Name: establishment of an oronasal fistula mice model
Uploaded: 2023-09-08T14:25:00.000+00:00
Duration: 3 min 51 s
Description: Watch this Scientific Journal Video about Investigating Wound Healing in Mice Models of Oronasal Fistulas at JoVE.com

This work was supported by the Research and Development Program, West China Hospital of Stomatology, Sichuan University (RD-02-202107), Sichuan Province Science and Technology Support Program (2022NSFSC0743), and Sichuan Postdoctoral Science Foundation (TB2022005) grant to H. Huang.

All animal procedures in this study were reviewed and approved by the Ethical Committee of the West China School of Stomatology, Sichuan University. Adult C57BL/6 mice (female) were used for the present study.
1. Surgical preparation
<ol>
	<li>Gather the necessary surgical instruments for the procedure: germinator, ophthalmologic cautery, microsurgical scissors, microsurgical tweezers, syringes, and needles (26 g x&#160; 0.63 inch) (Figure 1A,B) (see Table of Materials). 
	NOTE: Prior to the surgical procedure, autoclave the surgical instruments, including the ophthalmologic cautery, microsurgical tweezers, and microsurgical scissors, at 102.9 kPa (1.05 kg/cm2) and 121 &#176;C for 20 min.</li>
	<li>Collect the required surgical supplies: surgical drapes, latex gloves, sterile cotton, sterile sheets, sterile metal foil, foam board as the surgical platform, rubber bands (which can be obtained by tearing a medical latex glove), and tape (Figure 1C) (see Table of Materials). 
	NOTE: Use a separate set of supplies for each mouse, including syringes and sterile sheets for the surgical field.</li>
	<li>Clean the surgical area and apparatus (light source, foam board, and temperature maintenance device, see Table of Materials) using alcohol wipes. Cover the knobs and handles of instruments that may be required during the procedure with sterile metal foil.</li>
	<li>Aseptically open the individual instruments and carefully position them in the surgical area. Activate the germinator (see Table of Materials) and the lights for use during the procedure. Place the ophthalmologic cautery into the germinator and heat it to 250 &#176;C for 20 min.</li>
</ol>
2. Surgical procedure
<ol>
	<li>Perform fixation of the mouse and open the oral cavity following the steps below.
	<ol>
		<li>Select a female C57BL/6J mouse weighing 20-25 g and aged 8-12 weeks. House the mouse for 7 days before conducting any procedures.</li>
		<li>Anesthetize the mouse by intraperitoneal injection of Zoletil50 (80 mg/kg) and&#160;Xylazine (5 mg/kg) (see Table of Materials). Apply ophthalmic eye ointment to the eye of the mouse. Wait until there is no toe-pinch response. 
		&#8203;NOTE: The mouse is ready for the procedure when it is unable to turn over independently.</li>
		<li>Secure the mouse to a foam board lined with sterile sheets. Use tape to tie the mouse to the surgical platform in a supine position (Figure 2A).</li>
		<li>Open the oral cavity of the mouse. Position two&#160;needles (26 g x&#160; 0.63 inch) in front of the orbital ear plane and two more behind it. Place a rubber band around the needles and cross the incisors to hold the mouth open. Use microsurgical tweezers to prop open the corners of the mouth (Figure 2B). 
		NOTE: Ensure that the hard palate is clearly exposed. Fix the tongue under the rubber band to prevent obstruction of the field of view and burning during subsequent experiments.</li>
	</ol>
	</li>
	<li>Create the oronasal fistula (ONF) on the hard palate (Figure 3A-F).
	<ol>
		<li>Retrieve the ophthalmologic cautery, which has been heated to 250 &#176;C for 20 min. Place the cautery tip 1 mm away from the intersection of the midline of the palate and the line of the first premolar, creating a full-thickness mucosal injury to the hard palate in the midline. 
		NOTE: Avoid scalding the mouse&#39;s tongue.</li>
		<li>After a few seconds, remove the ophthalmologic cautery when the mucosa around the cautery tip turns white.</li>
		<li>Place the ophthalmologic cautery in the germinator and continue heating it to 250 &#176;C for 10 min. Repeat the previous step to enlarge the wound around the edges until it reaches a length of 2.0 mm and a width of 1.5 mm. 
		NOTE: Each extension should follow the edge of the last injury. Use a vernier caliper to measure the length and width of the injury. The injury should cover 10% of the palate.</li>
		<li>Use microsurgical scissors to remove any excess denatured soft tissue around the wound. Use sterile cotton to stop bleeding and prevent inhalation asphyxiation of the mouse. Measure the wound to ensure it forms a full-thickness hard palate mucosal injury measuring 2.0 x 1.5 mm in the midline.</li>
	</ol>
	</li>
</ol>
3. Post-operative care
<ol>
	<li>Administer Meloxicam to the mouse&#160;at the time of postoperative awakening, at a dose of 5 mg/kg/d for 3 days, subcutaneously29.</li>
	<li>Place the mouse on a temperature maintenance device until it fully regains consciousness. 
	&#8203;NOTE: Ensure that the mouse is positioned in a way that facilitates breathing. Rotate the mice every 10-15 min to prevent blood pooling or collapse of lung lobes. Once the mouse has warmed up, return it to its cage. Provide sterile jelly and irradiated feed at the bottom of the cage for the mice to consume.</li>
</ol>
4. Verification of the oronasal fistula creation
NOTE: The success of the oronasal fistula (ONF) creation is assessed on the 7th day following the surgical procedure.
<ol>
	<li>Prepare the necessary surgical supplies: rubber band, tape, syringes, surgical drapes, latex gloves, sterile sheets, sterile metal foil, and foam board.</li>
	<li>Wear surgical drapes and sterile gloves to maintain aseptic conditions. Disinfect the foam board, light source, and temperature maintenance device with alcohol.</li>
	<li>Induce general anesthesia by intraperitoneal injection of Zoletil50 (80 mg/kg). Wait until there is no toe-pinch response. Use the same method as described in step 2.1.3 and 2.1.4 to immobilize the mouse and expose the hard palate.</li>
	<li>Perform anatomical structural verification by ensuring that the septum is still visible at the wound site, indicating successful ONF creation (Figure 4A,B).</li>
	<li>Perform functional verification: close the oral cavity of the mouse and inject sterile water into its oral cavity using a sterile syringe. The successful creation of ONF is confirmed when fluid flows from the mouse&#39;s nostrils.</li>
	<li>Place the mouse on the temperature maintenance device (37 &#176;C) until it fully regains consciousness. Rotate the mice every 10-15 min to prevent blood pooling or collapse of lung lobes.</li>
</ol>

Development of an Effective Mice Model for Studying Oronasal Fistulas

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The authors have nothing to disclose.

Researchers have explored various materials, drugs, and novel techniques for treating ONF10,11,12,13,14,15,16,17. With advancements in surgical procedures, the incidence and recurrence of ONF have been reduced. However, due to the unique characteristics of the disease, the number of ONF patients in the clinic is limited, necessitating a standardized model for studying potential treatments. While several methods for creating ONF models have been described18,19,20,21,22,23, they were often brief and lacked detailed discussion of the experimental method. The formation of ONF has been verified through microscopic and histological studies that describe the histopathological features12,13,27. This protocol aimed to establish a reproducible mouse model of ONF to facilitate research.
Achieving uniform ONF creation posed a challenge. To ensure reproducibility, it was crucial to uniformly damage the palates of the mice. Controlling the diameter of the ONF, minimizing wound healing, and effectively stopping bleeding were key steps in creating ONF. Microsurgical scissors were used to remove excess soft tissue around the wound after using the ophthalmologic cautery, thereby minimizing changes in wound diameter during the healing phase. However, using microsurgical scissors to remove excess tissue carried the risk of significant bleeding and even death of the mice, contributing to higher mortality rates observed in other experiments12,13,27. In this protocol, the combination of microsurgical scissors and a hemostatic ophthalmic cautery was employed to denature and remove excess tissue, while sterile cotton was used to control bleeding. This method significantly reduced bleeding or even achieved complete hemostasis due to the cauterizing effect of the heated ophthalmic cautery.
An alternative method for creating the ONF model in mice has been reported, involving the use of a biopsy punch13,27. While this method provided better control over the wound diameter due to the consistent size of the punch, it had a high failure rate and posed challenges in managing the required force, potentially leading to the death of the mice. Controlling the depth and strength of ONF creation with this method was difficult, and determining whether the nasal septum had been reached was challenging. Additionally, controlling bleeding was problematic, and mice were at risk of suffocation due to severe bleeding during the experiment.
However, there are limitations to this experimental method. Firstly, the size of the wound cannot be controlled by the same fistula size in each mouse compared to a biopsy punch with a fixed diameter size. And measuring tools should be used to maximize the size of each fistula. The size of the palatal wound is critical to the experiment, as delayed healing of the wound is central to ONF formation. Therefore, determining an appropriate size for the palatal wound is important. If the wound is too small, it may heal quickly, not meeting the time requirements for subsequent experiments. Conversely, if it is too large, mice may die from excessive blood loss during surgery or experience difficulty eating post-surgery, leading to starvation. Hence, exploring the optimal size of the palatal wound is warranted. Nonetheless, the size (2.0 mm x 1.5 mm) used in the current experiment was deemed appropriate. In this protocol, we only use female mice, but either female or male mice can be chosen according to the design of study.

Oronasal fistula (ONF), an abnormal opening between the oral and nasal cavities, clinically manifests as a defect in a structural area from the alveolar process to the uvula, which commonly occurs as a complication following cleft palate repair1. Patients with ONF experience food reflux, articulation disorders, and impaired velopharyngeal function, significantly impacting their quality of life2,3,4. The rate of post-operative ONF ranges from 2.4% to 55% due to factors such as cleft width, Veau type, and surgical method5,6,7,8. Additionally, the recurrence rate after ONF repair is high, ranging from 0% to 43%9.
Several novel treatments have recently shown promise in the field of ONF, including different materials, drugs, and novel techniques10,11,12,13,14,15,16,17. Accurate evaluation of therapeutic effects is essential as it provides the basis for selecting and further developing ONF treatments. However, obtaining a valid assessment in the short term for therapeutic effects other than surgery is challenging, as the characteristics of ONFs vary among different patients. Therefore, establishing an ONF disease model is necessary to verify the effectiveness of these treatment methods.
For several decades, researchers have generated the oronasal fistula (ONF) model in various animal species, including rats18,19, piglets20,21, minipigs22, and dogs23, as these species possess a substantial hard palate suitable for surgical manipulation. However, mice have a genetic sequence and whole genome similar to that of humans, making them an important model for researching and developing new drugs24,25,26. Furthermore, mice offer little variation from batch to batch, making them a favorable choice for establishing the ONF model12,13,27.
However, the detailed steps for creating ONF were not described, and the stability of the ONF size was not taken into consideration. Additionally, the verification of ONF formation relied solely on observation28, without ensuring direct communication between the oral and nasal cavities. It was not demonstrated through other means, such as the mouse&#39;s loss of body weight due to difficulties in eating caused by the ONF. Furthermore, normal variation in wound size was not considered, which is crucial for studies on drugs or materials that promote or inhibit wound healing. Therefore, there is a strong need to establish a stable and validated ONF model.
The objective of this study was to develop a practical ONF model that addresses the aforementioned issues, with the hope that this protocol will serve as the foundation for future research on the mechanisms of palatal wound healing and novel treatments for ONF.

<table><tbody><tr><td>Germinator</td><td>Electron Microscopy Sciences&amp;nbsp;</td><td>66118-20</td><td>Heating and disinfection equipment</td></tr><tr><td>Latex gloves</td><td>Allmed</td><td></td><td>or similar</td></tr><tr><td>Lights</td><td>Olympus</td><td>A1813</td><td></td></tr><tr><td>Meloxicam</td><td>MedChemExpress</td><td>HY-B0261</td><td>crushed; 5 mg/kg</td></tr><tr><td>Microsurgical instruments (scissors and tweezers)</td><td>Jiangsu Tonghui Medical Devices Co.</td><td>M-Y-0087</td><td>Surgical instrument</td></tr><tr><td>Ophthalmologic cautery</td><td>Suqian Wenchong Medical Equipment Co.</td><td>1.00234E+13</td><td>Surgical instrument</td></tr><tr><td>Sterile cotton,</td><td>Yancheng Begu Technology Co.</td><td></td><td>or similar</td></tr><tr><td>Sterile metal foil</td><td>Biosharp</td><td></td><td>or similar</td></tr><tr><td>Sterile sheets</td><td>3M</td><td>XH003801129</td><td>or similar</td></tr><tr><td>Surgical drapes</td><td>Yancheng Begu Technology Co.</td><td></td><td>or similar</td></tr><tr><td>Syringes</td><td>Yancheng Begu Technology Co.</td><td>S-015301</td><td>or similar</td></tr><tr><td>Tape</td><td>Bkmamlab</td><td></td><td>or similar</td></tr><tr><td>Temperature maintenance device</td><td>Harvard Apparatus&amp;nbsp;</td><td>LE-13-2104</td><td></td></tr><tr><td>Zoletil50</td><td>Virbac</td><td></td><td>80 mg/kg</td></tr></tbody></table>

establishment of an oronasal fistula mice model

This study presents a method utilizing heated ophthalmologic cautery to develop a viable model for investigating oronasal fistulas. C57BL/6 mice were used to establish the oronasal fistula (ONF) model. To create the ONF, the mice were anesthetized, immobilized, and their hard palates were exposed. During the surgical procedure, a 2.0 x 1.5 mm full-thickness mucosal injury was induced in the midline of the hard palate using ophthalmologic cautery. It was crucial to control the size of the ONF and minimize bleeding in order to ensure the success of the experiment. Verification of the ONF model's effectiveness was conducted on the 7th-day post-operation, encompassing both anatomical and functional assessments. The presence of the nasal septum within the oral cavity and the outflow of sterile water from the nostrils upon injection into the oral cavity confirmed the successful establishment of the ONF model. The model demonstrated a practical and successful oronasal fistula, characterized by a low mortality rate, significant weight changes, and minimal variation in ONF size. Future studies may consider adopting this methodology to elucidate the mechanisms of palate wound healing and explore novel treatments for oronasal fistulas.

To assess the feasibility and stability of this experimental method, the same procedure was performed on ten mice, and observations were made regarding mortality, changes in wound size, body weight, and histologic analysis. The mice were euthanized on day 7.
The procedure exhibited a low mortality rate. The ophthalmologic cautery and germinator, depicted in Figure 1A-C, were the key instruments utilized in this experiment. The ONF model was created according to the provided protocol. Among the ten mice operated on, only one expired on the 7th day post-operation. The overall mortality rate throughout the experiment was approximately 10%.
The results revealed notable variability in the size of the ONF generated using the described method. On the day of surgery, all mice exhibited oval-shaped wounds measuring 2.0 mm in length and 1.5 mm in width. When assessing ONF formation on the 7th day after surgery, a significant variation in ONF size was observed (P = 0.0085) (Figure 5A,B).
The presence of ONF can result in complications such as food reflux and eating difficulties, potentially leading to changes in weight. Therefore, the body weight of the mice was also taken into account. The mice were weighed on the day of surgery (day 1) and on the 7th day (day 7) when ONF formation was examined. A significant reduction in weight was observed on day 7 compared to day 1 (P &#60; 0.001) (Figure 6A,B). The&#160; loss of their body weight was 25.16%.
For histologic analysis, both the wound and normal tissue were harvested from the mice on day 7. Isolated palates were used as samples for histology sectioning. They were placed in tissue embedding boxes and fixed using 4% paraformaldehyde and 10% formic acid decalcification reagent. The tissues were then embedded in paraffin, sectioned into 7 &#181;m slices along coronal planes, and stained with hematoxylin and eosin (H&#38;E). Histological analysis of the ONF revealed loss of hard palate mucosa, denuded bone, and ONF formation (Figure 7). Histology of the lungs was performed, and no&#160;abnormalities&#160;were detected between normal and ONF mice.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65578/65578fig01.jpg" /> 
Figure 1: Surgical instruments and supplies. (A) The germinator used for heating the ophthalmologic cautery. (B) Surgical instruments: ophthalmologic cautery, microsurgical scissors, microsurgical tweezers, syringes, and needles (26 g x&#160; 0.63 inch). (C) Surgical supplies: surgical drapes, sterile gloves, sterile cotton, sterile sheets, sterile metal foil, foam board, rubber bands, and tape. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65578/65578fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65578/65578fig02.jpg" /> 
Figure 2: Fixation of the mouse and opening of the oral cavity. (A) The forelimbs of the mouse were taped to secure it. (B) Syringe needles were inserted into the foam board, and a rubber band was placed over the needles. The mouse&#39;s oral cavity was opened using a rubber band and microsurgical tweezers. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65578/65578fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65578/65578fig03.jpg" /> 
Figure 3: Creation of the oronasal fistula. (A) Exposing the oral cavity. (B) Placing the tip of the ophthalmologic cautery on the midline portion of the hard palate. (C) Removing the ophthalmologic cautery. (D) Removing excess soft tissue around the wound using microsurgical scissors. (E) Stopping bleeding using sterile cotton. (F) Final formed palatal wound. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65578/65578fig03large.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/65578/65578fig04.jpg" /> 
Figure 4: Examination of the palatal wound on the 7th day after surgery. (A) Palatal wound on day 1. (B) Palatal wound on day 7. White arrows indicate the oronasal fistula (ONF). <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65578/65578fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/65578/65578fig05.jpg" /> 
Figure 5: Size of the palatal wound on Day 1 and Day 7. (A) Mean values for mice on days 1 and 7. (B) Significant difference verified using paired samples t-test. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65578/65578fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/65578/65578fig06.jpg" /> 
Figure 6: Weight of the mice on day 1 and day 7. (A) Mean values for mice on day 1 and day 7. (B) Significant difference verified using paired samples t-test. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65578/65578fig06large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/65578/65578fig07.jpg" /> 
Figure 7: Histological observation. Histological analysis of the ONF shows loss of hard palate mucosa, denuded bone, and ONF formation. (A) Oronasal fistula on day 7, magnification: 4x. (B) Oronasal fistula on day 7, magnification: 10x. (C) No-injury control, magnification: 4x. (D) No-injury control, magnification: 10x. Black arrow shows the location of the ONF. Scale bars: A,C = 200 &#181;m; B,D = 100 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/65578/65578fig07large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Investigating Wound Healing in Mice Models of Oronasal Fistulas at JoVE.com

Author Spotlight: Investigating Wound Healing in Mice Models of Oronasal Fistulas 

This article outlines a step-by-step procedure for establishing a mice model with an oronasal fistula. The oronasal fistula was created by employing heated ophthalmologic cautery to damage the midline portion of the hard palate, resulting in the formation of an opening between the oral and nasal cavities.

Establishment of an Oronasal Fistula Mice Model

This study presents a method utilizing heated ophthalmologic cautery to develop a viable model for investigating oronasal fistulas. C57BL/6 mice were used ...

establishment-of-an-oronasal-fistula-mice-model

Nanyang Technological University

Research

JoVE Journal

Medicine

1.0K Views.  Sichuan University. Ons onderzoek heeft zich gericht op het vaststellen van het stabiele en gestandaardiseerde muismodel van oronasale fistels. Dit protocol heeft ook de veranderingen in het lichaamsgewicht, de veranderingen in de wondgrootte en de histologische kenmerken van de orale neusfistelvorming aangetoond. De huidige experimentele uitdagingen bij het opstellen van oronasale fistelmodellen zijn de sterfte door verstikking of bloedverlies bij muizen en de standaardisatie van de wondgrootte.Ons proto...

Video: Auteur in de schijnwerpers: onderzoek naar wondgenezing bij muizen Modellen van oronasale fistels 

Heterotopic Mucosal Engrafting Procedure for Direct Drug Delivery to the Brain in Mice

Delivery of therapeutics into the brain is impeded by the presence of the blood-brain barrier (BBB) which restricts the passage of polar and high molecular weight compounds from the bloodstream and into brain tissue. Some direct delivery success in humans has been achieved via implantation of transcranial catheters; however this method is highly invasive and associated with numerous complications. A less invasive alternative would be to dose the brain through a surgically implanted, semipermeable membrane such as the nasal mucosa that is used to repair skull base defects following endoscopic transnasal tumor removal surgery in humans. Drug transfer though this membrane would effectively bypass the BBB and diffuse directly into the brain and cerebrospinal fluid. Inspired by this approach, a surgical approach in mice was developed that uses a donor septal mucosal membrane engrafted over an extracranial surgical BBB defect. This model has been shown to effectively allow the passage of high molecular weight compounds into the brain. Since numerous drug candidates are incapable of crossing the BBB, this model is valuable for performing preclinical testing of novel therapies for neurological and psychiatric diseases.

A mouse model of human endoscopic skull base reconstruction has been developed that creates a semipermeable interface between the brain and nose using nasal mucosal grafts. This method allows researchers to study delivery to the central nervous system of high molecular weight therapeutics which are otherwise excluded by the blood-brain barrier when administered systemically.

Delivery of therapeutics into the brain is impeded by the presence of the blood-brain barrier (BBB) which restricts the passage of polar and high ...

Facial Nerve Axotomy in Mice: A Model to Study Motoneuron Response to Injury

The goal of this surgical protocol is to expose the facial nerve, which innervates the facial musculature, at its exit from the stylomastoid foramen and either cut or crush it to induce peripheral nerve injury. Advantages of this surgery are its simplicity, high reproducibility, and the lack of effect on vital functions or mobility from the subsequent facial paralysis, thus resulting in a relatively mild surgical outcome compared to other nerve injury models. A major advantage of using a cranial nerve injury model is that the motoneurons reside in a relatively homogenous population in the facial motor nucleus in the pons, simplifying the study of the motoneuron cell bodies. Because of the symmetrical nature of facial nerve innervation and the lack of crosstalk between the facial motor nuclei, the operation can be performed unilaterally with the unaxotomized side serving as a paired internal control. A variety of analyses can be performed postoperatively to assess the physiologic response, details of which are beyond the scope of this article. For example, recovery of muscle function can serve as a behavioral marker for reinnervation, or the motoneurons can be quantified to measure cell survival. Additionally, the motoneurons can be accurately captured using laser microdissection for molecular analysis. Because the facial nerve axotomy is minimally invasive and well tolerated, it can be utilized on a wide variety of genetically modified mice. Also, this surgery model can be used to analyze the effectiveness of peripheral nerve injury treatments. Facial nerve injury provides a means for investigating not only motoneurons, but also the responses of the central and peripheral glial microenvironment, immune system, and target musculature. The facial nerve injury model is a widely accepted peripheral nerve injury model that serves as a powerful tool for studying nerve injury and regeneration.

We present a surgical protocol detailing how to perform a cut or crush axotomy on the facial nerve in the mouse. The facial nerve axotomy can be employed to study the physiological response to nerve injury and test therapeutic techniques.

The goal of this surgical protocol is to expose the facial nerve, which innervates the facial musculature, at its exit from the stylomastoid foramen and ...

Whole Mount Labeling of Cilia in the Main Olfactory System of Mice

The mouse olfactory system comprises 6-10 million olfactory sensory neurons in the epithelium lining the nasal cavity. Olfactory neurons extend a single dendrite to the surface of the epithelium, ending in a structure called dendritic knob. Cilia emanate from this knob into the mucus covering the epithelial surface. The proteins of the olfactory signal transduction cascade are mainly localized in the ciliary membrane, being in direct contact with volatile substances in the environment. For a detailed understanding of olfactory signal transduction, one important aspect is the exact morphological analysis of signaling protein distribution. Using light microscopical approaches in conventional cryosections, protein localization in olfactory cilia is difficult to determine due to the density of ciliary structures. To overcome this problem, we optimized an approach for whole mount labeling of cilia, leading to improved visualization of their morphology and the distribution of signaling proteins. We demonstrate the power of this approach by comparing whole mount and conventional cryosection labeling of Kirrel2. This axon-guidance adhesion molecule is known to localize in a subset of sensory neurons and their axons in an activity-dependent manner. Whole mount cilia labeling revealed an additional and novel picture of the localization of this protein.

Cilia of olfactory sensory neurons contain proteins of the signal transduction cascade, but a detailed spatial analysis of their distribution is difficult in cryosections. We describe here an optimized approach for whole mount labeling and en face visualization of ciliary proteins.

The mouse olfactory system comprises 6-10 million olfactory sensory neurons in the epithelium lining the nasal cavity. Olfactory neurons extend a single ...

Neuroscience

Establishment of a Rat Model of Superior Sagittal-Sinus Occlusion via a Thread-Embolism Method

The mechanisms contributing to the natural onset of cerebral venous sinus thrombosis (CVST) are mostly unknown, and a variety of uncontrollable factors are involved in the course of the disease, resulting in great limitations in clinical research. Therefore, the establishment of stable CVST animal models that can standardize a variety of uncontrollable confounding factors have helped to circumvent shortcomings in clinical research. In recent decades, a variety of CVST animal models have been constructed, but the results based on these models have been inconsistent and incomplete. Hence, in order to further explore the pathophysiological mechanisms of CVST, it is necessary to establish a novel and highly compatible animal model, which has important practical value and scientific significance for the diagnosis and treatment of CVST. In the present study, a novel Sprague-Dawley (SD) rat model of superior sagittal sinus (SSS) thrombosis was established via a thread-embolization method, and the stability and reliability of the model were verified. Additionally, we evaluated changes in cerebral venous blood flow in rats after the formation of CVST. Collectively, the SD-rat SSS-thrombosis model represents a novel CVST animal model that is easily established, minimizes trauma, yields good stability, and allows for accurately controlling ischemic timing and location.

Here, we establish a novel Sprague-Dawley (SD) rat model of superior sagittal sinus (SSS) thrombosis via a thread-embolization method, and the stability and reliability of the model were verified.

The mechanisms contributing to the natural onset of cerebral venous sinus thrombosis (CVST) are mostly unknown, and a variety of uncontrollable factors ...

Establishing a Transcranial Middle Cerebral Artery Occlusion Mice Model

A Modified Transcranial Middle Cerebral Artery Occlusion Model to Study Stroke Outcomes in Aged Mice

In experimental stroke research, middle cerebral artery occlusion (MCAO) with an intraluminal filament is widely used to model ischemic stroke in mice. The filament MCAO model typically exhibits a massive cerebral infarction in C57Bl/6 mice that sometimes includes brain tissue in the territory supplied by the posterior cerebral artery, which is largely due to a high incidence of posterior communicating artery atresia. This phenomenon is considered a major contributor to the high mortality rate observed in C57Bl/6 mice during long-term stroke recovery after filament MCAO. Thus, many chronic stroke studies exploit distal MCAO models. However, these models usually produce infarction only in the cortex area, and consequently, the assessment of post-stroke neurologic deficits could be a challenge. This study has established a modified transcranial MCAO model in which the MCA at the trunk is partially occluded either permanently or transiently via a small cranial window. Since the occlusion location is relatively proximal to the origin of the MCA, this model generates brain damage in both the cortex and striatum. Extensive characterization of this model has demonstrated an excellent long-term survival rate, even in aged mice, as well as readily detectable neurologic deficits. Therefore, the MCAO mouse model described here represents a valuable tool for experimental stroke research.

Author Spotlight: Innovative Techniques and Future Directions in Stroke Research 

This protocol demonstrates a unique mouse stroke model with a medium-sized infarct and an excellent survival rate. This model allows preclinical stroke researchers to extend the ischemia duration, use aged mice, and assess long-term functional outcomes.

In experimental stroke research, middle cerebral artery occlusion (MCAO) with an intraluminal filament is widely used to model ischemic stroke in mice. ...

Modified Arteriovenous Fistula Surgery to Establish the RV Volume Overload Mouse Model

Establishment and Confirmation of a Postnatal Right Ventricular Volume Overload Mouse Model

Right ventricular (RV) volume overload (VO) is common in children with congenital heart disease. In view of distinct developmental stages,the RV myocardium may respond differently to VO in children compared to adults. The present study aims to establish a postnatal RV VO model in mice using a modified abdominal arteriovenous fistula. To confirm the creation of VO and the following morphological and hemodynamic changes of the RV, abdominal ultrasound, echocardiography, and histochemical staining were performed for 3 months. As a result, the procedure in postnatal mice showed an acceptable survival and fistula success rate. In VO mice, the RV cavity was enlarged with a thickened free wall, and the stroke volume was increased by about 30%-40% within 2 months after surgery. Thereafter, the RV systolic pressure increased, corresponding pulmonary valve regurgitation was observed, and small pulmonary artery remodeling appeared. In conclusion, modified arteriovenous fistula (AVF) surgery is feasible to establish the RV VO model in postnatal mice. Considering the probability of fistula closure and elevated pulmonary artery resistance, abdominal ultrasound and echocardiography must be performed to confirm the model status before application.

Author Spotlight: Establishment and Confirmation of a Postnatal Right Ventricular Volume Overload Mouse Model  

This protocol presents the establishment and confirmation of a postnatal right ventricular volume overload (VO) model in mice with abdominal arteriovenous fistula (AVF), which can be applied to investigate how VO contributes to postnatal heart development.

Right ventricular (RV) volume overload (VO) is common in children with congenital heart disease. In view of distinct developmental stages,the RV ...

A Murine Maxillary Orthodontic Model Guide to Bone Remodeling Analysis

The Establishment of a Murine Maxillary Orthodontic Model

Due to the lack of reproducible protocols for establishing a murine maxillary orthodontic model, we present a reliable and reproducible protocol to provide researchers with a feasible tool to analyze mechanical loading-associated bone remodeling. This study presents a detailed flowchart in addition to different types of schematic diagrams, operation photos, and videos. We performed this protocol on 11 adult wide-type C57/B6J mice and harvested samples on postoperative days 3, 8, and 14. The micro-CT and histopathological data have proven the success of tooth movements coupled with bone remodeling using this protocol. Furthermore, according to the micro-CT results on days 3, 8, and 14, we have divided bone modeling into three stages: preparation stage, bone resorption stage, and bone formation stage. These stages are expected to help researchers concerned with different stages to set sample collection time reasonably. This protocol can equip researchers with a tool to carry out regenerative analysis of bone remodeling.

Author Spotlight: Insights into an Efficient Murine Maxillary Orthodontic Model Protocol

Here we demonstrate step-by-step a manageable, orthodontic tooth movement protocol operated on a murine maxillary model. With the explicit explanation of each step and visual demonstration, researchers can master this model and apply it to their experimental needs with a few modifications.

Due to the lack of reproducible protocols for establishing a murine maxillary orthodontic model, we present a reliable and reproducible protocol to ...

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 ...

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.

This study introduces the development of a molar extraction model in the murine mandible to provide a practicable model for studying alveolar bone ...

Biology

Establishing an Orofacial Muscle Fibrosis Model in Mice via a Freezing Injury

Freezing Injury in Mouse Masseter Muscle to Establish an Orofacial Muscle Fibrosis Model

Orofacial muscle constitutes a subset of skeletal muscle tissue, with a distinct evolutionary trajectory and development origin. Unlike the somite-derived limb muscles, the orofacial muscles originate from the branchial arches, with exclusive contributions from the cranial neural crest. A recent study has revealed that regeneration is also different in the orofacial muscle group. However, the underlying regulatory mechanism remains to be uncovered. Current skeletal muscle regeneration models mainly focus on the limb and trunk muscle. In this protocol, dry ice was used to induce freezing injury in the mouse masseter muscle and tibialis anterior muscle to create an orofacial muscle fibrosis model. The temporal dynamics of muscle satellite cells and fibro-adipogenic progenitors were different between the two muscles, leading to impaired myofiber regeneration and excessive extracellular matrix deposition. With the help of this model, a deeper investigation into muscle regeneration in the orofacial area could be carried out to develop therapeutic approaches for patients with orofacial diseases.

Author Spotlight: Exploring Orofacial Muscle Regeneration &#8211; Insights and Innovations

The goal of this protocol is to establish an orofacial muscle fibrosis model. Comparison of the histology between mice masseter and tibialis anterior muscle after freezing injury confirmed masseter muscle fibrosis. This model will facilitate further investigation into the mechanism underlying orofacial muscle fibrosis.

Orofacial muscle constitutes a subset of skeletal muscle tissue, with a distinct evolutionary trajectory and development origin. Unlike the somite-derived ...

Establishment of an Oronasal Fistula Mice Model

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