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

Summary

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.

Abstract

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.

Introduction

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 "river bank", 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 expanders^4,5,6. 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 model^1,13. 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 suture^16,17. 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 requirement^18,19. 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.

Protocol

All experimental procedures described here were approved by the Animal Care Committee of Shanghai Ninth People'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

1. Preparation of two retention holders.
   1. Use 0.014'' 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).
   2. Sterilize the wires and holders for the surgery in an autoclave, plasma sterilizer, or suitable cold sterilant (e.g., glutaraldehyde).
2. Preparation of the springs.
   1. 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.
   2. 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) if the condition is limited. 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.
   3. 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).

2. Sagittal suture expansion surgery

1. 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.
2. 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 the mice by subcutaneous injection.
3. Body position fixation: Put the mouse lying on the front and use surgical tapes to fix the limbs on the operating table.
4. 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.
5. Bond the retention holders after acid etching.
   1. 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.
   2. 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).
6. Reset the scalp flap.
   1. Manually put back the scalp flap (Figure 2C). Label the holders' position, cut two small holes in the scalp at the corresponding position of the bilateral retention holders, and then reset the flapped scalp.
   2. At the same time, pass the small loops through the holes to expose the skin's surface. Suture the curved incision with a 6-0 suture (Figure 2D). One to three days are allowed for the skin recovery. Administer meloxicam (1 mg/kg) to the mice by subcutaneous injection every 24 h 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.
7. Install the spring and guide wire.
   1. Cut the spring 1 mm longer than the distance between the two holders.
   2. Compress the selected pressure spring and place it between the small coils on both sides.
   3. 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.
   4. Check that the scalp under the spring area of the mouse is without any compression.
   5. 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).

3. Double labeling of mineralized bones

1. Preparation of the stock solution.
   1. Prepare a dilution solution with distilled water that contains 0.9% NaCl and 2% NaHCO₃.
   2. Use the diluent solution to prepare 20 mg/mL of Alizarin complex dihydrate and 10 mg/mL of Calcein (see Table of Materials).
   3. Adjust the pH to 7.4 using a pH measuring device. Flush the pH probe between measurements to ensure accurate readings.
   4. Put the dye into a sterile container and store it in a refrigerator at 4 °C; foil is used to keep light out.
2. 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.
3. 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.
4. Collect whole calvarial bones prepared for tissue clearing procedure a day after the second injection.

4. EdU staining

1. Intraperitoneal injection: Intraperitoneally inject EdU 2 h before euthanizing the mice (following institutionally approved protocols). The dose is 1 mg/10 g body weight.
2. 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 µmol/L final) and Sodium Ascorbate (100 mmol/L final, made fresh each use) (see Table of Materials).
3. 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).

5. Micro-computed tomography imaging

1. Tissue fixation and storage: Fix the calvarial bones in 4% paraformaldehyde (PFA) at 4 °C O/N and store them in 0.5% PFA before scanning.
2. Scanning and analysis: Scan the samples with a micro-computed tomography (µCT) imaging system with high resolution and a voxel size of 7 µm.

6. Preparation of work solution for PEGASOS tissue clearing

1. 4% polyformaldehyde (PFA): Dissolve 4 g of PFA powder in 1× PBS to 100 mL.
2. Cardiac perfusion solution: 0.02% Heparin, that is, 20 mg heparin powder dissolved in 1× 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.
3. 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.
4. Decolorization solution: 25% Quadrol, which is 250 mL of Quadrol (N, N, N', N' - 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 °C to increase its fluidity before configuration.
5. 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 °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 >9.5.
6. 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).
7. Transparent solution: BB-PEG solution, 75% BB + 22% PEG-MMA + 3% Quadrol (v/v), wherein BB is benzyl benzoate (BB).

7. Transparency of calvarial bones with the PEGASOS method

1. Anesthetize the mouse by intraperitoneal injection of anesthetics (step 2.1), and judge the depth of anesthesia of the mouse through the pinch reaction to ensure that there is no resistance reaction before cardiac perfusion.
2. Perform cardiac perfusion and fixation.
   1. 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.
   2. 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.
   3. 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.
   4. 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.
   5. Dissect and separate the tissues and organs, and fix them overnight with 4% PFA at 4 °C.
3. 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 °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.
4. Tissue decolorization: Place the fixed calvarial bones in 25% Quadrol solution (20 mL) in a shaking table at 37 °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.
5. EdU staining: Rinse the samples in 1× PBS for 5 min (three times), and then immerse them in the labeling cocktail for 1 day. After that, rinse the samples in 1× PBS for 1 h (three times).
6. Tissue degreasing and dehydration
   1. Place the tissues in 30% tB, 50% tB, and 70% tB solutions in sequence to perform gradient decreasing on a shaking table at 37 °C. Place in each concentration for about 2 h.
   2. Subsequently, dehydrate the samples in TB-PEG solution for 6 h on a shaking table at 37 °C.
      NOTE: The above processing time can be increased or decreased depending on the number of the tissue.
7. Tissue transparency: Place the completely dehydrated tissue in BB-PEG solution on a shaking table at 37 °C (2-4 h) until transparent. Currently, samples can be stored for up to 1-2 years.
   ​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.

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).

1. 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.
2. 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.
3. 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.
4. 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.
5. 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 "z-Galvo" can be selected when fine regulation is needed.
6. 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.

Results

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

Discussion

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...

Materials

Name	Company	Catalog Number	Comments
37% Acid etching	Xihubiom	E10-02/1807011
Alizarin red	Sigma-Aldrich	A3882
AUSTRALIAN WIRE	A.J.WILCOCK	0.014''
Benzyl benzoate	Sigma-Aldrich	B6630
Calcein green	Sigma-Aldrich	C0875
Copper(II) sulfate, anhydrous	Sangon Biotech	A603008
Dynamometer	Sanliang	SF-10N
EDTA	Sigma-Aldrich	E9884
EdU	Invitrogen	E104152
Laser Confocal Microscope	Leica	SP8
PBS	Sangon Biotech	E607008
PEG-MMA 500	Sigma-Aldrich	447943
PFA	Sigma-Aldrich	P6148
pH Meters	Mettler Toledo	S220
Quadrol	Sigma-Aldrich	122262
Sodium Ascorbate	Sigma-Aldrich	A4034
Sodium bicarbonate	Sangon Biotech	A500873
Sodium chloride	Sangon Biotech	A610476
Sodium hydroxide	Sigma-Aldrich	S5881
Spring	TAOBAO	0.2*1.5*1*7
Sulfo-Cyanine3 azide	Lumiprobe	A1330
tert-Butanol	Sigma-Aldrich	360538	Protect from light. Do not freeze.
Transbond MIP Moisture Insensitive Primer	3M Unitek	712-025
Transbond XT Light Cure Adhesive Paste	3M Unitek	712-035
Triethanolamine	Sigma-Aldrich	V900257
Tris-buffered saline	Sangon Biotech	A500027