What is a simple protocol for staining cells in suspension?

What is a simple protocol for staining cells in suspension?

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I am an engineering student studying how electric fields affect cells, specifically the phenomena of electroporation in living cells.

I know that electroporation is widely used for introducing genes into cells, but running a full transfection experiment is not so feasible in the lab I am working in.

My question is:

If I want to demonstrate the principle of electroporation, i.e. that cells open up and intake substances from the environment, by using some kind of a dye which i can later visualize, what could be a good model for this type of experiment? Specifically - what dye, what cells could be used for this?

You will get a lot of false-positives using the following method, and a real transfection of a fluorescent protein is always the way to go, because then you will prove that the transfection was really successful.

That said, you could try to use DAPI, followed by fluorescence microscopy or flow cytometry. DAPI is a very bright stain for DNA and cannot pass an intact cell membrane. If the cell membrane is disrupted, e. g. by necrosis, DAPI will enter the cell. I have never seen anyone trying to prove electroporation this way, but it will be worth a try. It could be that the time is too short for DAPI to enter the cell, so you'd need to tweak your setup.

I'd try the following: Prepare a cell solution with DAPI (something like 5 µM concentration should do). Split the solution into two electroporation cuvettes, perform the electroporation with one cuvette, leave the second one untouched as a control. After the electroporation, analyse the samples MOMENTARILY (means: within a few minutes) using a flow cytometer (if possible).

As for the cells, you could use anything. I suggest eukaryotic cells because they are more easy to visualize than bacteria. However, you would need a laminar flow hood for aseptic handling of cells. In theory you could also use bacteria, which are more easy to handle, but I'm not experienced with immunofluorescence in bacteria.

Cell Staining Protocol for Microscopy

Microscopy refers to the practice that involves the use of a microscope for the purposes of observing small scale structures that cannot be viewed using the naked eye and often cell staining is necessary as s tructures are difficult to discern due to insufficient contrast.

Cell staining is a technique used for the main purpose of increasing contrast through changing the color of some of the parts of the structure being observed thus allowing for a clearer view. There are a variety of stains that can be used in microscopy.

First of all, staining can be in-vivo or in-vitro. The difference between these is that whereas In-vivo staining refers to the staining of a biological matter while it is still alive, i n-vitro staining refers to a staining technique where the biological matter is non-living.

The following are common stains explaining techniques, preparations and procedures for each:

Haematoxylin and Eosin Staining

These are two stains used in the examination of thin slices of biological tissue. Contrast is created by the stains where Haematoxylin turns the nuclei blue while eosin turns the cytoplasm as well as other parts pink or red.

1- Measure 10 grams of haematoxylin crystals 500 ml of water (70- 80 degrees centigrade) and mix to dilute completely

2- In a separate flask, measure 20 grams of alum and mix with 500 ml of hot tap water (70- 80 degrees)

3- Mix the two mixtures together (1 and 2)

Whereas alum is the mordant, thymol prevents fungal growth.

5- The mixture is then kept in a translucent flask away from direct sunlight for one week. This is covered with a paper towel that allows for air circulation (early maturation). The solution is then put in to a dark flask and topped tightly after one week, and stored in a dark place for 3 weeks. (Late maturation)

1- Measure 10 grams of eosin crystals and add to mix in 1000ml of hot tap water (70- 80 degrees). This should be mixed to dilute and stored in a dark flask. This can be used directly.

1- A rehydrated section is stained in a solution of haematoxylin for 20 to 40 minutes

2- The section is then washed in tap water for about 3 minutes until it turns blue,

3- The section is the differentiated in 70percent ethanol that contains 1 percent of HCL for about 5 seconds to remove excess dye and allow the nuclear to emerge,

This is then washed in tap water,

5- Stain with eosin for 10 minutes,

6- Then wash for about 1 to 5 minutes in tap water,

7- Dehydration, clear and mount on a rack

Haematoxylin-Eosin Stain Kaposi's Sarcoma lesion - Cambridge University Press

Papanicolaou Staining

This is also referred to as pap-staining or pap smear. It is used for the purposes of examining cell samples that have been obtained from body fluids.

The technique involves the combination of chemicals that include:

o Light green SF yellowish,

ACCUMATE differentiating solution is made ready for use. A substitute of Scott's Tap water is prepared through mixing a part of Scott's Tap water substitute concentrate with 9 parts of deionized water. This is then followed by filtering papanicolaou staining system reagents before use.

1- Fix the slides in acetic fixative for 15 minutes,

2- Absolute alcohol for two minutes,

3- 70 percent of alcohol for 2 minutes,

4- in 50 percent for 2 minutes

5- Tap water for 2 minutes,

6- deep in haematoxylin for 4 minutes,

7- Briefly rinse in tap water,

8- Differentiate in acid alcohol for about 5 seconds,

10- Dehydrate using absolute alcohol two times,

11- Stain in orange G for ten seconds,

12- Rinse in absolute alcohol two times

13- Stain in E.A 50 for two minutes,

14- Stain in absolute alcohol two time,

15- Clear in xylene three times,

16- Mount the slide on xylene three times,

a) Presence of a psammoma body in absence of atypical cells in cervicovaginal smear (Papanicolaou stain, 200×) b) Serous ovarian cystoadenofibroma with parietal psammoma bodies (Hematoxylin and Eosin, 100×).

Pusiol et al. CytoJournal 2008 5:7

Acid and Basic Fuchsin Stain

Acid fuchsin is a magenta red acid dye that is largely used for plasma staining whereas basic fuchsin is a magenta basic dye largely used to stain the nucleus.

The technique is also referred to as acid fast staining. The acid fast bacteria have a waxy substance ( mycolic acid) on their cell wall that makes them impermeable to staining procedures.

The term acid fast is used since they resist decolourization with acid alcohol. Carbol fuchsin , the primary stain contains phenol, which helps solubilize the cell wall whereas heat is used to increase the penetration of the stain.

On using alcohol to decolorize, cells will be decolorized except for acid fast ones. Methylene blue is used as the counterstain to any cell that was decolorized. At the end of the procedure, acid fast cells remain red/pink while non-acid fast cells retain a blue color.

Preparation of carbol fuchsin by mixing two solutions:

Solution 1- 0.2 grams of basic fuchsin and 10 ml of 95 percent ethanol,

Solution 2- 5 grams phenol and 90 ml of distilled water,

1- Swab pulp larvae together so as to allow the pulp to spread over the slide and allow to dry

2- Heat fix by flaming over a burner several times,

3- Flood using 0.2 percent of carbol fuchsin for about 30 seconds,

4- Wash off the stain and then air dry/gently blot dry before examination

A photomicrograph of Mycobacterium smegmatis (pink) and Micrococcus luteus (blue) 1000x magnification. Mycobacterium smegmatis is acid-fast, retaining the carbol fuchsin dye, thus appearing pink. Micrococcus luteus is not acid-fast, loses the carbol fuchsin during decolorization, and is counter-stained with methylene blue.

Image from:

Wright's Stain

This is a Romanowsky type of metachromatic stain that is prepared by mixing specially treated methylene blue dye with eosin.

The acidic portion of the stain unites with the basic components of the cells such as hemoglobin, and thus they are referred to as eosinophilic and are stained pink or red.

The acidic components of the cell, such as the nucleic acids on the other hand take the basic dye and stain blue or purple.

PH has to be controlled using a buffer of 6.4 to 6.7 to avoid poor staining.

o M easure 1.0 grams of wright's stain powder and 400 ml of methanol (methyl alcohol),

o A dd a few glass beads to assist in dissolving and add the ethanol to the stain,

o M ix well at intervals until the powder has completely dissolved (do this by warming in 37 C water bath to aid in the dissolving),

o L abel the bottles and mark it as flammable and toxic,

o T ightly atop and store at room temperature in the dark

1- Prepare a fill of the sample and allow drying on a slide,

2- Prepare three containers, and fill one with one step Wright’s Stain and the other two with distilled water,

3- Keep the stain tightly covered when not in use to avoid evaporation (always replace the stain once it becomes insufficient)

4- Always replace distilled water once iridescent scum start forming on the surface, or when it starts turning blue,

5- Dip the slide in the stain for 15 to 20 seconds,

6- Dip the slide in distilled water in the second container for 15- 45 seconds,

7- Dip the slide in container 3 for 25 seconds using quick dips,

8- Wipe the back of the slide,

9- Dry the slide on a vertical position, on the absorbent surface and avoid blotting the smear,

10- Apply oil to examine microscopically,

These steps should be repeated two times for marrow smears.

1- Prepare the sample and allow to air dry,

2- Place the slide on a rack,

3- Apply the one step wright's stain using dropper bottles,

4- Wait for about 15-30 seconds and add similar volumes of distilled water,

5- Pour stain and water mixture off the slide,

6- Dip the slide in distilled water for about 25 seconds using quick dips,

7- Wipe the back side of the slide,

8- Dry the slide in a vertical position on the absorbent side and avoid blotting the smear,

These steps should be repeated twice for bone marrow samples.

Image from:

Gram Staining

This is one of the most common staining techniques.

It is largely used to differentiate bacteria species as either gram positive or gram negative. This is achieved through the chemical properties of bacterial cell walls, where different colors are displayed after staining.

This technique is based on the fact that the gram positive cell wall has a strong attraction for crystal violet following the addition of iodine as compared to the cell wall of gram negative.

Iodine is the mordant, and forms a complex with crystal violet, which is easily washed off from the gram negative cell wall using ethyl alcohol.

Cell staining in Microscopy is useful and necessary to highlight those structual elements of your sample/specimen to be properly observed. There are others that MicroscopeMaster may cover in the future so bookmark this page and revisit to see further staining information.

Learn more about Cell Culture, Cell Division, Cell Differentiation as well as Tissue Culture Types and Techniques. And take a look at the reasons to consider purchasing a Microscope Staining Kit.

Mallory F.B. A.M, M.D. S.D. Pathological Technique. Philadelphia & London. W.B. Sunders Company. Copyright, 1938, by W.B. Sunders Company. Reprinted June, 1942 and September, 1944. Pgs. 70 and 9

Johnson PL, Klein MN: Application of Papanicolaou stain to paraffin sections. Stain Technol 31:223, 1956

Delisle, G., and L. Tomalty. 2002. Mycobacterium tuberculosis. MicrobeLibrary, American Society for Microbiology, Washington, DC. is a participant in the Amazon Services LLC Associates Program, an affiliate advertising program designed to provide a means to earn fees by linking to and affiliated sites.

The material on this page is not medical advice and is not to be used for diagnosis or treatment. Although care has been taken when preparing this page, its accuracy cannot be guaranteed. Scientific understanding changes over time.

** Be sure to take the utmost precaution and care when performing a microscope experiment. MicroscopeMaster is not liable for your results or any personal issues resulting from performing the experiment. The MicroscopeMaster website is for educational purposes only.

Subculturing Suspension Cells

The following protocols describe general procedures for subculturing mammalian cells in suspension culture. Note that the procedure for passaging insect cells differs from that for mammalian cells on several crucial steps. For more information, refer to Notes on Subculturing Insect Cells.

For passaging your own cell line, we recommend that you closely follow the instructions provided with each product you are using in your experiments. The consequences of deviating from the culture conditions required for a particular cell type can range from the expression of aberrant phenotypes to a complete failure of the cell culture.

Passaging Suspension Cultures
Subculturing suspension cells is somewhat less complicated than passaging adherent cells. Because the cells are already suspended in growth medium, there is no need to treat them enzymatically to detach them from the surface of the culture vessel, and the whole process is faster and less traumatic for the cells. Replacement of growth medium is not carried out in suspension cultures instead, the cells are maintained by feeding them every 2 to 3 days until they reach confluency. This can be done by directly diluting the cells in the culture flask and continue expanding them, or by withdrawing a portion of the cells from the culture flask and diluting the remaining cells down to a seeding density appropriate for the cell line. Usually, the lag period following the passaging is shorter than that observed with adherent cultures.

Tissue Disaggregation

Mincing Tissues

After dissection of solid tissues from an organism, before introducing the digestive enzymes, tissues should be rinsed to clean off any blood or other unwanted material. The tissues should then be minced and dispersed with scissors, a scalpel, or a blade to increase total surface area. This increases contact between the enzymes and the surface of the tissues, leading to more efficient and complete digestion while shortening the time required for digestion.


Tissues hold cells together, supported by extracellular matrix and cell–cell junctions made up of a diverse set of proteins and other biological molecules which require specific enzymes for proper digestion and removal from the cell suspension. The digestive enzymes that play an essential role in the disaggregation of solid tissues are summarized in Table 1. The first category of enzymes to consider is those enzymes that break down the extracellular matrix. Dispase is a commonly used neutral protease isolated from bacteria, with a high level of enzymatic specificity for collagen IV and fibronectin 21, 22 . Dispase is useful in the detachment of cell colonies and the dissociation of tissue pieces into small clumps of cells, as it works to cleave attachments between cells and the extracellular matrix without affecting cell–cell junctions 23 . Caution should be used when digesting tissues with dispase, however, as it is able to cleave specific relevant surface molecules or antigens, such as those connected to T cell analysis 24, 25 . Therefore, omitting dispase from the digestion buffer may be helpful if a loss of epitopes is observed. Also effective in the disaggregation of the extracellular matrix is collagenase. Collagenase is able to break the peptide bonds present in collagen which helps to digest the extracellular matrix, releasing cells into suspension. It is important to note that purified collagenase enzymes are more effective than traditional collagenase naturally derived from bacteria as there is less variability in the composition of the purified enzyme, increasing the stability of the cells throughout the tissue digestion 26 . The final enzyme to consider in the digestion of the extracellular matrix in solid tissues is hyaluronidase. Hyaluronan, a structural proteoglycan in the extracellular matrix, is degraded by the hyaluronidase family of enzymes, which are produced in both bacterial and vertebrate organisms. These hyaluronidase enzymes cleave the β1,4-glycosidic bond present in the glycosaminoglycan portion of hyaluronan 27 , contributing to the digestion of the extracellular matrix.

-Breaks down extracellular matrix

-Cleaves attachments between cells and extracellular matrix

-Breaks down extracellular matrix

-Breaks peptide bonds present in collagen

-Breaks down extracellular matrix

-Cleaves glycosidic bonds in hyaluronan

-Prevents cell aggregation

-Cleaves cell–cell junctions

-Does not alter antigen expression as trypsin would

The next enzyme group to consider in the preparation of a single cell suspension includes the enzymes that break cell–cell junctions. Trypsin is a natural protease synthesized in the digestive system of vertebrate organisms. Although it is useful in degrading certain proteins present in cell–cell junctions, trypsin also has a very harsh effect on cell membrane proteins 1 . Also, trypsin has been shown to lead to free-DNA induced aggregation of cells, which indicates that cell lysis is occurring within the suspension 28 . Therefore, trypsin is traditionally avoided in the preparation of a single cell suspension from solid tissues for flow cytometry experiments. Papain is an alternative protease derived from the papaya plant. Papain is known to degrade the proteins that make up tight junctions between cells 29 . However, like trypsin, papain has been shown to lead to free-DNA induced aggregation of cells due to the cell lysis that occurs during enzymatic digestion 28 .

The final enzyme type to consider in the preparation of a single cell suspension is deoxyribonuclease (DNase) which acts to cleave the phosphodiester linkages of the DNA backbone. The two major types of DNase are DNase-I and DNase-II, which possess slightly different enzymatic functions. DNase-II is not suitable for the preparation of a single cell suspension as it plays a role in engulfment-mediated DNA degradation pathways involved in apoptosis 30, 31 . DNase-I is appropriate for tissue digestion and preparation of a single cell suspension as it prevents cell aggregation by degrading free-DNA released through dead cell lysis during the enzymatic digestion without initiating apoptotic pathways. Calcium chloride (CaCl2) acts as an enzyme activator of DNase-I and is therefore introduced into the digestion cocktail during enzymatic digestion. Calcium ions (Ca 2+ ) bind tightly to the DNase-I enzyme to stabilize its active conformation and allow for the proper degradation of free-DNA 32 .

To avoid the possible problems caused by adding the above enzymes to a tissue digestion cocktail, commercially available digestion cocktails have been developed and optimized. Accutase is a commercially available protease and collagenase blend which mimics the action of trypsin and collagenase but does so at a much lower concentration than is needed when using the standard enzymes. Accutase contains a mixture of enzymes with proteolytic, collagenolytic, and DNase activity and produces a higher total cell yield and improved overall antigen preservation when compared with using a cocktail of similar enzymes for tissue digestion 33, 34 . TrypLE is another commercially available enzyme cocktail containing purified, recombinant enzymes which mimic the activity of trypsin without altering the expression of cell surface antigens 35 . This product avoids the issues that arise with trypsin use in tissue digestion and allows for improved cell survival and more efficient single cell suspension preparation.

Enzymatic and Mechanical Dissociation

Enzymatic dissociation is carried out by introducing a digestion cocktail to minced, solid tissues and incubating at specific temperatures, based on the enzyme cocktail being used. Enzymes may be temperature specific, and therefore work with maximum speed and efficiency at a given temperature, commonly 37°C. Depending on the specific enzymes, enzymatic dissociation may also be carried out at 4°C or on ice. These lower temperatures will likely slow the reaction rate of the enzymes and extend the incubation period but can help to minimize cell death. Enzyme strength and enzyme concentration are the two most important factors to consider when choosing a digestion cocktail for the preparation of a single cell suspension for flow cytometry. Enzymes with high strength or high concentration may compromise cell surface markers present on the cells, which can affect the availability of these markers and the viability of the cells in further experiments. Therefore, lightly adherent cells such as lymphocytes should be isolated using a short digestion period with a gentle or mild enzyme to avoid these issues 36 . Determining the optimal strength and concentration of the enzymes being used in enzymatic dissociation is empirical and critical for proper isolation of cells and successful digestion of tissues.

Mechanical dissociation plays a role in the preparation of a single cell suspension from solid tissues throughout the enzymatic dissociation. To assist in the mechanical dissociation of the tissues, by which cells are released from the extracellular matrix into suspension, enzymatic digestion may be carried out on an orbital shaker. Following enzymatic dissociation, the suspension should be filtered in order to exclude any undigested tissue pieces or aggregates from the newly prepared single cell suspension.

16: Simple Stain

In order to stain the bacterial specimen for microscopy one must first prepare the smear on the slide. This basically involves three steps----transferring a liquid suspension of the bacterium on the slide, drying the smear, and then heating slightly to firmly attach the smear to the slide. Once this is done, the staining procedure begins.

Do NOT make your smear suspensions too thick. The dye will not penetrate well, and there will be far too many bacterial cells to see individual shapes and arrangements. One needs to be careful about thick smears when taking the specimen from an agar medium.

RECOMMENDATION: You may find it helpful to draw a circle (wax pencil is best) on the opposite side of the slide where you will spread your smear. This will help you later in locating the smear, sometimes a problem when moving the slide back and forth looking for your bacteria. The wax pencil is better than a marker because it will not wash off easily from the glass.


    Obtain a uniform suspension of cells: Follow the typsinization/trypsin neutralization protocol for the specific cell type. Place the cell suspension in a suitably-sized conical centrifuge tube. For an accurate cell count to be obtained, a uniform suspension containing single cells is necessary. Pipette the cell suspension up and down in the tube 5-7 times using a pipette with a small bore (5 ml or 10 ml pipette). For cells thawed from cryopreservation (in 1ml cryopreservation medium), pipette up and down 7-10 times using a one ml pipette.

For an accurate determination, the total number of cells overlying one 1 mm 2 should be between 15 and 50. If the number of cells per 1 mm 2 exceeds 50, dilute the sample and count again. If the number of cells per 1 mm 2 is less than 15, use a less diluted sample. If less dilute samples are not available, count cells on both sides of the hemocytometer (8 x 1 mm 2 areas).

Keep a separate count of viable and non-viable cells. If more than 25% of cells are non-viable, the culture is not being maintained on the appropriate amount of media. Reincubate the culture and adjust the volume of media according to the confluency of the cells and the appearance of the media. Include cells on top and left touching middle line. The cells touching middle line at bottom and right are not counted.

i. Trypan Blue is the "vital stain" excluded from live cells.
ii. Live cells appear colourless and bright (refractile) under phase contrast.
iii. Dead cells stain blue and are non-refractile.

  • %Cell Viability = [Total Viable cells (Unstained) / Total cells (Viable +Dead)] X 100.
  • Viable Cells/ml = Average viable cell count per square x Dilution Factor x 10 4 /
  • Average viable cell count per square = Total number of viable cells in 4 squares / 4.
  • Dilution Factor = Total Volume (Volume of sample + Volume of diluting liquid) / Volume of sample.
  • Total viable cells/Sample = Viable Cells/ml x The original volume of fluid from which the cell sample was removed.
  • Volume of media needed = (Number of cells needed/Total number of viable cells) x 1000.


The HTF isolation protocol presented here is a simple, fast method to obtain great number of cells (1.3 × 10 6 vimentin-positive fibroblasts) from a single 2–3 mm × 1 mm trabeculectomy biopsy within approx. 25 days. Our protocol may allow for easy setup of cell banks under laboratory conditions and for novel ophthalmologic drug testing in vitro. It was also demonstrated that 5% FGF-EMEM is a better choice than 10% DMEM for fast propagation of HTFs before the preparation of the experiments. However, in the case of the studies aiming to assess the effect of a tested agent on proliferation rate or type I collagen production ability, 5% FGF-EMEM should be used only for isolation of HTFs, and then 10% DMEM should be applied for experimental setup, as 5% FGF-EMEM significantly affects cell divisions and the ECM-forming ability of fibroblasts.

Specimen Preparation Protocols

Confocal microscopy was becoming more than just a novelty in the early 1980s due to the upswing in applications of widefield fluorescence to investigate cellular architecture and function. As immunofluorescence techniques, as well as the staining of subcellular structures using synthetic fluorophores, became widely practiced in the late 1970s, microscopists grew increasingly frustrated with their inability to distinguish or record fine detail in widefield instruments due to interference by fluorescence emission occurring above and below the focal plane. Today, confocal microscopy, when coupled to the application of new advanced synthetic fluorophores, fluorescent proteins, and immunofluorescence reagents, is one of the most sophisticated methods available to probe sub-cellular structure. The protocols described in this section address the specimen preparation techniques using synthetic fluorophores coupled to immunofluorescence that are necessary to investigate fixed adherent cells and tissue cryosections using widefield and confocal fluorescence microscopy.

Two days after tamoxifen induction, the epithelial cells in the interpapillary pit express random colors, indicating that multiple clones proliferated independently. However, after 84 days, each interpapillary pit was occupied by single-color cells, indicating that they are derived from monoclonal stem cells. (Nature Cell Biology 15, 511–518, 2013)

Image data courtesy of:
Hiroo Ueno(Ph.D.)
Department of Stem Cell Pathology, Kansai Medical University

Presented in Figure 1 is a laser scanning confocal image revealing the extensive filamentous actin network present in the smooth muscle tissue of a thin (8- micrometer) cryosection of rat diaphragm. The tissue cryosection was labeled with a cocktail containing Alexa Fluor 488 conjugated to phalloidin (staining actin) and Texas Red-X conjugated to wheat germ agglutinin (targeting lectins). In addition, nuclei in the specimen were counterstained with Hoechst 33342. Images were recorded in grayscale with an Olympus FluoView FV1000 coupled to a IX81 inverted microscope using Argon-ion (488 nanometer line), violet diode (405 nanometers), and green helium-neon (543 nanometers) lasers. During the processing stage, individual image channels were pseudocolored with RGB values corresponding to each of the fluorophore emission spectral profiles.

Staining Protocols

Triple-Staining Adherent Cells with MitoTracker, Phalloidin (or Phallacidin), and Nuclear Dyes

The vitality and physical properties of adherent cells grown on coverslips in Petri dishes can be determined using a combination of fluorescent stains. This protocol details a generalized procedure for staining a variety of cell types.

Staining Adherent Cells with Intermediate Filament Primary Antibodies and Synthetic Fluorophores

Fibroblast and epithelial cell lines derived from humans and laboratory animals produce brightly colored fluorescent specimens highlighting specific proteins in the intermediate filament network when stained.

Staining Cells and Tissue Cryosections with Tubulin Primary Antibodies, Phallotoxins, and Synthetic Fluorophores

Cell lines and mammalian tissue sections derived from humans and laboratory animals, including intestine, kidney, testes, muscle, liver, and the lungs, produce brightly colored fluorescent specimens.

Staining Adherent Cells with Cytokeratin Primary Antibodies and Synthetic Fluorophores

Epithelial cell lines derived from humans and laboratory animals produce brightly colored fluorescent specimens detailing the cytokeratin intermediate filament network when stained with keratin antibodies.

Triple-Staining Tissue Cryosections with Wheat Germ Agglutinin, Phalloidin, and Nuclear Dyes

A majority of the common tissue sections derived from laboratory animals, including intestine, kidney, testes, muscle, liver, and the lungs, produce brightly colored fluorescent specimens detailing a wide variety of anatomical features when stained.

Immunofluorescence with Brain Tissue Cryosections

Among the antibody targets often labeled in brain tissue sections are glial fibrillary acidic protein (GFAP), neuronal and axonal neurofilaments, class III beta-tubulin, microtubule-associated proteins, blood-brain barrier proteins, and antigens associated with specific diseases.

Immunofluorescence with Brain Tissue Floating Cryosections

Indirect immunofluorescence is a technique that enables the visualization of specific targets using a combination of primary and secondary antibodies. This protocol details a procedure for staining frozen brain floating cryosections greater than 30 micrometers in thickness.

Contributing Authors

Nathan S. Claxton, Gregory K. Ottenberg, John D. Griffin, Scott G. Olenych, and Michael W. Davidson - National High Magnetic Field Laboratory, 1800 East Paul Dirac Dr., The Florida State University, Tallahassee, Florida, 32310.

Flow Cytometry Protocol: Cell Surface Marker Staining

Note: This method is suitable for cell surface staining in flow cytometry.

Cell surface markers are expressed on the cell surface and can be used to define cell subtypes as well as function when they are labeled with fluorescent-labeled antibodies and analyzed by flow cytometry. As those surface proteins are accessible to the antibodies, they can be easily stained without permeabilization steps which are critical to intracellular staining.

  • PBS: Dissolve 8 g NaCl, 0.2 g KCl, 1.15 g Na2HPO4 and 0.2 g KH2PO4 in 800 mL distilled water. Adjust the pH to 7.4 with HCl and final volume to 1 liter with additional distilled H2O.
  • Cell staining buffer: Add 0.5% BSA and 0.05% Sodium Azide (NaN3) to PBS.
  • Fc receptor binding reagents
  • Conjugated primary antibodies
  • Conjugated secondary antibodies, if needed

Sample preparation:

1. Harvest the desired tissues and cells, prepare a single cell suspension and adjust the suspension to a concentration of 1 x 10 6 cells/mL in cell straining buffer.

Blocking Fc-receptors (optional):

2. Blocking Fc receptors is useful to reduce non-specific immunofluorescent staining. Add 1 μg of Fc receptors binding reagents per 1 x 10 6 cells and incubate for 10 minutes at room temperature.

  • For mouse cells, purified anti-mouse CD16/CD32 antibody specific for Fcγ R III/II (cat. no. CABT-45660RM) can be used to block the Fc receptors.
  • For human cells, purified human Fc receptor binding inhibitor solution can be used to eliminate non-specific staining.
  • In the absence of an effective/available blocking antibody for Fc receptors, an alternative approach is to block cells with excess irrelevant purified IgG from the same species and same isotype as the antibodies used for immunofluorescent staining.

Note: Do not wash excess blocking reagents from this reaction.

3. Add appropriately primary antibody with previously determined optimum concentration and vortex. Incubate on ice for 15 to 30 minutes.

Note: If using primary antibodies directly conjugated with fluorochrome, the incubation should be carried out in the dark, and then skip to Step 7.

4. Wash cells with cell staining buffer to remove any unbound antibodies. Centrifuge at 300-400 x g for 5 minutes at 4°C and discard supernatant.

6. Add an appropriate fluorescent-conjugated secondary antibody with recommended concentration. Incubate on ice for 15-20 min in the dark.

7. Repeat Step 4 three times.

8. Resuspend cell in 200-500 uL cell staining buffer for final flow cytometric analysis.

Immunofluorescence protocol (IF protocol)

Immunofluorescence is one of the widely used techniques in modern biology and medicine, and it is developed by Coons et al. (1950), and it is a combination of immunofluorescence technique and morphological technology to develop immune fluorescent cells (or tissue). The development of fluorescence immunoassay technology is very fast in recent years, especially in the field of medical, biological and environmental studies, it is widely used in the determination of endocrine hormones, growth factors, proteins, nucleic acids, neurotransmitters, receptors, in vivo drug and infectious sources, and so on, these subjects have been developed. According to the detection of different samples can be divided into three major categories.
There are two different immunofluorescence assay which include indirect immunofluorescence assay and direct immunofluorescence assay.For indirect immunofluorescence assay, the protocol mainly include tissue or cell preparation, tissue or cell fixation, serum blocking, primary antibody incubation, marked second antibody incubation, staining, result judgment and imaging. For direct immunofluorescence assay, there are only marked primary antibody been incubated without second antibody and other steps are same.
For direct immunofluorescence assay, specific fluorescent antibody was prepared by the combination of specific antibodies and fluorescein. It is the most simple and fast method for the examination of cell or tissue antigen. This method is highly specific and is commonly used in renal biopsy and pathogen examination. Its disadvantage is that a fluorescent antibody can only examine one kind of antigen, which is less sensitive. For indirect immunofluorescence assay, specific antibodies against the corresponding antigen, fluorescein labeled anti - antibody (anti - specific IgG fluorescent antibody) and the primary antibody
Although the basic steps and principles of immune fluorescence are the same, but because of the specific conditions are not the same, the detailed operation steps of each laboratory will not be exactly the same. For example, the use of the solution, the fixed liquid and antibody dilution liquid will be slightly different. Here to give a more common method, the detailed use of the operation can be based on the steps to adjust and change, and ultimately determine the most appropriate method for youself.

Indirect Immunofluorescence / IF

1. Prepare tissue or culture cells
2. Prepare tissue section or cells coverlip
3. Wash samples two times with PBS
4. Fix amples with 4% paraformaldehye in PBS for 15 min at room temperature(Note: Paraformaldehye is toxic, use only in fume hood)
5. Aspirate fixative, rinse two times in PBS for 5 min each
6. Permeabilize samples with 0.1-0.5% triton x-100 in PBS for 10 min(Note: Permeabilization is only required when the antibody needs access to the inside of the cells to detect the protein. These include intracellular proteins and transmembrane proteins whose epitopes are in the cytoplasmic region.
7. Aspirate triton x-100, rinse two times in PBS for 5 min each
8. Incubate samplels in 10% normal goat serum in PBS for 30 min at room temperature
9. Aspirate goat serum, incubate sections with primary antibody at appropriate dilution in PBS overnight at 4°C or 1 hour at 37°C(optimal condition should be confirmed in different laboratory)
10. Rinse three times in PBS for 5 min each
11. Incubate samplels with fluorochrome-conjugated secondary antibody at appropriate dilution in PBS for 1 hour at 37°C in dark(optimal condition should be confirmed in different l aboratory)
12. Rinse three times in PBS for 5 min each in dark
13. Incubate samples with 1 μg/ml DAPI
14. Mount samples with a drop of mounting medium

Direct Immunofluorescence / IF

1. Prepare tissue or culture cells
2. Prepare tissue section or cells coverlip
3. Wash samples two times with PBS
4. Fix amples with 4% paraformaldehye in PBS for 15 min at room temperature(Note: Paraformaldehye is toxic, use only in fume hood)
5. Aspirate fixative, rinse two times in PBS for 5 min each
6. Permeabilize samples with 0.1-0.5% triton x-100 in PBS for 10 min(Note: Permeabilization is only required when the antibody needs access to the inside of the cells to detect the protein. These include intracellular proteins and transmembrane proteins whose epitopes are in the cytoplasmic region.
7. Aspirate triton x-100, rinse two times in PBS for 5 min each
8. Incubate samplels in 10% normal goat serum in PBS for 30 min at room temperature
9. Aspirate goat serum, incubate sections with fluorochrome- conjugated primary antibody at appropriate dilution in PBS overnight at 4°C or 1 hour at 37°C(optimal condition should be confirmed in different laboratory)
10. Rinse three times in PBS for 5 min each in dark
11. Incubate samples with 1 μg/ml DAPI
12. Mount samples with a drop of mounting medium