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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E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Acute+Lung+Injury+in+Response+to+Intratracheal+Instillation+of+Lipopolysaccharide+in+an+Animal+Model+of+Emphysema+Induced+by+Elastase.">Acute Lung Injury in Response to Intratracheal Instillation of Lipopolysaccharide in an Animal Model of Emphysema Induced by Elastase.</a> Inflammation. 41 (1), 174-182 (2018).</li><li>Liu, F., Li, W., Pauluhn, J., Tr&#252;bel, H., Wang, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lipopolysaccharide-induced+acute+lung+injury+in+rats:+comparative+assessment+of+intratracheal+instillation+and+aerosol+inhalation.">Lipopolysaccharide-induced acute lung injury in rats: comparative assessment of intratracheal instillation and aerosol inhalation.</a> Toxicology. 304, 158-166 (2013).</li><li>Rittirsch, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Acute+Lung+Injury+Induced+by+Lipopolysaccharide+Is+Independent+of+Complement+Activation.">Acute Lung Injury Induced by Lipopolysaccharide Is Independent of Complement Activation.</a> Journal of Immunology. 180 (11), 7664-7672 (2008).</li><li>D'Alessio, F. R., 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=CD4+CD25+Foxp3++Tregs+resolve+experimental+lung+injury+in+mice+and+are+present+in+humans+with+acute+lung+injury.">CD4+CD25+Foxp3+ Tregs resolve experimental lung injury in mice and are present in humans with acute lung injury.</a> The Journal of Clinical Investigation. 119 (10), 2898-2913 (2009).</li><li>Ehrentraut, H., Weisheit, C., Scheck, M., Frede, S., Hilbert, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Experimental+murine+acute+lung+injury+induces+increase+of+pulmonary+TIE2-expressing+macrophages.">Experimental murine acute lung injury induces increase of pulmonary TIE2-expressing macrophages.</a> Journal of Inflammation. 15, 12 (2018).</li><li>Szarka, R. J., Wang, N., Gordon, L., Nation, P. N., Smith, R. 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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 border="0" cellpadding="0" cellspacing="0" style="width:1189px;" width="1189">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:653px;">1 ml syringes</td>
			<td style="width:289px;">BD, Franklin Lakes, NJ, USA</td>
			<td style="width:104px;">300013</td>
			<td style="width:143px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">10 ml syringes</td>
			<td>BD, Franklin Lakes, NJ, USA</td>
			<td>309110</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Anti-CD115 (c-fms) APC</td>
			<td>Thermo Fisher, Waltham, MA, USA</td>
			<td>17-1152-80</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Anti-CD11b (M1/70) - FITC</td>
			<td>Thermo Fisher, Waltham, MA, USA</td>
			<td>11-0112-81</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Anti-CD45 (30-F11) - eF450</td>
			<td>Thermo Fisher, Waltham, MA, USA</td>
			<td>48-0451-82</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Anti-F4/80 (BM-8) - PE Cy7</td>
			<td>Thermo Fisher, Waltham, MA, USA</td>
			<td>25-4801-82</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Anti-Gr1 (RB6-8C5)</td>
			<td>BD Biosciences, Franklin Lakes, NJ, USA</td>
			<td>552093</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Anti-Ly6C (HK1.4) PerCP-Cy5.5</td>
			<td>Thermo Fisher, Waltham, MA, USA</td>
			<td>45-5932-82</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Anti-Ly6G (1A8) APC/Cy7</td>
			<td>Bio Legend, San Diego, CA</td>
			<td>127623</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Buprenorphine hydrochloride</td>
			<td>Indivior UK Limited, Berkshire, UK</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">C57BL/6 mice, female, 10 - 12 weeks old</td>
			<td>Charles River, Wilmongton, MA, USA</td>
			<td></td>
			<td style="width:143px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">CaliBRITE APC-beads (6&#181;m)</td>
			<td>BD Biosciences, Franklin Lakes, NJ, USA</td>
			<td>340487</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:653px;">Canula 23 gauge 1&#39;&#39;</td>
			<td>BD, Franklin Lakes, NJ, USA</td>
			<td>300800</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:653px;">Canula 26 gauge 1/2&#39;&#39;</td>
			<td>BD, Franklin Lakes, NJ, USA</td>
			<td>303800</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Cell strainer 70 &#181;m</td>
			<td>BD Biosciences, Franklin Lakes, NJ, USA</td>
			<td>352350</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Collagenase Type I</td>
			<td>Sigma-Aldrich, St. Louis, MO, USA</td>
			<td>1148089</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Deoxyribonuclease II</td>
			<td>Sigma-Aldrich, St. Louis, MO, USA</td>
			<td>D8764&#160;</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Dulbecco&#39;s Phosphate Buffered Saline (PBS), sterile</td>
			<td>Sigma-Aldrich, St. Louis, MO, USA</td>
			<td>D8662</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Dulbecco&#8217;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 height="21">
			<td height="21" style="height:21px;">Ethylenediaminetetraacetic acid (EDTA) solution</td>
			<td>Sigma-Aldrich, St. Louis, MO, USA</td>
			<td>E7889</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">FACS tubes, 5 ml</td>
			<td>Sarstedt, N&#252;mbrecht, Germany</td>
			<td>551579</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Fetal calf serum (FCS)</td>
			<td>Sigma-Aldrich, St. Louis, MO, USA</td>
			<td>F2442</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Forceps</td>
			<td>Fine Science Tools, Heidelberg, Germany</td>
			<td>11049-10</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Isoflurane</td>
			<td>Baxter, Unterschlei&#223;heim, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Ketamine hydrochloride</td>
			<td>Serumwerk Bernburg, Bernburg, Germany</td>
			<td></td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Lipopolysaccharides (LPS) from Escherichia coli O111:B4</td>
			<td>Sigma-Aldrich, St. Louis, MO, USA</td>
			<td>L2630</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">LIVE/DEAD Fixable Dead Cell Green Kit</td>
			<td>Thermo Fisher, Waltham, MA, USA</td>
			<td>L23101</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Purified Rat Anti-Mouse CD16/CD32 (Mouse BD Fc Block&#8482;), Clone 2.4G2</td>
			<td>BD, Franklin Lakes, NJ, USA</td>
			<td>553141</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Red blood cell lysis buffer</td>
			<td>Thermo Fisher, Waltham, MA, USA</td>
			<td>00-4333-57</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">RPMI-1640, with L-glutamine and sodium bicarbonate</td>
			<td>Sigma-Aldrich, St. Louis, MO, USA</td>
			<td>R8758</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Scissors</td>
			<td>Fine Science Tools, Heidelberg, Germany</td>
			<td>14060-09</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Sodium azide (NaN3)</td>
			<td>Sigma-Aldrich, St. Louis, MO, USA</td>
			<td>S2002</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Spring scissors</td>
			<td>Fine Science Tools, Heidelberg, Germany</td>
			<td>15018-10</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Tissue forceps</td>
			<td>Fine Science Tools, Heidelberg, Germany</td>
			<td>11021-12</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:653px;">Tubes</td>
			<td>Eppendorf, Hamburg, Germany</td>
			<td>30125150</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">Venous catheter, 22 gauge</td>
			<td>B.Braun, Melsungen, Germany</td>
			<td>4268091B</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;">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...

Watch this Scientific Journal Video about Inducing Acute Lung Injury in Mice by Direct Intratracheal Lipopolysaccharide Instillation at JoVE.com

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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JoVE Journal

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24.4K 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 ...

Ischemia-reperfusion Model of Acute Kidney Injury and Post Injury Fibrosis in Mice

Ischemia-reperfusion induced acute kidney injury (IR-AKI) is widely used as a model of AKI in mice, but results are often quite variable with high, often unreported mortality rates that may confound analyses. Bilateral renal pedicle clamping is commonly used to induce IR-AKI, but differences between effective clamp pressures and/or renal responses to ischemia between kidneys often lead to more variable results. In addition, shorter clamp times are known to induce more variable tubular injury, and while mice undergoing bilateral injury with longer clamp times develop more consistent tubular injury, they often die within the first 3 days after injury due to severe renal insufficiency. To improve post-injury survival and obtain more consistent and predictable results, we have developed two models of unilateral ischemia-reperfusion injury followed by contralateral nephrectomy. Both surgeries are performed using a dorsal approach, reducing surgical stress resulting from ventral laparotomy, commonly used for mouse IR-AKI surgeries. For induction of moderate injury BALB/c mice undergo unilateral clamping of the renal pedicle for 26 min and also undergo simultaneous contralateral nephrectomy. Using this approach, 50-60% of mice develop moderate AKI 24 hr after injury but 90-100% of mice survive. To induce more severe AKI, BALB/c mice undergo renal pedicle clamping for 30 min followed by contralateral nephrectomy 8 days after injury. This allows functional assessment of renal recovery after injury with 90-100% survival. Early post-injury tubular damage as well as post injury fibrosis are highly consistent using this model.

We describe models of moderate and severe ischemia-reperfusion-induced kidney injury in which the mice undergo unilateral renal pedicle clamping followed by simultaneous or delayed contralateral nephrectomy, respectively. These models consistently give rise to renal dysfunction and post-injury fibrosis, but injury severity and survival are dependent on mouse background, age and surgical equipment.

Ischemia-reperfusion induced acute kidney injury (IR-AKI) is widely used as a model of AKI in mice, but results are often quite variable with high, often ...

Acute Brain Trauma in Mice Followed By Longitudinal Two-photon Imaging

Although acute brain trauma often results from head damage in different accidents and affects a substantial fraction of the population, there is no effective treatment for it yet. Limitations of currently used animal models impede understanding of the pathology mechanism. Multiphoton microscopy allows studying cells and tissues within intact animal brains longitudinally under physiological and pathological conditions. Here, we describe two models of acute brain injury studied by means of two-photon imaging of brain cell behavior under posttraumatic conditions. A selected brain region is injured with a sharp needle to produce a trauma of a controlled width and depth in the brain parenchyma. Our method uses stereotaxic prick with a syringe needle, which can be combined with simultaneous drug application. We propose that this method can be used as an advanced tool to study cellular mechanisms of pathophysiological consequences of acute trauma in mammalian brain in vivo. In this video, we combine acute brain injury with two preparations: cranial window and skull thinning. We also discuss advantages and limitations of both preparations for multisession imaging of brain regeneration after trauma.

Acute brain trauma is a severe injury that has no adequate treatment to date. Multiphoton microscopy allows studying longitudinally the process of acute brain trauma development and probing therapeutical strategies in rodents. Two models of acute brain trauma studied with in vivo two-photon imaging of brain are demonstrated in this protocol.

Although acute brain trauma often results from head damage in different accidents and affects a substantial fraction of the population, there is no ...

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 ...

Instillation and Fixation Methods Useful in Mouse Lung Cancer Research

The ability to instill live agents, cells, or chemicals directly into the lung without injuring or killing the mice is an important tool in lung cancer research. Although there are a number of methods that have been published showing how to intubate mice for pulmonary function measurements, none are without potential problems for rapid tracheal instillation in large cohorts of mice. In the present paper, a simple and quick method is described that enables an investigator to carry out such instillations in an efficient manner. The method does not require any special tools or lighting and can be learned with very little practice. It involves anesthetizing a mouse, making a small incision in the neck to visualize the trachea, and then inserting an intravenous catheter directly. The small incision is quickly closed with tissue adhesive, and the mice are allowed to recover. A skilled student or technician can do instillations at an average rate of 2 min/mouse. Once the cancer is established, there is frequently a need for quantitative histologic analysis of the lungs. Traditionally pathologists usually do not bother to standardize lung inflation during fixation, and analyses are often based on a scoring system that can be quite subjective. While this may sometime be sufficiently adequate for gross estimates of the size of a lung tumor, any proper stereological quantification of lung structure or cells requires a reproducible fixation procedure and subsequent lung volume measurement. Here we describe simple reliable procedures for both fixing the lungs under pressure and then accurately measuring the fixed lung volume. The only requirement is a laboratory balance that is accurate over a range of 1 mg&#8211;300 g. The procedures presented here thus could greatly improve the ability to create, treat, and analyze lung cancers in mice.

The goal of this paper is to describe simple methods that will greatly aid in the setup and analysis of mouse lungs with lung cancer or other pathologies. We present 3 protocols to simply and reliably carry out lung instillations, fixation, and lung volume measurements.

The ability to instill live agents, cells, or chemicals directly into the lung without injuring or killing the mice is an important tool in lung cancer ...

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 Pleural Effusion Model in Rats by Intratracheal Instillation of Polyacrylate/Nanosilica

Pleural effusion is a prevalent clinical finding of many pulmonary diseases. Having a useful animal pleural effusion model is very important to study these pulmonary diseases. Previous pleural effusion models paid more attention to biological factors rather than nanoparticles in the environment. Here, we introduce a model to make pleural effusion in rats by intratracheal instillation of polyacrylate/nanosilica, and a method of nanoparticle isolation in the pleural effusion. By intratracheal instillation of polyacrylate/nanosilica with concentrations of 3.125, 6.25 and 12.5 mg/kg&#8729;mL, the pleural effusion in rats presented on day 3, peaked at days 7-10 in 6.25 and 12.5 mg/kg&#8729;mL groups, then slowly decreased and disappeared on day 14. When the concentration of polyacrylate/nanosilica increased, the pleural effusion is produced more and faster. This pleural fluid was detected by ultrasound examination or CT chest scanning and confirmed by dissection of rats. Silica nanoparticles were observed in the rats' pleural effusion by transmission electron microscope. These results showed that the exposure to polyacrylate/nanosilica leads to the induction of pleural effusion, which was consistent with our previous report in humans. Additionally, this model is beneficial for the further study of nanotoxicology and the pleural effusion diseases.

Here, we present a protocol to construct a pleural effusion model in rats by intratracheal instillation of polyacrylate/nanosilica.

Pleural effusion is a prevalent clinical finding of many pulmonary diseases. Having a useful animal pleural effusion model is very important to study ...

Inducing Acute Liver Injury in Rats via Carbon Tetrachloride (CCl4) Exposure Through an Orogastric Tube

Acute liver injury (ALI) plays a crucial role in the development of hepatic failure, which is characterized by severe liver dysfunction including complications such as hepatic encephalopathy and impaired protein synthesis. Appropriate animal models are vital to test the mechanism and pathophysiology of ALI and investigate different hepatoprotective strategies. Due to its ability to perform chemical transformations, carbon tetrachloride (CCl4) is widely used in the liver to induce ALI through the formation of reactive oxygen species. CCl4 exposure can be performed intraperitoneally, by inhalation, or through a nasogastric or orogastric tube. Here, we describe a rodent model, in which ALI is induced by CCl4 exposure through an orogastric tube. This method is inexpensive, easily performed, and has minimal hazard risk. The model is highly reproducible and can be widely used to determine the efficacy of potential hepatoprotective strategies and assess markers of liver injury.

Inducing Acute Liver Injury in Rats via Carbon Tetrachloride CCl4 Exposure Through an Orogastric Tube

This protocol describes a common and feasible method of inducing acute liver injury (ALI) via CCl4 exposure through an orogastric tube. CCl4 exposure induces ALI through the formation of reactive oxygen species during its biotransformation in the liver. This method is used to analyze the pathophysiology of ALI and examine different hepatoprotective strategies.

Acute liver injury (ALI) plays a crucial role in the development of hepatic failure, which is characterized by severe liver dysfunction including ...

Intratracheal Instillation of Stem Cells in Term Neonatal Rats

Prolonged exposure to high concentrations of oxygen leads to inflammation and acute lung injury, which is similar to human bronchopulmonary dysplasia (BPD). In premature infants, BPD is a major complication despite early use of surfactant therapy, optimal ventilation strategies, and noninvasive positive pressure ventilation. Because pulmonary inflammation plays a crucial role in the pathogenesis of BPD, corticosteroid use is one potential treatment to prevent it. Nevertheless, systemic corticosteroid treatment is not usually recommended for preterm infants due to long-term adverse effects. Preclinical studies and human phase I clinical trials demonstrated that use of mesenchymal stromal cells (MSCs) in hyperoxia-induced lung injuries and in preterm infants is safe and feasible. Intratracheal and intravenous MSC transplantation has been shown to protect against neonatal hyperoxic lung injury. Therefore, intratracheal administration of stem cells and combined surfactant and glucocorticoid treatment has emerged as a new strategy to treat newborns with respiratory disorders. The developmental stage of rat lungs at birth is equivalent to that in human lungs at 26&#8722;28 week of gestation. Hence, newborn rats are appropriate for studying intratracheal administration to preterm infants with respiratory distress to evaluate its efficacy. This intratracheal instillation technique is a clinically viable option for delivery of stem cells and drugs into the lungs.

Described is a protocol for performing intratracheal transplantation of mesenchymal stromal cells (MSCs) through intratracheal injection in term neonatal rats. This technique is a clinically viable option for delivery of stem cells and drugs into neonatal rat lungs to evaluate their efficacy.

Prolonged exposure to high concentrations of oxygen leads to inflammation and acute lung injury, which is similar to human bronchopulmonary dysplasia ...

Intratracheal Administration of Dry Powder Formulation in Mice

In the development of inhalable dry powder formulations, it is essential to evaluate their biological activities in preclinical animal models. This paper introduces a noninvasive method of intratracheal delivery of dry powder formulation in mice. A dry powder loading device that consists of a 200 &#181;L gel loading pipette tip connected to an 1 mL syringe via a three-way stopcock is presented. A small amount of dry powder (1-2 mg) is loaded into the pipette tip and dispersed by 0.6 mL of air in the syringe. Because pipette tips are disposable and inexpensive, different dry powder formulations can be loaded into different tips in advance. Various formulations can be evaluated in the same animal experiment without device cleaning and dose refilling, thereby saving time and eliminating the risk of cross-contamination from residual powder. The extent of powder dispersion can be inspected by the amount of powder remaining in the pipette tip. A protocol of intubation in mouse with a custom-made light source and a guiding cannula is included. Proper intubation is one of the key factors that influences the intratracheal delivery of dry powder formulation to the deep lung region of the mouse.

Dry powder formulations for inhalation have great potential in treating respiratory diseases. Before entering human studies, it is necessary to evaluate the efficacy of the dry powder formulation in preclinical studies. A simple and noninvasive method of the administration of dry powder in mice through the intratracheal route is presented.

In the development of inhalable dry powder formulations, it is essential to evaluate their biological activities in preclinical animal models. This paper ...