We would like to acknowledge Dr. Oded Heyman for his help with animal positioning, Raphael Lieber for help with micro-CT analysis, and Prof. Andiara De Rossi Daldegan for advice on preforming the experiment. We would also like to acknowledge Dr. Sidney Cohen for critical reading and editing.
This work was supported by a grant from the Dr. Izador I. Cabakoff Research Endowment Fund to MK and IA, and a Yitzhak Navon fellowship from the Israel Ministry of Science and Technology to EG.

All methods described here have been approved by the Institutional Animal Care and Use Committee (IACUC) of The Hebrew University (Ethics no. MD-17-15093-5).
1. Animal anesthesia and positioning
<ol>
	<li>Prepare sterile solutions as described below.
	<ol>
		<li>Prepare 5 mL of 7 mg/mL of Ketamine and 0.09 mg/mL Medetomidine diluted in Phosphate Buffer Solution (PBS)/Saline.&#160;</li>
		<li>Prepare a sterile solution of Atipamezole (0.4 mg/mL) diluted in PBS/Saline (recommended to prepare an amount of 5 mL).</li>
		<li>Prepare a sterile solution of Mepivacaine (7.5 mg/mL) diluted in PBS/Saline for local anesthesia (one supplied vial of Mepivacaine is sufficient for a stock of 7.2 mL).</li>
	</ol>
	</li>
	<li>Use 6-8 weeks old female C57BL/6 mice for the experiment. Keep animals in a specific pathogen free (SPF) animal unit, with standard water and food provided by the facility and the animal feeding ad libitum.</li>
	<li>Weigh mice and inject intraperitoneally (IP) a volume of 10 &#181;L for each gram of the weight. For example, for a mouse that weighs 20 grams, 
	inject 200 &#181;L of Ketamine/Medetomidine solution. This will give a final concentration of (70 mg/kg) Ketamine and (0.9 mg/kg) Medetomidine 
	for each animal.</li>
	<li>In order to prevent eye ulceration due to dryness during the procedure, apply eye ointment containing Chloramphenicol (5%) on the animals&#39; eyes while they are under anesthesia. Keep the animals warm with a lamp while they are anesthetized. Check depth of anesthesia by checking pedal reflex (performing firm toe pinch).</li>
	<li>After the mouse is anesthetized, position the mouse laying on its right side (for a right handed researcher) on a closed-cell extruded polystyrene foam surface and attach feet to the surface using tape.</li>
	<li>Open the mouth by using small rubber bands around the incisors (1 for upper incisors, and 1 for lower incisors) held in place by long needles pinned into the closed-cell extruded polystyrene foam surface.</li>
	<li>Retract the right cheek by using forceps taped to the surface. Use forceps that are closed in steady state.</li>
	<li>Place the surface with mouse in a comfortable working position enabling access to mandibular molars, using proper magnification (a binocular microscope or clinical microscope) and light.</li>
	<li>Retract the tongue by using forceps or dental spatula and inject local anesthesia with a 27 Gauge needle into the mucobuccal fold adjacent to the first mandibular molar. 25-50 &#181;L is sufficient. Swelling of the tissue around the injection site should be visualized.</li>
</ol>
2. Pulp exposure
<ol>
	<li>Use a 1/4 round dental burr, or a same size diamond burr (a long shank is recommended) at a speed of 800 rounds per minute (rpm), attached to any suitable dental motor (for example an automatic torque reduction (ATR) motor).</li>
	<li>Drill on the occlusal part of the first right mandibular molar until the pulp horns are visible through the dentin. use short bursts of the motor to prevent area over-heating.</li>
	<li>Use a K- file or H-file #8 or #10 to pierce the pulp and insert the file into the pulp horns (mesial and distal) by breaking the dentin covering them. Files are sterilized by autoclave.</li>
	<li>Continue working with the files inside the pulp as deep as possible, while widening the openings with the files. Usually there will be bleeding visible from the pulp.</li>
	<li>Make sure to round sharp edges of the tooth with the burr and remove the tooth from occlusion, in order to reduce pain during the experiment. Clean debris during the procedure with a microbrush.</li>
	<li>Leave the contralateral tooth (left first mandibular molar) as a control.</li>
</ol>
3. End of clinical procedure
<ol>
	<li>Release mouse from the fixation state and inject Atipamezole IP (10 &#181;L for each gram of body weight. This will give a final dose of (4 mg/kg) Atipamezole.</li>
	<li>Inject buprenorphine diluted in PBS/saline (0.05 mg/kg) IP (recommended to prepare an amount of 3 mL on the day of the procedure). See Discussion for explanation why pain relief is necessary.</li>
	<li>Make sure the animals recover from anesthesia before leaving them. Give them soft food because they may be incapable of eating hard food due to tooth pain.</li>
</ol>
4. Post procedure follow-up (42 days)
<ol>
	<li>For the first 3 days after the procedure, weigh the animals and inject Buprenorphine (0.1 mg/kg) IP once a day (recommended to prepare an amount of 6mL).</li>
	<li>During the time of the experiment (42 days when inflammation builds up) monitor the animals by weight and general behavioral assessment 2-3 times a week.</li>
	<li>Deliver soft food to the animals during the course of the follow-up period. Wet the food with water and put it in a petri-dish on the floor of the cage.</li>
</ol>
5. Experiment termination and analysis
<ol>
	<li>When 42 days11 of follow-up have been completed , anesthetize animals IP with ketamine/xylazine at a concentration of 106.25 mg/kg Ketamine, and 75 mg/kg Xylazine (A stock solution of 42.5 mg/mL Ketamine, and 1.5 mg/mL Xylazine in a total volume of 2 mL is recommended) and preform cervical dislocation. 
	NOTE: There are different options for possible analyses in order to evaluate inflammation20,21. Here the analysis of the tissue by micro-CT and histological staining will be described.</li>
	<li>After euthanasia, use surgical scissors and forceps to cut out part of the jaw including the 3 molars (separately for each side- treated and non-treated control). By using forceps peel off as much as the soft tissue as possible, leaving mostly bone and teeth on the sample.</li>
	<li>Rinse the tissue briefly in PBS, and then put it in paraformaldehyde (PFA) 4% (diluted in PBS) for 24-48 hours for fixation. 
	CAUTION: Use PFA in a chemical hood, and according to its Material Safety Data Sheet (MSDS) guidelines.</li>
	<li>After 24-48 hours, rinse with PBS 3x in order to wash out the PFA.</li>
	<li>Micro-CT
	<ol>
		<li>For micro-CT analysis, take the harvested tissues (part of the mandible including 3 molars, in dimensions of approximately 4 mm x 5 mmx 1 mm) to a micro-CT scanner, place them in an 12 mm diameter tube filled with 1.5 mL of PBS oriented so that the buccal or lingual surfaces of the tooth and root are laying parallel to the bottom of the tube. Separate between the samples with sponge, and cover the top of the tube with flexible film.</li>
		<li>Open the scanning panel and choose the measurement number. Choose the control file with the following parameters: energy of 70 kV, intensity of 114 &#181;A, and resolution of 6 &#181;m3 voxel size.</li>
		<li>Click on scout view, and when the image appears, click reference line and mark the area of the samples for scanning. After each sample click add scan. When done marking the samples, go to the task list and choose start interact tasks. 
		NOTE: The same samples can subsequently be used for histology. It is important that they do not dry out during the CT-scan.</li>
	</ol>
	</li>
	<li>Use an appropriate computer program for micro-CT alignment and analysis.
	<ol>
		<li>Open the sample on the XY plane in the micro-CT evaluation. Choose three points on each sample for alignment: 
		The mesial apical constriction - defines the origin of the coordinate system. 
		The coronal part of the mesial canal - defines a point on the Z axis. 
		&#8203;The distal apical constriction - defines a point on the XZ plane.</li>
		<li>Use the measurement tool on the left panel, and draw a line reaching each point. Define each of the three points by an X, Y &#38; Z value, as it appears in the right panel.</li>
		<li>Define the area of interest in the sample by choosing task under &#181;CT evaluation, and click on 3D evaluation. A rectangle and a window with the dimensions per axis will appear on the screen. Match the dimensions of the rectangle to fit the area of interest on the XY axis by clicking in the corners with the middle button of the mouse. Choose the dimensions in the Z axis by inserting the first slice number of interest in the Z square under VOI Start, and the number of slices from the first until the last slice number of interest in the Z square under Dim.</li>
		<li>Click on applications under session manager, and choose DECterm. Wait a few seconds for a window to open, and then type the five steps of slanted script described below according to the following instructions (between the lines click enter): 
		ipl 
		isq_to_aim 
		aim1</li>
		<li>Copy the isq (file of interest) name: choose views under session manager and click microCTdata. Find the file being working on, and middle click on the file. Then middle click back in the window of the DECterm.</li>
		<li>Enter the X, Y &#38; Z values (with a space between them) which were determined under VOI when choosing region of interest.</li>
		<li>Enter the X, Y &#38; Z values (with a space between them) which were determined under dim when choosing region of interest.</li>
		<li>Wait until receiving a message: isq to aim completed. 
		(step II- alignment): 
		align z 
		aim1 
		aim2</li>
		<li>Enter the X, Y &#38; Z values (with a space between them) which were determined for the origin of the coordinate system.</li>
		<li>Enter the X, Y &#38; Z values (with a space between them) which were determined for the point on the z axis.</li>
		<li>Enter the X, Y &#38; Z values (with a space between them) which were determined for the point on the XZ plane.</li>
		<li>Wait until receiving a message: &#34;align_Z completed&#34; 
		(step III- header): 
		header 
		aim2 
		0 0 0 
		0 0 0</li>
		<li>Click enter. 
		(step IV- converting the file back from aim to isq): 
		toisq 
		aim2</li>
		<li>Copy the isq (file of interest) name: choose views under session manager and click microCT data. Find the file you are working on, and middle click on the file. Then middle click back in the window of the DECterm. Erase (using &#34;backspace&#34;) the &#34;.ISQ&#34; at the end of the file, and write:&#34;_new.ISQ&#34; to recognize the aligned file.</li>
		<li>Click enter | enter.</li>
		<li>Wait until receiving a message: &#34;toisq_from_aim completed&#34; 
		(step V- flip): 
		isq 
		a</li>
		<li>Copy the new isq (file of interest) name: choose views under session manager and click microCT data. Find the file being working on, and middle click on the file. Then middle click back in the window of the DECterm.</li>
		<li>Click enter | enter. Wait to receive a message: &#34;isq_to_aim completed&#34; 
		flip 
		a 
		aa 
		zx 
		wait to receive a message: &#34;flip_aim completed&#34; 
		from 
		aa</li>
		<li>Copy the new isq name: choose views under session manager and click microCT data. Find the file being working on, and middle click on the file. Then middle click back in the window of the DECterm. Erase (using &#34;backspace&#34;) the &#34;.ISQ&#34; at the end of the file, and write: &#34;_flip.ISQ&#34; to recognize the aligned file.</li>
		<li>Wait until receiving a message: &#34;from_aim_to_isq completed&#34;</li>
	</ol>
	</li>
	<li>Contouring
	<ol>
		<li>Choose the slice which represents approximately the middle of the tooth, in which the coronal pulp, radicular pulp of both canals, and both apical foramens are presented.</li>
		<li>Manually mark the contour of interest on the middle slice by clicking on the upper left contour panel- starting from the distal point of the distal root apex mark the apical border coronally to the radiopaque apical area, through the mesial border of the mesial root apex following the distal border of the mesial root and the mesial border of the distal root through the furcation region&#160;(See Figure 3 II,IV).</li>
		<li>Copy this contour (Ctrl+C, Ctrl+V) and adjust it accordingly to 5 slices positioned on either side of the middle slice.</li>
		<li>To calculate the tissue volume of the contour marked for each sample choose task under &#181;CT evaluation, and click on 3D evaluation. In the window of 3D evaluation click select near task and choose a filter to calculate the tissue volume (TV).</li>
	</ol>
	</li>
	<li>Histology
	<ol>
		<li>After the removal of the tissues from the micro-CT scanner, decalcify the tissues by putting each sample in a microcentrifuge tube with 1 mL of ethylenediaminetetraacetic acid (EDTA) 0.5 M pH 8.0 for 10 days. Change the EDTA solution every 3 days.</li>
		<li>Dehydration: Put samples in histological cassettes, and into increasing ethanol concentrations, an hour each, as follows: 70% ethanol (at this point the process can be stopped for several weeks, as long as the samples are kept in Ethanol), 90% ethanol 2x, 100% ethanol 2x, xylene 2x (caution: use xylene in a chemical hood, and according to its MSDS guidelines).
		<ol>
			<li>Transfer cassettes to liquid paraffin (60 &#176;C) and leave in chemical hood overnight for the xylene to evaporate.</li>
		</ol>
		</li>
		<li>Blocking: Use a histological blocking machine to embed the samples in paraffin. Be careful to embed them in a correct position (i.e., the crown and roots should be oriented parallel to the bottom part of the mold, in a way that the tooth is &#34;lying down&#34;) in order to get sagittal sections. The samples are now ready to be cut with a microtome.</li>
		<li>Cutting the sections: Start by cutting thick slices of ~20 &#181;m until reaching the relevant part of tissue. Once recognizing any part of the tooth (crown or roots) change to sections of 6 &#181;m, and change the angle of cutting according to the previous section.
		<ol>
			<li>For example, if the previous section included the crown but not the roots, change the angle of the block in the microtome so that the roots come closer to the knife, and the crown goes back (the parts of the tooth can be recognized with a naked eye on the block itself).</li>
			<li>Continue to adjust the angle until obtaining sagittal sections including coronal and radicular pulp, and the apical foramen. Cut in this orientation as many slices as possible, until the relevant tissue is all cut.</li>
		</ol>
		</li>
		<li>For staining protocols (e.g., Haemotoxylin and Eosin staining (H&#38;E), Brown &#38; Brenn staining, Tartrate-Resistant Acid Phosphatase (TRAP) staining, immunohistochemistry), see20,21.</li>
	</ol>
	</li>
</ol>

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L., Kadari, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preemptive+analgesia+and+local+anesthesia+as+a+supplement+to+general+anesthesia:+a+review.">Preemptive analgesia and local anesthesia as a supplement to general anesthesia: a review.</a> Anesthesia Progress. 52 (1), 29-38 (2005).</li><li>Song, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+a+Direct+Pulp-capping+Model+for+the+Evaluation+of+Pulpal+Wound+Healing+and+Reparative+Dentin+Formation+in+Mice.">Development of a Direct Pulp-capping Model for the Evaluation of Pulpal Wound Healing and Reparative Dentin Formation in Mice.</a> Journal of Visual Experimentalization. (119), (2017).</li><li>Yoneda, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+a+root+canal+treatment+model+in+the+rat.">Development of a root canal treatment model in the rat.</a> Scientific Reports. 7 (1), 3315 (2017).</li><li>AlShwaimi, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Regulatory+T+cells+in+mouse+periapical+lesions.">Regulatory T cells in mouse periapical lesions.</a> Journal of Endodontics. 35 (9), 1229-1233 (2009).</li></ol>

The authors have nothing to disclose

A method is introduced here for the induction of apical periodontitis in mice. The goal of the method is to exploit the apical periodontitis condition for studying mechanisms and consequences of this inflammatory process. Apical periodontitis was induced in 6-8 week old mice, an age in which the roots are fully developed24. In order to cause apical periodontitis in this model, the tooth pulp of mouse mandibular molars is exposed using a dental burr. Bacteria from the oral flora of the...

The goal of this method is to induce apical periodontitis in a mouse by contaminating the apex with the natural microflora, and to then study various characteristics of this pathological process.
Apical periodontitis (AP) is a common pathology of an inflammatory nature in the periodontal tissues surrounding the tooth root. This dental disease can cause severe pain and must be treated by a dentist. The treatment options include root canal treatment (primary or secondary), endodontic surgery, tooth extraction, or follow-up depending on the clinical and radiographic findings, and the opinion of the clinician. The mechanism of this inflammatory process, although studied for several decades1,2,3, is still not comprehensively understood. Considering the severity of this pathology, there is thus a clear need for research addressing its fundamental nature. Thus, systems where the study of AP is possible are of great scientific interest.
Since AP is a complex pathological process involving the local tissues and the immune system, in vitro studies are insufficient for a complete understanding of the processes. Study of clinical samples of this disease are also problematic due to ethical limitations and significant variability between different people and different clinical stages4,5, and hence the necessity of in vivo models. These models are based on the concept of exposing the dental pulp to contamination and observing the inflammatory reaction of the body to this stimulus in the periapical tissues6,7. Common in vivo models include rodents or larger animals such as dogs. Despite the clinical challenge in treating mice, which are very small animals with miniature teeth, the advantages of the mouse model are significant: practically, working with mice is technically simple in terms of facilities and is most cost effective, and scientifically, the mouse is a well-studied animal model with readily available genetic and molecular tools and a well-studied genome. Indeed, previous studies used a mouse model for studying inflammatory and bone resorption signals and cells involved in apical periodontitis8,9,10,11. Therefore, a clear protocol on how to use a mouse model for the study of AP is needed. Here, we describe such a protocol.
The protocol described here is has the big advantage of being appropriate to study knock-out (KO) mice and learn how the lack of a specific gene affects dental inflammation7,12. Other useful applications of this protocol include the study of effects of medications and systemic conditions on the development of apical periodontitis13, the effect of apical periodontitis on the development of osteonecrosis of the jaws14,15 and stem cell therapy for bone regeneration16.
This protocol can also be generalized as a model to study local inflammation. To study the inflammatory process, several mouse models have been developed, which include for example induced colitis or arthritis17,18. These models have systemic effects and have no built-in control in the same animal. Models for induced apical periodontitis, which include a contralateral control without inflammation, have the advantage of overcoming these limitations14,19.
The protocol described below is therefore useful for researchers who are interested in local inflammatory processes. The controlled nature of this inflammation, its confinement to a specific location, and the contralateral control tooth, all make this protocol valuable for studying the mechanisms involved in this process. Moreover, the protocol is useful for researchers interested in the clinical aspects of periapical inflammation. The mouse model is ideal to study different variables of the disease, in addition to the advantage of being able to easily perform genetic manipulations in the mouse model, to investigate the activity of specific genes in periapical inflammation.
Technically, the clinical procedure is challenging to carry out due to the small dimensions of the mice teeth. It will be beneficial to visualize this procedure in order to learn about positioning, equipment needed, and performance.

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			<td>CAS 104075-48-1</td>
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			<td>microbrushes- adjustable precision applicators</td>
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			<td>EMS</td>
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			<td></td>
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			<td>Chloramphenicol eye ointment (5%)</td>
			<td>Rekah pharmaceutical</td>
			<td>CAS 56-75-7</td>
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		<tr>
			<td>tweezers</td>
			<td>WAM</td>
			<td>Ref-CT</td>
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			<td>Eurovet Animal Health</td>
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			<td>Gadot</td>
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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.

A flow chart of the experimental steps is presented in Figure 1. As mentioned in the protocol, the mice are anesthetized, and their first mandibular molar on one side is drilled until pulp exposure, while the contralateral tooth is left as a control. Next, the teeth are left to be contaminated by the oral flora for 42 days, during which they are monitored and receive pain medication. After 42 days, mice are euthanized, and the teeth and adjacent jaw are taken...

Watch this Scientific Journal Video about Inducing Apical Periodontitis in Mice at JoVE.com

Inducing Apical Periodontitis in Mice

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 ...

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12.0K Views.  Hebrew University of Jerusalem. Here, we present a protocol to locally induce apical periodontitis in mice. We show how to drill a hole in the mouse'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.

Video: Inducing Apical Periodontitis in Mice

Heterotopic Heart Transplantation in Mice

The mouse heterotopic heart transplantation has been used widely since it was introduced by Drs. Corry and Russell in 1973. It is particularly valuable for studying rejection and immune response now that newer transgenic and gene knockout mice are available, and a large number of immunologic reagents have been developed. The heart transplant model is less stringent than the skin transplant models, although technically more challenging. We have developed a modified technique and have completed over 1000 successful cases of heterotopic heart transplantation in mice. When making anastomosis of the ascending aorta and abdominal aorta, two stay sutures are placed at the proximal and distal apexes of recipient abdominal aorta with the donor s ascending aorta, then using 11-0 suture for anastomosis on both side of aorta with continuing sutures. The stay sutures make the anastomosis easier and 11-0 is an ideal suture size to avoid bleeding and thrombosis.
When making anastomosis of pulmonary artery and inferior vena cava, two stay sutures are made at the proximal apex and distal apex of the recipient s inferior vena cava with the donor s pulmonary artery. The left wall of the inferior vena cava and donor s pulmonary artery is closed with continuing sutures in the inside of the inferior vena cava after, one knot with the proximal apex stay suture the right wall of the inferior vena cava and the donor s pulmonary artery are closed with continuing sutures outside the inferior vena cave with 10-0 sutures. This method is easier to perform because anastomosis is made just on the one side of the inferior vena cava and 10-0 sutures is the right size to avoid bleeding and thrombosis. In this article, we provide details of the technique to supplement the video.

The mouse heterotopic heart transplantation model has been proven by many investigators to be an important method for studying mechanisms of rejection and immune response. However, the techniques involved are still challenging. By modifying standard techniques we have had success with more than 1000 transplants.

The mouse heterotopic heart transplantation has been used widely since it was introduced by Drs. Corry and Russell in 1973. It is particularly valuable ...

Small Bowel Transplantation In Mice

Since 1990, the development of tacrolimus-based immunosuppression and improved surgical techniques, the increased array of potent immunosuppressive medications, infection prophylaxis, and suitable patient selection helped improve actuarial graft and patient survival rates for all types of intestine transplantation. Patients with irreversible intestinal failure and complications of parenteral nutrition should now be routinely considered for small intestine transplantation. However, Survival rates for small intestinal transplantation have been slow to improve compares increasingly favorably with renal, liver, heart and lung. The small bowel transplantation is still unsatisfactory compared with other organs. Further progress may depend on better understanding of immunology and physiology of the graft and can be greatly facilitated by animal models. A wider use of mouse small bowel transplantation model is needed in the study of immunology and physiology of the transplantation gut as well as efficient methods in diagnosing early rejection. However, this model is limited to use because the techniques involved is an extremely technically challenging. We have developed a modified technique. When making anastomosis of portal vein and inferior vena cava, two stay sutures are made at the proximal apex and distal apex of the recipient s inferior vena cava with the donor s portal vein. The left wall of the inferior vena cava and donor s portal vein is closed with continuing sutures in the inside of the inferior vena cava after, after one knot with the proximal apex stay suture the right wall of the inferior vena cava and the donor s portal vein are closed with continuing sutures outside the inferior vena cave with 10-0 sutures. This method is easier to perform because anastomosis is made just on the one side of the inferior vena cava and 10-0 sutures is the right size to avoid bleeding and thrombosis.  In this article, we provide details of the technique to supplement the video.

The mouse small bowel transplantation model has been recognized as an important tool to study mechanismes of immune rejection and screen new immunosuppressive drugs. However, this model is limited to use because the techniques involved is an extremely technically challenge. Now we introduce the modified technique.

Since 1990, the development of tacrolimus-based immunosuppression and improved surgical techniques, the increased array of potent immunosuppressive ...

Transverse Aortic Constriction in Mice

Transverse aortic constriction (TAC) in the mouse is a commonly used experimental model for pressure overload-induced cardiac hypertrophy and heart failure.1 TAC initially leads to compensated hypertrophy of the heart, which often is associated with a temporary enhancement of cardiac contractility. Over time, however, the response to the chronic hemodynamic overload becomes maladaptive, resulting in cardiac dilatation and heart failure.2 The murine TAC model was first validated by Rockman et al.1, and has since been extensively used as a valuable tool to mimic human cardiovascular diseases and elucidate fundamental signaling processes involved in the cardiac hypertrophic response and heart failure development. When compared to other experimental models of heart failure, such as complete occlusion of the left anterior descending (LAD) coronary artery, TAC provides a more reproducible model of cardiac hypertrophy and a more gradual time course in the development of heart failure. Here, we describe a step-by-step procedure to perform surgical TAC in mice. To determine the level of pressure overload produced by the aortic ligation, a high frequency Doppler probe is used to measure the ratio between blood flow velocities in the right and left carotid arteries.3, 4 With surgical survival rates of 80-90%, transverse aortic banding is an effective technique of inducing left ventricular hypertrophy and heart failure in mice.

Transverse aortic constriction (TAC) in the mouse is a commonly used experimental model to study mechanisms underlying cardiac hypertrophy and heart failure development. Here, we describe procedures to constrict the aorta to create a reproducible degree of cardiac hypertrophy in mice.

Transverse aortic constriction (TAC) in the mouse is a commonly used experimental model for pressure overload-induced cardiac hypertrophy and heart ...

Ambulatory ECG Recording in Mice

Telemetric ECG recording in mice is essential to understanding the mechanisms behind arrhythmias, conduction disorders, and sudden cardiac death. Although the surface ECG is utilized for short-term measurements of waveform intervals, it is not practical for long-term studies of heart rate variability or the capture of rare episodes of arrhythmias. Implantable ECG telemeters offer the advantages of simple surgical implantation, long-term recording of electrograms in ambulatory mice, and scalability with simultaneous recordings of multiple animals. Here, we present a step-by-step guide to the implantation of telemeters for ambulatory ECG recording in mice. Careful adherence to aseptic technique is required for favorable survival results with the possibility of implantation and recording from weeks to months. Thus, implantable ECG telemetry is a valuable tool for detection of critical information on cardiac electrophysiology in ambulatory animal models such as the mouse. 

Telemetric ECG has emerged as an essential tool in evaluating animal models for cardiac arrhythmias and sudden cardiac death. Here, we present a stepwise guide to telemetric ECG recordings for application in long-term ambulatory ECG monitoring in mice.

Telemetric ECG recording in mice is essential to understanding the mechanisms behind arrhythmias, conduction disorders, and sudden cardiac death. Although ...

Intravenous Injections in Neonatal Mice

Intravenous injection is a clinically applicable manner to deliver therapeutics. For adult rodents and larger animals, intravenous injections are technically feasible and routine. However, some mouse models can have early onset of disease with a rapid progression that makes administration of potential therapies difficult. The temporal (or facial) vein is just anterior to the ear bud in mice and is clearly visible for the first two days after birth on either side of the head using a dissecting microscope. During this window, the temporal vein can be injected with volumes up to 50 &#956;l. The injection is safe and well tolerated by both the pups and the dams. A typical injection procedure is completed within 1-2 min, after which the pup is returned to the home cage. By the third postnatal day the vein is difficult to visualize and the injection procedure becomes technically unreliable. This technique has been used for delivery of adeno-associated virus (AAV) vectors, which in turn can provide almost body-wide, stable transgene expression for the life of the animal depending on the viral serotype chosen.

Animal models of pediatric disease can experience early onset and aggressive disease progression. Clinically relevant therapy delivery to young mouse models can be technically difficult. This protocol describes a non-invasive intravenous injection method for newborn mice within the first two postnatal days of life.

Intravenous injection is a clinically applicable manner to deliver therapeutics. For adult rodents and larger animals, intravenous injections are ...

Inducing a Site Specific Replication Blockage in E. coli Using a Fluorescent Repressor Operator System

Obstacles present on DNA, including tightly-bound proteins and various lesions, can severely inhibit the progression of the cell&#39;s replication machinery. The stalling of a replisome can lead to its dissociation from the chromosome, either in part or its entirety, leading to the collapse of the replication fork. The recovery from this collapse is a necessity for the cell to accurately complete chromosomal duplication and subsequently divide. Therefore, when the collapse occurs, the cell has evolved diverse mechanisms that take place to restore the DNA fork and allow replication to be completed with high fidelity. Previously, these replication repair pathways in bacteria have been studied using UV damage, which has the disadvantage of not being localized to a known site. This manuscript describes a system utilizing a Fluorescence Repressor Operator System (FROS) to create a site-specific protein block that can induce the stalling and collapse of replication forks in Escherichia coli. Protocols detail how the status of replication can be visualized in single living cells using fluorescence microscopy and DNA replication intermediates can be analyzed by 2-dimensional agarose gel electrophoresis. Temperature sensitive mutants of replisome components (e.g. DnaBts) can be incorporated into the system to induce a synchronous collapse of the replication forks. Furthermore, the roles of the recombination proteins and helicases that are involved in these processes can be studied using genetic knockouts within this system.

Inducing a Site Specific Replication Blockage in E. coli Using a Fluorescent Repressor Operator System

We describe here a system utilizing a site-specific, reversible in vivo protein block to stall and collapse replication forks in Escherichia coli. The establishment of the replication block is evaluated by fluorescence microscopy and neutral-neutral 2-dimensional agarose gel electrophoresis is used to visualize replication intermediates.

Obstacles present on DNA, including tightly-bound proteins and various lesions, can severely inhibit the progression of the cell&#39;s replication ...

Evaluating Vascular Hyperpermeability-inducing Agents in the Skin with the Miles Assay

The primary function of the vascular endothelium in vertebrate organisms is to serve as a barrier between the blood and each tissue of the body, whereby the permeability of the endothelium to blood cells, plasma macromolecules, and water can be adapted according to the physiological need. In certain diseases, cytokines and growth factors are released that target the endothelial barrier to transiently increase vascular permeability; however, their prolonged presence may cause chronic vascular hyperpermeability and thereby tissue-damaging edema. The Miles assay is an in vivo technique that allows researchers to study vascular hyperpermeability through the proxy measurement of vascular leakage. Here, we provide a detailed protocol on how to perform this procedure in the mouse, which is the most widely used model organism to study mammalian physiology and pathology. The procedure involves the intravenous injection of Evans blue dye to label the circulating albumin followed by multiple intradermal injections of permeability-inducing agents and vehicle control solutions into opposing flanks of the mouse. Consequently, Evans blue dye gradually leaks into the dermis, where it accumulates and can be extracted for quantification as leakage induced by the permeability-inducing agent relative to the vehicle. The Miles assay can be performed in wild type or genetically modified mouse models and may be combined with drug administration to study molecular mechanisms that regulate vascular permeability and identify agents/targets capable of inducing or blocking hyperpermeability.

Here, we present a protocol to measure the vascular leakage induced by intradermal administration of permeability promoting agents into the murine skin. This technique can be used to determine the ability of molecules to promote or inhibit vascular leakage or to study the molecular mechanisms that regulate vascular permeability.

The primary function of the vascular endothelium in vertebrate organisms is to serve as a barrier between the blood and each tissue of the body, whereby ...

Culture Methods to Study Apical-Specific Interactions using Intestinal Organoid Models

The lining of the gut epithelium is made up of a simple layer of specialized epithelial cells that expose their apical side to the lumen and respond to external cues. Recent optimization of in vitro culture conditions allows for the re-creation of the intestinal stem cell niche and the development of advanced 3-dimensional (3D) culture systems that recapitulate the cell composition and the organization of the epithelium. Intestinal organoids embedded in an extracellular matrix (ECM) can be maintained for long-term and self-organize to generate a well-defined, polarized epithelium that encompasses an internal lumen and an external exposed basal side. This restrictive nature of the intestinal organoids presents challenges in accessing the apical surface of the epithelium in vitro and limits the investigation of biological mechanisms such as nutrient uptake and host-microbiota/host-pathogen interactions. Here, we describe two methods that facilitate access to the apical side of the organoid epithelium and support the differentiation of specific intestinal cell types. First, we show how ECM removal induces an inversion of the epithelial cell polarity and allows for the generation of apical-out 3D organoids. Second, we describe how to generate 2-dimensional (2D) monolayers from single cell suspensions derived from intestinal organoids, comprised of&#160;mature and differentiated cell types. These techniques provide novel tools to study apical-specific interactions of the epithelium with external cues in vitro and promote the use of organoids as a platform to facilitate precision medicine.

Here we present two protocols that allow the modeling of intestinal apical-specific interactions. Organoid-derived intestinal monolayers and Air-Liquid Interface (ALI) cultures facilitate the generation of well-differentiated epithelia accessible from both luminal and basolateral sides, whereas polarity-inverted intestinal organoids expose their apical side and are amenable to high throughput assays.

The lining of the gut epithelium is made up of a simple layer of specialized epithelial cells that expose their apical side to the lumen and respond to ...

Terminal H-reflex Measurements in Mice

The Hoffmann reflex (H-reflex), as an electrical analog to the stretch reflex, allows electrophysiological validation of the integrity of neural circuits after injuries such as spinal cord damage or stroke. An increase of the H-reflex response, together with symptoms like non-voluntary muscle contractions, pathologically augmented stretch reflex, and hypertonia in the corresponding muscle, is an indicator of post-stroke spasticity (PSS).
In contrast to rather nerve-unspecific transcutaneous measurements, here, we present a protocol to quantify the H-reflex directly at the ulnar and median nerves of the forepaw, which is applicable, with minor modifications, to the tibial and sciatic nerve of the hindpaw. Based on the direct stimulation and the adaptation to different nerves, the method represents a reliable and versatile tool to validate electrophysiological changes in spasticity-related disease models.

The clinical evaluation of spasticity based on the Hoffmann reflex (H-reflex) and using electrical stimulation of peripheral nerves is an established method. Here, we provide a protocol for a terminal and direct nerve stimulation for H-reflex quantification in the mouse forepaw.

The Hoffmann reflex (H-reflex), as an electrical analog to the stretch reflex, allows electrophysiological validation of the integrity of neural circuits ...

Establishing a Diaphyseal Femur Fracture Model in Mice

Bones have a significant regenerative capacity. However, fracture healing is a complex process, and depending on the severity of the lesions and the age and overall health status of the patient, failures can occur, leading to delayed union or nonunion. Due to the increasing number of fractures resulting from high-energy trauma and aging, the development of innovative therapeutic strategies to improve bone repair based on the combination of skeletal/mesenchymal stem/stromal cells and biomimetic biomaterials is urgently needed. To this end, the use of reliable animal models is fundamental to better understanding the key cellular and molecular mechanisms that determine the healing outcomes. Of all the models, the mouse is the preferred research model because it offers a wide variety of transgenic strains and reagents for experimental analysis. However, the establishment of fractures in mice may be technically challenging due to their small size. Therefore, this article aims to demonstrate the procedures for the surgical establishment of a diaphyseal femur fracture in mice, which is stabilized with an intramedullary wire and resembles the most common bone repair process, through cartilaginous callus formation.

This protocol describes a surgical procedure for the establishment of a diaphyseal fracture in the femur of mice, which is stabilized with an intramedullary wire, for fracture healing studies.

Bones have a significant regenerative capacity. However, fracture healing is a complex process, and depending on the severity of the lesions and the age ...