Here we provide an overview of different variables and techniques in immunofluorescence (IF) staining workflows. Also see our basic protocols for Immunofluorescence Staining of Cells for Microscopy, Cell Surface Staining for Flow Cytometry, and Intracellular Staining for Flow Cytometry.
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Culturing Cells for Microscopy
Cells can be imaged through tissue culture plastic with an inverted fluorescence microscope using objectives with low magnification and a long working-distance. High magnification oil-immersion objectives have short working distance and will not permit imaging through tissue culture plastic. Instead, coverglass should be used for imaging with oil-immersion objectives.
Cell culture-treated chambered coverglasses or coverglass-bottom 96-well plates are useful for culturing cells, and work very well for live cell imaging with an inverted fluorescence microscope. However, we have found that when fixing cells and permeabilizing cells in chambered coverglasses for immunofluorescence staining, the chambers can begin to leak. This can lead to artifacts from antibodies leaking from one well into neighboring wells. In our experience, coverglass-bottom 96 well plates withstand fixation and permeabilization conditions very well, even with alcohol, with no leakage between wells. Similarly, plastic chamber slides with removable chambers that allow coverslip mounting also may used, but they should be validated for fluorescence, because plastic can be autofluorescent.
Coverglass cultureware has the disadvantage of being quite costly. A low cost alternative is to culture cells on glass coverslips coated with poly-L-lysine or other extracellular matrix components. The coverslips are then mounted on standard microscope slides prior to imaging.
Fixation is the chemical treatment of cells or tissues to simultaneously kill and preserve cells by immobilizing proteins and other cellular components. Different fixation methods may be optimal for specific antigens or probes.
Formalin, formaldehyde, and paraformaldehyde, what’s the difference?
Aldehyde fixatives act by chemically cross-linking free amine groups on proteins. Formaldehyde is a commonly used fixative and is usually used in PBS or neutral saline at 2-4% for fixing cells, tissue specimens, or whole organisms by perfusion.
Formaldehyde is not stable in solution; with exposure to light and oxygen it polymerizes and precipitates. Formaldehyde solutions can be stabilized by the addition of methanol. Stabilized formaldehyde, also called formalin, is supplied as a 37-40% solution in water with 10-15% methanol. The classic fixative, 10% neutral buffered formalin (NBF), is commonly used for pathology studies. NBF contains ~3.7% formaldehyde/~1.5% methanol.
Many researchers prefer to use stabilizer-free formaldehyde, because methanol can permeabilize cell membranes and affect cell morphology. Paraformaldehyde is a polymerized formaldehyde solid that can be heated in alkaline water to generate formaldehyde. Fixative prepared in this way is commonly referred to as paraformaldehyde solution or PFA. While technically inaccurate, it serves to distinguish stabilizer-free formaldehyde from methanol-stabilized formaldehyde. Paraformaldehyde and formaldehyde are toxic and carcinogenic, and should be handled using laboratory safety precautions. We supply Paraformaldehyde, 4% in PBS, Ready-to-Use Fixative packaged under nitrogen as a convenient and safer alternative to preparing PFA solution from scratch.
Paraformaldehyde solution is usually diluted in 1X PBS to a final concentration of 2-4% for fixation. Generally cells or fresh-frozen cryosections are fixed at room temperature or 4°C for 10-30 minutes. Tissues and tissue slices may require longer fixation times. PFA diluted in PBS can be stored for at least one month at 4°C, protected from light.
Formaldehyde fixation alone does not completely permeabilize cellular membranes. For intracellular staining with antibodies, a separate permeabilization step is needed. However, formaldehyde can partially permeabilize membranes, especially after prolonged fixation at room temperature. Formaldehyde fixation also can contribute to background fluorescence (see Autofluorescence). Additionally, formaldehyde can mask antigens by cross-linking free amines in the antibody binding site, which can be avoided by using alcohol or acetone fixation.
Methanol and acetone fixation
Alcohols, such as methanol, and solvents, such as acetone, will fix cells by precipitating and dehydrating proteins, nucleic acids, and other cellular components. Alcohol and acetone fixation can preserve the morphology of some structures like microtubules better than aldehyde fixatives, but it also can disrupt membranes and other cellular structures. For example, methanol fixation cannot be used for phalloidin staining of filamentous actin. Pure acetone will rapidly melt polystyrene laboratory plastics, so a 1:1 mixture of methanol and acetone is often used to avoid this. Methanol and acetone can cause shrinkage of cells or tissues; acetic acid is sometimes added to these fixatives to reduce shrinkage.
Usually cells or fresh-frozen cryosections are fixed with pre-chilled methanol or acetone at -20°C for 5-10 minutes. Both alcohol and acetone fixatives can be stored in the freezer. They will not freeze, and so will be ready-to-use for fixation. Unlike aldehyde fixation, both alcohol and acetone fixation will permeabilize cells, so a separate permeabilization step is not needed.
Permeabilization & Blocking
Permeabilization is required to allow penetration of probes into fixed cells or tissues. Triton® X-100 and Tween®-20 are non-ionic detergents that are commonly used for permeabilization for immunofluorescence. Triton® X-100 is most commonly used for immunofluorescence, while Tween®-20 is traditionally used for western blots. However, Tween®-20 may be optimal for some intracellular targets for immunofluorescence. Cells can be permeabilized with PBS + 0.1-0.5% Triton® X-100 at room temperature or 4°C for 10 minutes, followed by incubation with a blocking buffer to prevent non-specific binding of antibodies. Alternatively, blocking buffer + 0.1% Triton® X-100 can be used for one-step blocking and permeabilization, as well as for antibody incubation steps. The detergent in blocking buffer acts as a surfactant for even spreading of buffer to further reduce non-specific antibody binding.
Other permeabilization agents may be useful for specific applications. Digitonin and saponin interact with cholesterol to form pores in membranes. Digitonin has been used to selectively permeabilize the plasma membrane while leaving intracellular membranes intact. Saponin has been used for reversible permeabilization of the plasma membrane. These agents may be useful for maintaining membrane morphology or for detection of membrane-associated probes or proteins that could be extracted by detergents like Triton® X-100 or Tween®-20. Because saponin permeabilization is reversible, it must be included during antibody incubation steps for intracellular staining.
Protein solutions or normal sera are used to dilute antibodies for binding as well as to block non-specific binding sites on cells before the antibody incubation steps. Traditionally, normal (non-immunized) serum from the same host as the secondary antibody is used for blocking in immunofluorescence protocols. However, purified proteins like BSA or gelatin at 1-5% are also effective blocking agents. Nonfat dry milk is commonly used for blocking membranes for western blots at 1-5%, but is not traditionally used for immunofluorescence. Normal goat serum, milk, and BSA should be avoided when using primary antibodies from goat or sheep, due to cross-reactivity of the blocking agent with secondary antibodies. Fish gelatin is a particularly versatile blocking agent for immunostaining of mammalian cells and western blots. It provides excellent blocking and does not cross-react with mammalian antibodies. After dilution to 1-2% in buffer, the gelatin will not solidify at low temperature, allowing antibody incubation to be performed in blocking buffer at 4°C.
Commercial blocking buffers that are specifically designed for immunofluorescence are also available, such as our TrueBlack® IF Background Suppressor System. These buffers contain blocking agents that suppress background from direct interaction of fluorescent dyes with samples, as well as non-specific antibody binding.
Sodium azide is usually added to blocking buffers at 2 mM or 0.02% final concentration to prevent microbial growth. However, azide should not be added to buffers for diluting HRP conjugates, because azide irreversibly inhibits HRP.
Blocking is generally not needed for staining with non-antibody probes like fluorescent phalloidin, lectins, bungarotoxin, or nuclear dyes, however, permeabilization may be required.
When selecting primary antibodies, check the supplier information to confirm that the antibody is suitable for your target species and application. When possible, test the antibody on a positive control cell line or tissue sample with high expression of the target protein to verify reactivity and specificity of staining. It’s usually advisable to validate a primary antibody using a secondary antibody (which will give more sensitive staining), before attempting direct immunofluorescence with a fluorescently labeled primary antibody, especially for tissue staining, where direct immunofluorescence can be challenging (see Staining tissue sections). It is usually necessary to perform a titration experiment to determine the optimal concentration of primary antibody for your sample type.
Polyclonal vs. monoclonal antibodies
Antibodies for research are generated by immunizing a host animal with a protein or peptide of interest to induce antibodies against the immunogen. The antibodies are then isolated from serum and purified by affinity chromatography to isolate those that specifically bind the protein of interest. Many different antibodies with different isotypes and binding sites may be generated in response to one antigen; if these are purified as a population, the resulting mix is called a polyclonal antibody. Polyclonal antibodies can show robust staining because they recognized multiple sites on a target protein. However, because the immune response of the host changes or time or between different animals, polyclonal antibodies can show lot-to-lot variability. Polyclonal antibodies can be produced in a wide range of host species, allowing flexibility for multiplexing with secondary antibodies.
Each B-cell produces a single antibody structure. Individual B-cells that secrete antibodies to a specific antigen can be isolated from a mouse, rabbit, or rat host and fused with an immortalized cell to create a cultured cell line called a hybrioma. Each hybridoma can be used to produce a single antibody clone, called a monoclonal antibody. With monoclonal antibodies, homogeneous batches of antibody can be isolated from cultured cells, allowing an antibody clone can be produced indefinitely with minimal lot-to-lot variability. Monoclonal antibodies recognize a single epitope in the target protein (usually 5-8 amino acids), so they can be used for highly specific protein analysis. However, monoclonal antibodies can be more susceptible to losing affinity after amino acid modifications (such as covalent labeling with a fluorescent dye) compared to polyclonal antibodies with heterogeneous antigen binding sites.
Recombinant monoclonal antibodies
In theory, immortalized hybridomas should produce a single antibody clone. However, many hybridomas have been found to express multiple heavy and light chains as a result of anomalies in the cloning process or mutations in the hybridoma over time (see Bradbury et al. 2018). As a consequence, antibodies that are thought to be monoclonal may in fact contain undesirable antibody variants with low affinity or non-specific binding. To avoid this possibility, the cDNA for the target-specific immunoglobulin can be cloned from a hybridoma and used for transient transfection of immortalized cells for antibody production. Antibodies produced by this method are called recombinant monoclonal antibodies, and currently are considered to be the most reliable antibody reagents in terms of specificity and lot-to-lot consistency. Browse our recombinant monoclonal antibodies.
Monospecific monoclonal antibodies
The majority of primary antibodies offered by Biotium have been validated by immunohistochemical staining of formalin-fixed, paraffin-embedded human tissues, which is usually a challenging assay condition. We also offer a growing number of antibodies validated by protein array validation, a proteomics-based approach to validation of antibodies using recombinant proteins. The antibody is bound to a HuProt™ Human Proteome Microarray displaying tens of thousands of unique, purified human proteins, representing ~81% of the human proteome. The level of binding of the monoclonal antibody (in combination with a fluorescently labeled secondary antibody) to each individual protein on the array is quantified and the signal strength is represented as the Z-score. The Z-score is described in units of standard deviations (SDs) above the mean value of all signals generated on that array. If targets are arranged in descending order of Z-score, the S-score is the difference (also in units of SDs) between the Z-scores, and represents the relative target specificity of the antibody for an individual protein. An antibody is considered monospecific for its intended target if its S-score for that protein is at least 2.5. For example, if a antibody binds to its intended target protein with a Z-score of 43 and to the second highest target with a Z-score of 14, then the S-score for the binding of the antibody to its intended target is 29. See our offering of monospecific monoclonal antibodies.
Secondary antibodies are antibodies that are raised in a host species to react with immunoglobulins from a different target species; they are used to detect primary antibodies in indirect immunofluorescence. Usually secondary antibodies react broadly with any antibody from the target species they are raised against, but secondary antibodies can be selected for more specific reactivity as well. Secondary antibodies are produced in a wide range of host species, most commonly goat and donkey. Typically secondary antibodies from the same species would be used together in a multiplex staining experiment (ie, goat anti-mouse would be paired with goat anti-rabbit to detect two primary antibodies raised in mouse and rabbit), but in our experience, donkey and goat secondary antibodies can be used together without cross-reacting with each other.
Highly cross-adsorbed secondary antibodies
Because secondary antibodies can cross-react with immunoglobulins from closely related target species, staining with primary antibodies from multiple species usually requires highly-cross adsorbed antibodies that have been selected for species specificity. Highly cross-adsorbed antibodies may also be required for staining tissues that contain endogenous IgG. This is commonly the case when rat tissue is stained with an anti-mouse secondary antibody, which can react with antibodies in blood vessels and immune cells in addition to the primary antibody. Using anti-mouse secondary antibodies that are pre-absorbed against rat serum proteins eliminates this problem. See our Goat Anti-Mouse IgG (Min x Rat), which is highly cross-adsorbed for minimal cross-reactivity to rat.
Isotype-specific secondary antibodies
The structure of the Fc portion of immunoglobulins, or isotype, varies with B-cell maturation status and antibody function. There are two subclasses of immunoglobulin light chain (kappa or lambda) and five subclasses of immunoglobulin heavy chain (IgG, IgM, IgA, IgD, and IgE). IgG is the most common antibody subclass in serum, and consequently the most common isotype for research antibodies. There are five IgG subclasses in mice: IgG1, IgG2a, IgG2b, IgG2c, and IgG3; IgG1 kappa is the most common isotype for mouse monoclonal antibodies used in research. Secondary antibodies generally react with all subclasses of immunoglobulin, but subtype specific secondary antibodies can be generated to specifically recognize individual Ig isotypes. Isotype-specific secondary antibodies can be useful for using multiple monoclonal antibodies from mouse of different isotypes in multiplexing assays.
F(ab’)2 fragment secondary antibodies are produced by enzymatic digestion of whole IgG to isolate the antigen-binding variable regions without the Fc constant region. Because of their smaller size, F(ab’2) fragments can penetrate thick tissue sections more efficiently than whole IgG. Because they lack the Fc region, they do not bind Fc receptors on immune cells, or interfere with detection using Fc-specific antibodies.
See our full selection of labeled secondary antibodies, including highly cross-adsorbed, isotype-specific, and F(ab’)2 fragment antibodies.
Direct vs. Indirect Immunofluorescence
Immunofluorescence (IF) can be performed with direct or indirect detection of the target antigen. Fluorescent antibody conjugates typically are labeled with 4-6 dye molecules. Direct IF uses a dye-conjugated antibody to fluorescently label the target antigen. With direct IF using monoclonal antibodies, only one antibody may bind to its target antigen, tagging it with 4-6 dye molecules. With indirect IF, an unlabeled primary antibody is bound to the target antigen, followed by binding of multiple fluorescent secondary antibodies to the primary antibody. Because each secondary antibody bears 4-6 dye molecules, this greatly increases the number of dyes associated with the target antigen, resulting in higher sensitivity relative to direct IF. However, direct IF offers the advantage of being able to stain a sample with multiple primary antibodies from the same host species simultaneously. Direct and indirect methods can be combined to exploit the advantages of both methods. See our Tech Tip for Combined Direct and Indirect Immunofluorescence to learn more.
Staining Tissue Sections
Sections of fresh, frozen, or fixed tissues can be prepared for staining using a variety of methods, including vibratome sectioning, cryosectioning, paraffin-embedding, or plastic embedding. Any of these types of sections can be stained with antibodies, though plastic-embedded sections are not well-suited for immunofluorescence and require specialized protocols. Paraffin and cryosections (5-15 um thickness) are mounted on glass slides. Thick tissue sections (40 um or thicker) are prepared with a vibratome; these cannot be affixed to slides and are left free-floating; staining of floating sections can be performed in multiwell plates. Thick sections typically require specialized protocols using much longer incubation and washing times to allow penetration of reagents compared to thin sections.
Paraffin sections are prepared from formalin-fixed tissue that has been dehydrated and embedded in wax. The sections must be deparaffinized and rehydrated prior to imaging by a series of incubations in xylene and ethanol solutions. Paraffin embedding is preferred for histopathology because it preserves tissue morphology very well. However, the extensive fixation and tissue processing can cross-link or damage antigens in the tissue, resulting in poor immunostaining. Antigen retrieval methods to reverse aldehyde cross-links, such as boiling in acidic or basic buffer, or protease treatment, may be necessary before sections can be stained with certain antibodies. Check the supplier information for your primary antibody to find the recommended antigen retrieval method.
Cryosections are prepared from fresh or fixed frozen tissue. Fresh-frozen tissue sections are preferred for immunostaining. While the tissue morphology may not be as well maintained in cryosections compare to paraffin sections, the preparation of tissues and sections is greatly simplified, with minimal modification or loss of antigens. Cryosections are stored frozen (usually at -80°C), and brought to room temperature and air dried before fixation (for fresh sections) and staining. Fresh-frozen sections must be fixed before staining, and can be fixed with fixatives commonly used for staining cells. When purchasing tissue cryosections from commercial sources, pay attention to how they are prepared. Some vendors provide cryosections that are pre-fixed with acetone, which may not be compatible with some stains like phalloidin.
Fixation, blocking and washing of sections on slides is conveniently done in Coplin jars or slide staining jars that can hold anywhere from 40 to 200 mL of solution. The slides are removed from the jars and placed on a flat surface for antibody incubation steps. After draining or wicking away excess buffer from the slides with a lab wipe, a small drop (50-100 uL) of antibody staining solution is placed directly on each section. A small square of Parafilm® can be placed on top of the antibody solution to spread it evenly over the section without air bubbles. A humidified chamber is often used for antibody incubation steps to prevent evaporation of the staining solution. Sections of plastic serological pipettes can be placed on top of damp paper towels in a plastic box with a lid to make a rack for the slides. The box can be covered in foil to protect slides from light during incubation with fluorescent conjugates.
Pap pens can be used to create hydrophobic barriers around sections to allow multiple sections on the same slide to be stained with different probes. The barriers are resistant to alcohols and heat, but can be removed with xylene. For best results, barriers should be applied to slides when dry, then allowed to dry thoroughly before immersing in buffer. Care must be taken to avoid contact of the barrier solution with the tissue section.
Fluorescence imaging of human tissue sections is often challenging, due to potentially lower endogenous protein expression compared to immortalized cell lines, higher autofluorescence, and potential loss of epitopes due to post-mortem tissue degradation or tissue processing. Staining with directly labeled primary antibodies may be suitable for only the most abundant targets. Primary antibody plus secondary antibody can provide enough signal amplification for sensitive detection of many antigens, but additional signal amplification using tyramides (see below) may be required for low-expressing targets.
Autofluorescence is a major source of non-specific background fluorescence in tissue sections and some primary cell types. Sources of autofluorescence include aldehyde fixatives, tissue components with endogenous fluorescence (including extracellular matrix proteins, red blood cells, and macrophages), and lipofuscin, which consists of highly autofluorescent granules of oxidized proteins and lipids that build up in the lysosomes of cells with age. When performing immunofluorescence staining of tissue samples, you should include an unstained control to determine the level of autofluorescence in your sample. While usually brightest in the blue and green wavelengths, autofluorescence has broad spectrum fluorescence that can make detection of specific fluorescence signal in tissues virtually impossible unless it is quenched or masked.
Many treatments have been reported to reduce autofluorescence, including quenching of aldehydes with ammonium sulfate and Tris, bleaching with sodium borohydride, and quenching of autofluorescence with blue or black dyes. The lipophilic dye Sudan Black B is highly effective at masking autofluorescence from lipofuscin, but has the drawback of introducing red fluorescent background. Our TrueBlack® Lipofuscin Autoflurescence Quencher was developed as an alternative to Sudan Black B; it effectively quenches lipofuscin autofluorescence with much lower background than Sudan Black B. TrueBlack® also can reduce autofluorescence from other sources such as red blood cells and extracellular matrix. See our Tech Tip: Battling Tissue Autofluorescence.
Tyramide signal amplification (TSA) is a powerful method for increasing the sensitivity of immunofluorescence staining. After binding to primary antibodies or to biotinylated primary antibodies, an HRP-conjugated secondary antibody or HRP-streptavidin mediates covalent coupling of fluorescent dye-tyramide to tyrosine-containing proteins in the vicinity of the reaction site. This enzymatic coupling reaction allows the target proteins to be labeled with large number of dye molecules, resulting in much stronger signal compared to using dye-labeled antibodies. TSA can be combined with conventional immunofluorescence protocols or performed sequentially for multi-color detection, see our Tech Tip for multicolor tyramide staining to learn more. Biotium offers CF® Dye Tyramide Amplification Kits as well as stand-alone tyramide reagents.
Antifade Mounting Medium
Mounting medium is a viscous liquid (often glycerol-based) that allows coverslips to be mounted on specimens on glass slides. This keeps the samples from drying out and allows imaging with oil immersion objectives. Because excitation light used in fluorescence microscopy can cause fluorophores to photobleach, antifade compounds are often included in mounting medium to prevent loss of signal during imaging.
Antifade mounting media are available in wet-set formulations that stay liquid during storage, requiring the coverslips to be sealed (see Sealing coverslips, below). Wet-set mounting media also can be added to chambered coverglass or coverglass-bottom 96-well plates, where samples are imaged through a coverglass substrate rather than with an overlaid coverslip. Hardset mounting medium is an alternative formulation that dries and hardens to form a durable seal between the sample and coverslip. Mounting media usually are formulated to have refractive indices similar to that of glass (~1.5), to avoid image distortion caused by bending of light by refraction as it passed through the glass into the medium during imaging.
The blue nuclear counterstain DAPI does not require washing after staining, and can be included directly in mounting medium for one-step counterstaining and mounting. However, using mounting media with DAPI has the potential drawback that the dye concentration is not as easy to adjust than if it used in a separate staining step. Also, care must be taken to avoid accidentally using mounting medium with DAPI in experiments where other blue dyes are used. Biotium offers wet-set EverBrite™ Mounting Medium and EverBrite™ Hardset Mounting Medium, both with or without DAPI. We also offer Drop-n-Stain EverBrite™ Mounting Medium, a less viscous wet-set formulation supplied in a dropper bottle, which is convenient for adding to slides, plates, or chambered coverglass.
Organic mounting medium (such as Permount™) is designed for mounting specimens stained with chromogenic substrates like DAB. These media contain xylene or other organic solvents, and require samples to be dehydrated (usually with ethanol or xylene) before mounting. They do not contain antifade reagents are are not commonly used for fluorescence, but some researchers report good results using these mountants with air-dried fluorescently-stained sections.
When using wet-set mounting medium to mount coverslips on slides, the edges of the coverslip must be sealed to keep the coverslip stationary during imaging, and to avoid leakage of mounting medium or drying of the specimen. Traditionally, nail polish is used for this purpose. However, nail polish has the disadvantage of inconsistency in performance between brands or batches, as well as requiring researchers to purchase it outside the lab. It also can contain alcohols that can leach into mounting medium during sealing and affect sample fluorescence.
Biotium offers CoverGrip™ Coverslip Sealant as an alternative to nail polish for sealing coverslips. It is insoluble in water and does mix with mounting medium, and is supplied in a convenient brush bottle for application, just like nail polish.
Fluorescence Cross-Talk & Bleed-Through
Fluorescence from one dye can bleed into adjacent channels during fluorescence imaging, especially if the signal is very bright, resulting in artifacts in fluorescence co-localization experiments. In any multi-color imaging experiment, single-stain controls should be included to determine whether there is fluorescence cross-talk or bleed-through of dye fluorescence between channels. Cross-talk can be avoided by choosing dyes with well-separated fluorescence emission for multicolor imaging. Our Fluorescence Spectra Viewer can be useful for evaluating dye separation for multicolor experiments. Probes should be titrated so the relative brightness of each dye is similar, if possible. Confocal microscopy imaging settings also can be optimized to minimize cross-talk by limiting cross-excitation during scanning, or by changing the emission cut-off for different dyes.
For DAPI bleed-through into the green channel, reduce the concentration of DAPI, or optimize confocal imaging settings to prevent cross-talk. UV excitation of DAPI and Hoechst also can cause these dyes to emit green or red fluorescence, a sometimes unexpected source of cross-talk; see our Tech Tip: Avoiding Artifacts from UV Photoconversion of DAPI and Hoechst. Using different counterstains, such as RedDot™2 Far-Red Nuclear Stain (for the Cy®5 channel), or NucSpot® 470 also can avoid this problem.
For common causes and solutions of low/no signal or high background/non-specific signal, see our Troubleshooting Tips for Fluorescence Staining.
Products for Immunofluorescence
|DAPI in H2O, 10 mg/mL||
|RedDot™2 Far-Red Nuclear Stain||
|NucSpot® 470, 1000X in DMSO||
|4% Paraformaldehyde in PBS, Ready-to-Use Fixative||
|TrueBlack® IF Background Suppressor System (Permeabilizing)||
|TrueBlack® Lipofuscin Autofluorescence Quencher||
|EverBrite™ Mounting Medium||
|EverBrite™ Hardset Mounting Medium|
|Drop-n-Stain EverBrite™ Mounting Medium|
|CoverGrip™ Coverslip Sealant||
|10X Phosphate Buffered Saline (PBS)||
|Permeabilization and Blocking Buffer (5X)||
|10X Fish Gelatin Blocking Agent||
|Fish Gelatin Powder||
|Bovine Serum Albumin, 30% Solution||
|Bovine Serum Albumin Fraction V||
|SuperHT Pap Pens||