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This unit describes protocols for infecting the mouse respiratory tract, and assaying virus replication and host response in the lung. Respiratory infections are the leading cause of acute illness worldwide, affecting mostly infants and children in developing countries. The purpose of this unit is to provide the readers with a basic strategy and protocols to study the pathogenesis and immunology of respiratory virus infection using the mouse as an animal model. The procedures include: (i) basic techniques for mouse infection, tissue sampling and preservation, (ii) determination of viral titers, isolation and analysis of lymphocytes and dendritic cells using flow-cytometry, and (iii) lung histology, immunohistochemistry and in situ hybridization.
The respiratory tract is remarkable for its extensive surface area (70 m2 in adult humans) which is in continuous contact with the external environment. The lung samples approximately 10,000 L of air every day and is exposed to a vast array of foreign particles. As a consequence, the respiratory system is a major portal by which microorganisms enter the body. When studying infection and immunity in this tract, the structurally and functionally distinct compartments must be appreciated as each compartment has distinct populations of both immune and parenchymal cells. The nasopharynx, or uppermost airway, is lined by both respiratory and olfactory epithelium. The respiratory epithelium of the mouse nasopharynx is columnar to cuboidal with cilia, and scattered goblet cells. The tracheal lining is similar but, unlike the human trachea, has a high percentage (50–60%) of non-ciliated, secretory Clara cells. The right lung of the mouse is divided into 5 lobes; the left lung is not divided. The lower respiratory tract is comprised of the branching conducting airways, which extend from the trachea to the terminal bronchioles, which open into the alveolar ducts. The bronchi and bronchioles of the mouse are lined predominantly by Clara cells, which are relatively rare in the human airway (Sternberg, 1997). This altered cell distribution must be kept in mind as it will alter the pattern of infection with some viral respiratory pathogens. Goblet cells are not found in the uninfected mouse lung. The bulk of the lung volume is made up of alveoli where gas exchange occurs. The alveolar ducts and spaces are lined primarily by flat type I pneumocytes overlying capillaries. The rare, plump type II cells produce surfactant and are thought to be the source of new type I cells (Maronpot et al., 1999).
Because efficient gas exchange requires that the alveoli be relatively free of fluid and inflammatory cells, the respiratory mucosal immune system must maintain a fine balance between eliminating pathogens and inhibiting inflammatory pathology. This requires discriminating between innocuous airborne antigens and pathogen-associated antigens, and inducing tolerance or immunity, respectively. Surveillance is maintained by a layer of dendritic cells underlying the mucosa of the larger conducting airways (Vermaelen and Pauwels, 2005; Weslow-Schmidt et al., 2007), and alveolar macrophages present within the air spaces of the lung. Additional inflammatory cells are recruited by chemokines secreted by epithelial and immune cells when pathogens are detected. The mouse model is permissive for some but not all human respiratory pathogens, and among those viruses which will infect both mouse and man, disease in the mouse may not necessarily mirror disease in the human host. This is an important consideration when undertaking infection experiments to model human diseases in the murine host.
This unit describes protocols for infecting the mouse respiratory tract, and assaying virus replication and host response in the lung. The reader is reminded that all procedures involving viral pathogens and infected tissues must be carried out in a biosafety cabinet prior to fixation.
Most intranasal infection protocols use relatively large volumes (30–100 µl) of inoculum, resulting in intratracheal delivery of virus. In this section we will discuss intranasal inoculation using both small and large volumes. Choice of route will vary with the virus studied, and the interest of the investigator in upper versus lower airway infection.
Avertin anesthesia (See REAGENTS section)
1 ml syringes and needles (25G2/8 to 27G) (BD)
Viral inoculum diluted in PBS or HBSS (Cellgro) to a volume of 30–100 µl/dose
Positive displacement pipet and tips (Rainin)
The use of a positive displacement pipette prevents aerosol contamination and isolates the virus sample from the pipette body. This precaution is extremely important if different virus mutants, strains or virus species are being used. You can change the titer of the inoculum by altering the amount of stock virus used and/or by adjusting the volume of diluent accordingly. We normally use volumes between 30 and 100 µl for mice 6 weeks of age or older, smaller volumes may be needed for younger animals.
Let the mouse recover for 5–10 minutes. It will have an increased respiratory rate for some minutes after intranasal inoculation. If a proper dose of anesthesia has been used, the mouse will regain consciousness and breathing will return to normal within 15 – 30 minutes.
This is a modification of the above intranasal infection. In this method, an inoculum is delivered specifically to the upper airway by using a smaller volume and administering the inoculum over several minutes (Gitiban et al., 2005; Visweswaraiah et al., 2002). Materials required are listed above except that the viral inoculum will be prepared such that each dose is contained within a 20 µl. volume. A timer is also required for this protocol.
The optimal method for determining virus titer will depend upon the virus you are using. In this section we outline generally applicable techniques for obtaining samples for testing, and examples of titering methods used for two viruses commonly used in our laboratory, RSV and influenza A virus.
2L nalgene beaker
Styrofoam insert (3–5 cm. in thickness, equal to the beaker in circumference)
70% ethanol or water
Dissecting scissors, forceps
Sterile plastic snap-cap tubes, preweighed, one for each sample
This primitive CO2 generation chamber is convenient as it can be sterilized after use, is portable, and can be placed within the biosafety cabinet.
Lung tissue (and adjacent lymph nodes if desired) harvested in this manner can be used for many purposes, but must be processed or frozen quickly to maintain its integrity. For determination of virus titers or cytokine levels, lungs should be placed in a sterile, plastic snap-cap tube (without liquid) and frozen immediately on dry ice. These samples can later be transferred to a −80°C freezer for storage.
Tissue homogenizer (E.g. PowerGen model 125, Fisher Scientific)
Decarbonated Dulbecco’s Modified Eagle Medium (dcDMEM).
It is important that samples not undergo repeated freeze-thaw cycles before virus titers are determined. You may homogenize you samples the day they are collected, then freeze (−80°) for assay at a later time. Or, freeze the samples as they are being collected, preparing homogenates and then setting up the plaque assay the same day. Viruses differ with respect to their ability to tolerate freezing, some pathogens may lose as much as a log of infectivity with each freeze-thaw cycle.
It is important to assure that biologically active organisms and molecules be reliably eliminated from all surfaces that will contact subsequent samples. In processing any significant number of samples, sterilization by autoclaving is impractical. The procedure described below will clean the apparatus and destroy any trace residual biologically active protein, nucleic acid or lipid.
Sterile 50 ml tissue culture tubes
0.1% SDS, 0.006 M sodium hypochlorite (commercial chlorine bleach diluted 1:100).
0.1% SDS, 0.001% coomassie blue
Run the homogenizer with the following solutions, taking care that the liquid covers all surfaces that will potentially contact subsequent samples.
Any virus recovered from mouse respiratory tract will, presumably, replicate to some degree in mouse cells. Different viruses, however, vary widely in the degree to which they produce “plaques” (countable lesions) on a cell monolayer. RSV, for instance, normally produces no obvious cytopathology in cultured mouse cells, and has historically been assayed on monolayers of human HEp2 cells or monkey Vero cells.
This procedure is for RSV; specifics will vary for other viruses.
1% Methylcellulose in MEM
10× Minimal Eagle’s Medium (Invitrogen)
10× Earle’s Balanced Salt Solution (Sigma-Aldrich)
Susceptible cell line
24 well plates
It is important that the dilution series be done as accurately as possible, since systematic errors will be amplified exponentially by the process of serial dilution. For this reason, care must be taken to avoid bubbles, and pipet tips changed between transfers. Care should be taken that the pipet tips should not carry any extra liquid as drops clinging outside of the pipet tip. HEBSA should be at room temperature, since a temperature differential between pipet tip and medium introduces a systematic inaccuracy. A multichannel pipettor can be used, but the above listed cautions must be observed with respect to each of the pipet tips at each step!
*In all these steps, it is important to prevent the monolayers from drying out. Aspirate with a minimum of suction; avoid exposing the cells to rushing air; remove/replace medium from no more than a few wells at a time.
The concentration of infectivity (commonly, but not quite correctly, referred to as “titer”) is estimated as:
(Number of plaques/well / Volume of inoculum/well) × Dilution.
E.g., the well inoculated from the 4th well in the dilution series above shows 35 plaques.
Inoculum volume = 0.100 ml
Dilution = 104
Concentration of infectious virus = (35/0.100)×104 = 3.5 × 106.
The standard error is estimated as the ratio as the square root of the total number of plaques counted. The relative standard error (i.e. expressed as a fraction of the concentration) is therefore the inverse of the square root of the number of plaques counted. In the above example, the standard error is (35)−1/2 = 16.9% If duplicate wells had been inoculated, yielding 34 and 36 plaques, for example, the estimated concentration would be the same, but the error would be reduced to (70)−1/2 = 12.0%
Many viruses (e.g. influenza virus, parainfluenza virus 5, Newcastle disease virus) do not produce visible plaques. Estimation of the infectivity of such viruses can be done by following the procedure given above, but staining the infected monolayers with fluorescent or enzymatically tagged antibodies. The procedure for immunofluorescent detection is outlined below. The procedure for enzyme-linked antibodies is similar, but details vary with the specific enzyme used.
Inverted fluorescent microscope
Primary antibody against viral antigen (primary antibody, e.g. rabbit anti-influenza)
Fluorophore-tagged secondary antibody (e.g. FITC-goat anti-rabbit IgG)
This section outlines the procedures for sampling and processing mouse tissues to obtain single cell suspensions that can be used for flow cytometry or other assays (ELISpots, migration assays, proliferation assays, and purification of cell subsets). The method used to process the organs differs depending on the cell population of interest. For the analysis of T cell lymphocytes the authors use mechanical digestion which is quick and convenient. For the analysis of dendritic cells, collagenase digestion is recommended. This procedure is necessary to separate dendritic cells form the extracellular matrix, and is followed by a short incubation in an EDTA solution, which helps to disrupt multicellular complexes.
Assorted syringes and needles (BD)
Terumo Surflo i.v. catheter 186 × 11/4”
15 ml and 50 ml centrifuge tubes (BD)
Serological pipettes (Costar)
Pipet-Aid (Drummond Scientific)
Tissue culture dishes (Falcon)
70 um cell strainers (BD Falcon)
HBSS no MgCl2, MgSO4, or CaCl2 (Cellgro)
Centrifuge (such as Sorvall Legend RT)
Hemacytometer (Hausser Scientific)
Trypan blue (MP Biomedicals)
Microscope (such as Zeiss Axiostar plus)
You can draw blood from the heart by sticking the needle of a 1 ml syringe into the apex of the left ventricle or by axillary bleed using a Pasteur pipet. Transfer the blood into a tube containing 5 ml HBSS/heparin.
With your forceps, hold of the skin of the neck and cut along the neck from the chin to the thorax. Expose the trachea and make a small incision using sharp scissors or a needle with a beveled edge. Insert only the plastic sleeve of a catheter in the trachea, and using a 1 ml syringe flush lungs 3 times with 1 ml HBSS. Repeat twice more using a total of 3 ml HBSS. Work slowly to avoid collapsing the lungs. As a general rule, you will pool the lavage from the mice that belong to the same experimental group due to the limited number of lymphocytes found in the airways. If the BAL specimen is to be used for cytokine assays, use only 1 ml of lavage fluid to maximize the concentration of mediators present.
Cut the skin of the mouse from the abdomen to the top of the thorax. Open the abdominal wall below the ribcage. Lift the sternum with tweezers and cut the diaphragm. Then cut away the lower part of the ribcage to expose the heart and lungs. Using fine tweezers harvest the lymph nodes. The mediastinal lymph nodes are located ventral to the trachea at the level of the thymus. The bronchial lymph nodes are situated at the bifurcation of the trachea. Next, harvest the lung. Place each organ in a labeled tube containing 5 ml HBSS.
If you are interested in analyzing lymphocytes a simple mechanical disruption of the tissues is enough. The spleen, lymph nodes and lung are gently pressed through a cell strainer mesh with a syringe plunger to achieve single cell suspensions. The strainers are repeatedly rinsed with HBSS to recover as many lymphocytes as possible. If you are interested in analyzing dendritic cells you will need to perform an enzymatic digestion of the organs. We mince the organs in a solution of 5 mg/ml collagenase A in MEM in a small dish and incubate them for 30 min at 37°C (see support protocol). The BAL samples do not need processing. All the samples are transferred to 15 ml conical tube if they are not already in one.
We use 5 ml for spleen and 1 ml lymph nodes and BAL. Lung cells need to be further purified using a gradient. Thus, lung cells are resuspended in 1.0 ml. of 80% Percoll (see support protocol 1).
Samples with low cell numbers such as BAL and lymph nodes have to be counted using the hemacytometer. This is a tedious and time consuming process, but it is essential in order to calculate absolute numbers of cells in each tissue using the frequencies obtained after flow cytometry analysis.
Percoll solutions cannot be filtered, so be extra careful and keep them sterile. We use the same gradient to process liver cells.
(in addition to basic protocol 3)
Percoll (GE Health Care)
10× HBSS no MgCl2, MgSO4, or CaCl2 (Cellgro)
HBSS no MgCl2, MgSO4, or CaCl2 (Cellgro)
(in addition to basic protocol 3)
Collagenase A solution
HBSS/2 mM EDTA solution
If you need to digest lymph nodes, transfer the organs to a 5 ml to a FACS tube with 200 µl collagenase solution, smash it up with a 1 ml syringe plunger, and incubate as indicated above.
This section describes a basic protocol for the staining of tissue lymphocytes to analyze antigen-specific CD8 T cells using flow cytometry. CD8 T cell responses are key for the control of viral infections (Doherty and Christensen, 2000; Wong and Pamer, 2003). The T cell receptor (TCR) on the surface of CD8 T cells recognizes peptides derived from viral antigens complexed with major histocompatibility complex (MHC) class I molecules on the surface of infected cells. CD8 T cells express a broad range of effector mechanisms that mediate resistance to infection: (i) direct cytolysis of target cells mediated by perforin, granzymes and Fas, (ii) secretion of cytokines such as TNFα and IFN-γ and (iii) secretion of chemokines that attract inflammatory cells at sites of infection. A prominent characteristic of respiratory virus infections, such as those mediated by influenza or respiratory syncytial virus, is that viral replication is limited to the lung epithelium. Thus, for viral clearance, the immune system must specifically target the mucosal surfaces of the respiratory tract. To accomplish this, antigen presenting cells migrate to the draining lymph nodes where they initiate a program of T cell proliferation and differentiation (Legge and Braciale, 2003; Vermaelen et al., 2001). Newly generated effector T cells return to the respiratory tract, complete their differentiation process, and terminate the infection (Cerwenka et al., 1999; Lawrence and Braciale, 2004). The effector CD8 T cells and memory cells that remain in the lung once infection is cleared are functionally different from their counterparts in lymphoid tissues (Hikono et al., 2006).
Fluorescence-activated flow cytometry is one of the most broadly used technologies in cell biology, immunology, hematology and oncology. It allows the simultaneous and rapid analysis of multiple parameters of each cell in a suspension. This technology provides the means to define and analyze specialized subpopulations of cells that may carry out distinct functions. In addition, the availability of MHC class I-tetrameric reagents (Altman et al., 1996) which can label epitope-specific CD8 T cells allows us to analyze antigen-specific T cells at the single cell level. Immunologists now routinely use “tetramers” to track and analyze T cell responses with a level of detail that was unimaginable 15-years ago.
Antibodies: Fc-Block (anti-mouse CD16/CD32 Fc II/III), anti-CD8α (FITC, Alexa 700, APC), anti-KLRG1 FITC (eBiosciences or BD Biosciences)
Tetrameric reagent: γHV68 ORF6487–495 /Kb APC (NIH Tetramer Core Laboratory)
Staining Wash Buffer (SWB)
1.1 ml microtubes (National Scientific Supply Co.)
The flow cytometry analysis on Figure 1 shows antigen-specific CD8 T cells gated using the tetrameric reagent ORF6487–495 /Kb, a murine γ-herpesvirus 68 (γHV68) peptide recognized by CD8 T cells when complexed with the MHC Class I allele Kb. This gate is used to plot histograms showing the expression profile of KLRG1 (killer lectin receptor G1). KLRG1 is a killer inhibitory receptor that is used to define terminally differentiated short-lived effector T cells (SLECs). As observed, the distribution of SLECs is different in lung and spleen: the majority of SLECs concentrate in spleen because by day 14 after γHV68 infection acute respiratory infection has been cleared and the virus infection is at the peak of latency phase establishment in the spleen in B cells, macrophages and dendritic cells (Flano et al., 2000).
Dendritic cells are professional antigen presenting cells essential for the generation of adaptive immune responses. Dendritic cells develop from bone marrow-derived precursors and are recruited from the blood circulation to peripheral organs, where they continuously sample their environment for foreign substances. Dendritic cells are found through the airways forming a network through the epithelium and also constitute a small fraction of the cells in the BAL (Vermaelen and Pauwels, 2005). They are able to take up and process antigens and migrate to the draining lymph nodes where they contribute to the initiation of adaptive immune responses. Pulmonary dendritic cells are key regulators of the local immune response to airborne pathogens, antigens and allergens (Holt et al., 2008).
In mice, the dendritic cells in the respiratory tract express high levels of MHC class II and CD11c (Wikstrom and Stumbles, 2007). They can be distinguished from the lung macrophages because dendritic cells are less autofluorescent and express higher levels of class II molecules on their cell surface (Jakubzick et al., 2006). As their secondary lymphoid organ counterparts, respiratory dendritic cells can be divided into two main subsets: conventional dendritic cells (CD11c+B220−) and a very small fraction of plasmacytoid dendritic cells (CD11c+B220+) (Weslow-Schmidt et al., 2007). Conventional respiratory dendritic cells can be further divided into two subsets on the basis of CD11b expression, with the predominant subset expressing high levels of CD11b. None of these subsets has been characterized extensively, but there is some evidence to suggest that they are functionally and phenotypically distinct (Belz et al., 2004; Julia et al., 2002; von Garnier et al., 2005). Other dendritic cell markers such as CD103, CD205 or CD8α can also be used to further define these populations.
Figure 2 presents a comparison between dendritic cells from spleen and lung stained with the same panel of fluorochrome-conjugated antibodies. The staining procedure is performed as described in the basic protocol 4 eliminating the steps 6 to 9 that are specific for the tetramer staining. As dendritic cells are a minor fraction of the total cells, the authors staining panel uses a dump channel with a mixture of antibodies (CD3, CD19 and NK1.1) to gate out lineage-positive cells (T cells, B cells and natural killer cells, respectively). The level of class II expression helps to differentiate dendritic cells from pulmonary macrophages. Then, conventional dendritic cells (CD11c+) are plotted against plasmacytoid dendritic cells (B220) and further analyzed using expression of CD11b versus CD8α. CD11b expression defines a subpopulation formerly known as “myeloid” dendritic cells and CD8α is expressed on “lymphoid” dendritic cells. The distribution of the subpopulations of dendritic cells is different between lung and spleen.
Respiratory dendritic cell staining antibody panel:
Examination of infected tissues under the microscope allows the investigator to assess many aspects of inflammation and disease that cannot be discerned from tissue homogenates. Standard hematoxylin and eosin (H&E) stained sections provide information about trafficking and localization of inflammatory cells, as well as the extent of necrosis/apoptosis.
2L nalgene beaker
Styrofoam insert (3–5 cm. in thickness, equal to the beaker in circumference)
70% ethanol or water
Dissecting scissors, forceps
10% buffered formalin (volume ≥ than 10x the volume of harvested tissues) in a closed container (Fisher Scientific)
5 ml. syringe and needles (25G2/8 to 27G) (BD)
Tissue cassettes (Fisher Scientific)
No. 2 pencil
Extended formalin fixation will inhibit binding of both antibodies and nucleic acid probes to tissue sections. Once fixed tissue is processed and embedded in paraffin, it is stable for many years at room temperature.
It is best to rely on the histology laboratory for the standard H&E stain as it requires many solutions that must be renewed often.,
An example of an H&E stained lung section is shown in the left-hand panel of Figure 3 and demonstrates the anatomy of the lung infected 24 hours earlier with influenza A virus. At this low magnification (photomicrograph is taken through a 5× objective) the anatomy of the lung can be clearly seen with large airways branching into smaller airways, and finally opening into the alveolar ducts and space. Very little inflammation can be appreciated at this time point.
You may wish to characterize the different types of inflammatory cells within the airspaces of the lung, and this can be done by using a portion of the BAL fluid, obtained as described above, to make cytospins.
Cytocentrifuge (Thermo Scientific)
Cytoslides (Thermo Scientific)
Disposable cytofunnels (Thermo Scientific)
Acetone (Fisher Scientific)
Immunohistochemistry relies upon the availability of specific antibodies which can bind with high affinity to antigens in tissue sections. Basically, this technique involves incubating tissues with a primary, antigen-specific antibody, followed by a conjugated secondary antibody that will bind to the Fc region of the primary antibody. For example, when mouse IgG is used to stain human tissue, a biotinylated secondary antibody recognizing mouse IgG can be used to detect the bound primary antibody. Avidin bound to horseradish peroxidase will specifically bind to biotin, and when chromogen is added it will be deposited at sites where primary antibody is bound.
4 µM tissue sections on Plus slides (Surgipath)
Coplin jars (Fisher Scientific)
3% H202 (McKesson)
Super sensitive wash buffer (Biogenex)
Primary antibody (determined by investigator)
Primary Antibody Dilution Buffer (Biomedia/Fisher Scientific) or use TBS with 1% BSA (Fraction V, Fisher Scientific) (w/v) as an antibody dilution buffer
Pap pen (Biocare Medical)
Biotinylated Secondary Antibody (determined by investigator)
Streptavidin-enzyme and substrate kits (Scytek or Dako)
Mayer’s Hematoxylin (Dako)
0.25% ammonium hydroxide
Permaslip (Alban Scientific)
This protocol is the most flexible, allowing you to use many different sources of primary and secondary antibody. Nonetheless, a more sensitive methodology is coming into use where, instead of a biotinylated secondary antibody and enzyme-conjugated avidin, an enzyme-linked polymer conjugated to a secondary antibody is used. The EnVision reagents from DAKO make use of this system, and although compatible reagents are not yet available for all primary antibodies, enhanced foreground and decreased background are routinely obtained with this technique.
Here we include a number of approaches to increasing the signal obtained with a particular primary antibody. As reagents are often limited when working with rodent models, it is often necessary to optimize results obtained with the antibodies available. For a detailed discussion of options available it is best to consult a manual devoted to the subject (Renshaw, 2007)
If a primary antibody gives a poor signal using the standard IHC protocol outlined above, “antigen retrieval” using a pressure cooker to heat the tissue sections may increase the foreground.
Coplin jars (Fisher Scientific)
0.5 M Tris at pH 10
Pressure cooker (any)
These conditions work for many antigens, but some will require less heat or more acidic conditions for optimal unmasking. Optimal conditions for a particular antigen must be worked out empirically.
Sometimes it is possible to use monoclonal antibodies to stain frozen tissues even when they are unable to detect antigen in fixed tissue sections.
5 ml syringe (BD)
Terumo Surflo i.v. catheter 186 ×11/4”
Plus Slides (Surgipath)
Tissue Tek Cryo-OCT embedding compound (Fisher Scientific)
Coplin jar (Fisher Scientific)
Acetone (Fisher Scientific)
Cryostat “chuck” (obtained from cryostat manufacturer)
This method describes the use of a non-radioactive RNA probe to detect mRNA in paraffin-embedded tissue sections. For this procedure you will need a probe that is labeled with digoxigenin (DIG). We use the DIG RNA Labeling Kit (Roche) to make the riboprobe and test its labeling efficiency. The following procedure takes two days to complete. For the first day all reagents and containers must be RNase free. Either treat all solutions with DEPC or make up the solutions with DEPC-treated water.
Coplin jars (Fisher Scientific)
Pap pen (Biocare Medical)
Diethylpyrucarbonate (DEPC)-treated water
100 mM glycine (Fisher Scientific)
Proteinase K (Sigma)
4% paraformaldehyde (Electron Microscopy Sciences)
100 mM Triethanolamine pH 8
Acetic Anhydride (Sigma)
Digoxigenin labeled riboprobe
ISH Buffers 1, 2, 3
Newborn calf serum
Anti-DIG alkaline phosphatase Fab fragment (Roche)
NBT/BCIP tablets (Roche)
Fast green FCF (Fluka)
Add the acetic anhydride to the TEA buffer just before use, and use a clean coplin jar for the 2nd incubation.
Avertin (tribromoethanol 20 mg/ml) is a short-acting anesthetic that is not available commercially and must be prepared. Sterile preparation procedures are essential. If Avertin is improperly prepared or stored in the light or at room temperature, it will break down within 24 hours into dibromoacetic acid and hydrobromic acid which are lethal. Freshly mixed solutions are strongly recommended for safe use. The solution can be kept as long as 4 months if it is stored in the dark at 4°C. The solution should be tested to ensure that it has a pH >5.
Glass Erlenmeyer flask
Hot plate with stirrer
Bottle-top filter 0.22 um (Millipore)
Work in the fume hood.
Add 10 ml tert-amyl-OH into glass bottle
Add 10 g 2,2,2-tribromoethanol to the alcohol
Stir until the solid goes into solution room temperature
Transfer to a hot plate on low heat
Very slowly add 250 ml distilled water to flask while stirring. Make sure that salts stay in solution before you add more water. This process takes several hours.
Bring solution up to 500 ml with distilled water.
Turn off the hot plate when all of the solid has gone into solution. Cover flask with aluminum foil to protect from the light, and stir overnight.
Check that the 2,2,2-tribromoethanol has not precipitated overnight. If necessary, return the flask to the hot plate and stir again on low heat until the solid has gone back into solution.
Adjust pH if necessary.
Bottle-top filter (0.22 µm).
Aliquot into 50 ml glass bottles, of darkened glass or foil-covered.
Store in the refrigerator.
Test it on mice.
The effective dose in mice is on the order of 25 mg/kg (approx 200–250 µl/animal) but this will vary by strain with 129 mice being very susceptible and requiring a lower dose, and C57BL6 mice requiring a higher dose. The optimal dose is best decided empirically with each new batch of Avertin made, and each mouse strain used.
The standard formulation of DMEM is buffered with bicarbonate, and therefore becomes alkaline when exposed to ambient atmosphere. For this reason, the medium used for the extended manipulations called for in these protocols is not supplemented with bicarbonate (Invitrogen). Adjust pH to neutral with sodium hydroxide).
Hybridization solution can be made up without salmon sperm DNA and stored at −20°C. When making up the hybridization buffer, dissolve the dextran sulfate in the water and Formamide before adding the other components.
Mix equal volumes of growth medium (DMEM + 25mM HEPES +5% Fetal calf serum + antibiotics*) and 1% methylcellulose in MEM. Mix thoroughly by repeated pipetting, to eliminate pockets of low viscosity.
Since the source of the virus – a live animal – cannot be “sterile”, there is a possibility of bacterial and fungal contamination. Antibacterial agents (e.g. penicillin/streptomycin) and antifungal agents (e.g. amphotericin B) may be added to prevent these from overgrowing the cultures.
Heat ddH2O in glass beaker with stirrer on hot plate to 60°C in fume hood.
Add paraformaldehyde powder and keep at 60°C.
Add 2 drops 2N NaOH, and the solution should clear within a few minutes.
Remove from heat and let cool to room temperature.
Add 20ml 10× PBS and bring volume up to 200ml with ddH2O.
Filter and store at 4°C.
The study of virus infections in vivo requires the analysis of many parameters by a large number of different methods. Because a single unit cannot encompass all of these methods in detail, we have focused on providing the reader with protocols for harvesting the relevant samples. Our suggested methods for detection of infectious virus, flow cytometry, and histology are given as examples of how the samples might be assayed. The assay techniques outlined here are relatively general, as we have chosen the most widely applicable techniques and reagents. It is up to the individual investigator to tailor these assays to meet their specific needs.
Although not explicitly described in this chapter, cytokine assays can be carried out using either BAL specimens (described in basic protocol 4) or lung homogenates. The same homogenate can be used to assay both virus titers and cytokines, although we have found that BAL samples generally have a lower background for cytokine measurement than do whole lung homogenates. ELISA assays are commercially available for many mouse cytokines and chemokines. A biological assay for IFN-α or –β activity is described in detail by Vogel et al. in Current Protocols in Immunology (Vogel et al., 1991), and this method can be used to assay lung BALs and homogenates, as well as serum samples.
Successful flow cytometry analysis of a given population requires the knowledge and experience to calibrate the instrument, standardize procedures and settings, and to analyze the data with a software package. However, the most important requirements are a good single cell suspension and a proven staining panel/protocol. The correct choice of reagents requires certain knowledge of the characteristics of each fluorochrome and of the antigens of interest. However, the basic approach described here can be used for multiple extracellular antigens, cell subpopulations and tissues with minor modifications.
High quality H&E sections are best obtained from a pathology core or hospital laboratory as production of formalin-fixed, paraffin-embedded tissue ‘blocks’ and slides from these blocks require a major investment in equipment and training which is not appropriate for most research laboratories. Nonetheless, if fixed tissue samples are submitted to the histotechnologist in tissue cassettes, taking care to explain the orientation needed for optimal slide interpretation, high quality H&E stained sections can easily be obtained. Additional sections for immunohistochemistry (IHC) or in situ hybridization (ISH) studies can also be prepared from these tissue blocks. IHC using virus-specific antibodies allows the investigator to visualize the extent of virus spread, and the cell type(s) targeted by a particular virus. This method can also be used to visualize the appearance of immune cell types in infected tissues over time, although this type of experiment may require the use of unfixed, frozen tissue sections. ISH is an alternative method if specific antibodies are not available, or the investigator wishes to determine the source of a secreted protein.
For human tissues there are a large number of commercially available monoclonal antibodies that can serve this purpose, and work well for staining formalin-fixed, paraffin-embedded tissue sections. Unfortunately this is not the case for mouse tissues, as there is a relatively small demand for these reagents. Many monoclonal antibodies that work well for staining cell suspensions prior to FACS analysis will not work well for staining tissue sections. The company selling the antibody may specify this, but usually this must be determined empirically. In some cases this can be overcome by using frozen sections rather than paraffin sections of fixed tissues, however frozen tissues are difficult to cut and the resulting histology is often suboptimal. In addition to the problem of low foreground signal, most monoclonal antibodies are made in mice, and background staining results when the anti-mouse secondary antibody recognizes endogenous antibody bound to mouse tissues or B cells. “Mouse to mouse” blocking reagents are available from a number of companies, but do not always solve the background problems. Our best results in mouse tissue have been obtained using polyclonal antisera raised in a non-mouse species such as hamster, goat, or rabbit. Fortunately, many such antisera, specific for a large number of viruses, are available from Millipore, and can be used to visualize infected cells in tissues using basic protocol 7. The right hand panel of Figure 3 demonstrates the utility of this technique with evidence of many infected airway lining cells (seen as red) following infection 24 hours earlier with a high dose (106 pfu) of influenza A virus. This is a demonstration of robust infection prior to the onset of visible pathology at early times post-inoculation.
We have included here a variety of different protocols to study respiratory virus infections. In addition, because of the many manipulations that are required in these protocols and the diverse technologies used, there are many different things that can go wrong. Thus, the investigator should refer to the specific protocols or to manual specific to each subject for more detailed information.
In these protocols, as with any in vivo experimental model, there will be variability between experiments and between individual animals at a given time point. As this is unavoidable, it is important to use sufficient numbers of animals such that a meaningful statistical analysis can be done. We typically use between 5 and 10 animals for each data point, and this is important for both objective measures such as cytokine production, as well as the more subjective evaluation of histopathology. When comparing more than 2 cohorts of animals, a simple student-t test is often not sufficient, and an analysis of variance must be performed to determine significance. For histologic analysis, differences between groups of animals will often be a matter of degree rather than a +/− situation, so it important to examine the slides blindly. We typically handle this by labeling each slide with only a number, and asking an unbiased examiner to determine whether the samples appear to segregate into groups. When no clear differences are present between 2 groups, we conclude that those are not significantly different. Although this system has worked well for us, it is important to use both qualitative and quantitative measures whenever possible, e.g. comparing BAL cytology and/or FACs analysis with the pattern of inflammation seen on tissue sections at each time point.
In planning an infection experiment, it is important to determine in advance which samples can be frozen (e.g. lungs for virus titers or BALs for ELISA), and which must be analyzed on the day that they are harvested. Tissues for histology must be fixed immediately, but cannot be processed until they have been fixed in formalin for several hours, but no more than 2 days. Lymphocyte analysis must be done immediately. Isolating immune cells from tissue samples and setting up a FACS-staining will take from 4 to 12 hr. depending on the number of tissue samples and the complexity of the staining. The samples can be run at the end of the day or, if fixed, during the following day. An ISH analysis will take at least one week, including the time needed to harvest, fix, process, and section tissues, with 2 days needed for staining.