An Animal Model of Pulpitis Based on an Outbred Mice Strain and Its Histological Analysis

Summary

Here, we present a protocol for creating a pulpitis model in NMRI (Naval Medical Research Institute) mice. Additionally, steps for making a jaw retraction clip and preparing slides for histology from the animal jaw are described. By following a standard protocol, variations in methodology and results can be avoided.

Abstract

The Naval Medical Research Institute (NMRI) mice have not been reported previously for creating a dental pulp inflammation model. In this protocol, we present an animal model employing NMRI mice and provide detailed histological procedures for the pulpitis model to ensure reproducibility. We included 70 NMRI mice that were randomly assigned to two groups: Group A (n=35), in which pulpitis was induced by the pulp exposure alone, and Group B (n=35), which received pulp exposure followed by Zymosan application. After the pulp exposure of the right maxillary molar, animals were sacrificed at designated time intervals (0, 6, 9, 12, 24, 48, and 72 h). The maxillary jaw was obtained for histology and quantitative histomorphological analysis. A time-dependent gradual increase in inflammation was observed in histological analysis for both groups. Both animal models allowed the study of inflammation up to 72 h. The technique presented in this paper is reproducible and will help standardize the histological study of pulpitis.

Introduction

Various animal models have been used to study pulpitis^1,2,3,4. Among these, mice models based on C57BL/6 and BALB/c are exclusively reported in pulpitis research^5,6,7,8. Both are inbred and have strict requirements for breeding^7,9. The NMRI mice used in the current protocol are an outbred strain, offering multiple advantages over the inbred strains. It has low set-up and maintenance costs. High reproduction, better immunity, and low mortality result in earlier and greater turnover of colonies¹⁰. Therefore, it is logistically and financially more feasible¹¹. Moreover, our results indicate that the disease profile of pulp inflammation produced in this model is similar to the previously reported models on inbred mice^5,6,8. Moreover, NMRI mice can withstand exposure of pulp to Zymosan for up to 72 h, developing an observable disease profile. This is a viable alternative to other inbred strains for studying pulpitis¹².

Zymosan is an extract of the cell wall of Saccharomyces cerevisiae, and it is mainly made up of chitin, mannans, mannoproteins, and ß-glucans¹³. Zymosan and Candida albicans share a similar cell wall structure and are detected by host innate immunity via a C-type lectin, pathogen recognition receptor, Dectin-1¹⁴. Dectin-1 is mainly expressed by dendritic cells and macrophages. It detects ß-glucans in fungal pathogens and uses the Syk/CARD9 pathway for the formation of inflammatory mediators¹⁵. Therefore, Zymosan is a viable alternative for studying the effect of exposure by C. albicans on pulp disease instead of using the organism¹⁶. This is supported by our previously published results, which show an early detection of Dectin-1 transcripts in Zymosan-exposed pulp. Similarly, Dectin-1 expressing CD68+ve macrophages were observed as well¹².

Histological techniques described in previous publications are often insufficient for novice researchers to reproduce with confidence. Consequently, extensive trial and error occur, leading to unnecessary animal loss during optimization of histological procedures. The present protocol standardizes the slide preparation technique and has been successfully applied to rats as well¹⁷. Here, all essential steps of the pulpitis model are described in detail, including a reproducible method for jaw retraction, induction of pulpitis by pulp exposure, induction of pulp inflammation using Zymosan, and subsequent histological analysis.

Protocol

The current animal study was designed in accordance with the ARRIVE guidelines. The Institutional Review Board for Animal Research and Ethics of Dow University of Health Sciences approved this animal study (Letter no. AR.IRB-06/DUHS/Approval/2016/05). The protocol included a total of 70 NMRI mice. The animals were obtained from the 'Central Animal House Facility of Dow Institute for Advanced Biological and Animal Research. The sample size of this protocol is based on a previous study⁸.

1. Animals

NOTE: The animals selected for the study weighed 26-30 g and were kept in standard cages under the following conditions: a temperature of ±21 °C, relative humidity of ±55%, and a 12 h light/dark ratio with freely available filtered water and standard feed (normal chow diet).There was regular monitoring for any sign of aggression, discomfort, or irritation. Diclofenac sodium (200 µL subcutaneously, 25 mg/mL) was used in the event of pain or discomfort. They were allowed to acclimatize for two weeks prior to any experimentation. Ten animals each were kept in appropriately labelled cages. For animal identification, a marker was placed on its tail.

1. Divide the animals (n=70) randomly into two groups: Group A (n=35), Group B (n=35).
2. In Group A, induce pulpitis by the exposure of the pulp only (See step 4.1.1 to step 4.1.9).
3. For Group B animal, apply Zymosan solution over exposed pulp (See step 4.2.1 to step 4.2.3).

2. Zymosan solution

1. Follow a previously published method for preparation of Zymosan solution as described below¹⁸.
2. Use Phosphate-buffered saline (PBS) to make a solution of 1 mg/mL from 1 mL of Zymosan.
3. Autoclave the solution at 121 °C for 15 min.
4. Allow the solution to cool to room temperature for 1 h.
5. Centrifuge the solution at 4 °C and 300 x g for 10 min.
6. Discard the supernatant and wash the pellet three times by re-centrifugation with ice-cold sterile PBS.
7. Store the solution at 4 °C until further use.
   NOTE: The solution must be used within 30 days, after which it may be discarded.

3. Jaw retraction clip

1. Use 0.7 mm stainless steel orthodontic wire to make a jaw retraction clip.
2. Take a wire of 100 mm length and make a loop of 5 mm diameter in its center.
3. Turn both parts of the wire 5 mm from the loop and 15 mm in total at right angles without sharp bends.
4. At a distance of 25 mm from the turn, make a notch and turn the remaining end outwards. Use Figure 1A for guidance.

4. Animal model

1. Group A (n = 35)
   1. Anesthetize animals by an intraperitoneal injection of 0.1 mL/20 g of Ketamine-Xylazine cocktail (87.5 mg/kg Ketamine + 12.5 mg/kg Xylazine).
   2. Use a 10x inclined long arm Stereomicroscope for all procedures (Figure 1B). Set the inclined arm in such a way that there is a direct view of the animal's retracted maxillary teeth. (Figure 1C)
   3. Squeeze the clip and place the notched part between the maxillary and mandibular incisors on either side of the clip.
   4. After releasing the clip, secure its ends with the help of adhesive tape (Figure 1C).
   5. Use a dental tweezer to expand the cheeks of the animal with your non-working hand. (Figure 1D,E).
      NOTE: Avoid excessive pressure. The injury from a tweezer can cause fatal bleeding and death of the animal.
   6. Use a #1/4 round carbide bur with an E-type slow-speed handpiece in an electric micro-motor under 10x magnification to half of its depth in the center of the right maxillary first molar to make a class 1 cavity. Shadow of pulp may be visible at this moment (Figure 1D,E).
   7. Run the bur gently at slow speeds in a sweeping motion. Use an electric motor to control the RPM.
   8. Use a K-type number 15 endodontic file to expose the pulp (Figure 2B).
      NOTE: The use of a #15 endodontic hand instrument must be extremely gentle, else a furcation perforation will ensue, which will only be evident once histological slides are prepared. The diameter of the #1/4 round carbide bur is 0.25 mm, and a #15K type file is 0.15 mm wide. Therefore, 0.15 mm will be the exposure size into the pulp (Figure 2C).
   9. Leave the exposed pulp open for induction of pulpitis
2. Group B (n = 35)
   1. Repeat all the steps given at step 4.1 with the addition of the following.
   2. Use a micropipette (0.2-2 µL) to place 0.2 µL of previously prepared Zymosan solution over the exposed pulp.
      NOTE: Care must be taken while dispensing the Zymosan solution. Inadvertent spilling of the solution into the oral cavity can cause death of the animal. Therefore, gauze is used to drape the tooth so that excess Zymosan solution gets absorbed by gauze (Figure 2D).
   3. Leave the treated pulp open for induction of pulpitis by Zymosan solution.

5. Sample preparation

1. Sacrifice five animals from each group at 0, 6, 9, 12, 24, 48, and 72 h by Carbon dioxide inhalation¹⁹.
2. Use a 7.25 L euthanasia chamber.
3. Do not pre-charge the chamber with CO₂.
4. Place the animals directly into the chamber.
5. Add 100% CO₂ at a fill rate between 30 and 70 % of the chamber volume per minute.
6. Keep the flow rate of CO₂ at 5 L/min.
7. Warm up the CO₂ and leave it to mix with the air present in the chamber.
8. Observe the animals for signs of unconsciousness within 23 minutes.
9. Monitor the animals for cessation of respiration and eyes to turn grey.
10. Keep the flow of CO₂ for at least one minute after the cessation of respiration.
11. Confirm the death of animals before removing them from the chamber. Make sure that there is no respiration and the eyes appear pale.
12. Separate the heads of animals immediately after euthanasia.
13. Use the untreated maxillary first molar of the left side of the animal sacrificed at 0 h as a control.
14. Use 10% buffered formalin in appropriately labelled 15 mL conical tubes to store the maxillary jaws for 2 days.

6. Histology

1. Decalcify the fixed jaws in 10% formic acid for 3 days.
   NOTE: After the third day, check whether a surgical blade can cut the jaw. If it is still not soft enough, immerse it again in 10% formic acid and check daily until it can be cut with a surgical blade.
2. Cut the jaw with a no. 22 blade. Make the cut parallel to the occlusal plane, along the sagittal plane, keeping the blade immediately on the palatal side of the palatal cusps of maxillary molars (Figure 3A).
3. Mark the occlusal surface of all teeth with India Ink.
   NOTE: This marking will help recognize the occlusal surface during sectioning (Figure 3B).
4. Embed the sectioned jaw in paraffin wax in a stainless-steel base mold casing by lying it flat against the base of the mold with the cut surface facing the base.
   NOTE: Embedding in this way ensures that the cut surface enters the microtome first, the sample orientation is correct, and that the sagittal sections are produced as shown in Figure 3E.
5. Remove the embedding cassette containing the embedded jaw from the base mold and place it in a microtome.
6. Make 3 µm thick sagittal sections and place them on a clean, charged microscope slide (Figure 3C).
7. Observe unstained slides under a compound microscope.
   NOTE: Keep observing the cut sections until the cavity is visible on the occlusal surface. If the sections are correctly made along the sagittal plane along the long axis of the maxillary first molar, one will observe the entire pulp space showing either one or two roots and pulp chamber (Figure 3E). From this point onwards, one can theoretically make 50 slides from one section since the size of exposure is 0.15 mm. (3 µm = 0.003 mm, 0.003 x 50 = 0.15)
8. Perform standard steps of Hematoxylin and Eosin staining procedures for the selected slides showing either one or two roots, along with the entire pulp chamber.
9. Observe the stained slides under a compound microscope.
10. Select five best slides per time interval that will allow observation of the maximum pulp space.

7. Statistical analysis

1. Perform statistical analysis using standard statistical software.
   NOTE: For quantitative histomorphological analysis, the observed slides from each time interval were scored according to a previously published criterion (Table 1)⁸. Kruskal-Wallis H test with Bonferroni correction at p=0.0018 was used to compare scores of different time intervals. The Mann-Whitney U test was used at p ≤ 0.05 to compare Groups A and B. All comparisons were carried out using standard statistical software.

Results

In this protocol, we present a detailed step-by-step procedure for two animal models of dental pulp inflammation in NMRI mice, along with their corresponding histological methods. Use of appropriate equipment is essential for the success of the protocol. In this regard, the 10x inclined long-arm stereomicroscope is the most crucial piece of equipment. The class 1 cavity preparation was performed using an electric micromotor at approximately one-third of its maximum speed (35,000 rpm), making the procedure manageable and ...

Discussion

The current protocol presents the development of a type 3 dental injury model (irreversible pulpitis) in NMRI mice for the first time in reported literature. It also details the histological analysis of Zymosan-induced pulp inflammation and provides a reproducible method for preparing histological slides from animal jaws.

Following this protocol, a type 3 dental injury model was produced in NMRI mice. Previous research work was performed on rats, rabbits, and dogs^1...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Base mold	Histo-line Laboratories, Italy	R4166C2:D23	For embedding the sample in paraffin
Carbide bur #1/4 round	Midwest, Wichita Falls, Texas	389101
Compound microscope Motic BA310	Motic Inc. Co. Ltd, Hongkong	https://moticmicroscopes.com/products/ba310e
Diclofenac sodium 25mg/mL	Indus Pharma (DYCLO)	https://sahar.ph/product/dyclo-diclofenac-sodium-25-mg-ml-im-iv/	200µL subcutaneously
E type slow speed hand piece Rose201-M4	Being Foshan, China	https://www.beingfoshan.com/en/h-pd-353.html
Embedding cassette	Histo-line Laboratories, Italy	R4004
Euthanasia Chamber	ConductScience, Illinois USA	CS-EUTH2A	7.25 Liters Capacity, For Mice and Rats [Up to 500g]
Falcon tubes 15 ml	Nest	188271
Formalin	Sigma-Aldrich, St. Louis, Missouri, United States	158127	buffered
Formic acid	BDH Laboratory Supplies Poole, England	28428	10%
Homogenizer	Omni Mixer Homogenizer, Omni International, Georgia USA	https://omni-inc.com/high-shear-lab-homogenizers/omni-th-0-2ml-100ml/omni-tissue-homogenizer-th
India Ink	Sigma-Aldrich, St. Louis, Missouri, United States	198285
K type number 15 endodontic file	Mani, Japan	https://www.mani.co.jp/en/product/dental_01.html
Ketamine-Xylazine cocktail	Brookes Pharma Pvt Ltd / Mylab (Pvt.) Ltd.		87.5mg/kg Ketamine + 12.5mg/kg Xylazine
Microtome HM 315 R	Microm, Germany	901100
Micro-motor Marathon-III	BT-Marathon, Saeyang Microtech Co.,Ltd. Korea	https://www.saeyang.com/theme/basic/sub/product/view.php?it_id=1752202111	Use at 1/3 of maximum speed
Micropipette 0.2-2 µL	Gilson, France	https://www.gilson.com/default/ pipetman-p2-0-2-2-micro-l-metal -ejector.html?srsltid=AfmBOoobE 0iSWY_902ZHGZ7AoJ6wzN5PI 0H4V4vkxHQgK1mq7Wg2kkbS	0.2-2 µL
Phosphate buffered saline	Sigma-Aldrich, St. Louis, Missouri, United States	P4417
Statistical Package for the Social Sciences	IBM, Armonk, New York	SPSS for Mac version 23.0
Stereomicroscope BMK-3 led	Meiji techno, Japan	https://meijitechno.com/product/bmk-3-led-stereo-microscope/
Surgical blade No. 22	Feather Surgical Blade, Feather Safety Razor Co Ltd., Osaka, Japan	FB. 12
Zymosan A	Sigma-Aldrich, St. Louis, Missouri, United States	Cat no Z4250	1mg/mL from 1 mL Zymosan