Mechanism and Regulation of Eukaryotic Cell Division · Cell division takes place in timely and...

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Mechanism and Regulation of

Eukaryotic Cell Division

January 2011

Zentrum für Molekulare Biologie (ZMBH)

Elmar Schiebel 1, Gislene Pereira 2, Oliver Gruss 1

1 ZMBH 2 DKFZ

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Table of Contents

Introduction ..............................................................................................................................3

The cell cycle .............................................................................................................................3

Model Systems ...........................................................................................................................5

Experiments and Protocols......................................................................................................6

Measuring the loss of chromosomes ..........................................................................................7

The use of GFP labelled proteins in cell cycle analysis .............................................................8

Background ................................................................................................................................8

Construction of GFP labelled proteins .....................................................................................10

Cassette PCR 11 Yeast transformation 12

Analysis of conditional lethal cell cycle mutants.....................................................................15

Background ..............................................................................................................................15

Fluctuation of protein levels throughout the cell cycle ............................................................18

Background ..............................................................................................................................18

Synchronisation of cells with alpha-factor...............................................................................22

The metaphase-anaphase transition..........................................................................................26

Background: The function of cohesin and separase in sister chromatid separation – according

to K. Nasmyth ..........................................................................................................................26

Induction of anaphase by cohesin cleavage .............................................................................29

Working with mammalian cells ...............................................................................................31

Background ..............................................................................................................................31

Western blotting .......................................................................................................................32

Immunofluorescence staining of cells ......................................................................................33

Purification of the anaphase promoting complex 40

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Introduction The cell cycle

We know for more than 100 years that life in general is based on cellular organisation.

This implies that the only way to make new cells is by division of already existing

ones. This very simple statement has important consequences. Reproduction, growth,

development as well as maintenance of a multi-cellular organism depend on the

ability of its cells to divide properly. Conditions, which interfere with faithful cell

division, like high doses of cytostatic drugs or ionizing radiation lead to death after a

very short exposure time.

In order to maintain the information for following divisions in both nascent

daughter cells after cell division, they need to inherit the information for their

structure and organisation. This information is encoded on the genomic DNA in

chromosomes. Duplication and faithful segregation of chromosomes to both

daughters are one of the central functions of cell division.

Cell division takes place in timely and spatially well defined series of

processes. For example, replication always has to precede chromosome segregation.

The time window, in which cells replicate their chromosomes is called S-Phase, while

the actual division of the chromosomes (Mitosis) and the division of the cytoplasm

(cytokinesis) is called M-Phase. Two Gap-Phases (G1 and G2) follow in between the

two central phases (see Fig. 1). They allow cells to grow and control entry into the

cell cycle (G1) or check the integrity of the DNA (DNA damage checkpoints in G1

and G2).

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Fig. 1: The eukaryotic cell cycle. The main function of cell division is the

duplication of the chromatin and its segregation to the two daughter cells.

Accordingly, the cell cycle in eukaryotes is divided into different phases: S-

Phase, M-Phase and the Gap-Phases (G1 and G2).

A complex network of regulatory proteins governs all steps of the cell cycle. In the

centre of this cell cycle control system are a family of heterodimeric kinases, the so-

called cyclin dependent kinases. The activation of cyclin-dependent kinase 1 (Cdk1)

by binding of mitotic cyclins for example starts M-Phase in higher eukaryotes. Active

Cdk1 directly phosphorylates target proteins, which in turn mediate the structural

rearrangements at the beginning of M-Phase, like e.g. chromosome condensation,

nuclear envelope breakdown and spindle assembly (Fig. 2).

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Fig. 2: A family of related Cyclin dependent kinases (Cdks) control specific

cell division processes. The activity of the kinases is regulated by the synthesis

and rapid degradation of their activating subunit, the cyclins.

Model Systems

Cell division is highly conserved among eukaryotes. Consequently, data from very

different model organisms have contributed to today’s knowledge of the eukaryotic

cell division cycle. The main model organisms of cell cycle control are:

Yeasts

Oocytes and Eggs of marine Invertebrates and Amphibia

Vertebrate tissue culture cells

The most important model organisms for the genetic analysis of cell division and cell

cycle control are the yeasts Schizosaccharomyces pombe (fission yeast) and

Saccharomyces cerevisiae (budding yeast).

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Experiments and Protocols: cell cycle analysis of yeast and mammalian

cells

Schedule

Over the next two weeks we will analysis conserved principles of cell cycle regulation

using the unicellular budding yeast Saccharomyces cerevisiae and human HeLa cells

as model organisms. We will use the powerful genetic potential of budding yeast to

study cell cycle mutants and analyse cell cycle checkpoints. We will also analyse cell

cycle progression of human HeLa cells in order to demonstrate that principals in cell

cycle regulation are conserved.

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Starts at day 1 (Monday). Measuring the loss of chromosomes – Linda/Sarah

A colony colour assay that measures chromosome stability is described in the

literature and can be used to study parameters that affect the mitotic maintenance of

yeast chromosomes, including ARS function, CEN function, spindle function and

chromosome size. A cloned ochre-suppressing form of a tRNA gene, SUP11, serves

as a marker on an in vitro-constructed chromosome [CF(CEN6 TRP1+ SUP11

CYHS2]). Diploid strains that are homozygous for an ochre mutation in ADE2 (ade2-

101) from red colonies. ade2-101/ade2-101 cells with one copy of the SUP11 gene

grow with pink color, and those carrying two or more copies of SUP11 from white

colonies. Loss of the [CF(CEN6 TRP1+ SUP11 CYHS2] chromosome in ade2-101

haploid cells causes a change of colony colour from white to red. This causes

sectoring of colonies. The degree of red sectors reflects the frequency of mitotic

chromosome loss.

- Haploid yeast strain (1513) YRN212 (MATalpha ura3-52 lys2-801 ade2-101 trp1∆1

cyhR [CF(CEN6 TRP1+ SUP11 CYHS2]).

- Haploid tub4-1 mutant (1586) YAS4 (YRN212 tub4-1).

Grow on SC-Trp plates at 23ºC.

ADE2: Phosphoribosylaminoimidazole carboxylase. It catalyzes a step in the 'de novo'

purine nucleotide biosynthetic pathway. A red pigment accumulates in mutant cells

deprived of adenine.

a. Chromosome loss assay

1. At day -1 (Sunday; done by Linda or Uschi): Cells of YRN212 and YAS4

are inoculated in SC-Trp medium at 23ºC.

2. At day 1 (Monday) inoculate 3x10 ml of YPD medium with 106 cells/ml

(OD600 of 0.1). Grow cells at 23ºC, 30ºC and 33ºC for 6 h. For each condition plate

out 100, 300 and 600 colonies on YPD plates. Incubate cells for 6 days at 23ºC. You

may want to incubate the plates for 1-3 days in the cold room. This enhances the

appearance of the red colour.

3. Count colonies according to appearance: i) red colonies, ii) white colonies,

iii) white colonies with red sectors. Give cells types in % of total. What are your

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conclusions? Does the tub4-1 mutation cause chromosome loss? How do you explain

chromosome loss in case of Tub4 malfunction?

Day 1 (Monday): The use of GFP labelled proteins in cell cycle analysis:

Linda/Sarah

Background

Green Fluorescent Protein (GFP)

Fig. 3: Structure of Green Fluorescent Protein (GFP)

Structure of the green fluorescent protein (GFP): The figure shows the overall

shape of the GFP protein. Eleven strands of sheets (green) form the walls of a

cylinder. Short segments of helices (blue) cap the top and bottom of the 'b-can' and

also provide a scaffold for the fluorophore, which is near the geometric centre of the

can.

GFP (Green Fluorescent Protein) is a protein produced by a jellyfish Aequorea

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victoria, which fluoresces in the lower green portion of the visible spectrum. The gene

for GFP has been isolated and has become a useful tool for making expressed proteins

fluorescent by creating chimeric genes composed of GFP or its different color variants

linked to genes of interest. The result is an in vivo fluorescent protein, which can be

followed in a living system.

Fig. 4: Structure of the GFP fluorophore

One initial problem with the use of GFP was the excitation and emission spectra of

the wild type protein. Wild type GFP has two excitation peaks: a major one at 395 nm

(in the long UV range) and a smaller one at 475 nm (blue) and an emission peak at

509 nm (green). For wild type GFP, exciting the protein at 395 nm causes fairly rapid

quenching of the fluorescence. Also most investigators use FITC filter sets to observe

GFP staining. To alleviate this problem, several mutants of the GFP gene were

constructed which have increased fluorescence, but perhaps more important, the

major excitation peak has been red-shifted to 490 nm with the emission staying at 509

nm. The mutant GFPs are better suited in combination with the FITC filter sets as

they have the same excitation range as FITC. Furthermore, the main laser line used

for FITC excitation is from the argon laser at 488 nm. There is no good commonly

used laser line near 395 nm in most confocal microscopes. One of the mutant GFPs

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which has a 5-6 times enhanced fluorescence has a serine to threonine substitution at

position 65 in the protein (S65T: See Heim et al., Nature, 373: 663-664 (1995)). Since

then, other mutations have been described that give further improvements in the

brightness of the emitted light (See Clontech's website).

For general fluorescence microscopy, investigators have been using normal

FITC filter sets for viewing GFP: Ex 485/20, Dichroic FT 510, Em 520-560. (485/20:

485 is the middle of the peak, /20 refers to the width of the band, 10 nm either side;

FT 510 blocks all light below 510 nm and allows anything above 510 nm to pass; the

emission filter Em allows light between 520-560 nm to pass).

In recent years, several different color forms of GFP have been produced

through mutagenesis and selection. These are in order from the shortest to the longest

emission spectra: blue, cyan, green, and yellow: FP or BFP, CFP, GFP and YFP.

GFP variants allow constructing double labeled specimens expressing two

fluorescently labeled proteins.

Construction of GFP labelled proteins

In yeast GFP labelled proteins are constructed by transforming yeast cells with PCR

cassettes containing GFP, a selection marker, and 45 bp flanking sequences that direct

the cassette through homologues recombination to the target site. In some cases, the

GFP gene fusion is non-functional like in the case of tubulin-GFP. However, tubulin-

GFP expressed “in trans” to the tubulin gene becomes incorporated into microtubules

and can therefore be used for the labelling of microtubules.

Today and tomorrow we will transform yeast cells with an integration plasmid

expressing TUB1-GFP-URA3, and cassettes for SPC42-GFP-His3MX6 (a marker for

the yeast spindle pole body (SPB, the yeast centrosome), and the cell cycle regulator

CDC14-GFP-His3MX6. The yeast transformants will be purified and then analysed

for the presence of the GFP-tagged protein. Finally, we will analyse the phenotype of

the yeast strains when incubated at restrictive conditions.

a. Prepare SPC42-GFP-His3MX6 and CDC14-GFP-His3MX6 by PCR

1. Day 1 (Monday): Set up PCR using plasmid pSM571-1 as template and

primers SPC42-S2/SPC42-S3 or primers CDC14-S2/CDC14-S3. In addition, digest

plasmid TUB1-GFP-URA3 with the restriction enzyme Stu1.

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PCR for Tag-cassettes

Stocks: Template DNA pSM571: 20 ng/ml Primers: 10 µM (Use S2/S3-primers) dNTPs mix (2 mM of each, ATP, TTP, GTP, CTP)

Cassette PCR with Phusion polymerase Condition for Buffer HF:

Buffer HF 5 µl

2 mM dNTPs 2.5 µl

10 µM Primer S2 1.25 µl

10 µM Primer S3 1.25 µl

Template pSM571 (60 ng/µl) 1 µl

Phusion (2 U/µl) 0.25 µl (add last)

Water 13.75 µl

Total 25 µl

Cycle conditions: 1. 30 sec at 98°C 2. 5 sec. at 98°C 3. 15 sec at 63°C 4. 50 sec at 72°C 5. Go to step 2 for 9 times 6. 5 sec at 98°C 7. 15 sec at 63°C 8. 50 sec at 72°C, increment of 8 sec per step 9. Go to step 6 for19 times 8. pause, 4°C X sec = is calculated based on length of expected product. Use 20 sec per 1 kb. Analyze 1 µl on 0.8% agarose gel (1 µl reaction+ 2 µl 6xSB + 9 µl water). Both PCR

products have 2313 bp. Why do they have identical sizes?

Store PCR reactions at -20°C.

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Restriction of plasmid TUB1-GFP-URA3 (6700 bp) with Stu1

TUB1-GFP-URA3 plasmid (1 µg/µl) 2 µl 10x Buffer NEB 4 2 µl water 16 µl StuI restriction enzyme 0.5 µl Mix and spin. Incubate for 1 h at 37ºC. Analyze 1 µl of the digestion on 0.8% agarose gel (1 µl reaction+ 2 µl 6xSB + 9 µl water). One restriction site – thus, how many fragments do you expect to see on the agarose gel? Size? b. Transformation of yeast cells with SPC42-GFP-His3MX6 and CDC14-GFP-His3MX6 cassette and TUB1-GFP-URA3 integration plasmid 1. Day 1 – Monday: inoculate 10 ml YPD or 10 ml YPG/R (ESM1105-1)

medium with following yeast strains:

- 2298: GPY511-1 (YPH499 cdc15-1 ∆ase1::KanMX6).

- 229: ESM1105-1 (ESM356-1 Gal1-CDC20-kanMX6) - YPG/R.

- 909: ESM1278-1 (YPH499 cdc15-1).

- 4809: YPH499

- 2978: ESM356-1

Grow cells at 23ºC over night.

2. Day 2 - Tuesday: Dilute 50 ml YPD or YPG/R (ESM1105-1) medium with

over night culture to OD600 of 0.2. Incubate for 3-4 hours at 23ºC.

Yeast Transformation

1. Transfer cells into a 50 ml sterile Falcon tube. Harvest cells by centrifugation

(3.000 g, 5 min, 20°C).

2. Discard supernatant. Wash cell pellet once with 25 ml sterile distilled water,

followed by 20 ml LiSorb.

3. Centrifuge for 1 min, 3.000 g, 20°C. Remove residual liquid.

4. Add 300 µl LiSorb, 50 µl carrier DNA (sheared salmon sperm DNA

(GIBCOBRL), heated for 5 min at 95°C and then cooled to 4°C) to the cell pellet.

Resuspend cells with a vortex mixer.

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Yeast cells were transformed as follows:

1. Mix 50 µl cells with DNA as outlined below. Incubate for 10 min at room

temperature.

Add Temp.

- GPY511-1 5 µl SPC42-GFP-His3MX6 23ºC

4 µl TUB1-GFP-URA3/Stu1 23ºC

5 µl CDC14-GFP-His3MX6 23ºC

- ESM1278-1 5 µl CDC14-GFP-His3MX6 23ºC


5 µl SPC42-GFP-His3MX6 23ºC

- YPH499 5 µl CDC14-GFP-His3MX6 30ºC

5 µl SPC42-GFP-His3MX6 30ºC


- ESM1105-1 4 µl TUB1-GFP-URA3/Stu1 30ºC

- ESM356-1 4 µl TUB1-GFP-URA3/Stu1 30ºC

Do not forget the “no DNA” control.

2. Add 300 µl LiPEG and mix using a vortex mixer. Incubate for 10 min at

room temperature. Then add 30 µl dimethyl sulfoxide (DMSO), mix and incubate for

10 min at 42˚C.

3. Harvest cells by centrifugation (3,000 g, 20°C, 3 min) and carefully remove

the supernatant by aspiration.

4. Resuspend cells in 300 µl sterile PBS. Mix using a vortex mixer. Plate cells

onto selective plates (SC-His or SC-Ura) with glucose or galactose/raffinose

(ESM1105-1, SC-Ura G/R).

5. Incubate plates for 3 days (until Thursday) at the indicated temperature (see

above).

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6. Purify transformants on Thursday by streaking them out for single colonies

on selective plates (six transformants per plate). Incubate until Saturday at 23ºC or

30ºC until single colonies come up. On Saturday/ prepare from a single colony a

patch on a selective plate. Incubate at the indicated temperature.

7. On Friday check transformants by fluorescence microscopy for the presence

of a GFP signal and the cellular distribution of the GFP-labelled protein.

b. Analysis of yeast strains (Day 6/7 – Monday and Tuesday) – for the group who

will do advanced imaging together with Sarah and Johanna: discuss the

experiment as soon as possible with them. You have an influence of the

experiments you want to do: possibilities are standard phenotype analysis

(described below), live cell imaging and FRAP.

1. Day 6: Take mutant and wild type cells and grow them over night in 5 ml

YPAD (23ºC, GPY511, ESM278, YPH499) or 5 ml YPAR/G (30ºC, ESM356-1,

ESM1105) medium at the indicated temperatures.

2. Dilute “YPAD” cells to OD600 of 0.2 with YPAD and grow at 23ºC for 2-4

h. Then shift cells to 37ºC for 3 hours (GPY511-1, ESM1278, GPY515 and YPH499).

For ESM1105 and ESM356-1, pellet cells and resuspend them in YPAD

medium at 30ºC for 3 hours.

3. Fix cells with formaldehyde (see c.).

c. Fixing GFP cells with formaldehyde and analysis of the phenotypes by

fluorescence microscopy

1. Take 1 ml of cell culture, 100 µl 1 M KPO4 pH 6.5, 100 µl formaldehyde

solution. Incubate for 10 min at room temperature.

2. Wash cells 2x with 10 ml PBS (by spinning at 3.000 rpm for 2 min in the

Eppendorf centrifuge).

3. Incubate cells for 30 min at room temperature in 100 µl DAPI (PBS + 1

µg/ml DAPI).

4. Wash cells with PBS (as before) until DAPI background is low (1-3 x).

Resuspend cells in a suitable volume PBS. Add 4 µl onto cover slip. Cover sample

with cover slip. Inspect cells by fluorescence microscopy.

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5. Describe phenotypes of cells.

Day 2 (Tuesday): Analysis of conditional lethal cell cycle mutants

Background

Conditional lethal mutants normally contain one or multiple point mutations in one

specific gene. They grow at the permissive temperature (frequently 23ºC) but fail to

grow at the restrictive temperature (for example 37ºC). In a series of ground braking

papers L Hartwell published the first cell division cycle mutants (cdc mutants). These

are conditional lethal mutants of which all cells of a culture arrest at the same phase in

the cell cycle when incubated at the restrictive temperature. According to the

appearance in the screen these mutants were named CDC1, …, CDC28, …, CDC31,

and so on. Similar screens were later performed in fission yeast and other organisms.

Because the nature of the mutated gene was first unclear, the cyclin dependent kinase

(CDK1) gene in budding yeast was named CDC28, while in fission yeast this gene

was named CDC2.

Hartwell LH, Culotti J, Reid B. Genetic control of the cell-division cycle in yeast. I. Detection of mutants. Proc Natl Acad Sci U S A. 1970 Jun;66(2):352-9. Hartwell LH. Genetic control of the cell division cycle in yeast. II. Genes controlling DNA replication and its initiation. J Mol Biol. 1971 Jul 14;59(1):183-94. Culotti J, Hartwell LH. Genetic control of the cell division cycle in yeast. 3. Seven genes controlling nuclear division. Exp Cell Res. 1971 Aug;67(2):389-401. Hartwell LH. Genetic control of the cell division cycle in yeast. IV. Genes controlling bud emergence and cytokinesis. Exp Cell Res. 1971 Dec;69(2):265-76.

Analysis of cdc mutants

Today, we will perform the Hartwell experiment.

Wild type cells:

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775, K699: MATa ade2-1 trp1-1 can1-100 leu2-3,112 his3-11,15 ura3. W303 background. 19126 YPH499: MATa ura3-52 lys2-801 ade2-101 trp1∆63 his3∆200 leu2∆1. S228c background.

Temperature sensitive cells:

K699 background 2152, K4386: MATa cdc23-1 ade2-1 trp1-1 can1-100 leu2-3,112 his3-11,15. 1089, K1993: MATa cdc15-2 ura3 leu2-3,112. 1099, K3451: MATa cdc12 ura3 can1-100 trp1-1 leu2-3,112. 829, SLJ250: MATa ∆bar1::HisG cdc14-1 ura3-1 leu2-3,112 trp1-1 his3-11,15 ade2-1 can1-100. 831, SLJ200: MATa ∆bar1::HisG tem1-3 ura3-1 leu2-3,112 trp1-1 his3-11,15 ade2-1 can1-100. 849, SLJ139: MATa cdc16 ∆bar1::hisG ura3-1 leu2-3,112 trp1-1 his3-11,15 ade2-1 can1-100. 4236, AY51: MATa spc42-10 ade2-1 his3-11,-15 ura3. 27653, US1197: MATa cdc4-13 ade2-1 trp1-1 can1-100 leu2-3,112 his3-11,15 ura3. 27652, US1198: MATa cdc4-12 ade2-1 trp1-1 can1-100 leu2-3,112 his3-11,15 ura3. 14439, DDY2402: MATalpha cse4-1 ade2-101 trp1∆901 leu2-3,112 his3-11,15 ura3. YPH499 background 8197, CLY169: YPH499 cdc5-10. 1544, ESM218: YPH499 tub4-1. 405, ESM23: YPH499 cdc31-2. 3054, YMK83: YPH499 spc110-2. 22575, LLY002-2: YPH499 scm3-1.

a. Shifting Cells to the Restrictive Temperature

1. At day -1 (Monday evening): inoculate 5 ml of YPAD medium in a 100 ml

flask with a small amount of yeast cells from plates. Shake the cultures at 23ºC, the

permissive temperature of the mutants, over night.

2. On Tuesday morning inoculate 10 ml of fresh YPAD medium with 500 µl

of the overnight culture. Incubate for 3 hours at 23ºC. Stain DNA of cells from 1 ml

of cell culture as described in b.

3. Transfer the other part of the culture to the 37ºC shaker and incubate for 2

hours. Stain DNA of 1 ml of cells as described in b.

b. DAPI Staining Protocol

Fixing cells with ethanol for DNA staining

1. Centrifuge 1 ml of cells of a culture in an Eppendorf tube (2 min, 6000 rpm,

room temperature). Remove supernatant.

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2. Re-suspend cell pellet in 500 µl of 70 % ethanol. Vortex well and keep cells

on ice until the experiment is finished.

3. Transfer 250 µl of the cell suspension in ethanol to a new Eppendorf tube

(remember to vortex well before transferring the cells).

4. Centrifuge (2 min, 6.000 rpm, room temperature) and re-suspend cells in

about 50 µl of PBS/DAPI.

PBS/DAPI: Dilute DAPI stock solution 10.000-fold in PBS: Add 2 µl of

DAPI stock (1 mg/ml) to 20 ml of PBS in a 50 ml Falcon tube. This solution can be

kept in the fridge (wrapped with aluminium foil around tube) for a couple of months.

5. Inspect cells by fluorescence microscopy. The DAPI staining will be fine

for about one day only. If cells form clumps, treat cells with a short burst in the bath

sonicator.

c. Analysis

Inspect wild type and mutants cells by phase contrast microscopy in the

presence of fluorescence light. Start with wild type cells. Analyse the size of the

daughter cell, the bud, and the localisation of the DAPI staining regions representing

the nuclei. Group cells into G1 (no bud, one DAPI), G1/S (small bud, one DAPI),

S/metaphase (medium to large bud, one DAPI) and anaphase cells (large bud, two

DAPI staining regions; one in the mother cell and one in the bud). Analyse about 50-

100 cells per time point per mutant.

What is the phenotype of the mutants? Are the conditional lethal mutants cell

cycle mutants?

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Day 3/4/5 (Wednesday/Thursday): Fluctuation of protein levels throughout the

cell cycle (Wednesday/Thursday supervised by Leontina and Lucia)

Background

Cell cycle progression is in part driven by the cell cycle dependent degradation of

cyclins and cell cycle inhibitors. As one principal, cell cycle regulators become

modified at epsilon amino groups of Lys residues by the small protein ubiquitin and

then degraded by the proteasome.

Ubiquitinylation

Ubiquitin is a compact globular protein with a C-terminus that extends away from the

main body of the protein into the aqueous space. It is the alpha-carboxyl group of the

C-terminal glycine, which forms isopeptide bonds with target proteins (which can

include another copy of ubiquitin). The isopeptide bond is formed with an (epsilon)

amino group in the side chain of a lysine residue of the target protein.

Activation of ubiquitin

Ubiquitin is activated by an "ubiquitin activating enzyme" or "E1". The E1 enzyme

hydrolyses ATP and forms a complex with the resulting ubiquitin adenylate (Eq. 2.1),

reminiscent of amino acyl adenylate formation in protein synthesis. Ubiquitin is then

transfer to the active cysteine of E1 to form a thiol ester between the C-terminus of

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ubiquitin and the thiol group of the E1. This occurs in concert with adenylation of an

additional ubiquitin. The thiol esters are readily cleaved by reducing agents such as

mercaptoethanol and dithiothreitol and also by hydroxylamine.

Eq. 2.2.1 E1 + ATP + Ub -----------> E1.Ub-AMP + PPi

Eq. 2.2.2 E1.Ub-AMP + Ub ----------> E1-s-co-Ub.AMP-Ub

The structure of a glycine-cysteine thiol ester:

Ubiquitin conjugation

Ubiquitin is not transferred from E1 to a target protein, but rather is transferred to a

family of ubiquitin-conjugation enzymes or ligases, which are named E2 enzymes. E2

enzymes are in most cases the proximal donors of ubiquitin to target proteins. E2

enzymes also have an active site cysteine, and ubiquitin is transferred from the E1 to

the E2 cysteine to form a thiol ester.

Eq. 2.3.1 E1-s-co-Ub.AMP-Ub + E2-SH -----> E2-s-co-Ub + E1.AMP-Ub

Ubiquitin is then transferred to the acceptor lysine of the target protein to form the

isopeptide bond.

Eq. 2.3.2 E2-s-co-Ub + Protein-NH2 -------> E2-SH + Protein-NH-CO-Ub

Multi-ubiquitin chains can be built up on a single lysine of the target protein by

isopeptide bond formation between the carboxyl group of gly76 of one ubiquitin with

the amino group of the side chain of lys48 of the preceding ubiquitin. Substitution of

lysine at position 48 with cysteine results in a mutant ubiquitin, which does not

support multi-ubiquitin chains and is incapable of targeting proteins for proteolysis.

The structures of Ub2 and Ub4 chains have been solved. It seems that four copies of

ubiquitin in a chain may be required to target proteins to the 26S proteasome as these

Ub4 chains exhibit certain structural characteristics that may be recognized by the

proteasome.

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Ubiquitin ligases - E3s

In vivo E2s require an additional protein factor, an E3, to transfer the ubiquitin onto

the protein. The E3 binds the target protein and the E2 enzyme, and therefore selects

the target protein. This implies that the E3 enzyme recognizes a motif in the substrate

protein that targets it for ubiquitinylation (for example: D- or KEN-box).

Fig. 5: Roles of E1, E2 and E3 in Ubiquitin modification of substrate proteins

E3: Anaphase-promoting complex (APC)

The APC is a complex of several proteins which is activated during mitosis to initiate

anaphase. The APC is an ubiquitin ligase or E3 protein that marks proteins for

degradation by the proteasome. Cell cycle progression is uni-directional and the

irreversibility of proteolysis contributes to this process. The APC triggers the

degradation of securin, cyclin B, polo kinase and other cell cycle regulators. The

activity of the APC is inhibited by the spindle checkpoint when not all kinetochores

are properly attached to the mitotic spindle.

There are two forms of the APC which target different sets of proteins and

which are regulated differently. One, which is active early in mitosis and is

responsible for triggering anaphase onset, contains the protein Cdc20. The other,

which is active later when cells exit mitosis, contains Hct1/Cdh1.

�

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Fig. 6: Different E3 ligases play important roles during the cell cycle

Analysis of the levels of Sic1 and Clb2 during the cell cycle

Budding yeast has one cyclin dependent kinase Cdk1 (or Cdc28) and nine cyclins

which associate with Cdk1 at different stages of the cell cycle. Cln1 to Cln3 bind to

Cdk1 in G1 phase, Clb5, 6 in S phase and the 4 mitotic cyclins Clb1-4 in mitosis. B-

type cyclins become inactivated through degradation at specific points in the cell

cycle. For example, Clb5 is degraded at the metaphase to anaphase transition and

Clb2 at the metaphase to anaphase transition and with mitotic exit.

Sic1 is a Cdk1-Clb inhibitor that inhibits Cdk1-Clb (but not Cdk1-Cln)

activity from late anaphase to G1/S. Degradation of Sic1 is facilitated by

phosphorylation. Ubiquitination in late G1 by the E3 enzyme SCF (see Fig. 6) targets

Sic1 for degradation by the proteasome.

Thus, the levels of Sic1 and Clb2 fluctuate during the cell cycle. Analysis of

the protein content of an asynchronous culture reflects the average of the protein

content of cells in different phases of the cell cycle. In order to measure protein levels

at a defined stage of the cell cycle, we have to use synchronized cells. In a

synchronised population, all cells are within a narrow window of the same cell cycle

phase.

Over the past 40 years different methods have been developed in order to

synchronize yeast populations. One way is to reversibly block the cell cycle in G1

using yeast mating pheromone alpha-factor. Alpha-factor binds to a receptor on the

surface of MATa cells and then triggers the mating signalling pathway. As an end

consequence of this signal transduction cascade, the Cdk1-Cln inhibitor Far1 blocks

Cdk1 activity and all cells of the culture arrest in G1 with low Cdk1 activity, high

Sic1 and low Clb1-6 protein levels. Upon washing the cells with culture medium to

22

remove alpha-factor, cells progress synchronously through S phase, G2 and mitosis.

Thus, most cells execute the same cell cycle event in a culture at the same time. It is

therefore possible to analyse cells on a biochemical level and make conclusions about

activity, localisation and protein levels of cell cycle regulators.

a. Synchronisation of cells with alpha-factor

1. On Tuesday, at day -1 inoculate 10 ml of filter sterilized YPAD medium

with cells of yeast strain YAS8 (YPH499 MATa sst1::hisG, No. 1810).

2. On Wednesday, at day 1, dilute culture to OD600 of 0.3 (50 ml culture).

Incubate cells for 3 h at 30ºC. Add 1 µg/ml (50 µl of 1 mg/ml in DMSO) alpha-factor

to the culture.

3. Incubate cells with alpha-factor for at least 2 h 15 min (or if necessary

longer) at 30ºC until >95% of cells show a mating projection. Check by microscopy.

Then, transfer cells into a 50 ml sterile Falcon tube. Sediment cells by centrifugation

(3.200 rpm, 2 min, room temperature). Discard supernatant and wash cell pellet twice

with 40 ml prewarmed 30ºC YPAD medium. Take up cells in 50 ml YPAD and

transfer cell suspension into a fresh Erlenmayer flask (t=0 min). Shake flask at 30ºC.

4. Take samples at 0, 30, 45, 60, 75, 90, 105, 120 and 135 min after the release

of the cell cycle block. For each time point measure OD600, then transfer 2 ml of

culture into 2 ml Eppendorf tube and centrifuge for 2 min at 14.000 rpm. Discard

supernatant. Freeze the cell pellet at -20°C. Analyze cells as described under b. – d.

“Yeast cell lysates using TCA.”

5. In addition, for each time point transfer 1 ml of cell culture in an Eppendorf

tube and centrifuge (2 min, 6.000 rpm, room temperature). Discard supernatant. Re-

suspend cells in 500 µl of 70 % ethanol. Vortex well and keep cells on ice. Follow e.

“DAPI staining protocol”.

6. In addition, for each time point, spin down 1 ml of sample and resuspend

pellet in 70% Ethanol, incubate at 4°C until the end of the time course. At the end of

the experiment incubate all samples for at least an additional 30 minutes at 4°C. At

this stage samples can be kept for days at 4 °C. Follow protocol f. “FACS analysis”.

23

b. Wednesday - Yeast cell lysates using TCA

1. Re-suspend cell pellet in 800 µl H20 and add immediately 150 µl of NaOH

1.85 M. Vortex well and keep tubes on ice for 10 min.

2. Add 150 µl of TCA 55% (w/w). Vortex well and keep tubes on ice for 10

min.

3. Centrifuge at 14.000 rpm for 15-20 min at 4°C. Remove supernatant by

aspiration. Centrifuge briefly for 10 seconds at room temperature and remove the

excess of supernatant by aspiration. Be careful! Do not remove protein pellet.

4. Add HU-DTT and vortex well until pellet is re-suspended. If supernatant

turns yellow, add 1-2 µl of 2 M Tris base to neutralize the pH.

Note: the amount of HU-DTT varies depending on the protein. Normally, re-

suspending 1 OD600 in 150 µl HU-DTT works well for most proteins. For less

abundant proteins (or larger ones) re-suspend 1 OD600 in 75 µl of HU-DTT.

HU-buffer (high-urea buffer): 8 M urea, 5% SDS, 200 mM Tris-Cl pH 6.8, 0.1 mM

EDTA, bromophenol blue, freeze at –20°C. Before use, adjust to 15 mg/ml DTT. HU-

DTT can be kept at -20°C for 3-4 weeks.

5. Heat samples for 15 min at 65°C (not at 95ºC !). The samples can be kept at

-20°C for several weeks.

c. Thursday - Loading SDS-PAGE gels

1. Heat samples in HU buffer for 10-15 min at 65°C. Centrifuge 14.000 rpm

for 2 min at room temperature (not at 4°C).

2. Load 15 µl per lane onto SDS-PAGE. Also load protein standards.

3. Run SDS-PAGE.

24

d. Thursday/Friday - Blotting

1. Assemble semi-dry blot. I = 120 mA per gel; const., U = 15 V (max); t =

115 min.

2. Stain nitrocellulose membrane with Poncau S. Mark MW markers. Wash membrane with PBS.

3. Block nitrocellulose membrane for 1 hour with 10% milk in PBS. 4. Add first antibody (rabbit anti-Clb2 (1:4000) or rabbit anti-Sic1 (1:1000)) in

5% milk/PBS-T. Incubate overnight until Friday. 5. Wash 3x with TBS-T for 5 min each. 6. Incubate filter with appropriate secondary antibody (goat anti rabbit-HRP

(1:5000) in 5% milk/PBS-T) for 1 hour. Wash 6x (5 min) with PBS-T. 7. ECL reaction: Mix solution A and B (1.5 ml each for one blot). Incubate

filter for 1 min. Put filter up side down onto foil. Removed excess liquid using a paper tissue. Cover the backside of the membrane with copy foil. Wrap bottom foil around the copy foil. Insert blot with MW markers up side up in a film cassette. Go to dark room. Expose blot with X-ray film in the dark for 1 min. Expose a second time according to the result of the first exposure.

When (relative to synchronisation time, budding, separated DAPI regions) does Clb2 and Sic1 accumulate? When are they degraded? How does this influence Cdk1-Clb activity? Draw a profile of Cdk1-Clb2 activity during the cell cycle.

e. Wednesday - DAPI Staining Protocol

1. Transfer 250 µl of the cell suspension in ethanol into a new Eppendorf tube

(remember to vortex well before transferring the cells).

2. Centrifuge (2 min, 6.000 rpm, room temperature) and re-suspend cells in

about 50 µl of PBS/DAPI.

PBS/DAPI: Dilute DAPI stock 10.000 in PBS: Add 2 µl of DAPI stock (1

mg/ml) to 20 ml of PBS in a 50 ml Falcon tube. This solution can be kept in the fridge

(wrap aluminium foil around tube) for a couple of months.

3. Inspect cells by fluorescence microscopy. DAPI staining is fine for about

one day only. Count about 50-100 cells per time point. Analyse unbudded G1, small

budded S phase, metaphase and anaphase cells. For the analysis, blot the distribution

of cell types (no, small and large buds; separated DAPI staining regions (anaphase

cells)) over time.

25

f. Wednesday and Thursday - FACS analysis

1. At each time point, spin down 1 ml of sample and resuspend pellet in 70%

ethanol, incubate at 4°C until the end of the time course. At the end of the experiment

incubate all samples for at least additional 30 minutes at 4°C. Samples will be kept at

4°C until Thursday at this stage.

2. On Wednesday, for FACS analysis, spin down samples at 6000 rpm for 2

minutes and resuspend cells in 500 µl of 50 mM Tris-HCl pH 7.8 buffer.

3. Spin down cells at 6000 rpm for 2 minutes, remove supernatant and

resuspend in 1 ml of 50 mM Tris-HCl pH 7.8 buffer.


resuspend in 800 µl 50 mM Tris-HCl pH 7.8 buffer.

5. Add 200 µl of 10 mg/ml RNAse A without DNAse and incubate overnight

at 37°C.

6. Next day (Thursday), spin down cells at 6000 rpm for 2 minutes and

resuspend the pellet in 500 µl of 5 mg/ml Pepsin freshly dissolved in 55 mM HCl and

incubate for 30 minutes at 37°C.


resuspend cells in 1 ml FACS-buffer.


resuspend cells in 500 µl of FACS-buffer.

9. Vortex cells well, and transfer to a new Eppendorf tube, add 1 µl propidium

iodide 1 mg/ml stock solution, incubate for at least 30 minutes at room temperature

(wrap tubes in aluminium foil).

10. Before analysis sonicate cells in bath sonicater for 10 s, transfer cells into

the FACS-tube and add 2 ml of 50 mM Tris-HCl pH 7.8. Read immediately after

dilution.

11. Do FACS analysis according to a preset program.

Stock Solutions 50 mM Tris-HCl: Dissolve 0.8 g Tris-HCl in 100 ml water, adjust pH to 7.8. RNAse A: Dissolve 30 mg RNAse in 3 ml water. Pepsin: Dilute 46 µl of 37 % (concentrated) HCl in 10 ml water. Dissolve 50 mg Pepsin in this 10 ml HCl solution.

26

FACS-Buffer: for 100 ml: 200 mM Tris-HCl pH 7.5 3.2 g 211 mM NaCl 1.2 g 78 mM MgCl2 1.6 g

Day 5 (Friday): The metaphase-anaphase transition

Background: The function of the cohesin complex and separase in sister

chromatid separation

In budding yeast Saccharomyces cerevisiae, as in all other eukaryotes, a multi-subunit

complex called cohesin complex is essential for holding sister chromatids together.

Cohesin is established during DNA replication and persists until the onset of

anaphase. Cohesin ensures correct attachment of sister chromatids to microtubules of

the mitotic spindle. Once chromosomes have achieved biorientation, cohesin resists

the tendency for sister chromatids to be split apart by microtubules until a protease,

called separase, cleaves the cohesin subunit Scc1. This disjoins the sister chromatids

and allows their movement to opposite spindle poles (see Annual Review of Cell and

Developmental Biology 2008 24:105-129).

To dissect the molecular mechanism of cohesion, the assembly of cohesin

from its four subunits has been investigated (Smc1, Smc3, Scc1, and Scc3). Using

EM, biochemistry and crystallography it was shown by the Nasmyth group that Smc

protomers fold up individually into rod-shaped molecules. Smc1 and Smc3 bind to

each other via heterotypic interactions between their dimerization domains to form a

V-shaped heterodimer. At the opposite end, the two heads of this Smc1/3 heterodimer

are connected by the cleavable Scc1 subunit. Cohesin therefore forms a large tripartite

proteinaceous ring, within which sister DNA molecules might be entrapped and

thereby held together (see Figure 7; panel a).

27

Fig. 7: Ribbon model of the yeast Smc1 head homodimer complex with C-

termini of Scc1 and of the cohesin ring. (a) Scc1 connects Smc1´s and Smc3´s

head domains to form a tripartite ring structure. (b) Two Smc1 head domains

(red, blue) dimerize by sandwiching two molecules of ATP analogues in-

between their interaction surfaces. Scc1‘s winged helix domain (green,

yellow) binds tightly to two ß-strands of Smc1.

28

Fig. 8: Models for cohesin chromosome tethering. In the ring model (a) soluble

chromatin-free single cohesin topologically entraps sister chromatids inside the ring-

shaped structure. Alternatively, the snap (b) and bracelet (c) models suggest that

chromatin-bound cohesins form oligomers mediated by the coiled coils or hinges,

respectively. In these models chromatids are held by a specific DNA-protein

interaction inside or outside the complex. The actual molecular mechanism for

chromosome tethering most likely is a hybrid of the models presented.

Separase is a cysteine protease responsible for triggering anaphase by hydrolysing

cohesin which is the protein responsible for binding sister chromatids during

metaphase. Separase is associated with securin, which prevents separase from

cleaving cohesin. Securin is ubiquitinated (APC) and hydrolysed by the proteasome.

This step activates separase.

29

Induction of anaphase by cohesin cleavage

Today (Friday) we will perform the Nasmyth experiment demonstrating that cohesin

cleavage can trigger anaphase. We will first arrest cells at the metaphase-anaphase

transition by depleting the APC specificity subunit Cdc20. Cells arrested in this way

have high Cdk1-Clb2/Clb5 activity, high levels of securing Pds1, no separase Esp1

activity, the phosphatase Cdc14 is entrapped in the nucleolus, the cohesion complex

holds the sister chromatinds together, the spindle is short (2 µm) and located in the

mother cell body.

The yeast strain K116 contains an engineered Scc1-Tev (TEV recognition site

at amino acid 268) cohesin subunit, which can be cleaved by the induction of the

protease Tev (the consensus Tev cleavage sequence is Glu-X-X-Tyr-X-Gln-Gly/Ser;

cleavage occurs between the conserved Gln and Gly/Ser residues; X is any amino

acid). As soon as Scc1-Tev is cleaved, the sister chromatids are disjointed and an

anaphase-like spindle will form.

Centromere DNA can be marked by inserting up to 100-200 copies of the tetO

DNA binding motif next to for example CEN5 as in case of K116 (yeast cells have 16

chromosomes – thus 16 centromere regions, which are only 125 bp in length). tetR-

GFP binds to the tetO and thereby labels the marked centromere region of living cells.

This way, the behaviour of centromeres can be followed by live cell imaging or of

fixed cells.

a. Induction of anaphase by cleavage of the cohesin complex

1. On Wednesday - day -2: Inoculate strain K116 (MATalpha scc1::HIS3

SCC1-TEV268-HA3::LEU2 pGal1-NLS-Myc9-TEVprotease-NLS2::TRP1 (10-fold

integrant by southern) ura3::3XURA3 tetO112 his3::HIS3tetR-GFP pMet3-HA3-

Cdc20::TRP1 ade2-1 can1-100 GAL, psi+) in 10 ml SR medium with adenine (100

mg/l) at 30ºC. Shake cells over night.

2. On Friday – day 1: Dilute cells in 20 ml SR medium with adenine to OD600

of 0.35 (8.00). Incubate for 2.5 hours at 30ºC (10.30). Then sediment cells by

centrifugation and take up cells in 10 ml YPAR medium with additional 2 mM

methionine and 2 mM cysteine to repress the MET3 promoter. Shake cells for 2-2.5

30

hours (12.30 – 13.00). Check after 1, 2 and 2.5 hours of whether cells are uniformly

arrested in metaphase (cells with a large bud).

3. As soon as a uniform cell cycle arrest is achieved, add 2% filter sterilized

galactose in order to induce expression of Gal1-Tev (t=0 min).

4. Take 1 ml samples after 0, 15, 30, 45, 60, 75 and 90 min. Fix cells with

formaldehyde (15.00 – Seminar !).

b. Fixing GFP cells with formaldehyde – after the seminar !

1. Take 1 ml of cell culture, 100 µl 1 M KPO4 pH 6.5, 100 µl formaldehyde

solution. Incubate for 10 min at room temperature.

2. Wash cells 2x with 10 ml PBS (by spinning at 3.000 rpm for 2 min in either

the Eppendorf centrifuge or the midifuge).

3. Incubate cells for 30 min at room temperature in 100 µl DAPI (PBS + 1

µg/ml DAPI).

4. Wash with PBS (as before) until DAPI background is low (1-3 x).

Resuspend cells in a suitable volume PBS. Add 4 µl onto cover slip. Cover sample

with cover slip. Inspect cells by fluorescence microscopy.

5. Do you see anaphase spindle elongation as indicated by the separation of

the centromere signal?

SR medium: Synthetic medium with 3% Raffinose. 6.7 g Bacto-yeast nitrogen base 30 g raffinose 100 mg adenine ad to 1 l and filter sterilize.

YPAR medium with additional 2 mM methionine and 2 mM cysteine

Mix A: Bacto Yeast extract 10 g Bacto Peptone 20 g Bi-dest. Water up to 700 ml (in 1 l bottle)

Autoclave. Mix B: Raffinose 30 g

Dissolve in 250 ml bi-dest. water (if necessary, heat up to 50°C to dissolve).

Adjust volume to 300 ml. Filter sterilise.

31

PART II: Group will be split in 2x3 working with mammalian cells (Conny, Ilaria and

Uschi). Two will perform advanced microscopy with Johanna and Sarah.

PART II: WORKING WITH MAMMALIAN CELLS Day 6-7 Monday and Tuesday: 6 students – supervised by Conny, Ilaria and

Uschi

Cell cycle analysis of mammalian cells

The goal of this part of the practical course is the analysis of cell cycle progression of

mammalian cells. Methods of choice are microscopy (immunostaining of fixed cells)

and the analysis of protein levels of marker proteins.

Methods:

Tissue culture work Cell culture is the process by which either prokaryotic or eukaryotic cells are grown

under controlled conditions. In practice the term "cell culture" refers to the culturing

of cells derived from multi-cellular eukaryotes, especially animal cells. The historical

development and methods of cell culture are closely related to those of tissue culture

and organ culture.

Maintaining cells in culture

Cells are grown and maintained at an appropriate temperature and gas mixture

(typically, 37°C, 5% CO2) in an incubator. Culture conditions vary widely for each

cell type. Variation of conditions for a particular cell type can result in different

phenotypes. For our purposes we use the very commonly used DME-medium

supplemented with 10% FCS and glutamine.

Media changes

The purpose of media changes is to replenish nutrients and avoid the build up of

potentially harmful metabolic biproducts and dead cells. In case of suspension

32

cultures, cells can be separated from the media by centrifugation and resuspended in

fresh media. For adherent cultures, the media can be removed directly by aspiration.

In our case, we will use the RPE-1 cells, which is a telomerase immortalized, diploid,

stable cell line.

Passaging of cells

Passaging or splitting of cells involves transferring a small number of cells into a new

vessel. Cells can be cultured for a longer time if they are split regularly, as it avoids

the senescence that is associated with prolonged high cell density. Suspension cultures

are easily passaged by diluting cells in fresh media. For adherent cultures, cells need

to be detached; this is done with a mixture of trypsin-EDTA. A small number of

detached cells will be used to seed a new culture.

Synchronization of mammalian cells

In general, cells in culture do not divide synchronously. Therefore, several methods

have been established to synchronize them. Most commonly thymidine, aphidicolin or

nocodazole are used for this purpose. The thymidine block arrests cells at the G1/S

boundary. After release by washing the cells, cells undergo DNA replication followed

by mitosis. Aphidicoline blocks the DNA synthesis in S-phase. Nocodazole blocks the

polymerization of tubulin and thereby activates the spindle assembly checkpoint that

blocks cell cycle progression by inhibiting APCCdc20 activity. Released cells will

reinstall the spindle apparatus and then synchronously undergo mitosis.

Protein analysis by Western Blotting To compare proteins levels at different points of the cell cycle, we will analyze

protein extracts of cells by immunoblotting. Immunoblotting is a method, which

detects specific proteins in a given sample or cell extract. It uses SDS-PAGE to

separate denatured proteins approximately according to their molecular weight. The

proteins are then transferred from the gel onto a membrane, where they are "probed"

with specific antibodies directed against the protein of interest. As a result, one can

examine the amount of a protein in a given sample for example changes of protein

levels during the cell cycle.

33

Immunofluorescence staining of cells Immunofluorescence is the labeling of antigens with fluorescent antibodies. This

technique is often used to visualize proteins within its cellular environment and to

determine cell cycle dependent changes in protein levels and localization.

Fluorescently labeled tissue culture cells are studied using a conventional

fluorescence or a confocal microscope. Most commonly, staining of cells by

immunofluorescence employs two sets of antibodies (like the immunoblot): a primary

antibody is used that is directed against the protein of interest. A subsequent

secondary antibody recognizing the primary antibody is coupled to a fluorescent dye.

Fluorescence Microscopy This method is of critical importance for modern life sciences, as it is extremely

sensitive, allowing the detection of single molecules. Cells can be fixed and analyzed

by indirect immunofluorescence with primary and secondary antibodies. Specific dyes

that for example bind to DNA are used to stain cellular structures (e.g. DAPI or

Hoechst). A fluorescence microscope is used to visualize the signal of immuno-

stained cells.

Experimental procedures 1st group:

Mitotic arrest/release of cells with nocodazole The aim of this experiment is to analyze different protein levels and localizations in

synchronized cells during the cell cycle. Cells can be arrested in mitosis by

nocodazole treatment, which is interfering with microtubule polymerisation.

Therefore, we will release the cells from nocodazole block and look at Cyclin A and

B and GAPDH protein levels at different time points by immunoblotting.

Additionally, we will analyze the centrioles (centrin-GFP) and the microtubules (anti-

tubulin) of cells via fluorescence microscopy.

Day -2 (Saturday):

1. Grow RPE-1 Centrin-GFP cells as monolayer in DMEM-F12 medium

supplemented with 10% FCS and 1% glutamine.

34

Day -1 (Sunday):

2. Add 100 ng/ml nocadozole in DMSO to the media (or previously mix fresh

media with nocadozole then add the media to the cells. Do not add large

volumes of DMSO to the cells since it is toxic!).

3. Incubate cells for 15-16 h.

4. Monday: (students will start at this point). Most of the cells should be in

mitosis. Mitotic cells are round-shaped and lightly attached to the surface. So

be very careful during the washing procedures!!!

5. Release the cells from the block by removing nocodazole:

a. Wash the cells twice with 1x PBS.

b. Add volume of pre-warmed media to the cells.

6. Start the kinetics and take samples at the following time points: 0 min, 20 min,

40 min, 60 min, 100 min, 140 min, 180 min.

a. For Western blot analysis:

i. Take cells (by adding 1 ml Trypsin solution, 5 min at 37°C) + media,

centrifuge briefly (1500 rpm, 4°C, 5 min)

ii. Resuspend cell pellet in 10 µl of 1x PBS and add 40 µl of Laemmli buffer,

resuspend and boil at 95°C for 5 min.

iii. Store cell extract at -20 °C for further analysis.

b. For Immunofluorescence:

i. Remove medium, wash with 1x PBS, add ice-cold methanol.

ii. Fix cells in methanol for 5 min at -20°C.

iii. Wash cells 1× with 1x PBS and store in 1x PBS at 4°C.

2nd group:

Inhibition of mitotic kinases and analysis of mitotic exit The goal of this experiment is to analyse the importance of various kinases for mitotic

exit. Therefore, we will arrest cells in prometaphase (activation of the spindle

assembly checkpoint by Eg5 inhibition) and inhibit the mitotic kinases Plk1, Aurora

A and Cdk1 with small molecule inhibitors. In order to follow the differences in cell

cycle progression, we will examine Cyclin B1 and GAPDH protein levels by

immunoblotting, because Cyclin B1 is known to be degraded at metaphase-anaphase

35

transition. Moreover, we will analyse the displacement of the centrosomal linker

protein C-Nap1, which is known to be phosphorylated and released from the

centrosomes in mitosis.

Day-2 (Saturday):

1. Grow RPE-1 Centrin-GFP cells as monolayer in DMEM-F12 medium


Day -1 (Sunday):

2. Add 5 µg/ml STLC in DMSO to the media (or previously mix the fresh media

with nocadozole then add the media to the cells. Do not add large volumes of

DMSO to the cells since it is toxic! STLC blocks the kinesin-5 motor Eg5).

3. Incubate cells for 15-16 h.



be very careful while the washing procedures!!!

5. Add following inhibitors to the medium:

1 - Untreated

2 - Plk1 inhibition (BI2536-1:1000)

3 - Aurora A inhibition (CPCA-1:1000)

4 - Cdk1 inhibition (RO3306-1:2000)

5 - Cdk1 inhibition + proteasome inhibitors (MG132 1:2500)

6. Incubate the cells for 2 h at 37°C in the incubator.

7. Collect cells:

a. For immunoblot analysis:

i. Take cells (by adding 1 ml Trypsin, 5 min at 37°C) + media and centrifuge

briefly (1500 rpm, 4°C, 5 min).

ii. Resuspend in 10 µl of 1x PBS and add 40 µl of Laemmli buffer to the cell

pellet; resuspend and boil at 95°C for 5 min.

iii. Store cell extract at -20°C for further analysis.

b. For immunofluorescence:



iii. Wash 1× with 1x PBS and store in 1x PBS at 4°C.

36

3rd group:

Specific inhibition of Plk1 kinase by analog-sensitive mutant In this part we will demonstrate the specific inhibition of the mitotic kinase Plk1 by

using a cell line carrying a particular Plk-as (analog-sensitive) allele. This is achieved

by the use of RPE-1 eGFP-Plk1-as cell line in which both copies of the Plk1 Locus

were disrupted and the Plk1-as allele was introduced by homologous recombination.

Thereby, the catalytic domain of the kinase has been enlarged to ensure the fitting of

bulky purines as inhibitor like 3MP-PP1. By using this purine analog we will

specifically inhibit Plk1 kinase and analyse its phenotype in mitosis.

Day-2 (Saturday):

1. Grow RPE-1 eGFP-Plk1-as cells as monolayers in DMEM-F12 medium


Day -1 (Sunday):

2. Add 5 µg/ml STLC or 10 µM 3MB-PP1 in DMSO to the media (or previously

mix the fresh media with the inhibitors, then add the media to the cells. Do not

add large volumes of DMSO to the cells since it is toxic!).

3. Incubate cells for 15-16 hrs.



be very careful while the washing procedures!!!

5. Release the cells from the block:

a. Wash the cells twice with 1x PBS.

b. Add equal volume of pre-warmed media to the cells.

6. Start the kinetics and take samples at the following time points: 0 min, 20 min,

40 min, 60 min, 100 min, 140 min, 180 min.

a. For immunoblot analysis:

i. Take cells (by adding 1ml Trypsin, 5min 37°C) + media and centrifuge

briefly (1500 rpm, 4°C, 5 min).

ii. Resuspend cells in 10 µl of 1x PBS and add 40 µl of Laemmli buffer,

resuspend and boil at 95°C for 5 min.

iii. Store cell extract at -20°C for further analysis.

b. For immunofluorescence:

37



iii. Wash 1× with 1x PBS and store in PBS at 4°C.

Day 1-2:

IMMUNOBLOT ANALYSIS

Day 1: SDS-PAGE

Protein samples will be separated under denaturing conditions by SDS-PAGE. Mini

gels (1 mm or 1.5 mm) from BioRad will be used with combs for 10 or 15 lane. The

following scheme will be applied to prepare two Mini gels.

Separating Gels Stacking Gels Compounds

6 % 8 % 10 % 4 % 30 % Acrylamide 2 ml 2.6 ml 3.35 ml 650 µ l ddH2O 5.4 ml 4.75 ml 4.05 ml 3.1 ml 1.5 M Tris-HCl pH 8.8 2.5 ml 2.5 ml 2.5 ml - 0.5 M Tris-HCl pH 6.8 - - - 1.25 ml 10 % SDS 100 µ l 100 µ l 100 µ l 50 µ l 10 % APS 50 µ l 50 µ l 50 µ l 25 µ l TEMED 5 µ l 5 µ l 5 µ l 5 µ l Table 1: Amounts of solutions required to prepare SDS-PAGE gels. Be aware that

acylamide is very toxic and be taken up through the skin. Ware gloves!

Appropriate amounts of each sample will be loaded on SDS-PAGE gel and run at

25 mA per gel for approximately 1 h in SDS running buffer. Unstained or prestained

broad range molecular weight markers (Fermentas) are used to compare protein sizes.

38

Fig. 9. Fermentas Protein

Ladder for SDS-PAGE.

After the SDS-PAGE, proteins will be transferred onto nitrocellulose membranes by

electro-blotting. We are going to transfer our proteins to membranes according to the

protocol described below. Transfer will take 1 h with 110 mA per blot. After

blocking, we will cut the membranes into three pieces and probe them with three

different antibodies, Cyclin A (around 60 kDa), Cyclin B (around 50 kDa), and as a

loading control GAPDH (around 36 kDa).

• Run a SDS-PAGE (10% resolving gel, 4% stacking gel).

• Prepare Whatman paper (6 × 9 cm) and PVDF blotting membrane (6 × 9 cm).

• Take out the gel and cut off the stacking gel.

• Activate the PVDF membrane in MeOH for 15 sec, wash for 5 min in water

and then for 5 min blotting buffer.

• Soak the Whatman paper in blotting buffer.

• Assemble a wet blot as described in the following scheme:

1. Put soaked 3 Whatman paper on the anode, get rid of air bubbles in the paper

by rolling a pipette over the paper.

2. Put the blotting membrane without any air bubbles.

3. Put the gel.

39

4. Again 3 piece of soaked Whatman paper.

• Get rid of the air bubbles in the sandwich!!!

• Close the blotting sandwich in the grid.

• Connect to the power supply.

• Switch on an run with a current of 110 mA .

• Transfer for 1.5 h.

• Disassemble the blotting sandwich.

Immunostaining of the immunoblot

• Block membrane with 5% milk in TBS-T (1x TBS- 0.1% Tween) for 30-60 min at

ambient temperature.

• Prepare fresh antibody solution (recommended dilution) in 5% milk in TBS-T

1st group: anti-Cyclin A (1:2000) (anti-mouse secondary antibody)

anti-Cyclin B1 (1:500) (anti-mouse secondary antibody)

anti-GAPDH (1:10000) (anti-rabbit-secondary antibody)

2nd group: anti-Cyclin B1 (1:500) (anti-mouse secondary antibody)

anti-GAPDH (1:10000) (anti-rabbit secondary antibody)

3rd group: anti-Cyclin A (1:2000) (anti-mouse secondary antibody)

anti-Cyclin B1 (1:500) (anti-mouse secondary antibody)

anti-GAPDH (1:10000) (anti-rabbit-secondary antibody)

• Incubate over night at 4°C .

• Next day, wash 3× with TBS-T for 5 min each.

Whatman paper 3x

Whatman paper 3x

Resolving gelBlotting membrane

+ Anode

- KatodeWhatman paper 3x

Whatman paper 3x

Resolving gelBlotting membrane

+ Anode

- Katode

40

• Incubate the membrane with secondary antibody (1:10000 in 5% milk in TBS-T)

for 1 h.

• Wash the blotting membrane 4× with TBS-T for at least 5 min.

• ECL-detection of the immuno-detectable protein:

o Mix solution A and B 1:1 (1.5 ml + 1.5 ml for each blot).

o Distribute it equally on the membrane.

o Incubate for 1 min.

o Put membrane up side down onto a foil.

o Remove excess liquid using a paper tissue.

o Cover the backside of the membrane with copy foil.

o Expose the blots at various time points with ImageQuant LAS 4000.

Immunofluorescence staining: • Permeabilization: treatment with 0.1% Triton-X-100 in 1x PBS for 10 min at

ambient temperature.

• Wash 1× with 1x PBS at ambient temperature.

• Block slides with 10% fetal calf serum (to saturate sides where no protein is

bound. It will reduce unspecific binding of the antibody to the glass slide!) for

30 min at room temperature.

• Wash 1× with 1x PBS.

• Incubation with first antibody:

1st group: anti-α-tubulin (1:1000)

2nd group: anti-C-Nap1 (1:100)

3rd group: anti-C-Nap1 (1:100) / anti-centrin (1:50)

o Prepare 3 wet chambers for the incubation of the slides: cut a Whatman

paper such that it fits in a big Petri dish; soak it with water and decant

excess water.

o Depending on the number of samples, prepare a master mix: Total Volume =

number of samples X × 15 µl. (But please prepare for 2 samples more master mix

solution, otherwise you will run out of the mastermix for the last coverslips!!)

o Example: to 500 µl add 1 µl of first antibody.

o Add 15 µl of antibody solution to the cells on the slide and cover it with

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parafilm.

o Incubate for 1h in the wet chamber at 37°C to allow binding of the antibody.

• Washing of the slides

o Take the parafilm off.

o Wash the slides in the tank 3× with 1x PBS.

• Incubate with the secondary antibody

o The secondary antibody is coupled to a fluorescent dye (here AlexaFluor).

o Dilute the antibody 1:500 as described.

o Incubate for 30 min as described above.

• Wash the slides as described above.

• Add 500 µl Hoechst-TBS-T (1:2000) to stain the DNA and incubate for 10 min.

• Wash the slides as described above.

• Add a drop of ProLong mounting medium on top of the slide and put the coverslip

onto the slide.

• Leave to dry at room temperature in the dark for at least 1 h or overnight.

• Analyse the slides by microscopy.

Mechanism and Regulation of Eukaryotic Cell Division · Cell division takes place in timely and...

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