4D Microscopy of Yeast

Summary

This protocol describes the analysis of fluorescently labeled intracellular compartments in budding yeast using multi-color 4D (time-lapse 3D) confocal microscopy. The imaging parameters are chosen to capture adequate signals while limiting photodamage. Custom ImageJ plugins allow labeled structures to be tracked and quantitatively analyzed.

Abstract

The goal of this protocol is to characterize how membrane compartments form and transform in live cells of budding yeast. Many intracellular compartments in yeast are dynamic, and a full understanding of their properties requires time-lapse imaging. Multi-color 4D confocal fluorescence microscopy is a powerful method for tracking the behavior and composition of an intracellular compartment on a time scale of 5-15 min. Rigorous analysis of compartment dynamics requires the capture of thousands of optical sections. To achieve this aim, photobleaching and phototoxicity are minimized by scanning rapidly at very low laser power, and the pixel dimensions and Z-step intervals are set to the largest values that are compatible with sampling the image at full resolution. The resulting 4D data sets are noisy but can be smoothed by deconvolution. Even with high quality data, the analysis phase is challenging because intracellular structures are often numerous, heterogeneous, and mobile. To meet this need, custom ImageJ plugins were written to array 4D data on a computer screen, identify structures of interest, edit the data to isolate individual structures, quantify the fluorescence time courses, and make movies of the projected Z-stacks. 4D movies are particularly useful for distinguishing stable compartments from transient compartments that turn over by maturation. Such movies can also be used to characterize events such as compartment fusion, and to test the effects of specific mutations or other perturbations.

Introduction

Compartments of the endomembrane system are in constant flux, and their full characterization requires live cell imaging. Described here is a protocol that employs 4D (time-lapse 3D) confocal microscopy to visualize fluorescently labeled compartments in budding yeasts. The method was developed to track the dynamics of secretory compartments in Pichia pastoris and Saccharomyces cerevisiae^1,2,3. This protocol focuses on S. cerevisiae, which has a nonstacked Golgi in which the individual cisternae are optically resolvable⁴. The unusual Golgi organization in S. cerevisiae enabled the demonstration by 4D microscopy that a Golgi cisterna initially labels with resident early Golgi proteins, and then loses those proteins while acquiring resident late Golgi proteins^3,5. This transition can be visualized by creating a strain in which the early Golgi protein Vrg4 is labeled with GFP while the late Golgi protein Sec7 is labeled with a monomeric red fluorescent protein. When individual cisternae are tracked, maturation is observed as a green-to-red conversion³. This type of analysis can provide valuable information about protein localization and compartment identity. For example, two proteins with slightly offset arrival and departure times might sometimes appear to label different compartments in static images, but can be seen in 4D movies to label the same compartment at different time points^6,7. Thus, 4D microscopy reveals phenomena that would not otherwise be evident.

Informative 4D microscopy of yeast compartments can be achieved with appropriate procedures and equipment². Whenever possible, fluorescent protein tagging is performed by gene replacement⁸ to avoid overexpression artifacts. Because intracellular structures are often very dynamic, 4D imaging is needed to ensure that a structure is tracked reliably over time. The protocol described here employs a laser scanning confocal microscope equipped with high sensitivity detectors. With this device, the entire cell volume of S. cerevisiae can be imaged by confocal microscopy approximately every 1-3 s, with 2 s intervals being typical. Data can be collected for up to 5-15 min depending on the labeling densities of the fluorophores and their photophysical properties. The main hurdle is to minimize photobleaching. For this purpose, the laser intensities are kept as low as possible, the confocal scan speed is maximized, and the optical parameters are configured to image at the Nyquist limit in order to capture the relevant information while avoiding excessive light exposure. These settings are also expected to alleviate phototoxicity, a factor that is often overlooked during live cell imaging^9,10,11. The resulting noisy data are processed with bleach correction and deconvolution algorithms to facilitate quantification of fluorescence intensities.

Even with high quality 4D movies, the analysis is tricky because yeast compartments tend to be numerous, heterogeneous, and mobile. Due to the intrinsic limitations of confocal microscopy and the non-optimal settings required for prolonged 4D imaging, fluorescent structures that are near each other are hard to resolve. This problem can be circumvented by focusing on the small number of fluorescent structures that remain optically resolvable for the duration of the labeling period, with the assumption that those structures are representative of the whole population of labeled compartments. Fluorescent compartments that can be reliably tracked are identified by viewing movies of projected Z-stacks and by creating a series of montages in which the optical sections for each time point are arrayed on a computer screen. This analysis employs custom ImageJ¹² plugins, which allow an individual structure to be tracked in isolation.

Recent methods papers covered the use of fluorescent proteins in yeast¹³ as well as the theory and practice of 4D confocal imaging of yeast cells². This protocol focuses on the key practical aspects of a 4D imaging experiment. It includes some enhancements to previously described procedures, as well as updated versions of the ImageJ plugin code and documentation. The example shown focuses on Golgi dynamics, but this protocol is equally suitable for imaging other yeast compartments.

Protocol

1. Preparation

1. Make nonfluorescent minimal NSD medium². The absence of riboflavin is expected to reduce background intracellular green fluorescence and associated phototoxicity.
2. Grow the yeast strain overnight at ~23 °C to logarithmic phase in 5 mL NSD in a 50 mL baffled flask with good aeration. About 3-4 h prior to analysis, dilute the yeast culture in fresh NSD so that the final OD₆₀₀ will be 0.5-0.8 at the time of imaging.
3. Prepare a 2 mg/mL solution of concanavalin A (ConA). If desired, freeze aliquots of this solution in liquid nitrogen and store at -80 °C.
4. Spin the ConA solution for 5 min at full speed in a microcentrifuge to remove particulate matter that may interfere with imaging. Then add 250 µL of the supernatant to a clean 35 mm coverglass bottom microscopy dish. After 15 min, wash 2-3 times with 2 mL of dH₂O and let dry.
5. Add 250 µL of the yeast culture to the ConA-coated dish, wait 10 min to allow the cells to adhere, and gently wash 2-3 times with 2 mL NSD. Cover the cells with 2 mL fresh NSD.

2. Imaging

1. Use a 63x or 100x oil immersion lens. A numerical aperture (NA) of 1.40 is sufficient, but a higher NA lens can also be used.
2. Format the frame size to 256 x 128 (width x height). If a larger frame size is needed, increasing the width will not reduce the scan speed as long as the confocal microscope is equipped with a resonant scanner.
3. Adjust the zoom factor to make the pixel size ~80 nm.
4. Use the maximum scan speed, which is typically on the order of 8 kHz. Turn on bidirectional X scanning if it is available, and if control experiments confirm that the scans from the two directions are in register.
5. Set the line accumulation to 4 or 6. Be sure to use accumulation (summing) instead of averaging.
6. Set the pinhole to 1.2 Airy units. Empirically, when imaging live yeast cells, this setting captures more photons than the standard choice of 1.0 Airy unit while causing no appreciable loss of resolution.
7. For each fluorescence channel, set the excitation wavelength, assign a high sensitivity detector, set the emission wavelength range, and turn on photon counting mode if available. The wavelength choices will depend on the fluorophores. As an example, excite GFP fluorescence at 485 nm and collect from 495-550 nm, and excite mScarlet fluorescence at 555 nm and collect from 562-625 nm.
8. Set the intensity of each laser to be as low as possible. This setting must be determined empirically. An appropriate intensity will result in capture of a noisy but interpretable image sequence, with the fluorescence signal bleaching no more than 50% by the end of a 5 min movie. On the confocal microscope used here (see Table of Materials), this intensity corresponds to a laser power setting on the order of 5%.
9. Use notch filters or time gating to avoid capturing reflected light from the coverslip. If time gating is available, set the gating window to 0.6-10.0 ns.
10. Turn on brightfield imaging and use a low sensitivity detector for data collection. Set the gain to a level that makes the cells clearly visible. Do not use differential interference contrast (DIC) because the prism will interfere with the capture of reliable fluorescence data.
11. Set the Z-step interval to 0.25-0.35 µm. Image the entire volume of the yeast cells by collecting about 20-25 optical sections per Z-stack. Specify the directionality of imaging such that "down" moves toward the coverslip.
12. For a typical movie, set the time interval between Z-stacks to 2 s and set the movie duration to 5 to 10 min. Depending on the compartment under study, reduce the interval to make shorter movies of relatively fast dynamics, or increase the interval to make longer movies of relatively slow dynamics.
13. Save the movie as a LIF file with the brightfield images in the last channel. Higher bit depths are unnecessary at this stage, and the processing pipeline is configured to accept 8-bit TIFF data.

3. Deconvolution

1. Launch the deconvolution software (see Table of Materials), using the Classic Maximum Likelihood Estimation algorithm.
2. Open the data set generated in step 2.13. Select "Deconvolution wizard". Inspect the values displayed in the "Parameter wizard". The imaging parameters should be detected and correctly displayed. Under "Refractive indexes", change the "Embedding med." value to 1.40 to approximate the yeast cytoplasm. Under “coverslip”, select “Launch editor” and “Estimate position” and “OK” to set the coverslip position. Select "Set all verified" and "Accept".
3. Proceed through the "Deconvolution wizard" for each fluorescence channel. Select "Manual" as the mode for background estimation. Inspect the raw data fluorescence intensity profile to determine an estimated background value, which is typically about 0.1.
4. In the deconvolution setup menu, enter a "Maximum iterations" value of 40 and turn off bleach correction. Enter an estimated SNR value, which is typically about 5.0. Click "Deconvolve".
5. If necessary, return to the original data file, adjust the background and SNR values, and repeat the deconvolution, until noise is sufficiently removed without eliminating genuine fluorescence from dim structures.
6. Merge the brightfield and deconvolved fluorescence channels. For subsequent movie editing steps in ImageJ, arrange the channels such that red is first, green (if present) is next, blue (if present) is next, and brightfield is last. Save the image sequence as an 8-bit TIFF file.

4. Bleach Correction and Movie Generation

1. Import the deconvolved image sequence into ImageJ and convert it into a hyperstack by choosing Image > Hyperstacks > Stack to Hyperstack. Select "xyzct" from the drop-down menu, and fill in the number of channels, Z-stack slices, and time frames (http://wiki.cmci.info/downloads/bleach_corrector).
2. Choose Image > Color > Split Channels.
3. Correct the fluorescence channels for photobleaching using the ImageJ plugin available from http://wiki.cmci.info/downloads/bleach_corrector. Once that plugin has been downloaded and installed, choose Plugins > EMBLtools > Bleach Correction. Select "Exponential Fit."
4. To merge the brightfield and bleach-corrected fluorescence channels into a hyperstack, choose Image > Color > Merge Channels. Save the resulting hyperstack as an 8-bit TIFF file.
5. Install the custom ImageJ plugins provided with this protocol. Follow the instructions in the accompanying document to view, edit, and quantify the image sequence, and to produce a final movie of the projected Z-stacks.

Results

The example given here documents and quantifies the maturation of two yeast Golgi cisternae as visualized by dual-color 4D confocal microscopy³. A yeast cell contains on the order of 10-15 Golgi cisternae, each of which matures over a time course of approximately 2-4 min. Maturation can be visualized by tagging the early Golgi marker Vrg4 with GFP and by tagging the late Golgi marker Sec7 with a red fluorescent protein such as mCherry or mScarlet. An individual cis...

Discussion

4D confocal imaging of yeast organelles requires careful tuning of multiple parameters. The major concern is photobleaching and phototoxicity. A typical 4D movie involves collecting thousands of optical sections, so the laser illumination must be kept as low as possible. Tandem fluorescent protein tags can be used to boost the signal without increasing expression of the tagged protein^16,17. Maximizing the scan speed helps to limit photodamage, and also allows Z-s...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
35 mm glass bottom dishes No. 1.5	MatTek	P35G-0.170-14-C	Imaging dishes
Concanavalin A powder	Sigma-Aldrich	C2010
Trolox	Vector Laboratories	CB-1000
Leica SP8 confocal microscope	Leica Microsystems
Leica Application Suite X	Leica Microsystems		Microscope software
Huygens Essential software, version 17.04	Scientific Volume Imaging		Deconvolution software
ImageJ	NIH		Image processing and analysis software