Name: inducing acute lung injury in mice by direct intratracheal lipopolysaccharide instillation
Uploaded: 2019-07-06T15:00:00.000+00:00
Duration: 11 min 7 s
Description: Watch this Scientific Journal Video about Inducing Acute Lung Injury in Mice by Direct Intratracheal Lipopolysaccharide Instillation at JoVE.com

The authors wish to thank Jan Kleiner and Susanne Schulz for providing technical support. The authors acknowledge the excellent support of the flow cytometry core facility at the medical faculty of Bonn University. The authors received no funding from any external organization. &#160;Part of the data given in the results section and depicted in Figure 3 has already been shown in a previous publication8.

This animal protocol was approved by the local committee for animal care (LANUV, Recklinghausen, Germany; protocol no. 84-02.04.2015) and was performed in accordance with the National Institutes of Health guidelines for the use of live animals (NIH publication No. 85-23, revised 1996).
1. ALI induction
<ol>
	<li>Use adult&#160;C57BL/6 mice at ages of about 10-12&#160;weeks. House the animals in individually ventilated cages with free access to water and standard rodent chow. However, it is possible to perform this approach on younger animals and with other mice strains.</li>
	<li>Store LPS (Escherichia coli O111:B4) in aliquots in concentrations of 5 mg/mL at -20 &#176;C. For intratracheal instillation, dilute LPS in sterile phosphate-buffered saline (PBS) to a final concentration of 2,000 &#181;g/mL.</li>
	<li>Weigh the mouse. Inject ketamine [120 mg/kg mouse bodyweight (BW)] and xylazine (16 mg/kg BW) intraperitoneally (lower one-third of the abdomen, paramedian), and wait until the onset of anesthesia.</li>
	<li>Check the depth of anesthesia by inducing a tactile stimulus. In case of insufficient anesthesia, repeat the injection with ketamine (30 mg/kg BW) and xylazine (4 mg/kg BW).</li>
	<li>Place the mouse in prone position on a temperature-controlled table to maintain a body temperature of 37 &#176;C.</li>
	<li>Apply sterile ophthalmic lubricant to prevent desiccation of corneas under anesthesia.</li>
	<li>Lift the head and hook incisors on a horizontal bar positioned approximately 5 cm above the table while the forepaws remain in close contact with the table. Super-extend the neck in a 90&#176; angle relative to the table (Figure 1). Hold the tongue with forceps to straighten the throat for easier intubation conditions.</li>
	<li>Cut a 22 gauge (G) venous catheter to a length of 20 mm. Gently insert the catheter in the vertical direction along the tongue&#8217;s root. Place a cold-light source on the skin above the larynx to help visualize the vocal chords and aim for the trachea. If resistance of the larynx occurs, retract the catheter a few millimeters before advancing again.</li>
	<li>Insert the catheter approximately 10 mm into the trachea. Ensure that the insertion is not too deep as this will result in unilateral instillation of fluid into the right or left main bronchus.</li>
	<li>Inject LPS (5 &#181;g/g BW) diluted in PBS using a pipette&#160;[injected volume depends on mouse bodyweight (e.g., 20 g bodyweight&#160;&#8594; use&#160;50 &#181;L of LPS solution)]. 
	NOTE: The mouse will typically respond with coughing or gasping to proper instillation of fluid into the trachea.</li>
	<li>Connect a syringe and add a bolus of 50 &#956;L air to assure that complete liquid volume is distributed in the lungs. Slowly remove the catheter.</li>
	<li>Keep the mouse&#39;s upper body in an upright position for 30 s to avoid leakage of the fluid from the trachea.</li>
	<li>In sham-operated animals, inject 50 &#181;L of sterile PBS intratracheally instead of LPS.</li>
	<li>Inject buprenorphine hydrochloride 0.08 mg/kg BW subcutaneously into the loose skin over the neck immediately following ALI induction and every 12 h thereafter, during the first 48 h.</li>
	<li>Maintain a body temperature of 37 &#176;C until full awareness is regained by keeping the mouse on the warming pad.</li>
	<li>Transfer the mouse into an individually ventilated cage with free access to food and water. Monitor the mouse regularly. Decreasing body temperature and respiratory depression indicates proper induction of ALI.&#160;</li>
</ol>
2. Blood sampling, bronchoalveolar lavage, organ harvesting
NOTE: Timing of euthanization depends on the scientific issue addressed. Usually, it is performed 12-72 h following LPS instillation3,4,9,10. Severity of ALI can be determined clinically by regular observation of body temperature and respiratory distress symptoms11.
<ol>
	<li>Induce anesthesia by placing the mouse in a chamber flooded with isoflurane. Use 3 vol% isoflurane with an oxygen flow of 1 L/min. Ensure deep narcosis by inducing a tactile stimulus. In case of insufficient depth of anesthesia, increase isoflurane up to 5 vol%.</li>
	<li>Sacrifice the mouse in deep anesthesia by atlanto-occipital dislocation.</li>
	<li>Fix the mouse with tape on an operation table and shortly disinfect the fur over the abdomen with 70% ethanol. Open the abdominal cavity carefully in the median line with scissors and tweezers. Remove parts of the intestine to achieve access to the vena cava inferior (IVC) right to the vertebral column and the abdominal aorta.</li>
	<li>Locate the kidney veins and insert a bent 23 G canula connected to a 1 mL syringe into the IVC directly below the confluence of the veins. Aspirate 250 &#181;L of blood and transfer into a 1.5 mL tube filled with 20 &#181;L of 0.5 M ethylenediaminetetraacetic acid (EDTA) solution. Shake gently to facilitate EDTA mixing and put the tube on ice.</li>
	<li>For bronchoalveolar lavage (BAL), prepare three 1 mL syringes with 0.5 mL of sterile PBS and 0.1 mL of air each. Shortly disinfect the fur of the throat with 70% ethanol and carefully expose the trachea with scissors and tweezers. Mobilize the trachea and wrap around a suture.</li>
	<li>Perform BAL: Puncture the trachea using micro-scissors and insert a 22 G venous catheter cut to a length of 20 mm. Fix the catheter with the suture and instillate 0.5 mL of sterile PBS and 0.1 mL of air. Aspirate the fluid after 60 s. Repeat the procedure with the additional two syringes and collect the whole aspirate in a 15 mL tube on ice.</li>
	<li>Carefully open the thorax with scissors and tweezers to harvest the lungs. Cut the diaphragm along the costal margin and cut through the ribs with two lateral incisions. Carefully avoid puncturing the lungs. Lift the sternum cranially and fix or remove it.</li>
	<li>Prepare two 10 mL syringes with 37 &#176;C warm PBS (without calcium and magnesium). Make a small incision into the left ventricle. Puncture the right ventricle with a 26 G canula and flush the pulmonary circulation with the prewarmed PBS. Be aware of the lungs turning pale during the procedure.&#160;</li>
	<li>Remove the right lobe of the lungs and cut it in two halves. Snap-freeze them in liquid nitrogen, followed by long-term storage at -80 &#176;C for further gene expression and protein analysis.</li>
	<li>Remove the whole left lung and homogenize it in a 48 well plate by mincing the tissue with scissor and tweezer. Incubate the tissue in 2 mL of digestion buffer [RPMI 1640 with 10% fetal calf serum (FCS) and 0.1% NaN3, collagenase I (1 mg/mL), and DNase II (7 mg/mL)] at 37 &#176;C for 60 min. Perform further homogenization by careful pipetting of the lung tissue pieces up and down.</li>
</ol>
3. Tissue preparation for FACS analysis
<ol>
	<li>Prepare fresh FACS buffer (Table 1): always use calcium- and magnesium-free PBS to reduce cation-dependent cell-to-cell adhesion and prevent clumping. Supplement with FCS (1%) to protect cells from apoptosis, prevent non-specific staining, and prevent cells from sticking to the FACS tubes. Include EDTA (0.5 mM) to prevent cation-based cell-to-cell adhesion when working with sticky and adherent cells like macrophages. Add sodium azide (0.1%), as it prevents bacterial contamination and photobleaching of fluorochromes and blocks antibody shedding.</li>
	<li>Transfer the blood samples (step 2.4) into 5 mL FACS tubes and gently mix the blood with 2 mL of red blood cell lysis buffer. Put the tubes on ice and terminate the reaction after 2 min by adding 2 mL of ice-cold PBS. Centrifuge the samples for 5 min at 400 x g and discard the supernatant. Resuspend the cell pellet with 60 &#181;L of FACS buffer and process for subsequent FACS staining according to previously described protocols12. 
	NOTE: Timing of euthanization of the mouse influences leukocyte count as part of the systemic inflammation. Therefore, it is recommended to adjust the cell number to 1 x 106 cells/60 &#181;L&#160;in this step to achieve the best staining results for flow cytometry analysis.</li>
	<li>Centrifuge BAL fluid (step 2.6) for 5 min at 400 x g. Aspirate the supernatant and freeze it in liquid nitrogen, followed by long-term storage at -80 &#176;C for further protein analysis. Resuspend BAL cell pellet with 2 mL of cold FACS buffer, then transfer the suspension into a 5 mL FACS tube using a 100 &#181;m mesh filter to restrain hairs.</li>
	<li>Again, centrifuge the sample for 5 min at 400 x g. Resuspend the pellet with 60 &#181;L of FACS buffer and process for subsequent FACS staining according to previously described protocols12. 
	NOTE: Timing of euthanization of the mouse influences leukocyte count in BAL as part of the inflammation. Therefore, it is recommended to adjust the cell number to 1 x 106 cells/60 &#181;L in this step to achieve the best staining results for flow cytometry analysis.</li>
	<li>Transfer the digested left lung tissue (step 2.10) into a 5 mL FACS tube using a 100 &#181;L mesh filter to extract clumps and terminate the digestion process by adding 2 mL of ice-cold FACS buffer. Centrifuge the sample for 5 min at 400 x g. Discard the supernatant and resuspend the pellet with 60 &#181;L of FACS buffer and process for subsequent FACS staining according to previously described protocols12. 
	NOTE: Timing of euthanization of the mouse influences leukocyte count in lung tissue as part of the inflammation. Therefore, it is recommended to adjust the cell number to 1 x 106 cells/60 &#181;L to achieve the best staining results for flow cytometry analysis.</li>
	<li>For FACS analysis, incubate cells with CD16/CD32 antibody at 4 &#176;C for 15 min to block non-specific binding of immunoglobulin to the Fc receptors. Add 20 &#181;L of blocking solution to 1 x 106 cells in 60 &#181;L in a 5 mL tube.</li>
	<li>Meanwhile, prepare a master mix with FACS buffer and antibodies as described in Table 2.</li>
	<li>After blocking, do not wash the cells. Add 20 &#181;L of antibody master mix per sample to obtain a final volume of 100 &#181;L. Incubate the samples for 20 min in the dark at 4 &#176;C.</li>
	<li>Wash each sample with 1 mL of FACS buffer and centrifuge for 5 min at 400 x g. Discard the supernatant and resuspend the pellet with FACS buffer to the appropriate cell concentration for FACS measurements. 
	NOTE: A cell number of 1 x 106 cells/500 &#181;L is suggested to achieve the best immune phenotyping results in FACS analysis with this protocol. However, it is recommended that antibodies have to be titrated individually.</li>
	<li>If required, add live/dead staining prior to the surface staining using specific commercially available kits8.</li>
	<li>Finally, add fixed numbers of commercially available fluorochrome-coupled calibration beads (3 x 105 beads in 20 &#181;L of FACS buffer) to each sample to determine absolute cell numbers12. The gating strategy for blood, BAL, and tissue cells is shown in Figure 2.</li>
</ol>

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The authors have nothing to disclose.

Minimal invasiveness, simple handling, and good reproducibility are key features of the presented approach to induce ALI in a small rodent model. The use of LPS instead of whole bacteria in animal models has advantages. It is a stable and pure compound and can be stored in lyophilized form until use. It is a potent stimulant for innate immune responses via the TLR4 pathway, and its biological activity may readily be quantified, facilitating the titration of disease severity with good reproducibility. Moreover, the use of...

Experimental animal models are indispensable in basic immune research. Administration of whole bacteria or microbial components has been frequently used in small animal models to induce local or systemic inflammation1. Lipopolysaccharide (LPS, or bacterial endotoxin) is a cell wall component and surface antigen of gram-negative bacteria (e.g., Enterobacteriaceae, Pseudomonas spp., or Legionella spp.). The thermostable and large molecule (molecular weight 1-4 x 106 kDa) consists of a lipid moiety (Lipid A), core region (oligosaccharide), and an O polysaccharide (or O antigen). Lipid A, with its hydrophobic fatty acid chains, anchors the molecule into a bacterial membrane and mediates (upon degradation of bacteria) the immunological activity and toxicity of LPS. Following binding to the LPS binding protein (LBP), LPS:LBP complexes ligate the CD14/TLR4/MD2 receptor complex located on the surface of many cell types, inducing a strong proinflammatory reaction with NF-&#954;B nuclear translocation and subsequent upregulation of cytokine expression2.
Acute lung injury (ALI) is defined as acute hypoxemic respiratory failure with bilateral pulmonary edema in the absence of heart failure3. Airway administration of LPS is a common way to induce pulmonary inflammation and ALI4,5,6,7. Although the sterile substance is not suitable to study pharmacologic interventions in infectious diseases, mechanistic immunological questions may be answered with adequate precision. Instillation of LPS into the trachea induces a robust inflammatory reaction with leukocyte invasion, upregulation of proinflammatory cytokines, and disruption of the alveolo-capillary barrier within hours to days, depending on the LPS dosage3,6,7.
The presented protocol describes a detailed step-by-step procedure to induce ALI in mice by intratracheal LPS instillation. The model has been validated by assessing cytokine expression, neutrophil granulocyte invasion, and intra-alveolar albumin leakage as previously described8.&#160;

<table><tbody><tr><td>1 ml syringes</td><td>BD, Franklin Lakes, NJ, USA</td><td>300013</td><td></td></tr><tr><td>10 ml syringes</td><td>BD, Franklin Lakes, NJ, USA</td><td>309110</td><td></td></tr><tr><td>Anti-CD115 (c-fms) APC</td><td>Thermo Fisher, Waltham, MA, USA</td><td>17-1152-80</td><td></td></tr><tr><td>Anti-CD11b (M1/70) - FITC</td><td>Thermo Fisher, Waltham, MA, USA</td><td>11-0112-81</td><td></td></tr><tr><td>Anti-CD45 (30-F11) - eF450</td><td>Thermo Fisher, Waltham, MA, USA</td><td>48-0451-82</td><td></td></tr><tr><td>Anti-F4/80 (BM-8) - PE Cy7</td><td>Thermo Fisher, Waltham, MA, USA</td><td>25-4801-82</td><td></td></tr><tr><td>Anti-Gr1 (RB6-8C5)</td><td>BD Biosciences, Franklin Lakes, NJ, USA</td><td>552093</td><td></td></tr><tr><td>Anti-Ly6C (HK1.4) PerCP-Cy5.5</td><td>Thermo Fisher, Waltham, MA, USA</td><td>45-5932-82</td><td></td></tr><tr><td>Anti-Ly6G (1A8) APC/Cy7</td><td>Bio Legend, San Diego, CA</td><td>127623</td><td></td></tr><tr><td>Buprenorphine hydrochloride</td><td>Indivior UK Limited, Berkshire, UK</td><td></td><td></td></tr><tr><td>C57BL/6 mice, female, 10 - 12 weeks old</td><td>Charles River, Wilmongton, MA, USA</td><td></td><td></td></tr><tr><td>CaliBRITE APC-beads (6&amp;micro;m)</td><td>BD Biosciences, Franklin Lakes, NJ, USA</td><td>340487</td><td></td></tr><tr><td>Canula 23 gauge 1&#039;&#039;</td><td>BD, Franklin Lakes, NJ, USA</td><td>300800</td><td></td></tr><tr><td>Canula 26 gauge 1/2&#039;&#039;</td><td>BD, Franklin Lakes, NJ, USA</td><td>303800</td><td></td></tr><tr><td>Cell strainer 70 &amp;micro;m</td><td>BD Biosciences, Franklin Lakes, NJ, USA</td><td>352350</td><td></td></tr><tr><td>Collagenase Type I</td><td>Sigma-Aldrich, St. Louis, MO, USA</td><td>1148089</td><td></td></tr><tr><td>Deoxyribonuclease II</td><td>Sigma-Aldrich, St. Louis, MO, USA</td><td>D8764&amp;nbsp;</td><td></td></tr><tr><td>Dulbecco&#039;s Phosphate Buffered Saline (PBS), sterile</td><td>Sigma-Aldrich, St. Louis, MO, USA</td><td>D8662</td><td></td></tr><tr><td>Dulbecco&amp;rsquo;s Phosphate Buffered Saline (PBS), without calcium chloride and magnesium chloride, sterile</td><td>Sigma-Aldrich, St. Louis, MO, USA</td><td>D8537</td><td></td></tr><tr><td>Ethylenediaminetetraacetic acid (EDTA) solution</td><td>Sigma-Aldrich, St. Louis, MO, USA</td><td>E7889</td><td></td></tr><tr><td>FACS tubes, 5 ml</td><td>Sarstedt, N&amp;uuml;mbrecht, Germany</td><td>551579</td><td></td></tr><tr><td>Fetal calf serum (FCS)</td><td>Sigma-Aldrich, St. Louis, MO, USA</td><td>F2442</td><td></td></tr><tr><td>Forceps</td><td>Fine Science Tools, Heidelberg, Germany</td><td>11049-10</td><td></td></tr><tr><td>Isoflurane</td><td>Baxter, Unterschlei&amp;szlig;heim, Germany</td><td></td><td></td></tr><tr><td>Ketamine hydrochloride</td><td>Serumwerk Bernburg, Bernburg, Germany</td><td></td><td></td></tr><tr><td>Lipopolysaccharides (LPS) from Escherichia coli O111:B4</td><td>Sigma-Aldrich, St. Louis, MO, USA</td><td>L2630</td><td></td></tr><tr><td>LIVE/DEAD Fixable Dead Cell Green Kit</td><td>Thermo Fisher, Waltham, MA, USA</td><td>L23101</td><td></td></tr><tr><td>Purified Rat Anti-Mouse CD16/CD32 (Mouse BD Fc Block&amp;trade;), Clone 2.4G2</td><td>BD, Franklin Lakes, NJ, USA</td><td>553141</td><td></td></tr><tr><td>Red blood cell lysis buffer</td><td>Thermo Fisher, Waltham, MA, USA</td><td>00-4333-57</td><td></td></tr><tr><td>RPMI-1640, with L-glutamine and sodium bicarbonate</td><td>Sigma-Aldrich, St. Louis, MO, USA</td><td>R8758</td><td></td></tr><tr><td>Scissors</td><td>Fine Science Tools, Heidelberg, Germany</td><td>14060-09</td><td></td></tr><tr><td>Sodium azide (NaN3)</td><td>Sigma-Aldrich, St. Louis, MO, USA</td><td>S2002</td><td></td></tr><tr><td>Spring scissors</td><td>Fine Science Tools, Heidelberg, Germany</td><td>15018-10</td><td></td></tr><tr><td>Tissue forceps</td><td>Fine Science Tools, Heidelberg, Germany</td><td>11021-12</td><td></td></tr><tr><td>Tubes</td><td>Eppendorf, Hamburg, Germany</td><td>30125150</td><td></td></tr><tr><td>Venous catheter, 22 gauge</td><td>B.Braun, Melsungen, Germany</td><td>4268091B</td><td></td></tr><tr><td>Xylazine hydrochloride</td><td>Serumwerk Bernburg, Bernburg, Germany</td><td></td><td></td></tr></tbody></table>

inducing acute lung injury in mice by direct intratracheal lipopolysaccharide instillation

Airway administration of lipopolysaccharide (LPS) is a common way to study pulmonary inflammation and acute lung injury (ALI) in small animal models. Various approaches have been described, such as the inhalation of aerosolized LPS as well as nasal or intratracheal instillation. The presented protocol describes a detailed step-by-step procedure to induce ALI in mice by direct intratracheal LPS instillation and perform FACS analysis of blood samples, bronchoalveolar lavage (BAL) fluid, and lung tissue. After intraperitoneal sedation, the trachea is exposed and LPS is administered via a 22 G venous catheter. A robust and reproducible inflammatory reaction with leukocyte invasion, upregulation of proinflammatory cytokines, and disruption of the alveolo-capillary barrier is induced within hours to days, depending on the LPS dosage used. Collection of blood samples, BAL fluid, and lung harvesting, as well as the processing for FACS analysis, are described in detail in the protocol. Although the use of the sterile LPS is not suitable to study pharmacologic interventions in infectious diseases, the described approach offers minimal invasiveness, simple handling, and good reproducibility to answer mechanistic immunological questions. Furthermore, dose titration as well as the use of alternative LPS preparations or mouse strains allow modulation of the clinical effects, which can exhibit different degrees of ALI severity or early vs. late onset of disease symptoms.

The described approach to induce ALI in mice was validated by assessing cytokine expression, neutrophil granulocyte infiltration, and alveolo-capillary barrier disruption 24 h and 72 h after LPS instillation. PBS-injected animals served as control. Intratracheal LPS administration induced a robust pulmonary proinflammatory response. Expression of TNF-&#945; in lung tissue was significantly upregulated, reaching a sustained and more than 50-fold increase compared to the control animals [RQ (TNF-&#945;/18s); 24 h: 53.7 (SD...

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Inducing Acute Lung Injury in Mice by Direct Intratracheal Lipopolysaccharide Instillation

Presented here is a step-by-step procedure to induce acute lung injury in mice by direct intratracheal lipopolysaccharide instillation and to perform FACS analysis of blood samples, bronchoalveolar lavage fluid, and lung tissue. Minimal invasiveness, simple handling, good reproducibility, and titration of disease severity are advantages of this approach.

Airway administration of lipopolysaccharide (LPS) is a common way to study pulmonary inflammation and acute lung injury (ALI) in small animal models. ...

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24.7K Views.  University Hospital Bonn. Presented here is a step-by-step procedure to induce acute lung injury in mice by direct intratracheal lipopolysaccharide instillation and to perform FACS analysis of blood samples, bronchoalveolar lavage fluid, and lung tissue. Minimal invasiveness, simple handling, good reproducibility, and titration of disease severity are advantages of this approach.

Video: Inducing Acute Lung Injury in Mice by Direct Intratracheal Lipopolysaccharide Instillation

Pressure Controlled Ventilation to Induce Acute Lung Injury in Mice

Murine models are extensively used to investigate acute injuries of different organs systems (1-34). Acute lung injury (ALI), which occurs with prolonged mechanical ventilation, contributes to morbidity and mortality of critical illness, and studies on novel genetic or pharmacological targets are areas of intense investigation (1-3, 5, 8, 26, 30, 33-36). ALI is defined by the acute onset of the disease, which leads to non-cardiac pulmonary edema and subsequent impairment of pulmonary gas exchange (36). We have developed a murine model of ALI by using a pressure-controlled ventilation to induce ventilator-induced lung injury (2). For this purpose, C57BL/6 mice are anesthetized and a tracheotomy is performed followed by induction of ALI via mechanical ventilation. Mice are ventilated in a pressure-controlled setting with an inspiratory peak pressure of 45 mbar over 1 - 3 hours. As outcome parameters, pulmonary edema (wet-to-dry ratio), bronchoalveolar fluid albumin content, bronchoalveolar fluid and pulmonary tissue myeloperoxidase content and pulmonary gas exchange are assessed (2). Using this technique we could show that it sufficiently induces acute lung inflammation and can distinguish between different treatment groups or genotypes (1-3, 5). Therefore this technique may be helpful for researchers who pursue molecular mechanisms involved in ALI using a genetic approach in mice with gene-targeted deletion.

A murine model for ventilator induced lung injury is an important tool to study an acute lung injury in vivo. Here, we report an easy applicable in situ model for acute lung injury using high-pressure mechanical ventilation to induce acute failure of the lung.

Murine models are extensively used to investigate acute injuries of different organs systems (1-34). Acute lung injury (ALI), which occurs with prolonged ...

The Utilization of Oropharyngeal Intratracheal PAMP Administration and Bronchoalveolar Lavage to Evaluate the Host Immune Response in Mice

The host immune response to pathogens is a complex biological process. The majority of in vivo studies classically employed to characterize host-pathogen interactions take advantage of intraperitoneal injections of select bacteria or pathogen associated molecular patterns (PAMPs) in mice. While these techniques have yielded tremendous data associated with infectious disease pathobiology, intraperitoneal injection models are not always appropriate for host-pathogen interaction studies in the lung. Utilizing an acute lung inflammation model in mice, it is possible to conduct a high resolution analysis of the host innate immune response utilizing lipopolysaccharide (LPS). Here, we describe the methods to administer LPS using nonsurgical oropharyngeal intratracheal administration, monitor clinical parameters associated with disease pathogenesis, and utilize bronchoalveolar lavage fluid to evaluate the host immune response. The techniques that are described are widely applicable for studying the host innate immune response to a diverse range of PAMPs and pathogens. Likewise, with minor modifications, these techniques can also be applied in studies evaluating allergic airway inflammation and in pharmacological applications.

The host immune response to pathogen infection is a tightly regulated process. Utilizing a lipopolysaccharide lung exposure model in mice, it is possible to conduct high resolution evaluations of the complex mechanisms associated with disease pathogenesis.

The host immune response to pathogens is a complex biological process. The majority of in vivo studies classically employed to characterize host-pathogen ...

Immunology and Infection

A Method for Generating Pulmonary Neutrophilia Using Aerosolized Lipopolysaccharide

Acute lung injury (ALI) is a severe disease characterized by alveolar neutrophilia, with limited treatment options and high mortality. Experimental models of ALI are key in enhancing our understanding of disease pathogenesis. Lipopolysaccharide (LPS) derived from gram positive bacteria induces neutrophilic inflammation in the airways and lung parenchyma of mice. Efficient pulmonary delivery of compounds such as LPS is, however, difficult to achieve. In the approach described here, pulmonary delivery in mice is achieved by challenge to aerosolized Pseudomonas aeruginosa LPS. Dissolved LPS was aerosolized by a nebulizer connected to compressed air. Mice were exposed to a continuous flow of LPS aerosol in a Plexiglas box for 10 min, followed by 2 min conditioning after the aerosol was discontinued. Tracheal intubation and subsequent bronchoalveolar lavage, followed by formalin perfusion was next performed, which allows for characterization of the sterile pulmonary inflammation. Aerosolized LPS generates a pulmonary inflammation characterized by alveolar neutrophilia, detected in bronchoalveolar lavage and by histological assessment. This technique can be set up at a small cost with few appliances, and requires minimal training and expertise. The exposure system can thus be routinely performed at any laboratory, with the potential to enhance our understanding of lung pathology.

We describe a method for inducing neutrophilic pulmonary inflammation by challenge to aerosolized lipopolysaccharide by nebulization, to model acute lung injury. In addition, basic surgical techniques for lung isolation, tracheal intubation and bronchoalveolar lavage are also described.

Acute lung injury (ALI) is a severe disease characterized by alveolar neutrophilia, with limited treatment options and high mortality. Experimental models ...

Direct Tracheal Instillation of Solutes into Mouse Lung

Intratracheal instillations deliver solutes directly into the lungs. This procedure targets the delivery of the instillate into the distal regions of the lung, and is therefore often incorporated in studies aimed at studying alveoli. We provide a detailed survival protocol for performing intratracheal instillations in mice. Using this approach, one can target delivery of test solutes or solids (such as lung therapeutics, surfactants, viruses, and small oligonucleotides) into the distal lung. Tracheal instillations may be the preferred methodology, over inhalation protocols that may primarily target the upper respiratory tract and possibly expose the investigator to potentially hazardous substances. Additionally, in using the tracheal instillation protocol, animals can fully recover from the non-invasive procedure. This allows for making subsequent physiological measurements on test animals, or reinstallation using the same animal. The amount of instillate introduced into the lung must be carefully determined and osmotically balanced to ensure animal recovery. Typically, 30-75 &mu;L instillate volume can be introduced into mouse lung.

Intratracheal instillations deliver solutes directly into the lungs. This procedure targets the delivery of the instillate into the distal regions of the lung, and is therefore often incorporated in studies aimed at studying alveoli. We provide a detailed survival protocol for performing intratracheal instillations in mice.

Intratracheal instillations deliver solutes directly into the lungs.  This procedure targets the delivery of the instillate into the distal regions of the ...

Biology

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 ...

Intra-tracheal Administration of Haemophilus influenzae in Mouse Models to Study Airway Inflammation

Here, we describe a detailed procedure to efficiently and directly deliver Haemophilus influenzae into the lower respiratory tracts of mice. We demonstrate the procedure for preparing H. influenzae inoculum, intra-tracheal instillation of H. influenzae into the lung, collection of broncho-alveolar lavage fluid (BALF), analysis of immune cells in the BALF, and RNA isolation for differential gene expression analysis. This procedure can be used to study the lung inflammatory response to any bacteria, virus or fungi. Direct tracheal instillation is mostly preferred over intranasal or aerosol inhalation procedures because it more efficiently delivers the bacterial inoculum into the lower respiratory tract with less ambiguity.

Intra-tracheal Administration of Haemophilus influenzae in Mouse Models to Study Airway Inflammation

We demonstrate the procedure for intra-tracheal inoculation of Haemophilus influenzae into the lower respiratory tracts of mice. This is a very useful tool to study signaling pathways that regulate airway inflammation in mouse models.

Here, we describe a detailed procedure to efficiently and directly deliver Haemophilus influenzae into the lower respiratory tracts of mice. We ...

An Improved Method for Rapid Intubation of the Trachea in Mice

Despite some anatomical and physiological differences, mouse models continue to be an essential tool for studying human lung disease. Bleomycin toxicity is a commonly used model to study both acute lung injury and fibrosis, and multiple methods have been developed for administering bleomycin (and other toxic agents) into the lungs. However, many of these approaches, such as transtracheal instillation, have inherent drawbacks, including the need for strong anesthetics and survival surgery. This paper reports a quick, reproducible method of intratracheal intubation that involves mild inhaled anesthesia, visualization of the trachea, and the use of a surrogate spirometer to confirm exposure. As a proof of concept, 8-12 week old C57BL/6 mice were administered either 2.0 U/kg of bleomycin or an equivalent volume of PBS, and both damage and fibrotic endpoints were measured post-exposure. This procedure allows researchers to treat a large cohort of mice in a relatively short period with little expense and minimal post-procedure care.

This article presents a rapid and simple method for administering bleomycin directly into the mouse trachea via intubation. Key advantages of this method are that it is highly reproducible, easy to master, and does not require specialized equipment or lengthy recovery times.

Despite some anatomical and physiological differences, mouse models continue to be an essential tool for studying human lung disease. Bleomycin toxicity ...

Open Tracheostomy Gastric Acid Aspiration Murine Model of Acute Lung Injury Results in Maximal Acute Nonlethal Lung Injury

Acid pneumonitis is a major cause of sterile acute lung injury (ALI) in humans. Acid pneumonitis spans the clinical spectrum from asymptomatic to acute respiratory distress syndrome (ARDS), characterized by neutrophilic alveolitis, and injury to both alveolar epithelium and vascular endothelium. Clinically, ARDS is defined by acute onset of hypoxemia, bilateral patchy pulmonary infiltrates and non-cardiogenic pulmonary edema. Human studies have provided us with valuable information about the physiological and inflammatory changes in the lung caused by ARDS, which has led to various hypotheses about the underling mechanisms. Unfortunately, difficulties determining the etiology of ARDS, as well as a wide range of pathophysiology have resulted in a lack of critical information that could be useful in developing therapeutic strategies.
Translational animal models are valuable when their pathogenesis and pathophysiology accurately reproduce a concept proven in both in vitro and clinical settings. Although large animal models (e.g., sheep) share characteristics of the anatomy of human trachea-bronchial tree, murine models provide a host of other advantages including: low cost; short reproductive cycle lending itself to greater data acquisition; a well understood immunologic system; and a well characterized genome leading to the availability of a variety of gene deletion and transgenic strains. A robust model of low pH induced ARDS requires a murine ALI that targets mainly the alveolar epithelium, secondarily the vascular endothelium, as well as the small airways leading to the alveoli. Furthermore, a reproducible injury with wide differences between different injurious and non-injurious insults is important.
The murine gastric acid aspiration model presented here using hydrochloric acid employs an open tracheostomy and recreates a pathogenic scenario that reproduces the low pH pneumonitis injury in humans. Additionally, this model can be used to examine interaction of a low pH insult with other pulmonary injurious entities (e.g., food particles, pathogenic bacteria).

This protocol induces acute lung injury in a mouse that has close fidelity to the pathogenesis of acid pneumonitis observed in humans. We generate a maximal acute nonlethal low pH lung injury and account for differences in rodent-human anatomic respiratory structure using an open tracheostomy coupled with circumferential pressure release.

Acid pneumonitis is a major cause of sterile acute lung injury (ALI) in humans. Acid pneumonitis spans the clinical spectrum from asymptomatic to acute ...

A Model of Self-limited Acute Lung Injury by Unilateral Intra-bronchial Acid Instillation

Selective intra-bronchial instillation of hydrochloric acid (HCl) to the murine left mainstem bronchus causes acute tissue injury with histopathologic findings similar to human acute respiratory distress syndrome (ARDS). The resulting alveolar edema, alveolar-capillary barrier damage, and leukocyte infiltration predominantly affect the left lung, preserving the right lung as an uninjured control and allowing animals to survive. This model of self-limited acute lung injury enables investigation of tissue resolution mechanisms, such as macrophage efferocytosis of apoptotic neutrophils and restitution of alveolar-capillary barrier integrity. This model has helped identify important roles for resolution agonists, including specialized pro-resolving mediators (SPMs), providing a foundation for the development of new therapeutic approaches for patients with ARDS.

Selective intra-bronchial acid instillation to the left lung in mice results in unilateral and self-limited acute lung injury that models human acute respiratory distress syndrome (ARDS) induced by gastric acid aspiration.

Selective intra-bronchial instillation of hydrochloric acid (HCl) to the murine left mainstem bronchus causes acute tissue injury with histopathologic ...

Visualizing Lung Cellular Adaptations during Combined Ozone and LPS Induced Murine Acute Lung Injury

Lungs are continually faced with direct and indirect insults in the form of sterile (particles or reactive toxins) and infectious (bacterial, viral or fungal) inflammatory conditions. An overwhelming host response may result in compromised respiration and acute lung injury, which is characterized by lung neutrophil recruitment as a result of the patho-logical host immune, coagulative and tissue remodeling response. Sensitive microscopic methods to visualize and quantify murine lung cellular adaptations, in response to low-dose (0.05 ppm) ozone, a potent environmental pollutant in combination with bacterial lipopolysaccharide, a TLR4 agonist, are crucial in order to understand the host inflammatory and repair mechanisms. We describe a comprehensive fluorescent microscopic analysis of various lung and systemic body compartments, namely the broncho-alveolar lavage fluid, lung vascular perfusate, left lung cryosections, and sternal bone marrow perfusate. We show damage of alveolar macrophages, neutrophils, lung parenchymal tissue, as well as bone marrow cells in correlation with a delayed (up to 36-72 h) immune response that is marked by discrete chemokine gradients in the analyzed compartments. In addition, we present lung extracellular matrix and cellular cytoskeletal interactions (actin, tubulin), mitochondrial and reactive oxygen species, anti-coagulative plasminogen, its anti-angiogenic peptide fragment angiostatin, the mitochondrial ATP synthase complex V subunits, &#945; and &#946;. These surrogate markers, when supplemented with adequate in vitro cell-based assays and in vivo animal imaging techniques such as intravital microscopy, can provide vital information towards understanding the lung response to novel immunomodulatory agents.

Combined ozone and bacterial endotoxin exposed mice show wide-spread cell death, including that of neutrophils. We observed cellular adaptations such as disruption of cytoskeletal lamellipodia, increased cellular expression of complex V ATP synthase subunit &#946; and angiostatin in broncho-alveolar lavage, suppression of the lung immune response and delayed neutrophil recruitment.

Lungs are continually faced with direct and indirect insults in the form of sterile (particles or reactive toxins) and infectious (bacterial, viral or ...

Inducing Acute Lung Injury in Mice by Direct Intratracheal Lipopolysaccharide Instillation

Summary

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Chapters in this video

Pressure Controlled Ventilation to Induce Acute Lung Injury in Mice

The Utilization of Oropharyngeal Intratracheal PAMP Administration and Bronchoalveolar Lavage to Evaluate the Host Immune Response in Mice

A Method for Generating Pulmonary Neutrophilia Using Aerosolized Lipopolysaccharide

Direct Tracheal Instillation of Solutes into Mouse Lung

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

Intra-tracheal Administration of Haemophilus influenzae in Mouse Models to Study Airway Inflammation

An Improved Method for Rapid Intubation of the Trachea in Mice

Open Tracheostomy Gastric Acid Aspiration Murine Model of Acute Lung Injury Results in Maximal Acute Nonlethal Lung Injury

A Model of Self-limited Acute Lung Injury by Unilateral Intra-bronchial Acid Instillation

Visualizing Lung Cellular Adaptations during Combined Ozone and LPS Induced Murine Acute Lung Injury