Histology Hacks for Cryosectioning
Histology laboratorians in the research world move fast, but those who perform cryosectioning know that these tissues require lighting speed and precision.
Cryosectioning is the artform of sectioning flash-frozen tissue, which aids the visualization of fine details in cells after staining.
Research environments require flexibility and adaptability for sectioning diverse tissues and materials. Cryosectioning is often the preferred methodology, providing superior antigenicity and more accurate detection of antigens in downstream applications. Automation has changed the game in cryosectioning, offering reproducibility and high-quality sectioning over manual methods.
In this webinar, Dr. Marla Rivera-Oliver shares some “Histology Hacks” to properly prepare tissue, an essential but often overlooked element of cryosectioning, and guide practitioners to achieve better cryosectioning outcomes in the research environment.
In this webinar, you will learn:
- Cryosectioning sample preparation tips and tricks
- Critical factors to get reproducible and high-quality cryosectioning results
- How automation improves cryosectioning techniques
It's nice to meet you!
Welcome to the Histology Hacks in Cryosectioning webinar.
I am Dr. Marla Rivera-Oliver, and I am very excited to share today some tips and tricks I've learned during my ten years of experience working on basic and translational research. I am based in the Dallas, Texas, area of the United States, and in my current role, I support customers from North Texas and Oklahoma. I completed a bachelor's in biology from the Interamerican University of Puerto Rico and a doctoral degree in Neurobiology from the University of Puerto Rico. I specialized in the spinal cord injury field and honed my skills in Cryosectioning, imaging, and in vivo electrophysiology. Then, I moved into a post-doctoral position at the Center for Discovery and Innovation of the City University of NY, where I grew my imaging, histology, and behavioral neuroscience expertise. Before joining Leica Biosystems, I joined the Leica Microsystems team as an account manager for microscopy. I supported the entire microscopy portfolio, from compound to confocal and electron microscopy. Since my bachelor's degree, the histology field has always been fascinating for me, where I had the opportunity to do my first tissue staining. Then during my doctorate studies, I had the chance to do a lot of histology and immunohistochemistry, which led me to publish a great article on spinal membrane receptors.
During our session, I will provide you with some great cryosectioning hacks I discovered while working in the spinal cord injury and locomotion laboratory in Puerto Rico and helped generate the quality data needed to publish this paper.
I worked mainly with mice and rat brain and spinal cord tissues to understand how the intrinsic mechanism of action of the spinal network affected locomotion, or the ability to walk, under certain circumstances like injury, alcohol exposure, and even caffeine.
To understand the physiology of the spinal network, I performed electrophysiology studies. To understand the anatomy and cell-to-cell interactions, I performed immunohistochemistry and other assays on previously sectioned tissue from the cryostat and the vibratome. The cryostat was key to achieving the thickness and quality of the tissue I needed and preserving its antigenicity.
Cryosectioning was the best solution as it was a fast and reliable way to get a tissue ready for staining, and the imaging quality was superior to any other technique I tried. During my learning process, I did a lot of troubleshooting with Cryosectioning, like working with uneven tissue and coiling sections.
The "hacks" we will discuss today have the potential for universal applications within any field that uses cryosection, as they are foundations of good practices. I hope you find inspiration and my talk provides value to your research project. So, let's jump right in!
During this session, we will be covering the following learning objectives:
- You will be able to identify three ways automation improves Cryosectioning techniques.
- You will be able to name three things essential to obtaining reproducible, high-quality sections with Cryosectioning.
- You will be able to name two “Histology Hacks” that can improve cryosectioning.
A bit of housekeeping:
There will be time for questions and answers at the closing of this presentation. Please feel free to add your questions to the chatbox, and we will answer as many as we have time for following this talk. Any questions we do not get to today will be forwarded to me by Labroots for later follow-up. In addition, please see my bio section for more information about me.
So with that said, let's start with the main question, What is cryosectioning?
What is cryosectioning (History/Timeline)
Cryosectioning, also called the "frozen section procedure," was first described by Dr. Louis B. Wilson in 1905, the chief of pathology at Mayo Clinic. Dr. Wilson was an expert at microscope-based research and a pioneer in visualizing diseased tissues. Shortly after arriving at the Mayo Clinic, Dr. Will Mayo said to him: "I wish you pathologists would find a way to tell us surgeons whether a growth is cancer or not while the patient is still on the table." Dr. Will was frustrated that it involved one operation in getting a tissue specimen, days to determine if it was malignant, anxiety for the patient who had to wait for answers, and a second operation if lab results showed the patient had cancer.
Dr. Wilson wanted to create a technique that offered rapid diagnoses of cancerous tumors or masses. Because there was no fast way to provide a cancer diagnosis at that time, he invented Cryosectioning. He was able to freeze, cut, stain, and give a diagnosis to surgeons within five minutes, often as quickly as two minutes.
Cryosectioning is widely used in modern science to perform microscopic analysis of numerous tissues with as many applications. The translational science field has embraced the technique to visualize fine details in cells. Let's talk about how that happens.
In 1920, the Spencer Automatic Freezing Microtome #880 arrived. This microtome is like that used by Dr. Wilson, although this particular model dates from 1920. But you can see how this cryosectioning technique gained popularity, and new inventions arose.
Currently, we have a wide variety of options available, from manual to automated Cryosectioning. Several labs have developed specialized techniques for different fields to enable a faster workflow and a high-quality throughput. But even though there are many techniques for Cryosectioning, they still all must confer the same goal: reproducibility. And with Cryosectioning, reproducibility could be a challenging task. There are at least three main factors that contribute to reproducibility. For me, those are Rate of Freezing, Temperature, and Cryopreservation.
3 Critical factors for reproducibility
The freezing rate, when the expansion of the water content by freezing stretches and penetrates cell membranes. Slow freezing promotes ice crystal formation and, thus, expansion. Note that the freezing rate, not the end temperature, is critical at first. Fast freezing depends on several factors: the percentage of the total surface in contact with the cold source, the volume of water, the starting temperature of the water in the specimen, and how cold the source is. Ideally, the cold source is at an extremely cold temperature over -80 degrees C and arranged to contact all surfaces.
Panel A shows the formation of ice crystals, a by-product of a slow rate of freezing.
Temperature: Temperature plays a big part in forming different types of ice that can affect tissue, and, ultimately, how it is sectioned.
For example, quick-frozen, or Vitreous or Amorphous ice is formed below -107 degrees C. This type of ice is semi-stable and forms Hexatagonal crystals and is the most common form of crystalline structure on Earth. At above -107 degrees C, Amorphous ice begins to restructure as cubic ice, which gradually extends by low temperature and pressure and will potentially change the morphology of your tissue. Understanding the types of ice formation at different temperatures will help you understand why it is important to section your specimens at below -171 degrees C.
Sectioning should be performed as soon as possible after biopsy and should begin as soon as the tissue is warm enough to cut. Tissue should never be left overnight or for long periods in a cryostat, even if there is no defrost cycle, as the conversion from vitreous ice to cubic will begin. Crystals will expand the ice and damage cells.
Let’s talk about workflow- 6 Steps
The six critical steps in Cryosectioning are: fixation/cryoprotection, specimen preparation, sample freezing/positioning, sectioning, staining and coverslipping, and finally, imaging.
I will now discuss each of the steps in more detail.
Fixation is optional but should be performed whenever is possible to prevent autolysis. The sample can be fixed before or after sectioning. The purpose of fixation is to preserve tissues permanently in as life-like a state as possible. Therefore, a variety of fixatives are available for use, depending on the type of tissue and features to be demonstrated. For example: aldehydes, mercurial, alcohols, oxidizing agents, and picrates. In my personal experience, I have always fixed my tissues with some form of aldehydes.
These examples illustrate the value of tissue fixation.
On the left panel, you can see an example of a Bronchiolo-alveolar Carcinoma - with 15 seconds drying time, using no fixation, and the arrows indicate where the cellular structure preservation is lost.
In contrast, on the right panel, the same type of tissue was immediately fixed with 95% ETOH. In this case, the arrows notate preserved tissue morphology, easily visualizing cellular membrane and nuclei.
Cryoprotection is optional but very well adopted into the research field. The sucrose solution protects the tissue and makes it less buoyant, so the tissue gains molecular weight and becomes easier to cut. A sucrose solution step for cryoprotection is very common in the research lab.
Step 2:Specimen Preparation
Immediately before cutting, samples should be grossed or trimmed with a razor blade. Avoid crushing artifacts by gently but firmly securing the specimen while cutting. This enables easy mounting onto the cutting device.
A smaller size is preferred as it can be frozen faster than a larger size. In preparing a frozen section, thickness is not as critical as in paraffin processing; it could be 1 cm or a little more.
If you'd like more information on grossing, we have great resources available on our website; please make sure to read "steps to better grossing" in our Knowledge Pathway for more details.
Step 3:Sample Freezing and Positioning
Pay attention to your fixation protocol.
Specimens for frozen sectioning can be prepared in a variety of ways. They can be frozen in a cryogen mixture such as isopentane cooled by dry ice, liquid nitrogen, or placed onto a thin layer of cryo compound on a specimen holder that is placed on the freezing shelf or a large cooled metal block placed inside the cryo chamber. The aim is to freeze the specimen as fast as possible to eliminate ice crystal formation or what is commonly called freeze artifact. My preferred method has always been to freeze with liquid nitrogen as it provides a fast and even freeze on the sample.
For the positioning of the tissue on the cryostat, the most critical edge of the tissue should be perpendicular or diagonal to the blade and not the first or last aspect of the tissue to meet the blade.
Frozen sections are less stable than paraffin or resin sections (which we won't cover in detail on this webinar) but provide superior preservation of antigenicity and, therefore, better detection of antigens by microscopy.
Move from fresh to freeze tissue in a smart way.
I prefer to section tissue at –20°c and thaw-mount sections on slides. Slides may be dried on a slide warmer at 37–40°c for a few minutes, and tissue sections should be stored at –70°c until use.
Step 5:Staining and Coverslipping
Staining and coverslipping can be performed manually or automated. This will depend on the needs of the laboratory and the volume of tissue.
The staining protocols are based on user preferences, and each lab has its own stain combinations. Some labs work strictly with hematoxylin and eosin stains, whereas others can use a wide variety of immunohistochemistry and in situ hybridization labels. Some labs will specialize in electron microscopy. There are certainly a variety of imaging needs, techniques, and combinations available. These protocols may be done manually, which will require much more time and effort, or become automated, which will help the staff but, more importantly, research, have the potential to decrease variability results.
Step 6:Cryosectioning and Imaging
Most, if not all, researchers who want to understand where a protein of interest is localized in a cell or tissue usually choose a fluorescent light microscopy (FML) approach. Compound fluorescence or confocal microscopes are good examples.
Using Cryosectioning for Immunohistochemisty (IHC )and Fluorescent In situ Hybridization (FISH) will allow for tissue with better antigenicity, make your cutting much faster, and will create beautiful thin sections (even monolayers) that will provide the level of resolution needed at the imaging stage.
On this image, panel A shows the results of a poorly preserved tissue which will lead to poor antigenicity. As a result, it can be challenging to identify or quantify data from that tissue section. Panel B shows a well-preserved tissue that conserves cell morphology and antigenicity.
We will talk in more detail about imaging later in the presentation. The common goal to strive toward is good quality in tissue preservation. One way to help the preservation of tissue for imaging is using automated Cryosectioning. Here, I propose three pillars for practicing automated Cryosectioning:
3 Pillars of Automation in Cryosectioning
Those 3 pillars are speed, accuracy and safety.
Pillar 1:Speed - In physics, speed means the time rate at which an object moves along a path. In Cryosectioning it means the time rate at which the specimen is moving along the path of the blade or knife, which will be translated into how many sections you will be able to cut in a certain amount of time. Automated Cryosectioning can allow for high-quality faster sectioning.
Pillar 2:Accuracy – As the cryostat has a pre-established speed rate, each section will be cut at the exact same rate producing the same accuracy per section.
Pillar 3:Safety – Safety is important for the user and the tissue. Safety protections may prevent any user accidents with the blade, and it is safe for the tissue as it will be cut with the highest precision, diminishing tissue loss.
But what key differences does an automated cryostat have compared to a manual instrument?
Automated Cryosectioning: 3 ways to improve
Three Of the Main Differences Are:
- Foot switch vs. wheel for sectioning (automates wheel turn velocity per rotation)
- Motorized handwheel vs manual- increase precision on blade movement. Motorized handwheel will also free both hands to manipulate sections.
- Preset counter – Don't worry about the number of sections or trimming levels already completed when following complex sectioning patterns. Section counter and section thickness totalizer does that automatically for you.
Cryostats 101: Understanding the cryosectioning instrument
Automation means using mechanization instead of a manual or hands-on process. In Cryosectioning, automation can consistently strive towards removing variability and improving consistency in cutting and ribboning of your specimen. For example: On a manual cryostat, the user sets the parameters and manually cuts, one by one, each section while collecting them on a slide. In comparison, the user selects the parameters and collects sections on automated cryostat, but the cryostat oversees cutting your specimen. Reproducibility and accuracy are key for accurate quantitative and qualitative results under the microscope.
Imaging is a world of possibilities; those possibilities will depend on tissue quality. One way to proceed with imaging is to trace the fluorescently tagged protein while it moved through the cell in real-time, which we call in vivo imaging (and which we won't discuss on this webinar). However, another strategy is to permeabilize chemically fixed cells or tissues and treat them with antibodies bearing a fluorescent compound (just to mention a few common fluorescence proteins been GFP, DAPI, Alexa Fluor mCherry, and others). These techniques include immunohistochemistry, fluorescent in-situ hybridization, and enzymatic degradation. And honestly, the first step for a successful fluorescence protein detection is having good tissue preservation.
This image illustrates well-preserved tissue under fluorescence light microscopy (panel A), having a higher level of detail under the confocal microscope (panel B) and the highest level of detail achieved under the electron microscope on panel C.
None of this tissue morphology and antigenicity would have been preserved if the sectioning protocol had not been perfectly created for the tissue needs.
So with that being said, how do you know if you have the correct protocol for your tissue? How can you understand what is happening when your results are not as expected? I also had those questions, and I will now share several histology hacks that helped me understand what was behind those undesirable results and how to overcome those challenges while cryosectioning.
Hack #1 - Pay attention to your fixation protocol.
Bad fixation will mean bad results.
Whenever you are using a fixation protocol, the fixation time for cells or tissue should last at least 6-8 hours, independent of how thick the sample is. This timeframe will allow for the covalent bonds to form and maintain the morphology of the tissue over time.
For animal tissue, it is preferable to apply the "whole-body perfusion" technique when available as it will ensure proper fixation of all tissues.
There are different approaches for fixation, the most common in research are 4% Formaldehyde (or PFA) for fluorescence light microcopy or a Glutaraldehyde solution for electron microscopy. Other techniques have been introduced to the field, like high-pressure freezing, which is a very interesting fixation protocol, but I have not tried myself.
And then, we cannot forget the sucrose solution that I like to include as part of my fixation discussion because after fixation, the specimen should be placed on a sucrose solution to cryoprotect your tissue during the freezing process and prevent the loss of morphology and antigenicity.
Panel A: well-fixed tissue showing good nuclear and cytoplasmic morphology with minimal shrinkage showing clearly defined basement membranes and cell margins.
Panel B: this is an example of poorly-fixed tissue showing inferior nuclear and cytoplasmic morphology with excessive shrinkage and poorly defined cell margins. Note the vacuolation and fragmentation of both nucleus and cytoplasm of cells of the distal tubule and retraction of the glomerulus due to shrinkage.
Let’s now move into the cryosectioning Hack #2:
Hack #2: Move from fresh to frozen tissue in a smart way.
Freeze a tissue sample up to 2 centimeters in diameter. Thicker samples won't freeze evenly and will coagulate with loss in antigenicity. Remember, preserving antigenicity and tissue morphology is our common goal.
Freezing the tissue should be a quick process using isopentane or liquid nitrogen (which is the most common) to prevent ice crystal formation, which will change morphological shape.
Histology Hack #3: Right temperature will save a lot of trouble when cryosectioning
I am talking about your instrument, your room, and even your floor temperature.
The temperature in the cryostat should be set for your type of sample, usually around 120 ˚C. If the temperature is too low, the block will be too rigid, and the sections won't be smooth and thin. If the temperature is too high, it won't achieve proper ribboning. Keep the cryostat door closed as much as possible to avoid temperature fluctuations.
Pay attention to the room temperature (which is usually around 68–72 ˚C), but low humidity is required. In humid regions like the Caribbean or West United States, we often find dehumidifiers in cryosectioning rooms. Room doors should be closed to avoid temperature fluctuations, and the cryostat should not be placed close to a window, air vent, or a place where sunlight can potentially change the instrument's temperature.
In summary, avoid as many temperature and humidity fluctuations as possible.
Cryosectioning Hack #4
Use The Right Tools to Achieve the Best Results
The use of charged slides will help with the section adhesion to slides.
Blades – use new blades each time you sit to cut. If the blade is dull, it won't cut thin and smooth sheets of tissue. Please remember to use a trimming blade and replace it with a new blade once closer to the specimen. You don't want to make your blade dull before starting to section your specimen. Ensure the specimen and blade are tightened securely and set to the desired temperature.
You will need to use brushes to help while collecting your sections and to keep your working area clean. Soft brushes should be used at all times, and one should try to avoid touching the section with the brush. Brushes can also help prevent the section's rolling while transferring to the slide.
Cryosectioning Hack #5
Proper Slide Storage
You may need to revisit your slides, store them the right way.
If fixation has been done correctly, your slides will remain intact for several months or even years. Temperature plays a huge role in proper tissue storage.
The finalized slides can be stored for months at 4 ˚C in a dark place such as a lab fridge, or stored for more extended periods in an ultra-low temperature freezer (preferably -20˚C) in a dark slide box.
Just remember to keep them stored in a low-humidity freezer, at a constant temperature and with no light.
Another tip is also an add-on to the cryostat called the CryoJane.
The CryoJane Tape-Transfer System is a method for Cryosectioning that makes it easy to produce high-quality frozen sections from difficult tissues. The CryoJane uses slides coated with an adhesive to help sections stick on the slide instead of the typically used brush. In addition, it can be used with most cryostats.
This system can provide sections as thin as two microns, with excellent quality characteristics such as being intact, wrinkle-free, and good adhesion to the microscope slide. It works by laying a piece of tape to the trimmed tissue block, and a section is cut and captured on the tape, which is then transferred to the adhesive-coated slide.
There are several videos on YouTube showing how the CryoJane has been used, type CryoJane on the search bar, and you will find the videos.
Now that we have discussed several hacks and tips about Cryosectioning, I believe it is time to compare how these hacks will impact your cryosection quality.
Here is a beautiful example of what can be achieved with the cryojane device by Pinskiy et al. The method is also suitable for larger brains, such as rats and monkeys. Workflow steps are explained on panel a.
On panel b from the Allen atlas, there are the most common sources of damage as of: folded areas (solid black arrows), as in a, b and c; torn sections (arrowheads, as in c), and sections with missing areas (dotted arrows, in d). Those common damages can be corrected by the use of this transfer tape technique.
The use of the CryoJane can help decrease tissue damage due to section collection.
In the same study from Dr. Pinskiy, we show here Thumbnail images of serial sections cut from a perfused mouse brain in three standard sectioning planes. Selected rows of serial sections cut in the coronal (top), sagittal (middle), and horizontal (bottom) planes. Sections were collected using the tape-transfer method, processed for Nissl stain, coverslipped, and imaged. In all cases, the sections appear of consistently high-quality.
Cryojane and tissue antigenicity
This is another example from the same study. Histochemical Panel A and B), Immunohistochemical staining (panels C, thru H), and the use of tape-transfer to section rat and macaque tissue is shown (Panel I-J).
Low-magnification views of coronal sections from mouse brains, reacted for (A) myelin Gallays silver stain technique, (C) tyrosine hydroxylase (TH), and (D) wisteria floribunda lectin (WFA), showing high antigenicity.
For each image, the arrow points to the region shown at higher magnification in (B), (E), and (F).
Tissue with native fluorescent expression is shown in (G) =for a mouse brain expressing Cre-dependent GFP in GAD2 cells.
Coronal sections of rat and monkey brains collected by the tape-transfer method are shown in (I) and (J). All sections are 20μm thick.
Today we have covered several topics on Cryosectioning in research. We talked about:
- What is Cryosectioning
- The 3 ways automation helps in cryosectioning
- The 5 steps of the cryosectioning workflow
- Hacks for better cryosectioning- and two bonus hacks!
- Additional Tips
This brings my presentation to an end. Thank you for attending, and I will be happy to answer any questions.
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