Supported by NIH HL-10342.

1. Preparing for the Procedure
One must first obtain and prepare the following items:
<ol>
	<li>The cannula. For intubation of 20-35 g mice, we use a 1 or 1.5 inch long, 20 gauge IV catheter (BD Insylte, Sparks, MD or Jelco Optiva, Carlsbad, CA). A new sterile catheter can be used for each mouse, but catheters can also be reused after sterilization by soaking in 70% ethanol overnight. Although neither the pharynx nor trachea of the mouse is sterile proper cleanliness procedures, including the use of sterile gloves and instruments, should be followed.</li>
	<li>The fiberoptic cable. We use &asymp;70 cm of 0.5 mm optical cable from Edmund Optics, but the length is not critical. It is important to make sure the fiber has its edge smoothed, since after cutting the cable to length with a razor, the edge is left relatively sharp, and it does not take much effort to pierce the tracheal wall. However, it is very easy to smooth this edge by holding the fiber about 2 cm from the end and then making small circles for a few seconds with the edges of the tip touching a piece of 1,000 grit emery paper (see demonstration in the video and Figure 1 of Macdonald, et al. 10). The other end is inserted through a rubber stopper. This is most easily accomplished by first pushing an 18-gauge needle through the stopper, inserting the optical fiber through the needle bore, then withdrawing the needle. The rubber stopper is connected to a 150 watt halogen light source (e.g. NCL-150, Volpi USA, or any other or light source, even less than 150 watts). It is important to make sure to use a stopper made of silicone rubber (or other heat resistant material), since ordinary rubber or cork may burn when located so close to the hot light source.</li>
</ol>
2. Performing the Intubation
<ol>
	<li>See Figures 1 and 2. Insert fiber optic cable through a short piece of silicone rubber tubing (&asymp;0.8 mm ID x 4 mm OD, Cole-Palmer, EW-96410-13). Tie this rubber tube fairly tight, while still allowing the fiber optic cable to be adjusted. Inserting the silicone tubing snugly in the Luer end of the cannula fixes the fiber optic cable's position inside the cannula. Adjust the position of the fiberoptic cable so that it extends through the cannula &asymp;4 mm in front of the cannula tip.</li>
	<li>Place the anesthetized mouse on a vertical support, suspended by its upper incisors (Figure 3). Most investigators find the best visualization with the ventral side of the mouse facing themselves. Very gently pull out the tongue and hold with thumb and forefinger. The middle finger is placed between the neck and plastic support. Traction on the tongue with the index finger and thumb is used to open the mouth, and to straighten the intubation path, the angle of the head is adjusted with the middle finger behind the neck shown in Figure 3.</li>
	<li>Using the fiber optic cable as a light source and introducer, push it through the visualized vocal cords. If the cords are not visible, gently pull harder on the tongue using the middle finger as support. When inserted, advance the cannula &asymp;5 mm further. Then, being very careful not to move the cannula, withdraw the fiber optic cable. Lie the mouse down and secure the cannula with a piece of tape and support the cannula hub on a piece of Plasticine (modeling clay), as shown in Figure 4.</li>
	<li>The procedure in step 3 cannot be easily taught or even demonstrated, since it is a solo operation. However, by subtle adjustments of the traction on the tongue and the support behind the head nearly all who try this soon find the right way to position the mouse to visualize the vocal cords.</li>
</ol>

<ol>
	<li>Brown, R. H., Walters, D. M., Greenberg, R. S., Mitzner, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+method+of+endotracheal+intubation+and+pulmonary+functional+assessment+for+repeated+studies+in+mice.">A method of endotracheal intubation and pulmonary functional assessment for repeated studies in mice.</a> J. Appl. Physiol. 87, 2362-2365 (1999).</li><li>Walters, D. M., Wills-Karp, M., Mitzner, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Assessment+of+cellular+profile+and+lung+function+with+repeated+bronchoalveolar+lavage+in+individual+mice.">Assessment of cellular profile and lung function with repeated bronchoalveolar lavage in individual mice.</a> Physiol. Genomics. 2, 29-36 (2000).</li><li>Rivera, B., Miller, S., Brown, E., Price, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+novel+method+for+endotracheal+intubation+of+mice+and+rats+used+in+imaging+studies.">A novel method for endotracheal intubation of mice and rats used in imaging studies.</a> Contemporary Topics in Laboratory Animal Science / American Association for Laboratory Animal Science. 44, 52-55 (2005).</li><li>Hamacher, J., 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=Microscopic+wire+guide-based+orotracheal+mouse+intubation:+description,+evaluation+and+comparison+with+transillumination.">Microscopic wire guide-based orotracheal mouse intubation: description, evaluation and comparison with transillumination.</a> Laboratory Animals. 42, 222-230 (2008).</li><li>Spoelstra, E. 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=A+novel+and+simple+method+for+endotracheal+intubation+of+mice.">A novel and simple method for endotracheal intubation of mice.</a> Laboratory Animals. 41, 128-135 (2007).</li><li>Zhao, X., 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+retrograde+intubation+in+mice.">A technique for retrograde intubation in mice.</a> Lab Animal. 35, 39-42 (2006).</li><li>Vergari, A., 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+new+method+of+orotracheal+intubation+in+mice.">A new method of orotracheal intubation in mice.</a> European Review for Medical and Pharmacological Sciences. 8, 103-106 (2004).</li><li>Vergari, A., Polito, A., Musumeci, M., Palazzesi, S., Marano, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Video-assisted+orotracheal+intubation+in+mice.">Video-assisted orotracheal intubation in mice.</a> Laboratory Animals. 37, 204-206 (2003).</li><li>Hastings, R. H., Summers-Torres, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Direct+Laryngoscopy+in+Mice.">Direct Laryngoscopy in Mice.</a> Contemporary Topics in Laboratory Animal Science / American Association for Laboratory Animal Science. 38, 33-35 (1999).</li><li>MacDonald, K. D., Chang, H. Y., Mitzner, W. <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+simple+method+of+mouse+lung+intubation.">An improved simple method of mouse lung intubation.</a> J. Appl. Physiol. 106, 984-987 (2009).</li><li>Ewart, S. L., Levitt, R. C., Mitzner, W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Respiratory+system+mechanics+in+mice+measured+by+end-inflation+occlusion.">Respiratory system mechanics in mice measured by end-inflation occlusion.</a> Journal of Applied Physiology. 79, 560-566 (1995).</li><li>MacDonald, K. D., McKenzie, K. R., Mitzner, W., Zeitlin, P. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lung+Mechanics+in+Heterozygous+CF+Mice+after+Repeated+LPS+Dosing.">Lung Mechanics in Heterozygous CF Mice after Repeated LPS Dosing.</a> Am. J. Respir. Crit. Care Med. 175 (4), A930 (2007).</li></ol>

None of the authors have any conflicts of interest to disclose.

The procedure described here has several advantages. First the apparatus is simple and relatively inexpensive.. The fabrication of the apparatus does not require any special tools or costly equipment. The use of a catheter introduces that also is the light source means that one never loses sight of the tracheal opening as the introducer approaches the tracheal opening. The use of a 0.5 mm introducer also serves to minimize trauma that might occur with an initial insertion of a larger cannula. We note here that a similar optical probe is available from a commercial vendor (Braintree Scientific, Braintree, MA). Their device uses a battery powered light source and optical fiber.
In the present work, we tested the procedure with repeat measurement of lung mechanics, but such intubation could just as easily be used to instill chemicals or cells into the lung, as has been described for repeated delivery of LPS 12. In addition, a prior report with a more primitive intubation procedure described the ability to do repeated BAL in individual mice 2, and this would be much more simply accomplished with the new intubation approach.
In practice, the method described here has been easily taught to fellows, students, and technicians who had never attempted intubation. In fact, during group training sessions, some of the students become sufficiently proficient to then teach some of the other students who had not yet tried it. This method thus has a considerable advantage as it minimizes the number of mice needed for practice and should allow minimal damage in repeated studies.
In doing the intubation, there are several practical issues that should be mentioned. It is important to be as gentle as possible with the retraction of the tongue in the initial opening of the mouth. If unprotected forceps are used it is easy to injure the tongue, and this can lead to death of the mouse. In first learning how to do the intubation, the most important thing is the use of the finger behind the neck to adjust the angle of the head to enable visualization of the tracheal opening. When done correctly, with sufficient traction on the tongue, the vocal cords can easily be seen. It is this initial visualization step that usually requires the most time, since once the tracheal opening is seen, it is relatively simple to insert the fiber cable and intravenous catheter. Initially if there is an issue with this visualization, the trainee is often not pulling on the tongue with sufficient force. Increasing this pull slightly will straighten the visualization path so the vocal cords can be seen. Hamacher, et al., described a unique intubation system with microscopic visualization 4. Their online video of this intubation is excellent and very instructive, although the means of positioning the head and neck is not entirely clear from the video and figure. While the system they describe seems to work very efficiently, it requires a dedicated microscope. Using the system and procedure we describe, the vocal cords and tracheal opening can be seen with the naked eye. In our original description of this method 10, we described a procedure to add a cone to the intubation cannula. This cone wedges into the narrow mouse pharynx and prevents the cannula from being inserted too deeply. We have found that this wedge is useful in teaching students the procedure, since it is very easy to insert the cannula to the carina or beyond, possibly penetrating through an airway wall. Simple instructions to fabricate the wedge can be found in that paper. However, once someone learns the procedure sufficiently well and where to position the cannula, this adaptation is no longer needed. 
Finally, we should note that we have only tested this procedure with the 20 g cannula in young adult mice of a few strains. In this situation, we have validated that the trachea and vocal cords can provide a very good seal around the cannula with normal ventilation pressures, i.e. there is minimal air leakage out of the lung with mechanical ventilation. Figure 6 shows results from on cannulation in a C57BL/6 mouse, where 3 increasing air boluses (0.5, 0.75, and 1 ml) were used to inflate the lungs. It is clear from this figure than pressure leaks are minimal to an airway pressure of at least 15 cmH2O. However, if one uses substantially younger or older mice, or mice from strains with a different lung anatomy, then it would be wise to confirm that there are minimal leaks. If there are, the procedure may then require use of a different size cannula.
In summary, the intubation procedure describe here is inexpensive to fabricate and simple to use, and it should enable most investigators and laboratory technicians to quickly learn to successfully intubate mice with relatively little experience.

In 1999, Brown et al. published a paper describing a method for intubation of the mouse lung 1. Such a technique has considerable utility in doing repeat pulmonary function or bronchoalveolar lavage in individual mice in longitudinal studies 2. Since that original paper, there have been several other papers that have described different approaches to mouse intubation 3-9. While all of these methods can be used successfully, they usually require considerable training or cost. One of the main issues with such intubation is that as the intubation cannula approaches nears the trachea pending insertion, the cannula itself blocks the light and hence the visualization of where it needs to go. Thus, the insertion becomes blind at the most critical time. In this paper we show how to simply and inexpensively eliminate this visualization problem, thereby ensuring successful intubation with relatively little training or experience.

<table border="1">
	<tbody>
		<tr>
			<td>Material</td>
			<td>Company</td>
			<td>Catalog #</td>
			<td>Comment</td>
		</tr>
		<tr>
			<td>Intubation cannula</td>
			<td>BD Insylte, Sparks, MD or Jelco Optiva, Carlsbad, CA</td>
			<td>&nbsp;</td>
			<td>1-in.-long, 20-gauge intravenous (IV) catheter</td>
		</tr>
		<tr>
			<td>Fiber-optic cable</td>
			<td>Edmund Optics, Barrington, NJ</td>
			<td>#NT02-542</td>
			<td>Approximately 2-ft length of 0.5-mm optical fiber (Communication grade plastic fiber). The edge of the fiber end that is inserted into the trachea should be gently rounded by holding the fiber &asymp;2 cm from the end and then making small circles while dragging the tip for a few seconds on 1,000-grit emory paper.</td>
		</tr>
		<tr>
			<td>Light Source</td>
			<td>Volpi</td>
			<td>NCL-150</td>
			<td>Although we use a 150-W halogen light source, any equivalent light source, even with much lower wattage can be used.</td>
		</tr>
		<tr>
			<td>Silicone rubber stopper</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>A #1 silicone rubber stopper fits the light source used in this demonstration. Different light sources may require a different size.</td>
		</tr>
		<tr>
			<td>Small silicone rubber tube</td>
			<td>Cole-Palmer</td>
			<td>EW-96410-13</td>
			<td>A &asymp;1.5 cm piece of silicone rubber tubing (0.8 inner diameter, 4 mm outer diameter)</td>
		</tr>
		<tr>
			<td>Angled support stand</td>
			<td>&nbsp;</td>
			<td>&nbsp;</td>
			<td>Ours is constucted from plexiglass, but any material to which a thread or wire can be affixed to hold the mouse at an almost vertical angle can be used.</td>
		</tr>
	</tbody>
</table>

a simple method of mouse lung intubation

A simple procedure to intubate mice for pulmonary function measurements would have several advantages in longitudinal studies with limited numbers or expensive animal. One of the reasons that this is not done more routinely is that it is relatively difficult, despite there being several published studies that describe ways to achieve it. In this paper we demonstrate a procedure that eliminates one of the major hurdles associated with this intubation, that of visualizing the trachea during the entire time of intubation. The approach uses a 0.5 mm fiberoptic light source that serves as an introducer to direct the intubation cannula into the mouse trachea. We show that it is possible to use this procedure to measure lung mechanics in individual mice over a time course of at least several weeks. The technique can be set up with relatively little expense and expertise, and it can be routinely accomplished with relatively little training. This should make it possible for any laboratory to routinely carry out this intubation, thereby allowing longitudinal studies in individual mice, thereby minimizing the number of mice needed and increasing the statistical power by using each mouse as its own control.

As an evaluation of the method, we used four 20 week old male BALB/c mice with average weight (&plusmn; SEM) of 27.7 &plusmn; 0.40 g. They were studied on five consecutive weeks, where the lung resistance was measured using a system as previously described 11. Each mouse was anesthetized with ketamine (100 &mu;g/g BW) and xylazine (15 &mu;g /g BW) in saline via IP injection. They were then intubated as described above. If there is any doubt whether the cannula is in the trachea and not the esophagus, this can be validated using a small dental mirror. Keep the mirror in a freezer, and when needed place in front of the Luer hub of the catheter. If the catheter is in the trachea, the exhaled breath will form a visible condensate on the mirror.
After intubation, we then connected the mice to the ventilator and measured Lung Resistance The mice were ventilated with a rate of 2 Hz and tidal volume of 0.2 ml, and respiratory resistance was measured by the inspiratory occlusion method as previously described11. Figure 5 shows 5 weekly measurements in each of the 4 mice. Reproducibility is excellent, showing that, at least at weekly intervals, there is no effect of the prior measurement. This is consistent with previously reported weekly assessments of mechanics and BAL cell profiles in individual BALB/c mice with a more difficult and potentially traumatic procedure 2.
<img alt="Figure 1" src="/files/ftp_upload/50318/50318fig1.jpg" /> 
 Figure 1. Picture showing items used for intubation. The fiberoptic cable is shown inserted into a silicone rubber stopper, with a small piece of silicone rubber tubing tied near the opposite end. A silicone rubber stopper is attached to the light source as shown in Figure 2.
<img alt="Figure 2" src="/files/ftp_upload/50318/50318fig2.jpg" /> 
 Figure 2. Picture showing rubber stopper connected to light source with other end of fiberoptic cable inserted through the intubation cannula. A simple support stand to hold the mouse during intubation is also shown on the left.
<img alt="Figure 3" src="/files/ftp_upload/50318/50318fig3.jpg" /> 
 Figure 3. Two perspectives showing the position of mouse readied for intubation.
<img alt="Figure 4" src="/files/ftp_upload/50318/50318fig4.jpg" /> 
 Figure 4. This figure shows an intubated mouse ready for ventilation. The tape around the mouth helps keep the cannula from moving. A small piece of Plasticine (modeling clay) provides a convenient rest to secure the cannula hub for connection to the ventilator.
<img alt="Figure 5" fo:content-width="3.5in" fo:src="/files/ftp_upload/50318/50318fig5highres.jpg" src="/files/ftp_upload/50318/50318fig5.jpg" /> 
 Figure 5. Lung resistance from each of 4 mice (in different colors) measured at 5 weekly intervals.
<img alt="Figure 6" src="/files/ftp_upload/50318/50318fig6.jpg" /> 
 Figure 6. Shown is a chart record of airway pressure in one intubated mouse following injections of 0.5, 0.75, and 1 ml. Each volume was held for 20-40 sec, then released before then next inflation. Although there may be a very slow leak after the stress relaxation recovery, this would have negligible effect on normal ventilation or assessment of dynamic pulmonary function measurements. <a href="https://www.jove.com/files/ftp_upload/50318/50318fig6large.jpg" target="_blank"> Click here to view larger figure</a>.

Watch this Scientific Journal Video about A Simple Method of Mouse Lung Intubation at JoVE.com

A Simple Method of Mouse Lung Intubation

This paper describes a striaghforward and efficient method of intubating mice for pulmonary function measurements or pulmonary instillation, that allows the mice to recover and be studied at later times. The procedure involves an inexpensive fiberoptic light source that directly illuminates the trachea.

A  simple procedure to intubate mice for pulmonary function measurements would  have several advantages in longitudinal studies with limited numbers or  ...

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50.5K Views.  Johns Hopkins Bloomberg School of Public Health. This paper describes a striaghforward and efficient method of intubating mice for pulmonary function measurements or pulmonary instillation, that allows the mice to recover and be studied at later times. The procedure involves an inexpensive fiberoptic light source that directly illuminates the trachea.

Video: A Simple Method of Mouse Lung Intubation

A Model of Disturbed Flow-Induced Atherosclerosis in Mouse Carotid Artery by Partial Ligation and a Simple Method of RNA Isolation from Carotid Endothelium

Despite the well-known close association, direct evidence linking disturbed flow to atherogenesis has been lacking. We have recently used a modified version of carotid partial ligation methods [1,2] to show that it acutely induces low and oscillatory flow conditions, two key characteristics of disturbed flow, in the mouse common carotid artery. Using this model, we have provided direct evidence that disturbed flow indeed leads to rapid and robust atherosclerosis development in Apolipoprotein E knockout mouse [3]. We also developed a method of endothelial RNA preparation with high purity from the mouse carotid intima [3]. Using this mouse model and method, we found that partial ligation causes endothelial dysfunction in a week, followed by robust and rapid atheroma formation in two weeks in a hyperlipidemic mouse model along with features of complex lesion formation such as intraplaque neovascularization by four weeks. This rapid in vivo model and the endothelial RNA preparation method could be used to determine molecular mechanisms underlying flow-dependent regulation of vascular biology and diseases. Also, it could be used to test various therapeutic interventions targeting endothelial dysfunction and atherosclerosis in considerably reduced study duration.

This describes a partial carotid ligation surgery, which causes disturbed flow conditions and subsequent atherosclerosis development (in two weeks) with intraplaque neo-vascularization (in four weeks) in the mouse common carotid artery. We also describe a novel method of RNA isolation from the carotid intima, providing high purity endothelial RNA.

Despite the well-known close association, direct evidence linking disturbed flow to atherogenesis has been lacking. We have recently used a modified ...

A Simple Guide Screw Method for Intracranial Xenograft Studies in Mice

The grafting of human tumor cells into the brain of immunosuppressed mice is an established method for the study of brain cancers including glioblastoma (glioma) and medulloblastoma. The widely used stereotactic approach only allows for the injection of a single animal at a time, is labor intensive and requires highly specialized equipment. The guide screw method, initially developed by Lal et al.,1 was developed to eliminate cumbersome stereotactic procedures. We now describe a modified guide screw approach that is rapid and exceptionally safe; both of which are critical ethical considerations. Notably, our procedure now incorporates an infusion pump that allows up to 10 animals to be simultaneously injected with tumor cells.
To demonstrate the utility of this procedure, we established human U87MG glioma cells as intracranial xenografts in mice, which were then treated with AMG102; a fully human antibody directed to HGF/scatter factor currently undergoing clinical evaluation2-5. Systemic injection of AMG102 significantly prolonged the survival of all mice with intracranial U87MG xenografts and resulted in a number of complete cures. 
This study demonstrates that the guide screw method is an inexpensive, highly reproducible approach for establishing intracranial xenografts. Furthermore, it provides a relevant physiological model for validating novel therapeutic strategies for the treatment of brain cancers.

In order to evaluate novel therapeutic paradigms for the treatment of glioma, physiological relevant models are essential. We utilize an implantable guide screw procedure for establishment of intracranial xenograft models that is more rapid and safer than stereotactic approaches.

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Noninvasive Intratracheal Intubation to Study the Pathology and Physiology of Mouse Lung

The use of a model that mimics the condition of lung diseases in humans is critical for studying the pathophysiology and/or etiology of a particular disease and for developing therapeutic intervention. With the increasing availability of knockout and transgenic derivatives, together with a vast amount of genetic information, mice provide one of the best models to study the molecular mechanisms underlying the pathology and physiology of lung diseases. Inhalation, intranasal instillation, intratracheal instillation, and intratracheal intubation are the most widely used techniques by a number of investigators to administer materials of interest to mouse lungs. There are pros and cons for each technique depending on the goals of a study. Here a noninvasive intratracheal intubation method that can directly deliver exogenous materials to mouse lungs is presented. This technique was applied to administer bleomycin to mouse lungs as a model to study pulmonary fibrosis.

The use of a model that mimics the condition of lung diseases in humans is critical for studying the pathophysiology and/or etiology of a particular disease and for developing therapeutic intervention. Here a noninvasive intratracheal intubation method that can directly deliver exogenous materials to mouse lungs is presented.

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A Modified Method for Heterotopic Mouse Heart Transplantion

Mice are often used as heart transplant donors and recipients in studies of transplant immunology due to the wide range of transgenic mice and reagents available. A difficulty is presented due to the small size of the animal and the considerable technical challenges of the microsurgery involved in heart transplantation. In particular, a high rate of technical failure early after transplantation may result from recipient death and post-operative complications such as hind limb paralysis or a non-beating heart. Here, the complete technique for heterotopic mouse heart transplantation is demonstrated, involving harvesting the donor heart and its subsequent implantation into a recipient mouse. The donor heart is harvested immediately following in situ perfusion with cold heparinized saline and transection of the ascending aorta and pulmonary artery. The recipient operation involves preparation of the abdominal aorta and inferior vena cava (IVC), followed by end-to-side anastomosis of the donor aorta with the recipient aorta using a single running 10-0 microsuture and a similar anastomosis of the donor pulmonary artery with the recipient IVC. Following the operation the animal is injected with 0.6 ml normal saline subcutaneously and allowed to recover on a 37&#160;&#176;C heating pad. The results from 227 mouse heart transplants are summarized with a success rate at 48 hr of 86.8%. Of the 13.2% failures within 48 hr, 5 (2.2%) experienced hind limb paralysis, 10 (4.4%) had a non-beating heart due to graft ischemic injury and/or thrombosis, while 15 (6.6%) died within 48 hr.

A technique is demonstrated for the microsurgical procedure for heterotopic transplantation of hearts in mice, including simplified methods for donor harvesting and recipient vessel anastomosis.

Mice are often used as heart transplant donors and recipients in studies of transplant immunology due to the wide range of transgenic mice and reagents ...

Implantation of Fibrin Gel on Mouse Lung to Study Lung-specific Angiogenesis

Recent significant advances in stem cell research and bioengineering techniques have made great progress in utilizing biomaterials to regenerate and repair damage in simple tissues in the orthopedic and periodontal fields. However, attempts to regenerate the structures and functions of more complex three-dimensional (3D) organs such as lungs have not been very successful because the biological processes of organ regeneration have not been well explored. It is becoming clear that angiogenesis, the formation of new blood vessels, plays key roles in organ regeneration. Newly formed vasculatures not only deliver oxygen, nutrients and various cell components that are required for organ regeneration but also provide instructive signals to the regenerating local tissues. Therefore, to successfully regenerate lungs in an adult, it is necessary to recapitulate the lung-specific microenvironments in which angiogenesis drives regeneration of local lung tissues. Although conventional in vivo angiogenesis assays, such as subcutaneous implantation of extracellular matrix (ECM)-rich hydrogels (e.g., fibrin or collagen gels or Matrigel - ECM protein mixture secreted by Engelbreth-Holm-Swarm mouse sarcoma cells), are extensively utilized to explore the general mechanisms of angiogenesis, lung-specific angiogenesis has not been well characterized because methods for orthotopic implantation of biomaterials in the lung have not been well established. The goal of this protocol is to introduce a unique method to implant fibrin gel on the lung surface of living adult mouse, allowing for the successful recapitulation of host lung-derived angiogenesis inside the gel. This approach enables researchers to explore the mechanisms by which the lung-specific microenvironment controls angiogenesis and alveolar regeneration in both normal and pathological conditions. Since implanted biomaterials release and supply physical and chemical signals to adjacent lung tissues, implantation of these biomaterials on diseased lung can potentially normalize the adjacent diseased tissues, enabling researchers to develop new therapeutic approaches for various types of lung diseases.

Recapitulation of the organ-specific microenvironment, which stimulates local angiogenesis, is indispensable for successful regeneration of damaged tissues. This report demonstrates a novel method to implant fibrin gels on the lung surface of living mouse in order to explore how the lung-specific microenvironment modulates angiogenesis and alveolar regeneration in adult mouse.

Recent significant advances in stem cell research and bioengineering techniques have made great progress in utilizing biomaterials to regenerate and ...

Intubation-mediated Intratracheal (IMIT) Instillation: A Noninvasive, Lung-specific Delivery System

Respiratory disease studies typically involve the use of murine models as surrogate systems. However, there are significant physiologic differences between the murine and human respiratory systems, especially in their upper respiratory tracts (URT). In some models, these differences in the murine nasal cavity can have a significant impact on disease progression and presentation in the lower respiratory tract (LRT) when using intranasal instillation techniques, potentially limiting the usefulness of the mouse model to study these diseases. For these reasons, it would be advantageous to develop a technique to instill bacteria directly into the mouse lungs in order to study LRT disease in the absence of involvement of the URT. We have termed this lung specific delivery technique intubation-mediated intratracheal (IMIT) instillation. This noninvasive technique minimizes the potential for instillation into the bloodstream, which can occur during more invasive traditional surgical intratracheal infection approaches, and limits the possibility of incidental digestive tract delivery. IMIT is a two-step process in which mice are first intubated, with an intermediate step to ensure correct catheter placement into the trachea, followed by insertion of a blunt needle into the catheter to mediate direct delivery of bacteria into the lung. This approach facilitates a &#62;98% efficacy of delivery into the lungs with excellent distribution of reagent throughout the lung. Thus, IMIT represents a novel approach to study LRT disease and therapeutic delivery directly into the lung, improving upon the ability to use mice as surrogates to study human respiratory disease. Furthermore, the accuracy and reproducibility of this delivery system also makes it amenable to Good Laboratory Practice Standards (GLPS), as well as delivery of a wide range of reagents which require high efficiency delivery to the lung.

Intubation-mediated Intratracheal IMIT Instillation: A Noninvasive, Lung-specific Delivery System

Intubation-mediated intratracheal (IMIT) instillation of reagents is an excellent, noninvasive method for studying respiratory disease, as well as a method for instilling therapeutic reagents directly into the lung. It is a rapid and highly reproducible method which is suitable for preclinical testing.

Respiratory disease studies typically involve the use of murine models as surrogate systems. However, there are significant physiologic differences ...

Mouse Pneumonectomy Model of Compensatory Lung Growth

In humans, disrupted repair and remodeling of injured lung contributes to a host of acute and chronic lung disorders which may ultimately lead to disability or death. Injury-based animal models of lung repair and regeneration are limited by injury-specific responses making it difficult to differentiate changes related to the injury response and injury resolution from changes related to lung repair and lung regeneration. However, use of animal models to identify these repair and regeneration signaling pathways is critical to the development of new therapies aimed at improving pulmonary function following lung injury. The mouse pneumonectomy model utilizes compensatory lung growth to isolate those repair and regeneration signals in order to more clearly define mechanisms of alveolar re-septation. Here, we describe our technique for performing mouse pneumonectomy and sham pneumonectomy. This technique may be utilized in conjunction with lineage tracing or other transgenic mouse models to define molecular and cellular mechanism of lung repair and regeneration.

Mouse pneumonectomy is a commonly employed model of compensatory lung growth. This procedure can be used in conjunction with lineage tracing or transgenic mouse models to elucidate underlying mechanisms.

In humans, disrupted repair and remodeling of injured lung contributes to a host of acute and chronic lung disorders which may ultimately lead to ...

Method of Isolated Ex Vivo Lung Perfusion in a Rat Model: Lessons Learned from Developing a Rat EVLP Program

The number of acceptable donor lungs available for lung transplantation is severely limited due to poor quality. Ex-Vivo Lung Perfusion (EVLP) has allowed lung transplantation in humans to become more readily available by enabling the ability to assess organs and expand the donor pool. As this technology expands and improves, the ability to potentially evaluate and improve the quality of substandard lungs prior to transplant is a critical need. In order to more rigorously evaluate these approaches, a reproducible animal model needs to be established that would allow for testing of improved techniques and management of the donated lungs as well as to the lung-transplant recipient. In addition, an EVLP animal model of associated pathologies, e.g., ventilation induced lung injury (VILI), would provide a novel method to evaluate treatments for these pathologies. Here, we describe the development of a rat EVLP lung program and refinements to this method that allow for a reproducible model for future expansion. We also describe the application of this EVLP system to model VILI in rat lungs. The goal is to provide the research community with key information and &#8220;pearls of wisdom&#8221;/techniques that arose from trial and error and are critical to establishing an EVLP system that is robust and reproducible.

Method of Isolated Ex Vivo Lung Perfusion in a Rat Model: Lessons Learned from Developing a Rat EVLP Program

Ex-Vivo Lung Perfusion (EVLP) has allowed lung transplantation in humans to become more readily available by enabling the ability to assess organs and expand the donor pool. Here, we describe the development of a rat EVLP program and refinements that allow for a reproducible model for future expansion.

The number of acceptable donor lungs available for lung transplantation is severely limited due to poor quality. Ex-Vivo Lung Perfusion (EVLP) has allowed ...

A Simple Device to Rapidly Prepare Whole Mounts of the Mouse Intestine

Preparing whole mounts of the mouse small intestine and colon for subsequent analysis or quantification can be time consuming and difficult. We describe the use of a simple device to cut and &#8216;roll&#8217; mouse intestines to rapidly prepare whole mount preparations of superior and uniform quality to that which can be achieved by hand. The device comprises a base that holds 4 stainless steel rods and a top, which acts a cutting guide. The rods are inserted into the lumen of the small intestine [divided into thirds] and the colon. The rods and samples are then placed over a piece of filter paper or card into the holding slots in the base of the device. The top of the device is then positioned and serves as a cutting guide. The two angled sections in the center of the top piece are used to guide a knife or scalpel and cut the intestines longitudinally on the top of the rods. Once the intestinal sections have been cut, the top is removed and the card,&#160;tissue and rods gently removed from the device and placed on the bench. The rods are then gently rolled sideways to flatten and stick the intestinal segments onto the underlying piece of filter paper or card. The final preparation can then be examined or fixed and stored for later analysis. The preparations are invaluable for the study of intestinal changes in normal or genetically modified mouse models. The preparations have been used for the study and quantification of the effects of inflammation (colitis), damage, pre-cancerous lesions (aberrant crypt foci (ACFs) and mucin depleted foci (MDFs)) and polyps or tumors.

The use of a simple device to cut and &#8216;roll&#8217; mouse intestines to rapidly prepare whole mount preparations is described.

Preparing whole mounts of the mouse small intestine and colon for subsequent analysis or quantification can be time consuming and difficult. We describe ...

Evaluation of Lung Metastasis in Mouse Mammary Tumor Models by Quantitative Real-time PCR

Metastatic disease is the spread of malignant tumor cells from the primary cancer site to a distant organ and is the primary cause of cancer associated death 1. Common sites of metastatic spread include lung, lymph node, brain, and bone 2. Mechanisms that drive metastasis are intense areas of cancer research. Consequently, effective assays to measure metastatic burden in distant sites of metastasis are instrumental for cancer research. Evaluation of lung metastases in mammary tumor models is generally performed by gross qualitative observation of lung tissue following dissection. Quantitative methods of evaluating metastasis are currently limited to ex vivo and in vivo imaging based techniques that require user defined parameters. Many of these techniques are at the whole organism level rather than the cellular level 3-6. Although newer imaging methods utilizing multi-photon microscopy are able to evaluate metastasis at the cellular level 7, these highly elegant procedures are more suited to evaluating mechanisms of dissemination rather than quantitative assessment of metastatic burden. Here, a simple in vitro method to quantitatively assess metastasis is presented. Using quantitative Real-time PCR (QRT-PCR), tumor cell specific mRNA can be detected within the mouse lung tissue.

This protocol describes how to use quantitative Real-time PCR (QRT-PCR) to detect tumor cell specific mRNA representing metastasis within the mouse lung tissue.

Metastatic disease is the spread of malignant tumor cells from the primary cancer site to a distant organ and is the primary cause of cancer associated ...