD), we expressed a dominant-negative retinoid receptor mutation in osteoblasts to specifically inhibit RAR transcriptional activity in mice. This approach allowed us to investigate the effects of VAD on cranial hypomineralization, mandibular deformity, and clavicular hypoplasia in clinical cases. In this study, microcomputed tomography (microCT) scanning of the craniomaxillofacial region of mice represented a valuable tool for studying the growth and development of this animal model. The manual estimation of images is both time-consuming and inaccurate. Hence, here, we present a straightforward, efficient, and accurate approach for segmenting and quantifying the microCT images of each craniomaxillofacial bone. MicroCT software was used to slice the mandible, frontal bone, parietal bone, nasal bone, premaxilla, maxilla, interparietal bone, and occipital bone of mice and measure their corresponding lengths and widths. This segmentation method can be applied to study growth and development in developmental biology, biomedicine, and other related sciences and allows researchers to analyze the effects of genetic mutations on individual craniofacial bones.

Here, we describe in depth a method, which is based on microcomputed tomography, to segment and measure 3D models of craniomaxillofacial bones in mice, for better assessment of craniomaxillofacial bone development in mice than what is possible with current methods.

To model craniofacial malformations caused by vitamin A deficiency (VAD), we expressed a dominant-negative retinoid receptor mutation in osteoblasts to ...

In Vivo Evaluation of Fracture Callus Development During Bone Healing in Mice Using an MRI-compatible Osteosynthesis Device for the Mouse Femur

Endochondral fracture healing is a complex process involving the development of fibrous, cartilaginous, and osseous tissue in the fracture callus. The amount of the different tissues in the callus provides important information on the fracture healing progress. Available in vivo techniques to longitudinally monitor the callus tissue development in preclinical fracture-healing studies using small animals include digital radiography and &#181;CT imaging. However, both techniques are only able to distinguish between mineralized and non-mineralized tissue. Consequently, it is impossible to discriminate cartilage from fibrous tissue. In contrast, magnetic resonance imaging (MRI) visualizes anatomical structures based on their water content and might therefore be able to noninvasively identify soft tissue and cartilage in the fracture callus. Here, we report the use of an MRI-compatible external fixator for the mouse femur to allow MRI scans during bone regeneration in mice. The experiments demonstrated that the fixator and a custom-made mounting device allow repetitive MRI scans, thus enabling longitudinal analysis of fracture-callus tissue development.

In Vivo Evaluation of Fracture Callus Development During Bone Healing in Mice Using an MRI-compatible Osteosynthesis Device for the Mouse Femur

The evaluation of tissue development in the fracture callus during endochondral bone healing is essential to monitor the healing process. Here, we report the use of a magnetic resonance imaging (MRI)-compatible external fixator for the mouse femur to allow MRI scans during bone regeneration in mice.

Endochondral fracture healing is a complex process involving the development of fibrous, cartilaginous, and osseous tissue in the fracture callus. The ...

HE Staining and Micro-CT to Evaluate Murine Trabecular Bone Microarchitecture

Trabecular Bone Microarchitecture Evaluation in an Osteoporosis Mouse Model

Bone microstructure refers to the arrangement and quality of bone tissue at the microscopic level. Understanding the bone microstructure of the skeleton is crucial for gaining insight into the pathophysiology of osteoporosis and improving its treatment. However, handling bone samples can be complex due to their hard and dense properties. Secondly, specialized software makes image processing and analysis difficult. In this protocol, we present a cost-effective and easy-to-use solution for trabecular bone microstructure analysis. Detailed steps and precautions are provided. Micro-CT is a non-destructive three-dimensional (3D) imaging technique that provides high-resolution images of trabecular bone structure. It allows for the objective and quantitative evaluation of bone quality, which is why it is widely regarded as the gold standard method for bone quality assessment. However, histomorphometry remains indispensable as it offers crucial cellular-level parameters, bridging the gap between two-dimensional (2D) and 3D assessments of bone specimens. As for the histologic techniques, we chose to decalcify the bone tissue and then perform traditional paraffin embedding. In summary, combining these two methods can provide more comprehensive and accurate information on bone microstructure.

Author Spotlight: An Economic and Efficient Method for Quantitative Evaluation of Bone Microarchitecture in a Murine Osteoporosis Model 

This protocol presents an economical and efficient method for quantitative evaluation of bone microarchitecture in a mouse model of osteoporosis by combining Hematoxylin-Eosin (HE) staining and micro-computed tomography (Micro-CT) techniques.

Bone microstructure refers to the arrangement and quality of bone tissue at the microscopic level. Understanding the bone microstructure of the skeleton ...

Establishing a Diaphyseal Femur Fracture Model in Mice

Bones have a significant regenerative capacity. However, fracture healing is a complex process, and depending on the severity of the lesions and the age and overall health status of the patient, failures can occur, leading to delayed union or nonunion. Due to the increasing number of fractures resulting from high-energy trauma and aging, the development of innovative therapeutic strategies to improve bone repair based on the combination of skeletal/mesenchymal stem/stromal cells and biomimetic biomaterials is urgently needed. To this end, the use of reliable animal models is fundamental to better understanding the key cellular and molecular mechanisms that determine the healing outcomes. Of all the models, the mouse is the preferred research model because it offers a wide variety of transgenic strains and reagents for experimental analysis. However, the establishment of fractures in mice may be technically challenging due to their small size. Therefore, this article aims to demonstrate the procedures for the surgical establishment of a diaphyseal femur fracture in mice, which is stabilized with an intramedullary wire and resembles the most common bone repair process, through cartilaginous callus formation.

This protocol describes a surgical procedure for the establishment of a diaphyseal fracture in the femur of mice, which is stabilized with an intramedullary wire, for fracture healing studies.

Bones have a significant regenerative capacity. However, fracture healing is a complex process, and depending on the severity of the lesions and the age ...

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

Summary

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Sequential In vivo Imaging of Osteogenic Stem/Progenitor Cells During Fracture Repair

Computed Tomography and Optical Imaging of Osteogenesis-angiogenesis Coupling to Assess Integration of Cranial Bone Autografts and Allografts

Combined In vivo Optical and µCT Imaging to Monitor Infection, Inflammation, and Bone Anatomy in an Orthopaedic Implant Infection in Mice

Adult Mouse Digit Amputation and Regeneration: A Simple Model to Investigate Mammalian Blastema Formation and Intramembranous Ossification

Three and Four-Dimensional Visualization and Analysis Approaches to Study Vertebrate Axial Elongation and Segmentation

Establishment of a Segmental Femoral Critical-size Defect Model in Mice Stabilized by Plate Osteosynthesis

Analysis of Craniomaxillofacial Malformations in Mice Using Three-dimensional Microcomputed Tomography

In Vivo Evaluation of Fracture Callus Development During Bone Healing in Mice Using an MRI-compatible Osteosynthesis Device for the Mouse Femur

Trabecular Bone Microarchitecture Evaluation in an Osteoporosis Mouse Model

Establishing a Diaphyseal Femur Fracture Model in Mice