This work was supported in part by the Intramural Research Program of the National Institutes of Health, National Institute of Environmental Health Sciences (Z01 ES102005).

All experiments were performed in accordance with the Animal Welfare Act and the U.S. Public Health Service Policy on Humane Care and Use of Laboratory Animals after review by the Animal Care and Use Committee of the NIEHS.
1. Preparation of K. pneumoniae Culture
Caution: Perform all steps in a biosafety level 2 (BSL2) hood or other BSL2 designated area and discard waste per institute BSL2 guidelines.
<ol>
	<li>For suspension growth of K. pneumoniae, thaw a glycerol stock of bacteria and inoculate a working culture by transferring 1 ml of K. pneumoniae stock to 50 ml of tryptic soy broth (TSB) in a 500 ml or larger flask.</li>
	<li>Make glycerol stocks by adding 900 &#181;l of cultured log-phase K. pneumoniae to 100 &#181;l of sterile glycerol and freezing at -20 &#176;C or -80 &#176;C. For long term storage, -80 &#176;C is recommended.</li>
	<li>Culture bacteria from step 1.1 by shaking flask overnight (14 - 18 hr) at 37 &#176;C at 200-225 rpm.In order to promote log phase growth, dilute 1-5 ml of this overnight culture into a flask containing 50 ml TSB in the morning and place back at 37 &#176;C with shaking for ~2.5 hr.</li>
	<li>Transfer 1 ml of K. pneumoniae culture from the last step into a 1.5 ml tube and spin at 7,500 x g for 2 min. A white pellet should be visible. Remove supernatant without disturbing the pellet, gently resuspend bacteria with pipette in 1 ml of sterile saline, and spin again at 7,500 x g for 2 min.
	<ol>
		<li>Perform this wash step 2x, bringing up the final, washed pellet in 1 ml of sterile saline.</li>
	</ol>
	</li>
	<li>Perform log-wise serial dilutions of this neat inoculum. For example, add 400 &#181;l of the inoculum into 3.6 ml of sterile saline serially to make a 10-1-10-9 dilution series.</li>
	<li>Determine the OD600 of washed K. pneumoniae by placing 1 ml of the 1:10 and 1:100 dilutions into disposable cuvettes and measuring the OD600, blanking first with sterile saline. Measure duplicates to ensure accuracy. An OD600 of 1.0 is roughly equivalent to 4-7 x 108 CFU/ml. Note that the relationship between OD and CFU/ml may not be linear.
	<ol>
		<li>Use the OD600 from the dilutions to calculate concentration of K. pneumoniae, applying the OD:CFU/ml conversion above, and factoring in the dilution.</li>
	</ol>
	</li>
	<li>Use the K. pneumoniae concentration to prepare the desired inoculum CFU dose in a &#60;100 &#181;l volume for delivery to mice (see below).</li>
	<li>Also plate 100 &#181;l of 10-6-10-9 dilutions and 100 &#181;l of sterile saline onto individual tryptic soy agar (TSA) plates at RT. Incubate at 37 &#176;C overnight and enumerate bacterial colonies in order to calculate exact concentration that was used in experiment and to control for contamination. 
	NOTE: Actual bacterial concentration of final inoculum may be &#177; 500 CFU/ml that predicted by absorbance. Experimenters should ideally confirm the OD600:CFU/ml relationship in pilot studies prior to use in vivo in mice, empirically readjusting the OD600:CFU/ml conversion based on their personal experience.</li>
</ol>
2. Murine Intratracheal (i.t.) Aspiration of K. pneumoniae from Oropharynx
<ol>
	<li>Ensure that mice are uniquely identified by tail mark, tattoo, or other approved identifier.</li>
	<li>Draw up dosing inoculum of bacteria using P200 pipette prior to removing mice from isoflurane in order to expedite procedure. For best results with i.t. aspiration, use a volume of &#60;100 &#181;l to prevent overflow. Herein, use 2000 CFU/50 &#181;l of K. pneumoniae.</li>
	<li>Anesthetize mice in a clear chamber with isoflurane gas (e.g., 2% isoflurane at flow rate of 1 L/min) or as per institutional guidelines. The number of mice to anesthetize together is determined by comfort level of the experimenter, typically 1-2&#160;at a time. Observe breathing and confirm level of anesthesia once deep breaths are visible and 2-3 sec can be counted between breaths. 
	NOTE: If there is concern about possible confounding effects of volatile (inhaled) anesthetics, anesthesia can be achieved instead with intraperitoneal injection of ketamine/xylazine. With the method described above, deep anesthesia only lasts a few min. If more prolonged anesthesia is induced, eye ointment should be used to prevent ocular desiccation.</li>
	<li>Once mouse is anesthetized, position it in a semirecumbent supine position, suspended by the maxillary incisors from a rubber band stretched between pegs on a slanted Plexiglas board (Figure 1).</li>
	<li>Gently pull the mouse tongue to the side with a pair of blunt non-ridged forceps and deposit dose into the oral cavity with a P200 pipette. Exercise great care to avoid inducing trauma to either the tongue or the oropharynx.</li>
	<li>While keeping the tongue retracted, gently occlude the nose with a gloved finger until the mouse inhales. Continue to cover the nose until the mouse has taken two or more inhalations, and no liquid is visible in the oral cavity. Covering the nose helps ensure that the mouse will inhale the bacteria into the lungs, as mice are obligate nose breathers.</li>
	<li>Remove the mouse from the inoculation board and return to its cage, placing the mouse on its back to prevent bedding or debris from blocking the nares while the mouse is recovering from anesthesia.</li>
	<li>Once all mice have been dosed and have awoken from anesthesia, house mice in a cubicle/room containing only mice that have been dosed with K. pneumoniae. Monitor mice daily, including body weights if going beyond 48 hr post-infection.</li>
	<li>Use daily mash and/or supplemental heat if allowing the mice to go beyond 48 hr.</li>
</ol>
3. Bronchoalveolar Lavage Fluid (BALF) Collection and Analysis
<ol>
	<li>Euthanize mice per institutional guidelines. Here, use a lethal injection of 150 mg/kg sodium&#160;pentobarbital &#160;or equivalent dose of commercial euthanasia solution&#160;followed by exsanguination, as this avoids possible complications to the lungs associated with CO2 inhalation and cervical dislocation.</li>
	<li>Position mouse on back and sterilize mouse by spraying fur with 70% ethanol particularly over the chest and neck area.</li>
	<li>Make a longitudinal cut just below the sternum, then holding the sternum with forceps, nick the diaphragm, allowing the lung to fall back into the chest cavity. Cut up through the chest cavity on either side of the vasculature and then up through the neck, exposing the trachea.</li>
	<li>As the trachea is surrounded by longitudinal muscles on either side, carefully cut through the middle and push tissue to the sides, as this avoids cutting the surrounding vasculature, which could introduce blood into the trachea. If the vasculature is nicked, clean the surrounding area with gauze prior to accessing the tracheal lumen.</li>
	<li>Use surgical scissors or a needle to nick the trachea about &#188; of the way down from the head and insert a cannula attached to a 1 ml syringe preloaded with 1 ml of 1x phosphate-buffered saline (PBS) caudally into the trachea. If using mice &#60;8 weeks of age or females, reduce the lavaging volume to 600-800 &#181;l.</li>
	<li>Push the volume into the lung slowly, allowing all lobes to inflate and then pull the volume back out with the syringe, repeating 3x. If PBS is coming out of the nostrils the cannula has not been pushed far enough into the trachea or inflation is occurring too quickly. Alternatively, if the lungs are not inflating well, the cannula has been pushed in too far, so withdraw slightly.</li>
	<li>Collect pooled washes in a 15 ml tube on ice.</li>
	<li>Spin the airspace lavage fluid at 1,200 x g for 5 min, collect supernatant into a 1.5 ml tube and freeze at -80 &#176;C. Subsequently, use the supernatant (i.e., BALF) for measurement of protein, cytokines, and for various other biochemical assays. The pellet represents cells from the bronchoalveolar space.</li>
	<li>If red blood cells (RBCs) are evident in the cell pellet based on color, lyse with 1 ml of ACK lysis buffer for 1 min followed by addition of 5 ml of 1x PBS to stop the lysis reaction and dilute the buffer. Handle all specimens identically, either treated or not with ACK lysis buffer.</li>
	<li>Spin cells at 1,200 x g for 5 min, decant the supernatant, and bring the cells up in 1 ml of 1x PBS.</li>
	<li>Vortex and count cells on a hemocytometer for enumeration of total airspace cells.</li>
	<li>Based on density of cells add approximately 80-150 &#181;l (~80 &#181;l for an infected mouse and ~150 for a naive mouse) to a slide chamber on a cytospin centrifuge. Spin cells onto microscope slides for subsequent staining to determine cell differential populations.</li>
	<li>Allow cells to dry on slides overnight. Stain cells with tri-stain (e.g., Hema 3 [see table]) by dipping the slides 7 times in fixative, 9 times in stain 1, and 7 times in stain 2 (no rinses between stains), and allow to dry. After stain has dried, coverslip slides and manually count differentials by microscope. Generally, neutrophils and monocyte/macrophages are easily distinguished by their size and nuclear morphology.</li>
</ol>
4. Determining Bacterial Load in Lung and Peripheral Tissues
<ol>
	<li>Before beginning: prepare dilution tubes with 900 &#181;l of 1x PBS for lung and spleen and 90 &#181;l of 1x PBS for blood. Label plates for culturing bacteria and set at room temperature; therefore, once tissue has been collected time will be minimized between homogenization and plating for bacterial growth. 
	Note: Due to the heterogeneous deposition of bacteria in the lungs that can occur with aspiration, it is recommended that individual mice be used for analysis of either total airway cellularity or total (bilateral) lung bacterial burden.</li>
	<li>Euthanize the mouse as per step 3.1 and subsequently collect blood from the left ventricle, placing into a heparinized tube to avoid clotting. Remove all lung lobes and place in a 15 ml tube containing 5 ml of 1x PBS, followed by removal of the spleen and placement into 2 ml of 1x PBS. Place tubes on ice. Take care to clean instruments with ethanol between each tissue/mouse.</li>
	<li>Homogenize the tissue on ice, preferably with disposable homogenizers that can be changed between each sample. Disposable homogenizer tips can be autoclaved for reuse between experiments.</li>
	<li>Once tissue has been homogenized, perform serial dilutions and plate. Plate samples of a given tissue/dilution in duplicate on a single TSA plate by drawing a dividing line on the plastic. Plating and dilution suggestions are shown below (in all cases, 10 &#181;l of sample is spread onto plate) and are based on a 24 hr post-infection necropsy. If analyzing 48-72 hr samples, a larger number of dilutions may need to be plated due to increased bacterial burden at later time points.
	<ol>
		<li>For Blood - dilute 1:10 by adding 10 &#181;l of blood into 90 &#181;l of sterile 1x PBS. Plate neat and 1:10 diluted samples in duplicate. Once blood has been plated, spin residual blood at 9,300 x g for 10 min to extract plasma for measurement of cytokines and chemokines.</li>
		<li>For Lung - dilute 1:10 - 1:100 by adding 100 &#181;l of homogenized lung into 900 &#181;l of sterile 1x PBS serially. Plate neat homogenate and all dilutions in duplicate.</li>
		<li>For Spleen - dilute 1:10-1:100 by adding 100 &#181;l of homogenized spleen into 900 &#181;l of sterile 1x PBS serially. Plate neat homogenate and both dilutions in duplicate.</li>
	</ol>
	</li>
	<li>Allow spread samples to dry briefly.</li>
	<li>Invert TSA plates and place in a static 37 &#176;C incubator overnight.</li>
	<li>Determine the number of colonizing bacteria by averaging the bacterial colonies from both sides of the plate and from each dilution. Factor in the initial dilutions of 5 ml for lung and 2 ml for spleen to determine total tissue CFUs.</li>
</ol>

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The authors declare that they have no competing financial interests.

Murine models of bacterial pneumonia, partnered with gene targeting and in vivo biological and pharmacological interventions, have provided critical insights into the pulmonary host defense response. Great advances have been made in particular in our understanding of the chemokines and adhesion molecules that govern recruitment of neutrophils to the infected airspace10,11. In vivo models of pneumonia, unlike cell-based or alternative approaches, have also provided key insights into endocrine ...

Community-acquired pneumonia remains the leading cause of death from infection in the U.S., with little overall change in mortality rates over the past 40 years despite improvements in vaccination and antibiotic strategies1,2. Despite the lack of perceptible progress at the public health level, in recent years dramatic advances have been made in our understanding of the molecular and cellular pathogenesis of pneumonia, with many of these steps forward made possible by the use of mouse models of lung infection. The genetic tractability of the mouse, the similarity of the murine and human immune systems, and the vast array of murine-targeted immunologic reagents that have become commercially available have together facilitated rapid progress of the field.
Mouse models of bacterial pneumonia described in the literature have generally relied upon one of four technical routes for pathogen inoculation: i) aerosolization; ii) intranasal delivery; iii) peroral delivery; and iv) surgical intratracheal injection (i.e., tracheotomy)3. All routes of infection have advantages and disadvantages3. In particular, relative exposure of the upper airway, potential for admixture of oronasal flora, requirements for general anesthesia, variability of the inoculum delivered to the distal lung, lobar distribution of the delivered pathogens, technical expertise requirements, and procedural morbidity vary widely across these approaches.
Commonly used peroral infection techniques include endotracheal (translaryngeal) cannulation via either a &#39;blind&#39; (non-visualized) approach, or under direct laryngeal visualization3-5. Both methods, while robust, require substantial training and also carry risk of trauma to the upper airway. In the present report, we describe a technically non-intensive, non-invasive, inexpensive, and rapid method of peroral infection, whereby bacteria (Klebsiella pneumoniae in the example provided) pipetted into the oropharynx of an anesthetized mouse are delivered to the lungs via aspiration (i.e., inhalation). We and others have used the aspiration pneumonia technique successfully6-9. This versatile and easily learned lung-delivery method can be extended to the delivery of many additional non-caustic agents to the lungs, including cytokines and other proteins, pathogen-associated molecules (e.g., lipopolysaccharide), cells (i.e., adoptive transfer), and toxins (e.g., bleomycin). In addition to discussing important technical considerations, we also describe an integrated, quantitative approach for assessing the subsequent host response to pneumonia, including downstream measurement of bacterial clearance (i.e., colony forming unit [CFU] quantitation in the lung and peripheral organs) and leukocyte accumulation in the airspace.

<table border="0" cellpadding="0" cellspacing="0" width="698">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Klebsiella pneumoniae, serotype 2</td>
			<td style="width:169px;">ATCC</td>
			<td style="width:119px;">43816</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Tryptic soy broth</td>
			<td style="width:169px;">Becton Dickenson</td>
			<td>211825</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Excel Safelet IV Catheters, 18G x 1 1/4&#34;</td>
			<td>Claflin Medical Equipment</td>
			<td style="width:119px;">MEDC-031122</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Hema 3 Solution 1</td>
			<td style="width:169px;">Fisher</td>
			<td style="width:119px;">23-122-937</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Hema 3 Solution 2</td>
			<td style="width:169px;">Fisher</td>
			<td style="width:119px;">23-122-952</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Hema 3 Fixative</td>
			<td style="width:169px;">Fisher</td>
			<td style="width:119px;">23-122-929</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">27&#189; gauge tuberculin syringes</td>
			<td style="width:169px;">Fisher</td>
			<td style="width:119px;">14-826-87</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">Lithium heparin plasma collectors</td>
			<td style="width:169px;">Fisher</td>
			<td style="width:119px;">2675187</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:243px;">L-shaped disposable spreaders</td>
			<td style="width:169px;">Lab Scientific</td>
			<td style="width:119px;">DSC</td>
			<td></td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;width:243px;">1x PBS, pH 7.4</td>
			<td style="width:169px;">prepared in-house</td>
			<td style="width:119px;">n/a</td>
			<td>Distilled water (5 L), NaCl (40 g), KCl (1 g), Na2HPO4 (5.75 g), KH2PO4 (1 g)&#160;&#160;&#160;</td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;width:243px;">ACK lysis buffer</td>
			<td style="width:169px;">prepared in-house</td>
			<td style="width:119px;">n/a</td>
			<td>NH4Cl (4.145 g), KHCO3 (0.5 g), EDTA (18.6 mg), bring up to 500 ml with distilled water and pH to 7.4</td>
		</tr>
	</tbody>
</table>

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.

C57BL/6 mice were infected with 2000 CFU of K. pneumoniae 43816 (serotype 2) via oropharyngeal aspiration into the lungs. At this dose, mice typically begin to show clinical symptoms 12-24 hr post-infection including lethargy, ruffled fur, and weight loss of 5-10% (Figure 2A). Within 48-72 hr post-infection, many of the mice show symptoms of illness and morbidity that is typically preceded by an average of 20% weight loss and results in hunched postures with decr...

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

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 ...

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Immunology and Infection

8.9K Views.  National Institutes of Health. 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.

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

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 ...

Preparation of Mouse Pituitary Immunogen for the Induction of Experimental Autoimmune Hypophysitis

Autoimmune hypophysitis is a chronic inflammation of the pituitary gland caused or accompanied by autoimmunity1. It has traditionally been considered a rare disease but reporting has increased markedly in recent years. Hypophysitis, in fact, develops not uncommonly as a "side effect" in cancer patients treated with antibodies that block inhibitory receptors expressed on T lymphocytes, such as CTLA-42 and PD-1 receptors. Autoimmune hypophysitis can be induced experimentally by injecting mice with pituitary proteins mixed with an adjuvant3. In this video article we demonstrate how to extract proteins from mouse pituitary glands and how to prepare them in a form suitable for inducing autoimmune hypophysitis in SJL mice.

Autoimmune hypophysitis can be reproduced in mice by injecting an extract of mouse pituitary proteins.

Autoimmune hypophysitis is a chronic inflammation of the pituitary gland caused or accompanied by autoimmunity1.  It has traditionally been considered a ...

Induction of Experimental Autoimmune Hypophysitis in SJL Mice

Autoimmune hypophysitis can be reproduced experimentally by the injection of pituitary proteins mixed with an adjuvant into susceptible mice1. Mouse models allow us to study how diseases unfold, often providing a good replica of the same processes occurring in humans. For some autoimmune diseases, like type 1A diabetes, there are models (the NOD mouse) that spontaneously develop a disease similar to the human counterpart. For many other autoimmune diseases, however, the model needs to be induced experimentally. A common approach in this regard is to inject the mouse with a dominant antigen derived from the organ being studied. For example, investigators interested in autoimmune thyroiditis inject mice with thyroglobulin2, and those interested in myasthenia gravis inject them with the acetylcholine receptor3. If the autoantigen for a particular autoimmune disease is not known, investigators inject a crude protein extract from the organ targeted by the autoimmune reaction. For autoimmune hypophysitis, the pathogenic autoantigen(s) remain to be identified4, and thus a crude pituitary protein preparation is used. In this video article we demonstrate how to induce experimental autoimmune hypophysitis in SJL mice.

This video shows how to induce autoimmune hypophysitis in SJL mice and how to assess its severity by histopathology.

Autoimmune hypophysitis can be reproduced experimentally by the injection of pituitary proteins mixed with an adjuvant into susceptible mice1. Mouse ...

Using Luciferase to Image Bacterial Infections in Mice

Imaging is a valuable technique that can be used to monitor biological processes. In particular, the presence of cancer cells, stem cells, specific immune cell types, viral pathogens, parasites and bacteria can be followed in real-time within living animals 1-2. Application of bioluminescence imaging to the study of pathogens has advantages as compared to conventional strategies for analysis of infections in animal models3-4. Infections can be visualized within individual animals over time, without requiring euthanasia to determine the location and quantity of the pathogen. Optical imaging allows comprehensive examination of all tissues and organs, rather than sampling of sites previously known to be infected. In addition, the accuracy of inoculation into specific tissues can be directly determined prior to carrying forward animals that were unsuccessfully inoculated throughout the entire experiment. Variability between animals can be controlled for, since imaging allows each animal to be followed individually. Imaging has the potential to greatly reduce animal numbers needed because of the ability to obtain data from numerous time points without having to sample tissues to determine pathogen load3-4.
This protocol describes methods to visualize infections in live animals using bioluminescence imaging for recombinant strains of bacteria expressing luciferase. The click beetle (CBRLuc) and firefly luciferases (FFluc) utilize luciferin as a substrate5-6. The light produced by both CBRluc and FFluc has a broad wavelength from 500 nm to 700 nm, making these luciferases excellent reporters for the optical imaging in living animal models7-9. This is primarily because wavelengths of light greater than 600 nm are required to avoid absorption by hemoglobin and, thus, travel through mammalian tissue efficiently. Luciferase is genetically introduced into the bacteria to produce light signal10. Mice are pulmonary inoculated with bioluminescent bacteria intratracheally to allow monitoring of infections in real time. After luciferin injection, images are acquired using the IVIS Imaging System. During imaging, mice are anesthetized with isoflurane using an XGI-8 Gas Anethesia System. Images can be analyzed to localize and quantify the signal source, which represents the bacterial infection site(s) and number, respectively. After imaging, CFU determination is carried out on homogenized tissue to confirm the presence of bacteria. Several doses of bacteria are used to correlate bacterial numbers with luminescence. Imaging can be applied to study of pathogenesis and evaluation of the efficacy of antibacterial compounds and vaccines.

Methods for bioluminescence imaging of bacterial infections in living animals are decribed. Pathogens are modified to express luciferase allowing optical whole body imaging of infections in live animals. Animal models can be infected with luciferase expressing pathogens and the resulting course of disease visualized in real-time by bioluminescence imaging.

Imaging is a valuable technique that can be used to monitor biological processes.  In particular, the presence of cancer cells, stem cells, specific ...

A Method for Labeling Vasculature in Embryonic Mice

The establishment of a functional blood vessel network is an essential part of organogenesis, and is required for optimal organ function. For example, in the thymus proper vasculature formation and patterning is essential for thymocyte entry into the organ and mature T-cell exit to the periphery. The spatial arrangement of blood vessels in the thymus is dependent upon signals from the local microenvironment, namely thymic epithelial cells (TEC). Several recent reports suggest that disruption of these signals results in thymus blood vessel defects 1,2. Previous studies have described techniques used to label the neonatal and adult thymus vasculature 1,2. We demonstrate here a technique for labeling blood vessels in the embryonic thymus. This method combines the use of FITC-dextran or Griffonia (Bandeiraea) Simplicifolia Lectin I (GSL 1 - isolectin B4) facial vein injections and CD31 antibody staining to identify thymus vascular structures and PDGFR-&beta; to label thymic perivascular mesenchyme 3-5. The option of using cryosections or vibratome sections is also provided. This protocol can be used to identify thymus vascular defects, which is critical for defining the roles of TEC-derived molecules in thymus blood vessel formation. As the method labels the entire vasculature, it can also be used to analyze the vascular networks in multiple organs and tissues throughout the embryo including skin and heart 6-10.

This article describes a method for labeling embryonic skin and thymus blood vessels.

The establishment of a functional blood vessel network is an essential part of organogenesis, and is required for optimal organ function. For example, in ...

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 ...

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 ...

Induction of Murine Intestinal Inflammation by Adoptive Transfer of Effector CD4+CD45RBhigh T Cells into Immunodeficient Mice

There are many different animal models available for studying the pathogenesis of human inflammatory bowel diseases (IBD), each with its own advantages and disadvantages. We describe here an experimental colitis model that is initiated by adoptive transfer of syngeneic splenic CD4+CD45RBhigh T cells into T and B cell deficient recipient mice. The CD4+CD45RBhigh T cell population that largely consists of na&#239;ve effector cells is capable of inducing chronic intestinal inflammation, closely resembling key aspects of human IBD. This method can be manipulated to study aspects of disease onset and progression. Additionally it can be used to study the function of innate, adaptive, and regulatory immune cell populations, and the role of environmental exposures, i.e., the microbiota, in intestinal inflammation. In this article we illustrate the methodology for inducing colitis with a step-by-step protocol. This includes a video demonstration of key technical aspects required to successfully develop this murine model of experimental colitis for research purposes.

Induction of Murine Intestinal Inflammation by Adoptive Transfer of Effector CD4+CD45RBhigh T Cells into Immunodeficient Mice

Here, we present a protocol to induce colonic inflammation in mice by adoptive transfer of syngeneic CD4+CD45RBhigh T cells into T and B cell deficient recipients. Clinical and histopathological features mimic human inflammatory bowel diseases. This method allows the study of the initiation of colonic inflammation and progression of disease.

There are many different animal models available for studying the pathogenesis of human inflammatory bowel diseases (IBD), each with its own advantages ...

Induction of Experimental Autoimmune Encephalomyelitis in Mice and Evaluation of the Disease-dependent Distribution of Immune Cells in Various Tissues

Multiple sclerosis is presumed to be an inflammatory autoimmune disease, which is characterized by lesion formation in the central nervous system (CNS) resulting in cognitive and motor impairment. Experimental autoimmune encephalomyelitis (EAE) is a useful animal model of MS, because it is also characterized by lesion formation in the CNS, motor impairment and is also driven by autoimmune and inflammatory reactions. One of the EAE models is induced with a peptide derived from the myelin oligodendrocyte protein (MOG)35-55 in mice. The EAE mice develop a progressive disease course. This course is divided into three phases: the preclinical phase (day 0 - 9), the disease onset (day 10 - 11) and the acute phase (day 12 - 14). MS and EAE are induced by autoreactive T cells that infiltrate the CNS. These T cells secrete chemokines and cytokines which lead to the recruitment of further immune cells. Therefore, the immune cell distribution in the spinal cord during the three disease phases was investigated. To highlight the time point of the disease at which the activation/proliferation/accumulation of T cells, B cells and monocytes starts, the immune cell distribution in lymph nodes, spleen and blood was also assessed. Furthermore, the levels of several cytokines (IL-1&#946;, IL-6, IL-23, TNF&#945;, IFN&#947;) in the three disease phases were determined, to gain insight into the inflammatory processes of the disease. In conclusion, the data provide an overview of the functional profile of immune cells during EAE pathology.

This manuscript describes the methods for induction and scoring of the experimental autoimmune encephalomyelitis (EAE) model, together with the assessment of immune cell distribution and mRNA cytokine levels in lymph nodes, spleen, blood and spinal cord using flow cytometry and quantitative PCR, respectively, at various disease phases.

Multiple sclerosis is presumed to be an inflammatory autoimmune disease, which is characterized by lesion formation in the central nervous system (CNS) ...