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High-Throughput Live Imaging of Microcolonies to Measure Heterogeneity in Growth and Gene Expression
Chapters
Summary April 18th, 2021
Yeast growth phenotypes are precisely measured through highly parallel time-lapse imaging of immobilized cells growing into microcolonies. Simultaneously, stress tolerance, protein expression, and protein localization can be monitored, generating integrated datasets to study how environmental and genetic differences, as well as gene-expression heterogeneity among isogenic cells, modulate growth.
Transcript
Our protocol for high throughput, yeast cells enables tracking of growth and gene expression for thousands of microcolonies simultaneously. Difference in between strains, between environments and even between genetically identical cells grown in a shared environment can be quantified. The protocol yields precise estimates of the average growth rate and average fluorescence intensity of gene expression reporters.
In addition, because so many individual microcolonies are assayed, precise estimates of entire distributions of growth and expression values are obtained. This protocol follows two main steps, the preparation of the experimental plate and the preparation of the cells for imaging. The cells must be plated immediately after dilution and mixing.
So we recommend setting up your experimental plate first. You'll notice the repeated mixing of cells throughout this protocol, which is extremely important in the steps up until plating. On the day of the experiment, all media and other solutions used for the assay must be filtered with a two micron filter, even if they're already sterile, in order to remove any crystals which may appear in microscope images.
This includes experimental media and the Concanavalin A solution. Next, prepare the microscope plate. In order to keep the glass surface of the wells clear of debris and scratches, always keep a lint and static free wipe underneath the plate.
Use sterile technique on a sterilized bench. Filter 25 milliliters of 1X Concanavalin A solution and pipette 200 microliters into each well of the microscope plate. We recommend stabilizing the pipette to ensure proper height.
Centrifuge the microscope plate with a lint and static free wipe underneath it for two minutes at 411 times G in order to remove any air bubbles which could prevent Concanavalin A from evenly coating the bottom of the wells. Let the plate containing the Concanavalin A solution rest for one to two hours. Be consistent across your experiments.
After the Concanavalin A incubation, clear the Concanavalin A from the 96 well plate either with a suction device or by forcefully dumping the solution out of the plate as shown here. Some droplets will remain. Next, wash the wells by adding 400 microliters of sterile water to each well.
Add the water immediately after removing the Concanavalin A solution. Do not let the plate sit dry. It is important to avoid touching the coated bottom of the plate with the pipette tips in order not to displace or remove Concanavalin A from the glass surface of the wells.
Remove the water in the same manner as the Concanavalin A solution and immediately add 185 microliters of experimental media to your microscope plate. Do not allow the plate to sit dry. The plate is now ready for the addition of 50 microliters of diluted cells.
Retrieve your pre-grown randomized 96-well plate. Your experimental question will determine if the cells are saturated or in log phase and the number of randomized replicates that should be used in the assay. Centrifuge your plate for two minutes at 411 times G to minimize the chance of cross-contamination when removing the foil covering.
Be sure never to tilt your plate when you are handling it. If yeast and media come into contact with the foil covering the plate, there can be contamination across wells. First prepare the plates you will conduct your serial dilutions into.
Add 90 microliters of experimental media to each of your dilution plates. Next prepare your cells. Be extremely careful when removing the foil from your 96-well plate to prevent droplets from one well from entering another.
The yeast will be concentrated at the bottom of the plate due to centrifugation. To ensure that cells are well mixed during your dilution, always add a small volume of cells to a larger volume of media. Then add another large volume of media to the mixture.
Be sure to pipette mix vigorously at every step. In this video, we performed dilutions for yeast cells at saturation and synthetic complete media, which we will dilute 10, 000 fold to achieve a final concentration of about 4, 000 cells per well in the microscope plate. To re-suspend the cells, pipette up and down vigorously and move the pipette tip around the well in a circular motion.
After mixing, there should be no traces of settled yeast remaining in any well as seen here in well A1.All other wells shown here have not been re-suspended for comparison. Count the cells in your pre-grown plate, for example, with the hemocytometer, and combine with any other strain that you will add to the same well in equal proportions. Here, we mix the cells from our pre-grown plate with a fluorescent reference strain at a one to one ratio.
Once your cell concentrations are standardized, add ten microliters of cells to the first serial dilution plate. Then add 100 microliters of media to a final volume of 200 microliters. With the addition of cells and media alike, pipette mix vigorously, moving the pipette tip around the wells in a circular motion.
From this point on, until the cells are added to the microscope plate, proceed quickly through the protocol. Next, add ten microliters of yeast from the first serial dilution plate into the second serial dilution plate. Then add 100 microliters of experimental media to that.
Mix well. If your yeast do not have a tendency to flocculate, then they are ready to be added to the microscope plate directly after this step. Proceed immediately to plating.
However, if the yeast strains you are using do have a moderate tendency to flocculate, the following optional sonication step will be necessary. If sonicating, first pipette mix the yeast solution once again, before the sonication. Place the diluted yeast onto the sonicator and submerge the 96-well pins into the solution of each well, but not touching the bottom of the well.
Set the sonication program, which is sufficiently strong to break apart flocculated yeast cells, but does not kill cells or cause elevated stress responses. Some testing may be required to identify the best sonication program for a given experiment. Once the program is complete, remove the cells and immediately proceed to the next step.
Add 15 microliters of yeast from the second serial dilution plate into the microscope plate. Pipette up and down to mix, but do so carefully to avoid disturbing the Concanavalin A attached to the bottom of the wells of the plate. Next, add 200 microliters of experimental media to a final volume of 400 microliters, to the microscope plate.
Pipette mix cautiously. Cover the plate with a breathable membrane. Make sure the edges of the plate are well-sealed.
Centrifuge the microscope plate with a lint and static free tissue underneath it for two minutes at 411 times G in order for the yeast cells to attach to the Concanavalin A at the bottom of the plate. The plate is now ready to be imaged. At the microscope, wipe the top and the bottom of the plate with a lint and static free wipe and blow compressed air onto the bottom of the plate to remove debris.
Place the plate onto the microscope. Make sure that the A1 well is in the top left-hand corner and that the microscope is heated to the correct temperature for cell growth. Also make sure that the plate is neatly tucked into the microscope and that the bottom of the plate is flat.
If the plate has a plastic cover, remove this before starting microscopy. This will be important to capture good images for downstream analysis. We load an automatically generated file with the X and Y coordinates and the focal positions to image the entire microscope plate.
If using a pre-generated file, spot check a few positions in the microscope plate to ensure cells are in the correct focal plane. It is helpful for the focal plane to be slightly above the cells so that they are surrounded by a dark rim and they are brighter than the background. Set the number of time points and the time point spacing that you will use for the experiment.
If you are collecting both growth rate and fluorescence intensity data, set up a fluorescence imaging program as well. Adjust the brightness in every channel to be imaged so that no position on the microscope plate is overexposed. Test the exposure you will use when you are not in live mode.
For fluorescent images, fluorescence should be visible to the naked eye. But otherwise we recommend setting low exposure values. Appropriately spaced colonies seeded by individual cells grow in a monolayer along the surface of the plate.
As a result, automated image analysis can be used to accurately calculate growth rate for many individual microcolonies. Fluorescent markers can be used to differentiate multiple strains growing in the same well. As a result, entire distributions of growth rates for individual microcolonies growing in a shared environment can be calculated.
The large sample sizes achievable with this experiment also allow for precise estimation of average differences between different strains or growth conditions. Here, growth rates for paired sets of colonies grown in single wells from a set of mutation accumulation lines and the in-well GFP reference control colonies are shown. If cells are observed clumping on the microscope plate, the clumps must be broken up by sonication prior to plating.
Cells in clumps that are not broken up will not grow in a planar microcolony along the plate surface, making correct calculation of growth rate impossible. Not all media is compatible with the microcolony growth rate assay. Some forms of media, including media that contains yeast extract or has a low pH, prevent binding of cells to Concanavalin A.This results in cells floating away from growing microcolonies over the course of the experiment and an erroneous growth rate measurements.
It is important to avoid plating cells too densely. When cells are plated too closely, faster growing microcolonies merge at earlier time points, shown here as a loss of color tracking. This effect can bias growth rate estimates as slow-growing colonies are less likely to merge with their neighbors before data from sufficient time points are collected.
Depending on growth conditions, a subset of cells may undergo a lag phase before beginning to grow. It is important to account for this when using colony areas to calculate growth rate rather than fitting a growth curve to all available time points. The described protocol can be used to investigate a wide range of questions related to yeast growth and evolution.
Its flexible microplate format means that it can be adapted to other yeast species, different growth conditions, other microbes, and potentially other cells in culture.
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