This study was supported by the University Synergy Innovation Program of Anhui Province (GXXT-2021-077) and the Anhui University of Science and Technology Graduate Innovation Fund (2021CX2120).

All procedures followed the guidelines of the National Institutes of Health&#39;s Guide for the Care and Use of Laboratory Animals (NIH Publication No. 8023, revised 1978) and were approved by the Institutional Animal Care and Use Committee at the Medical School of Anhui University of Science and Technology.
1. Managing and feeding mice
<ol>
	<li>Assign 20 healthy C57BL/6 male mice to the experimental or vehicle groups in a 1:1 ratio. Acclimatize the mice to the new environment for 1 week.</li>
	<li>Provide a constant light time of 12 h per day. Use a time control switch for precise timing.</li>
</ol>
2. Preparing the CSD suspension
<ol>
	<li>At least 1 day before nasal drips, grind the silica in an agate mortar for 0.5 h.</li>
	<li>Observe the size and shape of the crystal particles. Take representative photographs using scanning electron microscopy (SEM).
	<ol>
		<li>Use conductive tape to bind the particles to prepare the sample for SEM. Use a hair dryer to gently blow away the silicon particles that are not firmly bonded.</li>
		<li>Evacuate the sample chamber, turn on the high pressure, and capture the image. 
		NOTE: The working distance (WD) between the lens and the sample is 5.9 mm, the accelerating voltage is 2.0 kV, and the magnification (Mag) is 100,000x, using the SignalA detector. The particles are dispersed with irregular crystallization, and approximately 80% have a diameter of 1-5 &#181;m (Figure 1A).</li>
	</ol>
	</li>
	<li>Make a 20 mg/mL sterile CSD suspension. Dilute the CSD using sterile saline and mix it with an ultrasonic shaker (40 kHz, 80 W) at room temperature (RT) for 30 min.</li>
	<li>Stir and mix the CSD suspension thoroughly on a vortex mixer for 10 s before administering the nasal drips.</li>
</ol>
3. Administering nasal drips to mouse
<ol>
	<li>Rapidly anesthetize a mouse with 2% isoflurane at a dose of 3.6 mL/h in an anesthesia machine (Figure 1B, left panel). 
	NOTE: Anesthesia should be performed in a fume hood to avoid inhalation of the anesthetic by the technician. Ensure that the adequate depth of anesthesia by observing the change from rapid and irregular breathing to a slow and steady state in the mice.</li>
	<li>Drip 50 &#181;L of the CSD nasally within 4-8 s (Figure 1B, right).
	<ol>
		<li>For the nasal drip, place the head of the mouse on the researcher&#39;s metacarpophalangeal joint with the tip of the index finger.</li>
		<li>Keep the mouse in a prone position, with four fingers slightly flexed and the tip of the thumb lightly touching the lower lip of the mouse to straighten the airway. Avoid touching the pharynx to elicit the gag reflex.</li>
		<li>Aspirate 50 &#956;l of liquid using a 200 &#956;l pipette. Drop the liquid into the mouse&#39;s nasal cavity in three to four divided doses depending on the mouse&#39;s respiratory rate. Each instillation should consist of 15 to 20 &#956;l liquid. Administer the drips once every 3 days, 5x within 12 days. Treat the control mouse with an equal amount of saline.</li>
	</ol>
	</li>
	<li>Gently massage the heart area of the mouse 5x-10x for 5 s.
	<ol>
		<li>Hold the mouse&#39;s body with the palm, pinch the skin on the back of the neck with the thumb and index finger, and fix the mouse&#39;s hind limbs with the other fingers.&#160;Then, gently press the mouse&#39;s heart-beating area with the index finger of the other hand 5-10 times in a period of 5 s.</li>
	</ol>
	</li>
	<li>When the mouse&#39;s breathing has stabilized, place it in a recovery cage with a heating pad and observe until it recovers from anesthesia, then return the mouse to its home cage. Sacrifice the mouse at 31 days.</li>
</ol>
4. Collecting the lung tissues and preparing a paraffin section
<ol>
	<li>Inject 0.18 mL of 10% chloral hydrate intraperitoneally, ensuring that the mice do not respond to toe or tail stimulation (performed using the toothed forceps). Then, proceed to the next step.</li>
	<li>Fix the mouse limbs on a foam test board and spray with 75% alcohol to dampen the fur. Remove most of the thoracic ribs at the midline of the clavicle and open the thoracic cavity of the mice to expose the heart and lungs.</li>
	<li>Immediately cut open the right atrium with ophthalmic surgical scissors and slowly inject 20 mL of phosphate buffer (PBS) from the heart tip at the left atrial beat with the greatest amplitude to allow the whole blood to flow. Next, remove the lower lobe of the right lung and store it at -80 &#176;C for western blotting analysis.</li>
	<li>Keep perfusing 10 mL of 4% paraformaldehyde (PFA) in the same site after the PBS injection. Collect the remaining lung and preserve the sample in 30 mL of 4% PFA for pathological analysis.</li>
	<li>After 72 h of fixation, embed specimens in paraffin.
	<ol>
		<li>Dehydrate the tissues through a graded series of ethanol (EtOH) dilutions in deionized water (60%, 70%, 80%, 90%, 100%) for 1 h each at RT. Clear the sample in two washes of xylene for 1 h each.</li>
		<li>Infiltrate the samples with the melted paraffin wax by heating to 50 &#176;C for 2 h. Repeat this process in another cylinder. Cool the wax molds with tissues for 1 h to harden.</li>
		<li>After the wax has hardened and the tissue has been embedded, use a paraffin sectioning machine to slice the tissue at 5 &#181;m. The precise sectioning and slide mounting steps were previously described11. 
		&#8203;NOTE: Sufficient tissue perfusion is indicated by developing muscle twitching and tail twisting into an &#34;S&#34; shape or flexion after receiving 10 mL of 4% PFA.</li>
	</ol>
	</li>
</ol>
5. Performing hematoxylin and eosin (HE) staining
<ol>
	<li>Heat the paraffin-embedded tissues on a hot plate (60 &#176;C) for more than 4 h to allow for adhesion to the slides and improved deparaffination.</li>
	<li>Dewax and hydrate paraffin sections. Soak the slides with samples in xylene 2x for 30 min each time. Next, dip them in anhydrous ethanol, then 95%, 85%, 75% alcohol, and deionized water for 5 min, respectively.</li>
	<li>Perform hematoxylin and eosin staining. Stain the tissues in a hematoxylin staining bucket for 10 min. Rinse them with gently running water for 5 min. Then, dip the slides in the eosin staining bucket for 10 s.</li>
	<li>Dehydrate the samples in 75%, 85%, 95%, and anhydrous ethanol for 5 min each. Clear the tissue sections by immersing them in xylene for 5 min. Seal the section with approximately 60 &#181;L of neutral resin drops. Place the cover slide over the section and carefully lower it to avoid air bubbles.</li>
</ol>
6. Performing Masson staining
<ol>
	<li>Dewax and hydrate the paraffin samples, as mentioned in step 5.2. Then, stain the cell nuclei with 50% Weigert&#39;s hematoxylin for 10 min. Soak the tissue in acidic ethanol liquefaction for 10 s and rinse the tissue slides gently with running water for nuclei bluing. 
	NOTE: Prepare the Weigert&#39;s hematoxylin staining solution immediately before use.</li>
	<li>Stain the samples with drops of Lichun red staining solution (40 &#181;L for each tissue slide) for 7 min and wash them with a weak acid working solution (30% hydrochloric acid) for 1 min to remove the unbound Lichun red dye.</li>
	<li>Dip them in 95% alcohol for 20 s and dehydrate them 2x with anhydrous ethanol for 1-3 s each. After that, clear the tissues with xylene and seal them with 60 &#181;L of neutral resin drops, as mentioned in step 5.4.</li>
</ol>
7. Performing Sirius red staining
<ol>
	<li>Dewax and hydrate paraffin sections as mentioned in step 5.2.</li>
	<li>Infiltrate the sections for 1 h with Sirius red staining solution.</li>
	<li>Stain the cell nuclei of the samples for 8-10 min with Mayer hematoxylin staining solution. Rinse them gently with running water for 10 min. Then, dehydrate and clear the tissue slides. Seal them as mentioned in step 5.4.</li>
</ol>
8. Performing immunohistochemistry
<ol>
	<li>Dewax and hydrate the paraffin samples as described in step 5.2.</li>
	<li>Infiltrate the specimens with a 3 mg/mL EDTA antigen retrieval solution of about 30 mL. Boil for 20-30 min. Wash the tissues with deionized water, and then incubate them in phosphate-buffered solution containing 0.5% Tween-20 (PBST) for 5 min.</li>
	<li>Soak the samples for 15 min with 0.3% hydrogen peroxide solution to inactivate the endogenous peroxidase in the specimens. Wash them 3x with PBST for 5 min each time. 
	NOTE: The 0.3% hydrogen peroxide solution must be made fresh in a light-proof environment.</li>
	<li>Permeabilize the membrane of specimens for 15 min with 0.3% Triton-100 solution. Then, block with 30-40 &#181;L of 5% bovine serum albumin (BSA) for 1 h.</li>
	<li>Remove the blocking solution. Add diluted primary antibodies NF-&#954;B (dilution 1:200) and CD68 (dilution 1:1,000) and incubate the specimens overnight at 2-8 &#176;C in a microscope slide IHC wet box to prevent evaporation and light.</li>
	<li>The next day, transfer them to RT for 1 h. Then, wash them with PBST 3x for 5 min each.</li>
	<li>Incubate the samples for 1 h in rabbit anti-mouse horseradish peroxidase-labeled secondary antibodies (dilution ratio 1:500) at RT and wash them with PBST 3x for 5 min each. 
	NOTE: Both primary and secondary antibodies were diluted with 5% BSA.</li>
	<li>Incubate samples with the 3,3&#39;-Diaminobenzidine (DAB) substrate corresponding to the enzyme-labeled antibody for 5-20 min. Stop the reaction with deionized water when the optimal staining intensity is reached. 
	NOTE: The DAB solution needs to be prepared freshly and protected from light. The color development reaction should be observed in real-time under the microscope to determine when to stop staining. Positive specimens exhibit intense staining, while negative specimens do not develop color.</li>
	<li>Counterstain the samples for 30 s with Weigert hematoxylin. Next, rinse the tissues under running water for 1 min. Then, dehydrate, clear, and seal the tissue slides, as mentioned in step 5.4.</li>
</ol>
9. Performing western blotting analysis
<ol>
	<li>Lyse the lung tissues to extract proteins. Add 200 &#181;L of RIPA working solution to 20 mg of the lung tissue.</li>
	<li>Homogenize the tissue on ice for 5 min using a handheld electric grinder and incubate for 1 h on ice with gentle shaking. Afterward, centrifuge the homogenate at 4 &#176;C for 15 min at 14,800 x g.</li>
	<li>Collect the supernatant and determine the protein concentration with the BCA protein assay kit. Make the protein storage solution with RIPA at 6 &#181;g/&#181;L protein. Add 20 &#181;L of 5x loading buffer to the 80 &#181;L of the protein lysate. Disrupt the secondary protein structure by heating the protein-containing microcentrifuge tubes in a metal bath (100 &#176;C) for 20 min.</li>
	<li>After cooling, aliquot 100 &#181;L of the protein storage solution into each tube and store the samples in a -80 &#176;C refrigerator. Dilute the protein concentration to 2&#8211;3 &#956;g/&#956;L with 1x loading buffer before electrophoresis. 
	NOTE: The protein extraction process should be performed on ice. For the RIPA working solution, add 1 &#181;L of 100 mM phenylmethylsulfonyl fluoride (PMSF) to 99 &#181;L of RIPA to inhibit phosphorylated protein degradation.&#160; Prepare 1x loading buffer by diluting 5x loading buffer with RIPA at a ratio of 1:4.&#160;</li>
	<li>Add 20 &#181;g of samples to each well and run the gel. For 5% concentrated gels, use 80 V for 20 min to have the proteins electrophoresed down from the same starting point. For 10% isolated gels, run at 100 V for 1 h to allow the proteins of different molecular weights to be separated as much as possible.</li>
	<li>Pre-activate the PVDF membrane with methanol for 20 s. Transfer the proteins to the PVDF membrane using the wet transfer method with 400 mA current for 1-2 h. 
	NOTE: Ensure that the electrophoresis tank and the electrotransfer tank are horizontal. Cool the entire tank with ice, as the membrane transfer process generates much heat.</li>
	<li>Wash the membrane with TBST solution for 5 min each time, 5x. Then, block with 5% BSA or 5% skim milk for 1 h. Dilute the primary antibodies NF-&#954;B (1:1,000) and &#946;-actin (1:1,000) with 5% BSA. Submerge the strips in the antibody solution and shake the strips gently overnight at 2-8 &#176;C.</li>
	<li>Wash the strips with PBST. Next, dilute horseradish peroxidase (HRP)-conjugated goat anti-rabbit secondary antibody (1:10,000) and incubate them in the diluted secondary antibody for 1 h at RT with gentle shaking.</li>
	<li>Prepare an enhanced chemiluminescence (ECL) developer, drop it on the strip, and incubate for 3 min.</li>
	<li>Expose the strip to a gel imager for 20 s. Measure the gray value of the strip to assess the protein level by system software. Use &#946;-actin as an internal control.</li>
</ol>

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Morbidity and Mortality Weekly Report. 67 (30), 819-824 (2018).</li><li>Voelker, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Black+Lung+resurgence+raises+new+challenges+for+coal+country+physicians.">Black Lung resurgence raises new challenges for coal country physicians.</a> JAMA. 321 (1), 17-19 (2019).</li><li>Nakashima, K., 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=Regulatory+role+of+heme+oxygenase-1+in+silica-induced+lung+injury.">Regulatory role of heme oxygenase-1 in silica-induced lung injury.</a> Respiratory Research. 19 (1), 144 (2018).</li><li>Li, Y., 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=Minute+cellular+nodules+as+early+lesions+in+rats+with+silica+exposure+via+inhalation.">Minute cellular nodules as early lesions in rats with silica exposure via inhalation.</a> Veterinary Sciences. 9 (6), 251 (2022).</li><li>Salehi, F., 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=Immunological+responses+in+C3H/HeJ+mice+following+nose-only+inhalation+exposure+to+different+sizes+of+beryllium+metal+particles.">Immunological responses in C3H/HeJ mice following nose-only inhalation exposure to different sizes of beryllium metal particles.</a> Journal of Applied Toxicology. 29 (1), 61-68 (2009).</li><li>Yang, T., 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=Emodin+suppresses+silica-induced+lung+fibrosis+by+promoting+Sirt1+signaling+via+direct+contact.">Emodin suppresses silica-induced lung fibrosis by promoting Sirt1 signaling via direct contact.</a> Molecular Medicine Reports. 14 (5), 4643-4649 (2016).</li><li>Cornell, W. C., 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=Paraffin+embedding+and+thin+sectioning+of+microbial+colony+biofilms+for+microscopic+analysis.">Paraffin embedding and thin sectioning of microbial colony biofilms for microscopic analysis.</a> Journal of Visualized Experiments. (133), e57196 (2018).</li><li>Li, B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+suitable+silicosis+mouse+model+was+constructed+by+repeated+inhalation+of+silica+dust+via+nose.">A suitable silicosis mouse model was constructed by repeated inhalation of silica dust via nose.</a> Toxicology Letters. 353, 1-12 (2021).</li><li>Mu, M., 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=Coal+dust+exposure+triggers+heterogeneity+of+transcriptional+profiles+in+mouse+pneumoconiosis+and+Vitamin+D+remedies.">Coal dust exposure triggers heterogeneity of transcriptional profiles in mouse pneumoconiosis and Vitamin D remedies.</a> Particle and Fibre Toxicology. 19 (1), 7 (2022).</li><li>Kato, K., 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=Muc1+deficiency+exacerbates+pulmonary+fibrosis+in+a+mouse+model+of+silicosis.">Muc1 deficiency exacerbates pulmonary fibrosis in a mouse model of silicosis.</a> Biochemical and Biophysical Research Communications. 493 (3), 1230-1235 (2017).</li><li>Liu, F., 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++regulatory+T+cells+depletion+may+attenuate+the+development+of+silica-induced+lung+fibrosis+in+mice.">CD4+CD25+Foxp3+ regulatory T cells depletion may attenuate the development of silica-induced lung fibrosis in mice.</a> PLoS One. 5 (11), 15404 (2010).</li><li>Mansouri, N., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mesenchymal+stromal+cell+exosomes+prevent+and+revert+experimental+pulmonary+fibrosis+through+modulation+of+monocyte+phenotypes.">Mesenchymal stromal cell exosomes prevent and revert experimental pulmonary fibrosis through modulation of monocyte phenotypes.</a> JCI Insight. 4 (21), 128060 (2019).</li><li>Ohtsuka, Y., Wang, X. T., Saito, J., Ishida, T., Munakata, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+linkage+analysis+of+pulmonary+fibrotic+response+to+silica+in+mice.">Genetic linkage analysis of pulmonary fibrotic response to silica in mice.</a> The European Respiratory Journal. 28 (5), 1013-1019 (2006).</li></ol>

The authors declare no conflicts of interest.

Silicosis mouse models are crucial for studying the pathogenesis and treatment of silicosis. This protocol describes a method for preparing a model of silicosis in mice through repeated nasal exposure. This method allows for the study of the pathological characteristics of silicosis induced by different exposure times. Mice were anesthetized on a ventilator, and their respiratory rate was monitored. The initial short, fast breathing rate gradually slowed and deepened over time. The anesthesia caused the mice's muscles to...

Workers are inevitably exposed to irregular crystalline silica dust (CSD), which can be inhaled and is more toxic in numerous occupational contexts, including mining, pottery, glass, quartz processing, and concrete1,2. A chronic dust inhalation condition known as silicosis causes progressive lung fibrosis3. According to epidemiological data, the incidence of silicosis has been declining globally over the past few decades, but in recent years, it has been increasing and affecting younger people4,5,6. The underlying mechanism of silicosis presents a significant challenge for scientific research due to its insidious onset and protracted incubation period. It is still unknown how silicosis develops. Furthermore, no current medications can stop the progression of silicosis and reverse pulmonary fibrosis.
The current mouse models for silicosis involve tracheal ingestion of a mixed suspension of CSD. For example, administering CSD into the lungs by adopting the cervical trachea trauma after anesthesia does not comply with repeated human exposure to dye dust7. The impact of exposure to ambient dust on individuals can be studied by exposing them to CSD in the form of aerosols, which more accurately reflects the environmental concentrations of this toxic substance8. However, environmental CSD cannot simply be inhaled directly into the lungs due to the unique physiological structure of the mouse nose9. Moreover, the equipment associated with this technology is expensive, which has caused researchers to re-evaluate the mouse silicosis model10. By inhaling CSD suspension through a nasal drip five times within 2 weeks, it was possible to build a dynamic model of silicosis. This model is consistent and safe while being easy to use. It is important to note that this study allows for repeated inhalation of CSD in mice. The mouse silicosis model created through this procedure is expected to be more beneficial for research requirements.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>0.5 mL tube</td>
			<td>Biosharp</td>
			<td>BS-05-M</td>
			<td></td>
		</tr>
		<tr>
			<td>10% formalin neutral fixative</td>
			<td>Nanchang Yulu Experimental Equipment Co.</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Adobe Illustrator</td>
			<td>Adobe&#160;</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Alcohol disinfectant</td>
			<td>Xintai Kanyuan Disinfection Products Co.</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>CD68</td>
			<td>Abcam</td>
			<td>ab125212</td>
			<td></td>
		</tr>
		<tr>
			<td>Citrate antigen retrieval solution</td>
			<td>biosharp life science</td>
			<td>BL619A</td>
			<td></td>
		</tr>
		<tr>
			<td>DAB chromogenic kit</td>
			<td>NJJCBio</td>
			<td>W026-1-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Dimethyl benzene</td>
			<td>West Asia Chemical Technology (Shandong) Co</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Enhanced BCA protein assay kit</td>
			<td>Beyotime Biotechnology</td>
			<td>P0009</td>
			<td></td>
		</tr>
		<tr>
			<td>Hematoxylin and Eosin (H&#38;E)</td>
			<td>Beyotime Biotechnology</td>
			<td>C0105S</td>
			<td></td>
		</tr>
		<tr>
			<td>HRP substrate</td>
			<td>Millipore Corporation</td>
			<td>P90720</td>
			<td></td>
		</tr>
		<tr>
			<td>HRP-conjugated Affinipure Goat Anti-Rabbit IgG(H+L)</td>
			<td>Proteintech</td>
			<td>Sa00001-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Iceacetic acid</td>
			<td>West Asia Chemical Technology (Shandong) Co</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>ImageJ</td>
			<td>NIH</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>RWD Life Science</td>
			<td>R510-22</td>
			<td></td>
		</tr>
		<tr>
			<td>Masson&#39;s Trichrome stain kit</td>
			<td>Solarbio</td>
			<td>G1340</td>
			<td></td>
		</tr>
		<tr>
			<td>Methanol</td>
			<td>Macklin</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Microtubes</td>
			<td>Millipore</td>
			<td>AXYMCT150CS</td>
			<td></td>
		</tr>
		<tr>
			<td>NF-&#954;B p65</td>
			<td>Cell Signaling Technology</td>
			<td>8242S</td>
			<td></td>
		</tr>
		<tr>
			<td>Oscillatory thermostatic metal bath</td>
			<td>Abson</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraffin embedding machine</td>
			<td>Precision (Changzhou) Medical Equipment Co.</td>
			<td>PBM-A</td>
			<td></td>
		</tr>
		<tr>
			<td>Paraffin Slicer</td>
			<td>Jinhua Kratai Instruments Co.</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphate buffer (PBS)&#160;</td>
			<td>Biosharp</td>
			<td>BL601A</td>
			<td></td>
		</tr>
		<tr>
			<td>Physiological saline&#160;</td>
			<td>The First People&#39;s Hospital of Huainan City</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>Pipettes</td>
			<td>Eppendorf</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>PMSF</td>
			<td>Beyotime Biotechnological</td>
			<td>ST505</td>
			<td></td>
		</tr>
		<tr>
			<td>Polarized light microscope</td>
			<td>Olympus</td>
			<td>BX51</td>
			<td></td>
		</tr>
		<tr>
			<td>Precision balance</td>
			<td>Acculab</td>
			<td>ALC-110.4</td>
			<td></td>
		</tr>
		<tr>
			<td>Prism7.0</td>
			<td>GraphPad&#160;</td>
			<td>Version 7.0</td>
			<td></td>
		</tr>
		<tr>
			<td>PVDF membranes</td>
			<td>Millipore</td>
			<td>3010040001</td>
			<td></td>
		</tr>
		<tr>
			<td>RIPA lysis buffer</td>
			<td>Beyotime Biotechnology</td>
			<td>P0013B</td>
			<td></td>
		</tr>
		<tr>
			<td>RODI IOT intelligent multifunctional water purification system</td>
			<td>RSJ</td>
			<td>RODI-220BN</td>
			<td></td>
		</tr>
		<tr>
			<td>Scilogex SK-D1807-E 3D Shaker</td>
			<td>Scilogex</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>SDS-PAGE gel preparation kit</td>
			<td>Beyotime Biotechnology</td>
			<td>P0012A</td>
			<td></td>
		</tr>
		<tr>
			<td>Silicon dioxid</td>
			<td>Sigma</td>
			<td>#BCBV6865</td>
			<td></td>
		</tr>
		<tr>
			<td>Sirius red staining</td>
			<td>Nanjing SenBeiJia Biological Technology Co., Ltd.</td>
			<td>181012</td>
			<td></td>
		</tr>
		<tr>
			<td>Small animal anesthesia machine</td>
			<td>Anhui Yaokun Biotech Co., Ltd.</td>
			<td>ZL-04A</td>
			<td></td>
		</tr>
		<tr>
			<td>Universal Pipette Tips (0.1&#8211;10 &#181;L)</td>
			<td>KIRGEN</td>
			<td>KG1011</td>
			<td></td>
		</tr>
		<tr>
			<td>Universal Pipette Tips (100&#8211;1000 &#181;L)</td>
			<td>KIRGEN</td>
			<td>KG1313</td>
			<td></td>
		</tr>
		<tr>
			<td>Universal Pipette Tips (1&#8211;200 &#181;L)</td>
			<td>KIRGEN</td>
			<td>KG1212</td>
			<td></td>
		</tr>
		<tr>
			<td>Vortex&#160;mixer&#160;</td>
			<td>VWR</td>
			<td>NA</td>
			<td></td>
		</tr>
		<tr>
			<td>ZEISS GeminiSEM 500</td>
			<td>Zeiss Germany</td>
			<td>SEM 500</td>
			<td></td>
		</tr>
		<tr>
			<td>&#946;-actin</td>
			<td>Bioss</td>
			<td>bs-0061R</td>
			<td></td>
		</tr>
	</tbody>
</table>

a silicosis mouse model established by repeated inhalation of crystalline silica dust

Silicosis can be caused by exposure to respiratory crystalline silica dust (CSD) in an industrial environment. The pathophysiology, screening, and treatment of silicosis in humans have all been extensively studied using the mouse silicosis model. By repeatedly making mice inhale CSD into their lungs, the mice can mimic the clinical symptoms of human silicosis. This methodology is practical and efficient in terms of time and output and does not cause mechanical injury to the upper respiratory tract due to surgery. Furthermore, this model can successfully mimic acute/chronic transformation process of silicosis. The main procedures were as follows. The sterilized 1-5 &#181;m CSD powder was fully ground, suspended in saline, and dispersed in an ultrasonic water bath for 30 min. Mice under isoflurane-induced anesthesia switched from shallow rapid breathing to deep, slow aspiration for approximately 2 s. The mouse was placed in the palm of a hand, and the thumb tip gently touched the lip edge of the mouse's jaw to straighten the airway. After each exhalation, the mice breathed in the silica suspension drop by drop through one nostril, completing the process within 4-8 s. After the mice's breathing had stabilized, their chest was stroked and caressed to prevent the inhaled CSD from being coughed up. The mice were then returned to the cage. In conclusion, this model can quantify CSD along the typical physiological passage of tiny particles into the lung, from the upper respiratory tract to the terminal bronchioles and alveoli. It can also replicate the recurrent exposure of employees due to work. The model can be performed by one person and does not need expensive equipment. It conveniently and effectively simulates the disease features of human silicosis with high repeatability.

The potential pathogenesis of silicosis in mice was investigated using the proposed method. We found that the body weight of the mice in the experimental group decreased significantly relative to the control group and that the body weight recovered slowly after cessation of exposure. Due to the optimized dose used here, no mortality was observed in silica-exposed mice in this experiment. The technical roadmap of repeated nasal drip to CSD is shown in (Figure 1). The previously described proc...

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A Silicosis Mouse Model Established by Repeated Inhalation of Crystalline Silica Dust

This protocol describes a method for establishing a mouse model of silicosis through repeated exposure to silica suspensions via a nasal drip. This model can efficiently, conveniently, and flexibly mimic the pathological process of human silicosis with high repeatability and economy.

Silicosis can be caused by exposure to respiratory crystalline silica dust (CSD) in an industrial environment. The pathophysiology, screening, and ...

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Research

JoVE Journal

Medicine

2.0K Views.  Anhui University of Science and Technology. This protocol describes a method for establishing a mouse model of silicosis through repeated exposure to silica suspensions via a nasal drip. This model can efficiently, conveniently, and flexibly mimic the pathological process of human silicosis with high repeatability and economy.

Video: A Silicosis Mouse Model Established by Repeated Inhalation of Crystalline Silica Dust

A Mouse Model of in Utero Transplantation

The transplantation of stem cells and viruses in utero has tremendous potential for treating congenital disorders in the human fetus. For example, in utero transplantation (IUT) of hematopoietic stem cells has been used to successfully treat patients with severe combined immunodeficiency.1,2 In several other conditions, however, IUT has been attempted without success.3 Given these mixed results, the availability of an efficient non-human model to study the biological sequelae of stem cell transplantation and gene therapy is critical to advance this field. We and others have used the mouse model of IUT to study factors affecting successful engraftment of in utero transplanted hematopoietic stem cells in both wild-type mice4-7 and those with genetic diseases.8,9 The fetal environment also offers considerable advantages for the success of in utero gene therapy. For example, the delivery of adenoviral10, adeno-associated viral10, retroviral11, and lentiviral vectors12,13 into the fetus has resulted in the transduction of multiple organs distant from the site of injection with long-term gene expression. in utero gene therapy may therefore be considered as a possible treatment strategy for single gene disorders such as muscular dystrophy or cystic fibrosis. Another potential advantage of IUT is the ability to induce immune tolerance to a specific antigen. As seen in mice with hemophilia, the introduction of Factor IX early in development results in tolerance to this protein.14 
In addition to its use in investigating potential human therapies, the mouse model of IUT can be a powerful tool to study basic questions in developmental and stem cell biology. For example, one can deliver various small molecules to induce or inhibit specific gene expression at defined gestational stages and manipulate developmental pathways. The impact of these alterations can be assessed at various timepoints after the initial transplantation. Furthermore, one can transplant pluripotent or lineage specific progenitor cells into the fetal environment to study stem cell differentiation in a non-irradiated and unperturbed host environment.
The mouse model of IUT has already provided numerous insights within the fields of immunology, and developmental and stem cell biology. In this video-based protocol, we describe a step-by-step approach to performing IUT in mouse fetuses and outline the critical steps and potential pitfalls of this technique.

A Mouse Model of in Utero Transplantation

The mouse model of in utero transplantation is a versatile tool that can be used to study the potential clinical applications of stem cell transplantation and gene therapy in the fetus. In this protocol, we present a general approach to performing this technique

The transplantation of stem cells and viruses in utero has tremendous potential for treating congenital disorders in the human fetus.  For example, in ...

Mouse Model of Surgically-induced Endometriosis by Auto-transplantation of Uterine Tissue

Endometriosis is a chronic, painful disease whose etiology remains unknown. Furthermore, treatment of endometriosis can require laparoscopic removal of lesions, and/or chronic pharmaceutical management of pain and infertility symptoms. The cost associated with endometriosis has been estimated at 22 billion dollars per year in the United States1. To further our understanding of mechanisms underlying this enigmatic disease, animal models have been employed. Primates spontaneously develop endometriosis and therefore primate models most closely resemble the disease in women. Rodent models, however, are more cost effective and readily available2. The model that we describe here involves an autologous transfer of uterine tissue to the intestinal mesentery (Figure 1) and was first developed in the rat3 and later transferred to the mouse4. The goal of the autologous rodent model of surgically-induced endometriosis is to mimic the disease in women. We and others have previously shown that the altered gene expression pattern observed in endometriotic lesions from mice or rats mirrors that observed in women with the disease5,6. One advantage of performing the surgery in the mouse is that the abundance of transgenic mouse strains available can aid researchers in determining the role of specific components important in the establishment and growth of endometriosis. An alternative model in which excised human endometrial fragments are introduced to the peritoneum of immunocompromised mice is also widely used but is limited by the lack of a normal immune system which is thought to be important in endometriosis2,7. Importantly, the mouse model of surgically induced endometriosis is a versatile model that has been used to study how the immune system8, hormones9,10 and environmental factors11,12 affect endometriosis as well as the effects of endometriosis on fertility13 and pain14.

A description of the surgical induction of endometriosis in mice and rats by auto-transplantation of uterine tissue to the arterial cascade of the intestinal mesentery.

Endometriosis is a chronic, painful disease whose etiology remains unknown. Furthermore, treatment of endometriosis can require laparoscopic removal of ...

Mouse Model of Intraluminal MCAO: Cerebral Infarct Evaluation by Cresyl Violet Staining

Stroke is the third cause of mortality and the leading cause of disability in the World. Ischemic stroke accounts for approximately 80% of all strokes. However, the thrombolytic tissue plasminogen activator (tPA) is the only treatment of acute ischemic stroke that exists. This led researchers to develop several ischemic stroke models in a variety of species. Two major types of rodent models have been developed: models of global cerebral ischemia or focal cerebral ischemia. To mimic ischemic stroke in patients, in whom approximately 80% thrombotic or embolic strokes occur in the territory of the middle cerebral artery (MCA), the intraluminal middle cerebral artery occlusion (MCAO) model is quite relevant for stroke studies. This model was first developed in rats by Koizumi et al. in 1986 1. Because of the ease of genetic manipulation in mice, these models have also been developed in this species 2-3.
	Herein, we present the transient MCA occlusion procedure in C57/Bl6 mice. Previous studies have reported that physical properties of the occluder such as tip diameter, length, shape, and flexibility are critical for the reproducibility of the infarct volume 4. Herein, a commercial silicon coated monofilaments (Doccol Corporation) have been used. Another great advantage is that this monofilament reduces the risk to induce subarachnoid hemorrhages. Using the Zeiss stereo-microscope Stemi 2000, the silicon coated monofilament was introduced into the internal carotid artery (ICA) via a cut in the external carotid artery (ECA) until the monofilament occludes the base of the MCA. Blood flow was restored 1 hour later by removal of the monofilament to mimic the restoration of blood flow after lysis of a thromboembolic clot in humans. The extent of cerebral infarct may be evaluated first by a neurologic score and by the measurement of the infarct volume. Ischemic mice were thus analyzed for their neurologic score at different post-reperfusion times. To evaluate the infarct volume, staining with 2,3,5-triphenyltetrazolium chloride (TTC) was usually performed. Herein, we used cresyl violet staining since it offers the opportunity to test many critical markers by immunohistochemistry. In this video, we report the MCAO procedure; neurological scores and the evaluation of the infarct volume by cresyl violet staining.

The intraluminal middle cerebral occlusion model in mice is herein presented. The extent of cerebral infarct is evaluated by a neurologic score and cresyl violet staining, an alternative staining to TTC, offering the great advantage to test in parallel many interest markers.

Stroke is the third  cause of mortality and the leading cause of disability in the World. Ischemic  stroke accounts for approximately 80% of all strokes. ...

MRI Mapping of Cerebrovascular Reactivity via Gas Inhalation Challenges

The brain is a spatially heterogeneous and temporally dynamic organ, with different regions requiring different amount of blood supply at different time. Therefore, the ability of the blood vessels to dilate or constrict, known as Cerebral-Vascular-Reactivity (CVR), represents an important domain of vascular function. An imaging marker representing this dynamic property will provide new information of cerebral vessels under normal and diseased conditions such as stroke, dementia, atherosclerosis, small vessel diseases, brain tumor, traumatic brain injury, and multiple sclerosis. In order to perform this type of measurement in humans, it is necessary to deliver a vasoactive stimulus such as CO2 and/or O2 gas mixture while quantitative brain magnetic resonance images (MRI) are being collected. In this work, we presented a MR compatible gas-delivery system and the associated protocol that allow the delivery of special gas mixtures (e.g., O2, CO2, N2, and their combinations) while the subject is lying inside the MRI scanner. This system is relatively simple, economical, and easy to use, and the experimental protocol allows accurate mapping of CVR in both healthy volunteers and patients with neurological disorders. This approach has the potential to be used in broad clinical applications and in better understanding of brain vascular pathophysiology. In the video, we demonstrate how to set up the system inside an MRI suite and how to perform a complete experiment on a human participant.

Non-invasive imaging of the brain vasculature&#8217;s ability to dilate or constrict may allow a better understanding of cerebrovascular pathophysiology in various neurological diseases. The present report describes a reproducible and patient-comfortable protocol to perform vascular reactivity imaging in humans using magnetic resonance imaging (MRI).

The brain is a spatially heterogeneous and temporally dynamic organ, with different regions requiring different amount of blood supply at different time. ...

Dry Powder and Nebulized Aerosol Inhalation of Pharmaceuticals Delivered to Mice Using a Nose-only Exposure System

Obstructive respiratory diseases like asthma and chronic obstructive pulmonary disease (COPD) are currently treated by inhaled anti-inflammatory and bronchodilator drugs. Despite the availability of multiple treatments, both diseases are growing public health concerns. The majority of asthma patients are well controlled on current inhaled therapies but a substantial number of patients with severe asthma are not. Asthma affects an estimated 300 million people worldwide and approximately 20 percent have a severe form of the disease. In contrast to asthma, there are few effective therapies for COPD. An estimated 10% of the population has COPD and the trend in death rates is increasing for COPD while decreasing for other major diseases. Although developing drugs for inhaled delivery is challenging, the nose-only inhalation unit enables direct delivery of novel drugs to the lung of rodents for pre-clinical efficacy and safety/toxicology studies. Inhaled drug delivery has multiple advantages for respiratory diseases, where high concentration in the lung improves efficacy and minimizes systemic side effects. Inhaled corticosteroids and bronchodilators benefit from these advantages and inhaled delivery may also hold potential for future biologic therapies. The inhalation unit described herein can generate, sample for characterization, and uniformly deposit a drug aerosol in the lungs of rodents. This enables the pre-clinical determination of the efficacy and safety of drug doses deposited in the lungs of rodents, key data required before initiating clinical development.

The inhalation unit described herein can generate, sample for characterization, and uniformly deposit a drug aerosol in the lungs of rodents. This enables the pre-clinical determination of the efficacy and safety of drug doses deposited in the lungs; key data enabling clinical inhaled drug development.

Obstructive respiratory diseases like asthma and chronic obstructive pulmonary disease (COPD) are currently treated by inhaled anti-inflammatory and ...

Repeated Measurement of Respiratory Muscle Activity and Ventilation in Mouse Models of Neuromuscular Disease

Accessory respiratory muscles help to maintain ventilation when diaphragm function is impaired. The following protocol describes a method for repeated measurements over weeks or months of accessory respiratory muscle activity while simultaneously measuring ventilation in a non-anesthetized, freely behaving mouse. The technique includes the surgical implantation of a radio transmitter and the insertion of electrode leads into the scalene and trapezius muscles to measure the electromyogram activity of these inspiratory muscles. Ventilation is measured by whole-body plethysmography, and animal movement is assessed by video and is synchronized with electromyogram activity. Measurements of muscle activity and ventilation in a mouse model of amyotrophic lateral sclerosis are presented to show how this tool can be used to investigate how respiratory muscle activity changes over time and to assess the impact of muscle activity on ventilation. The described methods can easily be adapted to measure the activity of other muscles or to assess accessory respiratory muscle activity in additional mouse models of disease or injury.

This paper introduces a method for repeated measurements of ventilation and respiratory muscle activity in a freely behaving amyotrophic lateral sclerosis (ALS) mouse model throughout disease progression with whole-body plethysmography and electromyography via an implanted telemetry device.

Accessory respiratory muscles help to maintain ventilation when diaphragm function is impaired. The following protocol describes a method for repeated ...

Generation of a Chronic Obstructive Pulmonary Disease Model in Mice by Repeated Ozone Exposure

Chronic obstructive pulmonary disease (COPD) is characterized by persistent airflow limitation and lung parenchymal destruction. It has a very high incidence in aging populations. The current conventional therapies for COPD focus mainly on symptom-modifying drugs; thus, the development of new therapies is urgently needed. Qualified animal models of COPD could help to characterize the underlying mechanisms and can be used for new drug screening. Current COPD models, such as lipopolysaccharide (LPS) or the porcine pancreatic elastase (PPE)-induced emphysema model, generate COPD-like lesions in the lungs and airways but do not otherwise resemble the pathogenesis of human COPD. A cigarette smoke (CS)-induced model remains one of the most popular because it not only simulates COPD-like lesions in the respiratory system, but it is also based on one of the main hazardous materials that causes COPD in humans. However, the time-consuming and labor-intensive aspects of the CS-induced model dramatically limit its application in new drug screening. In this study, we successfully generated a new COPD model by exposing mice to high levels of ozone. This model demonstrated the following: 1) decreased forced expiratory volume 25, 50, and 75/forced vital capacity (FEV25/FVC, FEV50/FVC, and FEV75/FVC), indicating the deterioration of lung function; 2) enlarged lung alveoli, with lung parenchymal destruction; 3) reduced fatigue time and distance; and 4) increased inflammation. Taken together, these data demonstrate that the ozone exposure (OE) model is a reliable animal model that is similar to humans because ozone overexposure is one of the etiological factors of COPD. Additionally, it only took 6 - 8 weeks, based on our previous work, to create an OE model, whereas it requires 3 - 12 months to induce the cigarette smoke model, indicating that the OE model might be a good choice for COPD research.

This study describes the successful generation of a new chronic obstructive pulmonary disease (COPD) animal model by repeatedly exposing mice to high concentrations of ozone.

Chronic obstructive pulmonary disease (COPD) is characterized by persistent airflow limitation and lung parenchymal destruction. It has a very high ...

Isometric Contractility Measurement of the Mouse Mesenteric Artery Using Wire Myography

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and drugs. It is widely used in many studies on the physiological functions of vascular smooth muscle, the pathogenesis of vascular diseases such as hypertension, and the development of smooth muscle relaxant drugs. The mouse is a widely used model animal with a large pool of disease models and genetically modified strains. We introduced this method to measure the isometric contraction of mouse mesenteric artery in detail. A 1.4-mm segment of mouse mesenteric resistance artery was isolated and mounted on a myograph chamber by passing two steel wires through its lumen. After equilibration and normalization steps, the vessel segment was potentiated by a high-K+ solution twice prior to the contraction assay. As an example of the application of this method in drug development, we measured the relaxant effect of a novel natural substance, neoliensinine, isolated from a Chinese herb, embryos of the lotus seed (Nelumbo nucifera Gaertn.) on mouse mesenteric arteries. The vessel segments mounted on the myograph chamber were stimulated with a high-K+ solution. When the force tension reached a stable sustained phase, cumulative doses of neoliensinine were added to the chamber. We found that neoliensinine had a dose-dependent relaxant effect on smooth muscle contraction, thus suggesting that it bears potential activity against hypertension. In addition, as the vessel segment can survive at least 4 hours after mounting and maintain contractility induced by the high-K+ solution for many times, we suggest that the wire myograph system may be used for the time-consuming process of drug screening.

The wire myograph technique is used to investigate vascular smooth muscle functions and screen new drugs. We report a detailed protocol for measuring the isometric contractility of the mouse mesenteric artery and for screening new relaxants of vascular smooth muscle.

The wire myograph technique is used to assess the contractility of vascular smooth muscles in response to depolarization, GPCR agonists/inhibitors and ...

Vinyl Chloride and High-Fat Diet as a Model of Environment and Obesity Interaction

Vinyl chloride (VC), an abundant environmental contaminant, causes steatohepatitis at high levels, but is considered safe at lower levels. Although several studies have investigated the role of VC as a direct hepatotoxicant, the concept that VC modifies sensitivity of the liver to other factors, such as nonalcoholic fatty liver disease (NAFLD) caused by high-fat diet (HFD) is novel. This protocol describes an exposure paradigm to evaluate the effects of chronic, low-level exposure to VC. Mice are acclimated to low-fat or high-fat diet one week prior to the beginning of the inhalation exposure and remain on these diets throughout the experiment. Mice are exposed to VC (sub-OSHA level: &#60;1 ppm) or room air in inhalation chambers for 6 hours/day, 5 days/week, for up to 12 weeks. Animals are monitored weekly for body weight gain and food consumption. This model of VC exposure causes no overt liver injury with VC inhalation alone. However, the combination of VC and HFD significantly enhances liver disease. A technical advantage of this co-exposure model is the whole-body exposure, without restraint. Moreover, the conditions more closely resemble a very common human situation of a combined exposure to VC with underlying nonalcoholic fatty liver disease and therefore support the novel hypothesis that VC is an environmental risk factor for the development of liver damage as a complication of obesity (i.e., NAFLD). This work challenges the paradigm that the current exposure limits of VC (occupational and environmental) are safe. The use of this model can shed new light and concern on the risks of VC exposure. This model of toxicant-induced liver injury can be used for other volatile organic compounds and to study other interactions that may impact the liver and other organ systems.

The goal of this protocol was to develop a murine model of low-level toxicant exposure that does not cause overt liver injury but rather exacerbates pre-existing liver damage. This paradigm better recapitulates human exposure and the subtle changes that occur upon exposure to toxicant concentrations that are considered safe.

Vinyl chloride (VC), an abundant environmental contaminant, causes steatohepatitis at high levels, but is considered safe at lower levels. Although ...

Repeated Orotracheal Intubation in Mice

The literature describes several methods for mouse intubation that either require visualization of the glottis through the oral cavity or incision in the ventral neck for direct confirmation of cannula placement in the trachea. The relative difficulty or the tissue trauma induced to the subject by such procedures can be an impediment to an investigator&#8217;s ability to perform longitudinal studies. This article illustrates a technique in which physical manipulation of the mouse following the use of a depilatory to remove hair from the ventral neck permits transcutaneous visualization of the trachea for orotracheal intubation regardless of degree of skin pigmentation. This method is innocuous to the subject and easily achieved with a limited understanding of murine anatomy. This refined approach facilitates repeated intubation, which may be necessary for monitoring progression of disease or instillation of treatments. Using this method may result in a reduction of the number of animals and technical skill required to measure lung function in mouse models of respiratory disease.

The goal of this article is to describe a refined method of intubation of the laboratory mouse. The method is noninvasive and, therefore, ideal for studies that require serial monitoring of respiratory function and/or instillation of treatments into the lung.

The literature describes several methods for mouse intubation that either require visualization of the glottis through the oral cavity or incision in the ...