This study was supported by R01DE023348 (RHK) from NIDCR/NIH and the Faculty Research Grant (RHK) from the Council on Research of the Academic Senate of the Los Angeles Division of the University of California.

Mice were purchased from Jackson Laboratory and kept in a pathogen-free vivarium in the UCLA Division of Laboratory Animal Medicine (DLAM). The experiments were performed according to the approved institutional guidelines from the Chancellor&#39;s Animal Research Committee&#160;(ARC#2016-037).
1. Mouse Anesthetization
<ol>
	<li>Use eight-week-old female C57/BL6 mice (n = 3).</li>
	<li>Anesthetize the mice using ketamine (80-120 mg/kg of mouse weight)/xylazine (5 mg/kg of mouse weight) solutions and administer intraperitoneally (ip) at a dose of 10 mL/kg.</li>
	<li>Prepare ketamine (80 - 120 mg/kg)/xylazine (5 mg/kg) solutions and administer them intraperitoneally (ip) at a dose of 10 mL/kg.</li>
	<li>Confirm that the mice are fully anesthetized by performing a toe pinch.</li>
</ol>
2. Pulp-capping Procedure
<ol>
	<li>Place the mouth holder in the mouse&#39;s mouth.</li>
	<li>Secure the mouth holder on the table such that the head is facing upward.</li>
	<li>Place the microscope (10X) on top of the mouth so that the first maxillary molar is fully visible.</li>
	<li>Using the &#188;-round bur in a high-speed handpiece at 200,000 rpm, remove the enamel part of the tooth in the middle until the pulp is visible through the transparent dentin. Do not expose the pulp with the bur.</li>
	<li>Using a #15 endodontic K-file (diameter of 150 &#181;m), perforate through the dentin and expose the pulp. 
	NOTE: Special care should be taken so that the dentin debris does not get pushed into the pulp. This can be avoided by rotating the K-file quarterly and then pulling the K-file out.</li>
	<li>Mix MTA with sterile H2O according to the manufacturer&#39;s instructions. Deliver and place the MTA onto the exposed pulp with the tip of the explorer. Use the back side of the paper point (fine) to pack the MTA into the exposed pulp by gentle tapping. The thicker side of the paper point is flat and therefore allows for the proper condensation of the MTA into the exposed pulp.</li>
	<li>Etch the tooth for 15 s by placing the 35% phosphoric acid etchant where it just covers the tooth. Take special care to limit the placement of the etchant, as it may irritate gingival tissues. 
	NOTE: The etchant comes in a syringe and is used to roughen the tooth surfaces so that dental adhesives can flow in to mediate micromechanical bonding onto the tooth. Because they are viscous, it can be self-contained by applying small amounts directly onto the tooth.</li>
	<li>Use negative-pressured suction to remove the etchant. Use a cotton pellet that is lightly soaked with H2O to remove the residuals of the etchant. Repeat this step until the etchant is completely removed from the tooth.</li>
	<li>Using a compressed air duster, gently dry the tooth.</li>
	<li>Apply the dental adhesives using the backside of the paper point.</li>
	<li>Make the adhesive layer thin using compressed air for 3 s.</li>
	<li>Cure the dental adhesives for 20 s using the curing-light unit.</li>
	<li>Place the flowable composite in small amounts onto the tooth that was capped with MTA. Use the tip of the explorer to flow the composite into the tooth grooves.</li>
	<li>Cure the composite for 30 s using a light-curing unit to polymerize it. Confirm that the composite is fully cured and hard using the explorer.</li>
</ol>
3. Post-op Care
<ol>
	<li>Administer carprofen (5 mg/kg) subcutaneously (sc) immediately after the pulp-capping procedure.</li>
	<li>Place the mice on a heating pad at low power to keep the animals warm before they wake up.</li>
	<li>Return the mice to the vivarium for housing.</li>
</ol>
4. Tissue Procurement
<ol>
	<li>After 5 - 6 weeks, euthanize the mice by cervical dislocation under a complete anesthetic condition with isoflurane.</li>
	<li>Carefully remove the maxilla out of the base of the skull and put it into a 50-mL tube. Fix the entire maxilla that contains both the pulp-capped tooth and the contralateral uncapped tooth in 4% paraformaldehyde in PBS, pH 7.4, at 4 &#176;C overnight, and then store it in a 70% ethanol solution. 
	NOTE: Paraformaldehyde is toxic and carcinogenic. The proper use paraformaldehyde should be monitored as outlined in the standard operating procedures (SOP).</li>
	<li>Scan the mouse maxillae using the &#181;CT scan. To secure the maxillae during scanning, wrap the samples with gauze soaked with 70% ethanol and place them in the 15-mL cell culture tube.</li>
</ol>
5. &#181;CT Scanning
<ol>
	<li>Prepare the samples for &#181;CT scanning. Briefly, wrap the samples with gauze soaked with 70% ethanol and secure them in a generic 15-mL cell culture conical tube. Mount the tube onto the &#181;CT scanning stage, as outlined in the manufacturer&#39;s instructions.</li>
	<li>Set the X-ray source to a current of 145 &#181;A, a voltage of 55 kVp, and an exposure time of 200 ms.</li>
	<li>Perform image acquisition with the &#181;CT scanner at a 20-&#181;m resolution and with a 0.5 mm Al filter.</li>
	<li>Reconstruct the image and visualize it11.</li>
	<li>Once the &#181;CT scan is complete, start decalcification with 5% EDTA and 4% sucrose in PBS (pH 7.4) for 2 weeks.</li>
</ol>
6. Tissue Processing and Staining
<ol>
	<li>Embed the decalcified tissues in paraffin. Before embedding, trim the maxilla by making a sagittal cut immediately anterior to the first molar. While embedding, position this surface downward, such that the longitudinal section of the first molar is the cutting surface.</li>
	<li>Using the microtome, prepare 5 &#181;m-thick slides. The pulp-capping areas usually coincide with the distopalatal (DP) root, which can be used as a landmark. Determine the precise area of interest by examining the histology under the light microscope and comparing the &#181;CT images.</li>
	<li>For H&#38;E staining, deparaffinize and rehydrate the slides with xylene (2x) and serially diluted ethanol (100% EtOH 2x, 95% EtOH 2x, and 70% EtOH 1x).</li>
	<li>Rinse the slides with running tap water.</li>
	<li>Stain with Hematoxylin solution for 2.5 min and rinse with tap water.</li>
	<li>Dip the slides in 95% ethanol for 1 min.</li>
	<li>Stain with Eosin solution for 1 min and rinse with tap water.</li>
	<li>Dehydrate with serially diluted ethanol (70% EtOH 1x, 95% EtOH 2x, and 100% EtOH 3x) and xylene (3x).</li>
	<li>Mount the slides with mounting solution.</li>
</ol>

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The authors have nothing to disclose.

Currently, there are several different experimental models available to validate the in vivo effects of dental materials, scaffolds, or growth factors on odontogenic differentiation of dental pulp stem cells (DPSCs)13. These models include ectopic autologous transplantation of DPSCs into an organ, such as the renal capsule, or subcutaneous transplantation of DPSCs into immunocompromised mice with scaffolds14,15. However, these methods are limited in that their odontogenic effect on DPSCs is not performed in the orthotopic pulp environment. On the other hand, orthotopic transplantation into the pulp or pulp-capping procedures on a tooth are used in larger animals16,17. Although these models are valuable in evaluating odontogenic potential in the orthotopic environment, the use of those models is largely observational in nature, providing limited mechanistic insights on pulp wound healing and reparative dentin formation.
In this paper, we present a detailed method to perform pulp-capping in mice. This step-by-step procedure includes anesthetizing the mice, preparing the Class-I-like cavity, placing the pulp-capping materials, harvesting the maxillae, analyzing with the &#181;CT scan, and assessing tissue samples for reparative dentin formation. Our pulp-capping mouse model will be instrumental in investigating the fundamental molecular mechanisms of pulpal wound healing in vivo in the context of reparative dentin by enabling the use of transgenic or knockout mice, which are widely available in the research community.
Recent studies demonstrated several mouse models in which dentin formation was observed18,19. Saito et al. created a Class-I-like preparation without pulp exposure, which stimulates reactionary, not reparative, dentin formation. Both reactionary dentin and reparative dentin are classified as tertiary dentin, which forms following external stimulation to the tooth. However, unlike reactionary dentin, which is formed by existing odontoblasts, reparative dentin is formed by odontoblast-like cells, such as DPSCs, when the pulp becomes exposed and the odontoblastic layers are breached20. Therefore, it does not represent an actual pulp-capping procedure in the clinic. In another study, glass ionomer was used to cap the pulp exposure19. However, a clinical study showed that the glass ionomer induced chronic inflammation, but not reparative dentin21. In this regard, our pulp-capping mouse model better represents the actual pulp-capping procedure in patients.
It is noteworthy that when we harvested mice after more than 6 weeks, reparative dentin formation occurred throughout the pulp chamber and the root canals (Figure 4). Such an observation is rather unexpected, as we anticipated reparative dentin formation at the junction between the pulp-capping material and the pulp. However, potent mineralization of the pulp is also observed in clinical settings, especially in relatively young patients22. Because the 8-week-old mice used in this study are considered to be &#34;young adults&#34;23, a possibility exists whereby these mice still harbor significant odontogenic potential. Therefore, it would be worthwhile to examine the ageing effects of reparative dentin formation in mice.
Our histological examinations revealed that, although reparative dentin was clearly formed in the pulp-capped tooth, there were characteristics of both dentin and bone formation, as evidenced by the presence of dentinal tubules (red arrow) and osteocytes (black arrowheads) in the reparative dentin (Figure 5). Such observations suggest that reparative dentin formation may be induced by locally residing odontoblast-like dental pulp stem cells, as well as infiltrating mesenchymal stem cells from the surrounding bone.
Compared to the dentin-forming cells, we found no dentin-resorbing cells within the pulp, as determined by tartrate-resistant acid phosphatase (TRAP) staining (data not shown). Indeed, pulpal or periapical inflammation induces osteoclast formation on the bone surfaces around the tooth, but not on the dentin surfaces due to as-yet-unknown mechanisms24. Of note, there was a clear demarcation between the existing dentin and the newly formed reparative dentin (Figure 4). A previous study demonstrated a similar phenomenon; when a tooth is extracted in the presence of bisphosphonate or anti-RANKL antibody, both of which inhibit the functions of osteoclasts, there were clear demarcations between existing lamellar bone and newly formed woven bone25. This notion further supports the absence of dentin-resorbing cells in the pulp. Collectively, our established mouse model would provide unique opportunities to examine the mechanisms of pulp wound healing and reparative dentin formation in vivo.
There is a limitation to the pulp-capping mouse model. Genetic makeups between humans and mice are clearly different. The complete genome has been sequenced in humans and mice, and there is about 85% similarity in the protein-coding regions between mice and humans26,27. In line with this notion, it was suggested that findings related to pulp capping in animals do not necessarily reflect those in humans28. Nonetheless, animal models are extensively used in the research community to recapitulate human diseases in vivo, such as collagen-induced arthritis for rheumatoid arthritis29, ovariactomy-induced bone loss for osteoporosis30, lipopolysaccharide (LPS) administration for systemic shock31, and ligature placement for periodontitis32. As such, the pulp-capping mouse model will be essential to examine the molecular mechanisms of pulp wound healing and reparative dentin formation in vivo. Nonetheless, just like other animal models, interpretation and validation of the findings from the pulp-capping mouse model should be carefully evaluated.
In summary, the current study demonstrates successful pulp capping in mice. Unlike other known models, this pulp-capping mouse model will provide an invaluable research tool in the field of pulp regeneration and reparative dentin formation because it provides: 1) an opportunity to utilize widely available genetically engineered mouse strains to elucidate the underlying mechanisms at the molecular level and 2) an economically efficient way to obtain statistically significant results by increasing sample sizes. Further studies await, including objective quantification of reparative dentin formation in vivo, age-dependent effects of reparative dentin formation, evaluation of clinically available pulp-capping materials, and validation of molecular determinants that are required for proper pulpal wound healing and reparative dentin regeneration.

Dental caries are one of the most prevalent oral diseases and the leading cause of surgical interventions to dentitions in almost all individuals1,2. The prognosis of surgical interventions and restorations of a tooth largely depends upon proper pulpal response and successful wound healing. Indeed, dental caries that penetrate deeply through the enamel and dentin frequently lead to the exposure of the underlying pulp tissue that is often "capped" with dental materials, such as calcium hydroxide (Ca(OH)2) or hydraulic calcium-silicate cements (HCSCs), including mineral trioxide aggregates (MTA). The ultimate goal of such a pulp-capping procedure is to expedite pulpal wound healing by regenerating reparative dentin, a physical barrier that functions as a "biological seal" to protect the underlying pulp tissue and to increase the life expectancy of the tooth and the overall oral health. However, the underlying mechanism of pulpal wound healing and reparative dentin formation is not fully understood.
To better understand the mechanisms of pulpal wound healing and reparative dentin formation in vivo, several animals were previously used, including monkeys, dogs, and pigs3-5. Among them, rats are frequently used because they are relatively smaller in sizes compared to the other animals, but their teeth are large enough to perform direct pulp capping without any technical difficulties6-10. These animal models are ideal alternatives to human studies for examining pulpal responses and reparative dentin formation. However, their utilization is limited to observational studies at the cellular level, and they scarcely provide mechanistic insights during reparative dentin formation at the molecular level.
Recent technical advances in genetic engineering provided invaluable and indispensable research tools-mice that harbor a gene that is either overexpressed or deleted-that are instrumental to studying molecular mechanisms of human diseases in vivo. The numbers of different strains of transgenic or knockout mice that are strategically inducible in a cell-specific manner are continually growing in the scientific community. Therefore, examining pulpal wound healing and reparative dentin regeneration in these mice would greatly help to expedite our understanding of these processes at the molecular level. However, the use of mice is significantly dampened, as performing a pulp-capping procedure on a mouse tooth is technically challenging due to its miniature size. Here, we present our reproducible method of performing direct pulp capping in mice for the evaluation of pulpal wound healing and reparative dentin formation in vivo.

<table><tbody><tr><td>BM-LED stereo microscope</td><td>MEIJI Techno</td><td></td><td>Microscope&amp;nbsp;</td></tr><tr><td>Optima MCX-LED&amp;nbsp;</td><td>Bien Air Dental</td><td>1700588-001</td><td>Electic motor engine</td></tr><tr><td>isoflurane</td><td>Henry schein animal health</td><td>NDC 11695-0500-2</td><td></td></tr><tr><td>1/4 round bur</td><td>Brasseler</td><td>001092T0</td><td></td></tr><tr><td>Endodontic K-file</td><td>Roydent</td><td>98947</td><td></td></tr><tr><td>ProRoot MTA</td><td>Dentsply</td><td>PROROOT5W</td><td>MTA</td></tr><tr><td>Paper point</td><td>Henry schein</td><td>100-3941</td><td></td></tr><tr><td>Ultra-Etch</td><td>Ultradent product Inc.</td><td></td><td>Phosphoric acid etchant</td></tr><tr><td>OptiBond SoloPlus</td><td>Kerr</td><td>29669</td><td>Adhesives</td></tr><tr><td>Coltolux LED</td><td>Coltene/whaledent Inc.</td><td>C7970100115</td><td>Curing light unit</td></tr><tr><td>Characterization tint</td><td>Bisco</td><td>T-14012</td><td>Flowable composite</td></tr><tr><td>Skyscan</td><td>Breuker</td><td>1275</td><td>uCT scanner</td></tr><tr><td>Microm</td><td>Thermo</td><td>HM355S</td><td>Microtome</td></tr><tr><td>Hematoxyline-1</td><td>Thermo Scientific</td><td>7221</td><td></td></tr><tr><td>Eosin-Y</td><td>Thermo Scientific</td><td>7111</td><td></td></tr><tr><td>Cytoseal 60</td><td>Thermo Scientific</td><td>8310-16</td><td>Mounting solution</td></tr></tbody></table>

development of a direct pulp-capping model for the evaluation of pulpal wound healing and reparative dentin formation in mice

Dental pulp is a vital organ of a tooth fully protected by enamel and dentin. When the pulp is exposed due to cariogenic or iatrogenic injuries, it is often capped with biocompatible materials in order to expedite pulpal wound healing. The ultimate goal is to regenerate reparative dentin, a physical barrier that functions as a "biological seal" and protects the underlying pulp tissue. Although this direct pulp-capping procedure has long been used in dentistry, the underlying molecular mechanism of pulpal wound healing and reparative dentin formation is still poorly understood. To induce reparative dentin, pulp capping has been performed experimentally in large animals, but less so in mice, presumably due to their small sizes and the ensuing technical difficulties. Here, we present a detailed, step-by-step method of performing a pulp-capping procedure in mice, including the preparation of a Class-I-like cavity, the placement of pulp-capping materials, and the restoration procedure using dental composite. Our pulp-capping mouse model will be instrumental in investigating the fundamental molecular mechanisms of pulpal wound healing in the context of reparative dentin in vivo by enabling the use of transgenic or knockout mice that are widely available in the research community.

Here, we showed the step-by-step procedures to perform pulp capping on mice teeth. One of the key aspects of pulp capping in mice is to have the appropriate apparatus. In this regard, having the microscope with a 10X power magnification is essential (Figure 1A). To create a Class-I-like preparation in the tooth, we used a ¼-round burr in an electric high speed handpiece at 200,000 rpm (Figure 1B). Alternatively, any other engines, including those that use compressed air, can be used to prepare a tooth.
In Figure 2A-2E, representative steps for performing pulp capping are shown. The Class-I-like preparation was performed (Figure 2B). Because a water spray may drown the mice during the procedure, its use is not recommended. For this reason, it is essential to prepare the tooth with gentle and intermittent strokes to prevent overheating. The use of compressed air is also recommended to provide cooling effects. While exposing the pulp with an endodontic file, take caution not to push the dentin debris into the pulp, as this may interfere with the data interpretation in reparative dentin formation (Figure 2C). This can be avoided by using the compressed air. Similarly, the MTA should be placed on the exposed pulp without pushing too far into the pulp. MTA placement can be achieved by using the backside of the paper point with gentle tapping motions (Figure 2D). Following MTA placement, the tooth should be cleaned using H2O-soaked cotton pellets to remove any remaining MTA in the grooves, which may interfere with the binding of the composite onto the tooth. Conventional methods of composite restoration are used, including etching the tooth surfaces, priming and bonding with adhesives, and placing and curing the flowable composites (Figure 2E).
Six weeks after pulp capping, the mice were harvested, and the top view was photographed to confirm that the composite was still intact (Figure 3A). µCT scanning showed significant recession of pulpal space in the pulp-capped group (Figure 3B), suggesting that reparative dentin was formed in the pulp. The decalcified tissues samples were subjected to H&amp;E staining to further examine histologically the formation of reparative dentin in vivo. In the control group, odontoblastic layers (OB) were prominently evident around the edges of the dentin (Figure 4A - 4C). In contrast, the pulp-capping group had significant amounts of reparative dentin (RD) formed in the pulpal space (Figure 4D - 4F). Interestingly, a closer examination revealed that reparative dentin (RD) exhibited a typical characteristic of dentin (e.g., striated lines representing dentinal tubules, red arrow), as well as that of bone (e.g., osteocytes representing trapped osteoblasts, black arrowheads). When we stained for Dentin Matrix Protein 1 (DMP1), a marker for odontogenic differentiation12, we found a significant increase in DMP1 expression in the pulp of the pulp-capped tooth when compared to that of the uncapped tooth (Figure 5A and 5B), indicating that reparative dentin was formed within the pulp.
<img alt="Microscopy setup with light source; optical analysis and sample examination equipment." src="/files/ftp_upload/54973/54973fig1.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1: The Equipment Setup for the Pulp-capping Procedure. (A) A microscope (10X) for visualizing the mouse tooth. (B) The high-speed handpiece and the electric motor engine for preparing a Class I preparation to expose the pulp. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54973/54973fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Dental restoration process; stages from initial tooth to composite placement in clinical procedure." src="/files/ftp_upload/54973/54973fig2.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2: Representative Steps in the Capping Procedure. (A) Unprepared tooth on the maxillary first molar in a mouse. (B) Initial enamel removal using the quarter-round bur. (C) Pulp exposure using the endodontic file. (D) MTA placement in the exposed pulp. (E) Composite restoration placement on the tooth. The bar represents 500 µm. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54973/54973fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Dental pulp therapy comparison; microscopy image; pulp-capped vs. control; clinical study results." src="/files/ftp_upload/54973/54973fig3.jpg" data-altupdatedat="1747911167000" data-alt="Figure 3"> 
Figure 3: Clinical Presentation and µCT Scanning of the Pulp-capped and Uncapped Tooth in Mice. (A) Occlusal view of the mouse maxillae on the pulp-capped tooth (left) and uncapped tooth (right). (B) The cross-sectional µCT images of the maxillae. The bar represents 500 µm. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54973/54973fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Histological examination, odontoblast differentiation, dentin formation, microscope imagery." src="/files/ftp_upload/54973/54973fig4.jpg" data-altupdatedat="1747911167000" data-alt="Figure 4"> 
Figure 4: Histological Evidence of Reparative Dentin Formation In Vivo. (A-C) H&amp;E staining of the uncapped maxillary first molar in a mouse at 100X, 200X, and 400X. (D-E) H&amp;E staining of the pulp-capped maxillary first molar in a mouse at 100X, 200X, and 400X. The bar represents 100 µm (OB = odontoblast layers; RD = reparative dentin; black arrowheads = osteocytes; red arrow = dentinal tubules). <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54973/54973fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Histology slide comparison, stained tissue sections A and B, cellular structure analysis." src="/files/ftp_upload/54973/54973fig5.jpg" data-altupdatedat="1747911167000" data-alt="Figure 5"> 
Figure 5: Immunohistochemical Staining of DMP1. (A) DMP1 staining of the uncapped maxillary first molar in a mouse at 400X. (B) DMP1 staining of the pulp-capped maxillary first molar in a mouse at 400X. The bar represents 100 µm. <a href="https://cloudflare-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54973/54973fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Development of a Direct Pulp-capping Model for the Evaluation of Pulpal Wound Healing and Reparative Dentin Formation in Mice at JoVE.com

Development of a Direct Pulp-capping Model for the Evaluation of Pulpal Wound Healing and Reparative Dentin Formation in Mice

We describe a step-by-step method of performing direct pulp capping on mice teeth for the evaluation of pulpal wound healing and reparative dentin formation in vivo.

Dental pulp is a vital organ of a tooth fully protected by enamel and dentin. When the pulp is exposed due to cariogenic or iatrogenic injuries, it is ...

development-direct-pulp-capping-model-for-evaluation-pulpal-wound

Nanyang Technological University

Research

JoVE Journal

Medicine

11.2K Views.  The UCLA School of Dentistry.  We beschrijven een stap voor stap methode om direct pulp aftopping op muizen tanden voor de evaluatie van pulpa wondgenezing en vorming herstellende dentine in vivo. 

Video: Development of a Direct Pulp-capping Model for the Evaluation of Pulpal Wound Healing and Reparative Dentin Formation in Mice

Murine Model of Wound Healing

Wound healing and repair are the most complex biological processes that occur in human life. After injury, multiple biological pathways become activated. Impaired wound healing, which occurs in diabetic patients for example, can lead to severe unfavorable outcomes such as amputation. There is, therefore, an increasing impetus to develop novel agents that promote wound repair. The testing of these has been limited to large animal models such as swine, which are often impractical. Mice represent the ideal preclinical model, as they are economical and amenable to genetic manipulation, which allows for mechanistic investigation. However, wound healing in a mouse is fundamentally different to that of humans as it primarily occurs via contraction. Our murine model overcomes this by incorporating a splint around the wound. By splinting the wound, the repair process is then dependent on epithelialization, cellular proliferation and angiogenesis, which closely mirror the biological processes of human wound healing. Whilst requiring consistency and care, this murine model does not involve complicated surgical techniques and allows for the robust testing of promising agents that may, for example, promote angiogenesis or inhibit inflammation. Furthermore, each mouse acts as its own control as two wounds are prepared, enabling the application of both the test compound and the vehicle control on the same animal. In conclusion, we demonstrate a practical, easy-to-learn, and robust model of wound healing, which is comparable to that of humans.

A murine model of cutaneous wound healing that can be used to assess therapeutic compounds in physiological and pathophysiological settings.

Wound healing and repair are the most complex biological processes that occur in human life. After injury, multiple biological pathways become activated. ...

A Co-Culture Method to Study Neurite Outgrowth in Response to Dental Pulp Paracrine Signals

Tooth innervation allows teeth to sense pressure, temperature and inflammation, all of which are crucial to the use and maintenance of the tooth organ. Without sensory innervation, daily oral activities would cause irreparable damage. Despite its importance, the roles of innervation in tooth development and maintenance have been largely overlooked. Several studies have demonstrated that DP cells secrete extracellular matrix proteins and paracrine signals to attract and guide TG axons into and throughout the tooth. However, few studies have provided detailed insight into the crosstalk between the DP mesenchyme and neuronal afferents. To address this gap in knowledge, researchers have begun to utilize co-cultures and a variety of techniques to investigate these interactions. Here, we demonstrate the multiple steps involved in co-culturing primary DP cells with TG neurons dispersed on an overlying transwell filter with large diameter pores to allow axonal growth through the pores. Primary DP cells with the gene of interest flanked by loxP sites were utilized to facilitate gene deletion using an Adenovirus-Cre-GFP recombinase system. Using TG neurons from the Thy1-YFP mouse allowed for precise afferent imaging, with expression well above background levels by confocal microscopy. The DP responses can be investigated via protein or RNA collection and analysis, or alternatively, through immunofluorescent staining of DP cells plated on removable glass coverslips. Media can be analyzed using techniques such as proteomic analyses, although this will require albumin depletion due to the presence of fetal bovine serum in the media. This protocol provides a simple method that can be manipulated to study the morphological, genetic, and cytoskeletal responses of TG neurons and DP cells in response to the controlled environment of a co-culture assay.

We describe the isolation, dispersion and plating of dental pulp (DP) primary cells with trigeminal (TG) neurons cultured atop overlying transwell filters. Cellular responses of DP cells can be analyzed with immunofluorescence or RNA/protein analysis. Immunofluorescence of neuronal markers with confocal microscopy permits the analysis of neurite outgrowth responses.

Tooth innervation allows teeth to sense pressure, temperature and inflammation, all of which are crucial to the use and maintenance of the tooth organ. ...

Developmental Biology

Murine Excisional Wound Healing Model and Histological Morphometric Wound Analysis

The murine excisional wound model has been used extensively to study each of the sequentially overlapping phases of wound healing: inflammation, proliferation and remodeling. Murine wounds have a histologically well-defined and easily recognizable wound bed over which these different phases of the healing process are measurable. Within the field, it is common to use an arbitrarily defined &#8220;middle&#8221; of the wound for histological analyses. However, wounds are a three-dimensional entity and often not histologically symmetrical, supporting the need for a well-defined and robust method of quantification to detect morphometric defects with a small effect size. In this protocol, we describe the procedure for creating bilateral, full-thickness excisional wounds in mice as well as a detailed instruction on how to measure morphometric parameters using an image processing program on select serial sections. The two-dimension measurements of wound length, epidermal length, epidermal area, and wound area are used in combination with the known distance between sections to extrapolate the three-dimension epidermal area covering the wound, overall wound area, epidermal volume and wound volume. Although this detailed histological analysis is more time and resource consuming than conventional analyses, its rigor increases the likelihood of detecting novel phenotypes in an inherently complex wound healing process.

This protocol describes how to generate bilateral, full-thickness excisional wounds in mice and how to subsequently monitor, harvest, and prepare the wounds for morphometric analysis. Included is an in-depth description of how to use serial histological sections to define, precisely quantify and detect morphometric defects.

The murine excisional wound model has been used extensively to study each of the sequentially overlapping phases of wound healing: inflammation, ...

Live Tracking of Dental Cells via Two-Photon Microscopy

Using Ex Vivo Live Imaging to Investigate Cell Divisions and Movements During Mouse Dental Renewal

The continuously growing mouse incisor is emerging as a highly tractable model system to investigate the regulation of adult epithelial and mesenchymal stem cells and tooth regeneration. These progenitor populations actively divide, move, and differentiate to maintain tissue homeostasis and regenerate lost cells in a responsive manner. However, traditional analyses using fixed tissue sections could not capture the dynamic processes of cellular movements and interactions, limiting our ability to study their regulations. This paper describes a protocol to maintain whole mouse incisors in an explant culture system and live-track dental epithelial cells using multiphoton timelapse microscopy. This technique adds to our existing toolbox for dental research and allows investigators to acquire spatiotemporal information on cell behaviors and organizations in a living tissue. We anticipate that this methodology will help researchers further explore mechanisms that control the dynamic cellular processes taking place during both dental renewal and regeneration.

Author Spotlight: Understanding Dynamic Cellular Behaviors in Adult Mouse Dental Tissue Renewal and Repairment 

Ex vivo live imaging is a powerful technique for studying the dynamic processes of cellular movements and interactions in living tissues. Here, we present a protocol that implements two-photon microscopy to live track dental epithelial cells in cultured whole adult mouse incisors.

The continuously growing mouse incisor is emerging as a highly tractable model system to investigate the regulation of adult epithelial and mesenchymal ...

Assessment of Acute Wound Healing using the Dorsal Subcutaneous Polyvinyl Alcohol Sponge Implantation and Excisional Tail Skin Wound Models.

Wound healing is a complex process that requires the orderly progression of inflammation, granulation tissue formation, fibrosis, and resolution. Murine models provide valuable mechanistic insight into these processes; however, no single model fully addresses all aspects of the wound healing response. Instead, it is ideal to use multiple models to address the different aspects of wound healing. Here, two different methods that address diverse aspects of the wound healing response are described. In the first model, polyvinyl alcohol sponges are subcutaneously implanted along the mouse dorsum. Following sponge retrieval, cells can be isolated by mechanical disruption, and fluids can be extracted by centrifugation, thus allowing for a detailed characterization of cellular and cytokine responses in the acute wound environment. A limitation of this model is the inability to assess the rate of wound closure. For this, a tail skin excision model is utilized. In this model, a 10 mm x 3 mm rectangular piece of tail skin is excised along the dorsal surface, near the base of the tail. This model can be easily photographed for planimetric analysis to determine healing rates and can be excised for histological analysis. Both described methods can be utilized in genetically altered mouse strains, or in conjunction with models of comorbid conditions, such as diabetes, aging, or secondary infection, in order to elucidate wound healing mechanisms.

Here, two murine wound healing models are described, one designed to assess cellular and cytokine wound healing responses and the other to quantify the rate of wound closure. These methods can be used with complex disease models such as diabetes to determine mechanisms of various aspects of poor wound healing.

Wound healing is a complex process that requires the orderly progression of inflammation, granulation tissue formation, fibrosis, and resolution. Murine ...

Immunology and Infection

Digital Planimetry for Assessing Wound Closure Kinetics in a Mouse Model

Chronic wounds, due to their high prevalence, are a serious global health concern. Effective therapeutic strategies can significantly accelerate healing, thereby reducing the risk of complications and alleviating the economic burden on healthcare systems. Although numerous experimental studies have investigated wound healing, most rely on qualitative observations or quantitative direct measurements. The objective of this study was to standardize an indirect wound measurement method using digital planimetry, incorporating digital scaling and segmentation. This approach addresses the lack of detailed, step-by-step methodologies for accurate wound assessment. A photodocumentation booth was designed and constructed, and computer-assisted digital planimetry tools were employed to minimize variability in measurements of the wound area, perimeter, and the distance from the wound center to its edges. A circular traumatic wound (5 mm in diameter) was created on the dorsal midline at the shoulder blade level of male CD1 mice (n = 4, 10 weeks old, 30-35 g). Wound evolution was photodocumented for 14 days using the custom-designed photo booth, which controlled lighting conditions, focal distance, and subject positioning. Scaling and wound measurements were performed using segmentation in ImageJ software, and statistical analysis was conducted using statistical analysis software. The kinetics of wound closure showed a slight increase in wound size and perimeter between day 0 and day 2, followed by a gradual decrease until complete closure by day 14. The photodocumentation booth and computer-assisted digital planimetry enabled quantitative measurements with minimal variability. In conclusion, these tools provide a reliable and reproducible method for evaluating wound closure kinetics in pre-clinical models.

Wounds represent a global health challenge. This study developed a standardized photo booth utilizing digital planimetry to minimize wound measurement variability. Monitoring wounds in mice over 14 days revealed an initial increase in wound area and perimeter, followed by gradual closure. This methodology may aid in evaluating wound closure kinetics in pre-clinical models.

Chronic wounds, due to their high prevalence, are a serious global health concern. Effective therapeutic strategies can significantly accelerate healing, ...

Bioengineering

Creating a Pulp Inflammation Mouse Model Using a Novel Mouth-Gag 

Establishment of a Murine Pulp Exposure Model with a Novel Mouth-Gag for Pulpitis Research

Pulpitis, a common cause of natural tooth loss, leads to necrosis and loss of bioactivity in the inflamed dental pulp. Unraveling the mechanisms underlying pulpitis and its efficient treatment is an ongoing focus of endodontic research. Therefore, understanding the inflammatory process within the dental pulp is vital for improving pulp preservation. Compared to other in vitro experiments, a murine pulpitis model offers a more authentic and genetically diverse context to observe the pathological progression of pulpitis. However, using mice, despite their cost-effectiveness and accessibility, poses difficulties due to their small size, poor coordination, and low tolerance, complicating intraoral and dental procedures. This protocol introduces a novel design and application of a mouth-gag to expose mouse pulp, facilitating more efficient intraoral procedures. The mouth-gag, comprised of a dental arch readily available to most dentists and can significantly expedite surgical preparation, even for first-time procedures. Micro-CT, hematoxylin-eosin (HE) staining, and immunofluorescence staining were used to identify changes in morphology and cell expression. The aim of this article is to help researchers establish a more reproducible and less demanding procedure for creating a pulp inflammation model using this novel mouth-gag.

Author Spotlight: Enhancing Dental Pulp Research with Improved Mouse Models

This article presents a streamlined protocol for establishing a pulpitis model in mice using an innovative mouth-gag, followed by subsequent histological analysis.

Pulpitis, a common cause of natural tooth loss, leads to necrosis and loss of bioactivity in the inflamed dental pulp. Unraveling the mechanisms ...

Inducing Apical Periodontitis in Mice

The mechanisms involved in local induced inflammation can be studied using several available animal models. One of these is the induction of apical periodontitis (AP). Apical periodontitis is a common pathology of an inflammatory nature in the periodontal tissues surrounding the tooth root. In order to better understand the nature and mechanism of this pathology it is advantageous to perform the procedure in mice. The induction of this odontogenic inflammation is achieved by drilling into the mouse tooth until the dental pulp is exposed. Next, the tooth pulp remains exposed to be contaminated by the natural oral flora over time, causing apical periodontitis. After this time period, the animal is sacrificed, and the tooth and the jaw bone can be analyzed in various ways. Typical analyses include micro-CT imaging (to evaluate bone resorption), histological staining, immunohistochemistry, and RNA expression. This protocol is useful for research in the field of oral biology to better understand this inflammatory process in an in vivo experimental setting with uniform conditions. The procedure requires a careful handling of the mice and the isolated jaw, and a visual demonstration of the technique is useful. All technical aspects of the procedures leading to induced apical periodontitis and its characterization in a mouse model are demonstrated.

Here, we present a protocol to locally induce apical periodontitis in mice. We show how to drill a hole in the mouse&#39;s tooth and expose its pulp, in order to cause local inflammation. Analysis methods to investigate the nature of this inflammation, such as micro-CT and histology, are also demonstrated.

The mechanisms involved in local induced inflammation can be studied using several available animal models. One of these is the induction of apical ...

Biology

Optimized Protocol for Mouse Wound Healing Model with Stem Cell-Derived Exosomes

Mouse Wound Models and Preparation of Single-Cell Suspensions 

The management of acute full-thickness skin injuries presents a considerable challenge in clinical practice. The complexity of wound healing, involving diverse cell populations and the intricate wound microenvironment, complicates the assessment of treatment effects and their interactions using traditional experimental approaches. Single-cell transcriptome sequencing technology has revolutionized the understanding of cellular functions and mechanisms during wound healing. However, the variability in methodologies for creating mouse wound models introduces confounding factors that can impact experimental results. Furthermore, the process of dissociating mouse skin tissues presents significant technical hurdles, highlighting the critical need for high-quality single-cell suspensions for accurate transcriptome sequencing. Therefore, this study aims to optimize the construction of mouse wound models to accurately assess changes in wound closure while eliminating variables that could influence the subsequent sequencing result. Additionally, the preparation of single-cell suspensions was performed using both enzyme preparation protocols and test kit protocols to optimize cost and effectiveness.

Mouse Wound Models and Preparation of Single-Cell Suspensions

This study presents two crucial experimental techniques: the construction of a murine wound model for assessing wound closure and the preparation of single-cell suspensions from murine skin to preserve cellular viability.

The management of acute full-thickness skin injuries presents a considerable challenge in clinical practice. The complexity of wound healing, involving ...

The Establishment of a Murine Mandibular Molar Extraction Socket Healing Model

This study introduces the development of a molar extraction model in the murine mandible to provide a practicable model for studying alveolar bone regeneration and intramembranous ossification. C57/J6 mice were used to extract the mandibular first molar to establish this model. They were euthanized, and the bilateral mandibles harvested, at 1 week and 4 weeks post-surgery, respectively. Subsequent serial stereoscopic harvest, histological assessment, and immunofluorescence staining were performed to demonstrate successful surgery. Immediately after surgery, the stereoscopic images displayed an empty extraction socket. The hematoxylin and eosin (H&#38;E) at 1 week and Masson staining at 4 weeks post-surgery showed that the area of the original root was partially and fully filled with bone trabeculae, respectively. The immunofluorescence staining showed that, compared with the homeostasis side, the Sp7 expression increased at 1 week post-surgery, suggesting vigorous osteogenesis in the alveolar fossa. All these results demonstrated a practicable murine tooth extraction socket healing model. Upcoming studies revealing the mechanisms of jawbone defect healing or socket healing could adopt this method.

This protocol demonstrates step-by-step details of how to extract the mandibular first molar in the mouse. It provides an alternative method for researchers focusing on jawbone healing and regeneration.

Development of a Direct Pulp-capping Model for the Evaluation of Pulpal Wound Healing and Reparative Dentin Formation in Mice

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