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

Summary

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.

Abstract

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 µm 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.

Introduction

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 concrete^1,2. 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 people^4,5,6. 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.

Protocol

All procedures followed the guidelines of the National Institutes of Health'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

1. 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.
2. Provide a constant light time of 12 h per day. Use a time control switch for precise timing.

2. Preparing the CSD suspension

1. At least 1 day before nasal drips, grind the silica in an agate mortar for 0.5 h.
2. Observe the size and shape of the crystal particles. Take representative photographs using scanning electron microscopy (SEM).
   1. 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.
   2. 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 µm (Figure 1A).
3. 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.
4. Stir and mix the CSD suspension thoroughly on a vortex mixer for 10 s before administering the nasal drips.

3. Administering nasal drips to mouse

1. 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.
2. Drip 50 µL of the CSD nasally within 4-8 s (Figure 1B, right).
   1. For the nasal drip, place the head of the mouse on the researcher's metacarpophalangeal joint with the tip of the index finger.
   2. 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.
   3. Aspirate 50 μl of liquid using a 200 μl pipette. Drop the liquid into the mouse's nasal cavity in three to four divided doses depending on the mouse's respiratory rate. Each instillation should consist of 15 to 20 μl liquid. Administer the drips once every 3 days, 5x within 12 days. Treat the control mouse with an equal amount of saline.
3. Gently massage the heart area of the mouse 5x-10x for 5 s.
   1. Hold the mouse's body with the palm, pinch the skin on the back of the neck with the thumb and index finger, and fix the mouse's hind limbs with the other fingers. Then, gently press the mouse's heart-beating area with the index finger of the other hand 5-10 times in a period of 5 s.
4. When the mouse'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.

4. Collecting the lung tissues and preparing a paraffin section

1. 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.
2. 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.
3. 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 °C for western blotting analysis.
4. 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.
5. After 72 h of fixation, embed specimens in paraffin.
   1. 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.
   2. Infiltrate the samples with the melted paraffin wax by heating to 50 °C for 2 h. Repeat this process in another cylinder. Cool the wax molds with tissues for 1 h to harden.
   3. After the wax has hardened and the tissue has been embedded, use a paraffin sectioning machine to slice the tissue at 5 µm. The precise sectioning and slide mounting steps were previously described¹¹.
      ​NOTE: Sufficient tissue perfusion is indicated by developing muscle twitching and tail twisting into an "S" shape or flexion after receiving 10 mL of 4% PFA.

5. Performing hematoxylin and eosin (HE) staining

1. Heat the paraffin-embedded tissues on a hot plate (60 °C) for more than 4 h to allow for adhesion to the slides and improved deparaffination.
2. 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.
3. 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.
4. 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 µL of neutral resin drops. Place the cover slide over the section and carefully lower it to avoid air bubbles.

6. Performing Masson staining

1. Dewax and hydrate the paraffin samples, as mentioned in step 5.2. Then, stain the cell nuclei with 50% Weigert'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's hematoxylin staining solution immediately before use.
2. Stain the samples with drops of Lichun red staining solution (40 µL 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.
3. 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 µL of neutral resin drops, as mentioned in step 5.4.

7. Performing Sirius red staining

1. Dewax and hydrate paraffin sections as mentioned in step 5.2.
2. Infiltrate the sections for 1 h with Sirius red staining solution.
3. 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.

8. Performing immunohistochemistry

1. Dewax and hydrate the paraffin samples as described in step 5.2.
2. 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.
3. 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.
4. Permeabilize the membrane of specimens for 15 min with 0.3% Triton-100 solution. Then, block with 30-40 µL of 5% bovine serum albumin (BSA) for 1 h.
5. Remove the blocking solution. Add diluted primary antibodies NF-κB (dilution 1:200) and CD68 (dilution 1:1,000) and incubate the specimens overnight at 2-8 °C in a microscope slide IHC wet box to prevent evaporation and light.
6. The next day, transfer them to RT for 1 h. Then, wash them with PBST 3x for 5 min each.
7. 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.
8. Incubate samples with the 3,3'-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.
9. 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.

9. Performing western blotting analysis

1. Lyse the lung tissues to extract proteins. Add 200 µL of RIPA working solution to 20 mg of the lung tissue.
2. 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 °C for 15 min at 14,800 x g.
3. Collect the supernatant and determine the protein concentration with the BCA protein assay kit. Make the protein storage solution with RIPA at 6 µg/µL protein. Add 20 µL of 5x loading buffer to the 80 µL of the protein lysate. Disrupt the secondary protein structure by heating the protein-containing microcentrifuge tubes in a metal bath (100 °C) for 20 min.
4. After cooling, aliquot 100 µL of the protein storage solution into each tube and store the samples in a -80 °C refrigerator. Dilute the protein concentration to 2–3 μg/μL with 1x loading buffer before electrophoresis.
   NOTE: The protein extraction process should be performed on ice. For the RIPA working solution, add 1 µL of 100 mM phenylmethylsulfonyl fluoride (PMSF) to 99 µL of RIPA to inhibit phosphorylated protein degradation.  Prepare 1x loading buffer by diluting 5x loading buffer with RIPA at a ratio of 1:4. 
5. Add 20 µg 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.
6. 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.
7. 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-κB (1:1,000) and β-actin (1:1,000) with 5% BSA. Submerge the strips in the antibody solution and shake the strips gently overnight at 2-8 °C.
8. 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.
9. Prepare an enhanced chemiluminescence (ECL) developer, drop it on the strip, and incubate for 3 min.
10. 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 β-actin as an internal control.

Representative Results

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 procedures included CSD suspension preparation, isoflurane-induced anesthesia, nasal drip, and thoracic massage¹². We demonstrated collagen deposition and myofibroblast differentiation following 4 weeks of static feeding¹². We have used this dust exposure method to study the underlying mechanism of pulmonary fibrosis caused by coal dust. The new subsets of macrophage and the intervention effect of vitamin D were found through single-cell transcriptome technology¹³. In this study, the progression of pulmonary fibrosis in mice was significantly accelerated by exposure to CSD, which caused damage to bronchial peripheral elastic fibers (Figure 2 and Figure 3). Silicosis nodules are composed of macrophages that contain CSD. Fibrosis nodular is enriched with many CSD, and CD68-positive macrophages actively engulf these particles. These deposited CSDs promote the formation of fibrous foci. As mentioned previously, after being exposed to CSD for 4 weeks, mice acquired obvious lesions, such as collagen deposition in silicon lung nodules and damage to lung tissue structure. There was also modest injury of the tissue surrounding the bronchi¹². The biological process of ER stress caused by CSD is related to NF-κB, which is involved in the inflammatory response (see Figure 4). Overall, these findings show that the proposed approach can effectively simulate the development of mouse silicosis.

Figure 1: Crystalline silica dust (CSD) particles below five microns were used for nasal drip. (A) Scanning electron microscopy (SEM) of CSD showed that the particles were irregularly shaped. (B) CSD suspension was used to prepare the mouse silicosis model by nasal drip. The machine on the left is used for isoflurane anesthesia, and the right panel outlines the key points that lead to the nasal drip working. Please click here to view a larger version of this figure.

Figure 2: Sirius red staining showed fibrosis in mouse lungs by nasal drip CSD for 1 month. (A) Sirius red staining was performed to measure collagen deposition in lung tissue following CSD or saline treatment (left upper and left lower in panes). Polarizing microscopy revealed three different types of collagen fibers (red, yellow, and green), of which type 1 collagen fibers shown in red are a risk factor for silicosis. However, no significant fibrosis was found in the vehicle group (right up and low panels). (B) Fibrosis score (FS) is a semi-quantitative evaluation index based on Sirius red staining¹² that significantly differs from the control (***P < 0.0001). Scale bar = 200 µm. Please click here to view a larger version of this figure.

Figure 3: Immunohistochemistry staining of CD68 in CSD-treated mouse lung to monitor the role of macrophages in a silicotic nodule forming. A typical silica nodule is characterized by liquefied necrosis after phagocytosis of CSD in the center, surrounded by macrophages in the periphery (HE staining). In addition, CSD was enriched in the nodules, accompanied by fibrosis (Masson staining). The immunohistochemistry staining of CD68 revealed that macrophages were widely present in lung tissue. Furthermore, these macrophages had ingested CSD (seen under the polarized light microscopy, right panel), which caused severe lung injury. Scale bar = 50 µm. Please click here to view a larger version of this figure.

Figure 4: The NF-κB expression in the CSD-treated mouse lung. (A) Immunohistochemistry staining was performed on the lung tissue. The CSD-treated mice lung on the right pane showed high NF-κB staining compared with the Vehicle group on the left. Scale bar = 50 µm. (B) Representative western blot showed that the CSD-treated mice have an increased NF-κB expression in the lung. Fresh lung tissue lysis was subjected to western blotting. (C) The NF-κB difference between the Sil and Veh groups was significant (** P < 0.01). Band intensity was measured using Image J. Please click here to view a larger version of this figure.

Discussion

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 relax, leading to deep breathing and allowing them to inhale CSD during a time window of slow, deep breathing. During this process, the operator holds the mandible of the mouse with his thumb and straightens its neck to prevent the liquid from entering the digestive tract, a method of respirable dust exposure that does not cause pain and is non-invasive. This method can meet the individual needs of repeated implementation to study dust exposure. Kato et al. in 2017 used a single large dose (125 mg/mL, 40 µL) via an oropharyngeal drip to model pulmonary fibrosis, and we refer to this concentration to try to explore the pulmonary changes induced by multiple small doses through the nasal cavity¹⁴.

There are several challenges associated with post-anesthesia nasal drips from a technical standpoint. During this process, the operator holds the mandible of the mouse with their thumb to straighten its neck and to prevent the liquid from entering the digestive tract. If the mice do not achieve deep anesthesia or the operator is not skilled and misses the time window for slow deep breathing, the effect of the nasal drip and the pathological characteristics of the model will not be as expected. Therefore, to avoid over-anesthetized death and under-anesthetized modeling failure, those who work with isoflurane-anesthetized mice must be adequately trained and master the critical characteristics of isoflurane-anesthetized mice before performing the nasal drip. Moreover, pressing the chest of the mice after the nasal drip, commonly called massaging, promotes the travel of CSD to reach the terminal bronchi and even the alveolar walls of the lungs. Mice anesthetized to a deep level were prone to asphyxiation when they received a high dose of nasal drips. However, if their chest was rapidly compressed, the survival rate increased. Last but not least, the creation of silicosis models necessitates mice with a sound physical foundation, and some studies have illustrated that mice older than 10-12 weeks of age have poor tolerance and an increased mortality rate after repeated exposure to CSD.Despite its advantages, this model has several disadvantages. One disadvantage is that mouse does not directly inhale dust particles through airflow. To address this issue, we have limited the amount of CSD suspension inhaled in a single breath and the frequency of repeated administration of CSD. Furthermore, we have set up vehicle groups. The data reveals that inhaling a small amount of liquid only temporarily disrupts ventilation, which will not damage the lungs of the mice¹². The vehicle control mice will quickly absorb the inhaled saline liquid without impacting the mice's lung function, body weight, or basal activity. On the contrary, repeated exposure to CSD through nose inhalation can create the desired dynamic model of silicosis in mice¹². Because different exposure frequencies affect the fibrosis process in models with repeated exposures, we do not have a clear time point. Usually, for a one-time exposure of silica dust, day 7 is still in the early stages the silicosis. Days 7-14 correspond to the inflammatory activity stage, days 14-28 match the fibrosis transformation stage, and the time after day 28 corresponds to the fibrosis formation stage^12,15. However, the pathology at these time points can vary depending on the dose.

The nasal drip method has several advantages over traditional endotracheal drips or surgically exposed tracheostomy approaches^16,17, such as reducing tracheotomy-related infections and postoperative care. This approach can also satisfy the requirement for repeated CSD exposure during one experiment and does not require expensive equipment. In addition, the nasal drip method is a more realistic representation of dust exposure, as small particles enter the lungs from the upper respiratory tract and travel to the terminal bronchioles and alveoli.Since we use multiple small doses of silica drips, the silica dust dose can enter the lung to the maximum extent and continuously stimulate the organism. Thus, this modeling approach is developed to simulate the recurrent exposure of employees to CS in their work environment and to explore the pathological process of silicosis.

Creating mouse models of silicosis via nasal exposure to CSD can be applied to repeated exposures. Therefore, this animal model can be used to study the effects of exposure frequency and dosing on silicosis progression and the dynamic pathological features and mechanisms of silicosis. Moreover, combined exposure with other substances or medications is also very feasible.

Materials

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