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Vital dyes routinely used for staining cultured cells can also be used to stain and image live tissue slices ex-vivo. Staining tissue with vital dyes allows researchers to collect structural and functional data simultaneously and can be used for qualitative or quantitative fluorescent image collection. The protocols presented here are useful for structural and functional analysis of viable properties of cells in intact tissue slices, allowing for the collection of data in a structurally relevant environment. With these protocols, vital dyes can be applied as a research tool to disease processes and properties of tissue not amenable to cell culture based studies.
Staining and imaging cells with vital fluorescent dyes is a highly versatile approach widely used among cell biologists. The diversity of commercially available vital dyes allows for microscopic analysis of multiple cellular and subcellular parameters individually or combined. Parameters amenable to study using fluorescent probes include structure, organization, and localization of organelles and subcellular compartments, as well as functional measurements, such as reactive oxygen species (ROS) production, mitochondrial membrane potential, and pH. While vital dyes are popular tools among cell biologists, cell culture studies designed to model human disease are intrinsically limited; they lack the complex structure, cell type heterogeneity, and physical history of diseased tissues. Isolated reports of ex-vivo tissue staining appear in the literature, but these studies utilize few dyes and typically rely on two-photon microscopy for imaging.
In this unit methods are presented for staining and imaging live tissue slices ex-vivo for qualitative and quantitative data collection. The most basic procedure is suitable for collecting morphology and localization data from tissue using dyes that stain mitochondria and DNA simultaneously with detection of green fluorescent protein (GFP) tagged proteins (see Basic Protocol). Alternate dye sets may be used as a quality control measure for tissue viability and to examine cell death in tissue samples ex-vivo (see Alternate Protocols 1 and 2). Following these specific protocols is a more generalized method useful for a wide variety of dyes, which can be selected to fit the desired purpose (see Alternate Protocol 3). This protocol has been applied successfully to a range of tissue types, including human biopsy tissue. Two additional protocols present alternate methods for collecting quantitative data; one requires that images are collected at a constant depth from the surface of the tissue (see Alternate Protocol 4), while the other allows for depth independent acquisition, but requires empirical determination of the rates of signal decay as a function of depth within the tissue imaged (see Alternate Protocol 5). Data collected in this manner can be used to quantify changes in functional properties within three-dimensional structures within tissue. Physiological states such as cellular and mitochondrial ROS production can be examined using a slight variation on the basic technique (see Alternate Protocol 6). The staining steps in the presented protocols require little prior experience, while the image collection requires moderate experience with confocal microscopy, for which they are designed.
A variety of transgenic mouse lines exist where protein targets of interest are tagged with fluorescent protein conjugates or fluorescent proteins are expressed under the control of regulatory units of interest. A range of spectral variants have arisen over the years since the original cloning of GFP to allow for greater versatility and the use of multiple transgenic markers at once. The use of transgenically expressed fluorescent proteins allows for temporal and spatial investigation of protein expression or promoter activation using standard techniques such as western blotting of homogenized tissue and immunostaining of sectioned fixed samples or direct imaging of aldehyde fixed samples. Samples from fluorescent protein expressing animals have also been subjected to imaging with two-photon microscopy for analysis of localization within intact tissue sections. The following protocol provides a method for staining live GFP expressing tissue slices with dyes specific for mitochondria and DNA. This method provides a simple and highly accessible method for analyzing the localization and level of GFP in living tissue in relation to nuclei and mitochondria in living tissue slices. This technique is particularly applicable to investigation of dynamic subcellular processes such as mitophagy, where autophagosomes, which are commonly labeled with transgenic GFP tagged LC3(1), are assembled around mitochondria to target them for destruction via the lysosome(2,3). In this method a slice of freshly excised tissue is viably stained and imaged by confocal microscopy.
Damaged cells can be easily distinguished from intact cells with the use of cell-impermeable DNA binding dyes, such as ethidium homodimer-1 (EthD-1). Alternatively, dyes that are activated and retained only in intact cells provide a positive marker for cell viability. Calcein AM is a cell-permeable molecule that is converted by intracellular esterases into a calcium binding, highly fluorescent molecule. The use of these dyes together in staining of live tissue slices ex-vivo provides both positive and negative indicators of cell viability, allowing for the collection of data related to cell mortality in disease, as well as a powerful quality control method for tissue viability.
Alternate Protocol 1 provides a simple method for determining whether cell membranes have been compromised. This method is only one possible set of dyes that provides viability data, and may not be the best method for every scenario. Another useful dye set for examination of viability is a combination of the DNA dye Hoechst 33342, Sytox Green, and tetramethylrhodamine ester (TMRE). Hoechst 33342 is cell permeable and stains all nuclei regardless of membrane status. Sytox Green is similar to EthD-1 in that it cannot cross intact membranes and is a specific stain for the nuclei of cells with compromised membranes. TMRE is a membrane potential dependent mitochondrial dye. Mitochondrial membrane potential is often lost prior to cell death via apoptosis (REF), and when cells die via necrosis. Examining membrane potential provides an additional measure of viability not directly related to the cell membrane. Furthermore, staining with TMRE allows for examination of changes to mitochondrial function (as membrane potential) and mitochondrial structure in parallel with cellular viability. While the Sytox stain is extremely bright and highly specific, this method is not useful when tissue contains nuclear localized GFP.
The method provided in Basic Protocol 1 is a specific staining set designed for imaging the subcellular localization of GFP and the general tissue structure and morphology of tissue expressing GFP. This approach can also be applied to non-fluorescent protein expressing tissues as well as tissues expressing any spectral variant of GFP. Using the more generalized approach described below, dye sets can be selected based on the parameters of interest, the spectral compatibility of the dyes, and the available laser and filter sets on the confocal microscope used.
The methods in Basic Protocol 1 and Alternate Protocols 1–3 are all designed for the collection of data related to structure and localization but are not sufficient for collecting quantitative fluorescent intensity data. In order to collect quantitative data using confocal microscopy of a thick sample multiple variables must be accounted for. These include penetration of the dyes, signal attenuation of each dye as a property of depth, and signal crossover from one channel into the others. There are two approaches to collecting quantitative data: collecting data at constant depth in order to eliminate depth related variables, or collecting data at multiple, but defined, depths in order to empirically determine the relative rates of signal attenuation among the dyes and allow for normalized depth-independent collection of data. The first approach is utilized in this protocol.
The methods presented thus far are sufficient for collecting localization and structural information and for collecting quantitative data at a constant depth of imaging. In some circumstances it may be desirable to collect quantitative data across a set of z-plane images. In order to quantify data with depth of imaging as a variable the rate of signal attenuation with depth must be empirically determined for each of the dyes used. This information can be collected simultaneously with the experimental data and accounted for during image analysis.
The preceding protocols are designed for dyes that stain their respective targets without the need for active respiratory activity. While many stains that depend on functional cellular events may work while staining on ice it may be necessary for some targets to perform the staining at ambient or physiological temperature. Furthermore, the reactions necessary for the conversion of some probes into fluorescent molecules may not occur at a useful rate at 4°C. ROS sensitive dyes are one such set of probes that are poorly activated at low temperature and in the absence of active cellular metabolism. Staining at ambient or physiological temperatures requires that the staining buffer more closely match the natural environment of the tissue and provide appropriate nutrients and oxygen to support respiratory activity for the duration of the staining period. Raising the temperature also causes the tissue to become more fragile, especially weak and fatty tissues such as brain, requiring greater care during staining and handling.
Requires 10X HBSS (purchased from Invitrogen, catalog number 14065056). Dilute to 1X with sterile water. Supplement with 8.9mM sodium bicarbonate (100X purchased from Sigma, catalog number S8761) and 1% (w/v) Bovine Serum Albumin (BSA – Sigma catalog number A2153) or 5% (v/v) Fetal Bovine Serum (FBS – Invitrogen catalog number 10082-147). pH to 7.4. If this is prepared in advance cell culture grade antiobiotics, such as a penicillin/streptomycin (Invitrogen catalog number 15070-063), can be added, as directed by the manufacturer, to prevent bacterial growth. Store in 4°C for up to 6 months.
Dilute the following into 500mL sterile deionized water: 8g of NaCl, 0.2g of KCl, 1.44g of Na2HPO4, and 0.24g of KH2PO4. pH to 7.4, add sterile deionized water to 1L. Can also be made and prepared as 10X by dissolving in a total of 100mL sterile water.
Dilute Hoechst 33322 (Sigma catalog number B2261) to 1mg/mL in 1X PBS. Store protected from light at 4°C, up to 6 months, or long term in aliquots at −20°C.
Dissolve the 50μg contents of one MitoTracker Deep Red FM Special Packaging vial (Invitrogen catalog number M22426) with 459.9μL of DMSO. Store at −20 or −80°C protected from light.
Dissolve the 25mg contents of one bottle of tetramethylrhodamine ethyl ester perchlorate (Sigma catalog number 87917) with 4.8mL of DMSO to make a 10mM (50,000X) master stock solution. Store this long term in aliquots at −20°C protected from light.
Further dilute this to 200μM (1,000X) by diluting 1:50 in DMSO. Also store at −20° protected from light.
Requires 500mL low glucose DMEM with L-glutamine and pyruvate (Invitrogen catalog number 11885-084). Supplement 500mL bottle with 8.9mM sodium bicarbonate (100X purchased from Sigma, catalog number S8761) and 1% (w/v) Bovine Serum Albumin (BSA – Sigma catalog number A2153) or 5% (v/v) Fetal Bovine Serum (FBS – Invitrogen catalog number 10082-147).
Add cell culture grade antibiotics, such as a penicillin/streptomycin (Invitrogen catalog number 15070-063) to 1X to preserve unused solution for later use. Store at 4°C.
A variety of techniques exist for the study of cell and molecular processes involved in the etiology or progression of disease. Standard approaches include observation of morphologic characteristics by light or electron microscopy and detection of markers of disease state using immunological or mass spectroscopy based methods. These approaches work well when suitable disease markers exist or when structural defects, such as physical aberrations or detectable mislocalization of proteins, are present. Conversely, cellular processes involving short lived or unstable mediators are often very difficult to study in the context of their role in disease. Stable markers may not exist for important dynamic properties of the process under examination, such as flux. Examples of such parameters involved in disease include the production and neutralization of ROS, the regulation or disruption of mitochondrial membrane potential, the localization and concentration of biologically active ions, and cellular events such as autophagy and lysosomal maturation. These processes have been implicated in a wide range of clinical pathologies, from neurodegenerative diseases such as Parkinson’s and Alzheimer’s(5–7), to metabolic diseases such as diabetic nephropathy and diabetic cardiac hypertrophy(8–12). Mitochondrial dysfunction, as measured by mitochondrial ROS production or decreased mitochondrial membrane potential, has also been strongly implicated in the processes of cellular and organism aging(13–16).
While ROS and mitochondrial dysfunction are not directly amenable to study using classic histological and immunological techniques, several approaches have been developed to study these dynamic cellular events. Vital dyes and fluorescent protein markers provide a robust method for studying a variety of parameters in live cells in culture. These dyes allow for quantitative and descriptive analyses, provided that a suitable cell culture system exists. Unfortunately, while cell culture models can provide powerful insights into kinetic or unstable biological processes they often lack important structural information and contextual relevance with regards to the tissue they model. Even the most robust models of specific cell types are often highly derived, and nearly all cell culture models necessarily ignore the heterogeneity of cell types that exist within tissue. Furthermore, cells in culture typically lack the structural properties of the corresponding cells in situ and nearly always lack the cell-to-cell interactions that occur in vivo.
Isolated reports of imaging live, excised tissues appear in the literature (17–21), but these examples are sporadic, have utilized relatively few dyes, and typically rely on the use of two-photon microscopy. The methods here provide an approach applicable to a range of tissues using a variety of vital dyes. These techniques provide a valuable way to investigate the role of complex biological properties such as mitochondrial function, ROS, and autophagy, in disease and aging and a versatile tool potentially applicable to a broad spectrum of parameters amenable to fluorescent microscopy. Furthermore, while two-photon microscopy would greatly enhance the depth of imaging, the methods described in this Unit do not require a two-photon microscope, thus making them more widely accessible.
Careful tissue handling is the most important parameter in each of these protocols. Quality of staining, imaging, and data all depend on the viability and structural integrity of the tissue slices; the ability to collect high-quality images of control tissue(s) should be demonstrated prior to attempting to collect experimental data. If viability is poor (as compared to those in Figure 2) a number of potential factors must be considered. These include the amount of time the tissue spent between separation from (or death of) the donor, time and temperature of incubation in buffer, the care with which the tissue was sectioned, and the total time before imaging. Potential solutions may include altering the collection or euthanasia protocol, utilizing tissue cutting blocks (see Basic Protocol 1 Materials) or a vibratome (non listed here), incubating in cold buffer, and limiting the size of the experiment in order to allow for quicker processing and imaging.
In order to select suitable dyes there are three major factors that must be considered: (1) the structures or parameters of interest, (2) the spectral compatibility of the dyes with the laser and filter sets available on the microscope, and (3) the spectral compatibility each dye with the other dyes in the set. A set of four dyes can generally be used in combination, filling the spectral space of blue, green, red, and far-red dyes.
Proper selection of excitation and emission settings is also crucial to success. A set of suggested excitation wavelengths and emission filter sets are provided in Table 1. If unlisted dyes are used, or if the microscope available is not equipped with the filters or lasers suggested, optical settings appropriate to the experiment should be carefully determined. Whenever possible, cultured cells should be relied upon to test dyes and determine optimal settings.
Autofluorescence is always a potential problem in fluorescent imaging experiments. Commercially available vital dyes are typically bright and live tissue tends to have much lower autofluorescence than fixed and processed tissue slices, but tissue autofluorescence can interfere with data collection and image analysis. Extracellular matrix(17,22), NADPH(23,24), lipofuscin(25,26), and advanced glycation end products (AGEs)(27,28) are a few examples of naturally occurring substances that can interfere with imaging. In order to prevent misinterpretation of data every experiment must include a negative control to demonstrate the lack of (or at least much smaller) signal in the absence of staining. Such a control should be included for every tissue type and condition (including age), as these confounding factors vary by tissue, treatment, and source (age, species, etc).
The methods listed in basic protocol 1 and alternate protocols 1 and 3 provide high resolution,
The nuclei of non-viable cells stain brightly red with EthD-1. Non-viable cells do not stain with Calcein AM, whereas intact cells stain brightly green throughout their cytoplasm. Dead cells are positive for EthD-1 and negative for Calcein AM. All cell nuclei stain brightly blue-white after staining with Hoechst 33342. The dependence of viability on depth of optical sectioning is illustrated in Figure 2. As can be seen, high viability is typically seen at 5–10um depth, although results may depend on tissue type and even the direction of sectioning (see Figure 2B1).
The nuclei of non-viable cells stain brightly with the green Sytox nuclear dye (Figure 2C1-2). Mitochondria of dead cells lose their red TMRE staining and become weakly fluorescent or undetectable. Cells may lose mitochondrial function without losing membrane integrity. In this case the cells would show weak or no mitochondrial TMRE signal but would not show Sytox Green staining. The reverse is also possible, allowing cells to be determined viable or nonviable by multiple criteria.
Tissue collection and processing should take 30–60 min, but are dependent on the skills and tools available to the researcher and can vary greatly depending on circumstances and complexity of the experiment undertaken.
Staining and destaining tissues should take <45 min.
Image collection and data analysis are crucially dependent on the complexity of the experiment and the experience of the researcher. Imaging generally lasts 2–4 hours, but can last 6–8 hours for complicated imaging. Image analysis depends wholly on what is being analyzed and how the data is approached and can take as little as 30 min or if the analysis is difficult many hours.
LC3-GFP mice were generously provided by Dr. Al La Spada at the University of California, San Diego. The human colon sample in Figure 2 was collected with human subjects approval by Dr Teri Brentnall at the University of Washington Medical Center. This work was funded by NIH grants PO1 AG001751, P30 AG013280, RO1 HL101186 and T32 AG000057.
NIH ImageJ software - http://rsbweb.nih.gov/nih-image/