You want the best slides possible for your staining experiments. A lot of hard work is done to make sure your sample is preserved under the proper conditions after harvesting, grossing, and staining. At the end of the staining protocol when it’s time to view your slides and image under a microscope, or save them for a later date, it’s important your tissue samples are protected. That’s where coverslipping comes into play.  

Although coverslipping is the last step in the staining process, its importance cannot be understated. Properly coverslipping your slides ensures that your tissue specimens are preserved for optimal imaging and viewing.  

In this blog post, we will walk you through how to properly coverslip your slides to protect your samples. Be sure to also watch the video at the end of the blog post to see how to mount your slides. 

Keep your slides in buffer before coverslipping 

When you reach the end of your staining process, the protocol usually requires you to run your slides through a series of clearing agents, such as xylene. If you are not going to immediately coverslip your slides, you should leave them in the solution. This will prevent your slides from drying out and losing the stain you worked so hard to obtain.  

For some IHC protocols, such as H&E, leaving your slides in the solution for hours after staining is okay. If you’re doing an IF protocol, you should aim to coverslip as soon as possible to preserve your fluorescent signal. 

Remove excess buffer 

Once you are ready to coverslip your slides, you should remove the excess clearing agent. This can be done by removing your slide from the solution and gently dabbing the slide vertically against an absorbent material such as a Kimwipes™, paper towel, etc. You can also use the same absorbent material to remove excess from the back of the slide so it is easier to handle. 

Cover your tissue sample with mounting media 

After removing the excess clearing agent, you should immediately mount your slides with the appropriate mounting media. When adding the media, take extra care not to introduce air bubbles in order to prevent having bubbles in your images when you photograph them later. You should also be mindful of how much mounting media you’re using. You should use enough mounting media to cover the specimen, but not so much that excess media will leach outside of the coverslip.  

Gently apply coverslip 

It is important when applying the coverslip that you do so as gently as possible. Using too much force can tear your specimen and cause artifacts on the slide. To gently apply the coverslip, hold the top and bottom of the slide in one hand between your thumb and index finger. You can do this vertically or horizontally. Using your other hand, angle the bottom edge of the coverslip to a 45-degree angle at the bottom of the slide. Slowly and gently lower the coverslip onto the slide so it covers the sample.  

In using this technique, not only do you reduce the possibility of damaging the tissue, but you reduce the air bubbles that might occur underneath the coverslip. If there are any air bubbles present, you can gently press the coverslip to expel them, but be careful not to press too hard or do this directly on the sample as this can damage the tissue. 

Optional step: Depending on the mounting media you use, you may need to seal your coverslip. If you use a mounting media such as VECTASHIELD® Antifade Mounting Media, sealing the coverslip is not necessary. 

In just a few short steps, you can properly coverslip your slides and have peace of mind that your samples are preserved for whatever your experimental needs are. For other tips and tricks for your slides, check out our Staining Sample Prep Guide as well as other blog posts. 

Check out this video to see how to properly coverslip your slide and get high quality slides for viewing. 

Antibodies are blood proteins produced in the immune system in response to a specific antigen. This specific antigenicity is what makes antibodies such a powerful tool in immunochemistry. The earliest reference to antibodies goes back to the 1890s. Emil von Behring and Shibasabura Kitasato discovered that transferring serum from a diphtheria-immunized animal to animals infected with diphtheria altered the course of the disease (1). Since the 1890s groundbreaking strides have been made in the world of antibodies and their applications in life science and health science fields. With a wide variety to choose from, there are important considerations to make when selecting your own primary antibody. Keep reading to learn how to choose a primary antibody for your workflow. 

Antigen of Interest 

An antibody is classified as a primary antibody if it binds directly to the target antigen of interest. In comparison, a secondary antibody is an antibody whose antigen is another antibody. The primary antibody must be raised against the target antigen, otherwise, there would not be any successful detection. The sample processing should also be addressed. For example, some antibodies only recognize proteins in their denatured state, thereby revealing the epitope that would otherwise be obscured. Other antibodies may require proteins in their folded state. Additionally, some primary antibodies may require an antigen retrieval step to reverse cross-links introduced by formalin-fixed, paraffin-embedded tissues. These characteristics of the antibodies should be considered during experimental design. 

Clonality 

Next, it is important to choose the clonality of the primary antibody—whether it will be monoclonal or polyclonal. In monoclonal antibody production, individual B-cells are isolated from an immunized animal, cloned, and grown to produce homogenous antibodies (2). These monoclonal antibodies not only target the same antigen, but a single epitope, aka binding site, on the antigen. In comparison, polyclonal antibodies are purified from the serum of an immunized animal, so the antibodies have a mixture of epitope targets of the same antigen (3). A monoclonal primary antibody will give more specificity to the target with less crosstalk with other proteins. Additionally, since the monoclonal antibodies are derived from cultured cells, there is less variability from lot to lot. The benefit of polyclonal antibodies is greater tolerability in target protein conformation (fixation type, temperature, pH, etc.) and higher sensitivity. 

Host Species 

The host species of the primary antibody will have important implications if a secondary antibody is used in the detection system. In this case, it is most common to choose a host species different from the species of tissue being tested. For example, when detecting an antigen in rabbit tissue, a mouse-raised primary antibody is an appropriate host species option. Specific considerations must be taken when the primary antibody is raised in the same species as the tissue to prevent cross reactivity between the secondary antibody and the tissue. For example, if a mouse-raised primary antibody is used on mouse tissue, the secondary antibody, which must be anti-mouse, will detect endogenous mouse Ig in the tissue sample, leading to non-specific detection and potential false positives. This is addressed with certain blocking reagents, but experimental designs can be simplified by choosing an alternate host species of primary antibody. 

Class 

In mammals there are 5 main classes of antibodies: IgG, IgM, IgA, IgD, and IgE, which each have a unique structure. Physiologically, each class of antibody has unique biological properties, locations, and antigenicity (4). In immunochemistry staining, it is important to note the class of the primary antibody because the secondary antibody must be directed against the class of the primary antibody. This is especially important in monoclonal antibodies. 

Labeling of the Antibody 

Labels are conjugated to antibodies to make visualization of the target antigen possible. Labeling can occur on the primary antibody, secondary antibody, or subsequent amplifying reagents. The most typical labels are either a reporter enzyme for immunohistochemistry or a fluorophore for immunofluorescence. These allow for either color development or fluorescent signal at the site of the antigen. 

A primary antibody can be directly conjugated with a label for visualization, removing the need for secondary antibodies. While possible, it is more likely to use a conjugated secondary antibody or amplification reagent, so the primary antibody can remain unconjugated. This increases sensitivity and allows the primary antibody to be used in lower concentration, therefore saving money on an expensive reagent. 

There are many factors to consider when choosing a primary antibody and this is not an exhaustive list. Understanding each of these characteristics is important when developing an immunochemistry protocol and selecting a primary antibody.  

We hope this blog post gave you valuable insights on primary antibodies. For more resources, check out our IHC and IF Resource Guides, and be sure to stay tuned to the blog for more tips and tricks.  

References 

1. Kaunitz JD. 2018. The Development of Monoclonal Antibodies. Digestive Diseases and Sciences. 
2. Köhler G, et al. 1975. Continuous Cultures of Fused Cells Secreting Antibody of Predefined Specificity. Nature. 
3. Leenaars M, et al. 2005. Critical Steps in the Production of Polyclonal and Monoclonal Antibodies: Evaluation and Recommendations. Institute for Laboratory Animal Research Journal. 
4. Schroeder HW, et al. 2010. Structure and Function of Immunoglobulins. The Journal of Allergy and Clinical Immunology. 

Back in my days as an undergraduate student, I learned to use nail polish to create a hydrophobic barrier on my immunohistochemistry (IHC) and immunofluorescence (IF) slides. The idea was to apply a thin layer of polish around the tissue to reduce the volume of reagents required to complete the experiments. But it just didn’t work that well, at least not for me. I didn’t have the fine motor skills necessary to draw a thin line using a thick brush. I’d smudge polish all over the slide and sometimes on top of the tissue section. Even if I managed to spare the tissue, the polish layer would often get too wide or thick, and I would have to peel it off before applying the coverslip. The entire process was time consuming and frustrating. In my opinion, trying to use nail polish to create a hydrophobic barrier added more trouble than value.   

Needless to say, I was grateful when the first PAP pens—pens that deliver a hydrophobic liquid—started to hit the market. After the liquid dries, it forms a thin film that effectively repels fluids, helping reagents stay confined within the delineated area. The quality of the available PAP pens has improved over the years, and now you can’t go wrong. They are straightforward to use, and creating a hydrophobic barrier adds little time to the overall workflow as they can dry in as few as 2 minutes. 

PAP pens bring many benefits to IHC and IF workflows. Here are 5 reasons you should add the ImmEdge^® Hydrophobic Barrier PAP Pen from Vector Laboratories to your list of essential lab tools as well as a video tutorial on how to use ImmEdge pens.  

ImmEdge PAP pens save reagents and reduce the cost of running IHC and IF experiments 

The purpose of creating hydrophobic barriers is to reduce reagent usage and the cost of IHC and IF experiments. The math is simple: Use half of the volume of reagents to completely cover tissue sections means a 50% savings. But no percentage of savings makes the risk of damaging your tissue and ruining your experiment worth it. PAP pens eliminate the hassle and make the goal of saving reagents and research money possible. 

ImmEdge PAP pens give you flexibility 

With PAP pens, you can physically isolate different tissue sections on the same slide and incubate them with different reagents. This can be handy in many scenarios, such as when comparing different primary antibodies against the same epitope. During the visualization process, you don’t need to switch between several slides and can simply slide across 2–6 tissue sections. As a result, imaging the sections and comparing the results from each antibody is easier and quicker. The same applies to titration experiments. Specimen visualization and image acquisition will flow much faster when you don’t need to keep switching between multiple slides.  

ImmEdge PAP pens are compatible with buffers and reagents in the staining workflow 

Unlike nail polish, most PAP pens are resistant to alcohol and acetone. Therefore, you can use PAP pens to create hydrophobic barriers at different steps of your workflow knowing you’ll get consistent results. Most PAP pens are also insensitive to detergents (i.e., Tween^® 20, Triton X-100) and compatible with enzymes and fluorescence-based detection systems. In addition, hydrophobic barriers won’t interfere with fluorophores and antibodies. Removing the hydrophobic barrier before applying the coverslip is unnecessary. Still, if you need to clean the PAP pen film for any reason, you can use any clearing reagent, such as xylene. Be sure to check the manufacturer’s info sheet to confirm the compatibility of a specific PAP pen.  

ImmEdge PAP pens are heat-stable  

The ability to endure high temperatures makes PAP pens applicable to other techniques, such as in-situ hybridization. Adding hydrophobic barriers to in-situ hybridization brings the same cost-reduction benefits as in IHC and IF experiments. However, the selected barrier method needs to offer stability at the hybridization temperature indicated in your protocol. Heat-resistant PAP pens also give you the flexibility of applying the hydrophobic barriers before performing heat-induced epitope retrieval.  

ImmEdge PAP pens are environmentally friendly 

In addition to all the benefits above, ImmEdge Hydrophobic Barrier PAP Pens are also an environmentally friendly option. They deliver high-quality results, but are free of ozone-depleting hydrocarbons. You can rest assured that your choice to save reagents and research funding won’t come at the expense of the environment.  

Creating hydrophobic barriers has become much easier since I first started running immunostaining experiments. Using PAP pens effectively reduces reagent usage and experiment costs without impacting your workflow. If you want to learn other strategies to optimize your staining workflow, check out the Staining Sample Preparation Guide and stay tuned for other tips and tricks from the blog. 

Not convinced how easy it is to use the ImmEdge PAP Pen? Watch the video below to see how it’s done. 

It can be frustrating to get to the end of an immunohistochemistry (IHC) or immunofluorescence (IF) experiment just to find out the staining didn’t work. While there are some standard steps to take to solve problems you might come across, sometimes you need to look outside the box. Here we discuss some miscellaneous factors to focus on when an IHC or IF protocol performs unexpectedly. 

This blog post is the third of a three-part series intended to help researchers navigate different parts of the IHC and IF staining processes that might present challenges. You can check out the first part of the series, “Background staining in IHC and IF“, and the second the second part of the series, “Absence of positive staining in IHC and IF“. 

Species-on-species 

With modern techniques, antibodies can be raised in a variety of species: mouse, rat, goat, horse, and more. Species choice is important to minimize cross-reactivity, but sometimes it is inevitable to work with species-on-species reagents and tissues. For example, it is problematic to use a mouse primary antibody on mouse tissue because the anti-mouse secondary antibody will recognize endogenous mouse immunoglobulins in addition to the primary antibody (1,2). Certain kits, such as Vector M.O.M.® (Mouse on Mouse) Immunodetection Kits will block endogenous mouse immunoglobulins, making it possible to fluorescently detect one or more mouse primary antibodies on mouse tissue.  

Unwanted species interaction can also occur with xenograft models, for example the study of a human tumor grown in a mouse model (3). While the tissue itself is human derived, it will be vascularized with mouse blood vessels and contain mouse immunoglobulins. Applying an anti-mouse secondary antibody may produce cross-reactivity with the xenograft tissue, leading to potential false positives. In this situation, similar M.O.M. reagents can ameliorate the issue. Anti-rat and anti-hamster secondary antibodies also have a degree of cross-reactivity with mouse tissue because they are related rodent species. To find out how to eliminate unwanted binding of a closely related species, check out our first blog post from this series. 

Counterstain 

Counterstaining IHC/IF samples spatially orients the target within its surroundings, which can help guide researchers as they analyze their results. Ensure judicious use of a counterstain, as heavy staining may mask weakly expressed antigens. Additionally, too similar of a substrate color can obscure target antigens causing false positives or false negatives. To navigate these challenges, make sure the counterstain and initial detection systems provide adequate contrast and have compatible protocol parameters such as solubility and heat resistance, if applicable. Researchers should run counterstain-only controls to validate their chosen counterstain performance. Running a titer can help determine the best counterstain protocol prior to proceeding with the antigen targets.  

Multiplexing 

IHC and IF protocols can be adapted to detect more than one target in a sample, allowing detailed analysis of different targets in parallel. With these more advanced experiments, keep in mind some not-so-obvious steps which may negatively affect your stain if not addressed.  

Watch out for potential incompatibilities between secondary antibodies. While it is usually no problem to use two secondary antibodies raised in the same species, issues may arise with some antibody combinations. For example, a secondary antibody raised in horse in the initial labeling reagents and an anti-horse secondary in the subsequent labeling reagents would lead to cross reactivity. To address this without changing antibodies, run the staining workflow sequentially through to the initial system development prior to the subsequent labeling system. 

 
In an avidin/biotin detection system, it can be necessary to block endogenous avidin and biotin to mitigate background noise (4). This is typically done before the first primary antibody incubation, but when detecting multiple antigens, it is necessary to block before each primary antibody to prevent the subsequent set of labeling reagents from interacting with the prior set of labeling reagents.  

Once the detection systems are determined, it is best to optimize single labeling protocols prior to combining them into a multiplex protocol (1). This optimization has a second benefit: the stained sections from the single label protocols can be used as controls to compare the multiplex staining quality. 

Microscopy setup 

IHC slides are typically viewed under a bright-field microscope. There are some parameters to keep in mind, like the brightness adjustments and condenser, but there are more specific parameters to check off with fluorescent microscopy in IF. It is necessary to define the proper excitation and emission spectrum for targets. Each fluorophore emission spectra should ideally have a narrow peak with minimal overlap to optimize each signal distinctly (5). Ensure the appropriate filters are chosen to maximize the target fluorophore emission intensity while minimizing the unwanted signal from other fluorophores. 

Miscellaneous 

Sometimes even the miscellaneous category has a miscellaneous category. Here are some extra steps to review when assessing why the staining did not work.  

IHC and IF kits are widely available and offer a convenient number of compatible products for a protocol, but not all kits cover every step. Ensure every step from beginning to end is accounted for, including all the necessary ancillary components, and confirm compatibility throughout the full workflow. 

Each product will have specific storage conditions and expiry dates. Using a product outside of these specifications can negatively impact the stain results. Take care to store all reagents as recommended and use them within the denoted expiry date. Key factors to keep in mind are storage temperature, protection from light, and reduction of freeze-thaw cycles. Additionally, while a product concentrate may be stable for many months, the working solution may need to be made fresh prior to each use. 

While this is not an exhaustive list for why a staining did not turn out as expected, it contains great clues for the detective’s journey to solving the case. For more tips and tricks to improve your staining, check out our IHC and IF Resources, and stay tuned for more help on the blog. 

References

1. Van der Loos, CM. 2007. Multiple Immunoenzyme Staining: Methods and Visualizations for the Observation With Spectral Imaging. Journal of Histochemistry & Cytochemistry.  
2. Goodpaster T, et al. 2013. A Flexible Mouse-On-Mouse Immunohistochemical Staining Technique Adaptable to Biotin-Free Reagents, Immunofluorescence, and Multiple Antibody Staining. Journal of Histochemistry & Cytochemistry.  
3. Cekanova M, et al. 2014. Animal Models and Therapeutic Molecular Targets of Cancer: Utility and Limitations. Drug Design, Development and Therapy.  
4. Bussolati G, et al. 2003. Retrieved Endogenous Biotin: A Novel Marker and a Potential Pitfall in Diagnostic Immunohistochemistry.  
5. Evans K, et al. 2013. Chapter 9.11 – Hematology. The Immunoassay Handbook (Fourth Edition).