Manipulation and Analysis of Cell Cycle-Dependent Processes in Budding Yeast

Summary

This protocol details two methods of yeast cell cycle arrest and optional release, and elaborates on the use of fluorescence microscopy to study cell cycle-dependent processes in S. cerevisiae.

Abstract

Eukaryotic cells follow a conserved cell cycle that regulates diverse processes, including DNA maintenance and organelle homeostasis. Studying cellular processes in a cell cycle-dependent manner is often necessary to properly interpret experimental results. There are chemical and genetic methods available to produce cell cycle synchronization in cultured cells across a wide swath of organisms, including vertebrate models, enabling the study of cell cycle-dependent processes. However, among model organisms, budding yeast remains a powerhouse for cell cycle analysis due to its particularly robust synchronization methods, short generation time, and genetic tractability. Yeast shares core cell cycle machinery with other eukaryotes, which has enabled landmark discoveries in cell cycle regulation. This protocol details methods for cell cycle analysis in yeast, focusing on G1 arrest-release and mitotic arrest-release experiments, including strain construction, culture preparation, and microscopy. PCR tagging methods for producing suitable strains for cell cycle arrests and fluorescence microscopy are presented. A G1 arrest is achieved using the peptide pheromone α-factor, and brief washes result in synchronous release and cell cycle progression. Samples are taken at different time points following release into the cell cycle and fixed for microscopy. A second method arrests yeast cells in mitosis by depleting the cell cycle regulator Cdc20 to achieve a metaphase-arrested population, as well as optional release into anaphase. Samples are fixed and prepared for imaging pre- and post-release, and are imaged and analyzed. Image analysis focuses on cataloging dynamic localization and population abundance changes of proteins in the cell cycle. These synchronization methods are suitable for diverse cell cycle manipulations, and while their use in imaging fixed cells is highlighted here, they can be adapted for many other analyses, including live cell imaging as well as biochemical and molecular assays.

Introduction

Eukaryotic cell division is highly regulated through a program called the cell cycle. The highly conserved and dynamic processes occurring in the cell cycle make it interesting to study in-and-of-itself, but also have wide-spread implications that inform investigations of other cell biological processes-for example, many organelles undergo dramatic remodeling during cell division, and the abundance and localization of many proteins is highly regulated throughout^1,2,3. Although there are some additional layers of complexity present in metazoan systems compared to yeast, including more variety of regulatory kinases and proteins, as well as additional input from external signaling in metazoans, cell division is overall very highly conserved in all eukaryotes^4,5. Much fundamental work establishing key insights into cell cycle progression and crucial regulatory steps was done in yeast, and it remains an extremely attractive organism for studying cell cycle-related questions for a multitude of reasons^6,7. Chief among these is the wide array of methods available to manipulate the cell cycle, allowing for arrest at various stages and synchronization of cells. Additional advantages include yeast's short cell cycle (~90-150 min depending on temperature and media conditions), the ease of genetic manipulation, and the large breadth of conditional alleles that enable study in an otherwise wild-type context.

Progression through the different phases of the cell cycle is driven by the activity of several key kinases, including the ubiquitous Cyclin-dependent kinase (Cdk), Polo-like kinase, and Aurora kinases. Cell cycle checkpoints regulate progression through the cell cycle, ensuring that cells do not progress into the next stage of division unless specific conditions are met^8,9. These checkpoints, and the transitions they regulate, can be hijacked by researchers to achieve long term arrest, or transient arrest and release to synchronize a population of cells (Figure 1). For example, the cell cycle commitment checkpoint (Restriction point or Start in yeast) ensures that the cell has enough nutrients to commit to division before starting DNA replication¹⁰. This checkpoint can be manipulated in haploid yeast to achieve a G1 arrest, taking advantage of the yeast mating program that interacts with the cell cycle. In baker's yeast (Saccharomyces cerevisiae), haploid cells have one of two mating types, MATa or MATα, which is genetically determined by a mating type locus. Yeast of one mating type are sensitive to the mating pheromone of the other mating type and respond by activating a MAP kinase signaling cascade that ultimately phosphorylates and stabilizes the Cdk inhibitor Far1, which prevents cell cycle entry by binding to the Cln1/2-Cdk complex and blocking its activity^11,12,13. Cells responding to mating pheromones can be visually determined by the presence of a mating projection or shmoo (Figure 1). Thus, chemically synthesized α-factor can be added to cultures of MATa cells to achieve a G1 arrest. Another cell cycle transition that researchers can target to synchronize yeast cells is the metaphase-to-anaphase transition. During mitosis, the Spindle Assembly Checkpoint (SAC) monitors the attachment of kinetochores to microtubules and inhibits the anaphase promoting complex/cyclosome (APC/C) by sequestering its activator, Cdc20, when unattached kinetochores are detected. The APC/C is an E3 ligase complex that is essential for anaphase entry. When the SAC is satisfied, Cdc20 then binds the APC/C, and APC/C ubiquitinates several proteins, primarily securin and cyclin B, which targets them for degradation, and promotes chromosome segregation and mitotic exit^14,15. Although Cdc20 itself is not a component of the SAC, its inhibition is the target of SAC activation; therefore, inducible depletion of Cdc20 can be leveraged to arrest a yeast culture in metaphase (Figure 1)¹⁶.

This protocol encompasses two methods of yeast cell cycle manipulation, as well as an example application of each method. In the first, α-factor is used to arrest cells in G1, followed by a wash step to release cells from the arrest, resulting in synchronous cell cycle re-entry. The synchronized cells will continue through the entire cell cycle, enabling researchers to track and study cells in each cell cycle stage. In this method, the dynamic localization of a GFP-tagged protein of interest, Stu2, is monitored throughout, and its association with the mitotic spindle is tracked. This method also leverages a tagged spindle pole body (SPB) component, Spc110-mCherry, which enables measurements of mitotic spindle length and is an excellent proxy for cell cycle progression¹⁷. This method demonstrates how cell cycle arrest and release enable synchronization of cell populations, facilitating the study of a protein or other component of interest throughout the cell cycle.

In the second method, a mitotic arrest is accomplished via depletion of Cdc20 using an auxin-inducible degron (AID) system¹⁸. In cells expressing TIR1 (an F-box protein of plant origin), any protein with an AID tag will be ubiquitinated and degraded upon the addition of auxin. Here, we use an AID-tagged Cdc20 to enable inducible degradation of Cdc20, thereby causing an arrest in metaphase. This cell population is then monitored for the recruitment of the tagged SAC component Bub1-GFP to kinetochores, using the tagged kinetochore protein Mtw1-mCherry as a fiducial marker. Although Bub1 plays SAC-independent roles at the kinetochores and localizes to kinetochores in early mitosis, its persistent kinetochore localization in arrested cells signals improper kinetochore-microtubule attachments¹⁹. We showed this by treating cells with the microtubule poison, nocodazole, which disrupts kinetochore-microtubule attachments. This experiment highlights how processes specific to a distinct cell cycle phase can be studied using highly tunable arrest methods. We further demonstrate that auxin can be washed out of cdc20-AID cultures, allowing release into anaphase and entry into a new cell cycle, though in our experience, these cultures do not release as synchronously as an α-factor arrest-release. These two arrest-release methods are both effective and easy to implement, but many other methods are available to researchers in yeast to arrest cells at different cell cycle stages^20,21,22,23. Further details of these methods can be found in the Discussion.

Protocol

1. Construction of strains for cell cycle analysis and imaging

1. Design primers to C-terminally tag the gene of interest using Pringle Tagging Plasmids (pFA6a plasmids)²⁴. In short, design F2 and R1 primers that, when used with a pFA6a-series plasmid, produce a PCR product that can be directly transformed into yeast to generate a 'tagged' version of the gene of interest. Use the primer pairs and plasmids contained in Table 1 to tag the genes of interest in this protocol. Plasmid and primer design strategies for C-terminal tagging of yeast genes are described in detail by Longtine et al.²⁴.
   NOTE: This method can be used to produce fluorescently tagged proteins for imaging, or Auxin Inducible Degron (AID) tagged proteins to facilitate cell cycle arrest and other experiments enabled by degradation of a target protein¹⁸. For example, to tag CDC20 with the AID (IAA7) tag, the forward tagging primer consists of 25-40 bases of the CDC20 gene immediately before the stop codon, excluding the STOP codon (5' TACAAGGAGGCCCTCTAGTAC
   CAGCCAATATTTGATCAGG 3'), and also the Pringle Plasmid F2 adapter sequence (5' cggatccccgggttaattaa 3') appended to the 3' end to produce the final primer sequence (5' TACAAGGAGGCCCTCTAGTAC
   CAGCCAATATTTGATCAGG cggatccccgggttaattaa 3'). The reverse CDC20 tagging primer contains the reverse complement of the 25-40 base pairs immediately after the STOP codon (5' ATTATATGCCTTGACATGAA
   CTTTTATTTTTTTTATTTTA 3') and the Pringle Plasmid R1 adapter sequence (5' gaattcgagctcgtttaaac 3') to produce the final primer sequence (5' ATTATATGCCTTGACATGAACTTTTATT
   TTTTTTATTTTA gaattcgagctcgtttaaac 3'). It is also important to note that using PCR primers shorter than 40 base pairs will reduce the efficiency of integration into the yeast genome.
2. Dilute primers in molecular biology grade water to 10 µM.
3. Perform PCR with the F2 and R1 plasmid pair and the appropriate tagging plasmid using standard methods, following the thermal cycler program contained in Table 2, then confirm the size of the PCR product by agarose gel electrophoresis.
4. Inoculate the yeast strain to transform into 25 mL of Yeast extract Peptone Adenine Dextrose (YPAD) culture and grow overnight at 23 °C. For constructing cdc20-AID yeast (that will degrade Cdc20 upon treatment with auxin), the transformed strain may either already contain OsTIR1 expressed from an ectopic locus, or OsTIR1 may be crossed in later¹⁸.
5. The following morning, check the OD600 of the overnight culture. If culture OD <2.0 proceed to step 1.6. If OD > 2.0, dilute to ~0.2-0.4 and allow for 2-3 cycles of growth.
6. Harvest cells at OD600 = ~0.4-2.0 to ensure they are in logarithmic growth phase. Harvest 5 ODs of cells per transformation (e.g., 5 mL of OD600 = 1.0). If doing more than one transformation of the same strain, cells can be combined at this stage. Harvest cells by centrifuging at 3,000 x g for 5 min at 23 °C.
7. Discard supernatant and wash the cell pellet with 25 mL of water, followed by centrifuging at 3,000 x g for 5 min at 23 °C.
8. Discard the supernatant and resuspend cells in 1 mL of 0.1 M Lithium Acetate (LiAc) and transfer to 1.6 mL tube(s). If doing more than one transformation, split the 1 mL resuspension into multiple tubes (e.g., for 2 transformations split 2x 500 µL; for 3 transformations split 3x 333 µL, etc.).
9. Centrifuge cells at 1,000 x g for 2 min at 23 °C and discard the supernatant.
10. Add the following to the cell pellet (in this order): i) 240 µL of PEG (50%), ii) 36 µL of 1.0 M LiAc, iii) 20 µL = 50 µg of Salmon Sperm DNA pre-boiled for 5 min, iv) 54 µL of water.
11. Add 10 µL of PCR product (obtained in step 1.3) to the cells and vortex for 1 min, ensuring everything is well mixed.
12. Incubate cells at 30 °C for 30 min in a roller drum in a warm room or a thermoshaker at 1000 rpm.
13. Heat shock cells at 42 °C for 15 min in a water bath.
    NOTE: The optimal heat shock time might differ depending on the yeast strain background. This protocol is optimized for the W303 strain background.
14. Centrifuge cells at 1,000 x g for 2 min at 23 °C. Remove supernatant and gently resuspend in 100 µL of water.
15. For transformations using KanMX or other drug selection, plate cells on YPAD and incubate the plates at 23 °C, then replica plate to drug plate the following day.
    NOTE: Allowing a day of growth on YPAD is critical to allow for expression of drug resistance genes in transformed cells.
16. For transformations using auxotrophic markers (URA, HIS, etc.), plate cells directly on selective medium.
17. Incubate plates at 23 °C for 3 days to allow the yeast transformant colonies to grow.
18. Verify the correct tagging of the gene by genomic PCR or by western blotting. These methods are described in detail by Burnette²⁵ and Chu et al²⁶.
    NOTE: For producing strains with multiple tagged alleles, repeated transformation is not recommended without back-crossing, as transformation can be a mutagenic process. To produce the strain containing Stu2-GFP and Spc110-mCherry, for instance, we recommend transforming strains separately to produce Stu2-GFP and Spc110-mCherry, then combining these alleles into one haploid strain by yeast cross, sporulation, tetrad dissection, and genotypic analysis. More information on yeast crosses is available from Stansfield and Stark²⁷ and Morin et al.²⁸. To perform an α-factor arrest-release, the haploid strain utilized must be MATa.

2. Yeast culture and cell cycle synchronization: α-factor arrest-release

1. Inoculate yeast into 25 mL of YPAD culture overnight at 23 °C, to be at OD600 = 0.5-2.0 the next morning.
   1. If cells are above OD600 = 2.0 in the morning, dilute back to OD600 = 0.2-0.4 and let them grow for at least 1 cell cycle (2-3 h at 23 °C).
2. Dilute cells back to OD600 = 0.5 and add α-factor to a final concentration of 1 µg/mL (stock 10 mg/mL in DMSO). Cells ideally would have a bar1 mutation (e.g. bar1-1), which renders them more sensitive to α-factor. BAR1 wild type cells are less sensitive to α-factor and require a concentration of α-factor that is at least an order of magnitude higher to arrest properly, which is less economical.
3. At 2.5-3.5 h post-arrest, check the degree of arrest by counting the percentage of non-budded shmooed cells. When 90-95% of cells are shmooed (arrested in G1), proceed to release. Timing will differ depending on the yeast strain, growth medium, and temperature.
4. Release cells from α-factor arrest by spinning down in centrifuge at 3,000 x g for 3-5 min at 23 °C, and then pouring off supernatant.
5. Wash cells by resuspending the pellet in 25 mL of YPAD + 1% DMSO.
6. Repeat steps 2.4-2.5 twice for a total of three washes, changing the used tube once after the first wash.
7. Add YPAD to cells to bring the final volume to 25 mL and transfer cells into a new flask.
   NOTE: Using a new flask is critical to eliminate trace amounts of α-factor left that might prevent the release.
8. Take time point (0 min) sample and fix (see fixation instructions in steps 4.1 to 4.4).
9. Continue taking time point samples every 15 min if growth takes place at 23 °C. This timing will change at other temperatures based on how fast the cells grow.
10. At time point 60 min, judge the synchrony of the release by checking for uniform small buds (bud size < 25% mother cell size) under a light microscope.
11. Optional) At 60 min post-release, add α-factor again to a final concentration of 1 µg/mL to prevent progression into the next cell cycle. Confirm synchronous cell release by verifying uniform small-budded morphology before addition.
12. Continue taking time point samples up to 180 min, or as long as desired. At 23 °C, metaphase occurs ~45-60 min post-release, anaphase occurs ~60-90 min post-release, and most cells will be in the new G1 by 120 min post-release.

3. Yeast culture and cell cycle synchronization: Cdc20 depletion arrest-release

1. Inoculate an overnight culture of yeast at 23 °C to be at an OD600 = 0.5-2.0 the next morning.
   1. If cells are above OD600 = 2.0 the next morning, dilute back to OD600 = 0.2-0.4 and let them grow for at least 1 cell cycle (2-3 h at 23 °C).
2. Dilute back to OD600 = 0.5 and add auxin to a final concentration of 500 µM from 1 M stock (500x) to all cultures. For cultures receiving other drugs (nocodazole), add them concurrently with auxin at the appropriate concentration (e.g., ~10 µg/mL nocodazole).
3. At 2 h post-arrest, check the degree of arrest by counting the percentage of large-budded cells (large-budded is determined as the presence of bud > 25% mother cell size). When 90-95% of cells are arrested in mitosis, proceed to either collecting samples or releasing cells from arrest, depending on the experimental application. If 90% of cells are not arrested, allow the culture to continue growing, and check it every 15-30 min. Timing will differ depending on the yeast strain, growth medium, and temperature.
   NOTE: It is not advisable to allow arrests to proceed for too long (greater than 4 h) if releasing, as prolonged arrests may reduce cell viability.
4. To release from Cdc20 depletion arrest, harvest culture by centrifugation at 3000 x g for 3-5 min at 23 °C.
5. Wash cells by resuspending the pellet in 25 mL of YPAD + 1% DMSO, then harvesting by centrifugation at 3,000 x g for 3-5 min at 23 °C.
6. Repeat steps 3.4-3.5 for a total of 2 washes.
7. Add YPAD to cells to bring the final volume to 25 mL and transfer the cells to a new flask.
   NOTE: Using a new flask is critical to remove trace amounts of auxin that might prevent a release.
8. Take time point 0 and fix (see fixation instructions in steps 4.1 to 4.4).
9. Continue taking timepoints every 10-15 min for the desired length of the time course. At 23 °C, most cells would have released from the arrest after 50-70 min.
   NOTE: Cells do not release as synchronously from this Cdc20 depletion arrest as they do with α-factor arrest. However, it can be useful to study anaphase or mitotic exit events. Additionally, a technical limitation of this arrest-release is that in later time points after the release, it is more difficult to differentiate between metaphase cells that have not yet released and the cells that released earlier and have reached the metaphase of their next cell cycle. If needed, this can be overcome by either doing live cell imaging to track individual cells as they release from the arrest or by adding α-factor to the culture (assuming the cells are MATa) to arrest cells in G1 after mitotic exit.

4. Yeast fixation

1. Centrifuge 1 mL (or 0.5-1.5 ODs) of culture for 1 min at max speed (~20,000 x g) at 23 °C.
2. Aspirate off supernatant and resuspend cells in 500 µL of fixative solution (3.7% formaldehyde in 0.1 M potassium phosphate (KPi) pH 6.4).
3. Incubate cells in the fixative solution for 2-15 min at
   23 °C. For time course experiments, it is generally advisable to fix cells for 15 min to align the timing of centrifuging cells when taking samples.
   NOTE: Fixation timing should be optimized for specific fluorophores and proteins of interest.
4. Centrifuge cells for 1 min at max speed at 23 °C, aspirate off supernatant, and resuspend in 500 µL of 0.1 M KPi (pH 6.4). Cells in KPi can be stored at 4 °C until ready to image.
   NOTE: Fixed cell fluorescence signal can last for up to 2 weeks, depending on the fluorophore, however, for quantification of fluorescent signals, collect images within 7 days.

5. Preparing slides for imaging

1. When prepared to image, centrifuge cells for 1 min at max speed at 23 °C and discard supernatant. Resuspend cells in 10-100 µL of Triton/DAPI/sorbitol solution (1.2 M sorbitol, 1% triton, 0.1 M KPi, pH 7.5, 2 µg/mL DAPI).
2. Pipette ~0.8 µL of cells directly onto a clean coverslip. Use a pipette tip to gently spread the cell droplet into an approximately 1 cm² circle in the center of the coverslip.
3. Place the coverslip onto a microscopy slide. Use a Kimwipe to gently press around the slide to distribute the sample.
4. Seal the edges of the coverslip with nail polish.
5. Optionally store in a cool dark location for up to 4-6 h before imaging to help cells settle and prevent excess motion.

6. Imaging

1. Take slide to a microscope equipped with a 60x/1.42 oil objective and a Red, Green, Blue, Far Red laser and filter set. Higher magnification objectives (up to 100x) can also be used.
2. Focus the microscope on the yeast cells adhered to the coverslip.
   NOTE: If yeast cells are moving too much, slides may be left in a cool dark location for 20-60 min to settle.
3. Once the yeast are in focus, tune exposure settings of each channel to produce a signal to noise ratio of at least 3:1, ensuring no fluorescent puncta are over-exposed.
   NOTE: For example, on an imaging system we use with a scientific CMOS camera, using 60x 550 objective and immersion oil with a refractive index of n = 1.516, setting max signal of the red channel to 1000-2000 au, the green channel to 3000 au, the blue channel to 3000 au and the DIC channel to 4000-5000 au produces high quality images.
4. Adjust the acquisition settings to take 14-20 0.2 µm Z-stacks to span 2-4 µm total.
5. For microscopes equipped with post-processing modules, select Deconvolution and Quick Projection for each stack.
6. Collect Z-stacks of cells, taking enough images to capture ~100-200 cells for each condition or time point.
7. Refer to the Supplemental File 1 for software instructions with screenshots from the imaging software.

7. Image analysis

1. Open projected images in ImageJ and adjust brightness and contrast to view different channels. If the microscope used for imaging does not have post-processing modules, perform max projection of the Z-stacks at this stage.
2. To determine cell cycle progression, count 100 cells per time point and classify them as containing one or two nuclei. Alternatively, calculate the spindle length or distance between spindle pole bodies to define cell cycle state. The distance between Spc110 foci < 1.5 µm corresponds with metaphase cells, and the distance > 1.5 µm corresponds with anaphase cells.
3. To calculate intensities of protein puncta (e.g., Stu2-GFP), use the freehand selection tool to draw around the boundary of the signal of interest. Add this signal to ROI (Region of Interest) manager by pressing T or using the Menu. In ROI manager select Measure to calculate the intensity of each selected puncta. To view changes in puncta signal over time, plot average intensity with 95% confidence interval on a line graph for each time point.
4. To calculate the percentage of cells with Bub1-GFP puncta, ensure the green channel minimum and maximum brightness settings are the same across different images. Cells with a Bub1-GFP puncta showing overlap with an Mtw1-mCherry puncta are counted as Bub1 localized to the kinetochore; otherwise, the cell is counted as negative. Count ~100 cells per condition. Aravamudhan et al.²⁹ contains more information on Bub1 localization to kinetochores.
5. To calculate the percentage of large-budded mononuclear cells, count the number of cells with a large bud and one nucleus (one DAPI signal) in the population and divide by the total number of cells.
6. Refer to the Supplemental File 1 for the software instructions with screenshots from the imaging software.

Results

Analyzing changes in cell cycle-dependent protein localization by fluorescence microscopy can be readily accomplished using the methods we describe here. Our group has long been interested in the dynamic regulation and function of the mitotic spindle. In yeast, spindle pole bodies (marked by component Spc110) function as microtubule organizing centers from which microtubule filaments emanate to create the structure of the mitotic spindle³⁰. The microtubule binding ...

Discussion

Utilizing cell cycle synchronization in budding yeast enables studying important mechanisms for a variety of cellular processes. The use of G1 arrest-releases with α-factor treatment allows synchronous progression of a population of cells through the stages of the cell cycle, and as we showed, can reveal dynamic localization patterns of cellular regulators like Stu2³⁶. Cell cycle arrests can also be accomplished using genetic means via depletion of cell cycle regulators like Cdc20, w...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
?-factor	University of Utah Core Synthesis Facility		Sequence: WHWLQLKPGQPMY
1.5 mL Eppendorf Tubes	Axygen	MCT-175-C
10mM dNTP mix	Thermo Scientific	R0193
50 mL conical tubes	greiner bio-one	227 261
5x Phusion HF reaction buffer	New England BioLabs	B0518S
Acetic Acid, Glacial	Fisher Chemical	BP2401C-212
adenine hemisulfate salt	Sigma-Aldrich	A9126-100G
Agar, Granulated	Apex Chemicals and Reagents	20-275
agarose	Apex Bioresearch Products	20-102GP
Autoclave Amsco Century Steam Sterilizer	Steris	SV-1262
autoclaved DI water
Auxin	Sigma-Aldrich	Cat#I3750-5G-A; CAS: 87-51-4
Cargille Laser Liquid	Cargille Laboratories	20130
D-Sorbitol	Sigma-Aldrich	S1876-500G
DAPI (40 ,6-Diamidino-2-Phenylindole, Dihydrochloride)	Molecular Probes	Cat#D1306
Deoxyribonucleic acid sodium salt from salmon testes	Sigma-Aldrich	D1626
Dextrose	Fisher Chemical	D16-10
Disodium Ethylenediamine Tetraacetate	Fisher Chemical	S811-10
DMSO	Thermo Scientific	20688
FIJI/ImageJ2 vs 2.14.0/1.54f	ImageJ2	https://imagej.net/software/fiji/
Fixed Speed Vortex Mixer	VWR	https://dabos.com/product/vortex-mixers-vwr-fixed-speed-vortex-mixer-00001-24763?srsltid=AfmBOoo5TH0aoExvrrrphDaFt8XAsDqLvkjxtEUj1QWlFbWh7_gwzMObLT4&gQT=2
Fluorescent microscope DV Ultra	Leica	https://www.leica-microsystems.com/c/am/lsr-w/fluorescence-microscope-wf/?nlc=20250214-SFDC-022570&utm_source=google&utm_medium=cpc&utm_campaign=25-AM-LSR-L3-LSPO-LSWF-SE-Google-Ads-WF-Thunder-Search&utm_content=text_ad&utm_term=fluorescence%20microscopes&gad_source=1&gad_campaignid=170130111&gbraid=0AAAAADrbsAF-dGDbxzgT8m_cvXSlf4BB0&gclid=CjwKCAjwmenCBhA4EiwAtVjzmkMJUGFksaHezZvlBUlbbS1tR8RqXP24dbSRzcRgTT8RmJy7nyeThBoC3yQQAvD_BwE	Serial #: NV01063. No longer supported
Formaldehyde	Fisher Chemical	Cat#F79-500
gel apparatus	Thermo Scientific	Owl EasyCast B1
GeneRuler DNA Ladder Mix	Fermentas	SM0333
glass beads	Fisher Scientific	11312A
Glass Slides	VWR	48300-026
Innova 2300 Platform Shaker	New Brunswick	NB-2300
Kimwipes	Kimtech	06-666
Laboratory centrifuge for 1.5 mL tubes	Eppendorf	2525
Laboratory centrifuge for 50 mL tubes	Eppendorf	5804
Lithium acetate dihydrate	Sigma-Aldrich	L4158-250G
Master cycler nexus X2	eppendorf	https://www.eppendorf.com/us-en/Products/PCR/Thermocyclers/Mastercycler-nexus-X2-p-PF-82586
Micro-pipettes p2, p20, p200 and p1000 and corresponding tips	Rainin	L-2XLS+R, L-20XLS-R, L-200XLS-R, L-1000XLS-R
Microscope Cover Glass	Fisher Scientific	12541014
Nocodazole	Calbiochem	Cat#487928; CAS: 31430-18-9; Lot#B35705
Orange G	Sigma-Aldrich	O7252
PEG	Hampton Research	HR2-591
Peptone granulated	Fisher Bioreagents	BP9725-5
Phusion HF DNA Polymerase	New England BioLabs	M0530L
Pipet-X	Rainin	PX-100R
Potassium phosphate, dibasic	Thermo Scientific	424195000
Potassium phosphate, monobasic	Thermo Scientific	424200025
power source	Bio-Rad	23786
Start Acquire Ultra 1.2.2	softWoRx Cytiva	Obtain with DV Ultra
Tris Base	Fisher Bioreagents	BP152-10
Triton X-100	Sigma-Aldrich	9002-93-1
tube rotator	VWR	10136-084
water bath	VWR	WBE10A11B
Water, Ultra Pure	Apex Bioresearch Products	18-194
Yeast extract Granulated	Fisher Bioreagents	BP9727-5