Introduction to Immunofluorescence
A review of fluorochromes and the specialized microscope used in immunofluorescence techniques will begin this presentation. A typical (human skin and kidney) clinical specimen will be followed from receipt in the laboratory, through freezing, cryomicrotomy and staining. Direct and indirect IF procedures will be discussed in detail. Technical comments of the many steps required for this technology will be given to allow the technologist a better understanding of the detail of attention required for this technology. Photomicrographs of finished slides will conclude this presentation.
- Proper specimen handling.
- The importance of attention to detail.
- Reagents and supplies required for IF staining.
- Antibody staining patterns.
JAMES BURCHETTE, HT (ASCP) (Ret.):
Today we will have several learning objectives. We will review the proper handling of specimens, discuss the importance of attention to detail. This is very important in producing a quality product, all be it immunofluorescence (IF), or any laboratory technique. We will look at reagents and supplies required for immunofluorescent staining, and have images to look at for antibody staining pattern recognition.
What we're going to do is follow a specimen, follow our specimens from receiving to the final product. We will perform direct and indirect manual IF techniques on frozen tissue. There will be a brief discussion of formalin fixed, paraffin-embedded immunofluorescent technique. Throughout the presentation there are numerous technical notes, and there will be no flow, or molecular pathology discussed.
What is immunofluorescence?
It one of the most simple, simplest of all immunohistochemical techniques to perform. It is filtered light that excites a fluorochrome, a fluorescing compound, that is attached to an antibody. The most frequently use fluorochromes are fluorescein, or FITC, rhodamine, phycoerythrin (PE), Dylight, Texas Red. There are many Cy labels, Cy2, Cy3 are quite popular, and many Alexa Fluor fluorescent labels as well.
Some advantages and disadvantages of immunofluorescence. Advantages, It's a very rapid process. It has strong clinical implications. A clinician can get results back on a biopsy much faster than submitting formalin-fixed material and waiting for the processing, cutting, staining, and so forth. It's an easily performed technique, and there are many other clinical and research applications, especially using multicolor staining. We will just be discussing single staining in this presentation.
Some disadvantages, and I really don’t view these as disadvantage, but some places could look at like this. It does require an investment and a specialty microscope. It does require a designated viewing area, usually in a darkened area, a darkened room. It requires a microtome, a cryomicrotome, and you do need a technologist that has the ability to perform antibody dilution. I know of no company that sells ready-to-use fluorescent-labeled antibodies. And the clinical application, it does require a control tissue bank, and an ultra-low freezer for storage of antibodies and tissue blocks. And it does require your institution to be committed to this technology.
Here we have a fluorescence microscope. Power source here that I’ve got the arrow on, power source, a light source, digital camera, of course, your oculars and your objectives, a barrier shield to protect you from UV light, and in this area here you will have your bank of different excitation filters that are required for specific fluorochrome labels. And then a laptop or a PC to record and view the images as well.
Use log showing bulb hours; some microscopes actually will have a clock that will record how many hours are on a bulb. After a period of time the bulb loses intensity and needs to be changed. You should always wear gloves and safety glasses when changing the bulbs. The gloves prevent oil from your finger getting on the bulb and etching into the glass of the bulb. Keep a scheduled maintenance report file for CAP, or your accreditation inspections. I found that a Congo red histology special stain is a good slide to keep around the microscope because it fluoresces so well, and it can be used for microscope alignment.
Photography in the old days was used, was performed with 35 mm film that then had to be developed; whereas, today digital imaging is so much easier. You want to be aware of photo bleaching, so don’t burn out the area of interest by leaving the light source and the objective on directing the stream of light through the tissue. If the phone should ring, or you get interrupted, make you leave the objective on the tissue section with the light source on, it will burn that out. Do put multiple sections on the slide so you can find that optimal area for photography, for capturing the image.
A couple of notes on the history of immunofluorescence, the Fluorescence Foundation is a good source for reviewing fluorochromes and the history. Dr. Coons labeled the first antibodies in 1941 with fluorescein, thus giving birth to the field of immunofluorescence. And not much has really changed today since his early development of this technology.
Most of our tissue specimens we want to receive in the fresh state. In the clinical arena, the most commonly submitted specimens are skin, kidney, and then other biopsies you may receive are heart, lung, liver, and conjunctiva biopsies. If you’re in a reference lab, or even in the institutions where biopsies are taken when the laboratory is closed, shipping and storage of these fresh tissues is somewhat problematic, so you can use Michel’s Transport Media for preserving these tissue antigens and it allows convenient ambient temperature shipping or holding until you can process the specimen in the lab. Michel’s Transport Solution is a saturated ammonium sulphate solution which fixes tissues-bound immunoglobulins. Fresh specimens are immersed in a volume 20 X greater than the specimen size, and specimens can typically be held for up to one week at room temperature. There's no need to refrigerate these specimens. Upon receiving the specimen in the Michel’s Transport Media, you must wash the sections, the tissue samples in the buffer wash solution that removes the saturated ammonium sulphate, then it's ready for freezing and subsequent microtomy.
Here are two examples of dividing a renal biopsy. Sometimes you will receive a renal biopsy, the pathologist is not in the lab, or you’ve been instructed to divide the sample yourself. So you always want to handle the sections for immunofluorescence first to prevent contamination from your formalin or your glutaraldehyde for electron microscope (EM) samples. Always clean your forceps.
The most common embedding media is OCT (optimal cutting temperature compound), and you should have a cryostat chuck with a bed of OCT already frozen sitting in the bottom of the cryostat on the freezing bar. Place your sample on there, put more OCT on top of it, and use the heat extraction bar in the cryostat to rapidly freeze that block. Another freezing technique is using liquid nitrogen-cooled methyl butane. It's the same technique you use for freezing muscle biopsies. Dry ice acetone slurry is another low-temperature freezing alternative, and there are commercial freezing baths and proprietary solutions as well. I have a gelatin listed here as an embedding media, and I used that at Duke for 34 years, and actually it was used before my time in the immunofluorescence lab for many years prior to that. I will go into detail more on the gelatin embedding in just a moment. So you want to make sure your renal biopsy specimen pieces are embedded close to each other. That allows you to trim a block, giving a smaller cut surface area. That allows you to group your sections on the slide in a much tighter group, thus making it easier for the pathologist to locate the slide of the sections, makes it easier for you when handling the sections, and also the amount of antibody you need to put on the slide to cover the sections. As with all histologic technique, the skin is embedded on edge.
Embedding and Cryo-sectioning
A few comments about the embedding gelatin, you do not want to use house, kitchen-type gelatin, you want to use a 275 Bloom strength, which I actually had that on the next screen, but a rapid freezing technique, such as the isopentane and liquid nitrogen must be used. You do not want to use a slow freezing technique, like in a cryostat on the cold bar like for OCT. The gelatin will take on a totally different consistency and won't cut, so you must use a rapid-freezing technique. Small embedding molds must be used. It doesn’t work with large specimens and large embedding molds. Adjust your cryostat temperature to minus 15 to 16, and once you get the knack of cutting gelatin you can take serial sections. They're so easy, they just roll right off. An anti-roll device does not work with gelatin, you need to use a brush that’s been feathered down to just a few fibers to guide the section off the block as it goes across the knife blade. And warm water for your permanent slide, you’re hematoxylin and eosin (H&E) to wash off the embedding gelatin that’s surrounding the tissue section. It doesn’t interfere with the staining, but would leave a background on the glass.
So, 7.5% embedding gelatin using a 275 Bloom strength gelatin, that relates to how hard the gelatin sets up. Dissolve this 100 mL of water, 800 mL of water on a hot plate with a stir bar, add your 7.5 grams of gelatin, allow it to cool, and add 1 mL of 0.2% thimerosal, which is also known as Merthiolate, and store at 4 degrees Centigrade. I would keep a small plastic bottle with gelatin sitting on a hot plate, not boiling hot, but just warm enough to keep it liquified, or you can keep the gelatin in a water bath to keep it liquified. Do keep the cap tightly closed so it doesn’t evaporate, and evaporation can lead to a thicker gelatin solution and cause problems.
So, we've frozen our tissue, received our tissue, and have frozen it, and now we're ready to cut it in the cryostat. You want to prepare your slides, pre-label your slides with accession number, antibody, patient name, whatever your institution or lab requires. Use a permanent ink marker, because pencil will give graphite debris and it will smudge during handling. Like I said, trim your block down to a smaller size. Cut at 3 to 4 microns, and maintain order throughout the entire process, during your cutting, staining, cover-slipping, placement in the folder. It helps with trouble-shooting, and maintaining order in the whole process. If you snap freeze in a low-temperature freezing system, allow your blocks to acclimate to the cryostat temperature. They can be too hard and need to acclimate. So use a slow, even cutting stroke, and I use room-temperature slides to pick up the sections. You want to quickly pick up the sections and keep the frost wiped up on the blade holder. If the frost builds up on the blade holder you can have your sections stick to it, and then when you lift them up it causes a stingy, distorted artifact on your tissue sections. Again, place multiple sections on your slide and group them close together.
This illustrates a cutting sequence. I have slides labeled, sitting on top of the cryostat. I'll use a polychrome slide, and more about polychrome in a moment, but it's a quick reference, quick stain, quick reference slide to identify your tissue, or if you’re cutting a renal biopsy looking for glomeruli, it saves a lot of time from having to perform an H&E, check it under the scope, and again, more on that later. But you’ll have your polychrome and a permanent H&E slide for your first sections, and then your antibody slides just go one, two, three, four, five, six, seven, and then go back and put another single section on through the entire panel. That way you have multiple levels. This does show a pre-cut positive control on each of these slides. More on controls later.
Here's an example of a polychrome methylene blue on a frozen section of kidney. Our glomerulus, the mesangium is this matrix inside the glomerulus. Surrounding the glomerulus we have our basement membrane, or Bowman’s capsule, and then we have our proximal and distal tubules out here.
Like I said, this is a very rapid stain for frozen sections. It's available commercially, but it's also easy to make for itself. I’ve never had it go bad. It improves with age, like a fine wine. You do want to filter before use to remove any particulates. It is a little stinky when you make it up, but it's worth making yourself. You get a much better product. The procedure is you cut your section, pick it up on a slide, dip the slide in the stain for five seconds, rinse the excess stain from the slide with water, and then you can either wipe the back of the slide and look at it under the scope as is, or you can put a cover glass on the slide, mounting with water. And the sections are not permanent. A polychrome can be rendered into an H&E by placing it in the H&E fixative, and then proceeding with the frozen section H&E stain of choice.
Here is my frozen section fixative formula. It contain methanol, formaldehyde, and acetic acid. For optimal results, cut your frozen section and immediately pick it up, and immediately place it into the fixative for 60 to 75 seconds. Rinse it in warm tap water and proceed with your staining procedure of choice.
Okay, we've cut our sections after freezing, now we want to scribe the tissue sections. You do that so you know where the tissue sections are when you’re handling the slides. The pathologist, when the pathologist is looking at the slides in the fluorescence room, the circle will actually refract a little bit of light allowing him to know those sections in that circle. When you’re scribing the slides with your diamond-point pen or your carbide-tipped pen, you want to support the slide in this area here, along the sides, so you’re supporting the slide to prevent it from snapping across and breaking. Scribe the slides while the block is in the cryostat in case you happen to break one, and eventually everybody does at one time or another, that way the block is still faced up to the knife edge. You can pick up new sections on another slide and keep moving forward.
Positive controls must be run in the clinical lab. You can use a known case that has come though the lab that’s positive for a particular antigen; tonsil, spleen, lymph node can be used. If you have an autopsy service and you know of a patient that succumbs from lupus, and has lupus nephritis, multiple, multiple blocks of kidney can be frozen and used for a control tissue. I would always pre-cut a box of slides and store them in an ultra-low temperature freezer. When removing those from the freezer, you want to lay them out flat so they rapidly thaw and any condensation that forms on the slide quickly evaporates. And you can use one set of controls per batch of slides being run.
We've cut our tissue block now, let's seal the cut surface of the block with some OCT to prevent the tissue from becoming freezer burned. Some people wrap the frozen block in aluminum foil. I don’t adhere to that, because my logic is the heat from your fingers transmitting through the aluminum foil partially thaws the block, the frozen tissue block. Your major distributors that sell the ultra-low freezers all stock file systems, stainless steel racks, and fiberboard boxes. You can put the beginning and ending accession number on these boxes for retrieval of specimens at a later time.
So we're getting to where we're going to start our staining. Some of the reagents we need will be, of course, our fluorochrome-labeled antibodies, some antibody diluent, our procedural buffer, mounting media to coverslip the slides with, and acetone to be used as our section fixative. Our fluorescein conjugated antibodies are the most commonly used ones in the clinical arena for your direct IF panels on kidney and skin. Some of the more common antibodies are anti-human IgG, IgM, IgA, kappa lambda, albumin, fibrinogen, C3 and C1q. There's others that can go in there, such as IgE, it's all what your pathologist wants in their panel, but these are more of the common antibodies used.
Storage of antibodies, we want to keep them frozen at ultra-low temperatures. Use polypropylene tubes, polystyrene tubes will crack on you when frozen. They will shatter when centrifuged at ultra-high speed centrifugation. Centrifuge if needed, if you notice any debris on your sections, label your thawed tubes with the date they were thawed, and store the test tube. And again, I emphasize, maintain order. These two schematics show our tissue section with the antigen that we're trying to demonstrate in our tissue section. Here is our primary anti-human antibody with a fluorochrome label attached to the FC receptor. So our primary antibody would be applied to the tissue section, wherever its target site is it would bind to. We rinse the slide with buffer, coverslip it, and we would see a signal where the antibody is bound.
Our indirect technique uses an unlabeled primary antibody, followed by anti-species specific secondary antibody. Example, if this is a rabbit primary, the secondary is going to be anti-rabbit, again, with a fluorochrome tag attached to it. As I mentioned earlier, the primary antibodies, fluorescein-labeled, fluorochrome-labeled primary antibodies are virtually unknown out there that are pre-diluted, so we have to dilute them ourselves. I grew up in the immunopathology lab where we made everything ourselves, especially in the early days when there weren’t ready-to-use reagents. So here's a formula for a simple antibody-diluting buffer that I used for years. You want to use a good grade bovine albumin, and prepare a 1% solution with sodium azide as a preservative.
Procedural buffers for our technique can either be Tris-buffered saline, or phosphate-buffered saline. Both are available as a commercial ready-to-use, or a 10X, 20X concentrate that you can then dilute in your laboratory. Of course, you can always prepare them from stock reagents or in-house preparations.
Okay, we're ready to start staining, so some materials that we need are some slide wipe tissues, some Pasteur glass pipettes, and I like the 5 ¾ inch glass Pasteur pipettes with a bulb that you actually place on the pipette. I feel it gives you more control over the dropping of the antibody onto to slide, as compared to a plastic transfer pipette. We will need a squirt bottle to hold some procedural buffer, a waste cup to catch our rinsing of the slides, a carboy to hold our bulk buffer, a timer, a fume hood for our acetone fixation, and also it's good for drying your slides in, Coplin jars, diamond-tipped pen, a humidity-controlled incubation chamber, an absorbent towel to drain the excess buffer onto, a test tube rack to hold our tubes of antibody, charged slides for our sections, and of course, cover glass, and more on cover slipping a little bit later.
As I said, there's a lot of wiping and handling of the slides during the procedure. And you can take a Kim-Wipe tissue and fold, this is a whole Kim-Wipe, so fold in the center line, then fold again in the center line, fold it again, and fold it again until you come up with a little final square. You have multiple layers of tissue that are very absorbent, and the corners here work really well when you’re working around the circles on your slide, removing the excess buffer prior to putting your antibody on. So in this old picture here of this young man that has hair and no gray, you can see his incubation chamber here, some moistened towels with water creating the humidity. His test tube rack with antibodies, a squirt bottle with PBS, a waste cup, Coplin jar, some slides are drying over here about to be fixed with acetone, and a little bottle embedding gelatin on a HISTO/Orientator little heating block.
So we're going to look at our direct and indirect procedures. We've cut our slides, air dried them for 30 minutes, a minimum of 30 minutes. And you can prop them up on a rack, against a test tube rack inside a hood, and lower the sash so the air blows across them. They dry very rapidly like that. We want to place our slides in fresh acetone, room-temperature acetone for five minutes. Using cold acetone, as some protocols call for, can lead to a lot of different issues; such as, condensation forming in the acetone, using a Coplin jar containing acetone inside the cryostat, and you’re working in the cryostat, you’re breathing fumes. So I did a comparison of room temp versus cold acetone, and there's no change with using room temp acetone. In fact, that’s what we always used for fixing lymph nodes when doing frozen section immunophenotyping. So it works perfectly well for immunofluorescence. Following acetone fixation, we want to remove the slides, allow them to air dry, five minutes is more than enough, place them in our protocol buffer. From this step on you do not want the sections on the slide evaporating or drying. It will cause auto-fluorescence and a background. So we're ready to proceed with our staining protocol.
So 1 through 4 is what I just explained, staining with step 5, we're applying our, in this case a fluorescein-labeled antibody. And we're going to incubate 30 minutes. At the conclusion of that 30 minutes we will rinse our slides with our buffer, and then place them in a Coplin jar with fresh buffer. And we want to coverslip, and I'm using a glycerol Tris buffer cover-slipping medium that I prepare myself. And store the tissue sections at 4 degrees in a closed holder to prevent the light from fading any of the fluorescence. Here's an example of a skin biopsy from a patient with lupus, with C1q antibody. The granular staining pattern at the base of the dermis, the epidermis, the dermal epidermal junction. This is a piece of debris right here and here. Debris happens. It can come from dust, from your finger nails, squamous epithelial cells. For our indirect, two-step indirect immunofluorescence, 1 through 4 is just as we’ve previously discussed. Starting with step 5 we want to apply an unlabeled primary antibody, incubate 30 minutes, rinse that antibody from the slide, do a buffer wash, apply our species specific, anti-species secondary antibody that’s labeled with your label of choice, incubate 30 minutes, rinse, wash, coverslip, store or view. So here's an example of an indirect. We have an unlabeled rabbit anti-human herpes simplex virus primary antibody. Followed up with goat anti-rabbit. So rabbit is our host species of our primary that this antibody was made in. And our secondary has to be anti-rabbit. In this case our host species is goat, but we're using goat, anti-rabbit IgG, FITC-labeled secondary. Here's an example of that in a HSCD infection in the brain, showing some infected astrocytes.
Another commonly used indirect immunofluorescence these days is using the C4d antibody for rejection. So we have a mouse anti-human C4d primary. Our secondary is goat anti-mouse IgG with an Alexa Fluor label, or a goat anti-mouse with a Cy2 label. And here's an example of peritubular capillaries in a renal transplant case of antibody-mediated rejection expressing the C4d antibody. And just for comparison, I put in this immunohistochemical stain with a DAB chromogen for C4d.
So, a couple of troubleshooting topics. Drying your slides post-microtomy, you will have poor staining if you don’t dry them, so make sure you do that minimal 30-minute drying. Allowing your buffer to dry on the slide after fixation can lead to auto-fluorescence and stain edge staining artifact. Fixation, use fresh acetone. Freeze, thaw of the tissue sections can alter the integrity of that biopsy. Staining debris, unbound fluorescein products, also cover slipping, too much mounting media, it leaching out, oozing out from under the coverslip and getting on top of the coverslip and making a mess.
So real quick here, we're starting to get short on time, so I'm going to have to speed things up. Indirect immunofluorescence on formalin-fixed paraffin embedded; perform your pre-treatment, heat retrieval, enzyme retrieval, whatever is needed for that particular antigen. Following pre-treatment we have a buffer wash, a non-immune serum blocking. We want to apply the primary antibody for 30 to 60 minutes, buffer rinse and wash. Apply a specific labeled secondary antibody. You can make 30-minutes buffer rinse, wash, coverslip, and again, store at 4 degrees in a closed folder until ready to view.
Proteolytic enzyme digest can be trypsin, pepsin, proteinase K. Heat-induced epitope retrieval can be your typical high and low pH used on paraffin sections for immunohistochemistry. Quenching of auto-fluorescence with sodium borohydride, or copper sulphate, and you can Google these for more information.
A couple of additional comments on troubleshooting on formalin-fixed paraffin-embedded material, I can emphasize enough, use the non-immune blocking serum prior to the application of the primary antibody. When using a mouse monoclonal antibody, always use an anti-mouse isotype-specific labeled secondary antibody. An example there would be the mouse anti-human E-cadherin is an IgG-1 kappa isotype. So you would want an anti-mouse IgG-1 kappa isotype. In this case our host species is goat, and normal goat serum would be the type of non-immune serum that we would use as our serum blocking step. That alone is the best tip I can give anybody for performing immunofluorescence on formalin-fixed material.
IF mounting media, there are commercial solutions available, such as Glycerogel or Vectashield. Some have a counter stain, some don’t. Some contain an anti-fade chemical property to reduce fluorescence fading. There are in-house preparations. None of these are permanent. Here is my glycerol Tris mounting media for fluorescent studies, 0.25 M Tris buffer and glycerol. You want to mix this up well ahead of time to allow any air bubbles to dissipate.
When cover slipping, do not allow your tissue sections to dry. Mount from your procedural buffer, drain the edge of the slide on a paper towel to absorb off the excess mounting media. There's no need to seal with nail polish or Vaseline. It's a wasted step and just makes a mess. And again, maintain order throughout the entire process. Use the smallest coverslip you can to keep the pathologist’s fingers away from the cover glass. They have a tendency to put their fingers on the cover glass and move it, and make a mess with it. And, “Can you re-coverslip that slide for me.” “Damn, I worked hard to get that thing just right, and there you go, messing it up,” so use a small cover glass. A little note for your folders, use a post-it note, put your accession number and date down on the bottom and stick this inside the flap of the folder. That way when the folders are stacked up inside the refrigerator you can rapidly look down the stack and find a particular case to retrieve.
As I mentioned earlier, Congo red is a fluorescing compound. Calcafluor White is a product used in textiles that has an immediate reaction that recognizes yeast, fungi, and parasites.
So real quick, a few staining examples. A section of skin for immunoglobulin A, a granular deposit at the dermal, epidermal junction, a C1q, complement 1q, and a section of skin for lupus, again, a granular pattern. A more linear staining pattern with C3 and a case of Bullous pemphigoid in skin, again, localizing at the dermal, epidermal junction. Pemphigus localizes in the intercellular or intraepidermal spaces in the epidermis. And another example of pemphigus with immunoglobulin gamma in the epidermis of skin. You can also receive oral mucosal biopsies and for pemphigus, pemphigoid. Vasculitis disease for IgM in skin. Serum indirect, where the patient’s serum is serially diluted and applied to a section of monkey esophagus. So these desmosome-type cells here are the same type of cells that the antibody in the patient’s serum would bind to in a skin section. And serum indirect for pemphigus in the intraepidermal area in this stratified squamous epithelium in esophagus. Kidney immunoglobulin gamma in this linear staining pattern, staining these mesangial deposits in a glomerulus. IgG in membranous glomerulonephritis, and a granular pattern of C3 in post infection glomerulonephritis. And this is I think a very pretty staining pattern. It's a linear staining pattern of IgG in glomerulus with Goodpasture’s syndrome.
Some references: Good buffer and protocol references in Lab Rat, and IHC World. Pathology Student is a great site. Subscribe to that. A few references from projects that I’ve been involved with that used immunofluorescence techniques. And even though this reference here says Immunocytochemical Detection of Lymphocyte Surface Antigens in Fixed Tissue Sections, that was actually Michel’s Transport Media used there, not formalin-fixed material.
I'm sorry I had to kind of speed things up, but I was running out of time. I thank you very much for sitting in and hearing me speak on this. I hope I was able to provide to some insight to help you with your immunofluorescence. Thank you very much.
Questions and Answers
Can you please explain and the serum indirect procedure?
Yes. A patient has circulating autoantibodies in their system. A red stopper tube of blood is drawn from the patient. It's allowed to clot. We would then spin it down, take off the serum, dilute it 1:2, 1:4, 1:8, 1:16, 1:32, 1:64, 1:128, and each one of those dilutions would be applied to a section of monkey esophagus. The section would incubate with the patient serum, be washed off, then we would follow it up with a secondary antibody, such as goat anti-human, or rabbit anti-human IgG that was labeled with fluorescein. And wherever our patient serum would bind to that cell type in the tissue for that particular disease would then bind with the anti-human antibody, and we would get a signal there.