Imaging the actin cytoskeleton in live budding yeast cells

Summary

Although budding yeast, Saccharomyces cerevisiae, is widely used as a model to organism in biological research, studying cell biology in yeast was hindered due to its small size, rounded morphology, and cell wall. However, with improved techniques, researchers can acquire high-resolution images and carry out rapid multidimensional analysis of a yeast cell. As a result, imaging in yeast has emerged as an important tool to study the cytoskeletal organization, function, and dynamics. This chapter describes techniques and approaches for visualizing the actin cytoskeleton in live yeast cells.

1. Introduction

Thirty-six years ago, the actin cytoskeleton was first visualized in fixed budding yeast cells with fluorescent phalloidin [1]. Subsequently, fluorescence imaging became a useful tool to study actin-containing structures in yeast. Many elements of actin cytoskeletal structure and function are conserved within the eukaryotic lineage. Nevertheless, unlike mammalian cells, budding yeast contains a single actin gene, making it a convenient model organism for studying actin organization and function. The actin cytoskeleton is responsible for functions in yeast as in other eukaryotic cells including cell polarity, cell division, endocytosis and organelle transport.

The actin cytoskeleton of yeast consists of two major structures that are maintained throughout the cell cycle: actin patches and cables. Actin patches are endocytic vesicles that are coated with actin filaments that are nucleated by the Arp2/3 complex. Other patch constituents include conserved proteins such as: clathrin and its adaptors, endocytic adaptors, nucleation promoting factors (NPFs), kinases, and actin bundling proteins (e.g. Sac6p/fimbrin) [2]. Early in the G1 phase of the cell cycle, actin patches become concentrated at the nascent bud site, reflecting cell polarization towards the small growing daughter cell during bud emergence. Patches remain concentrated in the bud during isotropic growth [3, 4]. Finally, when the bud cell reaches a similar diameter to the mother cell, actin patches re-distribute to the bud neck and facilitate cytokinesis [3, 4]. Thus, the distinctive polarity of actin patches can be used as an indicator for the function of the cell polarization machinery.

Actin cables are bundles of actin filaments that extend through the entire length of the yeast cell and reach the mother distal tip. They are essential for yeast cell division because they act as tracks for transport of cellular constituents including mitochondria, Golgi, vacuoles, secretory vesicles, spindle alignment elements, and mRNA from mother to developing daughter cells during cell division [5]. They are also critical for transport of vesicles and other cargos that contribute to cytokinesis.

Actin cables are highly dynamic. They undergo retrograde flow, movement from the bud toward the mother cell tip. In yeast, as in mammalian cells, retrograde actin cable flow (RACF) is driven by the pushing force of actin cable assembly and pulling force of a type II myosin protein, Myo1p [6, 7]. Actin cable assembly occurs in the bud tip and bud neck. During this process, actin filaments are polymerized by formin proteins (Bni1p and Bnr1p) and assembled into bundles by Sac6p (fimbrin), Abp140p, and Bnr1p. Actin cables are further stabilized by two tropomyosin isoforms (Tpm1p and Tpm2p).

Although RACF occurs in the retrograde direction, from daughter to mother cell, actin cable-dependent cargo transport is primarily in the anterograde direction, from mother to daughter cell. Thus, mitochondria and other cargos are effectively “swimming upstream”, against the opposing force of RACF as they move into the daughter cells along actin cables. As a result, actin cable dynamics play a role in promoting inheritance of fitter mitochondria by yeast daughter cells, which in turn, affects daughter cell fitness and lifespan [8]. Therefore, studying actin cable dynamics is important for understanding yeast cell biology.

Imaging techniques and technical methods have evolved over time to allow for efficient live-cell imaging of actin dynamics in yeast cells. Here, we describe the steps to visualize and characterize actin movement: tagging of yeast genes at their chromosomal loci with fluorescent proteins; determining functionality of tagged proteins; optimizing imaging methods; and quantitative analysis of actin dynamics.

2. Detection of cellular structures using targeted fluorescent protein

Live imaging of the yeast actin cytoskeleton can be carried out by tagging endogenous proteins with fluorescent proteins or expression of tagged heterologous actin-binding peptides. The most commonly used fluorescent protein is GFP and its variants; however, other fluorescent proteins can be used. Here, we describe how to select a fluorescent protein for imaging studies and tag a protein at its chromosomal locus.

2.1. Choosing a fluorescent protein

The green fluorescent protein (GFP) revolutionized live-cell imaging in cell biology. GFP was first cloned from the jellyfish Aequorea victoria, and has been used as a foundation for developing variants with a range of colors and molecular properties. Moreover, combining laboratory mutagenesis with novel fluorescent proteins, such as mCherry from Discosoma sp. coral and Teal from Clavularia sp. coral, has produced fluorescent proteins with new colors, improved brightness, faster folding, and decreased oligomerization [9].

Fluorescent proteins that are used for live-cell imaging must be photostable and able to withstand repeated excitation. In addition, the emission of the selected fluorescent protein must be bright enough to highlight structures of interest above background fluorescence and detector noise, while being spectrally distinct from other labels being used. Fluorescent protein brightness is determined by the protein’s intrinsic brightness (the product of extinction coefficient and quantum yield), and the behavior of the fluorescent protein upon ectopic expression (rate of folding and stability). The amount of signal that is excited and detected depends on the imaging system’s light sources, lenses, filters and detectors, and their compatibility with the fluorescent protein excitation and emission spectra. For more information on fluorescent protein physical and spectral properties, https://www.fpbase.org/ is an invaluable resource. Lastly, the fluorescent protein must not be toxic or disrupt the cellular behavior or protein function to which it is fused.

2.1.1. Tagging endogenous proteins

Uniquely in yeast, homologous recombination methods are commonly used to tag a protein of interest at its chromosomal locus. This involves the use of an insertion cassette, a double-stranded linear DNA that contains the fluorescent tag and a selection marker, which is inserted into the target site. It is produced by PCR using a tagging vector as a template. Templates for tagging vectors encompass a variety of fluorescent proteins (e.g. GFP, mCherry, mCitrine), epitopes (e.g. HA, myc), or affinity tags (e.g. GST, TAP, His) that are linked with one of many selectable markers that confer drug resistance or auxotrophic rescue. Families of tagging vectors have been constructed to share PCR-priming sequences. Therefore, a single set of primers made from one template family can be used to insert different tags associated with that template for a given target gene. Additionally, template vectors are available to regulate expression levels from endogenous promoters, constitutively active promoters (e.g. AHD1, GPD1), or regulatable promoters (e.g. GAL1), which can be used for N-terminal tagging. Fluorescent protein and epitope tags can also be used for biochemical techniques including affinity purification, immunoprecipitation, western blot analysis, and immunofluorescence.

Some cassettes, like the pOM family, permit the removal of the selectable marker after tagging [10]. These cassettes are constructed with a marker flanked by LoxP sites, allowing removal of the marker by bacteriophage Cre recombinase that is conditionally expressed from a plasmid [11]. Removing the selectable marker is useful in several situations: 1) to create an N-terminal tag with no separation between the tag and the endogenous promoter of the tagged protein; 2) to insert a tag internally within the coding region of the gene of interest; 3) for multiple rounds of tagging at the same locus; and 4) for use in yeast strains with a limited number of selectable markers. Commonly available tagging vectors are listed in Table 1.

Table 1:

Yeast tagging cassette vectors

Plasmid family	Tag position	Promoter	Tags	Markers
pFA6a 1	C terminal	endogenous	GFP(S65T) 3xHA 13xMyc GST GFPEnvy2 GFPIvy2 GFPγ2	TRP1 kanMX6 HIS3MX6
pFA6a-PGAL1 1	N terminal or internal	GAL1	GFP(S65T) 3xHA GST	TRP1 kanMX6 HIS3MX6
pUR 3	C terminal	endogenous	DsRed	HIS3 URA3(K.l.)
pYM 4	C terminal	endogenous	yEGFP EGFP EBFP ECFP EYFP DsRed, DsRedI RedStar, RedStar2 eqFP611 FlAsH 1xHA, 3xHA, 6xHA 3xMyc, 9xMyc 1xMyc+7xHis TAP Protein A	kanMX4 hphNT1 natNT2 HIS3MX6 klTRP1
pKT 5	C terminal	endogenous	yEGFP yECFP yEVenus yECitrine yESapphire yEmCFP6 yEmCitrine tdimer27 yECitrine+3xHA yECitrine+13xMyc yECFP+3xHA yECFP+13xMyc	KanMX SpHIS5 CaURA3
pOM 8	N terminal or internal	endogenous9	yEGFP 6xHA 9xMyc Protein A TEV-ProteinA TEV-GST-6xHis TEV-ProteinA-7xHis	kanMX6 URA3(K.l.) LEU2(K.l.)
pCY 10	C-terminal	endogenous	Cerulean yECFP yEmCFP yEGFP Venus yEVenus yECitrine yEmCitrine mCherry mEos2 yEc-MYC yEHA yEFlAsH	HygromycinB Zeocin

¹[40]

²[16]

³[41]

⁴[42]

⁵[43]

⁶monomeric version

⁷tandem dimer of DsRed

⁸[10]

⁹After Cre-mediated removal of auxotrophic marker

¹⁰[44]

Tagging vector primers should be designed with sequences that hybridize with both the tagging vector and the target site for homologous recombination within the yeast chromosome. The insertion cassette is produced by PCR using the tagging vector as a template. The resulting amplified DNA is transformed into yeast using a standard protocol [12]. Recombinants that carry the inserted tag are selected using the marker from the insertion cassette, followed by PCR-based screening to ensure the correct insertion size, and finally sequencing to confirm accuracy of the coding sequence.

2.2. Visualization of actin cytoskeleton dynamics

Live-cell imaging of actin dynamics permits analysis of RACF velocity as well as cargo trafficking, such as the movement of mitochondria along actin cables (Fig. 1) [13, 14] Here, we will describe the two methods we use to visualize actin dynamics in living yeast cells.

Fig. 1. Colocalization of the actin cytoskeleton with cargo.

Yeast cells were grown in YPG and washed in SC medium prior to imaging. Left panels: Actin cables visualized with Abp140p-GFP. Middle panels: Two cellular structures, actin patches (top) and mitochondria (bottom), visualized by tagging Abp1p and Cit1p, respectively, with mCherry. Images were captured using standard GFP filters and 300 ms exposure time for GFP and standard DsRed filters and 200 ms exposure time for mCherry. Right panels: merged images showing partial colocalization of actin patches and mitochondria along actin cables. Scale bar = 1 μm.

2.2.1. Visualization of actin dynamics with the fluorescently tagged actin bundling protein, Abp140p

Tagging the actin-encoding ACT1 gene at its C-terminus with GFP compromises actin function: plasmid-borne ACT1-GFP does not rescue the loss of the endogenous ACT1 gene [15]. Generating a functional fusion protein with actin is challenging, because every surface of the actin protein is involved in important protein-protein interactions, including nucleation, polymerization, interaction with motor proteins and other force generators, cable assembly, capping, cross-linking, and severing. As an alternative, actin-binding proteins are often more tolerant of tagging with fluorescent proteins. For example, tagging actin-binding proteins like Abp140p and Abp1p with C-terminal GFP produces a fluorescent signal that localizes to actin cables and actin patches, respectively (Fig. 1) [6, 7]. Other actin proteins that can be tagged with GFP while maintaining normal function are listed in Table 2.

Table 2:

GFP-tagged actin proteins in budding yeast

Actin-containing structure	GFP tagged proteins
Actin patch (early)	Las17p [45, 46],Sla1p [46, 47], Pan1p [46, 48], End3p [49], Sla2p [46, 50], Bzz1p [51], Vrp1 [52]
Actin patch (mid)	Myo3p [53], Myo5p [54], Bbc1p [51],
Actin patch (late)	Abp1p [15, 46, 55, 56], Arp2p [45], Arp3p [45], Arc15p [46, 57], Sac6p [56, 58], Cap1p [59], Cap2p [60], Scp1p [61], Cof1p [62]
Actin cable (vegetative)	Abp140p [6], Lifeact (Abp140 aa 1–17) [18, 31]

Even when protein function is preserved, fluorescent protein fusions may require optimization for successful imaging. For example, Abp140p-GFP produces only a low signal in the cells propagated using glucose-based media. However, this can be improved by culturing cells in a non-fermentable carbon source, such as lactate. Cells grown in lactate medium produce cables that are thicker, and therefore more visible (Note 1). Alternatively, improved signal to noise ratio can be achieved by tagging actin-binding proteins with GFPEnvy. GFPEnvy is an improved version of GFP with brighter signal and photostable, which was constructed by combining mutations found in Superfolder GFP and GFPγ [16]. Tagging Abp140p with GFPEnvy improves signal intensity and reduces photobleaching during time-lapse videos. GFP and GFPEnvy do not disrupt actin dynamics when fused to Abp140p; however, other fluorophore proteins such as td-Tomato, compromise the function of Abp140p. Moreover, although GFPEnvy was produced from a monomeric form of GFP, recent studies indicate that it can dimerize in vitro [17]. Thus, it is important to test whether actin dynamics are preserved when using fluorescent protein tags.

2.2.2. Visualization of actin dynamics with the fluorescently tagged actin-binding peptide Lifeact

An alternative probe for visualizing actin dynamics is, Lifeact, an actin-binding peptide, tagged with a fluorescent protein. Lifeact consists of the first 17 amino acids of Abp140p, and is sufficient to mediate actin localization of fluorescent proteins without disrupting dynamics [18]. Lifeact has been used across many eukaryotic organisms, including fungi, plants, and invertebrates [19–25]. However, there are some caveats to using an exogenous construct to visualize the actin cytoskeleton. Lifeact fails to label some actin structures, such as stress-induced cofilin-bound actin rods [26]. Moreover, Lifeact overexpression can alter actin dynamics, generating artifacts. For example, overexpressing Lifeact in Drosophila germline cells causes loss of cortical actin integrity leading to fertility defects [27]. In fission yeast, highly expressed Lifeact-mCherry or mEGFP-Lifeact affects actin patch formation and cytokinesis [28]. The structure of the Lifeact–F-actin complex indicates that dosage-dependent effects are caused by the reduced binding of cofilin to actin filaments [28–31]. Therefore, expression conditions should be optimized for each application.

Initial reports of Lifeact in budding yeast indicated that it can label actin-containing structures, but its fluorescence intensity was extremely weak, preventing long-term imaging [18, 31]. Our lab has resolved this issue by integrating Lifeact conjugated to GFPEnvy, rather than GFP, into the yeast genome for stable expression.

The stable genomic expression of Lifeact-GFPEnvy (LA-Envy) at the HO locus does not interfere with cell growth (Fig. 2A). Moreover, our previous studies indicate that Abp140p-GFP labels a small number of punctate structures (e.g. Fig. 2C) that either are not actin patches or are a subset of actin patches. On the other hand, we find that LA-Envy localizes to both punctate and filamentous structures, which co-localize with phalloidin-stained actin patches and cables (Fig. 2B). Equally important, we find that the expression of LA-Envy does not result in depolarization of the actin cytoskeleton. The presence of >5 actin patches in the mother cell is a hallmark of cell depolarization and we found LA-Envy expressing cells contain < 5 punctate structures per mother cell (Fig. 2C).

Fig. 2. Visualizing actin cable dynamics with Lifeact-GFPEnvy.

(A) Maximum growth rates and growth curves of untransformed cells (WT), and cells expressing Lifeact-GFPEvny or Abp140p-GFPEnvy in YPD or SC media at 30 °C. Maximum growth rates were calculated for 2 hrs intervals (n = 3–6 cultures for each independent experiment; statistical significances were calculated by unpaired Student’s t-test. Error bars represent SEM (standard error of the mean). (B) Representative images of fixed mid-log phase yeast cells in lactate medium. Left panels: F-Actin structures stained with Alexa568-phalloidin. Middle panels: Lifeact or Abp140p tagged with GFPEnvy. Right panels: Merged images, showing that actin cables visualized by Lifeact-GFPEnvy or Abp140p-GFPEnvy co-localize with phalloidin. White arrows indicate punctate structures (actin patches) in mother cells. Lifeact-GFPEnvy reveals more punctate structures than Abp140p-GFPEnvy. Scale bars,1 μm. (C) Left panels: Quantification of punctate actin structures or actin cables in mid-log phase cells expressing Lifeact-GFPEnvy or Abp140p-GFPEnvy in lactate or SC media at 30°C. n > 30 cells/trial. Statistical significances were calculated by unpaired t-test (*p < 0.05; **p < 0.01; ****p < 0.0001). Error bars represent SEM. Right panels: Representative images of live yeast cells expressing Lifeact-GFPEnvy or Abp140p-GFPEnvy grown in the indicated media. Scale bars, 1 μm.

When we compared labeled cables in cells expressing LA-Envy and Abp140p-GFPEnvy, we observed fewer cables in LA-Envy-expressing cells in both lactate and SC medium, compared to Abp140p-GFPEnvy-expressing cells (Fig. 2C). However, there was no detectable difference in actin cable number in phalloidin-stained cells expressing these two probes (data not shown). Taken together, our results suggest that LA-Envy and Abp140p-GFPEnvy label both patches and cables, but with different affinities; patches are more likely to be labeled with LA-Envy and cables with Abp140p-GFPEnvy. At least in the case of cables, the difference does not appear to be an artifact of fusion protein expression, but rather arises from a difference in binding that renders some structures below the threshold of detection. It remains to be determined why this phenomenon occurs, but we speculate that it results from competitive binding between actin cables and patches.

Overall, we find that Lifeact-Envy is the first actin probe that labels both actin cables and patches in live-cells. Table 2 lists known functional fluorescent protein-labeled actin-binding proteins that serve as probes for actin patches or actin cables in budding yeast. Finally, Table 3 lists suggested imaging conditions for common fluorescent protein-labeled actin-binding proteins.

Table 3:

Suggested imaging conditions for methods described in this chapter

Fluorescent protein	Excitation/Emission	Light source	Exposure time
Abp140p-GFP	488/507	LED 470 nm (45%) or Metal-Halide Lamp + standard GFP Filter	250 ms
Abp140p-GFPEnvy	488/507	LED 470 nm (45%) or Metal-Halide Lamp + standard GFP Filter	250 ms
Lifeact-GFPEnvy	488/507	LED 470 nm (45%) or Metal-Halide Lamp + standard GFP Filter	250 ms
Abp1p-mCherry	587/610	LED 550 nm (45%) OR Metal-halide lamp + standard DsRed filter	200 ms

3. Materials

3.1. Yeast growth media

1. Amino acid supplements: For every 1 liter of synthetic complete medium, supplement with all of the following. For dropout medium, omit one or more components to select for the desired strains: 10 mL adenine (2 mg/mL stock in 0.05 M HCl), 10 mL uracil (2 mg/mL stock in 0.5% NaHCO₃), 10 mL arginine (1 mg/mL stock in H₂O), 10 mL histidine (1 mg/mL stock in H₂O), 10 mL leucine (10 mg/mL stock in H₂O), 10 mL lysine (3 mg/mL stock in H₂O), 10 mL methionine (2 mg/mL stock in H₂O), 10 mL phenylalanine (5 mg/mL stock in H₂O), 10 mL tryptophan (2 mg/mL stock in H₂O), 30mg tyrosine.

2. Synthetic complete medium (SC): For every 1 liter, dissolve 6.7 g yeast nitrogen base without amino acids, 20 g glucose, and amino acid supplements as needed in 800 mL distilled H₂O. Adjust pH to 5.5 with NaHCO₃ and bring volume to 1 L with distilled H₂O. Sterilize by autoclaving.

3. Yeast peptone-dextrose medium (YPD): For 1 liter, dissolve 10 g yeast extract, 20 g bacto-peptone, and 20 g glucose in distilled H₂O and bring volume to 1 L with distilled H₂O. Sterilize by autoclaving.

4. Lactate medium: For 1 liter, combine 3.0 g yeast extract, 0.5 g glucose, 0.5 g CaCl₂, 0.5 g NaCl, 0.5 g MgCl₂, 1 g KH₂PO₄, 1 g NH₄Cl, 22 mL 90% lactic acid, and 7.5 g NaOH pellets. Dissolve ingredients in 800mL distilled H₂O. Adjust pH to 5.5 with NaOH. Bring volume to 1 L with distilled H₂O.

5. Synthetic raffinose medium: For 1 liter, combine 6.7 g of yeast nitrogen base without amino acids and amino acid supplements as needed. Dissolve ingredients in 600 mL distilled H₂O. Adjust pH to 5.5 with NaHCO₃. Bring volume to 900 mL with distilled H₂O. Supplement after autoclaving with 100 mL filter-sterilized 20% raffinose.

6. Synthetic glycerol medium (SG): For 1 liter, combine 6.7 g of yeast nitrogen base without amino acids, 30 mL of glycerol, 0.5 g glucose, and amino acid supplements as needed. Dissolve ingredients in 800 mL distilled H₂O. Adjust pH to 5.5 with NaHCO₃.

7. Synthetic galactose medium: For 1 liter, combine 6.7 g of yeast nitrogen base without amino acids and amino acid supplements as needed. Dissolve ingredients in 600 mL distilled H₂O. Adjust pH to 5.5 with NaHCO₃. Bring volume to 950 mL with distilled H₂O. Supplement after autoclaving with 50 mL filter-sterilized 40% galactose.

3.2. Reagents for PCR amplification

1. Betaine solution 5M, PCR reagent (Sigma, B0300)

2. KAPA HiFi HotStart Ready Mix (2x) (Kapa Biosystems)

3. Sterile distilled H₂O

4. Forward and reverse primers (diluted to 50 μM each)

3.3. Reagents for yeast transformation and marker excision

1. 50% w/v Polyethylene glycol 3350 in H₂O

2. 1 M lithium acetate in H₂O

3. 0.1 M lithium acetate in H₂O

4. Carrier DNA: 2 mg/mL single-stranded DNA from salmon testes in H₂O. Store at −20°C. (Sigma, D7656)

5. DNA for transformation (1 PCR reaction as described in protocol, or 1μg)

6. Sterile distilled H₂O for resuspending cells before plating

7. Selective plates for growth of cells containing plasmid or integrated DNA

4. Methods

4.1. Modification of yeast genes at their chromosomal loci

4.1.1. Primer design

Generally, primers should be designed with 60 or more bases. The 5’ end of each primer consists of at least 40 bases with perfect homology to the target site. As an example, for C-terminal tagging with a fluorescent protein, the 5’ end of the forward primer should contain the 40–45 bases directly upstream of the stop codon, while the 5’ end of the reverse primer should contain the reverse complement of the 40–45 bases directly downstream of the stop codon. The 3’ ends will consist of 18–25 bases complementary to the sequences that will be inserted from the vector, which include the open reading frame of the fluorescent protein, the transcription termination site, and the selectable marker (Note 2).

For further preservation of protein function and folding when tagging with a fluorescent protein, one can insert a short linker between the gene of interest and the fluorescent tag. For instance, GFP fluorescence can often be further optimized by increasing the length of the linker between the target gene and GFP molecule (e.g. by adding five alanines). The linker length and amino acid composition may need to be adjusted for different target proteins. Typically, proteins with processive enzymatic functions, such as polymerases and telomerases, benefit from poly-glycine linkers [32]. When C-terminally tagging, the linker should be incorporated into the forward primer by including codons encoding the appropriate amino acids between the 40 bases of homology to the target gene and the 20 bases corresponding to the template. A similar strategy can be used in the reverse primer for N-terminal tagging.

4.1.2. Amplification of insertion cassette from tagging vector

1. Prepare the PCR reaction: For 50 μL, 10 μL Betaine 5M (Sigma, B0300), 25 μL Kapa HiFi HotStart ReadyMix (2x), 12 μL sterile distilled H₂O, 1 μL each of forward and reverse primers, 1 μL template DNA. A single 50-μL PCR reaction will provide sufficient DNA for a single yeast transformation (minimally 1 μg of DNA) if a high-fidelity, high-yield polymerase is used.

2. Amplify the cassette using the following thermocycler conditions: Initial denaturation cycle at 95°C for 2–5 minutes; 35 cycles of: 98°C for 20 s, 60–75°C (Note 3) for 15 s, and 72°C for 60 s/kb; final extension cycle at 72°C for 1–5 minutes.

3. Use the PCR reaction directly in the lithium acetate transformation protocol described below. Alternatively, amplified DNA can be purified using any commercially available PCR purification kit (but see Note 4).

4.2. Lithium acetate transformation of yeast

The lithium acetate transformation protocol is the most commonly used method for yeast transformations [12]. The following protocol is for one transformation reaction. A negative control containing no DNA should always be carried out in parallel.

1. Grow yeast to mid-log phase (0.2–0.5 OD₆₀₀).

2. Transfer 10⁷ cells from the culture to a 1.5 mL microfuge tube.

3. Pellet cells (30 s, 6000 x g) and wash with 500 μL 0.1 M lithium acetate. Resuspend in 240 μL of 50% w/v polyethylene glycol 3350.

4. Add 36 μL 1.0 M lithium acetate, 25 μL of carrier DNA, and 1 μg (50 μL) of DNA to be transformed (PCR-amplified insertion cassette or plasmid DNA) or 50 μL of H₂O for negative control.

5. Vortex vigorously and incubate in a water bath at 30°C for 30 min.

6. Heat shock in a water bath at 42°C for a minimum of 20 min and a maximum of 3 hrs.

7. Concentrate cells by mild centrifugation (30 s, 6000 x g).

8. For transformations using auxotrophic markers, resuspend cell pellet gently in 100 μL of sterile distilled H₂O and plate on appropriate selective media.

9. For transformations using drug resistance markers (e.g. KanMX6), resuspend cell pellet gently in 500 μL of YPD or synthetic complete medium with dropouts as needed to maintain plasmids. Allow cells to recover at 30°C for 2–4 hrs. Concentrate and plate cells as described in steps 7–8 on appropriate selective medium containing drug.

4.3. Marker excision by Cre recombinase

If using a LoxP-containing vector such as the pOM family for tagging a gene of interest, the selectable marker can be removed by the following method.

1. Verify tagging by PCR and sequencing, as described below.

2. Using methods described above, transform the tagged strain with a plasmid that encodes Cre recombinase under control of a galactose-inducible promoter [33]. Select transformants on appropriate medium selecting for the Cre plasmid.

3. Grow cells to mid-log phase in 5 mL liquid medium in a 50-mL conical-bottom tube. Use medium that selects for the Cre plasmid.

4. Pellet cells (5 min, 4000 x g) and resuspend in 5 mL galactose medium.

5. Incubate cells in galactose medium in a shaking incubator at 30°C for 12–16 hrs.

6. Plate an aliquot of cells on non-selective (YPD or SC) medium.

7. Screen colonies for loss of the tagging cassette marker by replica plating on selective plates.

8. To induce dropout of the Cre plasmid, grow cells in non-selective liquid medium for 24–48 hrs, plate on non-selective medium, and screen for loss of the plasmid by replica plating on selective plates (Note 5).

4.4. Validating and characterizing fluorescent protein-tagged cytoskeletal proteins

4.4.1. PCR screening and sequencing

Validating genomic tag insertion at the target locus can be done using a PCR screening method. Use isolated genomic DNA from transformants and untransformed cells as templates for the PCR reaction, and compare fragment sizes amplified by a primer pair that hybridizes up- and downstream of the tagging locus. Sequencing should then be carried out to verify that tagged genes do not carry any mutations.

4.4.2. Verification of protein expression and function

Initial steps to confirm successful expression of tagged proteins in transformed cells can be performed through visual inspection. Prepare cells for short-term imaging as described below and screen several colonies for consistent fluorescent protein expression.

We recommend using Western blot analysis to study expression of tagged proteins. Benefits of protein expression monitoring include 1) comparing the expression level of tagged protein to that of its native form; 2) troubleshooting protein expression when the levels fall below the threshold of detection by microscopy; and 3) identifying degradation products that may hinder proper localization and function.

It is important to evaluate the localization and functions of tagged constructs and confirm that these are preserved, when knowledge of the gene of interest is available. Tagging proteins with fluorescent proteins may alter their function and/or localizations by disrupting native protein-protein interactions, or by inducing oligomerization through the fluorescent protein. Evaluating whether function is preserved in fluorescent protein-tagged strains can be done by examining their cell growth rates and loss of any characterized phenotypes of the gene of interest. Fluorescent protein-tagged strains with compromised protein function will exhibit similar phenotypes to cells bearing mutations in that gene. For a positive indicator of function, a plasmid-borne version of the fluorescent protein-tagged protein can be used to rescue a wild-type phenotype in yeast cells with a genomic deletion of the gene of interest (Note 6). After confirming proper localization and preserved function, strains can be used for live-cell analysis. If, on the other hand, target protein localization and/or function is found to be compromised by a fluorescent protein tag, an alternative approach is to fuse the target protein to a smaller epitope tag such as myc or HA and visualize the protein in fixed cells using immunofluorescence.

4.5. Wide-field imaging of living cells

4.5.1. Equipment for wide-field imaging

Budding yeast are relatively small in comparison to other eukaryotic organisms. The diameter of mother cells is typically 4–6 μm. Therefore, a microscope equipped with a high-magnification, high-numerical aperture objective lens and a sensitive camera are required to resolve small intracellular structures with good spatial resolution.

Light throughput and sensitivity of an imaging system are imperative for live-cell imaging because short exposure times and low illumination intensity are needed to reduce photobleaching and phototoxicity. Short exposure times are also important for improving temporal resolution when imaging dynamic structures. Additional time resolution improvement can be achieved by using a direct trigger connection between the acquisition computer and imaging components, and maximizing the computer’s processor speed, random access memory (RAM), and storage disk speed.

Examples of epifluorescence microscope setups that can be used for wide-field live yeast imaging include the following:

• Inverted AxioObserver.Z1 microscope (Carl Zeiss Inc., Thornwood, NY) equipped with a metal halide lamp and Colibri LEDs for excitation; 100x/1.3 EC Plan-Neofluar oil-immersion objective; and an Orca ER cooled CCD camera with 1280 × 1024 pixel resolution (Hamamatsu, Bridgewater, NJ). ZEN software (Carl Zeiss) is used to control the camera and microscope hardware.

• Upright Axioskop 2 microscope (Carl Zeiss Inc., Thornwood, NY) equipped with a pE-4000 LED illumination system (CooLED, Andover, UK) for excitation; 100x/1.4 Plan-Apochromat objective (Carl Zeiss Inc., Thornwood, NY); and an Orca ER cooled CCD camera. NIS Elements 4.60 Lambda software (Nikon, Melville, NY) is used to control the camera and microscope hardware.

4.5.2. Multi-Color Imaging

Live-cell imaging is not limited to visualizing one organelle at a time. Multiple organelles can be imaged at once with the use of different fluorescent proteins or vital dyes, allowing the analysis of the interaction between organelles or morphological changes within multiple organelles. As described earlier, S. cerevisiae is amenable to integration of a wide variety of fluorescent proteins at multiple chromosomal loci. This can be achieved by sequential tagging through multiple homologous recombination events or through crossing different lines.

The limiting factor for multi-color imaging is identifying a compatible collection of distinct fluorescent proteins that are sufficiently bright, spectrally distinct, and photostable, and that do not compromise protein function. The most commonly paired fluorescent proteins for 2-color imaging are GFP (or GFPEnvy) and red fluorescent protein (RFP), such as mCherry. These fluorescent proteins have distinct spectra and display consistently bright and robust signals. For 3-color imaging, the most commonly combined fluorescent proteins are GFP, RFP, and blue fluorescent protein (BFP) [34]. However, BFP requires UV excitation, which can lead to organelle fragmentation, reactive oxygen species production, and cell death [35]. As an alternative, our lab developed a trio of fluorescent proteins that are photostable, fluorescently distinct and well tolerated by yeast: mTFP1/mCitrine/mCherry [36].

Multiple fluorescent proteins can be detected either simultaneously or sequentially. Simultaneous detection requires either splitting the emitted signal into multiple detectors, or dividing the camera area into multiple regions using an image splitter (e.g. DualView, Optical Insights, Santa Fe, NM). Although this approach maximizes speed of acquisition, it entails a significant risk of spectral bleedthrough. Therefore, in most cases, sequential imaging offers sufficient speed and more reliable differentiation between labels. There are three strategies to achieve sequential imaging: 1) changing excitation and emission filters, 2) LED excitation switching, and 3) laser excitation switching. Changing excitation and emission filter sets requires a motorized microscope stand that rotates the filter turret, or separate filter wheels for excitation and emission. Rotating the filter turret allows the use of specialized cubes for each channel, which will result in the best signal. However, turret rotation may induce vibration in the system, and is also quite slow (> 1 s) compared to that of separate filter wheels (tens or hundreds of ms).

A faster, more stable approach is switching the excitation wavelength directly at the light source. This is accomplished using either lasers or narrow-spectrum, high intensity light-emitting diodes (LEDs), such as those listed in the example microscope configurations above. These systems can be configured with multiple wavelengths that match the spectra of commonly used fluorescent proteins and are controlled individually by electrical signals with virtually no lag time. Signal can be collected through a multi-band dichroic/emission filter cube, eliminating any mechanical motion. Thus, these systems are preferred for multi-color imaging of dynamic structures and for excitation ratio imaging.

4.5.3. Preparing cells for general short-term imaging

For short-term visualization, concentrated cells can be deposited directly on a slide. However, cells on a microscope slide should not be imaged for more than 10 min. After this point, cells exhibit signs of stress and significant decreases in viability. The following protocol can be used for general imaging of cells expressing fluorescent proteins. Special procedures for imaging actin cable dynamics are detailed in the next section.

1. Grow yeast cells overnight to mid-log phase (OD₆₀₀ = 0.2–0.5) in 5 mL of appropriate liquid medium in a 50-mL conical-bottom tube at 30°C with shaking (Notes 7).

2. Transfer 1 mL of mid-log phase culture to a 1.5-mL microfuge tube and concentrate cells by centrifugation at ~6,000 x g for 30 s (Note 8).

3. Without disturbing the pellet, remove almost all of the supernatant, leaving a volume of supernatant in the tube approximately twice the volume of the pellet.

4. Resuspend pellet in residual medium and transfer ~1.5 μL of the cell suspension to a cleaned microscope slide (Note 9).

5. Apply a #1.5 (170 μm thick) coverslip, taking care to avoid creating bubbles between slide and coverslip (Note 10).

6. Acquire images. Prepare a fresh slide from the concentrated cell suspension after 10 min.

4.5.4. Preparing cells for short time-lapse imaging of actin cable dynamics

Short-term imaging can be used to determine the velocity of actin cable movement. Concentrated cells in lactate medium are applied directly to a slide. This protocol can be used to follow actin cable dynamics using fluorescent proteins conjugated to either Abp140p or Lifeact.

1. The morning before the day of the experiment, prepare a pre-culture. Inoculate cells from a colony into 5 mL of appropriate medium in a 50-mL conical-bottom tube and incubate 4–5 hrs in a 30°C shaking incubator.

2. From the pre-culture, inoculate an overnight culture with fresh media to reach mid-log phase (OD₆₀₀ 0.2–0.5) the following day.

3. On the day of the experiment, transfer 1 mL of culture to a 1.5-mL microfuge tube and concentrate cells by centrifugation at ~6,000 x g for 30 s (Note 9).

4. Without disturbing the pellet, remove almost all of the supernatant, leaving a volume of supernatant in the tube approximately twice the volume of the pellet.

5. Resuspend pellet in residual medium and transfer ~1.5 μL of the cell suspension to a cleaned microscope slide.

6. Apply a #1.5 (170 μm thick) coverslip, taking care to avoid creating bubbles between slide and coverslip (Note 10).

7. Acquire time-lapse series at a single focal plane, at 500 ms intervals, for 30 s (6s 11 and 12).

8. Analyze cable velocity as described in Section 4.8.

4.5.5. Optimizing imaging conditions and preventing toxicity

During live-cell imaging, extended illumination of cells can lead to phototoxicity by two primary mechanisms: 1) direct cell damage by photons with wavelengths close to the UV; and 2) free radical and reactive oxygen species production by the interaction between photons and cellular molecules. The following criteria can be used to check for photodamage to cytoskeletal function.

• Cytoskeletal structures: intense or long excitation can cause actin cable depolymerization.

• Dynamics: While behavior of photodamaged cytoskeletal components has not been characterized, photodamage can be suspected if there are alterations in cytoskeletal velocity or other motility parameters in wild-type cells.

Even if cells remain healthy throughout the imaging process, excessive light exposure still can cause photobleaching of fluorophores. Phototoxicity and photobleaching can be reduced by the following strategies. Resolution, sensitivity, and toxicity must always be balanced. However, in general, longer exposures at low intensity are less phototoxic than shorter exposures at high intensity.

1. Reduce excitation intensity. For LED and/or lasers, adjust intensities through acquisition software. Alternatively, add a neutral density filter.

2. Reduce exposure time. However, be sure to maintain enough dynamic range in pixel intensities to produce an interpretable image.

3. Reduce spatial or time resolution. For example, increase the time interval and/or the z-series interval.

To increase signal-to-noise ratio without increasing light exposure:

1. Apply binning in the camera. This will result in decreased spatial resolution, but this can be alleviated by adding a projection tube before the camera.

2. Increase camera gain.

3. Reduce camera readout speed, if possible.

4. Use a filter set with a broader spectral window or more efficient coatings to increase throughput.

4.5.6. Special considerations for quantitative analysis

Two criteria must be met for quantitative fluorescence imaging: 1) emitted fluorescence must be proportional to the number of fluorescent molecules present; and 2) recorded pixel intensities must be proportional to the amount of light emitted by the sample. Under well-controlled conditions, these criteria are met at sufficient levels and can be used to measure many biological factors.

Absolute quantification of fluorescent molecules by imaging is a serious challenge due to many variables that complicate the linearity of fluorescence emission and pixel intensities [37]. However, proper controls and normalization can allow reliable measurements of relative changes in the volume and intensity of fluorescent structures, providing a powerful tool for hypothesis testing. Minimally, when comparing intensities, the following variables must be controlled in any wide-field fluorescence imaging experiment:

• Age and concentration of fluorescent probe

• Sample age, unless photobleaching is known to be negligible

• Objective lens

• Illumination spectrum and intensity, including filters, field aperture setting, and light source alignment

• Camera model, binning, exposure time, gain, offset, readout speed, and bit depth

Imaging parameters should always be adjusted to produce pixel values above the background level of the detector and sample, but below saturation if intensities are compared between groups, adjust imaging parameters using the brightest samples, to avoid saturating the detector. In addition, sample size (number of cells imaged) must be sufficient to determine the variability within the population, and fields of view must be selected without conscious or unconscious bias.

4.6. Deconvolution of wide-field fluorescence images

Typically, the resolution of wide-field fluorescence z-series can be further improved by deconvolution. Deconvolution is a computational method for removing out-of-focus light from wide-field fluorescence images. As a result, images are sharper with a higher signal:noise ratio and enhanced three-dimensional information. Various algorithms are available, however, only some allow for quantitation after processing [38]. Deconvolution can be executed by algorithms provided by commercial software such as Volocity (Quorum Technologies) or by free open-source software, such as Fiji/ImageJ. We have found that Volocity and Fiji/ImageJ produce comparable results. For best deconvolution results, use z-series acquired with an interval of 0.2–0.3 μm.

Volocity Deconvolution

1. Acquire z-series images using general short-term imaging conditions described above.

2. Load datasets in Volocity and verify that the x, y, and z scale are correctly set.

3. For each channel, generate a new calculated PSF. When prompted, enter the numerical aperture of the objective lens and the maximum wavelength of fluorescence emission (e.g. 507 nm for GFP and 620 nm emission wavelength for mCherry).

4. Using the calculated PSF for each channel, perform iterative deconvolution with 60 iterations, 100% confidence limit (Note 13). Number of iterations can be adjusted according to signal intensity and noise level.

Fiji/ImageJ Deconvolution

1. Download and install Diffraction PSF 3D (https://imagej.net/Diffraction_PSF_3D) and Iterative Deconvolve 3D (https://imagej.net/Iterative_Deconvolve_3D) plugins from the Fiji website (https://imagej.net/Fiji/Downloads).

2. Create a calculated PSF by using Diffraction PSF 3D: Plugins > Diffraction PSF 3D. Set up PSF parameters by entering immersion medium refractive index, objective lens numerical aperture, emission wavelength, image pixel size, image z step size, image width, image height, and image z slice number. The following are typical settings using our microscopy setup:

   • refractive index: 1.518 (if using immersion oil)
   • numerical aperture: 1.40
   • emission wavelengths: 507 nm or 610 nm for GFPEnvy and RFP, respectively
   • longitudinal spherical aberration at maximum aperture: 0
   • image pixel spacing: 64.50 (substitute your own spacing in nm if applicable)
   • slice spacing: 300 (substitute your own spacing in nm if applicable)
   • image width: 1343 (substitute your own image size in pixels if applicable)
   • image height: 1023 (substitute your own image size in pixels if applicable)
   • image depth: 21 (substitute your own z-stack size if applicable)
   • normalization: sum of pixel values =1
   • Name your PSF with the corresponding wavelength for easy recognition.

3. Deconvolve selected images: Plugins > Iterative Deconvolve 3D. Select the image and corresponding PSF, set Wiener filter gamma to 0, maximum iterations to 60, and delta for termination to 0. The deconvolution will be terminated automatically if iterations do not produce better resolution.

4. Save the deconvolved file.

4.7. Quantification of fluorescent signals

To quantify the amount of actin either within the whole cell or sub-regions of the cell, two common measurements of fluorescence signal intensity can be executed: mean and integrated intensity. Mean fluorescence intensity (mean gray value) is the average value of pixels in a cell or region of interest (ROI). Mean intensity can be a proxy for concentration of a fluorophore, but is not a good indicator of the total amount of a fluorophore, unless the cells or compartments are of similar size. For example, a small bud and a large bud may have the same number of labeled actin structures, but the small bud will have a higher mean intensity because the structures are crowded into a smaller volume. In scenarios where ROI sizes vary greatly, it is best to approximate the total amount of a fluorescent probe by calculating the integrated density (integrated intensity), which is the sum of all of the pixel values in the region. Thus, the integrated density takes into consideration not only the mean pixel value, but the area of the ROI. Here, we show how to obtain intensity measurements using Volocity and Fiji/ImageJ. For more sensitive and accurate measurements, background should be measured and subtracted from mean intensity values. To correct integrated intensity for background, subtract the product of the background intensity and the area of the ROI.

Using Volocity to measure integrated intensity

1. From the Measurements tab, create a protocol by adding the following steps in order: Find Objects Using Intensity, Exclude Objects by Size, Clip Objects to ROIs, Measure Objects.

2. For images with multiple channels, make sure that the correct channel is selected under the Find Objects Using Intensity tab.

3. Under Exclude Objects by Size, select a size criterion that will exclude background pixels or other artifacts of image capture or deconvolution. This value will depend on imaging conditions and must be determined empirically.

4. Under the Measure Objects tab, click the gear icon to select which measurements you wish to use. Check Intensity and Volume Measurements.

5. Define the ROI you wish to measure. Volocity allows rectangular, circular, and freehand ROIs. Use the Zoom function to make it easier to outline the desired area.

6. Adjust the threshold under the Find Objects Using Intensity tab by manipulating the lower and upper limits. We suggest maximizing the upper limit to include all pixels that have high intensity values. The lower limit should be adjusted to remove enough background to include only signal that is found within the cellular structure of interest.

7. All measurements will appear in a tab below the image. Individual objects will be separated (for example, if mitochondrial fluorescence intensity is being measured and there are 3 mitochondrial structures, three measurements will be provided, each color-coded to depict which structure they represent). These measurements can be copied into a spreadsheet application such as Microsoft Excel or saved within Volocity by clicking Make Measurement Item under the Measurement tab.

8. In the results table, the Sum column reports the integrated intensity of each object. The sum of all of these values is the integrated intensity within the ROI.

Using Volocity to measure mean intensity

1. Define the ROI you wish to measure including background ROIs (outside the cell). Volocity allows rectangular, circular, and freehand ROIs. Use the Zoom function to make it easier to outline the desired area.

2. Click Measure Object tab to measure the signal intensity.

3. All measurements will appear in a tab below the image. These measurements can be copied into a spreadsheet application such as Microsoft Excel or saved within Volocity by clicking Make Measurements under the Measurement tab.

4. Record the mean gray value (total fluorescence of area divided by area pixels).

5. Calculate corrected intensity measurements: subtract the background mean intensity from the mean intensity of each measured ROI.

Using Fiji/ImageJ to measure mean or integrated intensity

1. From the Analyze menu, choose Set Measurements. Check the following: Area, Mean Gray Value, Integrated Density, Display Label.

2. Subtract background intensity in the image:

   1. Generate a maximum projected image: Image > stacks > Z project. Include all slides by selecting the start and stop slice. Select “Average Intensity” as the projection type.
   2. Use the Rectangle selection tool to select a background region (outside the cells). Measure the mean intensity in this region.
   3. Subtract the mean background mean intensity from the original image: Process > Math > Subtract. Enter the mean background intensity from the last step. Process all slides. Using the ROI tool, define the ROI you wish to measure.

3. Using one of the ROI tools, define the ROI you wish to measure.

4. From the Analyze menu, choose Measure (or press M).

5. Copy or save the measurements that appear in the results window.

4.8. Analysis of actin dynamics

The methods described here employ Fiji/ImageJ [39] as the tool of choice in our own laboratory. Similar functions are available in most image analysis and processing software packages.

Tracking movements involves marking the position of an object at successive timepoints. For structures with elongated, irregular or dynamic shapes, like actin cables, a point can be defined on the object of interest and can be used as the reference point for velocity measurements. For example, when tracking actin dynamics with Abp140p-GFP-labeled actin, heterogeneity of Abp140p binding produces bright dots along the actin cable, and these areas serve as fiduciary marks to track movement [6]. Tracking can provide several quantitative measurements including velocity, distance traveled, and direction of movement. Here, we show how to calculate velocity.

4.8.1. Measuring retrograde actin cable flow velocity

1. Identify individual cells displaying cable motility by rapidly scrolling through the movie. Configure the Point Tool on the Fiji toolbar to Add to Overlay and Label Points. Then mark and label each cell of interest using the Point Tool.

2. Capture and save a screenshot showing the numbered cells of interest: Plugin > Utilities > Capture screen.

3. For each cell of interest, crop the movie to include only that cell, and save a copy with the cell number.

4. To track actin cable movement, use the Fiji Manual Tracking plugin.

5. Manually enter the microscope and image parameters, including time interval (e.g. 0.5 s) and x/y calibration (pixel size in μm). To generate an overlay showing the track, also check the following boxes: Tracking > Show path; Drawing > Overlay Dots & Lines.

6. Configure the Point Tool on the Fiji toolbar to Auto-measure and Auto-Next Slice.

7. Set the movie to the first timepoint to be analyzed, and click Add track.

8. Begin tracking a cable by clicking on a point of interest: either the leading or trailing end, or a fiduciary mark.

9. Continue marking the motile structure for the duration of observable movement.

10. Once tracking is finished, click End track. This will produce two results windows:

    1. The tracked path drawn as an overlay on the image. Use this to confirm that the movement of the actin cable matches the manual tracing.
    2. The Results table, giving position, slice number, distance, and velocity.
11. Manually calculate the incremental velocity by the following (Note 14):

    1. $Velocity=\frac{TotalDistance}{TotalTime(Slice\#\mathrm{*}VideoTimeInterval)}$

5. Notes

1. Abp140p-GFP signal intensity is higher in yeast propagated using non-fermentable carbon sources. Lactate medium is the preferred choice as it is not autofluorescent and provides more nutrients than synthetic complete medium with glycerol (SG). However, fragmentation of mitochondria may occur in lactate medium. If this is a problem, use glycerol-based media, such as SG.

2. The sequences for the 3’ ends are often provided by the designers of the tagging plasmid. In many cases, the forward primer will include the sequence of the polylinker upstream of the fluorescent protein tag while the reverse primer will include the reverse complement of the last few nucleotides of the selection marker, or some sequence downstream of the selection marker.

3. The second step of the 35-cycle procedure is the primer annealing phase. The temperature used should matched to the melting temperature of the DNA. For primers that have secondary structures, or when primer sequences cannot be manipulated to make the melting temperatures comparable, betaine, DMSO, or both can be added to the PCR reaction to a final concentration of 1 M and 5%, respectively (10 μL and 5 μL, respectively for a 50 μL reaction).

4. For most PCR reactions, we find that product purification is unnecessary. While purification will eliminate some contaminants from the PCR reaction, the accompanying loss of DNA from the process often outweighs the benefits of performing the purification. We specifically find that when using the KAPA HiFi HotStart or Platinum Taq DNA Polymerase High Fidelity PCR reaction kits, purification to eliminate contaminants is unnecessary.

5. The galactose-inducible Cre recombinase plasmids are CEN plasmids, which can often be dropped out by continuous growth on non-selective medium. Alternatively, pSH47 can be used as this galactose-inducible Cre recombinase expression plasmid contains the URA3 selectable marker which can be removed from cells by counter-selection using 5-fluoroorotic acid (5-FOA) plates after marker selection.

6. When performing rescue/plasmid complementation assays, it is important to express the fluorescent protein-tagged protein under the endogenous promoter of the gene. This will avoid unexpected artifacts, such as overexpression of a partially functional protein rescuing wild-type phenotypes.

7. For live-cell imaging, avoid using medium containing yeast extract (e.g. YPD, YPG) as the yeast extract is autofluorescent and will require repeated washing prior to imaging. Instead, use synthetic complete or lactate medium.

8. Avoid excessive/extended centrifugation, which causes actin cable depolymerization and mitochondrial fragmentation.

9. Remove as much glass debris and dust from the slide before adding the cell suspension to ensure a single-cell layer is formed when applying the cover slip.

10. Excess volume of cell suspension can cause cells to float and move during image acquisition, while insufficient volume can compress cells or cause uneven spreading. Slight pressure should be applied to the edges of the coverslip to stably trap cells between the slide and coverslip and avoid movement of cells during image acquisition. Take care not to put too much pressure, or cells may burst.

11. Actin cable dynamics are quite sensitive; acquire no more than four 30 s datasets per slide. Beyond this point, visible actin motility is greatly reduced due to stress-induced disorganization of the actin cytoskeleton.

12. Videos are captured within a single z-plane; therefore, the initial z-slice placement is the focal plane for the entire video. Due to the 3D shape of a yeast cell, actin cables are visually more apparent either just above or just below the center of the cell.

13. Artifacts can occasionally appear in deconvolved images. These may include halo-like structures, amplification of noise pixels, or the disappearance of structures that are visible in the raw data. If artifacts appear, repeat the deconvolution with fewer iterations. However, if artifacts do not appear, but the image is still blurry or faint, repeat with more iterations. If deconvolution fails to exceed the 95% confidence limit, verify that optics are clean, light source is aligned, and cells are mounted under a #1.5 coverslip. Also make sure that images are acquired with pixel values at least in the middle of the detector range, and not saturating the detector.

14. Fiji/ImageJ velocity is defined as the distance covered between two frames and intensity of the selected pixel or volume. This definition is not a relevant measurement for determining RACF speed, thus manually calculating the velocity of actin cable movement is recommended.

Fig. 3. Tracking actin dynamics: retrograde actin cable flow.

Yeast cells expressing Lifeact-GFPEnvy were propagated in lactate medium and imaged every 500 ms for 30 s. Selected frames are shown. Green dots follow a fiduciary mark on an actin cable within a mother cell that undergoes retrograde movement. Scale bar = 1 μm.