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7 Things You Didn’t Know About Imaging Cytometry

Written By: Tim Bushnell, PhD

We are visual creatures. We seek to capture and describe the world around us. Some of the earliest evidence of this is found in the cave paintings that have been found around the world, like this painting of a horse found in the caves in Lascaux, France.

Figure 1: Cave painting of a horse. From Wiki.

With the development of reliable microscopes, such as those developed by the dutch draper Antonie van Leeuwenhoek. For the first time, we were able to see what we could not see before, probing the unseen and learning in great detail how organisms worked.

Figure 2: Antonie van Leeuwenhoek and his microscope. The figures represent his drawings of red blood cells

Over time, the field of cytometry, that is the analysis of biological processes at the whole-cell level, has exploded and expanded in so many different directions. The flow cytometry can be thought of as a microscope with very poor resolution. The power of flow cytometry lies in its ability to analyze thousands of cells through many dimensions, providing an amazingly detailed understanding of the cell. However, do the resolution, it is not possible to tell where these signals are located.

In 1979, Dr. Leon Wheeless from the University of Rochester published this paper which described an imaging flow cytometer.

Figure 3: An early imaging flow cytometer.

This system combined the speed of a traditional flow cytometer with the ability to roughly image a cell, as shown below. The cells were focused enabling the group to capture two different images at two different angles. In this case, they were able to use this system for some very basic whole-cell imaging. They identified the barrier of the cell, the nucleus of the cell, and were able to make measurements for the DNA an RNA content, as shown below.

Figure 4: Data from the imaging flow cytometer.

Fast forward about 20 years and the ensuing development of lasers, detectors and flow cytometers. IN 2004, this paper was published, which described the multispectral imaging cytometry, the ImageStream 100, developed by a company called Amnis (now owned by Luminex). This system was able to take up to 6 multispectral images of cells in flow. The instrument layout is shown below.

Figure 5: The ImageStream 100 optical layout.

With this system, the cells were hydrodynamically focused and passed an interrogation point. The light was collected and separated using a fanned array of filters, which reflected the light onto different regions of a custom build CCD camera chip. To keep the cells in focus, the system uses a bead, aptly called the speed bead, and using two PMTs makes adjustments to the system to keep the cells in focus. Additionally, these beads are used to calculate the velocity of the cells. This is important because the system used a technique called time-delay integration (TDI) to capture the images. TDI was developed for observing moving objects at low light levels.

This system allowed for a host of new applications, from using morphology to characterize cells, to studying cell-cell interactions, phagocytosis, co-localization, and nuclear translocation to name just a few. Several years later, Amnis released the ImageStreamX. The new system had two cameras and more lasers, allowing for up to 12 multispectral images to be captured at the rate of 300 cells per second.

The imaging flow cytometer is an enabling technology, allowing for new experiments that previously couldn’t be performed by traditional flow. Since the cells are visualized, it allows for better counting of events of interest. Recently, it has also been shown to be useful for measuring EVs as illustrated by this article.

When preparing for an imaging flow experiment, there are several important considerations that need to be taken into account to ensure the highest quality data is generated.

1. Lasers are collinear.

One of the first important considerations is that the excitation lasers are co-linear. This has a dramatic impact on the fluorochrome choice. Below is a chart from an instrument showing some of the fluorochromes that can be used on the ImageStream and potential issues that may arise.

Figure 6: ISX laser and filter configuration, highlighting potential issues.

If one were to use PerCP-Cy5.5 on the ImageStream, it would be detected in Channel 5 as well as Channel 11. PerCP has a wide excitation profile, as shown below.

Figure 7: Excitation profile of Per-CY5.5 plotted using this tool.

So fluorochrome choice is a critical parameter in panel design. Avoiding tandem dyes and those dyes that have wide excitation profiles is recommended.

2. The detector is not a PMT.

Unlike a PMT, the CCD chip in the ImageStream system does not have an adjustable voltage to attenuate the sensitivity. Rather, fluorescence signal intensity is governed by the laser power. This requires careful stain balance to ensure successful results. A very bright fluorochrome should not be on the same laser as one substantially less bright. Likewise, care must be taken when using fluorescent proteins. A very bright GFP excited with high laser power can negatively impact multiple channels cuasing significant issues when trying to resolve other fluorochromes.

If the fluorochrome is too bright, it will saturate the camera, and show up as red on your images.

Figure 8: Consequences of saturation.

On the left are the uncompensated images, showing the saturation of the green signal. Anywhere there is red on the image, the pixels are saturated. This can be shown in the plot of fluorescence area versus the peak fluorescent hight on the right.

The three rules of compensation still apply,and saturated events need to be removed from the analysis, so during assay development and optimization, pay close attention to the laser power and stain balance.

3. Need a specific cell concentration.

Instead of using standard culture tubes, samples are loaded into the ISX in microcentrifuge tubes. As a rule of thumb, one to two million cells in 60 microliters is a good volume to use. While on the topic of tubes, the use of siliconized treated tubes is a good idea for intracellular staining. This helps to improve the pelleting of the fixed cells.

4. Use high-grade formaldehyde.

There are a variety of protocols and reagents available for fixation and permeabilization. Each has their pro’s and con’s, and every lab has its favorite reagent. To get high-quality fixation, it is recommended to use EM grade formaldehyde. The methanol-free formaldehyde packed in ampoules is a good source of the fixative. Dilute the fixative to the working concentration, typically 1 to 4% (although for most applications, 1-2% is sufficient), and fix away.

It should be remembered that formaldehyde can crosslink to itself in the absence of stabilizers,. As formaldehyde crosslinks, it reduced the effectiveness of the fixative/. Make sure to expire the solution within about 2 weeks. If the whole ampoule is not used at once, if it can be stored under nitrogen, or other inert gas, it will extend its live.

5. You will need to perform compensation.

It’s not possible to escape the need for compensation using the ImageStream. Uncompensated cells captured using the ImageStream are good to visualize spectral spillover as shown below.

Figure 9: Uncompensated images captured on the IS100. Data courtesy of David Basiji.

Figure 9 shows the results of spectral spillover into the other channels on the system. Take the CY3 control, for example. This is measured in the orange channel, but due to the width of the spectrum, it is measured in the green and red channels as well.

Figure 10: Cy3 emission profile with the green, orange and red filters shown.

Figure 11: Cells after compensation

After compensation, it is easy to see the signal of the individual fluorochromes is found in the appropriate channel. One thing to remember when compensating on the ImageStream is to turn off the brightfield image.

When compensating, after collecting the individual tubes, the files can be easily concatenated into one master file and using time as a parameter, each individual compensation control can be identified using a time plot. It helps save some time in the process.

6. Masking technique.

One of the powers of using imaging flow cytometry is that the cells are visualized so that we can identify where the signal is in relation to other signals, the cell surface, the nucleus, etc. To further refine the analysis, the software calculates a variety of ‘Masks’ or regions of interest, that can be used to define where in or on the cell the data can be extracted from.

Figure 12: Different masks as visualized on a cell.

In the software, a mask is displayed as a bluish overlay on a cell. The default mask is shown in the upper left. The other figures show how this mask can be manipulated. For example, the system can dilate or erode the system mask, if a nuclear dye is used, a nuclear specific mask can be generated (great for translocation studies), while threshold masks allow for showing only regions above a given threshold. All in all, these masks can help define where the signal of interest is.

7. Make complex mass.

What makes masking very valuable is the fact that masks can be combined using boolean logic to create a whole new mask. For example, say that you were interested in only looking at the signal that was located outside the nucleus. If the cells were labeled with a nuclear dye (like Draq-5) and a membrane dye (like DIL), it becomes a matter of defining the gates and putting them together to create a new gate.

Figure 13: Complex masking

Shown in the top figure is the system mask on the three images. If a nuclear mask is generated (Draq-5), and subtracted from the system mask in the DIL channel, this will result in a ringed region displayed on the BF image. Now it becomes possible to ask specific questions about the fluorescence in this region only.

It has been said that “a picture is worth a thousand words.” If that’s true, then how many words does the ImageStream write about visualized cells? Traditional applications like phagocytosis have been improved by morphological and fluorescent data, as well as the ability to define specific regions of interest. But these factors have also opened the door to new methods like nuclear translocation and co-localization. ImageStream exists at the intersection of traditional fluorescent flow and fluorescent microscopy, drawing from the best of both.

To learn more about 7 Things You Didn’t Know About Imaging Cytometry, and to get access to all of our advanced materials including 20 training videos, presentations, workbooks, and private group membership, get on the Flow Cytometry Mastery Class wait list.

Tim Bushnell, PhD


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