Creating and using settings on the Aurora
- olivertburton
- Mar 17
- 9 min read
Today's post is very simple--we're just looking at how to create reusable settings on the Cytek Aurora. If you've used this machine, you've probably done this already. I'm putting up the workflow we use, though, because creating settings is an important step in getting reproducible data with longitudinal studies. As far as I can tell, this is necessary (or at least very helpful) to get stable unmixing using library controls, a topic for an upcoming post. It also saves you time.
While this is geared towards the Aurora, creating templates with your desired voltage settings on the ID7000 will probably accomplish something similar.
Right, so here's the workflow. We're going to look at how to do this using mouse splenocytes and compensation beads.
If you haven't already done so, run the instrument QC and be sure it passes!
Now, always start creating new settings from the CytekAssaySettings (CAS). These are updated based on the QC. The fluorescence detector voltages are adjusted by the QC to hit specific targets, and we are not going to modify those. Changing the fluorescence detector voltages is unlikely to be necessary for >99% of immunology studies, so unless you're working with a reporter that's going off scale, I would not recommend opening this can of worms.
What we are going to change is the scatter voltages. By default, the CAS has FSC very high, presumably to get the QC beads in the desired position. This value is way off for cells.
Before we run anything, though, we're going to "air gate" our samples. This means we're going to draw gates as targets for where we want our samples to appear on FSC and SSC. We will then adjust the voltages to get the cells to fall into these positions. Why this way? Why not move the gate to where the cells are? If we move the gate, the cells are going to be in a different position every time we create settings, so potentially a different position with each experiment. That reduces our reproducibility between experiments. We can pretty quickly end up in a situation where the granulocytes up off the scatter plot (FSC/SSC too high) or the smallest lymphocytes are farther to the left (FSC/SSC too low). Comparing experiments with different scatter distributions will give different cell compositions during analysis if not carefully adjusted for.
How this might look:
On the left, I've got a gate for "Cells", which is actually a mouse lymphocyte gate that should be renamed. On the right, we have a plot of FSC vs. SSC-B (violet laser SSC). Because we have two side scatters, we can use them to display the cells differently. This is really useful if you want to have a clear view of the lymphocyte population boundaries (making the lymphocytes take up a lot of the screen) while also getting the granulocytes on screen. That's quite useful for whole blood or tissue work. To do this, we'll scale one SSC to get the lymphocytes big while setting the other to compress them down.
So, at this stage, draw the gates for the populations you want to study and place them where you want the cells to appear (roughly). If you're working with compensation beads, you can add a specific gate for beads if you want. We're going to add scatter thresholds to remove debris and noise, so I've draw those in as boxes. This template for creating settings can be saved as a worksheet.
In the video below, you can watch the workflow for getting from the CAS to settings that bring the cells into the gates. We start by running cells, then adjusting down the FSC to get the cells on screen. SSC usually needs to be increased for lymphocytes. Once the cells are centered, increase the FSC threshold. I like to set the threshold about halfway between 0 and the lower edge of the cells. You want it to be high enough to exclude most of the debris because these events just add to your file size and can add a lot if you're working with tissues or have any erythrocytes present. Setting a second threshold on SSC is optional and not as important on the Aurora as on other machines like the Symphony. This can help exclude additional noise (and there should be no meaningful information down there anyway). If you set two thresholds, use the Boolean "And" rather than "Or".
Adjust your SSC-B to get the other desired cell types on screen, if you want.
While this is geared towards the Aurora, creating templates with your desired voltage settings on the ID7000 will probably accomplish something similar.
Right, so here's the workflow. We're going to look at how to do this using mouse splenocytes and compensation beads.
If you haven't already done so, run the instrument QC and be sure it passes!
Now, always start creating new settings from the CytekAssaySettings (CAS). These are updated based on the QC. The fluorescence detector voltages are adjusted by the QC to hit specific targets, and we are not going to modify those. Changing the fluorescence detector voltages is unlikely to be necessary for >99% of immunology studies, so unless you're working with a reporter that's going off scale, I would not recommend opening this can of worms.
What we are going to change is the scatter voltages. By default, the CAS has FSC very high, presumably to get the QC beads in the desired position. This value is way off for cells.
Before we run anything, though, we're going to "air gate" our samples. This means we're going to draw gates as targets for where we want our samples to appear on FSC and SSC. We will then adjust the voltages to get the cells to fall into these positions. Why this way? Why not move the gate to where the cells are? If we move the gate, the cells are going to be in a different position every time we create settings, so potentially a different position with each experiment. That reduces our reproducibility between experiments. We can pretty quickly end up in a situation where the granulocytes up off the scatter plot (FSC/SSC too high) or the smallest lymphocytes are farther to the left (FSC/SSC too low). Comparing experiments with different scatter distributions will give different cell compositions during analysis if not carefully adjusted for.
How this might look:
On the left, I've got a gate for "Cells", which is actually a mouse lymphocyte gate that should be renamed. On the right, we have a plot of FSC vs. SSC-B (violet laser SSC). Because we have two side scatters, we can use them to display the cells differently. This is really useful if you want to have a clear view of the lymphocyte population boundaries (making the lymphocytes take up a lot of the screen) while also getting the granulocytes on screen. That's quite useful for whole blood or tissue work. To do this, we'll scale one SSC to get the lymphocytes big while setting the other to compress them down.
So, at this stage, draw the gates for the populations you want to study and place them where you want the cells to appear (roughly). If you're working with compensation beads, you can add a specific gate for beads if you want. We're going to add scatter thresholds to remove debris and noise, so I've draw those in as boxes. This template for creating settings can be saved as a worksheet.
In the video below, you can watch the workflow for getting from the CAS to settings that bring the cells into the gates. We start by running cells, then adjusting down the FSC to get the cells on screen. SSC usually needs to be increased for lymphocytes. Once the cells are centered, increase the FSC threshold. I like to set the threshold about halfway between 0 and the lower edge of the cells. You want it to be high enough to exclude most of the debris because these events just add to your file size and can add a lot if you're working with tissues or have any erythrocytes present. Setting a second threshold on SSC is optional and not as important on the Aurora as on other machines like the Symphony. This can help exclude additional noise (and there should be no meaningful information down there anyway). If you set two thresholds, use the Boolean "And" rather than "Or".
Adjust your SSC-B to get the other desired cell types on screen, if you want.
Once you have your cells in the desired positions, save the settings. Save them with a meaningful name. Scatter changes with fixatives and cell types, so I tend to save mine with a short description of what the cells and fixatives are. These settings should be valid any time I go back to using those cells with those conditions. As a bonus, using standardized settings means you'll notice if you've accidentally done something different to your cells.
Importantly, these settings are updated with the QC like the CAS. This means that if the FSC and SSC detector sensitivity on your machine changes, the voltage values you see in your saved settings will change. This is a good thing. This is what you want. This enables your cells to fall in the same position (your target gate) every time. You can validate this yourself using beads. In practice, you may see slight variations in the positions of your cells relative to your target gates, which, as far as I can tell, are all down to differential handling of the samples prior to reaching the cytometer. These variations are typically very, very minor and will be less than you'd get by trying to set new voltage each time, in my opinion.
If you plan to use compensation beads as well as cells, you'll probably want slightly different settings for the beads. If you've just created settings for your cells, you can modify these for the beads rather than starting over. All you need to change is the FSC and SSC (increasing FSC and decreasing SSC, typically). You can either set these such that the beads fall in your target "bead" gate, or you can adjust the scatter voltages such that the beads fall in the lymphocyte/cell gate, as in the example below. If you want, you can reduce the scatter thresholds since the beads don't contain much debris.
This second approach helps a little if you are using both cells and beads for unmixing. When you go to unmix the data, you can set a single gate that can be applied to both cells and beads because they're in the same position.
Once done, save these modified settings, e.g., "Compensation beads".
Now, when you go to run a new experiment using the same cells treated the same way, you can just recall your saved settings. You don't need to go through this process again.
If you're running both cells and beads in your Reference Group (single color controls), you can set specific settings for each sample. If you tell the Aurora to switch settings for your bead samples, it can automatically run all your cells on your "Mouse lymphocytes fix/perm" settings and all your beads on "Compensation beads" settings. That means you can walk away and do something else.
Note that you can also set different stopping criteria for individual samples. This allows you to set a "bead" stopping gate for bead samples and a "lymphocyte" gate for cells, for instance. Additionally, you can stop after a couple thousand events for beads, or set specific cell controls with rare markers (e.g., ST2) to stop with higher event numbers (e.g., 100,000). This allows you to get enough events where needed and speed up the acquisition where you can.
How often should you update your settings by re-creating them from CAS? I don't know because they're good for long enough that this is hard to track. To be cautious, I'd suggest you might want to re-do them every 6 months to a year for settings used for independent studies. If you're doing longitudinal human studies, you might be better off not changing the settings for the length of the study. If your instrument undergoes a major change (e.g., new laser, replaced DAC board, other optics hardware replacement), check and probably change your settings.
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