The host immune response to pathogen infection is a tightly regulated process. Utilizing a lipopolysaccharide lung exposure model in mice, it is possible to conduct high resolution evaluations of the complex mechanisms associated with disease pathogenesis.
The host immune response to pathogens is a complex biological process. The majority of in vivo studies classically employed to characterize host-pathogen interactions take advantage of intraperitoneal injections of select bacteria or pathogen associated molecular patterns (PAMPs) in mice. While these techniques have yielded tremendous data associated with infectious disease pathobiology, intraperitoneal injection models are not always appropriate for host-pathogen interaction studies in the lung. Utilizing an acute lung inflammation model in mice, it is possible to conduct a high resolution analysis of the host innate immune response utilizing lipopolysaccharide (LPS). Here, we describe the methods to administer LPS using nonsurgical oropharyngeal intratracheal administration, monitor clinical parameters associated with disease pathogenesis, and utilize bronchoalveolar lavage fluid to evaluate the host immune response. The techniques that are described are widely applicable for studying the host innate immune response to a diverse range of PAMPs and pathogens. Likewise, with minor modifications, these techniques can also be applied in studies evaluating allergic airway inflammation and in pharmacological applications.
Pulmonary infections associated with pathogenic bacteria species are a common cause of global morbidity and mortality. Determining the mechanisms that drive the host immune response to these pathogens will promote the development of novel prevention strategies and therapeutic agents that will attenuate the impact of these infections. The overall goal of the protocol described here is to provide the user with a flexible method to evaluate the host innate immune response to pathogen infection using a pathogen associated molecular pattern (PAMP) as a surrogate for live bacteria. The majority of previous studies evaluating the host innate immune response to bacteria have focused on peritoneal models due to the relative ease of execution. While these models are highly useful and have resulted in significant advances in the field of host-pathogen interactions and systemic inflammation, the data generated from these models are not always appropriate for studies involving the respiratory system. Here, a pulmonary model of acute lung inflammation is proposed as a practical and clinically relevant expansion of the classical intraperitoneal (i.p.) injection models. The proposed technique allows for the local assessment of the innate immune response in an organ specific model system.
The methods described here are designed to provide a simple and robust technique to allow users to evaluate the host immune response to LPS, which is a common PAMP. The methods are based on intratracheal (i.t.) instillation of LPS, which induces a robust innate immune response in the lungs of mice and mimics many of the pathophysiological features observed in human patients suffering from respiratory infections and acute lung injury1. A primary advantage of this technique is that it allows the user to evaluate the host immune response without the confounding factors and safety concerns associated with conducting in vivo studies using live bacteria. Likewise, the oropharyngeal i.t. administration route of exposure described in this protocol has significant advantages over other commonly utilized techniques, including intranasal (i.n.) administration and surgical i.t. administration. For example, oropharyngeal i.t. administration allows relatively accurate dosaging and lung deposition compared to i.n. administration, which typically suffers from increased variability of lung deposition due to the loss of agents in the nasal cavity and sinuses2-4. The i.t. administration route circumvents these cavities and allows direct access to the trachea and airway. Likewise, the surgical i.t. approach is a significantly more morbid administration method and requires extensive training to master. The protocols described here also include a description the common techniques and surrogate markers used to evaluate inflammation progression and end with a protocol describing the proper techniques for preparing the lungs for histopathology assessments. These protocols are focused on minimizing the number of mice required for each study by maximizing the data generated from each individual animal.
The protocols described are highly flexible and can be readily modified to evaluate a diverse range PAMPs and damage associated molecular patterns (DAMPs). Furthermore, with a few additional modifications, these protocols can also be applied to studies evaluating allergic airway disease progression or host-pathogen interactions with live bacteria, viruses or fungi5-10.
All studies were conducted under the approval of the Institutional Care and Use Committee (IACUC) for Virginia Tech and in accordance with the National Institutes of Health Guide for the Care and Use of Laboratory Animals.
1. Intratracheal (i.t.) Inoculation of LPS Using Oropharyngeal Administration
2. Serum and Bronchoalveolar Lavage Fluid (BALF) Collection
3. Histopathology Preparation
4. Cytokine Evaluation
5. Differential Staining and BAL Cellularity Evaluation
6. Histopathology Evaluation
The cell walls of gram-negative bacteria are composed of LPS, which is highly abundant in the environment. Inhalation of LPS in sensitive human populations exacerbates airway reactivity and is capable of triggering a robust immune response11. LPS is also a common PAMP used in mouse models to elicit a robust innate immune response. In the protocol described here, the mice received an i.t. dose of LPS isolated from E. coli (serotype 0111:B4) using oropharyngeal i.t. administration. In models of LPS exposure, both body temperature and weight are typical surrogate markers of animal morbidity and disease progression. The LPS challenge will cause a significant decrease in body temperature within the first 6 hr, which will gradually increase back to baseline over the course of 24 hr (Figure 1A). However, body weight will steadily decrease over the course of 24 hr (Figure 1B), where it will peak and gradually recover over the next 48-72 hr. Thus, body temperature is a more appropriate marker to evaluate the early stages of inflammation initiation; whereas, body weight is more reliable during later stages of pathogenesis and recovery.
Airway LPS exposure results in a significant influx of leukocytes to the lungs. Within the first 6 hr, cells associated with the host innate immune response can be observed in the BALF (Figure 1C). The BALF cellularity continues to increase over the next 24-48 hr (Figure 1C), where the immune response peaks and subsequently enters a period of inflammation resolution. By 24 hr, a significant number of neutrophils are present in the lungs and can be observed in the BALF following differential staining (Figures 1D and 1E). BALF cellularity assessments provide a robust and quantifiable technique to characterize the cells associated with the host immune response in the lungs. The increase in BALF cellularity is consistent with a significant influx of neutrophils into the airways, blood vessels and in the lung parenchyma (Figures 1F and 1G). Lung histopathology can be effectively evaluated using a semi-quantitative scoring system (Figure 1F). It is possible to accurately score the histopathology at specific landmarks in the lungs of different animals when the lungs are accurately inflated to the same size with fixative and sections processed to reveal the maximum longitudinal visualization of the intrapulmonary main axial airway. The gravity based inflation protocol, described here, is designed to facilitate this scoring system. LPS induces a significant increase in perivascular, peribroncheolar and parenchymal inflammation (Figure 1G).
LPS administration induces high levels of local and systemic pro-inflammatory mediators, including several cytokines associated with the innate immune response. Local cytokine levels can be assessed in the BALF using conventional techniques, such as ELISA. Common cytokines that are up-regulated in the lungs following LPS administration include TNF-α, IL-1β, and IL-6 (Figure 2A). Systemic cytokine levels can be evaluated in the serum using the same techniques as described for the BALF assessments (Figure 2B). It is not uncommon for some mediators to be present in the local microenvironment, but absent in the serum. For example, TNF-α is routinely found at high levels in the lungs following LPS, but is not typically found systemically under the conditions described in this protocol (Figures 2A and 2B; below the level of detection).
Figure 1. Oropharyngeal Intratracheal LPS Administration Increases Morbidity and Airway Inflammation in Mice. Male mice received 1 mg/kg of E. coli LPS (serotype 0111:B4) i.t. and disease pathogenesis was evaluated over the course of 24 hr. A-B) LPS induces a significant reduction in body temperature within 6 hr of administration; whereas body weight loss is more apparent at the later time points. C) LPS induces a significant increase in total BALF cellularity. D-E) Neutrophils are the dominate cell type present in the lungs 24 hr following LPS administration, as revealed by differential staining from cells isolated from the BALF. These data are typically illustrated as the percent of each cell type present in the BALF. F) Lung histopathology can be evaluated using a semi-quantitative scoring system based on two independent evaluations of the inflammation surrounding the intrapulmonary main axial airway. G) A significant amount of perivascular and peribroncheolar cuffing, mild parenchymal inflammation and some slight alveolar occlusion is typically observed 24 hr following lung LPS exposure. Click here to view larger image.
Figure 2. Airway Exposure to LPS Results in Increased Levels of Local and Systemic Cytokines. A) Following LPS exposure, local levels of a broad spectrum of pro-inflammatory cytokines can be observed in the BALF within the first 24 hr, including TNF-α, IL-1β, and IL-6. These cytokines can be evaluated using ELISA, as shown here 24 hr following LPS exposure. B) Several cytokines can also be detected systemically in the serum. However, there are some notable exceptions, including TNF-α, which are found in high levels in the BALF but not in the serum. Cytokines of interest in the serum should be evaluated empirically prior to large scale evaluation. Click here to view larger image.
Figure 3. Lung Inflation Using Gravity Displacement Results in Reduced Histopathology Variability and Improved Visualization. The gravity displacement method of fixation described in this protocol allows for optimal histopathology evaluation compared to uninflated lungs or manual inflation. A-B) Lungs fixed by gravity inflation demonstrate uniform features allowing for accurate scoring, reduced alveolar damage, enhanced visualization, and higher resolution evaluations of inflammation. C-D) Manual inflation of the lungs typically results in nonuniform areas of inflation. C) These nonuniform areas are often partially inflated, resulting in collapsed areas that are commonly mistaken as pathological features by novice reviewers. D) Manual inflation also results in areas of the lungs that are over-inflated, which results in extensively damaged alveolar spaces. E) Uninflated lungs demonstrate collapsed alveolar spaces and areas that are difficult to resolve and evaluate without extensive training. Likewise, due to the organ being collapsed this technique does not allow visualization of the lungs as they appear in situ. Click here to view larger image.
The most critical steps for successfully evaluating the host immune response in mouse lungs is as follows: 1) choose the appropriate mouse strain and sex for the model being evaluated; 2) optimize PAMP delivery to the lungs; 3) correctly collect and process the BALF; and 4) properly fix and prepare of the lungs for histopathological assessments.
The choice of mouse strain is an important factor in evaluating the host immune response. C57Bl/6 mice are typically considered the optimal mouse background for studying innate immunity due to their Th1 skewing and robust response to most PAMPs. Likewise, BALB/c mice are typically used for studying allergic disease models due to their Th2 skewing. A third commonly used strain in lung models are mice on the 129SvEv background, due to their common use in generating genetically modified animals. In all cases, caution should be taken during experimental design and whenever possible, age and sex matched liter-mate control animals should be used for studies comparing genetically modified mice with wild type animals. Typically, for the LPS protocol described here, 6-12 week old sex matched C57Bl/6 mice should be used and should weigh a minimum of 20 g. It is possible that LPS exposure will result in high levels of morbidity and mortality in sensitive mouse strains or genotypes. If this occurs, it may be necessary to adjust several aspects of this protocol, including reducing the LPS dose, using larger animals, and switching the gender of the animals used in the experiment. Small scale experiments should be conducted initially to determine animal sensitivity and the conditions for each experiment should be based on the most susceptible genotype or experimental condition.
Oropharyngeal i.t. administration has been found to be more accurate than other forms of agent delivery to the lungs2. This is in large part due to the direct access to the airway and circumventing issues associated with nasal and sinus cavity deposition. However, as with all animal procedures, i.t. administration requires extensive manual dexterity and practice to achieve proficiency. Improper technique can result in inefficient lung deposition and in some cases result in animal injury. Typically, Evans Blue Dye (EBD) can be effectively utilized as either a training tool for this procedure or to troubleshoot potential issues associated with deposition4. EBD can be administered i.t. and subsequently extracted from the tissue using formamide. The quantity of EBD can be calculated using absorption levels compared against a standard curve. In our hands, typical EBD recovery ranges between 90-98%. The bulk of the unrecovered dye is expected to be associated with leakage into the esophagus. Due to its accuracy, the i.t. administration technique is ideal for delivering a diverse range of dose sensitive agents to the lungs, such as pharmaceutical agents or infectious organisms.
Analysis of the BALF and BALF cellularity can provide a significant amount of insight into the overall progression of the host immune response. Inflammatory mediators released locally in the lungs following stimulation can be effectively quantified in the BALF using common immunology techniques, such as ELISA and Western Blot. Likewise, the cellular composition of the BALF can be evaluated using either differential cell staining or flow cytometry. Developing the proper technique and manual dexterity is the most critical part of performing the bronchoalveolar lavage (BAL). One of the most common issues that occur during the BAL is failure to fully withdraw the maximum volume of fluid originally placed in the lungs. This is commonly associated with an improperly secured cannula or when needles are used in place of actual cannulas. Specialized tracheal cannulas are commercially available. It is also important that the motion and force used to insert and withdraw the saline is smooth and consistent throughout the procedure. The protocol described here has been optimized for differential staining of cells collected in the BALF. Differential staining is a modified Wright Giemsa stain and is a highly effective technique to conduct morphology based cell identification. This staining technique allows for the differentiation of neutrophils and eosinophils based on their unique granule staining properties. Monocyte derived cells are also easy to identify and are commonly observed in the BALF. These include macrophages and dendritic cells. Likewise, T-cells and B-cells are also commonly observed. However, these monocytic cells and lymphocytes are often difficult for most researchers to accurately differentiate based on morphology alone. The utilization of flow cytometry can add much higher resolution to these evaluations. However, low cell numbers recovered from control animals is often a limiting factor. Thus, if flow cytometry is to be used, the experimental design should include additional negative control animals to increase cell recovery.
Lung histopathology assessments are another critical component of this protocol and allow for the direct visualization of disease progression (Figures 3A and 3B). When the lungs are properly prepared, histopathology can be accurately evaluated, quantified, and characterized. The most critical step in preparing the lungs for histopathology is properly inflating them with fixative. Manual methods of inflation typically result in lungs that are over-inflated, under-inflated or partially-inflated, which results in suboptimal visualization and morphology assessments (Figures 3C and 3D, respectively). Likewise, lungs that are not inflated prior to fixation are very difficult to accurately evaluate and often results in highly variable histopathology scores (Figure 3E). The gravity method of inflation discussed here provides a highly reproducible method of inflation with minimal variability. Using this technique, it is possible to evaluate specific landmarks in the lungs that are consistent between experimental animals. Furthermore, gravity inflation using an inflation stand has been shown to inflate the lungs at a fixative pressure of 20 mm12. This technique allows the visualization of the lungs in their most physiologically relevant size and shape and has been shown to be highly effective for the evaluation of sensitive pathophysiological processes12. It is critical that the inflation stand be set at the proper height from the mouse to generate the proper pressure. If suboptimal inflation is observed, the height of the inflation stand should be determined empirically. The use of a commercially available small rodent lung inflation stand, which will ensure the proper height, is recommended.
This procedure has been optimized for the delivery of LPS and other PAMPs to the lungs of mice. Once these techniques are mastered, additional studies can also be initiated using modified protocols to effectively evaluate host-pathogen interactions using live pathogens. Likewise, the techniques described here are highly versatile and can be applied to any study interested in assessing the clinical and physiological relevance of an experimental or pharmaceutical agent. Because of the accuracy in the dosing, this technique is also ideal for in vivo studies that require a high level of precision in agent delivery.
This procedure has been optimized for the delivery of LPS and other PAMPs to the lungs of mice. Once these techniques are mastered, additional studies can also be initiated using modified protocols to effectively evaluate host-pathogen interactions using live pathogens. Likewise, the techniques described here are highly versatile and can be applied to any study interested in assessing the clinical and physiological relevance of an experimental or pharmaceutical agent. Because of the accuracy in the dosing, this technique is also ideal for in vivo studies that require a high level of precision in agent delivery.
The authors have nothing to disclose.
The authors thank the VA-MD Regional College of Veterinary medicine for providing core and technical support for this project. This work is supported by an NIH Career Development Award (K01DK092355).
C57Bl/6J | The Jackson Laboratory | Stock 000664 |
Compact Scale | Ohaus Scale Corporation | 71142845 |
Small Animal Rectal Thermometer | Braintree Scientific | TH 5 |
Rectal Probe for Rodents | Braintree Scientific | RET 3 |
Ear Punch | Braintree Scientific | EP-S 901 |
Lipopolysaccharide from E. coli 0111:B4 | InvivoGen | LPS-EB |
1x Phosphate Buffered Saline | Life Technologies | 10010-023 |
Isoflurane | Baxter | 40032609 |
Intratrachael Administration and Lung Inflation Stand | ICAP Manufacturing | n/a |
Rodent Intubation Stand | Braintree Scientific | RIS 100 |
Scissors (blunt/sharp) | Fisher Scientific | 13-806-2 |
forceps (straight) | Fisher Scientific | 22-327-379 |
forceps (45º, curved) | Fisher Scientific | 10-275 |
Scissors (blunt/blunt) | Fisher Scientific | 08-940 |
Pipette (200 µl Capacity) | Gilson | F123601 |
Ethanol | Sigma | 459844 |
1 ml Syringe | BD Medical | 301025 |
10 ml Syringe | BD Medical | 301604 |
27 G x 0.5 in. needle | BD Medical | 305109 |
Refrigerated Microcentrifuge | Fisher Scientific | 13-100-676 |
1.2 mm Tracheal Cannulae with Luer-adapter | Harvard Apparatus | 732836 |
Hank's Balanced Salt Solution | Life Technologies | 14025-076 |
4-0 Silk Braided Surgical Suture | Ethicon | A183 |
Luer to Tube Connector Kits | Harvard Apparatus | 721406 |
Luer Stopcock Kit | Harvard Apparatus | 721664 |
Tygon formula E-3603 laboratory tubing | Sigma | R-3603 |
Formalin solution, neutral buffered, 10% | Sigma | HT501128-4L |
Mouse IL-1β OptEIA ELISA Kit | BD Biosciences | 559603 |
Mouse IL-6 OptEIA ELISA Kit | BD Biosciences | 550950 |
Mouse TNF-α OptEIA ELISA Kit | BD Biosciences | 560478 |
Hemacytometer | Hausser Scientific | 3520 |
Hemacytometer Cover Glasses | Thermo Scientific | 22-021-801 |
Trypan Blue | Thermo Scientific | SV3008401 |
Cytology Funel Clips | Fisher Scientific | 10-357 |
Cytology Funels | Fisher Scientific | 10-354 |
Filter Cards | Fisher Scientific | 22-030-410 |
Microscope Slides | Fisher Scientific | 12-544-1 |
Cover Glasses | Fisher Scientific | 12-540A |
Cytospin Cytocentrifuge | Thermo Scientific | A78300003 |
Diff Quick Staining Kit | Fisher Scientific | 47733150 |
Permount Mounting Medium | Fisher Scientific | SP15-500 |