Research reported in this publication was supported by JSPS KAKENHI JP21K09915 (MO) and the National Institute of General Medical Sciences;&#160;T34GM145509 (MM) and the National Institute of Dental and&#160;Craniofacial Research; R01DE025255 and R21DE032156&#160;(XH); R01DE029709, R21DE028715 and R15DE027851&#160;(TK); R01DE027648 and K02DE029531 (MS).

All procedures described in this protocol have been performed in accordance with guidelines and regulations for the use of vertebrate animals approved by the Institutional Animal Care Use Committee (IACUC) at Augusta University and at Nova Southeastern University which is accredited by the Association for Assessment and Accreditation of Laboratory Animal Care International (AAALAC). Note that Dr. Suzuki was employed by Augusta University where the mouse dental fluorosis experiments were completed.
1. Extraction of mandibular incisors in a mouse dental fluorosis model
<ol>
	<li>Feed fluoride-free diets to C57BL/6 mice (5-week-old, male) from 1 week prior to fluoride until termination of fluoride treatment.</li>
	<li>Prepare fluoride water by adding NaF in distilled water followed by vacuum filtration using a 0.2 &#181;m filter. Give animals fluoride water as NaF (0 ppm and 125 ppm; N=5/group) ad libitum for 6 weeks. Replace fluoride water with a freshly prepared batch every 2 days.</li>
	<li>After 6 weeks of fluoride water treatment, euthanize animals with CO2 followed by decapitation.</li>
	<li>Extract the hemi mandibular with incisor from each mouse. To collect the hemi mandibular with incisor, cut the muscles around the mandibular jaw without applying excessive force.</li>
	<li>Place the hemi mandibular in PBS and keep it at 4 &#176;C until &#956;-CT analysis (optional). Separate the incisor from the mandibular using a scalpel (#15) and scissors without damaging or breaking the specimen.</li>
	<li>Wash the isolated incisor with PBS and perform dehydration by immersing it in increasing strength of alcohol (70% and 100% ethanol) for 2-3 h. 
	NOTE: If the tissue (e.g., pulp) is not sufficiently dehydrated, resin impregnation is likely to be inhibited and subsequent evaluation will likely be inadequate.</li>
	<li>After dehydration with ethanol, embed the incisor horizontally in resin. Continue to step 3.</li>
</ol>
2. Extraction of maxillary alveolar bones in a mouse ligature-induced periodontal bone resorption (L-PBR) model
<ol>
	<li>Administer 0.8 mL of ketamine (100 mg/mL) + 0.1 mL of Xylazine (100 mg/mL) + 9.1 mL of PBS intraperitoneally (i.p.) to mouse (C57BL/6, 8-12-week-old, male) as anesthetics. The dosage is 0.01 mL/g (weight). Apply ophthalmic ointment to both eyes to prevent dryness under anesthesia.</li>
	<li>Place the anesthetized mouse on a heating pad for 5-10 min. Assess responses to tail/toe pinches and the intactness of the ocular reflex. Confirm that the mouse is unresponsive to the noxious stimuli and the reflex is absent.</li>
	<li>Place the mouse on the treatment table and keep the mouth open by means of a ligature 5-0 silk suture tied to a magnetic post on the treatment table.</li>
	<li>Under a surgical microscope, wind the ligature (Braided silk suture 6-0) around one side of the maxillary second molar (single layer)&#160;using micro needle holders. Minimize individual differences in analysis by using one side as the treatment side and the other side as the control.</li>
	<li>Tie the ligature and make a knot on the palate side. After making a knot, cut the remaining ligature as short as&#160;possible so that the excessive ligature does not interfere with chewing or eating. This is important to ensure that the ligature will not loosen by chewing during the subsequent observation period. 
	NOTE: Do not leave the animal unattended until it has regained sufficient consciousness to maintain sternal recumbency. Do not return the animal that has undergone surgery to the company of other animals until fully recovered. Maintain sterile conditions during survival.</li>
	<li>Feed diet and water to mice ad libitum for 2 weeks. After 2 weeks of ligation, euthanize mice with CO2 followed by decapitation.</li>
	<li>Extract both side maxillae (ligature side and control side) with molars from each mouse. To collect maxillae with molars, cut the muscles and bone around the maxillary jaw using scissors without applying excessive force. Place each maxilla in PBS and keep it at 4 &#176;C until &#181;CT analysis (optional).</li>
	<li>Separate the alveolar bone with molars (1st to 3rd) from the maxilla using a scalpel (#15) and scissors without damaging or breaking the specimen.</li>
	<li>Wash the isolated alveolar bone with PBS then dehydrate and degrease by immersion in increasing strength of alcohol (70% and 100% ethanol) for 2-3 h. 
	&#8203;NOTE: If&#160;the tissue (e.g., pulp and bone) is not sufficiently dehydrated, resin impregnation is likely to be inhibited and subsequent evaluation will likely be inadequate.</li>
	<li>After dehydration with ethanol, embed the alveolar bone horizontally in resin. Continue to step 3.</li>
	<li>Optional: Perform &#181;CT evaluation before microhardness testing.
	<ol>
		<li>Before microhardness testing, perform nondestructive structural analysis (e.g., &#181;CT) using the same sample for microhardness testing as a complementary evaluation (Figure 1). Structural information (3D image, mineral density, volume) by &#181;CT could support to evaluate sample mechanical properties and quality that may affect microhardness results.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/66583/66583fig01.jpg" /> 
Figure 1: Representative &#956;CT images of enamel in control and fluoride-treated mice incisors. (A) Representative &#956;CT sagittal image of mandibular incisor. (B-D) &#956;CT coronal images of control incisor (NaF 0 ppm). (E-G) &#956;CT coronal images of incisor treated with NaF (125 ppm). Representative enamel mineral density (EMD) is shown (g/cm3). <a href="https://www.jove.com/files/ftp_upload/66583/66583fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
3. Embedding samples in resin
<ol>
	<li>Continue from step 1.7 (dental fluorosis model) or step 2.10 (L-PBR model).</li>
	<li>Coat the inside surface of the mounting cup (1 inch) with a thin layer of petrolatum. Mix resin (cold setting embedding resin) according to instructions. Pour the resin and hardener into the provided plastic cup in a 15:2 volume ratio and carefully mix with a wooden spatula for at least 2 min. Avoid air bubbles.</li>
	<li>Place dehydrated and degreased incisor (Figure 2A) or alveolar bone with molars (Figure 2B) oriented horizontally and parallel to the bottom of the mounting cup (1 specimen per cup).</li>
	<li>Pour the mixed resin (just enough resin, about 1.5 mL) into the mounting cup to completely cover the specimen. Avoid adding more resin than necessary, as excess resin will impede the polishing process (Figure 2C,D). Place the mounting cup containing specimen on a hot plate at 50 &#176;C for at least 8 h to promote resin polymerization. This procedure contributes to holding the specimen in a stable position. 
	NOTE: Depending on the sample size, adjust the amount of resin to completely cover the specimen. Do not fill too much resin, otherwise more time will be needed to remove superfluous resin.</li>
	<li>After curing, remove the resin containing the specimen from the mounting cup. Remove burrs and arrange the specimen&#39;s plane and the opposite side plane as parallel and flat using an advanced grinder-polisher with rough water-resistant abrasive paper (Grit 60/P60 and 120/P120) under water flooding. Keep the height of specimen to approximately 3 mm for incisor and alveolar bone (Figure 2E,F). 
	&#8203;NOTE: When the specimen is analyzed by SEM following the microhardness measurement, the thickness of the sample should be about 3 mm so that subsequent SEM observation will not be affected. Smaller samples are more difficult to manipulate with the grinder. For the samples intended for microhardness only, the specimen height can increase to about 10-20 mm.</li>
	<li>Trim the external shape to make a rectangular solid resin block and round corners (approximately, width 30 mm, length 10 mm for incisor (Figure 2G) and width 10 mm, length 5 mm, for alveolar bone (Figure 2H)) using a precision sectioning saw.</li>
	<li>Once the rough shape correction is complete, remove debris and particles from the resin block using an ultrasonic cleaner (about 1 min). Continue to step 4.</li>
</ol>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/66583/66583fig02.jpg" /> 
Figure 2: Flow of resin-embedding and polishing procedure. (A) Dehydrated and degreased incisor. (B) Dehydrated and degreased alveolar bone in L-PBR. (C, D) Incisors and alveolar bone immersed in resin. (E, F) By cutting off the resin, it is easier to polish the target tissue surface. (G, H) Resin corners rounded for the polishing process. Abbreviations:&#160;L-PBR =&#160;ligature-induced periodontal bone resorption.&#160;<a href="https://www.jove.com/files/ftp_upload/66583/66583fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
4. Polishing of specimens
NOTE: Polishing of specimens is done manually using waterproof abrasive papers (from rough to finer) on an advanced grinder-polisher under water flooding.
<ol>
	<li>Place a rough water-resistant abrasive paper (Grit 600/P1200) on the grinder. Place the trimmed and cleaned resin block (from step 3.7) on the rough water-resistant abrasive paper.</li>
	<li>While pouring water, hold the resin block and polish the specimen&#39;s evaluation surface on the grinder-polisher (Speed 1-10 x g). At this time, be careful to hold the resin block so that the evaluation surface is parallel to the ground. To keep the evaluation surface intact, check the surface with the naked eye or under&#160;a microscope. 
	NOTE: Note that the grinder rotates clockwise, and uniform pressure can lead to an unparallel surface. To obtain a parallel surface, keep the glider rotational speed constant and press the specimen carefully for a few seconds, and then rotate the specimen 180&#176; to press for the same amount of time. Rough abrasive paper may remove not only resin but also specimen.</li>
	<li>Change the abrasive paper to the Grit 800/P2400 and place the resin block on it. Repeat step 4.2.</li>
	<li>Remove debris and particles from the resin block using an ultrasonic cleaner (about 1 min). 
	NOTE: Before proceeding, using an ultrasonic cleaner to remove any surface debris to prevent clogging is recommended.</li>
	<li>Next, perform serial polishing using finer abrasive papers; polishing order is 12 &#181;m, 9 &#181;m, 3 &#181;m, 1 &#181;m and 0.3 &#181;m.</li>
	<li>Place a lapping film (12 &#181;m) on the grinder-polisher table without rotation and place the resin block on the lapping film. 
	NOTE: In this experiment, the grinder table is suitable to get a flat surface condition under water flooding. Alternatively, a large plane mirror (or similar one) that provides parallelism can also be used.</li>
	<li>Under water cooling, carefully polish the specimen&#39;s evaluation surface on the lapping film by hand. Move the sample vertically, horizontally, and diagonally for the same number of seconds under water injection with strokes of 2 to 3 cm (1 inch). When the polishing procedure is properly achieved, the resin specimen will stick&#160;to the lapping film.</li>
	<li>Remove debris and particles as in step 4.4. Change the abrasive paper to the next size according to the serial polishing order (from 12 &#181;m to 0.3 &#181;m) and place the resin block on it.</li>
	<li>While pouring water, hold the resin block and carefully polish the specimen&#39;s surface on the lapping film by hand. Remove debris and particles as in step 4.4.</li>
	<li>Repeat steps 4.5 - 4.8 to complete the final polishing (0.3 &#181;m). After completing the final polishing (0.3 &#181;m), the specimen should have a mirror finish surface (Figure 3A).</li>
	<li>Clean the surface of the specimen with ethanol (100%) to degrease and dehydrate and store resin blocks at room temperature until microhardness testing. During storage, avoid excessive moisture and dust. Continue to step 5.</li>
</ol>
5. Vickers microhardness test
NOTE: Indentation of a mirror finish surface specimen is done using a microhardness tester. Testing is performed with a load of 25 g for 10 s with a Vickers tip.
<ol>
	<li>Vickers microhardness test for incisors (dental fluorosis model) 
	NOTE: Enamel can be divided into three layers from outside (oral cavity side) to inside (pulp side); namely, the superficial layer, the middle layer, and the deep layer (dentin-enamel junction, DEJ) (Figure 3B)16. In this protocol, three enamel layers are tested.
	<ol>
		<li>Set loading force to 25 g and loading duration to 10 s. Place the resin block on the stage.</li>
		<li>Indent 6 points in each enamel layer (superficial, middle and DEJ) and dentine in each region (cervical, middle and tip; Figure 3B).</li>
		<li>Measure the length of the two diagonals (d1 and d2; Figure 3B) to calculate the Vickers microhardness value (HV; Figure 4).</li>
	</ol>
	</li>
	<li>Vickers microhardness test for alveolar bone (L-PBR model)
	<ol>
		<li>Set loading force to 25 g and loading duration to 10 s. Place the resin block on the stage.</li>
		<li>Indent 3-6 points in each mesial and distal side of alveolar bone from the alveolar crest. Indent alveolar bones between 1st and 2nd molar (white square), and 2nd and 3rd molar. 
		NOTE: In this protocol, 6 points in each mesial and distal side (total 12 points) were evaluated for the control (intact) bone, and 3 points in each side (total 6 points) were evaluated for L-PBR. The number of indentation points depends on the conditions of the lesion (e.g., too much bone loss limits the indentation area).</li>
	</ol>
	</li>
</ol>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/66583/66583fig03v2.jpg" /> 
Figure 3: Evaluation regions of microhardness in mandibular incisor. (A) Mirror finish surface sample containing mandibular incisor. (B) Indentations in each region; cervical, middle, and tip (NaF 0 ppm). (C) Three enamel layers; from DEJ, Inner, Middle, and Outer enamel. Abbreviations: D = dentin, E = enamel, DEJ = dentin enamel junction <a href="https://www.jove.com/files/ftp_upload/66583/66583fig03largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/66583/66583fig04.jpg" /> 
Figure 4: Vickers microhardness of enamel treated with or without NaF. The microhardness of dentin and three enamel layers were evaluated in each region, cervical, middle, and tip region. (A-C) Control and (D-F) NaF (125 ppm) treatment. Data are presented as mean &#177; SD. Significant differences were evaluated by one-way ANOVA with Tukey&#39;s post-hoc test. p values &#60; 0.05 were considered statistically significant. **p &#60; 0.005, ***p &#60; 0.0005, ****p &#60; 0.0001 <a href="https://www.jove.com/files/ftp_upload/66583/66583fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Microhardness Protocol for Evaluating Rodent Oral Disease

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The authors declare no conflict of interest.

Microhardness is performed to evaluate mechanical properties of hard tissues like tooth and bone. To date, divergent microhardness measurement methods have been reported. Most of the measurement information, especially sample preparations and the indentation sites are likely to be insufficient. This study focused on the microhardness protocol for enamel and alveolar bone in dental fluorosis and periodontal diseases models. To obtain consistent and accurate results, the critical steps in this protocol are orientation of t...

Hardness is one of the mechanical properties (e.g., elasticity, hardness, viscoelasticity, and fracture behavior) and is commonly used to characterize the ability to resist compression deformation and fracture of a local area of a material. The static indentation hardness test is the most used method, including Vickers hardness and Knoop hardness1. The Vickers hardness test is implemented by pressing a diamond indenter into the surface under a fixed testing load. The indenter is pyramid-shaped, with a square base and an angle of 136&#176; between opposite faces. The length of both diagonals formed on the test surface is measured, and the average is used to calculate the hardness, which is determined by the ratio F/A (where F is the force and A is the surface area of the indentation). The Vickers microhardness number (HV=F/A) is usually expressed in kilograms-force (kgf) per mm2 indentation, with 1 HV &#8776; 0.1891 F/d2 (N/mm2). The Knoop hardness also consists of a diamond square pyramid indenter formed by two unequal opposite angles. The Knoop hardness number (HK) equals the ratio of applied load to the projected contact area. Hardness tests are classified into micro-indentation (microhardness) tests and macro-indentation tests, depending on the force applied to the test material. Micro-indentation tests typically use loads in the range 0.01-2 N (about 1-203 gf); meanwhile, macro-indentation tests use over 10 N (10119 gf)1.
To evaluate features of dental hard tissues in oral diseases, including tooth and alveolar bone, micro-CT (&#181;CT) and scanning electron microscopy (SEM) are used for structural analysis. &#181;CT provides 3D imaging information (volume and mineral density)2, and SEM produces microstructure images (enamel prism and bone lacuna-canalicular)3. Complementarily to structural analysis by &#181;CT and SEM, microhardness is one of the informative parameters to evaluate how structural changes alter the mechanical properties of tooth and alveolar bone in oral diseases, e.g., enamel malformation and periodontal bone resorption. The Vickers microhardness value of human enamel (HV = 283-374) is about 4 to 5 times higher than that of dentin (HV = 53-63)4,5. In rodent dental fluorosis models, enamel microhardness significantly decreases in mouse incisors treated with fluoride (HV = 136) compared to control enamel (HV = 334)6,7. This suggests that fluorosed enamel is softer and weaker with lower mineral content and higher protein content than found in non-fluorosed enamel. Microhardness is used to evaluate bone mechanical properties. Several previous studies have examined the mechanical behavior of human bone from different anatomic sites, including long bone microhardness8,9,10. The mean microhardness of human fluorosed femurs showed a significant decrease (HV = 222.4) compared to non-fluorosed femurs (HV = 294.4)11. Despite being a useful parameter, there is a scarcity of literature describing microhardness (either Vickers12 or Knoop13,14) of alveolar bone in oral diseases.
To date, divergent microhardness measurement methods have been reported. Since microhardness values vary15 depending on sample preparation (polishing and flat surface) and indentation site, diverse protocols can cause discrepancies among studies. Standardization of the microhardness testing protocol is essential for consistent and accurate evaluation in oral disease models. In the present study, we demonstrate a standardized protocol for microhardness analysis in tooth and alveolar bone in mouse dental fluorosis model and periodontal bone resorption model.

<table><tbody><tr><td>Braided Silk Suture 6-0</td><td>Teleflex</td><td></td><td></td></tr><tr><td>Canica Small Animal Surgery System</td><td>Kent Scientific Corporation&amp;nbsp;</td><td>SURGI 5001</td><td></td></tr><tr><td>CarbiMet PSA 120/P120</td><td>Buehler</td><td>30080120</td><td></td></tr><tr><td>CarbiMet PSA 60/P60</td><td>Buehler</td><td>36080060</td><td></td></tr><tr><td>CarbiMet PSA 600/P1200</td><td>Buehler</td><td>36080600</td><td></td></tr><tr><td>Castroviejo Micro Needle hilder</td><td>F.S.T</td><td>12060-01</td><td></td></tr><tr><td>Epofix cold setting embeding Resin</td><td>Electron Microscopey Science</td><td>CAT-1237</td><td></td></tr><tr><td>Fisherbrand 112xx Series Advanced Ultrasonic Cleaner</td><td>Fisher Brand</td><td>FB11201</td><td></td></tr><tr><td>Fluoride-free Rodent diet</td><td>Bio Serv</td><td>F1515</td><td>&amp;nbsp;AIN-76A, 1/2&quot; Pellets</td></tr><tr><td>in-vivo microCT Skyscan 1176</td><td>Bruker</td><td></td><td></td></tr><tr><td>Isomet 1000 Precison saw</td><td>Buehler</td><td>MA112180</td><td></td></tr><tr><td>Lapping film 0.3&amp;micro;m</td><td>Maruto instrument co, LTD. Japan</td><td>26-4203</td><td>Alternative&amp;nbsp;&lt;br/&gt; A3-0.3 SHT, 3M USA</td></tr><tr><td>Lapping film 1&amp;micro;m</td><td>Maruto instrument co, LTD. Japan</td><td>26-4206</td><td>Alternative&lt;br/&gt; A3-1 SHT, 3M USA</td></tr><tr><td>Lapping film 12&amp;micro;m</td><td>Maruto instrument co, LTD. Japan</td><td>26-4211</td><td>Alternative&lt;br/&gt; A3-12 SHT, 3M USA</td></tr><tr><td>Lapping film 3&amp;micro;m</td><td>Maruto instrument co, LTD. Japan</td><td>26-4204</td><td>Alternative&lt;br/&gt; A3-3 SHT, 3M USA</td></tr><tr><td>Lapping film 9&amp;micro;m</td><td>Maruto instrument co, LTD. Japan</td><td>26-4201</td><td>Alternative&lt;br/&gt; A3-9 SHT, 3M USA</td></tr><tr><td>Leica wild microscope&amp;nbsp;</td><td>Leica</td><td>LEIC M690</td><td></td></tr><tr><td>Metaserv 2000 Variable speed Grinder polisher</td><td>Buehler</td><td>No: 557-MG1-1160</td><td></td></tr><tr><td>MicroCut PSA 1200/P2500</td><td>Buehler</td><td>36081200</td><td></td></tr><tr><td>MicroCut PSA P4000</td><td>Buehler</td><td>36084000</td><td></td></tr><tr><td>Microhardness tester, ALPHA-MHT-1000Z&amp;nbsp;</td><td>PACE Technologies</td><td></td><td></td></tr><tr><td>SamplKups &amp;nbsp;1 inch</td><td>Buehler</td><td>No: 209178</td><td></td></tr><tr><td>Sodium Fluoride</td><td>Fisher Scientific</td><td>S299-100</td><td></td></tr><tr><td>West cott Stitch Scissor</td><td>JEDMED</td><td>Cat. #25-1180</td><td></td></tr><tr><td>ZooMed Repti Thern Undertank heater (U.T.H)</td><td>Zoo Med Laboratories, Inc.</td><td>RH-4</td><td></td></tr></tbody></table>

microhardness measurements on tooth and alveolar bone in rodent oral disease models

The mechanical property, microhardness, is evaluated in dental enamel, dentin, and bone in oral disease models, including dental fluorosis and periodontitis. Micro-CT (&#181;CT) provides 3D imaging information (volume and mineral density) and scanning electron microscopy (SEM) produces microstructure images (enamel prism and bone lacuna-canalicular). Complementarily to structural analysis by &#181;CT and SEM, microhardness is one of the informative parameters to evaluate how structural changes alter mechanical properties. Despite being a useful parameter, studies on microhardness of alveolar bone in oral diseases are limited. To date, divergent microhardness measurement methods have been reported. Since microhardness values vary depending on the sample preparation (polishing and flat surface) and indentation sites, diverse protocols can cause discrepancies among studies. Standardization of the microhardness protocol is essential for consistent and accurate evaluation in oral disease models. In the present study, we demonstrate a standardized protocol for microhardness analysis in tooth and alveolar bone. Specimens used are as follows: for the dental fluorosis model, incisors were collected from mice treated with/without fluoride-containing water for 6 weeks; for ligature-induced periodontal bone resorption (L-PBR) model, alveolar bones with periodontal bone resorption were collected from mice ligated on the maxillary 2nd molar. At 2 weeks after the ligation, the maxilla was collected. Vickers hardness was analyzed in these specimens according to the standardized protocol. The protocol provides detailed materials and methods for resin embedding, serial polishing, and indentation sites for incisors and alveolar. To the best of our knowledge, this is the first standardized microhardness protocol to evaluate the mechanical properties of tooth and alveolar bone in rodent oral disease models.

Dental fluorosis model: Figure 1 shows representative &#956;CT images of incisors in control and fluoride-treated mice. In the control (Figure 1B-D), the cervical region showed lower enamel mineral density (EMD) of 1.188 g/cm3 (Figure 1B) compared to the middle (1.924 g/cm3) and tip (1.819 g/cm3; Figure 1C,D). In the fluoride-treated ena...

Watch this Scientific Journal Video about Author Spotlight: Establishing an Accurate Microhardness Testing Protocol for Craniofacial Tissues at JoVE.com

Author Spotlight: Establishing an Accurate Microhardness Testing Protocol for Craniofacial Tissues

Microhardness is a mechanical property and an informative parameter for evaluating hard tissue pathophysiology. Here, we demonstrate a standardized protocol (sample preparation, polishing, flat surface, and indentation sites) for microhardness analysis in tooth and alveolar bone in rodent oral disease models, namely, dental fluorosis, and ligature-induced periodontal bone resorption.

Microhardness Measurements on Tooth and Alveolar Bone in Rodent Oral Disease Models

The mechanical property, microhardness, is evaluated in dental enamel, dentin, and bone in oral disease models, including dental fluorosis and ...

microhardness-measurements-on-tooth-alveolar-bone-rodent-oral-disease

Research

JoVE Journal

Biology

597 Views.  Nova Southeastern University. Microhardness is a mechanical property and an informative parameter for evaluating hard tissue pathophysiology. Here, we demonstrate a standardized protocol (sample preparation, polishing, flat surface, and indentation sites) for microhardness analysis in tooth and alveolar bone in rodent oral disease models, namely, dental fluorosis, and ligature-induced periodontal bone resorption.

Video: Author Spotlight: Establishing an Accurate Microhardness Testing Protocol for Craniofacial Tissues

Meal Duration as a Measure of Orofacial Nociceptive Responses in Rodents

A lengthening in meal duration can be used to measure an increase in orofacial mechanical hyperalgesia having similarities to the guarding behavior of humans with orofacial pain. To measure meal duration unrestrained rats are continuously kept in sound attenuated, computerized feeding modules for days to weeks to record feeding behavior. These sound-attenuated chambers are equipped with chow pellet dispensers. The dispenser has a pellet trough with a photobeam placed at the bottom of the trough and when a rodent removes a pellet from the feeder trough this beam is no longer blocked, signaling the computer to drop another pellet. The computer records the date and time when the pellets were taken from the trough and from this data the experimenter can calculate the meal parameters. When calculating meal parameters a meal was defined based on previous work and was set at 10 min (in other words when the animal does not eat for 10 min that would be the end of the animal&#39;s meal) also the minimum meal size was set at 3 pellets. The meal duration, meal number, food intake, meal size and inter-meal interval can then be calculated by the software for any time period that the operator desires. Of the feeding parameters that can be calculated meal duration has been shown to be a continuous noninvasive biological marker of orofacial nociception in male rats and mice and female rats. Meal duration measurements are quantitative, require no training or animal manipulation, require cortical participation, and do not compete with other experimentally induced behaviors. These factors distinguish this assay from other operant or reflex methods for recording orofacial nociception.

A lengthening in meal duration represents orofacial nociceptive behavior in rodents similar to the guarding behavior of humans with orofacial pain. Eating is a behavior that requires no training or animal manipulation, requires cortical participation, and is not competing with other experimentally induced behaviors, distinguishing this assay from alternative reflex or operant measurements.

A lengthening in meal duration can be used to measure an increase in orofacial mechanical hyperalgesia having similarities to the guarding behavior of ...

Behavior

In situ Compressive Loading and Correlative Noninvasive Imaging of the Bone-periodontal Ligament-tooth Fibrous Joint

This study demonstrates a novel biomechanics testing protocol. The advantage of this protocol includes the use of an in situ loading device coupled to a high resolution X-ray microscope, thus enabling visualization of internal structural elements under simulated physiological loads and wet conditions. Experimental specimens will include intact bone-periodontal ligament (PDL)-tooth fibrous joints. Results will illustrate three important features of the protocol as they can be applied to organ level biomechanics: 1) reactionary force vs. displacement: tooth displacement within the alveolar socket and its reactionary response to loading, 2) three-dimensional (3D) spatial configuration and morphometrics: geometric relationship of the tooth with the alveolar socket, and 3) changes in readouts 1 and 2 due to a change in loading axis, i.e. from concentric to eccentric loads. Efficacy of the proposed protocol will be evaluated by coupling mechanical testing readouts to 3D morphometrics and overall biomechanics of the joint. In addition, this technique will emphasize on the need to equilibrate experimental conditions, specifically reactionary loads prior to acquiring tomograms of fibrous joints. It should be noted that the proposed protocol is limited to testing specimens under ex vivo conditions, and that use of contrast agents to visualize soft tissue mechanical response could lead to erroneous conclusions about tissue and organ-level biomechanics.

In situ Compressive Loading and Correlative Noninvasive Imaging of the Bone-periodontal Ligament-tooth Fibrous Joint

In this study, the use of an in situ loading device coupled with micro-X-ray computed tomography for fibrous joint biomechanics will be discussed. Experimental readouts identifiable with an overall change in joint biomechanics will include: 1) reactionary force vs. displacement, i.e. tooth displacement within the alveolar socket and its reactionary response to loading, 2) three-dimensional (3D) spatial configuration and morphometrics, i.e.&#160;geometric relationship of the tooth with the alveolar socket, and 3) changes in readouts 1 and 2 due to a change in loading axis, i.e. concentric or eccentric loads.

This study demonstrates a novel biomechanics testing protocol. The advantage of this protocol includes the use of an in situ loading device coupled to a ...

Bioengineering

A Mouse Model for Pathogen-induced Chronic Inflammation at Local and Systemic Sites

Chronic inflammation is a major driver of pathological tissue damage and a unifying characteristic of many chronic diseases in humans including neoplastic, autoimmune, and chronic inflammatory diseases. Emerging evidence implicates pathogen-induced chronic inflammation in the development and progression of chronic diseases with a wide variety of clinical manifestations. Due to the complex and multifactorial etiology of chronic disease, designing experiments for proof of causality and the establishment of mechanistic links is nearly impossible in humans. An advantage of using animal models is that both genetic and environmental factors that may influence the course of a particular disease can be controlled. Thus, designing relevant animal models of infection represents a key step in identifying host and pathogen specific mechanisms that contribute to chronic inflammation.
Here we describe a mouse model of pathogen-induced chronic inflammation at local and systemic sites following infection with the oral pathogen Porphyromonas gingivalis, a bacterium closely associated with human periodontal disease. Oral infection of specific-pathogen free mice induces a local inflammatory response resulting in destruction of tooth supporting alveolar bone, a hallmark of periodontal disease. In an established mouse model of atherosclerosis, infection with P. gingivalis accelerates inflammatory plaque deposition within the aortic sinus and innominate artery, accompanied by activation of the vascular endothelium, an increased immune cell infiltrate, and elevated expression of inflammatory mediators within lesions. We detail methodologies for the assessment of inflammation at local and systemic sites. The use of transgenic mice and defined bacterial mutants makes this model particularly suitable for identifying both host and microbial factors involved in the initiation, progression, and outcome of disease. Additionally, the model can be used to screen for novel therapeutic strategies, including vaccination and pharmacological intervention.

Animal models have proven to be invaluable tools in defining host and pathogen specific mechanisms that contribute to the development of chronic inflammation. Here we describe a mouse model of oral infection with the human pathogen Porphyromonas gingivalis and detail methodologies to assess the progression of inflammation at local and systemic sites.

Chronic inflammation is a major driver of pathological tissue damage and a unifying characteristic of many chronic diseases in humans including ...

Immunology and Infection

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.

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 ...

Medicine

Robust Ligature-Induced Model of Murine Periodontitis for the Evaluation of Oral Neutrophils

The main advantages of studying the pathophysiology of periodontal disease utilizing murine models are the reduced cost of animals, array of genetically modified strains, the vast number of analyses that can be performed on harvested soft and hard tissues. However, many of these systems are subject to procedural criticisms. As an alternative, the ligature-induced model of periodontal disease, driven by the localized development and retention of a dysbiotic oral microbiome, can be employed, which is rapidly induced and relatively reliable. Unfortunately, the variants of ligature-induced murine periodontitis protocol are isolated to focal regions of the periodontium and subject to premature avulsion of the installed ligature. This minimizes the amount of tissue available for subsequent analyses and increases the number of animals required for study. This protocol describes the precise manipulations required to place extended molar ligatures with improved retention and usage of a novel rinse technique to recover oral neutrophils in mice with an alternative approach that mitigates the aforementioned technical challenges.

This article presents a protocol for establishing a ligature-induced model of murine periodontitis involving multiple maxillary molars, resulting in larger areas of the involved gingival tissue and bone for subsequent analysis as well as reduced animal usage. A technique to assess oral neutrophils in a manner analogous to human subjects is also described.

The main advantages of studying the pathophysiology of periodontal disease utilizing murine models are the reduced cost of animals, array of genetically ...

Induction of Periodontitis via a Combination of Ligature and Lipopolysaccharide Injection in a Rat Model

Periodontitis (PD) is a highly prevalent, chronic immune-inflammatory disease of the periodontium, that results in a loss of gingival soft tissue, periodontal ligament, cementum, and alveolar bone. In this study, a simple method of PD induction in rats is described. We provide detailed instructions for placement of the ligature model around the first maxillary molars (M1) and a combination of injections of lipopolysaccharide (LPS), derived from Porphyromonas gingivalis at the mesio-palatal side of the M1. The induction of periodontitis was maintained for 14 days, promoting the accumulation of bacteria biofilm and inflammation. To validate the animal model, IL-1&#946;, a key inflammatory mediator, was determined by an immunoassay in the gingival crevicular fluid (GCF), and alveolar bone loss was calculated using cone beam computed tomography (CBCT). This technique was effective in promoting gingiva recession, alveolar bone loss, and an increase in IL-1&#946; levels in the GCF at the end of the experimental procedure after 14 days. This method was effective in inducing PD, thus being able to be used in studies on disease progression mechanisms and future possible treatments.

Induction of Periodontitis via a Combination of Ligature and Lipopolysaccharide Injection in a Rat Model

In this study, a rat model of induction of periodontitis is presented via a combination of retentive ligature and repetitive injections of lipopolysaccharide derived from Porphyromonas gingivalis, over 14 days around the first maxillary molars. The ligation and LPS injection techniques were effective in inducing peridontitis, resulting in alveolar bone loss and inflammation.

Periodontitis (PD) is a highly prevalent, chronic immune-inflammatory disease of the periodontium, that results in a loss of gingival soft tissue, ...

3D Imaging of PDL Collagen Fibers during Orthodontic Tooth Movement in Mandibular Murine Model

Orthodontic tooth movement is a complex biological process of altered soft and hard tissue remodeling as a result of external forces. In order to understand these complex remodeling processes, it is critical to study the tooth and periodontal tissues within their 3D context and therefore minimize any sectioning and tissue artefacts. Mouse models are often utilized in developmental and structural biology, as well as in biomechanics due to their small size, high metabolic rate, genetics and ease of handling. In principle this also makes them excellent models for dental related studies. However, a major impediment is their small tooth size, the molars in particular. This paper is aimed at providing a step by step protocol for generating orthodontic tooth movement and two methods for 3D imaging of the periodontal ligament fibrous component of a mouse mandibular molar. The first method presented is based on a micro-CT setup enabling phase enhancement&#160;imaging of fresh collagen tissues. The second method is a bone clearing method using ethyl cinnamate that enables imaging through the bone without sectioning and preserves endogenous fluorescence. Combining this clearing method with reporter mice like Flk1-Cre;TdTomato provided a first of its kind opportunity to image the 3D vasculature in the PDL and alveolar bone.

We present a protocol for generating orthodontic tooth movement in mice and methods for 3D visualization of the collagen fibers and blood vessels of periodontal ligament without sectioning.

Orthodontic tooth movement is a complex biological process of altered soft and hard tissue remodeling as a result of external forces. In order to ...

Analyzing Alveolar Bone Remodeling to Explore Skeletal Mechanical Biology

Using Inducible Osteoblastic Lineage-Specific Stat3 Knockout Mice to Study Alveolar Bone Remodeling During Orthodontic Tooth Movement

The alveolar bone, with a high turnover rate, is the most actively-remodeling bone in the body. Orthodontic tooth movement (OTM) is a common artificial process of alveolar bone remodeling in response to mechanical force, but the underlying mechanism remains elusive. Previous studies have been unable to reveal the precise mechanism of bone remodeling in any time and space due to animal model-related restrictions. The signal transducer and activator of transcription 3 (STAT3) is important in bone metabolism, but its role in osteoblasts during OTM is unclear. To provide in vivo evidence that STAT3 participates in OTM at specific time points and in particular cells during OTM, we generated a tamoxifen-inducible osteoblast lineage-specific Stat3 knockout mouse model, applied orthodontic force, and analyzed the alveolar bone phenotype. 
Micro-computed tomography (Micro-CT) and stereo microscopy were used to access OTM distance. Histological analysis selected the area located within three roots of the first molar (M1) in the cross-section of the maxillary bone as the region of interest (ROI) to evaluate the metabolic activity of osteoblasts and osteoclasts, indicating the effect of orthodontic force on alveolar bone. In short, we provide a protocol for using inducible osteoblast lineage-specific Stat3 knockout mice to study bone remodeling under orthodontic force and describe methods for analyzing alveolar bone remodeling during OTM, thus shedding new light on skeletal mechanical biology.

Author Spotlight: Comparing Alveolar and Long Bone Remodeling to Explore OTM Model Potential 

This study provides a protocol for using inducible osteoblast lineage-specific Stat3 knockout mice to study bone remodeling under orthodontic force and describes methods for analyzing alveolar bone remodeling during orthodontic tooth movement, thus shedding light on skeletal mechanical biology.

The alveolar bone, with a high turnover rate, is the most actively-remodeling bone in the body. Orthodontic tooth movement (OTM) is a common artificial ...

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

Trabecular Bone Microarchitecture Evaluation in an Osteoporosis Mouse Model

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

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

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

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

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 ...

Microhardness Measurements on Tooth and Alveolar Bone in Rodent Oral Disease Models

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Meal Duration as a Measure of Orofacial Nociceptive Responses in Rodents

In situ Compressive Loading and Correlative Noninvasive Imaging of the Bone-periodontal Ligament-tooth Fibrous Joint

A Mouse Model for Pathogen-induced Chronic Inflammation at Local and Systemic Sites

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

Robust Ligature-Induced Model of Murine Periodontitis for the Evaluation of Oral Neutrophils

Induction of Periodontitis via a Combination of Ligature and Lipopolysaccharide Injection in a Rat Model

3D Imaging of PDL Collagen Fibers during Orthodontic Tooth Movement in Mandibular Murine Model

Using Inducible Osteoblastic Lineage-Specific Stat3 Knockout Mice to Study Alveolar Bone Remodeling During Orthodontic Tooth Movement

Trabecular Bone Microarchitecture Evaluation in an Osteoporosis Mouse Model

Inducing Apical Periodontitis in Mice