This work was supported by AI101263 from the National Institutes of Health (T.V.B), and by a grant from The Hartwell Foundation (M.L.M). M.L.M. is a member of the Duke Comprehensive Cancer Center.

NOTE: All experimental procedures involving the use of rats were done in accordance with protocols approved by the Animal Care and Use Committee of Duke University.
1. Preparation of the Tracheal Intubation Cannula
<ol>
	<li>Cut off the needle end of a 2 inch 14 G angiocathether needle with cutting pliers.</li>
	<li>Pinch the lumen of the needle closed with needle-nosed pliers.</li>
	<li>Cut the pinched section down to the edge of the open-lumen portion with cutting pliers, and then cut the edges on both sides of the remaining end at 30 - 45&#176; angle to the needle edge.</li>
	<li>Place the bottom 3 - 4 mm of the cannula into the open area of straight slip-joint pliers and pull upwards until a slight upward curve has been formed at the end. 
	NOTE: This will facilitate directing the catheter into the trachea (Figure 1).</li>
	<li>Use fine sandpaper to smooth down the edges at the end of the cannula.</li>
	<li>Return the blunted instrument into the angiocatheter tubing. Ensure that the tubing is slightly shorter than the blunted needle.</li>
</ol>
2. Pre-Surgical Procedures
<ol>
	<li>Set up the site for the procedure by placing an operating microscope set to 10X magnification over the operating area.</li>
	<li>Place a warming pad on the operating area and cover with a clean absorptive pad.</li>
	<li>Set up a heart rate and blood oxygenation monitor near the surgical field.</li>
	<li>Weigh the rat for weight-based medication dosing (e.g., postoperative analgesia, antibiotics, or lymphocyte depleting antibody). 
	NOTE: Typical adult rats weigh between 350 - 450 g.</li>
	<li>Sedate the rat with vaporized 3% isoflurane-O2 (3 L/min) using an induction chamber vented to a waste anesthetic gas scavenging system containing activated charcoal. Allow 5 min for anesthesia induction to provide deep anesthesia necessary for endotracheal intubation.</li>
	<li>Administer Carprofen or Meloxicane (4.4 mg/kg) subcutaneously to the rat prior to proceeding with intubation and surgery.</li>
	<li>Use electric clippers to shave the neck and chest of the anesthetized rat prior to intubation.</li>
</ol>
3. Intubation
<ol>
	<li>Prepare the intubation setup by rubbing a small amount of lubrication (e.g., K-Y gel) on the end of the intubation cannula. Turn on the ventilator to begin the flow of isoflurane gas through the connector tubing.</li>
	<li>Transfer the rat to the intubation apparatus and suspend the rat on the metal bar by its upper incisor teeth (Figure 2A). 
	NOTE: Certain intubation apparatuses may be purchased online. The model shown here is homemade using an acrylic sheet molded around sections of a 2 x 4 inch lumbar piece using a butane torch, and reinforcements are attached with chloroform. The bar (or metal wire) is attached by drilling holes over masking tape placed on the side of the acrylic to prevent splintering or cracking.</li>
	<li>Trans-illuminate the rat&#8217;s neck by positioning a flexible high-intensity light source 1 - 2 cm from the ventral surface of the neck (Figure 2B).</li>
	<li>Use a pair of student standard pattern forceps to pull the tongue gently upwards and to the side of the bottom teeth. Grip the tongue between the thumb and forefinger of one hand while placing the flat inside surface of one side of the forceps against the lower end of the tongue. Press ventrally to expose the epiglottis and aperture of the larynx (Figure 2C).</li>
	<li>Visualize the vocal cords and guide the upturned, blunted end of the intubation cannula anteriorly through the open glottis into the trachea until the hub of the angiocatheter touches the incisors.</li>
	<li>Remove the metal stylet, and attach the anesthesia tubing to the angiocatheter opening to begin ventilation with isoflurane. 
	NOTE: The typical oxygen flow rate to the ventilator is 3 L/min with 3% isoflurane.
	<ol>
		<li>Set the ventilator at a rate of 60 respirations/min in a volume controlled ventilator mode that achieves a pressure of ~12 - 14 mmHg. Use a positive end-expiratory pressure (PEEP) of 3 cm H2O.</li>
	</ol>
	</li>
	<li>Observe bilateral chest wall expansion in sync with the ventilator to ensure proper placement of the endotracheal tube.</li>
	<li>Apply veterinarian-recommended eye ointment to the rat&#8217;s eyes to prevent dryness while under anesthesia.</li>
	<li>Apply veterinarian ointment on the rat&#8217;s eyes to prevent dryness while under anesthesia.</li>
	<li>Secure the endotracheal tube to the head of the rat with a strip of cloth tape. Ensure a firm connection between the endotracheal tube and the anesthetic inflow tubing.</li>
	<li>Attach the blood oxygenation and heart rate monitor to the rat&#8217;s foot and begin monitoring. Confirm proper anesthetization by confirming there is no reaction to toe-pinch.</li>
</ol>
4. Thoracotomy and Thymectomy
<ol>
	<li>Follow standard aseptic technique for the entirety of the procedure.
	<ol>
		<li>Clean the working area and operating table and disinfect with a 70% ethanol solution.</li>
		<li>Use sterile surgical gloves during the procedure, and autoclave all instruments and materials used during the procedure.</li>
		<li>Apply providone-iodine to the entire chest area and allow it to dry. Then clean the surface of the skin with 70% ethanol on gauze. Cover the rat with clear plastic wrap cutting a hole to expose the sterile operating field.</li>
	</ol>
	</li>
	<li>Identify the suprasternal notch in the upper thoracic region. Make a 2 cm midline incision through the skin starting 2 - 3 mm above the suprasternal notch and extending distally midline along the sternum using blunt-tipped Shea scissors.</li>
	<li>Perform a 1.5 cm&#160;median sternotomy from the suprasternal notch again using blunt-tipped Shea scissors. Keep the bottom edge of the scissors just underneath the sternum and advance slowly.</li>
	<li>Insert a small Alm retractor just underneath the separated sternum and open to reveal the pre-tracheal strap muscles (sternohyoid and sternothyroid muscles). Separate the pre-tracheal strap muscles using blunt Graefe forceps.&#160; NOTE: At this point, the trachea can be seen, and the intubation tube should be visualized inside the trachea.</li>
	<li>Place the prongs of the&#160;small Alm retractor underneath the separated strap muscles and sternum. Open the retractor to expose the superior aspect of the thymus.</li>
	<li>Use fine Dumont forceps to release the lateral edges of the thymus tissue and expose the lower thymic lobes.</li>
	<li>Pull the thymus gently superiorly into the open incision site, taking care to avoid contact with the superior vena cava, subclavian and carotid vessels, and to minimize disruption of the delicate pleural lining between the thymus and the lungs.</li>
	<li>As the thymic vessels are dissected and revealed, use micro-scissors to divide them. Use a cotton swab to hold pressure on the vessels for hemostasis if needed.</li>
	<li>Deliver the lower thymic lobes into the incision and then sharply lyse the posterior attachments. Remove the intact thymus and carefully inspect the removed thymus for any missing sections. 
	NOTE: Small thoracic lymph nodes may be visualized surrounding the thymus and are often difficult to differentiate from thymic tissue. These will be discrete round nodes with a similar appearance to thymic tissue but will not be in continuity with the thymus.</li>
	<li>Remove the Alm retractor and close the sternohyoid and sternothyoid muscles with two interrupted 5-0 Maxon sutures.</li>
	<li>Apply 2 drops of surgical grade cyanoacrylate tissue adhesive across the sutures to seal the mediastinum under positive pressure ventilation. 
	NOTE: This will decrease the incidence of pneumothorax and hematoma.</li>
	<li>Close the sternum with two interrupted 4-0 silk sutures on a cutting needle. Insert the needle through gaps between the ribs, taking care to guide the needle just under the sternum to avoid the underlying muscle layer.</li>
	<li>Close the skin layer with a running 4-0 Nylon suture. Discontinue the isoflurane at this point to shorten the post-operative anesthesia recovery period.</li>
	<li>Clean the incision site and surrounding skin with saline-moistened gauze.</li>
	<li>Apply several drops of Bupivicaine (0.25%) for local anesthesia, followed by 1 - 2 drops of cyanoacrylate glue over the incision to seal the incision.</li>
	<li>Continue to ventilate until the rat shows signs of independent respiratory effort and begins to move its extremities. Extubate the rat and allow it to recover under close observation in a cage placed on a warming pad.</li>
	<li>Administer Buprenorphine (0.01 - 0.05 mg/kg) subcutaneously to the rat immediately post-operatively and repeat every 8-12 hr&#160;for 48 hr&#160;post-operatively (normally 5 total doses). At 24 and 48 hr&#160;post-operatively, administer Carprofen or Meloxicane (4.4 mg/kg) subcutaneously (3 total doses including the pre-operative dose).</li>
	<li>Continue to monitor until the rat is moving around the cage. At any signs of respiratory distress, perform re-intubation and re-exploration of the chest.
	<ol>
		<li>Do not leave an animal unattended until it has regained sufficient consciousness to maintain sternal recumbency.</li>
		<li>Do not return an animal that has undergone surgery to the company of other animals until fully recovered.</li>
	</ol>
	</li>
	<li>Remove the skin layer sutures 1 week post-operatively.</li>
</ol>

<ol>
	<li>Hard, G. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Thymectomy+in+the+neonatal+rat.">Thymectomy in the neonatal rat.</a> Lab. Anim. 9 (2), 105-110 (1975).</li><li>Kobayashi, 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=A+technique+for+complete+thymectomy+in+adult+rats.">A technique for complete thymectomy in adult rats.</a> J. Immunol. Methods. 1759 (94), 0 (1994).</li><li>Delrivi&#232;re, L., Gibbs, P., Kobayashi, E., Kamada, N., Gianello, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=New+technique+of+complete+thymectomy+in+adult+rats+without+tracheal+intubation.">New technique of complete thymectomy in adult rats without tracheal intubation.</a> Microsurgery. 18 (1), 6-8 (1998).</li><li>Walker, K. G., Jacques, B. C., Tweedle, J. R., Bradley, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+simpler+technique+for+open+thymectomy+in+adult+rats.">A simpler technique for open thymectomy in adult rats.</a> J. Immunol. Methods. 175 (1), 141 (1994).</li><li>Na, N., Zhao, D., Huang, Z., Hong, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+improved+method+for+rat+intubation+and+thymectomy.">An improved method for rat intubation and thymectomy.</a> Chin. Med. J. (Engl). 124 (17), 2723-2727 (2011).</li><li>Kastl, S., 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=Simplification+of+rat+intubation+on+inclined+metal+plate).">Simplification of rat intubation on inclined metal plate).</a> Adv. Physiol. Educ. 28 (1-4), 29-34 (2004).</li><li>Weksler, B., Ng, B., Lenert, J., Burt, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+simplified+method+for+endotracheal+intubation+in+the+rat.">A simplified method for endotracheal intubation in the rat.</a> J. Appl. Physiol. 76 (4), 1823-1825 (1994).</li><li>Jou, I. 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=Simplified+rat+intubation+using+a+new+oropharyngeal+intubation+wedge.">Simplified rat intubation using a new oropharyngeal intubation wedge.</a> J. Appl. Physiol. 89 (5), 1766-1770 (2000).</li><li>Rivard, A. L., 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=Rat+intubation+and+ventilation+for+surgical+research.">Rat intubation and ventilation for surgical research.</a> J. Invest. Surg. 19 (4), 267-274 (2006).</li><li>Zhao, D., 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=A+model+of+isolated,+vascular+whole+thymus+transplantation+in+nude+rats.">A model of isolated, vascular whole thymus transplantation in nude rats.</a> Transplant Proc. 44 (5), 1394-1398 (2012).</li><li>AbuAttieh, 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=Affinity+maturation+of+antibodies+requires+integrity+of+the+adult+thymus.">Affinity maturation of antibodies requires integrity of the adult thymus.</a> Eur. J. Immunol. 42 (2), 500-510 (2012).</li><li>Reeves, J. P., Reeves, P. A., Chin, L. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Survival+surgery:+removal+of+the+spleen+or+thymus.">Survival surgery: removal of the spleen or thymus.</a> Curr. Protoc. Immunol. 1 (Unit 1.10), (2001).</li><li>Siemionow, M., Izycki, D., Ozer, K., Ozmen, S., Klimczak, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Role+of+thymus+in+operational+tolerance+induction+in+limb+allograft+transplant+model.">Role of thymus in operational tolerance induction in limb allograft transplant model.</a> Transplantation. 81 (11), 1568-1576 (2006).</li><li>Satoh, 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=Immunosuppressive+effect+of+long-term+drainage+of+thoracic+duct+on+immunological+memory+in+adult+thymectomized+rats.+Transplant.">Immunosuppressive effect of long-term drainage of thoracic duct on immunological memory in adult thymectomized rats. Transplant.</a> Proc. 37 (4), 1947-1948 (2005).</li><li>Groen, H., Klatter, F., Pater, J., Nieuwenhuis, P., Rozing, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Temporary,+but+Essential+Requirement+of+CD8++T+Cells+Early+in+the+Pathogenesis+of+Diabetes+in+BB+Rats+as+Revealed+by+Thymectomy+and+CD8+Depletion.">Temporary, but Essential Requirement of CD8+ T Cells Early in the Pathogenesis of Diabetes in BB Rats as Revealed by Thymectomy and CD8 Depletion.</a> Clin. Dev. Immunol. 10 (2-4), 141-151 (2003).</li></ol>

The authors have nothing to disclose.

The present protocol for complete thymectomy provides a minimally invasive approach with a three layer incision closure with surgical glue application designed to minimize complications. Complete removal of the thymus was demonstrated by the loss of na&#239;ve T cells and by IHC staining of mediastinal lymphoid tissue for cytokeratin.
The procedure of adult rat thymectomy has been complicated by mortality rates ranging from 1.5 - 20% due to peri-operative complications, most of which are the a...

Since the early 1960s, the thymus has been recognized for its critical role in the development of central immunological tolerance. Rodent thymectomy has proven to be an essential procedure in defining the role of the thymus in lymphocyte differentiation, self-tolerance, and immunotolerance in the setting of allograft transplantation and tumor metastasis. Removal of the rat thymus allows for studies involving T-cell depletion or the adoptive transfer of defined T cell populations without the re-emergence of native na&#239;ve T cells.
Thymectomies in neonatal rodents can be accomplished using a suction technique with reliable outcomes1. In adult rats, this technique is associated with an approximate 20% mortality rate and frequently results in an incomplete thymectomy2. To consistently achieve complete thymectomy in adult rats, open exposure of the mediastinum through a median sternotomy is required. However, this procedure is associated with complications that include tracheal injury, hemorrhage and pneumothorax leading to an overall mortality rate ranging from 1.5 - 6%2-4.
Over the last two decades improvements in thymectomy techniques have decreased perioperative complications and have improved survival rates. Endotracheal intubation allowing for positive pressure ventilation has lessened pneumothorax rates5. Methods of intubation previously described range from open exposure to the trachea to less invasive methods using direct vocal cord visualization. Complications associated with the intubation procedure include tracheal injury, vocal cord rupture, unintended esophageal intubation, and hemorrhage resulting from cardiac puncture or laceration of the superior vena cava. In addition, close proximity of the lower thymic lobes to the pleural lining can result in pneumothorax.
Here we describe a technique of thymectomy through a minimally invasive 2 cm skin incision following a simple method of endotracheal intubation using a blunted-end angiocatheter and transcutaneous tracheal illumination. The thymectomy involves a 1.5&#160;cm sternotomy and a three-layer closure with surgical glue application to seal the mediastinum and minimize the incidence of hemorrhage and respiratory complications. This method reliably results in complete thymectomy as evidenced by the disappearance of CD4+ and CD8+ na&#239;ve T cells following thymectomy and the absence of thymic tissue on IHC staining. Operative times and peri-procedural mortality are kept to a minimum.

<table border="0" cellpadding="0" cellspacing="0" width="718">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Name of Reagent/ Equipment</td>
			<td style="width:117px;">Company</td>
			<td style="width:152px;">Catalog Number</td>
			<td style="width:167px;">Comments/Description</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">2&#34; 14-gauge angiocatheter</td>
			<td style="width:117px;"></td>
			<td style="width:152px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Operating microscope</td>
			<td style="width:117px;">Zeiss</td>
			<td style="width:152px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Warming pad</td>
			<td style="width:117px;"></td>
			<td style="width:152px;"></td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:283px;">Heart rate and blood oxygenation monitor for rodents with foot sensors</td>
			<td style="width:117px;">Harvard Apparatus</td>
			<td style="width:152px;">ST1 72-8010, ST1 72-8098 (Rat foot sensor)</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Intubation apparatus (plastic with metal bar at the top)</td>
			<td style="width:117px;"></td>
			<td style="width:152px;"></td>
			<td style="width:167px;">See Figure 2&#160;</td>
		</tr>
		<tr height="67">
			<td height="67" style="height:67px;width:283px;">Small animal anesthesia system with induction box, isoflurane tank and O2 tank</td>
			<td style="width:117px;">Harvard Apparatus</td>
			<td style="width:152px;">ST1 72-6420</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="105">
			<td height="105" style="height:105px;width:283px;">Small animal ventilator with tubing</td>
			<td style="width:117px;">CWE</td>
			<td style="width:152px;">12-02000 (ventilator)and 12-04000 (external valve assembly for mice/rats)</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">High-intensity Fiber-Optic Illuminator</td>
			<td style="width:117px;">Dolan Jenner</td>
			<td style="width:152px;">EEG 2823M</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Student standard pattern forceps</td>
			<td style="width:117px;">Fine Science Tools</td>
			<td style="width:152px;">91100-16</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Fine straight scissors</td>
			<td style="width:117px;">Fine Science Tools</td>
			<td style="width:152px;">14060-09</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Blunt-tipped Shea scissors</td>
			<td style="width:117px;">Fine Science Tools</td>
			<td style="width:152px;">14105-12</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Small Alm retractor (for sternum)</td>
			<td style="width:117px;">Fine Science Tools</td>
			<td style="width:152px;">17008-07</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Blunt Graefe forceps</td>
			<td style="width:117px;">Fine Science Tools</td>
			<td style="width:152px;">11050-10</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Fine Dumont forceps</td>
			<td style="width:117px;">Fine Science Tools</td>
			<td style="width:152px;">11254-20</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">5-0 Maxon sutures</td>
			<td style="width:117px;">Ethicon</td>
			<td style="width:152px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">4-0 Silk sutures (with cutting needle)</td>
			<td style="width:117px;">Ethicon</td>
			<td style="width:152px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">6-0 Nylon suture</td>
			<td style="width:117px;">Ethicon</td>
			<td style="width:152px;"></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:283px;">Cyanoacrylate glue (Endermil)</td>
			<td style="width:117px;"></td>
			<td style="width:152px;"></td>
			<td></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:283px;">Lubrication gel</td>
			<td style="width:117px;">Akorn Animal Health</td>
			<td style="width:152px;">NDC 17478-162-35</td>
			<td></td>
		</tr>
	</tbody>
</table>

complete thymectomy in adult rats with non-invasive endotracheal intubation

Thymectomy in neonatal rodents is an established and reliable procedure for immunological studies. However, in adult rats, complications of hemorrhage and pneumothorax from pleural disruption can result in a significant mortality rate. This protocol is a simple method of rat thymectomy that utilizes a mini-sternotomy and endotracheal intubation. Intubation is accomplished with a non-invasive and easily reproducible method and allows for positive pressure ventilation to prevent pneumothorax and a controlled airway that allows sufficient time for careful thymus dissection to minimize pleural disruption. A 1.5 cm sternal incision decreases contact with mediastinal vessels and pleura, while still providing full visualization of the thymus. Following exposure of the mediastinum, the thymus is removed by blunt dissection under magnification. The pleural space is then sealed by suture closure of the pre-tracheal muscles followed by the application of surgical glue. The thorax is then closed by suture closure of the sternum, followed by suture closure of the skin. All thymectomies were complete as evidenced by immunohistochemical (IHC) staining of mediastinal tissue, and absence of na&#239;ve T-cells by flow cytometry, and the procedure had a 96% survival rate. This method is suitable when complete thymectomy with minimal complications is desired for further immunological studies in athymic adult rats.

This procedure was performed on adult Lewis rats (n = 26). The mean operation time was 15 &#177; 3 min. There was no intraoperative mortality. The mean intubation time &#8211; from placement of rat on the intubation apparatus to establishment of ventilation &#8211; was 45 &#177; 5 sec. 24 rats had a full recovery from the operation with no evidence of respiratory difficulties or hemorrhage through post-operative day (POD) 14. One rat developed respiratory difficulty on POD 4 and underwent re-operation to explore the medi...

Watch this Scientific Journal Video about Complete Thymectomy in Adult Rats with Non-invasive Endotracheal Intubation at JoVE.com

Complete Thymectomy in Adult Rats with Non-invasive Endotracheal Intubation

Rodent thymectomy is a valuable technique in immunological research. Here, a protocol for complete thymectomy in adult rats using a mini-sternotomy along with non-invasive intubation and positive pressure ventilation to minimize perioperative morbidity and mortality is described.

Thymectomy in neonatal rodents is an established and reliable procedure for immunological studies. However, in adult rats, complications of hemorrhage and ...

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Immunology and Infection

16.4K Views.  Duke University Medical Center. Rodent thymectomy is a valuable technique in immunological research. Here, a protocol for complete thymectomy in adult rats using a mini-sternotomy along with non-invasive intubation and positive pressure ventilation to minimize perioperative morbidity and mortality is described. 

Video: Complete Thymectomy in Adult Rats with Non-invasive Endotracheal Intubation

Non-invasive Imaging of Leukocyte Homing and Migration in vivo

Two-photon (2P) microscopy is a high resolution imaging technique that has been broadly adapted by biologists. The value of 2P microscopy is that it provides rich spatiotemporal information regarding cell behaviors within intact tissues and in live mice. Leukocyte recruitment plays a significant role in host defense against infection and when unchecked, can contribute to inflammatory and autoimmune diseases. Studying leukocyte recruitment in vivo is technically challenging since cells are moving rapidly within vessels located deep within light scattering tissues. To date, most intravital imaging studies require surgical preparation to expose the blood vessels and tissues. To avoid the tissue damage and inflammation induced by surgery itself, here, we describe a non-invasive single-cell imaging approach that can be used to study leukocyte trafficking in the mouse footpad and phalanges. We discuss the technical aspects of our 2P imaging preparation and walk the reader through a typical experiment from initial set up to execution and data collection.

Non-invasive Imaging of Leukocyte Homing and Migration in vivo

Here, we describe a non-invasive two-photon (2P) microscopy approach to study leukocyte homing in the mouse footpad. We discuss the technical aspects of our tissue imaging preparation and walk the reader through a typical experiment from initial set up to execution and data collection.

Two-photon (2P) microscopy is a high resolution imaging technique that has been broadly adapted by biologists.  The value of 2P microscopy is that it ...

Non-invasive Imaging of Disseminated Candidiasis in Zebrafish Larvae

Disseminated candidiasis caused by the pathogen Candida albicans is a clinically important problem in hospitalized individuals and is associated with a 30 to 40% attributable mortality6. Systemic candidiasis is normally controlled by innate immunity, and individuals with genetic defects in innate immune cell components such as phagocyte NADPH oxidase are more susceptible to candidemia7-9. Very little is known about the dynamics of C. albicans interaction with innate immune cells in vivo. Extensive in vitro studies have established that outside of the host C. albicans germinates inside of macrophages, and is quickly destroyed by neutrophils10-14. In vitro studies, though useful, cannot recapitulate the complex in vivo environment, which includes time-dependent dynamics of cytokine levels, extracellular matrix attachments, and intercellular contacts10, 15-18. To probe the contribution of these factors in host-pathogen interaction, it is critical to find a model organism to visualize these aspects of infection non-invasively in a live intact host. 
The zebrafish larva offers a unique and versatile vertebrate host for the study of infection. For the first 30 days of development zebrafish larvae have only innate immune defenses2, 19-21, simplifying the study of diseases such as disseminated candidiasis that are highly dependent on innate immunity. The small size and transparency of zebrafish larvae enable imaging of infection dynamics at the cellular level for both host and pathogen. Transgenic larvae with fluorescing innate immune cells can be used to identify specific cells types involved in infection22-24. Modified anti-sense oligonucleotides (Morpholinos) can be used to knock down various immune components such as phagocyte NADPH oxidase and study the changes in response to fungal infection5. In addition to the ethical and practical advantages of using a small lower vertebrate, the zebrafish larvae offers the unique possibility to image the pitched battle between pathogen and host both intravitally and in color. 
The zebrafish has been used to model infection for a number of human pathogenic bacteria, and has been instrumental in major advances in our understanding of mycobacterial infection3, 25. However, only recently have much larger pathogens such as fungi been used to infect larva5, 23, 26, and to date there has not been a detailed visual description of the infection methodology. Here we present our techniques for hindbrain ventricle microinjection of prim25 zebrafish, including our modifications to previous protocols. Our findings using the larval zebrafish model for fungal infection diverge from in vitro studies and reinforce the need to examine the host-pathogen interaction in the complex environment of the host rather than the simplified system of the Petri dish5.

The rapid development, small size and transparency of zebrafish are tremendous advantages for the study of innate immune control of infection1-4. Here we demonstrate techniques for infecting zebrafish larvae using the fungal pathogen Candida albicans by microinjection, methodology recently used to implicate phagocyte NADPH oxidase activity in control of fungal dimorphism5.

Disseminated candidiasis caused by the pathogen Candida albicans is a clinically important problem in hospitalized individuals and is associated with a 30 ...

Non-invasive Optical Imaging of the Lymphatic Vasculature of a Mouse

The lymphatic vascular system is an important component of the circulatory system that maintains fluid homeostasis, provides immune surveillance, and mediates fat absorption in the gut. Yet despite its critical function, there is comparatively little understanding of how the lymphatic system adapts to serve these functions in health and disease1. Recently, we have demonstrated the ability to dynamically image lymphatic architecture and lymph "pumping" action in normal human subjects as well as in persons suffering lymphatic dysfunction using trace administration of a near-infrared fluorescent (NIRF) dye and a custom, Gen III-intensified imaging system2-4. NIRF imaging showed dramatic changes in lymphatic architecture and function with human disease. It remains unclear how these changes occur and new animal models are being developed to elucidate their genetic and molecular basis. In this protocol, we present NIRF lymphatic, small animal imaging5,6 using indocyanine green (ICG), a dye that has been used for 50 years in humans7, and a NIRF dye-labeled cyclic albumin binding domain (cABD-IRDye800) peptide that preferentially binds mouse and human albumin8. Approximately 5.5 times brighter than ICG, cABD-IRDye800 has a similar lymphatic clearance profile and can be injected in smaller doses than ICG to achieve sufficient NIRF signals for imaging8. Because both cABD-IRDye800 and ICG bind to albumin in the interstitial space8, they both may depict active protein transport into and within the lymphatics. Intradermal (ID) injections (5-50 &mu;l) of ICG (645 &mu;M) or cABD-IRDye800 (200 &mu;M) in saline are administered to the dorsal aspect of each hind paw and/or the left and right side of the base of the tail of an isoflurane-anesthetized mouse. The resulting dye concentration in the animal is 83-1,250 &mu;g/kg for ICG or 113-1,700 &mu;g/kg for cABD-IRDye800. Immediately following injections, functional lymphatic imaging is conducted for up to 1 hr using a customized, small animal NIRF imaging system. Whole animal spatial resolution can depict fluorescent lymphatic vessels of 100 microns or less, and images of structures up to 3 cm in depth can be acquired9. Images are acquired using V++ software and analyzed using ImageJ or MATLAB software. During analysis, consecutive regions of interest (ROIs) encompassing the entire vessel diameter are drawn along a given lymph vessel. The dimensions for each ROI are kept constant for a given vessel and NIRF intensity is measured for each ROI to quantitatively assess "packets" of lymph moving through vessels.

Recently developed imaging techniques using near-infrared fluorescence (NIRF) may help elucidate the role the lymphatic system plays in cancer metastasis, immune response, wound repair, and other lymphatic-associated diseases.

The lymphatic  vascular system is an important component of the circulatory system that  maintains fluid homeostasis, provides immune surveillance, and ...

Culturing Microglia from the Neonatal and Adult Central Nervous System

Microglia are the resident macrophage-like cells of the central nervous system (CNS) and, as such, have critically important roles in physiological and pathological processes such as CNS maturation in development, multiple sclerosis, and spinal cord injury. Microglia can be activated and recruited to action by neuronal injury or stimulation, such as axonal damage seen in MS or ischemic brain trauma resulting from stroke. These immunocompetent members of the CNS are also thought to have roles in synaptic plasticity under non-pathological conditions. We employ protocols for culturing microglia from the neonatal and adult tissues that are aimed to maximize the viable cell numbers while minimizing confounding variables, such as the presence of other CNS cell types and cell culture debris. We utilize large and easily discernable CNS components (e.g. cortex, spinal cord segments), which makes the entire process feasible and reproducible. The use of adult cells is a suitable alternative to the use of neonatal brain microglia, as many pathologies studied mainly affect the postnatal spinal cord. These culture systems are also useful for directly testing the effect of compounds that may either inhibit or promote microglial activation. Since microglial activation can shape the outcomes of disease in the adult CNS, there is a need for in vitro systems in which neonatal and adult microglia can be cultured and studied.

We outline methods for the efficient and quick isolation/culture of viable microglia from the neonatal cerebral cortex and adult spinal cord. The dissection and plating of cortical microglia can be accomplished within 90 minutes, with the subsequent microglial harvest taking place ~ 10 days following the initial dissection.

Microglia are the resident macrophage-like cells of the central nervous system (CNS) and, as such, have critically important roles in physiological and ...

Non-Invasive Model of Neuropathogenic Escherichia coli Infection in the Neonatal Rat

Investigation of the interactions between animal host and bacterial pathogen is only meaningful if the infection model employed replicates the principal features of the natural infection. This protocol describes procedures for the establishment and evaluation of systemic infection due to neuropathogenic Escherichia coli K1 in the neonatal rat. Colonization of the gastrointestinal tract leads to dissemination of the pathogen along the gut-lymph-blood-brain course of infection and the model displays strong age dependency. A strain of E. coli O18:K1 with enhanced virulence for the neonatal rat produces exceptionally high rates of colonization, translocation to the blood compartment and invasion of the meninges following transit through the choroid plexus. As in the human host, penetration of the central nervous system is accompanied by local inflammation and an invariably lethal outcome. The model is of proven utility for studies of the mechanism of pathogenesis, for evaluation of therapeutic interventions and for assessment of bacterial virulence.

Non-Invasive Model of Neuropathogenic Escherichia coli Infection in the Neonatal Rat

Here, a procedure is described for the establishment of systemic infection in the neonatal rat with cultures of Escherichia coli K1. This non-invasive procedure permits colonization of the gastrointestinal tract, translocation of the pathogen to the systemic circulation, and invasion of the central nervous system at the choroid plexus.

Investigation of the interactions between animal host and bacterial pathogen is only meaningful if the infection model employed replicates the principal ...

Non-invasive Imaging of the Innate Immune Response in a Zebrafish Larval Model of Streptococcus iniae Infection

The aquatic pathogen, Streptococcus iniae, is responsible for over 100 million dollars in annual losses for the aquaculture industry and is capable of causing systemic disease in both fish and humans. A better understanding of S. iniae disease pathogenesis requires an appropriate model system. The genetic tractability and the optical transparency of the early developmental stages of zebrafish allow for the generation and non-invasive imaging of transgenic lines with fluorescently tagged immune cells. The adaptive immune system is not fully functional until several weeks post fertilization, but zebrafish larvae have a conserved vertebrate innate immune system with both neutrophils and macrophages. Thus, the generation of a larval infection model allows the study of the specific contribution of innate immunity in controlling S. iniae infection.
The site of microinjection will determine whether an infection is systemic or initially localized. Here, we present our protocols for otic vesicle injection of zebrafish aged 2-3 days post fertilization as well as our techniques for fluorescent confocal imaging of infection. A localized infection site allows observation of initial microbe invasion, recruitment of host cells and dissemination of infection. Our findings using the zebrafish larval model of S. iniae infection indicate that zebrafish can be used to examine the differing contributions of host neutrophils and macrophages in localized bacterial infections. In addition, we describe how photolabeling of immune cells can be used to track individual host cell fate during the course of infection.

Non-invasive Imaging of the Innate Immune Response in a Zebrafish Larval Model of Streptococcus iniae Infection

Here, we present a protocol for the generation and imaging of a localized bacterial infection in the zebrafish otic vesicle.

The aquatic pathogen, Streptococcus iniae, is responsible for over 100 million dollars in annual losses for the aquaculture industry and is capable of ...

A Non-invasive and Technically Non-intensive Method for Induction and Phenotyping of Experimental Bacterial Pneumonia in Mice

Although community-acquired pneumonia remains a major public health problem, murine models of bacterial pneumonia have recently facilitated significant preclinical advances in our understanding of the underlying cellular and molecular pathogenesis. In vivo mouse models capture the integrated physiology and resilience of the host defense response in a manner not revealed by alternative, simplified ex vivo approaches. Several methods have been described in the literature for intrapulmonary inoculation of bacteria in mice, including aerosolization, intranasal delivery, peroral endotracheal cannulation under &#39;blind&#39; and visualized conditions, and transcutaneous endotracheal cannulation. All methods have relative merits and limitations. Herein, we describe in detail a non-invasive, technically non-intensive, inexpensive, and rapid method for intratracheal delivery of bacteria that involves aspiration (i.e., inhalation) by the mouse of an infectious inoculum pipetted into the oropharynx while under general anesthesia. This method can be used for pulmonary delivery of a wide variety of non-caustic biological and chemical agents, and is relatively easy to learn, even for laboratories with minimal prior experience with pulmonary procedures. In addition to describing the aspiration pneumonia method, we also provide step-by-step procedures for assaying the subsequent in vivo pulmonary innate immune response of the mouse, in particular, methods for quantifying bacterial clearance and the cellular immune response of the infected airway. This integrated and simple approach to pneumonia assessment allows for rapid and robust evaluation of the effect of genetic and environmental manipulations upon pulmonary innate immunity.

Several methods have been described in the literature for modeling bacterial pneumonia in mice. Herein, we describe a non-invasive, inexpensive, rapid method for inducing pneumonia via aspiration (i.e., inhalation) of a bacterial inoculum pipetted into the oropharynx. Downstream methods for assessment of the pulmonary innate immune response are also detailed.

Although community-acquired pneumonia remains a major public health problem, murine models of bacterial pneumonia have recently facilitated significant ...

Non-invasive In Vivo Fluorescence Optical Imaging of Inflammatory MMP Activity Using an Activatable Fluorescent Imaging Agent

This paper describes a non-invasive method for imaging matrix metalloproteinases (MMP)-activity by an activatable fluorescent probe, via in vivo fluorescence optical imaging (OI), in two different mouse models of inflammation: a rheumatoid arthritis (RA) and a contact hypersensitivity reaction (CHR) model. Light with a wavelength in the near infrared (NIR) window (650 - 950 nm) allows a deeper tissue penetration and minimal signal absorption compared to wavelengths below 650 nm. The major advantages using fluorescence OI is that it is cheap, fast and easy to implement in different animal models.
Activatable fluorescent probes are optically silent in their inactivated states, but become highly fluorescent when activated by a protease. Activated MMPs lead to tissue destruction and play an important role for disease progression in delayed-type hypersensitivity reactions (DTHRs) such as RA and CHR. Furthermore, MMPs are the key proteases for cartilage and bone degradation and are induced by macrophages, fibroblasts and chondrocytes in response to pro-inflammatory cytokines. Here we use a probe that is activated by the key MMPs like MMP-2, -3, -9 and -13 and describe an imaging protocol for near infrared fluorescence OI of MMP activity in RA and control mice 6 days after disease induction as well as in mice with acute (1x challenge) and chronic (5x challenge) CHR on the right ear compared to healthy ears.

Non-invasive In Vivo Fluorescence Optical Imaging of Inflammatory MMP Activity Using an Activatable Fluorescent Imaging Agent

This paper explains the application of fluorescent imaging using an activatable optical imaging probe to visualize the in vivo activity of key matrix metalloproteinases in two different experimental models of inflammation.

This paper describes a non-invasive method for imaging matrix metalloproteinases (MMP)-activity by an activatable fluorescent probe, via in vivo ...

Analysis of Lymphocyte Extravasation Using an In Vitro Model of the Human Blood-brain Barrier

Lymphocyte extravasation into the central nervous system (CNS) is critical for immune surveillance. Disease-related alterations of lymphocyte extravasation might result in pathophysiological changes in the CNS. Thus, investigation of lymphocyte migration into the CNS is important to understand inflammatory CNS diseases and to develop new therapy approaches. Here we present an in vitro model of the human blood-brain barrier to study lymphocyte extravasation. Human brain microvascular endothelial cells (HBMEC) are confluently grown on a porous polyethylene terephthalate transwell insert to mimic the endothelium of the blood-brain barrier. Barrier function is validated by zonula occludens immunohistochemistry, transendothelial electrical resistance (TEER) measurements as well as analysis of evans blue permeation. This model allows investigation of the diapedesis of rare lymphocyte subsets such as CD56brightCD16dim/- NK cells. Furthermore, the effects of other cells, cytokines and chemokines, disease-related alterations, and distinct treatment regimens on the migratory capacity of lymphocytes can be studied. Finally, the impact of inflammatory stimuli as well as different treatment regimens on the endothelial barrier can be analyzed.

Analysis of Lymphocyte Extravasation Using an In Vitro Model of the Human Blood-brain Barrier

Here, we describe a human blood-brain barrier model enabling to investigate lymphocyte transmigration into the central nervous system in vitro.

Lymphocyte extravasation into the central nervous system (CNS) is critical for immune surveillance. Disease-related alterations of lymphocyte ...

A Non-invasive Way to Isolate and Phenotype Cells from the Conjunctiva

Traditionally, ocular surface cytology is studied with techniques such as spatula technology and brush technology. The problem with these techniques is that they may induce traumatic lesions on the surface of the eye, which can progress to scarring, eyelid deformity, limbal stem cell deficiency and in some cases, cause great discomfort to the subject. To avoid these clinical problems, impression cytology (IC) was developed to diagnose dry eye disease and later neoplasia, atopic disease, vernal keratoconjunctivitis and keratoconjunctivitis sicca. Typically, clinicians manually cut filter papers into required shapes and apply these to the ocular surface. Here, we describe how to perform IC using a commercially available medical device. This technique is explained here followed by immunophenotyping by flow cytometry. This technique requires less manual handling and causes less injury to the ocular surface.

The exposed normal ocular surface consists of cornea and conjunctiva. Epithelial cells, goblet cells and immune cells are present in the conjunctiva. Here, a non-invasive, technique of impression cytology is described using an impression cytology device and flow cytometry to analyze immune cells in the conjunctiva.

Traditionally, ocular surface cytology is studied with techniques such as spatula technology and brush technology. The problem with these techniques is ...