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

The dental pulp, an integral part of the tooth, is responsible for multiple essential functions such as nutrient supply, dentin formation, sensory function, and defense reactions¹. Nevertheless, the dental pulp, surrounded by hard tissue, is susceptible to injuries and damages from deep caries, pulpitis, trauma, or subsequent therapies^2,3. The absence of functional dental pulp increases the risk of tooth fragility⁴. Moreover, the loss of pulp vitality in young permanent teeth can adversely affect tooth maturation, and current denture techniques fail to restore the neural feedback offered by healthy pulp⁴. This situation has led researchers to explore alternative solutions for managing inflamed pulp beyond mere removal.

In 2007, Murray et al. initiated the application of tissue engineering in regenerative endodontics, thereby sparking increased interest in pulp preservation and regeneration⁵. However, inflamed pulp tissue poses a challenge as cells release inflammatory factors such as IL-6, which recruit inflammatory cells and results in cell necrosis, loss of pulp vitality, and complications in functional recovery^6,7. Understanding inflammation and the associated cell death is, therefore, crucial for advancements in the preservation of vital pulp. There are a number of experiments that have been conducted to explore the molecular biology of the inflame pulp in vivo or in vitro^8,9. Though in vitro experiments like 2D or 3D cell cultures have been developed for years and are becoming mature and widely used to test reactions of pulp cells to inflammatory factors, these experiments cannot reflect the interaction between pulp tissue and the systemic immune system¹⁰. If the phenomenon being studied is derived from cells of other tissue origin like immune, vascular and nervous system, then pure pulp cell culture will lead to a dead end. Therefore, in vivo experiments are very necessary and referential.

Mice have increasingly become a common choice in inflammation research in vivo due to their cost-effectiveness, high fertility, and vitality. However, a comprehensive protocol for mice pulpitis model is currently absent, which can serve as a reference. The small size of mice and their sensitivity to stimulation pose significant challenges during experimental procedures. Observing the minuscule teeth concealed deep within the mouse mouth often necessitates the use of a cantilever microscope, notwithstanding the more common presence of desktop microscopes in laboratories. The absence of a mouth opener requires assistance from others. To address this, the group has devised a mouth-gag using readily available materials which aims to provide a standardized and reproducible protocol for constructing the mice pulpitis model. This article details the procedure, covering preoperative preparation, immobilization, pulp exposure surgery, and sample collection on C57 mice. This protocol recommends the use of the mouth-gag, providing information on its structure, production, and application to facilitate other researchers in replicating the procedure.

The experimental procedures in this study were approved by the ethical committee of the West China School of Stomatology, Sichuan University (WCHSIRB-D-2021-125). Adult C57BL/6 mice were obtained from Gempharmatech Experimental Animals Company, Chengdu, China. The whole crown of the maxillary first molar erupts 21 days after birth. Mice for surgery should be older than 21 days with normal vitality¹¹. Here, 6 to 8-week-old mice were used for modeling. Figure 1 is a flow diagram showing the protocol used.

1. Preoperative preparation (Figure 2)

1. Obtain the following instruments: Stereoscopic microscope, fixing plate, medical tape, mouth-gag, minimally invasive dental burr with a diameter of 0.6 mm, dental high-speed dental handpiece, 8# C+ file, heating pad, 1 mL syringe, sterile cotton ball, eye forceps.
2. Obtain the following drugs: anesthesia mix, veterinary ointment.

2. Preparation of the mouth-gag

1. Weigh and anesthetize the mouse by intraperitoneal injection of anesthesia mix solution (10% ketamine hydrochloride + 5% xylazine + 85% sterile isotonic saline) at 0.007 mL/g of body weight and confirm proper anesthetization through the toe pinch method. Apply ophthalmic lubricating ointment on the eyes to prevent eyes injury because of desiccation during operation.
   NOTE: Medical hats, masks, gloves and overalls and other basic protection are necessary. Make sure both surgery environment and mouse chamber are clean and safe. A heat pad for thermal support throughout the procedure is necessary.
2. Prepare the mouth-gag as described below (Figure 3).
   1. Obtain the following materials: Orthodontic arch wire with diameter of 8 µm, young loop bending plier, heavy wire cutter, marker pen, rubber cap with a length of 3 mm and a cross section diameter of 1 mm.
   2. First, straighten the arch wire with left hand for fixation and thumb, index finger of the right-hand bend slightly against the arc of the wire. Repeat this action several times, it will facilitate the bending to the correct three-dimensional Angle.
   3. Use the Yong loop bending plier to bend the top edge (Figure 3G, a-i) of the trapezoid (Figure 3C, a-l-k-b) about 8 mm long at the midpoint of the bow wire. Make sure that point a (Figure 3) is on the edge of the plier beak.
   4. Hold the pliers with left hand, clamp the free end of the bow wire with right thumb and forefinger, and bend the bow wire from point a to make an angle of about 120°. Duplicate the previous action at point i (Figure 3G). Check whether the arch wire is on one plane by putting it on a horizontal table without prying.
   5. Leave about 9 mm of length on each side (Figure 3D, a-b, l-k) and bend the free-end to a 75° angle by using the same skill as step 2.2.4, while ensuring that each edge is on one plane. Bend this acute angle with the tip of the plier beak.
   6. Find point c about 5 mm from point b. Follow the same skill to bend a 105° angle on point c. Bend another 105° angle at point d 5mm from point c. Leave about 4.5 mm from point d and find point e. Bend the free end at point e to form an angle of about 100 -105° (Figure 3E).
      NOTE: The 6-8-week-old C57 mice we used were about 20 g. The spacing of 5 mm could not only jam the upper and lower jaws of the mice without moving, but also would not press the skin of the mice and cause discomfort. If other species or ages of mice are used, please adjust the length of c-d and i-h parts according to the actual situation (Figure 3E,G).
   7. Bend an extra tongue depressor for mandibular part (Figure 3G, j-i-h-g).
   8. Duplicate bending steps of a-b-c part on l-k-j part. Clamp i-k and k-j parts at the same time and bend the free end at point j to make it vertical to i-k-j plane. Clamp point i which is 5 mm from point j, bend the free end to make it parallel to both i-k-j plane and c-d part (Figure 3H).
   9. Leave 5 mm length from point i, at point h, bend the arch vertical to i-h part and parallel to j-i-h plane. Find point g at 5mm from point h. Clamp j-i-h-g plane and bend the free end symmetrical to k-j part. Then free end after point f should be symmetrical to point e-free end (Figure 3H).
   10. Put rubber caps on the free end (Figure 3F).

3. Immobilization

1. Fix the mouse supine on the fixation plate with limbs secured by skin tape. Compress the free ends of mouth-gag with thumb and index finger.
2. Fix the mouse front incisors in the trapezoidal groove of two arms. Ensure that the arm with tongue depressor is for mandibula. Adjust the mouth-gag to make sure mouse's tongue is immobilized but not ischemic.

4. Tooth assessment

1. Ensure the maxillary first molar for surgery should be free of dental caries, trauma and odontogenesis. Ensure there is no redness, swelling, or fistula on the surrounding gingiva. Ensure the opposite teeth are healthy and available to act as the healthy control group.

5. Pulp exposure

1. Use dental burr to drill at the occlusal side of maxillary first molar at a speed of 20,000 rpm. Ensure that the enamel is removed. Keep the operation with dental handpiece only in shallow layer of dentine to prevent excessive thermal stimulation on dental pulp tissue¹².
2. At the same time, prevent overheating by using a syringe to drop normal saline on the tooth every 3 min during the operation.
3. Put an 8# or 10# C+ file on the lowest position of the drilled pit and pierce through the last dentine to expose the pulp chamber. It will be an obvious feeling of falling when local dentine is thoroughly removed. Do not prob too deep, or dental pulp tissue might be brought out of pulp chamber.
4. Clean the fragments around the tooth. Take off the mouth-gag; the surgery is finished. Use the opposite maxillary first molar as the control without operation.

6. Post-operative care

1. After surgery, administer carprofen (5 mg/kg) subcutaneously and place the mouse on the chemothermal heating pad in a prone position until recovery from anesthesia. Feed the mice and provide drinking water. The recovery process should be supervised. No other animals should be in the same chamber until the mouse is fully recovered.

7. Sample collection and storage

1. Euthanize the mouse with cervical dislocation under deep anesthetic condition 24 h after operation or any other time point as per experiment⁹. Cut the skeleton muscles attached to the maxilla and zygoma with ophthalmic scissors. Remove skeleton, frontal bone and soft tissue and take out the lamina gnathostegite with maxillary molars.
   NOTE: According to He et al., it's commended that pulpitis sample should be collected less than 72 h after surgery to avoid extensive necrosis in dental pulp tissue¹³.
2. Sagittally divide the gnathostegite in half and store the tissue in 4% paraformaldehyde in PBS, pH 7.4, at 4 °C for 24h-fixation.

8. Histological analysis

1. Wash the tissue with phosphate buffered saline (PBS) and decalcify them in daily-changed decalcification solution of 5% EDTA and 4% sucrose in PBS, pH 7.4, at 4 °C for 2-4 weeks¹⁰.
2. Embed the 1/2 gnathostegite in paraffin and make sure the sagittal face without teeth is at the bottom of tissue cassettes.
3. Cut the paraffin block to 5 µm thick slices with a paraffin microtome. Adjust the angle of the paraffin block according to the proximal, distal, upper as well as lower relationship observed under the microscope to ensure the complete crown pulp and perforation of the first molar can be cut.

The procedure described above was performed on the right maxillary first molar of 3, 6-8 weeks old C57BL/6 mice, while the left maxillary first molars were preserved as control. Histology and immunofluorescence results from blank control, 12 h pulpitis and 24 h pulpitis samples were utilized for demonstration.

Following protocol of CT analysis from Goldman et al.¹⁵, pulp exposure was confirmed through micro-CT and reconstruction modeling in Figure 4 A-C. Sagittal slices of the maxillary first molars, both from the control and surgery sides, underwent HE staining (Figure 5). Pulp tissue necrosis and cell morphology disintegration at the wound margin were shown. The necrosis was mainly concentrated in the pulp tissue near the perforation, and the shape of the pulp tissue on the unopened side was normal. At 24 h, most of the pulp tissues, including the root pulp, were morphologically intact. (Figure 5).

The expression of IL-6¹⁴ was low in control, and a small amount of IL-6 could be observed around the wound at 12 h, while the expression of IL-6 was significantly increased at 24 h. Moreover, the expression of IL-6 was mainly concentrated in the wound margin and the middle pulp horn (Figure 6). In Figure 6 D,E, number of IL-6+ dental pulp cells and ratio of IL-6+ cells to total dental pulp cells increases over time in three time points. It can be considered that the area gradually develops inflammation and aggravates after pulp exposure.

We invited 5 colleagues who have never performed surgery on maxillary teeth of C57 mice to calculate their time needed to immobilize the upper and lower jaw of mouse (Stage 1) and expose mouse maxillary first molar (Stage 2) following the traditional procedure with two rubber band referring to the protocol published by Goldman et al. or our protocol with the mouth-gag¹⁵. Time for placing the drill correctly on the mouse maxillary first molar under microscope was also calculated for analysis. The results in Figure 3 J,K suggested that the time of mouth fixation and finding mice maxillary first molar with the mouth-gag were significantly shortened compared with the traditional way(P<0.05). The use of mouth-gag can improve operation efficiency and reduce operation difficulty.

Figure 1: Flow diagram of pulp exposure procedure. (A) After fixation with mouth-gag, the maxillary first molar of the mouse should be fully exposed under the microscope. (B) Use a high-speed dental handpiece with minimally invasive dental burr to remove occlusal enamel and superficial dentin of the first molar but be careful not to penetrate the dentin directly to avoid excessive influence on the pulp caused by overheating. (C) Use 8# C+ file to penetrate the remaining dentin and expose the pulp. (D) Samples were collected 24 h after surgery. HE and immunofluorescence staining proved that a pulpitis model could be established at this time point. Please click here to view a larger version of this figure.

Figure 2: Equipment for pulp exposure procedure. (A) Stereoscopic microscope and motor of dental high-speed dental handpiece. (B) The 8# C+ file, minimally invasive dental burr, mouth-gag, tweezers, and dental high-speed dental handpiece. Please click here to view a larger version of this figure.

Figure 3: Making the mouth-gag. (A) Bend the wire to make two working arms of the mouth-gag. (B-C) Steps of bending tongue spatula for mandible. (D-E) No tongue spatula for maxilla. (F, H) Rubber caps or bent ends are needed to protect injury from being pricked. (G) Three views of the mouth-gag for reference. (I) Clinical use of mouth-gag. (J) The average time for beginners to fix the upper and lower jaws (Stage 1) using traditional fixation method and mouth-gag, respectively. (K) The average time for beginners to clearly find the maxillary first molar (Stage 2) using traditional fixation method and mouth-gag, respectively. Please click here to view a larger version of this figure.

Figure 4: Micro-CT analysis of operated fist molar with perforation marked by red dotted line. (A) Sagittal plane of tooth, ensure that no perforation on pulp-chamber floor exists. (B) Coronal plane. Corresponding to the perforation in (A), complete penetration of the enamel dentin circled with red dotted line can be observed. (C) Occlusal plane of CT reconstruction model. The location of perforation circled with red dotted line is confirmed in a more intuitive way and the floor of the pulp chamber can be observed through perforation. (D-G) Intraoperative photos of the treatment. (D) Check the maxillary first molar of the mouse to ensure there are no cavities or malformation. (E) Using a minimally invasive burr to remove enamel and shallow dentin. As can be seen in the image, a pit (circled with dashed white line) can be observed on the occlusal face of the tooth without perforation. (F) Using 8# C+ file to penetrate the remaining dentin, the file can get stuck in the tooth without hand support. (G) When the file is removed, the tooth has a pink perforation, indicating successful pulp exposure. Please click here to view a larger version of this figure.

Figure 5: Hematoxylin-eosin staining. (A) Holistic view of untreated first molar for control. (A-1,2,3) High-magnification figures corresponding to 1, 2, 3 in panel (A). The shape of dental pulp tissue at 3 positions were intact, and odontoblasts were in an orderly arrangement. (B) Holistic view of teeth 12 h after surgery. (B-1,2,3) High-magnification figures corresponding to 1, 2, 3 in panel (B). The shape of dental pulp tissue at position 1 and 2 were generally intact. Necrosis could be observed near the perforation. (C) Holistic view of teeth 24 h after surgery. (C-1,2,3) High-magnification figures corresponding to 1, 2, 3 in panel (C). Necrosis extends from a single pulp horn to nearby pulp tissue. But most of the pulp tissues, including the root pulp are morphologically intact. Please click here to view a larger version of this figure.

Figure 6: Immunofluorescence staining. (A) Holistic view of untreated first molar for control. (A-1,2,3) High-magnification figures corresponding to 1, 2, 3 in panel (A). Nearly no expression of IL-6 could be observed. (B) Holistic view of teeth 12 h after surgery. (B-1, 2, 3) High-magnification figures corresponding to 1, 2, 3 in panel (B). Increasing IL-6 concentrated in tissue near the perforation shown in B-3. No obvious changes were observed in B-1,2. (C) Holistic view of teeth 24 h after surgery. (C-1, 2, 3) High-magnification figures corresponding to 1, 2, 3 in panel (C). Expression of IL-6 was significantly increased in C-2,3. (D) Total amount of IL-6⁺ dental pulp cells in control, 12 h pulp exposure, 24 h pulp exposure immunofluorescence staining results. (E) Ratio of IL-6+ cells to total amount of dental pulp cells in control, 12 h pulp exposure, 24 h pulp exposure immunofluorescence staining results. Please click here to view a larger version of this figure.

As the solitary soft tissue within teeth, dental pulp fulfills a crucial role in maintaining bioactivity of tooth but remains highly sensitive. The preservation of this vital pulp has become the preferred initial approach in recent endodontic treatments, necessitating a comprehensive understanding of the inflammatory mechanisms of dental pulp¹⁶. The spatiotemporal fluctuation of the inflammatory microenvironment and interactions between resident cell types in pulpitis complicates its investigation through in vitro studies¹¹. Instead, in vivo studies offer benefits by replicating the physiological environment found in humans. Utilizing experimental mice, specifically those with overexpressed or knocked down genes, provides an effective instrument for hypothesis validation. The frequently used C57 mice in laboratories, however, pose challenges due to their small size and lack of coordination, rendering the application of stimuli to their teeth problematic¹⁷. To address this issue, a comprehensive explanation of novel mouth-gag is needed to assist researchers in performing procedures within the oral cavities of mice. Moreover, this article outlines the protocol for establishing a pulpitis model through pulp exposure on the mice's first molar using the mouth-gag, thereby offering a valuable guide for subsequent research.

After numerous iterations, a scalable mouth-gag that is straightforward to construct was successfully designed. The dimensions and a three-view schematic of the mouth-gag are provided in Figure 3. The protocol significantly simplified the wire bending process by adopting a trapezoidal design in lieu of an arc. The mouth-gag uses a 0.8 mm diameter orthodontic arch wire, which balances the need to prevent slippage from the mouse's mouth and provide sufficient opening force. Moreover, the orthodontic arch wire's elasticity safeguards the temporomandibular joint of the mice. The mouth-gag is compact, resistant to corrosion, and can be stored in a 50 mL centrifuge tube with alcohol for repeated use. Despite a mouse's biting pressure, the mouth-gag remains stable in the mouth, allowing researchers to operate unassisted under a microscope. It should be noted that the mouth-gag's size can be adjusted to fit different mouse body sizes. If the mouth is stretched beyond the mouse's limit, the mouth-gag should be immediately removed to avoid injury to the temporomandibular joint (TMD) or the maxillofacial muscles.

He et al.'s report indicates that necrosis can be detected 24 h post pulp exposure, with the majority of pulp tissue becoming necrotic after 72 h¹⁸. Hence, it is essential to collect the pulp tissue within this 72 h window to avoid invalid experimental conclusions due to excessive dead cells. When inserting the C+ file into the pulp chamber, repeated rotation and deep pushing must be avoided to prevent excessive damage to the pulp tissue. If heavy bleeding occurs during the procedure, it is recommended to gently remove the blood using a small cotton ball to prevent coughing. The operation should only be conducted on one side of the mouse maxilla due to the potential negative impact on model accuracy from simultaneous modeling on both sides, which may cause dietary intake complications. Post-surgery, it is advised not to administer anti-inflammatory drugs or antibiotics to the mice.

The mouth-gag, while effective in maintaining the mouse's mouth open, has limitations concerning the surgical site. The mandibular posterior teeth, protected by the tongue pressed with a tongue depressor, are often not clearly visible. Therefore, the procedure is only suitable for operations on the maxilla teeth or mandibular anterior teeth. If the surgery exceeds 20 min, it is recommended to give the mouse a break every 10 min since the mouth-gag's stability depends on antagonizing the mice's occlusal force. C57 mice, chosen for their rapid reproduction and availability, are sensitive to various stimuli; hence, a slight overdose of drugs or stimuli can be lethal. Furthermore, their teeth's small size requires a higher skill level for tissue slicing.

In conclusion, pulp inflammation and necrosis represent pressing challenges in pulp regeneration. This study offers a comprehensive demonstration of creating a pulpitis model in mice, with immunofluorescence results verifying the inflammatory factors. This article proposes a novel, convenient mouth-gag design, which provides the operator with unobstructed sight by keeping the mouse's mouth open without tongue interference. However, operations on the mandibular posterior teeth remain a challenge. Considering the advantages of using experimental mice, endodontic modeling in mice holds significant promise, and it is planned to anticipate further reductions in the technical threshold.