We are grateful to the Laboratory Animal Center, Beijing Clinical Research Institute, and Beijing Friendship Hospital for providing animal care. This work was supported by the National Key Technologies R&#38;D Program (No. 2015BAI13B09), Beijing Natural Science Foundation (No. 7232035), National Natural Science Foundation of China (No. 82171823, 82374190), and Distinguished Young Scholars from Beijing Friendship Hospital (No. yyqcjh2022-4).

Animal care and experimental procedures were conducted in accordance with the Guiding Principles in the Care and Use of Animals (China) and were approved by the Ethics Review Committee of Beijing Friendship Hospital (NO. 20-2056). C57BL/6 mice (8-12 weeks old) were used for the study.
1. Preparation for surgery
<ol>
	<li>Fast all mice for 12 h prior to the planned modeling to meet the standardization of experimental conditions (Figure 1A).</li>
	<li>Prepare and sterilize surgical instruments.</li>
	<li>Prepare the anesthetic tribromoethanol and the heating pad. Monitor the heating pad to prevent overheating and keep it at a consistent temperature (37.5 &#176;C).</li>
</ol>
2. Anesthesia
<ol>
	<li>Anesthetize each mouse with tribromoethanol.
	<ol>
		<li>Dilute tribromoethanol to prepare a 20 mg/mL solution in normal saline. Administer 0.2 mL of the tribromoethanol working solution per 10 g of body weight via intraperitoneal injection. Use ophthalmic ointment on mice&#39;s eyes to prevent dryness while under anesthesia.</li>
		<li>Ensure mice are completely anesthetized within 5 min and remain anesthetized for 20 min.</li>
	</ol>
	</li>
	<li>Assess the depth of anesthesia by observing the inability of the mouse to remain upright and check for muscle relaxation. Observe the loss of voluntary movement, blink reflex, and response to reflex stimulation (toe or tail pinch with firm pressure).</li>
	<li>Assess the respiratory rate and pattern of the mouse by monitoring the movement of the chest wall and abdomen. Ensure the breathing rate is ~ 55-65 breaths/min under optimal anesthesia.</li>
	<li>Place the anesthetized mouse on a board and secure it with tape.</li>
	<li>After the mice were completely anesthetized, use an electric hair shaver to remove hair from the abdomen. After shaving, wipe off all loose hair on the abdomen using a saline-moistened cotton ball. 
	NOTE: Ensure proper ventilation and use of a fume hood or biosafety cabinet when handling tribromoethanol to minimize exposure risks. During the procedure, wear appropriate personal protective equipment, including gloves and lab coats, to prevent accidental contact with chemicals and biological materials.</li>
</ol>
3. Surgery
<ol>
	<li>Fully extend the limbs to expose the abdomen. Ensure that the mouse&#39;s head is positioned to maintain a clear airway (Figure 1B).</li>
	<li>Disinfect the skin of the surgical area twice using a cotton ball soaked with 75% alcohol. After disinfection, use a dry, sterile medical gauze to remove excess alcohol from the abdomen. Administer lidocaine (10 mg/kg, subcutaneously) at the intended site prior to the incision.</li>
	<li>Use a sterile scalpel to make an incision in the skin and lift the rectus abdominis muscle in the middle of the abdomen using tweezers.
	<ol>
		<li>Make a small incision along the median line of the rectus abdominis muscle, taking care to avoid injury to various organs in the abdomen.</li>
		<li>Ensure the approximate range of the incision is as follows. Ensure that the upper margin of the incision is 6-8 mm from the xiphoid process of the sternum, the lower margin is 6-8 mm from the external genitalia, and the length of the incision is ~1 cm.</li>
	</ol>
	</li>
	<li>Place a piece of sterile surgical drape or sterile gauze&#160;on both sides of the abdominal incision. Use hemostatic forceps to fix the sterile gauze on the upper and lower edges of the incision, exposing the incision (Figure 1C). 
	NOTE: In the sham group, the incision was covered with wet gauze for 5 min without any surgical operation.</li>
	<li>Properly fix the sterile drape or gauze, then use two cotton swabs pre-moistened with saline to gently press against both sides of the abdominal wall adjacent to the incision. Squeeze out a small amount of the intestinal tube through the incision and expose it by placing it on the sterile drape or gauze (Figure 1D).</li>
	<li>Completely moisten two sterile cotton swabs with normal saline. Gently grasp the intestinal tissue with the moistened cotton swabs and carefully remove the small intestine.
	<ol>
		<li>Locate the caecum and then take out the intestine with the sterile cotton swabs until 2 cm before the stomach to avoid touching the pancreas. Extend the intestine from the proximal end of the pancreato-duodenal ligament to the distal end of the ileocecal region.</li>
	</ol>
	</li>
	<li>Apply consistent pressure along the entire small intestine from the proximal to the distal ends. Ensure uniform force application for 5 min until small bleeding spots emerge on the intestinal surface (Figure 1E).</li>
	<li>After 5 min, use pre-moistened sterile cotton swabs to carefully place all the small intestines back into the abdominal cavity, following the normal physiological anatomical position of the small intestine.</li>
	<li>Gently massage the abdomen across the sterile gauze and abdominal wall for 3-5 s to ensure that the bowel is restored to its natural anatomical position and to prevent artificial mechanical intestinal obstruction or mesenteric torsion after surgery.</li>
	<li>Inject 100 &#181;L of saline into the abdominal cavity to replace lost fluids during the operation and lubricate the abdominal tissue.</li>
	<li>Close the abdomen using 6-0 surgical suture and perform a two-layer closure at the abdominal incision. Begin by closing the muscle layer with continuous sutures, taking care to avoid damage to the abdominal organs. Lift the rectus abdominis muscle during suturing (Figure 1F).</li>
	<li>After completely closing the rectus abdominis muscle, suture it completely with simple intermittent sutures using a 6-0 surgical suture. Maintain a stitch length of ~0.5 mm and a needle spacing of ~2 mm.</li>
	<li>After closing the abdominal incision, gently wipe the area near the incision with dry sterile medical gauze to keep it dry and clean from blood, tissue fluid, or normal saline.</li>
	<li>Apply a small amount of medical incision adhesive to and around the incision to avoid splitting after surgery.</li>
	<li>After the medical incision adhesive has completely dried, transfer the mice to a paper towel, and place the cage paper towel on a heated blanket maintained at a constant temperature of 37.5 &#176;C until the mice fully recover. Do not leave the animals unattended until they have regained sufficient consciousness to maintain sternal recumbency.</li>
	<li>Transfer the awake mice to the animal feeding barrier environment and allow them to drink freely until the detection time.</li>
</ol>
4. Gastrointestinal transit assay
<ol>
	<li>Administrate 200 &#181;L of FITC-dextran solution (5 mg/mL in PBS) to the fasted mice via gavage 22.5 h after surgery (Figure 1A).</li>
	<li>Euthanize the mice using a method approved by the Institutional Animal Care and Use Committee (IACUC), such as CO2 inhalation followed by cervical dislocation.
	<ol>
		<li>To isolate the digestive tract, make a midline incision from the diaphragm to the pelvis, carefully removing the surrounding connective tissues and organs.</li>
		<li>Separate the digestive tract from the surrounding tissues and organs, ensuring that the entire tract from the stomach to the anus is included.</li>
	</ol>
	</li>
	<li>Divide the digestive tract into 15 equal segments (from stomach to colon), numbered sequentially from 1 to 15 based on physiological position (sequence: St -S1:S10-Ce-L1:L3) (Figure 2A).</li>
	<li>Cut each segment of intestinal tissue with its content into pieces measuring 1-4 mm and place them into separate 1.5 mL centrifuge tubes containing 1 mL of DPBS.</li>
	<li>Vortex the centrifuge tubes for 10 s to homogenize the samples.</li>
	<li>Centrifuge the tubes at 500 &#215; g for 1 min. Collect the supernatant for FITC fluorescence quantification using a multimode microplate reader.</li>
	<li>Describe the gastrointestinal transit by calculating the geometric center (GC) of the FITC-dextran and calculate using the following formula: (&#931;[% FITC per segment x segment number])/100.</li>
</ol>
5. Paraffin embedding and hematoxylin and eosin (HE) staining
<ol>
	<li>Euthanize the mice by CO2 inhalation followed by cervical dislocation after 24 h. Perform paraffin embedding and gastrointestinal assays on the same animal.</li>
	<li>Open the abdominal cavity and collect small intestinal tissue for further analysis.</li>
	<li>Fix the entire bowel with intestinal content in Carnoy&#39;s fixative (60% methanol + 10% acetic acid + 30% chloroform) at 4 &#176;C for 2 h.</li>
	<li>Deacidify the tissue by rinsing twice with methanol for 30 min each time.</li>
	<li>Replace methanol with ethanol twice for 30 min each time.</li>
	<li>Deacidify the tissue by rinsing twice with methanol for 30 min each time. To achieve tissue transparency with xylene, immerse the tissue in xylene and incubate for 1 h.</li>
	<li>Embed the tissue in paraffin wax using a paraffin embedding machine. Cut the embedded tissue into 4 &#181;m thick slices using a microtome. 
	NOTE: Divide the small intestine into different waxes for improved performance. Properly label the segments to identify their location.</li>
	<li>Perform HE staining using the HE staining kit. Seal the slides and examine them under a microscope for analysis.</li>
</ol>
6. Immune cell isolation and flow cytometry
<ol>
	<li>Prepare the following solutions.
	<ol>
		<li>Predigestion solution: Prepare Hank&#39;s balanced salt solution without Ca2+ and Mg2+ with 1 mM dithiothreitol (DTT) and 10 mM ethylenediaminetetraacetic acid (EDTA).</li>
		<li>Digestion solution: Prepare a solution containing 200 &#181;g/mL DNAase I, 500 &#181;g/mL collagenase IV, 4% FBS, and 100 &#181;M HEPES buffer in RPMI 1640.</li>
		<li>Magnetic-activated cell sorting works (MACS) buffer: Prepare a buffer containing 5 g/L BSA and 2 mM EDTA in PBS without Ca2+ or Mg2+.</li>
	</ol>
	</li>
	<li>Remove fecal contents by washing the intestines with normal saline. Remove Peyer&#39;s patches and connective tissue of the small intestine. Cut the small intestines open longitudinally and then cut into 1 cm segments in ice-cold PBS.</li>
	<li>Wash the intestine segments with predigestion solution at 37 &#176;C for 20 min on an orbital shaker. Collect the predigestion solution and filter it through 70 &#181;m cell strainers to obtain intraepithelial lymphocytes (IELs).</li>
	<li>Digest the segments in the digestion solution for 30 min and then filter the suspension through 70 &#181;m cell strainers to obtain lamina propria lymphocytes (LPLs).</li>
	<li>Centrifuge the LPL cells at 500 &#215; g for 10 min, and discard the supernatant.</li>
	<li>Wash and suspend the cells with DPBS (sodium azide, Tris, and protein-free) at a concentration of 1 x 106 cells/100 &#181;L.</li>
	<li>Incubate the cells with a fluorescent viability dye (1:500) for 15 min at room temperature (RT) in the dark.</li>
	<li>Suspend the cells with MACS buffer and centrifuge them at 500 &#215; g for 5 min.</li>
	<li>Count the stained cells using a hemocytometer or automated cell counter. Resuspend the cells at a concentration of 1 x 106 cells/100 &#181;L of MACS buffer. Perform antibody staining (1:200-1:400) by adding the appropriate volume of antibody to the cell suspension and incubating at 4 &#176;C for 15 min.</li>
	<li>Wash the cells as described in step 6.8.</li>
	<li>Detect the samples using flow cytometry and analyze data with the associated flow cytometry software7,8.</li>
</ol>

Murine Model for Studying Postoperative Ileus through Intestinal Manipulation

<ol>
	<li>Chapman, S. J., Pericleous, A., Downey, C., Jayne, D. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Postoperative+ileus+following+major+colorectal+surgery.">Postoperative ileus following major colorectal surgery.</a> Br J Surg. 105 (7), 797-810 (2018).</li><li>Farro, G., 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=CCR2-dependent+monocyte-derived+macrophages+resolve+inflammation+and+restore+gut+motility+in+postoperative+ileus.">CCR2-dependent monocyte-derived macrophages resolve inflammation and restore gut motility in postoperative ileus.</a> Gut. 66 (12), 2098-2109 (2017).</li><li>Wouters, M. M., Vicario, M., Santos, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+mast+cells+in+functional+GI+disorders.">The role of mast cells in functional GI disorders.</a> Gut. 65 (1), 155-168 (2016).</li><li>Engel, D. 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=T+helper+type+1+memory+cells+disseminate+postoperative+ileus+over+the+entire+intestinal+tract.">T helper type 1 memory cells disseminate postoperative ileus over the entire intestinal tract.</a> Nat Med. 16 (12), 1407-1413 (2010).</li><li>Schwartz, C., Voehringer, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+murine+basophils+by+flow+cytometry+and+histology.">Identification of murine basophils by flow cytometry and histology.</a> Methods Mol Biol. 2163, 367-375 (2020).</li><li>Funk, E. K., Weissbrod, P., Horgan, S., Orosco, R. K., Califano, J. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preclinical+experience+with+a+novel+single-port+platform+for+transoral+surgery.">Preclinical experience with a novel single-port platform for transoral surgery.</a> Surg Endosc. 35 (8), 4857-4864 (2021).</li><li>Nemoto, S., Mailloux, A. W., Kroeger, J., Mul&#233;, J. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=OMIP-031:+Immunologic+checkpoint+expression+on+murine+effector+and+memory+T-cell+subsets.">OMIP-031: Immunologic checkpoint expression on murine effector and memory T-cell subsets.</a> Cytometry A. 89 (5), 427-429 (2016).</li><li>Brandi, J., Wiethe, C., Riehn, M., Jacobs, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=OMIP-93:+A+41-color+high+parameter+panel+to+characterize+various+co-inhibitory+molecules+and+their+ligands+in+the+lymphoid+and+myeloid+compartment+in+mice.">OMIP-93: A 41-color high parameter panel to characterize various co-inhibitory molecules and their ligands in the lymphoid and myeloid compartment in mice.</a> Cytometry A. 103 (8), 624-630 (2023).</li><li>Swidsinski, A., Weber, J., Loening-Baucke, V., Hale, L. P., Lochs, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatial+organization+and+composition+of+the+mucosal+flora+in+patients+with+inflammatory+bowel+disease.">Spatial organization and composition of the mucosal flora in patients with inflammatory bowel disease.</a> J Clin Microbiol. 43 (7), 3380-3389 (2005).</li><li>Bialkowska, A. B., Ghaleb, A. M., Nandan, M. O., Yang, V. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Improved+Swiss-rolling+technique+for+intestinal+tissue+preparation+for+immunohistochemical+and+immunofluorescent+analyses.">Improved Swiss-rolling technique for intestinal tissue preparation for immunohistochemical and immunofluorescent analyses.</a> J Vis Exp. (113), e54161 (2016).</li><li>van Bree, S. H., 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=Novel+method+for+studying+postoperative+ileus+in+mice.">Novel method for studying postoperative ileus in mice.</a> Int J Physiol Pathophysiol Pharmacol. 4 (4), 219-227 (2012).</li></ol>

The authors have nothing to disclose.

The success of surgery relies on several critical steps. First, maintaining consistency during intestinal intramural (IM) surgery is imperative to induce extensive injury to the small intestine. Proper pressure applied during the IM procedure and the resulting hyperemic effect on the intestine are crucial for surgical success. The observation of the entire digestive tract turning pink and displaying red hemorrhagic spots after rubbing with a cotton swab served as an indicator of a successful operation. Additionally, ensu...

Postoperative ileus (POI) is a syndrome that poses a significant challenge in the field of human health, particularly in the management of patients undergoing abdominal surgery. Characterized by delayed recovery of gastrointestinal motility, POI contributes to prolonged hospital stays and increased health care costs, yet no established definition, etiology, or treatment exists1. Recent research has shed light on the pivotal role of immune cells in the progression of POI2,3,4, yet further investigation is required to elucidate the underlying mechanisms involved.
In this protocol, we introduce a murine model of POI induced by intra-abdominal surgery, which closely mimics the impact of abdominal surgery on the digestive tract. Our goal was to provide a standardized method for modeling POI in mice, enabling researchers to investigate its pathophysiology and explore novel therapeutic interventions.
The rationale behind the development and utilization of this technique lies in the need for reliable preclinical models to study POI. Traditional approaches to studying POI often lack translational relevance or fail to capture the complex interplay of factors contributing to the condition. By introducing a murine model that closely replicates the clinical scenario, researchers can more accurately investigate the mechanisms underlying POI and test potential therapeutic interventions in a controlled experimental setting.
Compared to alternative techniques, the murine model of POI presented in this protocol offers several advantages. Initially, we integrated our experimental findings with recent advancements to establish a standardized and reproducible protocol for inducing POI in experimental animals. This protocol facilitates consistent assessment of gastrointestinal transit function. Second, employing histological staining and flow cytometry enabled the assessment of tissue swelling, immune cell proliferation, and activation, yielding valuable insights into the inflammatory processes underlying POI5.
In the broader context of the literature, establishing a murine model of POI contributes to the expanding body of research aimed at comprehending the pathophysiology of this condition. By bridging the gap between basic science and clinical practice, preclinical models play a pivotal role in developing novel therapeutic strategies for POI6. Moreover, the availability of standardized animal models enhances the reproducibility and comparability of research findings across different laboratories. However, this POI model relies on mechanical stimulation during the surgical procedure. Other forms of stimulation-induced ileus may not be suitable for this model. Additionally, researchers should consider factors such as animal welfare regulations, ethical considerations, and resource availability when planning experiments using this model.
In summary, the introduction of a murine model of POI signifies a noteworthy advancement in preclinical research on this debilitating condition. Additionally, we employed H&#38;E staining and flow cytometry to assess tissue swelling and immune cell proliferation and activation. The establishment of a murine POI model would facilitate the discovery of POI mechanisms and promote the development of novel therapies for POI.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>1 M HEPES</td>
			<td>Thermo</td>
			<td>15630080</td>
			<td></td>
		</tr>
		<tr>
			<td>APC anti-mouse I-A/I-E (MHC-II)</td>
			<td>Biolegend</td>
			<td>107614</td>
			<td></td>
		</tr>
		<tr>
			<td>APC anti-mouse TCRb</td>
			<td>Biolegend</td>
			<td>109212</td>
			<td></td>
		</tr>
		<tr>
			<td>APC/Cy7 anti-mouse CD4</td>
			<td>Biolegend</td>
			<td>100414</td>
			<td></td>
		</tr>
		<tr>
			<td>APC/Cy7 anti-mouse Ly6G</td>
			<td>Biolegend</td>
			<td>127624</td>
			<td></td>
		</tr>
		<tr>
			<td>Brilliant Violet 421 anti-mouse CD69</td>
			<td>Biolegend</td>
			<td>104545</td>
			<td></td>
		</tr>
		<tr>
			<td>Brilliant Violet 421 anti-mouse F4/80</td>
			<td>Biolegend</td>
			<td>123132</td>
			<td></td>
		</tr>
		<tr>
			<td>Brilliant Violet 785 anti-mouse/human CD44</td>
			<td>Biolegend</td>
			<td>103041</td>
			<td></td>
		</tr>
		<tr>
			<td>BUV395 anti-mouse CD8a</td>
			<td>BD</td>
			<td>563786</td>
			<td></td>
		</tr>
		<tr>
			<td>BUV737 anti-mouse CD3e</td>
			<td>BD</td>
			<td>612771</td>
			<td></td>
		</tr>
		<tr>
			<td>Collagenase IV</td>
			<td>Sigma-Aldrich</td>
			<td>C5138</td>
			<td></td>
		</tr>
		<tr>
			<td>Culture Microscope</td>
			<td>CKX53</td>
			<td>Olympus</td>
			<td></td>
		</tr>
		<tr>
			<td>Deoxyribonuclease I from bovine pancreas (DNase I)</td>
			<td>Sigma-Aldrich</td>
			<td>DN25-5G</td>
			<td></td>
		</tr>
		<tr>
			<td>DL-Dithiothreitol solution</td>
			<td>Sigma-Aldrich</td>
			<td>43816-10ML</td>
			<td></td>
		</tr>
		<tr>
			<td>EDTA</td>
			<td>Sigma-Aldrich</td>
			<td>EDS-100G</td>
			<td></td>
		</tr>
		<tr>
			<td>FITC anti-mouse CD45</td>
			<td>Biolegend</td>
			<td>147709</td>
			<td></td>
		</tr>
		<tr>
			<td>FITC-dextran (70 kWM)</td>
			<td>Sigma-Aldrich</td>
			<td>FD70-100MG</td>
			<td>Gastrointestinal Transit Assay</td>
		</tr>
		<tr>
			<td>HE staining kit</td>
			<td>solarbio</td>
			<td>G1120</td>
			<td></td>
		</tr>
		<tr>
			<td>PE anti-mouse CD11b</td>
			<td>Biolegend</td>
			<td>101208</td>
			<td></td>
		</tr>
		<tr>
			<td>PE anti-mouse PD-1</td>
			<td>Biolegend</td>
			<td>114118</td>
			<td></td>
		</tr>
		<tr>
			<td>PE/Cy7 anti-mouse CD11c</td>
			<td>Biolegend</td>
			<td>117318</td>
			<td></td>
		</tr>
		<tr>
			<td>Percoll</td>
			<td>GE (Pharmacia)</td>
			<td>17-0891-01</td>
			<td></td>
		</tr>
		<tr>
			<td>Symphony A5 Flow cytometer</td>
			<td>BD</td>
			<td>-</td>
			<td>Immune cell detection and sorting</td>
		</tr>
		<tr>
			<td>Tribromoethanol</td>
			<td>Sigma-Aldrich</td>
			<td>T48402</td>
			<td>Anesthesia</td>
		</tr>
		<tr>
			<td>Varioskan LUX</td>
			<td>Thermo</td>
			<td>N16699</td>
			<td>Multimode microplate reader</td>
		</tr>
		<tr>
			<td>Zombie Aqua Fixable Viability kit</td>
			<td>Biolegend</td>
			<td>423102</td>
			<td>Fluorescent viability dye&#160;</td>
		</tr>
	</tbody>
</table>

postoperative ileus murine model

Most patients experience postoperative ileus (POI) after surgery, which is associated with increased morbidity, mortality, and hospitalization time. POI is a consequence of mechanical damage during surgery, resulting in disruption of motility in the gastrointestinal tract. The mechanisms of POI are related to aberrant neuronal sensitivity, impaired epithelial barrier function, and increased local inflammation. However, the details remain enigmatic. Therefore, experimental murine models are crucial for elucidating the pathophysiology and mechanism of POI injury and for the development of novel therapies.
Here, we introduce a murine model of POI generated via intestinal manipulation (IM) that is similar to clinical surgery; this is achieved by mechanical damage to the small intestine by massaging the abdomen 1-3 times with a cotton swab. IM delayed gastrointestinal transit 24 h after surgery, as assessed by FITC-dextran gavage and fluorescence detection of the segmental digestive tract. Moreover, tissue swelling of the submucosa and immune cell infiltration were investigated by hematoxylin and eosin staining and flow cytometry. Proper pressure of the IM and a hyperemic effect on the intestine are critical for the procedure. This murine model of POI can be utilized to study the mechanisms of intestinal damage and recovery after abdominal surgery.

In this protocol, POI was surgically induced by intestinal manipulation (IM), which is similar to the effect of clinical surgery. In the sham group, an incision was made without the IM. POI mice were sacrificed 24 h post-POI surgery along with sham control mice. The critical function of the digestive tract, content transit function, was detected by gavage of FITC-dextran. The POI model was considered successful because the FITC intensity increased in the proximal part of the small intestine&#160;(Fig...

Postoperative Ileus Murine Model

We describe a murine model of postoperative ileus generated via intestinal manipulation. Gastrointestinal transit function, pathologic changes, and immune cell activation were assessed 24 h after surgery.

Most patients experience postoperative ileus (POI) after surgery, which is associated with increased morbidity, mortality, and hospitalization time. POI ...

Universidad de Cartagena UDEC

Research

JoVE Journal

Immunology and Infection

419 Views.  Capital Medical University. We describe a murine model of postoperative ileus generated via intestinal manipulation. Gastrointestinal transit function, pathologic changes, and immune cell activation were assessed 24 h after surgery.

Video: Postoperative Ileus Murine Model

Murine Model of CD40-activation of B cells

Research on B cells has shown that CD40 activation improves their antigen presentation capacity. When stimulated with interleukin-4 and CD40 ligand (CD40L), human B cells can be expanded without difficulties from small amounts of peripheral blood within 14 days to very large amounts of highly-pure CD40-B cells (&gt;109 cells per patient) from healthy donors as well as cancer patients1-4. CD40-B cells express important lymph node homing molecules and can attract T cells in vitro5. Furthermore they efficiently take up, process and present antigens to T cells6,7. CD40-B cells were shown to not only prime na&iacute;ve, but also expand memory T cells8,9. Therefore CD40-activated B cells (CD40-B cells) have been studied as an alternative source of immuno-stimulatory antigen-presenting cells (APC) for cell-based immunotherapy1,5,10. In order to further study whether CD40-B cells induce effective T cell responses in vivo and to study the underlying mechanism we established a cell culture system for the generation of murine CD40-activated B cells. Using splenocytes or purified B cells from C57BL/6 mice for CD40-activation, optimal conditions were identified as follows: Starting from splenocytes of C57BL/6 mice (haplotype H-2b) lymphocytes are purified by density gradient centrifugation and co-cultured with HeLa cells expressing recombinant murine CD40 ligand (tmuCD40L HeLa)11. Cells are recultured every 3-4 days and key components such as CD40L, interleukin-4, -Mercaptoethanol and cyclosporin A are replenished. In this protocol we demonstrate how to obtain fully activated murine CD40-B cells (mCD40B) with similar APC-phenotype to human CD40-B cells (Fig 1a,b). CD40-stimulation leads to a rapid outgrowth and expansion of highly pure (&gt;90%) CD19+ B cells within 14 days of cell culture (Fig 1c,d). To avoid contamination with non-transfected cells, expression of the murine CD40 ligand on the transfectants has to be controlled regularly (Fig 2). Murine CD40-activated B cells can be used to study B-cell activation and differentiation as well as to investigate their potential to function as APC in vitro and in vivo. Moreover, they represent a promising tool for establishing therapeutic or preventive vaccination against tumors and will help to answer questions regarding safety and immunogenicity of this approach12.

In this video, we demonstrate the procedure of CD40-activation and expansion of murine B cells from splenocytes of C57BL/6 mice, which can be used as a model antigen-presenting cell (APC) to study induction of immunity.

Research on B cells has shown that CD40 activation improves their antigen presentation capacity. When stimulated with interleukin-4 and CD40 ligand ...

Murine Model of Allergen Induced Asthma

Asthma is a major cause of morbidity and mortality, affecting some 300 million people throughout the world.1 More than 8% of the US population has asthma, with the prevalence increasing.2 As with other diseases, animal models of allergic airway disease greatly facilitate understanding of the underlying pathophysiology, help identify potential therapeutic targets, and allow preclinical testing of possible new therapies. Models of allergic airway disease have been developed in several animal species, but murine models are particularly attractive due to the low cost, ready availability, and well-characterized immune systems of these animals.3 Availability of a variety of transgenic strains further increases the attractiveness of these models.4 Here we describe two murine models of allergic airway disease, both employing ovalbumin as the antigen. Following initial sensitization by intraperitoneal injection, one model delivers the antigen challenge by nebulization, the other by intratracheal delivery. These two models offer complementary advantages, with each mimicking the major features of human asthma.5
The major features of acute asthma include an exaggerated airway response to stimuli such as methacholine (airway hyperresponsiveness; AHR) and eosinophil-rich airway inflammation. These are also prominent effects of allergen challenge in our murine models,5,6 and we describe techniques for measuring them and thus evaluating the effects of experimental manipulation. Specifically, we describe both invasive7 and non-invasive8 techniques for measuring airway hyperresponsiveness as well as methods for assessing infiltration of inflammatory cells into the airways and the lung. Airway inflammatory cells are collected by bronchoalveolar lavage while lung histopathology is used to assess markers of inflammation throughout the organ. These techniques provide powerful tools for studying asthma in ways that would not be possible in humans.

Experimental mouse models of allergic asthma offer new possibilities for studying disease pathogenesis and developing new therapeutics. These models are well suited to measuring factors governing the allergic immune response, airway inflammation, and pulmonary pathophysiology.

Asthma is a major cause of morbidity and mortality, affecting some 300 million people throughout the world.1 More than 8% of the US population has asthma, ...

Measurement of Tactile Allodynia in a Murine Model of Bacterial Prostatitis

Uropathogenic Escherichia coli (UPEC) are pathogens that play an important role in urinary tract infections and bacterial prostatitis1. We have recently shown that UPEC have an important role in the initiation of chronic pelvic pain2, a feature of Chronic prostatitis/Chronic pelvic pain syndrome (CP/CPPS)3,4. Infection of the prostate by clinically relevant UPEC can initiate and establish chronic pain through mechanisms that may involve tissue damage and the initiation of mechanisms of autoimmunity5.
A challenge to understanding the pathogenesis of UPEC in the prostate is the relative inaccessibility of the prostate gland to manipulation. We utilized a previously described intraurethral infection method6 to deliver a clinical strain of UPEC into male mice thereby establishing an ascending infection of the prostate. Here, we describe our protocols for standardizing the bacterial inoculum7 as well as the procedure for catheterizing anesthetized male mice for instillation of bacteria.
CP/CPPS is primarily characterized by the presence of tactile allodynia4. Behavior testing was based on the concept of cutaneous hyperalgesia resulting from referred visceral pain8-10. An irritable focus in visceral tissues reduces cutaneous pain thresholds allowing for an exaggerated response to normally non-painful stimuli (allodynia). Application of normal force to the skin result in abnormal responses that tend to increase with the intensity of the underlying visceral pain. We describe methodology in NOD/ShiLtJ mice that utilize von Frey fibers to quantify tactile allodynia over time in response to a single infection with UPEC bacteria.

Infection of the prostate may be a contributing factor in mediating pelvic pain in chronic prostatitis. We describe the procedure for preparation of standardized bacterial inoculum, instillation of bacteria into the urethra of male mice and methodology for measuring tactile allodynia in mice over time.

Uropathogenic Escherichia coli (UPEC) are pathogens that play an important role in urinary tract infections and bacterial prostatitis1. We have recently ...

Organoids as Model for Infectious Diseases: Culture of Human and Murine Stomach Organoids and Microinjection of Helicobacter Pylori

Recently infection biologists have employed stem cell derived cultures to answer the need for new and better models to study host-pathogen interactions. Three cellular sources have been used: Embryonic stem cells (ESC), induced pluripotent stem cells (iPSC) or adult stem cells. Here, culture of mouse and human gastric organoids derived from adult stem cells is described and used for infection with the gastric pathogen Helicobacter pylori. Human gastric glands are isolated from resection material, seeded in a basement matrix and embedded in medium containing growth factors epidermal growth factor (EGF), R-spondin, Noggin, Wnt, fibroblast growth factor (FGF) 10, gastrin and transforming growth factor (TGF) beta inhibitor. In these conditions, gastric glands grow into 3-dimensional organoids containing 4 lineages of the stomach. The organoids expand indefinitely and can be frozen and thawed similarly as cell lines. For infection studies, bacteria are microinjected into the lumen of the organoids. Infected organoids are processed for imaging. The described methods can be adapted to other organoids and infections with other bacteria, viruses or parasites. This allows the study of infection-induced changes in primary cells.

Stem cell derived cultures harbor tremendous potential to model infectious diseases. Here, the culture of mouse and human gastric organoids derived from adult stem cells is described. The organoids are microinjected with the gastric pathogen Helicobacter pylori.

Recently infection biologists have employed stem cell derived cultures to answer the need for new and better models to study host-pathogen interactions. ...

Femur Window Chamber Model for In Vivo Cell Tracking in the Murine Bone Marrow

Bone marrow is a complex organ that contains various hematopoietic and non-hematopoietic cells. These cells are involved in many biological processes, including hematopoiesis, immune regulation and tumor regulation. Commonly used methods for understanding cellular actions in the bone marrow, such as histology and blood counts, provide static information rather than capturing the dynamic action of multiple cellular components in vivo. To complement the standard methods, a window chamber (WC)-based model was developed to enable serial in vivo imaging of cells and structures in the murine bone marrow. This protocol describes a surgical procedure for installing the WC in the femur, in order to facilitate long-term optical access to the femoral bone marrow. In particular, to demonstrate its experimental utility, this WC approach was used to image and track neutrophils within the vascular network of the femur, thereby providing a novel method to visualize and quantify immune cell trafficking and regulation in the bone marrow. This method can be applied to study various biological processes in the murine bone marrow, such as hematopoiesis, stem cell transplantation, and immune responses in pathological conditions, including cancer.

Femur Window Chamber Model for In Vivo Cell Tracking in the Murine Bone Marrow

The protocol describes a novel murine femur window chamber model that can be used to track movement of cells in the femoral bone marrow in vivo. Intravital multiphoton fluorescence microscopy is used to image three components of the femoral bone marrow (vasculature, collagen matrix, and neutrophils) over time.

Bone marrow is a complex organ that contains various hematopoietic and non-hematopoietic cells. These cells are involved in many biological processes, ...

A Murine Model of Group B Streptococcus Vaginal Colonization

Streptococcus agalactiae (group B Streptococcus, GBS), is a Gram-positive, asymptomatic colonizer of the human gastrointestinal tract and vaginal tract of 10 - 30% of adults. In immune-compromised individuals, including neonates, pregnant women, and the elderly, GBS may switch to an invasive pathogen causing sepsis, arthritis, pneumonia, and meningitis. Because GBS is a leading bacterial pathogen of neonates, current prophylaxis is comprised of late gestation screening for GBS vaginal colonization and subsequent peripartum antibiotic treatment of GBS-positive mothers. Heavy GBS vaginal burden is a risk factor for both neonatal disease and colonization. Unfortunately, little is known about the host and bacterial factors that promote or permit GBS vaginal colonization. This protocol describes a technique for establishing persistent GBS vaginal colonization using a single &#946;-estradiol pre-treatment and daily sampling to determine bacterial load. It further details methods to administer additional therapies or reagents of interest and to collect vaginal lavage fluid and reproductive tract tissues. This mouse model will further the understanding of the GBS-host interaction within the vaginal environment, which will lead to potential therapeutic targets to control maternal vaginal colonization during pregnancy and to prevent transmission to the vulnerable newborn. It will also be of interest to increase our understanding of general bacterial-host interactions in the female vaginal tract.

A Murine Model of Group B Streptococcus Vaginal Colonization

The purpose of this protocol is to imitate human group B Streptococcus (GBS) vaginal colonization in a murine model. This method may be used to investigate host immune responses and bacterial factors contributing to GBS vaginal persistence, as well as to test therapeutic strategies.

Streptococcus agalactiae (group B Streptococcus, GBS), is a Gram-positive, asymptomatic colonizer of the human gastrointestinal tract and vaginal tract of ...

Stenosis of the Inferior Vena Cava: A Murine Model of Deep Vein Thrombosis

Deep vein thrombosis (DVT) and its devastating complication, pulmonary embolism, are a severe health problem with high mortality. Mechanisms of thrombus formation in veins remain obscure. Lack of mobility (e.g., after surgery or long-haul flights) is one of the main factors leading to DVT. The pathophysiological consequence of the lack of mobility is blood flow stagnation in venous valves. Here, a model is described that mimics such flow disturbance as a thrombosis-driving factor. In this model, partial flow restriction (stenosis) in the inferior vena cava (IVC) is created. Closure of about 90% of the IVC lumen for 48 h results in development of thrombi structurally similar to those in humans. The similarities are: i) most of the thrombus volume is red, i.e., consists of red blood cells and fibrin, ii) presence of a white part (lines of Zahn), iii) non-denuded endothelial monolayer, iv) elevated plasma D-Dimer levels, and v) possibility to prevent thrombosis by low molecular weight heparin. Limitations include variable size of thrombi and the fact that a certain percentage of wild-type mice (0 - 35%) may not produce a thrombus. In addition to visual observation and measurement, thrombi may be visualized by non-invasive technologies, such as ultrasonography, which allows for monitoring the dynamics of thrombus development. At shorter time points (1 - 6 h), intravital microscopy may be applied to directly observe events (e.g., recruitment of cells to the vessel wall) preceding thrombus formation. Use of this method by several teams around the world has made it possible to uncover basic mechanisms of DVT initiation and identify potential targets that might be beneficial for its prevention.

We describe here stenosis in the inferior vena cava as a murine model of deep vein thrombosis. This model recapitulates blood flow restriction, one of the major triggers of venous thrombosis in humans.

Deep vein thrombosis (DVT) and its devastating complication, pulmonary embolism, are a severe health problem with high mortality. Mechanisms of thrombus ...

In Vivo Imaging of Reactive Oxygen Species in a Murine Wound Model

The generation of reactive oxygen species (ROS) is a hallmark of inflammatory processes, but in excess, oxidative stress is widely implicated in various pathologies such as cancer, atherosclerosis and diabetes. We have previously shown that dysfunction of the Nuclear factor (erythroid-derived 2)-like 2 (Nrf2)/ Kelch-like erythroid cell-derived protein 1 (Keap1) signaling pathway leads to extreme ROS imbalance during cutaneous wound healing in diabetes. Since ROS levels are an important indicator of progression of wound healing, specific and accurate quantification techniques are valuable. Several in vitro assays to measure ROS in cells and tissues have been described; however, they only provide a single cumulative measurement per sample. More recently, the development of protein-based indicators and imaging modalities have allowed for unique spatiotemporal analyses. L-012 (C13H8ClN4NaO2) is a luminol derivative that can be used for both in vivo and in vitro chemiluminescent detection of ROS generated by NAPDH oxidase. L-012 emits a stronger signal than other fluorescent probes and has been shown to be both sensitive and reliable for detecting ROS. The time lapse applicability of L-012-facilitated imaging provides valuable information about inflammatory processes while reducing the need for sacrifice and overall reducing the number of study animals. Here, we describe a protocol utilizing L-012-facilitated in vivo imaging to quantify oxidative stress in a model of excisional wound healing using diabetic mice with locally dysfunctional Nrf2/Keap1.

In Vivo Imaging of Reactive Oxygen Species in a Murine Wound Model

We describe a non-invasive in vivo imaging protocol that is streamlined and cost-effective, utilizing L-012, a chemiluminescent luminol-analog, to visualize and quantify reactive oxygen species (ROS) generated in a mouse excisional wound model.

The generation of reactive oxygen species (ROS) is a hallmark of inflammatory processes, but in excess, oxidative stress is widely implicated in various ...

A Murine Model of Dengue Virus-induced Acute Viral Encephalitis-like Disease

Dengue virus (DENV), an arthropod-borne virus transmitted by mosquitoes, may cause the severe disease known as dengue hemorrhagic fever, which is characterized by lethal complications due to plasma leakage, ascites, pleural effusion, respiratory distress, severe bleeding, and organ impairment. A few cases of DENV infection present neurological manifestations; however, studies have not explored DENV-induced neuropathogenesis further. In this study, we present a protocol to use an immunocompetent outbred ICR (Institute of Cancer Research) mouse for investigating the induction of central nervous system (CNS) infection with DENV, followed by the progression of acute viral encephalitis-like disease.

Here, we present a protocol for creating an immunocompetent ICR (Institute of Cancer Research) murine model of central nervous system infection to display the development of neuropathy. Monitoring acute viral encephalitic disorders by identical disease scores could be performed for showing dengue virus-induced neuropathy in vivo.

Dengue virus (DENV), an arthropod-borne virus transmitted by mosquitoes, may cause the severe disease known as dengue hemorrhagic fever, which is ...

An Advanced Murine Model for Nonalcoholic Steatohepatitis in Association with Type 2 Diabetes

Obesity is associated with chronic low-grade inflammation and insulin resistance, contributing to an increasing prevalence of chronic metabolic diseases, such as type 2 diabetes and nonalcoholic steatohepatitis (NASH). Recent research has established that pro-inflammatory immune cells infiltrate obese hypertrophic adipose tissue and liver. Given the emerging importance of immune cells in the context of metabolic homeostasis, there is a critical need to quantify and characterize their modification during the development of type 2 diabetes and NASH. However, animal models that induce pathophysiological features typical of human NASH are sparse.
In this article, we provide a detailed protocol to identify immune cell subsets isolated from liver and adipose tissue in a reliable mouse model of NASH, established by housing high-fat diet (HFD) mice under non-specific pathogen-free (SPF) conditions without a barrier for at least seven weeks. We demonstrate the handling of mice in non-SPF conditions, digestion of the tissues and identification of macrophages, natural killer (NK) cells, dendritic cells, B and T cell subsets by flow cytometry. Representative flow cytometry plots from SPF HFD mice and non-SPF mice are provided. To obtain reliable and interpretable data, the use of antibodies, accurate and precise methods for tissue digestion and proper gating in flow cytometry experiments are critical elements.
The intervention to restore physiological antigen exposure in mice by housing them in non-SPF conditions and unspecific exposure to microbial antigens could provide a relevant tool for investigating the link between immunological alterations, diet-induced obesity and related long term complications.

A simple and reliable diet-induced rodent animal model for nonalcoholic steatohepatitis (NASH) is described, achieved through non-SPF housing of the animals and administration of a specific high-fat diet. We describe identification of hepatic and adipose immune cell subsets to recapitulate human immunological conditions by exposing mice to environmental germs.

Obesity is associated with chronic low-grade inflammation and insulin resistance, contributing to an increasing prevalence of chronic metabolic diseases, ...

Postoperative Ileus Murine Model

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Chapters in this video

Murine Model of CD40-activation of B cells

Murine Model of Allergen Induced Asthma

Measurement of Tactile Allodynia in a Murine Model of Bacterial Prostatitis

Organoids as Model for Infectious Diseases: Culture of Human and Murine Stomach Organoids and Microinjection of Helicobacter Pylori

Femur Window Chamber Model for In Vivo Cell Tracking in the Murine Bone Marrow

A Murine Model of Group B Streptococcus Vaginal Colonization

Stenosis of the Inferior Vena Cava: A Murine Model of Deep Vein Thrombosis

In Vivo Imaging of Reactive Oxygen Species in a Murine Wound Model

A Murine Model of Dengue Virus-induced Acute Viral Encephalitis-like Disease

An Advanced Murine Model for Nonalcoholic Steatohepatitis in Association with Type 2 Diabetes