The authors would like to acknowledge the National Institute of Biological Sciences, Beijing for the assistance. This work was supported by Beijing Municipal Natural Science Foundation (No. Z17110200040000).

All procedures used in this protocol were carried out in accordance with the recommendations in the Guidelines for the Care and Use of Laboratory Animals of the National Institute of Biological Sciences, Beijing. 8 week-old CD-1 male mice were housed in a specific pathogen free (SPF) facility until the experiments were conducted. Surgeries were performed using completely anesthetized mice (i.e., without any toe pinch responses). After surgery, mice were kept in a warm, humid room with sufficient food and fresh water. Mice were sacrificed using an overdose of anesthetic delivered by intraperitoneal injection.
1. Mouse PNX Surgery
<ol>
	<li>Fully anesthetize the mice with sodium phenobarbital (120 mg / kg body weight) and buprenorphine (0.1 mg / kg body weight) via intraperitoneal (I.P.) injection. Perform surgery when mice no longer react to toe pinching.</li>
	<li>Remove hair on the left thorax of the mice with chemical depilatory treatment (~3 x 3 cm2 area).</li>
	<li>Secure each mouse on an intubation platform with its ventral side facing the operator (Figure 1A).</li>
	<li>Pull out the mouse tongue and illuminate the vocal cords with a small animal laryngoscope containing a notch for guiding catheters18 (Figure 1A).</li>
	<li>Distinguish the vocal cords by observing the movements of the vocal cords during breathing. Gently insert a 20 G intravenous intubation cannula into the trachea at an anterior angle19.</li>
	<li>Place mice in a right lateral recumbent position and connect the cannula to a mechanical ventilator (e.g., pressure controlled; see the Table of Materials). Check the insertion of the cannula into the trachea by observing the breathing movements of the mouse chest (Figure 1B).</li>
	<li>Set the inspiratory pressure of the ventilator to 12 cm H2O and set the respiratory rate to 120 breaths per min (Figure 1B).</li>
	<li>Decontaminate the skin in the surgical area with betadine and 70% ethanol.</li>
	<li>Make a 2 - 3 cm posterolateral thoracotomy incision in the space at the 5th intercostal space, cut through skin and muscles with Noyes Spring Scissors (cutting edge: 14 mm; tip diameter: 0.275 mm) (Figure 2B, C). Surgical instruments used for thoracotomy procedure are sterilized prior to use.</li>
	<li>Make a 1.5 cm incision at the 5th intercostal space to expose the left lung (Figure 2D, E). During the operation, use a high temperature cauterizer to stop bleeding.</li>
	<li>Lift one-third of the left lung lobe from the chest with blunt tip forceps (Figure 2F), and then use a cotton swab to pull out the entire left lung (Figure 2G).</li>
	<li>Identify the pulmonary artery and bronchi of the left lung lobe (Figure 2G).</li>
	<li>Tightly ligate the bronchi and vessels at the hilum with a silk surgical suture and cut out the left lung lobe at 3 - 4 mm from the ligation (Figure 2H, I). 
	NOTE: Be careful not to cut off the suture knots on the left hilum, which can cause pneumothorax (i.e., air or gas in the cavity of the thorax).</li>
	<li>Close the chest wall with 1 suture, and then stitch the muscle layer and the skin layer sequentially, using 5 - 6 interrupted sutures. Leave a 3 - 4 mm gap between each suture (Figure 2M, 2N). 
	NOTE: Keep the surgical suture needle away from the heart; inadvertent cardiac puncture will result in immediate death.</li>
	<li>Disinfect the surgical area with povidone-iodine.</li>
	<li>After the surgical operation, place the mouse on a 38 &#176;C thermal pad and connect the mouse to the ventilator until spontaneous breathing movements commence (Figure 2O).</li>
</ol>
2. Prosthesis Implantation
<ol>
	<li>Perform steps 1.1 - 1.13 of the PNX procedure (that is, up to the point when the left lung lobe of the mouse is removed).</li>
	<li>Clamp the center of the silicone prosthesis (customer made, 12 mm in length, 3 mm in thickness, 7 mm in width, 0.2 g, ellipsoid-shape) using blunt forceps (Figure 2J). Sterilize silicone prosthesis prior to insertion.</li>
	<li>Hold the rib with forceps with one hand to expose the thoracic cavity, and then insert the prosthesis into the left empty thoracic cavity with another hand. 
	NOTE: The insertion angle is approximately 45 degrees between the frontal plane of the prosthesis and the thoracic surface (Figure 2K, L). Be very gentle when inserting the prosthesis. Excessive force will result in pleural rupture.</li>
	<li>Adjust the orientation of the prosthesis with blunt forceps to ensure that the prosthesis occupies the left empty thoracic cavity.</li>
	<li>Perform steps 1.14 - 1.16 of the mouse PNX procedure.</li>
</ol>
3. Measurement of ILV
<ol>
	<li>Prepare a custom device (&#34;inflation tube&#34;) that consists of a plunger removed from a disposable serological pipette (10 mL), a 40 cm long flexible tube with a needle adapter, a flow rate control valve, and an 18 G needle. After assembly, secure the pipette on a board with tape (Figure 3A). The distance between the top of the pipette and the experimental bench must be at least 30 cm.</li>
	<li>Prepare fresh 4% paraformaldehyde (PFA) fixation solution by dissolving 20 g PFA in 500 mL pre-heated 1x phosphate buffered saline (PBS) in a 55 &#176;C water bath, shaking manually once every 10 min until the solution is clear. After cooling to room temperature, filter the solution with a 0.45 &#181;m filter. 
	CAUTION: Wear appropriate personal protective equipment (PPE) when handling PFA.</li>
	<li>Sacrifice mice with an overdose injection of anesthetic (0.8% phenobarbital sodium, 1,000 U/mL heparin).</li>
	<li>Secure each mouse on a polystyrene dissection plate and spray it with 70% alcohol.</li>
	<li>Carefully open the mouse chest and cut out the sternum using scissors to thoroughly expose the lung lobes.</li>
	<li>Remove excessive tissue using scissors to expose the trachea. Make sure to separate the trachea from the esophagus.</li>
	<li>Cut the abdominal aorta and insert a 25-gauge needle into the right ventricle of heart; connect the needle to a 20 mL syringe prior to this insertion. Slowly push 1x PBS into the heart to remove blood cells until lungs turn white. Typically, 5 - 10 mL PBS is required to clear the pulmonary blood vessels.</li>
	<li>Fill the custom-constructed inflation tube with 4% fresh PFA and remove all the bubbles from the inflation tube.</li>
	<li>Insert the 18-gauge needle of the inflation tube into the trachea and clip the trachea with vessel clips to avoid fluid leakage.</li>
	<li>Inflate lungs with 4% PFA at a constant transpulmonary pressure of 25 cm/H2O2,20. Incubate the lungs at room temperature for 2 h to achieve fully expanded lungs. This &#34;pre-fix&#34; step is critical for preserving lung morphology.
	<ol>
		<li>By monitoring the inflation tube, record the value of the initial 4% PFA volume and record the final volume. The internal lung volume equals the initial 4% PFA volume minus the final 4% PFA volume.</li>
	</ol>
	</li>
	<li>Ligate the trachea and using scissors, gently dissect out the lungs (keeping the lungs intact) from surrounding connective tissues. Be very gentle to avoid damaging the lungs.</li>
	<li>Incubate the lungs in a 50-mL conical tube filled with 4% PFA for 12 h at 4 &#176;C with gentle shaking on an shaker (50 rpm). Proceed to tissue processing and staining (see section 4).</li>
</ol>
4. Tissue Embedding, Sectioning, and Hematoxylin &#38; Eosin (H&#38;E) Staining
<ol>
	<li>After fixation, use Noyes Spring Scissors to trim the heart and excessive connective tissues off the lungs. Gently separate the individual lung lobes by cutting off the bronchus that connects the lung lobes to the trachea.</li>
	<li>Extensively wash the lung lobes 3 - 4 times in 50 mL 1x PBS (30 min/wash) on an orbital shaker (50 rpm).</li>
	<li>Following the final wash, cryoprotect the lung lobes by immersing them in a 30% sucrose solution (in 1x PBS) at 4 &#176;C until the tissue sinks to the bottom of the 50-mL conical tubes (approximately 12 h).</li>
	<li>Prior to embedding and cryosectioning the tissues, remove the lung lobe samples from the tubes with forceps, retain the accessory lobes for the histological analysis, dab the remaining sucrose solution from the surface of the accessory lobe samples, and then thoroughly immerse the sample into a Petri dish containing optimal cutting temperature (O.C.T) compound for approximately 30 min.</li>
	<li>Freeze the O.C.T-embedded accessory lobe samples in liquid nitrogen using cryomolds. Position the largest surface area of the lobe parallel to the bottom of the mold.</li>
	<li>Prepare a total of three 10-&#181;m-thick sections for each sample during cryosectioning for histological analysis. Discard the first 1 mm of tissue, collect one 10-&#181;m-thick section, discard 0.5 mm of tissue, collect another section, discard 0.5 mm of tissue, and collect the third (final) section.</li>
	<li>Air dry the sections for 1 h before performing H&#38;E staining.</li>
	<li>Perform H&#38;E staining
	<ol>
		<li>Wash the sections in 3 - 4 changes of tap water and then stain the sections in fresh hematoxylin for 2 min; rinse the section under running tap water; immerse the section two times in a 1% HCl-70% ethanol solution to remove excess hematoxylin.</li>
		<li>Stain the section in fresh eosin for 3 min; dehydrate the sections with two successive 30 s washes in 95% ethanol and two 30 s washes with 100% ethanol; clear the sections in xylene for 30 s, repeat the clearing step once in fresh xylene; mount the slides with mounting medium using glass coverslips.</li>
	</ol>
	</li>
</ol>
5. Quantification of MLI
<ol>
	<li>Acquire digital images of the H&#38;E stained accessory lobe sections (20X magnification) using a bright-field microscope.</li>
	<li>To quantify the MLI, select a total of 15 non-overlapping views (1,000 &#181;m x 1,000 &#181;m) randomly from the suitable areas (without arteries and veins, major airways, and alveolar ducts) of 3 sections.</li>
	<li>Place a grid with 10 evenly-distributed vertical lines and 10 equally-distributed horizontal lines of defined length (1,000 &#181;m) on the chosen areas of view using a ruler tool; each line is thus spaced 100 &#181;m apart (Figure 4B).</li>
	<li>Define the value of one intercept as the linear length between two adjacent alveolar epithelia. Measure the values of all intercepts along each 1,000 &#181;m length line.</li>
	<li>For each grid, quantify the values of all intercepts among the 10 horizontal 1,000 &#181;m length lines and the 10 vertical 1,000 &#181;m length lines. 
	NOTE: MLI is the average value of the intercept lengths from a total of 15 grids analyzed from among the 3 sections prepared for each of the accessory lobes.</li>
</ol>
6. Calculation of ISA
<ol>
	<li>Calculate the ISA using Equation 1 (see the Introduction). Refer to section 3 for the measurement of ILV and refer to section 5 for the quantification of MLI.</li>
</ol>

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The authors have nothing to disclose.

In this protocol, we provide detailed descriptions about the measurement of pulmonary parameters after mouse left lung PNX and prosthesis implantation. ISA is now considered to be a key metric for the assessment of respiratory function in many pulmonary diseases and in injury-induced alveolar regeneration. However, although the pulmonary research community is in agreement about the utility of ISA as a useful metric, to date, there has been little consideration of the standardization of the measurement of ILV and MLI, the...

The fundamental function of the lung is the exchange of oxygen and carbon dioxide between blood vessels and the atmosphere. Lung diseases such as bronchopulmonary dysplasia (BPD), chronic obstructive pulmonary disease (COPD), and acute respiratory infections, result in decreased ISA2. Researchers studying lung disease have developed several quantitative methods to evaluate morphological changes in lungs, including MLI, ILV, number of gas exchange units, ISA, and lung tissue compliance2,3. Pioneering studies by Weibel et al.4 and Duguid et al.5 together established that ISA can be used as a direct measure of lung gas-exchange capacity in human lungs and can be used as a criterion to determine emphysema severity. A number of studies published in the last five years have used lung morphological parameters (e.g., ISA and MLI) to assess morphological and functional changes in the lungs of mice during development6 and during recovery from injury PNX1,7. ISA is calculated using Equation 18,9:
<img alt="Equation" src="/files/ftp_upload/56114/56114eq1.jpg" />
, where ILV is the internal lung volume and MLI is an intermediary parameter that represents the pulmonary peripheral airspace size10.
PNX, the surgical removal of one or more lung lobes, has been widely reported to induce alveolar regeneration in many species, including humans11, mice1, dogs12, rats13, and rabbits14,15. A study of mice lungs at fourteen days post-PNX showed that both the expansion of pre-existing alveoli and the de novo formation of alveoli contribute to the restoration of ISA, ILV, and the number of alveoli in the remaining lung tissues1. We and others have shown that the insertion of materials such as sponge, wax, or a custom-shaped prosthesis into the empty thoracic cavity following PNX (i.e., prosthesis implantation) impairs alveolar regeneration. It is now firmly established that mechanical force functions as one of the most important factors for initiating alveolar regeneration1,16,17. Such studies have highlighted the effectiveness of using ISA values from PNX-treated and Prosthesis-implanted lungs as a criterion to quantitatively evaluate alveolar regeneration.
Observer bias is known to significantly influence measured values for lung morphological parameters (e.g., MIL and ILV). Standardized protocols can be used to obviate this bias in determining both ILV and MLI, which are the two parameters used in the calculation of ISA. Here, we provide highly-detailed, standardized protocols for measuring these lung parameters. Importantly, the ability to accurately quantify ISA promises to improve the reliability and reproducibility of studies of lung function in injury-induced alveolar regeneration models and should facilitate mechanistic discoveries in multiple pulmonary diseases.

<table><tbody><tr><td>Low cost cautery kit</td><td>Fine Science Tools</td><td>18010-00</td><td></td></tr><tr><td>Noyes scissors</td><td>Fine Science Tools</td><td>15012-12</td><td></td></tr><tr><td>Standard pattern forceps</td><td>Fine Science Tools</td><td>11000-12</td><td></td></tr><tr><td>Castroviejo Micro Needle Holders</td><td>Fine Science Tools</td><td>12060-01</td><td></td></tr><tr><td>Vessel clips</td><td>Fine Science Tools</td><td>18374-44</td><td></td></tr><tr><td>I. V. Cannula-20 gauge</td><td>Jinhuan Medical Product Co., LTD.</td><td>29P0601</td><td></td></tr><tr><td>Surgical suture</td><td>Jinhuan Medical Product Co., LTD.</td><td>F602</td><td></td></tr><tr><td>Mouse intubation platform</td><td>Penn-Century, Inc</td><td>Model MIP</td><td></td></tr><tr><td>Small Animal Laryngoscope</td><td>Penn-Century, Inc</td><td>Model LS-2-M</td><td></td></tr><tr><td>TOPO Small Animal Ventilator</td><td>Kent Scientific</td><td>RSP1006-05L</td><td></td></tr><tr><td>Thermal pad</td><td>Stuart equipment</td><td>SBH130D</td><td></td></tr><tr><td>Pentobarbital sodium salt</td><td>Sigma</td><td>P3761</td><td></td></tr><tr><td>Heparin sodium salt</td><td>Sigma</td><td>H3393</td><td></td></tr><tr><td>Hematoxylin Solution</td><td>Sigma</td><td>GHS132</td><td></td></tr><tr><td>Eosin Y solution, alcoholic</td><td>Sigma</td><td>HT110116</td><td></td></tr><tr><td>10 mL Pipette</td><td>Thermo Scientific</td><td>170356</td><td></td></tr><tr><td>Paraformaldehyde</td><td>Sigma</td><td>P6148</td><td></td></tr><tr><td>O.C.T Compound</td><td>Tissue-Tek</td><td>4583</td><td></td></tr><tr><td>cryosection machine</td><td>Leica</td><td>CM1950</td><td></td></tr><tr><td>Disposable Base Molds</td><td>Fisher HealthCare</td><td>22-363-553</td><td></td></tr><tr><td>18 gauge needle</td><td>Becton Dickinson</td><td>305199</td><td></td></tr><tr><td>Povidone iodine</td><td>Fisher Scientific</td><td>19-027132</td><td></td></tr><tr><td>70% ethanol</td><td>Fisher Scientific</td><td>BP82011</td><td></td></tr><tr><td>Infusion sets for single use</td><td>Weigao</td><td>SFDA 2012 3661704</td><td></td></tr><tr><td>Phosphate buffered saline</td><td>Gibco</td><td>10010023</td><td></td></tr><tr><td>Tapes</td><td>3M Scotch</td><td>8915</td><td></td></tr><tr><td>Cotton pad</td><td>Vinda</td><td>Dr.P</td><td></td></tr><tr><td>Silicone prosthesis</td><td>Custom made</td><td></td><td></td></tr><tr><td>Brightfield microscope</td><td>Olympus</td><td>VS120</td><td></td></tr><tr><td>Ruler tool</td><td>Adobe Photoshop</td><td></td><td></td></tr></tbody></table>

a standardized method for measuring internal lung surface area via mouse pneumonectomy and prosthesis implantation

Pulmonary morphology, physiology, and respiratory functions change in both physiological and pathological conditions. Internal lung surface area (ISA), representing the gas-exchange capacity of the lung, is a critical criterion to assess respiratory function. However, observer bias can significantly influence measured values for lung morphological parameters. The protocol that we describe here minimizes variations during measurements of two morphological parameters used for ISA calculation: internal lung volume (ILV) and mean linear intercept (MLI). Using ISA as a morphometric and functional parameter to determine the outcome of alveolar regeneration in both pneumonectomy (PNX) and prosthesis implantation mouse models, we found that the increased ISA following PNX treatment was significantly blocked by implantation of a prosthesis into the thoracic cavity1. The ability to accurately quantify ISA is not only expected to improve the reliability and reproducibility of lung function studies in injured-induced alveolar regeneration models, but also to promote mechanistic discoveries of multiple pulmonary diseases.

We performed here an experiment with a PNX-treated group and a prosthesis implantation (Prosthesis-implanted) group. These groupings are the same as the groupings used in a previously-published study from our research group14.
The mouse PNX and prosthesis implantation procedures are shown in Figure 2. 8 week-old CD-1 male mice are used for the surgeries and for the quantifica...

Watch this Scientific Journal Video about A Standardized Method for Measuring Internal Lung Surface Area via Mouse Pneumonectomy and Prosthesis Implantation at JoVE.com

A Standardized Method for Measuring Internal Lung Surface Area via Mouse Pneumonectomy and Prosthesis Implantation

Internal lung surface area (ISA) is a critical criterion for assessing lung morphology and physiology in lung diseases and injury-induced alveolar regeneration. We describe here a standardized method that can minimize the measurement bias for ISA in both lung pneumonectomy and prosthesis implantation mouse models.

Pulmonary morphology, physiology, and respiratory functions change in both physiological and pathological conditions. Internal lung surface area (ISA), ...

a-standardized-method-for-measuring-internal-lung-surface-area-via

Research

JoVE Journal

Developmental Biology

13.1K Views.  Peking University. Internal lung surface area (ISA) is a critical criterion for assessing lung morphology and physiology in lung diseases and injury-induced alveolar regeneration. We describe here a standardized method that can minimize the measurement bias for ISA in both lung pneumonectomy and prosthesis implantation mouse models. 

Video: A Standardized Method for Measuring Internal Lung Surface Area via Mouse Pneumonectomy and Prosthesis Implantation

A Simple Method of Mouse Lung Intubation

A simple procedure to intubate mice for pulmonary function measurements would have several advantages in longitudinal studies with limited numbers or expensive animal. One of the reasons that this is not done more routinely is that it is relatively difficult, despite there being several published studies that describe ways to achieve it. In this paper we demonstrate a procedure that eliminates one of the major hurdles associated with this intubation, that of visualizing the trachea during the entire time of intubation. The approach uses a 0.5 mm fiberoptic light source that serves as an introducer to direct the intubation cannula into the mouse trachea. We show that it is possible to use this procedure to measure lung mechanics in individual mice over a time course of at least several weeks. The technique can be set up with relatively little expense and expertise, and it can be routinely accomplished with relatively little training. This should make it possible for any laboratory to routinely carry out this intubation, thereby allowing longitudinal studies in individual mice, thereby minimizing the number of mice needed and increasing the statistical power by using each mouse as its own control.

This paper describes a striaghforward and efficient method of intubating mice for pulmonary function measurements or pulmonary instillation, that allows the mice to recover and be studied at later times. The procedure involves an inexpensive fiberoptic light source that directly illuminates the trachea.

A  simple procedure to intubate mice for pulmonary function measurements would  have several advantages in longitudinal studies with limited numbers or  ...

Medicine

In vivo Measurement of the Mouse Pulmonary Endothelial Surface Layer

The endothelial glycocalyx is a layer of proteoglycans and associated glycosaminoglycans lining the vascular lumen. In vivo, the glycocalyx is highly hydrated, forming a substantial endothelial surface layer (ESL) that contributes to the maintenance of endothelial function. As the endothelial glycocalyx is often aberrant in vitro and is lost during standard tissue fixation techniques, study of the ESL requires use of intravital microscopy. To best approximate the complex physiology of the alveolar microvasculature, pulmonary intravital imaging is ideally performed on a freely-moving lung. These preparations, however, typically suffer from extensive motion artifact. We demonstrate how closed-chest intravital microscopy of a freely-moving mouse lung can be used to measure glycocalyx integrity via ESL exclusion of fluorescently-labeled high molecular weight dextrans from the endothelial surface. This non-recovery surgical technique, which requires simultaneous brightfield and fluorescent imaging of the mouse lung, allows for longitudinal observation of the subpleural microvasculature without evidence of inducing confounding lung injury.

In vivo Measurement of the Mouse Pulmonary Endothelial Surface Layer

The endothelial glycocalyx/endothelial surface layer is ideally studied using intravital microscopy. Intravital microscopy is technically challenging in a moving organ such as the lung. We demonstrate how simultaneous brightfield and fluorescent microscopy may be used to estimate endothelial surface layer thickness in a freely-moving in vivo mouse lung.

The endothelial glycocalyx is a layer of proteoglycans and associated glycosaminoglycans lining the vascular lumen. In vivo, the glycocalyx is highly ...

A Permanent Window for Investigating Cancer Metastasis to the Lung

Metastasis, accounting for ~90% of cancer-related mortality, involves the systemic spread of cancer cells from primary tumors to secondary sites such as the bone, brain, and lung. Although extensively studied, the mechanistic details of this process remain poorly understood. While common imaging modalities, including computed tomography (CT), positron emission tomography (PET), and magnetic resonance imaging (MRI), offer varying degrees of gross visualization, each lacks the temporal and spatial resolution necessary to detect the dynamics of individual tumor cells. To address this, numerous techniques have been described for intravital imaging of common metastatic sites. Of these sites, the lung has proven especially challenging to access for intravital imaging owing to its delicacy and critical role in sustaining life. Although several approaches have previously been described for single-cell intravital imaging of the intact lung, all involve highly invasive and terminal procedures, limiting the maximum possible imaging duration to 6-12 h. Described here is an improved technique for the permanent implantation of a minimally invasive thoracic optical Window for High-Resolution Imaging of the Lung (WHRIL). Combined with an adapted approach to microcartography, the innovative optical window facilitates serial intravital imaging of the intact lung at single-cell resolution across multiple imaging sessions and spanning multiple weeks. Given the unprecedented duration of time over which imaging data can be gathered, the WHRIL can facilitate the accelerated discovery of the dynamic mechanisms underlying metastatic progression and numerous additional biologic processes within the lung.

Here, we present a protocol for the surgical implantation of a permanently indwelling optical window for the murine thorax, which enables high-resolution, intravital imaging of the lung. The permanence of the window makes it well-suited to the study of dynamic cellular processes in the lung, especially those that are slowly evolving, such as metastatic progression of disseminated tumor cells.

Metastasis, accounting for ~90% of cancer-related mortality, involves the systemic spread of cancer cells from primary tumors to secondary sites such as ...

Cancer Research

Automated Measurement of Pulmonary Emphysema and Small Airway Remodeling in Cigarette Smoke-exposed Mice

COPD is projected to be the third most common cause of mortality world-wide by 2020(1). Animal models of COPD are used to identify molecules that contribute to the disease process and to test the efficacy of novel therapies for COPD. Researchers use a number of models of COPD employing different species including rodents, guinea-pigs, rabbits, and dogs(2). However, the most widely-used model is that in which mice are exposed to cigarette smoke. Mice are an especially useful species in which to model COPD because their genome can readily be manipulated to generate animals that are either deficient in, or over-express individual proteins. Studies of gene-targeted mice that have been exposed to cigarette smoke have provided valuable information about the contributions of individual molecules to different lung pathologies in COPD(3-5). Most studies have focused on pathways involved in emphysema development which contributes to the airflow obstruction that is characteristic of COPD. However, small airway fibrosis also contributes significantly to airflow obstruction in human COPD patients(6), but much less is known about the pathogenesis of this lesion in smoke-exposed animals. To address this knowledge gap, this protocol quantifies both emphysema development and small airway fibrosis in smoke-exposed mice. This protocol exposes mice to CS using a whole-body exposure technique, then measures respiratory mechanics in the mice, inflates the lungs of mice to a standard pressure, and fixes the lungs in formalin. The researcher then stains the lung sections with either Gill&#8217;s stain to measure the mean alveolar chord length (as a readout of emphysema severity) or Masson&#8217;s trichrome stain to measure deposition of extracellular matrix (ECM) proteins around small airways (as a readout of small airway fibrosis). Studies of the effects of molecular pathways on both of these lung pathologies will lead to a better understanding of the pathogenesis of COPD.

The goal of this protocol is to provide automated methods to quantify chronic lung pathologies in a murine model of COPD. The protocol includes exposing mice to cigarette smoke (CS), measuring pulmonary function, inflating the lungs, and using morphometry methods to measure emphysema and small airway remodeling in mice.

COPD is projected to be the third most common cause of mortality world-wide by 2020(1).  Animal models of COPD are used to identify molecules that ...

Mouse Pneumonectomy Model of Compensatory Lung Growth

In humans, disrupted repair and remodeling of injured lung contributes to a host of acute and chronic lung disorders which may ultimately lead to disability or death. Injury-based animal models of lung repair and regeneration are limited by injury-specific responses making it difficult to differentiate changes related to the injury response and injury resolution from changes related to lung repair and lung regeneration. However, use of animal models to identify these repair and regeneration signaling pathways is critical to the development of new therapies aimed at improving pulmonary function following lung injury. The mouse pneumonectomy model utilizes compensatory lung growth to isolate those repair and regeneration signals in order to more clearly define mechanisms of alveolar re-septation. Here, we describe our technique for performing mouse pneumonectomy and sham pneumonectomy. This technique may be utilized in conjunction with lineage tracing or other transgenic mouse models to define molecular and cellular mechanism of lung repair and regeneration.

Mouse pneumonectomy is a commonly employed model of compensatory lung growth. This procedure can be used in conjunction with lineage tracing or transgenic mouse models to elucidate underlying mechanisms.

In humans, disrupted repair and remodeling of injured lung contributes to a host of acute and chronic lung disorders which may ultimately lead to ...

Measurement of the Pressure-volume Curve in Mouse Lungs

In recent decades the mouse has become the primary animal model of a variety of lung diseases. In models of emphysema or fibrosis, the essential phenotypic changes are best assessed by measurement of the changes in lung elasticity. To best understand specific mechanisms underlying such pathologies in mice, it is essential to make functional measurements that can reflect the developing pathology. Although there are many ways to measure elasticity, the classical method is that of the total lung pressure-volume (PV) curve done over the whole range of lung volumes. This measurement has been made on adult lungs from nearly all mammalian species dating back almost 100 years, and such PV curves also played a major role in the discovery and understanding of the function of pulmonary surfactant in fetal lung development. Unfortunately, such total PV curves have not been widely reported in the mouse, despite the fact that they can provide useful information on the macroscopic effects of structural changes in the lung. Although partial PV curves measuring just the changes in lung volume are sometimes reported, without a measure of absolute volume, the nonlinear nature of the total PV curve makes these partial ones very difficult to interpret. In the present study, we describe a standardized way to measure the total PV curve. We have then tested the ability of these curves to detect changes in mouse lung structure in two common lung pathologies, emphysema and fibrosis. Results showed significant changes in several variables consistent with expected structural changes with these pathologies. This measurement of the lung PV curve in mice thus provides a straightforward means to monitor the progression of the pathophysiologic changes over time and the potential effect of therapeutic procedures.

Here we present a protocol to simply and reliably measure the lung pressure-volume curve in mice, showing that it is sufficiently sensitive to detect phenotypic parenchymal changes in two common lung pathologies, pulmonary fibrosis and emphysema. This metric provides a means to quantify the lung&#8217;s structural changes with developing pathology.

In recent decades the mouse has become the primary animal model of a variety of lung diseases. In models of emphysema or fibrosis, the essential ...

Instillation and Fixation Methods Useful in Mouse Lung Cancer Research

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

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

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

Intraperitoneal Glucose Tolerance Test, Measurement of Lung Function, and Fixation of the Lung to Study the Impact of Obesity and Impaired Metabolism on Pulmonary Outcomes

Obesity and respiratory disorders are major health problems. Obesity is becoming an emerging epidemic with an expected number of over 1 billion obese individuals worldwide by 2030, thus representing a growing socioeconomic burden. Simultaneously, obesity-related comorbidities, including diabetes as well as heart and chronic lung diseases, are continuously on the rise. Although obesity has been associated with increased risk for asthma exacerbations, worsening of respiratory symptoms, and poor control, the functional role of obesity and perturbed metabolism in the pathogenesis of chronic lung disease is often underestimated, and underlying molecular mechanisms remain elusive. This article aims to present methods to assess the effect of obesity on metabolism, as well as lung structure and function. Here, we describe three techniques for mice studies: (1) assessment of intraperitoneal glucose tolerance (ipGTT) to analyze the effect of obesity on glucose metabolism; (2) measurement of airway resistance (Res) and respiratory system compliance (Cdyn) to analyze the effect of obesity on lung function; and (3) preparation and fixation of the lung for subsequent quantitative histological assessment. Obesity-related lung diseases are probably multifactorial, stemming from systemic inflammatory and metabolic dysregulation that potentially adversely influence lung function and the response to therapy. Therefore, a standardized methodology to study molecular mechanisms and the effect of novel treatments is essential.

The incidence of obesity is rising and increases the risk of chronic lung diseases. To establish the underlying mechanisms and preventive strategies, well-defined animal models are needed. Here, we provide three methods (glucose-tolerance-test, body plethysmography, and lung fixation) to study the effect of obesity on pulmonary outcomes in mice.

Obesity and respiratory disorders are major health problems. Obesity is becoming an emerging epidemic with an expected number of over 1 billion obese ...

Immunology and Infection

Pre-Conditioning the Airways of Mice with Bleomycin Increases the Efficiency of Orthotopic Lung Cancer Cell Engraftment

Lung cancer is a deadly treatment refractory disease that is biologically heterogeneous. To understand and effectively treat the full clinical spectrum of thoracic malignancies, additional animal models that can recapitulate diverse human lung cancer subtypes and stages are needed. Allograft or xenograft models are versatile and enable the quantification of tumorigenic capacity in vivo, using malignant cells of either murine or human origin. However, previously described methods of lung cancer cell engraftment have been performed in non-physiological sites, such as the flank of mice, due to the inefficiency of orthotopic transplantation of cells into the lungs. In this study, we describe a method to enhance orthotopic lung cancer cell engraftment by pre-conditioning the airways of mice with the fibrosis inducing agent bleomycin. As a proof-of-concept experiment, we applied this approach to engraft tumor cells of the lung adenocarcinoma subtype, obtained from either mouse or human sources, into various strains of mice. We demonstrate that injuring the airways with bleomycin prior to tumor cell injection increases the engraftment of tumor cells from 0-17% to 71-100%. Significantly, this method enhanced lung tumor incidence and subsequent outgrowth using different models and mouse strains. In addition, engrafted lung cancer cells disseminate from the lungs into relevant distant organs. Thus, we provide a protocol that can be used to establish and maintain new orthotopic models of lung cancer with limiting amounts of cells or biospecimen and to quantitatively assess the tumorigenic capacity of lung cancer cells in physiologically relevant settings.

We describe a method to significantly enhance orthotopic engraftment of lung cancer cells into the murine lungs by pre-conditioning the airways with injury. This approach may also be applied to study stromal interactions within the lung microenvironment, metastatic dissemination, lung cancer co-morbidities, and to more efficiently generate patient derived xenografts.

Lung cancer is a deadly treatment refractory disease that is biologically heterogeneous. To understand and effectively treat the full clinical spectrum of ...

Real-Time Monitoring of Cellular Dynamics in Mouse Lung Transplantation

Intravital Two-Photon Microscopy of the Transplanted Mouse Lung

Complications after lung transplantation are largely related to the host immune system responding to the graft. Such immune responses are regulated by crosstalk between donor and recipient cells. A better understanding of these processes relies on the use of preclinical animal models and is aided by an ability to study intra-graft immune cell trafficking in real-time. Intravital two-photon microscopy can be used to image tissues and organs for depths up to several hundred microns with minimal photodamage, which affords a great advantage over single-photon confocal microscopy. Selective use of transgenic mice with promoter-specific fluorescent protein expression and/or adoptive transfer of fluorescent dye-labeled cells during intravital two-photon microscopy allows for the dynamic study of single cells within their physiologic environment. Our group has developed a technique to stabilize mouse lungs, which has enabled us to image cellular dynamics in na&#239;ve lungs and orthotopically transplanted pulmonary grafts. This technique allows for detailed assessment of cellular behavior within the vasculature and in the interstitium, as well as for examination of interactions between various cell populations. This procedure can be readily learned and adapted to study immune mechanisms that regulate inflammatory and tolerogenic responses after lung transplantation. It can also be expanded to the study of other pathogenic pulmonary conditions.

Author Spotlight: Advancing Lung Transplant Immunology Through Intravital Imaging

Here, we present a protocol to intravitally image the transplanted mouse left lung using two-photon microscopy. This represents a valuable tool for studying cellular dynamics and interactions in real-time following murine lung transplantation.

Complications after lung transplantation are largely related to the host immune system responding to the graft. Such immune responses are regulated by ...

A Standardized Method for Measuring Internal Lung Surface Area via Mouse Pneumonectomy and Prosthesis Implantation

Podsumowanie

Przeglądaj więcej filmów

A Simple Method of Mouse Lung Intubation

In vivo Measurement of the Mouse Pulmonary Endothelial Surface Layer

A Permanent Window for Investigating Cancer Metastasis to the Lung

Automated Measurement of Pulmonary Emphysema and Small Airway Remodeling in Cigarette Smoke-exposed Mice

Mouse Pneumonectomy Model of Compensatory Lung Growth

Measurement of the Pressure-volume Curve in Mouse Lungs

Instillation and Fixation Methods Useful in Mouse Lung Cancer Research

Intraperitoneal Glucose Tolerance Test, Measurement of Lung Function, and Fixation of the Lung to Study the Impact of Obesity and Impaired Metabolism on Pulmonary Outcomes

Pre-Conditioning the Airways of Mice with Bleomycin Increases the Efficiency of Orthotopic Lung Cancer Cell Engraftment

Intravital Two-Photon Microscopy of the Transplanted Mouse Lung