0) chromosome segregation in mitosis and meiosis – an introduction

CONTENTS:
0) CHROMOSOME SEGREGATION IN MITOSIS AND MEIOSIS – AN INTRODUCTION
1) BACTERIAL EXPRESSION AND AFFINITY PURIFICATION USING THE PET-SUMO
SYSTEM
2) IN VITRO EXPRESSION OF A FLUORESCENTLY TAGGED PROTEIN
3) ISOLATION OF XENOPUS TESTES AND SPERM NUCLEI
4) PREPARATION OF CSF-EXTRACTS FROM XENOPUS EGGS AND THEIR USE TO
STUDY EXIT FROM MEIOSIS II/MITOSIS
5) ANALYSIS OF CELL CYCLE REGULATORS BY MICRO-INJECTION EXPERIMENTS
INTO XENOPUS EMBRYOS
6) MICROSCOPY OF FLUORESCENTLY-LABELED MARKER PROTEINS IN LIVING
CELLS
7) CHARACTERIZATION OF ISOGENIC AND INDUCIBLE TRANSGENIC CELL LINES BY
FLOW CYTOMETRY AND CHROMOSOME SPREADING
8) INSTRUCTIONS AND QUESTIONS FOR THE PROTOCOL
9) APPENDIX (MARKER; HEMOCYTOMETER; TIME TABLE)
ad 0) CHROMOSOME SEGREGATION IN MITOSIS AND MEIOSIS – AN INTRODUCTION
In a typical eukaryotic cell division cycle the replication of chromosomes in S-phase is timely
separated by gap-phases (G2 and G1) from their segregation in M-phase (Figure 1). (G1-, Sand G2-phase together are commonly referred to as interphase.) By light microscopy Mphase appears as the most dramatic and beautiful phase of the cell cycle and, hence, has
fascinated cell biologists ever since it was first described by Walther Flemming around 1890
(Figure 1). It can be subdivided into mitosis, the segregation of the chromosomes (or nuclear
division), and cytokinesis, the division of the cytoplasm. Entry into prophase of mitosis
involves dramatic changes like nuclear envelope breakdown, spindle formation and
chromosome condensation. During prometaphase, sister chromatids attach via their
kinetochores to microtubules emanating from opposite poles of the spindle apparatus and
congress towards its equatorial plane. Once all chromosomes have properly aligned in
metaphase, sister chromatids spring apart and are being pulled towards opposite ends of the
mitotic spindle, a process that represents the visible hallmark of anaphase. Disassembly of
the spindle, decondensation of chromosomes, and reformation of the nuclear envelope in
telophase conclude mitosis and are followed by cytokinesis, thereby giving rise to two
identical daughter cells.
Chromosome segregation has to be highly accurate since any mistake has fatal
consequences for the cell. Chromosome missegregation during mitosis most likely is an early
step in tumorigenesis. Furthermore, many tumors exhibit persistent chromosome instability
(CIN) and often there is a direct correlation between the degree of aneuploidy and the
malignancy of tumors, which manifests in poor prognosis. Meiosis, which leads to the
formation of haploid gametes, is characterized by two consecutive nuclear divisions without
intercepting replication. Mistakes in chromosome segregation during this specialized cell
cycle results in sterility or trisomies like Downs syndrome.
Figure 1: Vertebrate mitosis as seen by Walther Flemming. Cartoon depicts cohesin, which consists of at least
four subunits, Smc1 and –3 and Scc1 and –3. The former belong to the conserved family of SMC (structural
maintenance of chromosomes) proteins and form long anti-parallel coiled coils, which interact with each other via
a flexible hinge region. The latter interact with the ATPase head domains of the Smc-heterodimer thus closing the
other end of the cohesin ring.
Protein phosphorylation and degradation drive the cell cycle
From the time of their generation in S-phase (replication) until their segregation in anaphase
of mitosis, sister chromatids are held together by virtue of a ring-shaped multi-protein
complex, cohesin, which likely encloses the two DNA double strands in its middle (Figure 1).
During this time a cohesin cleaving protease, separase, is held inactive by association with
its inhibitor securin (Figure 2). Only in late metaphase, when all chromosomes have properly
aligned, is securin degraded via the ubiquitin-proteasome system (UPS). The ubiquitin-ligase
(E3) responsible for specific degradation of securin is the anaphase-promoting complex or
cyclosome (APC/C) in conjunction with its accessory protein Cdc20. It recognizes a so-called
destruction box (DB) characterized by the motif Arg-x-x-Leu (x = any amino acid) in the
aminoterminal region of securin and other substrates. Once activated by the removal of
securin, proteolytically active separase now cleaves the Scc1/Rad21 subunit of cohesin,
thereby opening the ring and triggering anaphase.
Key cell cycle players are regulated in their activities by phosphorylation and
dephosphorylation. Cyclin-dependent kinases (CDKs) are serine/threonine-specific protein
kinases of crucial importance for this form of cell cycle control. In higher eukaryotes various
CDKs associate with various cyclins and become active at different times and locations,
thereby allowing for substrate specificity and, more importantly, specific integration of
regulatory signals. Cdk1 in conjunction with cyclin B1 can be considered the master regulator
of mitosis and meiosis. For example, activation of this kinase is necessary and sufficient to
drive cell-free extracts into a mitosis-like state. Conversely, inactivation of Cdk1 by
degradation of its regulatory subunit cyclin B1 via UPS directly correlates with exit from
mitosis. Interestingly, the specificity factor here is the same APC/CCdc20 that mediates the
destruction of securin. Thus, activation of this crucial ubiquitin ligase at the end of metaphase
couples chromosome segregation with subsequent exit from mitosis.
Figure 2: A simplified model of late mitotic events in vertebrates. A surveillance mechanism, the spindle assembly
checkpoint, keeps APC/CCdc20 inhibited until the last chromosome has properly attached to the spindle apparatus.
ad 1) BACTERIAL EXPRESSION AND AFFINITY PURIFICATION USING THE PET-SUMO
SYSTEM
As a result of fertilization and rise in intracellular Ca2+, calcium/calmodulin-dependent kinase
II (CaMKII) becomes active and phosphorylates XErp1, thereby creating a binding site for
polo like kinase 1 (called Plx1 in Xenopus). Plx1 binds via its so-called polo box domain
(PBD) to pre-phosphorylated XErp1, is simultaneously activated by conformational change
and then phosphorylates additional sites on XErp1, thereby creating a phospho-degron,
which results in SCF-dependent degradation of the APC/C inhibitor (Figure 3b). However,
even during a CSF-arrest the levels and activities of XErp1, APC/C, and cyclin B1 have to be
modulated.
Because cyclin B1 is constantly produced in eggs, some of it has constantly to be
degraded, too, simply to prevent hyper-activation of Cdk1 during a prolonged CSF-arrest,
which would counteract the rapid exit from meiosis II upon fertilization. Cdk1-dependent
phosphorylation of XErp1’s N-terminal half contributes to its SCF-dependent degradation. In
addition, Cdk1-dependent phosphorylation of residues in the C-terminal half weaken the
binding of XErp1 to APC/C (Figure 3a). Both these mechanisms, by which Cdk1 limits its own
activity, are counteracted by Mos, a meiosis-specific mitogen-activated protein kinase kinase
kinase (MAPKKK). The Mos-triggered MAPK cascade culminates in activation of ribosomal
S6 kinase (Rsk), which phosphorylates XErp1, thereby creating a landing pad for protein
phosphatase 2A (PP2A). Upon its recruitment to XErp1, PP2A now acts as an antagonist of
Cdk1 (Figure 3a). It de-phosphorylates N- and C-terminal residues, thus increasing XErp1’s
stability and APC/C-inhibitory potential. Capitalizing on these findings, it has been shown that
a C-terminal fragment of XErp1 is resistant against SCF-dependent degradation but retains
its ability to inhibit APC/CCdc20, especially when C-terminal inhibitory Cdk1 phosphorylation
sites are additionally mutated. You will affinity purify this same XErp1 fragment from
expression cultures of E. coli and later test how it influences the release of a CSF-extract into
interphase and the cell cycles of micro-injected early frog embryos.
Figure 3: Regulating the cytostatic factor in vertebrate female meiosis II (modified from Wu and Kornbluth, 2008).
(A) Regulation of XErp1 (Emi2) stability and activity during CSF-arrest. N- and C-terminal phosphorylations by
Cdk1 result in degradation and inactivation, respectively, of XErp1. These Cdk1 effects are antagonized by Mos,
Rsk, and PP2A. (B) Phosphorylation-dependent degradation of XErp1 as a result of fertilization-induced rise in
intracellular Ca2+.
The genetic code is degenerate. The existing codon options are used in an unequal
frequency in different species (Figure 4). Consequently, tRNA supply shortages can limit
heterologous expression. For the lab course the C-terminal 160 amino acids (position 491651; 18,3 kDa) of a XErp1 mutant with two inhibitory Cdk1 phosphorylation sites changed to
alanine (Thr- 454,551-Ala) have been expressed in two E. coli strains, BL21 and Rosetta 2,
both of which are deficient in lon and ompT proteases for enhanced protein stability. The
difference between both strains is that Rosetta 2 (but not BL21) cells carry an additional
plasmid that drives the over-expression of tRNAs for seven codons rarely used by E. coli but
frequently found in vertebrates (AUA, AGG, AGA, CUA, CCC, CGG, and GGA). Additional
features of this plasmid are a chloramphenicol resistance gene (Cam) for selection and a
p15a ori, which is compatible with colE1 replicons used in typical expression plasmids.
Figure 4: The codon quality of a section of the coding sequence for His6-Sumo-XErp1(491-651) is given in relative
adaptiveness values for the originating organism Xenopus laevis (top) and E. coli (bottom). (For each amino acid
the codon with the highest frequency value is set to 100% relative adaptiveness. All other codons for the same
amino acid are scaled accordingly.) Codons with a relative adaptiveness <20% and <10% are shown in grey and
red, respectively. Images were created with a graphical codon usage analyser (for further details see:
http://gcua.schoedl.de/sequential_v2.html).
The XErp1 fragment is expressed in fusion with an N-terminal His6-Sumo-tag (11,3 kDa). The
His6-tag allows affinity purification of the 29,6 kDa expression product on chelating agarose
(IMAC = immobilized metal ion affinity chromatography). Sumo is a small modifier, which is
conjugated to other proteins just like the related ubiquitin. However, it does not mark the
corresponding substrates for destruction but rather changes their cellular localization or
activity. Like ubiquitin, Sumo is removed from traget proteins and recycled once it has fulfilled
its task. Therefore, eukaryotic cells contain very active and highly Sumo-specific deconjugating proteases. In E. coli expressions, the well-expressed Sumo is not only used to
drive expression and improve the solubility of attached proteins but primarily because a
Sumo protease can be used to fully remove the tag at the end of the purification. This can be
of great advantage because tags frequently interfere with biological activity or crystallization
behavior of recombinant proteins.
Experiment:
• You will be given over-night cultures of either BL21 (groups 1-4) or Rosetta 2 (groups 5-8),
in which expression of His6-Sumo-XErp1(491-651) has been induced by IPTG. The optical
density (OD600) will be indicated on each flask.
• Assuming that the OD600 was not a dimension-less number but instead had the unit ml-1 (i.e.
would relate to 1 ml), harvest in a 1,5 ml tube the equivalent of 0.2 OD600 by centrifugation in
a table top centrifuge (1 minute at full speed). Immediately, remove all supernatant by
pipetting and then resuspend the cell pellet in 40 µl SDS-sample buffer (1x). This is your
sample “after induction” (AI). You will also receive from your tutor a corresponding sample
before IPTG addition (BI) for comparison.
• Harvest 250 ml of expression culture by centrifugation in a JA-14 rotor (Beckman) for 10
minutes at 6.000 rpm and 4°C. During this time…
• Prepare 50 ml of lysis buffer and cool on ice:
1x PBS; (additional) 0,4 M NaCl; 5 mM imidazole; 5 mM β-mercaptoethanol.
• Prepare 15 ml of wash buffer and keep cool until use (next day):
1x PBS; (additional) 0,4 M NaCl; 25 mM imidazole; 5 mM β-mercaptoethanol.
• When centrifugation is done: Decant all supernatant (collect in beakers for later sanitation)
and store cell pellet on ice until lysis.
• Resuspend your cells in ice cold 40 ml lysis buffer. Pipette up and down until all clumps
have resolved. Then: While cooling your samples, lyse the cells in a micro-fluidizer (Avestin,
EmulsiFlex C5; groups 1, 3, 5, 7) or by sonication (groups 2, 4, 6, 8) according to your tutor’s
instructions. When the yellow/brownish suspension changes from milky/cloudy to
clear/opalescent, lysis is usually complete.
• Remove any unlysed cells and debris by centrifugation in a JA-20 rotor (Beckman) for 10
minutes at 15.000 rpm and 4°C. During this time…
• Equilibrate Ni2+-NTA agarose (250 µl of packed beads) by washing it with 3x 1 ml lysis
buffer in a 2 ml plastic column. Then: Close bottom of column with a yellow cap leaving the
beads covered with approximately 250 µl of lysis buffer.
• When the centrifugation is done: Collect the clarified supernatant into 50 ml conical tube.
This is your crude extract (CE). Take out 5 µl and combine with 5 µl SDS-sample buffer
(2x).
• Resuspend your Ni2+-NTA agarose by pipetting up and down with a cut-off tip and transfer
into 50 ml conical tube containing your crude extract. Rinse and save the empty column.
• Rotate over night (or for at least 3 hours) at 4°C to keep beads in suspension.
• Pellet beads by centrifugation in a clinical centrifuge (swing out rotor) for 5 minutes at 800
rpm and 4°C. Take out 5 µl of the supernatant (SN) and combine with 5 µl SDS-sample
buffer (2x). Only after taking this sample: Pipette off supernatant and discard. Be sure not to
loose any Ni2+-NTA beads in this step and rather leave some supernatant behind.
• Resuspend beads in residual supernatant with a cut-off tip and transfer into in an empty 2
ml plastic column positioned with a column holder in a 50 ml conical tube on ice.
• Wash the beads with 5x 1,5 ml cold wash buffer. (You may let the buffer drain completely in
each step because due to capillary forces, the Ni2+-NTA beads do not easily run dry.)
• After all wash buffer has drained: Put the tip of your column in a 1,5 ml tube and collect
bound protein by eluting the beads with 3x 0,15 ml pre-cooled elution buffer. After carefully
mixing your 0,45 ml of eluate (EL), take out 10 µl and combine with 10 µl SDS-sample buffer
(2x).
• Heat all samples for 5 minutes to 95°C. Spin briefly to collect all liquid at the bottom of each
tube.
• Load pre-cast gradient SDS-PAGs with 10 µl of each sample according to the following
scheme and run at 20 mA (constant current) in SDS running buffer:
• Groups1, 2, 5, 6 only: Add 1 µl of His-tagged Sumo protease His6-SenP2 (SP) to your
eluate. Also: Take 5 µl SenP2 and combine with 5 µl SDS-sample buffer (2x).
• All groups: Using a 18-gauge needle and a 1 ml syringe, quantitatively transfer your eluate
into a dialysis unit (cut-off: 3,5 kDa) pre-wetted in cold dialysis buffer (Fig. 5). Dialyse for at
least 3 hours (or over night) at 4°C while stirring. Change buffer after 1 hour.
Figure 5: How to use “slide-a-lyzers”.
• Re-collect your eluate (Fig. 5) and estimate the volume of your final retenate (FR).
Remove any precipitated protein by centrifugation (1 minute, full speed). Take out 10 µl and
combine with 10 µl SDS-sample buffer (2x).
• Save some of the dialysis buffer as a reference (negative control in later experiments)!
• For its later use in egg extract and micro-injection experiments: Aliquot your final retenate,
snap-freeze in liquid nitrogen (wear eye protection!) and store at -20°C.
• Heat all SDS samples for 5 minutes to 95°C. Spin briefly to collect all sample at the bottom
of each tube.
• Load pre-cast gradient SDS-PAGs with 10 µl of each sample according to the following
scheme:
All gels:
• When bromophenol blue front reaches the bottom of the gels: Stop electrophoresis,
disassembly plates, Coomassie-stain, and then destain the PAGs.
• Put gels on white light box and take a picture with a digital CCD camera system (LAS3000,
Fuji) according to your tutor’s instructions.
Solutions provided:
• 10x PBS (0.2 M sodium phosphate buffer pH 7,4; 1.5 M NaCl)
• 5 M NaCl
• 1 M imidazole
• 14,4 M β-mercaptoethanol (toxic!)
• elution buffer (1x PBS, 200 mM imidazolec(in high concentrations imidazole acts like a
base!), 5 mM β-mercaptoethanol, pH re-adjusted to 7,0 with HCl)
• dialysis buffer (10 mM Hepes/KOH, pH 7,0; 80 mM KCl; 0,5 mM DTT)
• His6-SenP2 (1 mg/ml in dialysis buffer)
• Coomassie stain (0.25% Coomassie Blue R-250 in 50% methanol, 10% acidic acid)
• Coomassie destain (30% methanol, 7% acidic acid)
• SDS running buffer (25 mM Tris base; 210 mM glycine; 0,1% SDS)
• 4x SDS sample buffer (240 mM TrisHCl pH 6,8; 40% glycerol; 8% SDS; 0.04%
bromophenol blue; 5% (0,72 M) β-mercaptoethanol)
ad 2) IN VITRO EXPRESSION
OF A FLUORESCENTLY TAGGED PROTEIN
Traditionally, eukaryotic or prokaryotic in vitro
translation systems in rabbit reticulocyte lysate, wheat
germ extract or the E. coli S30 extract system
capitalize on the addition of radioactive amino acid(s).
In contrast, the FluoroTect-Green-Lysin-system
(Promega) allows the co-translational fluorescent
labeling of proteins through the use of a modified
charged lysine transfer RNA labeled with the
fluorophore BODIPY-FL, which has an absorbance
maximum of 502nm and an emission maximum of
510nm (Figure 6). Synthesized proteins are resolved
by conventional SDS-PAGE followed by fast, direct
Figure 6: FluoroTect GreenLystRNA from Promega.
fluoroimaging of the non-radioactive gel.
You will produce fluorescently labeled securin by coupled in vitro transcription/
translation (IVT) in wheat germ extract. On the corresponding plasmid, the expression of
securin is driven by a strong promotor recognized by viral SP6-RNA polymerase. To increase
the signal intensity and, hence, sensitivity of detection, the otherwise wild type ORF of
securin was extended at its 3’end to code for additional eight Lys residues. Experience has
shown that C-terminal tags (in contrast to N-terminal tags) do usually not interfere with the
degradation of APC/C-substrates via the UPS. Later in the course, you will therefore use your
IVT to follow APC/C activity in a Xenopus egg extract.
Experiment:
• Pour 15% SDS-PAG.
• Given that from an added
expression plasmid an
sensitive mRNA is transcribed in the first step, work
as careful as possible to
avoid contamination of your
samples with RNAses, e.g. wear gloves, use sterile pipette tips from a box (not a bag), sterile
1,5 ml tubes,…
• Add 2,5 µl of the securin-Lys8 encoding expression plasmid (0,2 mg/ml) into a fresh 1,5 ml
tube and store on ice.
• In a 2nd 1,5 ml tube, assemble the master mix on ice by combining the following:
12,5 µl
wheat germ extract
1 µl
TNT reaction buffer
1 µl
FluoroTect GreenLys tRNA
0,5 µl
amino acids without methionine
0,5 µl
amino acids without leucine
6 µl
H 2O
0,5 µl
SP6-RNAPol.
0,5 µl
RNase inhibitor (RNasin)
• Mix by carefully pipetting up and down and then, using the same pipette tip, transfer 17,5 µl
into the prepared tube with the securin plasmid and mix again. Label your sample with FS for
“fluorescent securin”. Save the remainder of the master mix – this is your negative
control (NC).
• Incubate both tubes at 30°C for 90 minutes.
• Take out 1 µl each and combine with 10 µl SDS sample buffer (1x)
• For its later use in egg extract: Snap-freeze your FS in liquid nitrogen (wear eye protection!)
and store at -20°C.
• Heat both SDS samples: 5 minutes, 95°C.
Spin briefly.
• Load 10 µl each onto self-prepared 15%
SDS-PAGs according to the scheme on left.
• Store the remainder of the samples at -20°C.
• When bromophenol blue front reaches the
bottom of the gels: Stop electrophoresis and
perform fluoroimaging with a digital CCD
camera system (LAS4000, Fuji) according to
your tutor’s instructions.
ad 3) ISOLATION OF XENOPUS TESTES AND SPERM NUCLEI
• Prepare the following buffers and store on ice: • 50 ml of 1x MMR; • 2 ml of 1x XN buffer
with 50% glycerol; • 100 ml of 1x XN buffer.
• Using a container with a closing lid, prepare 1 liter of 0.05% benzocaine (ethyl-paminobenzoate) in tab water (why not de-ionized?).
• Put 2 male frogs into this benzocaine solution; they will be dead after 15 minutes.
• Put a petri dish on ice and fill it with 1xMMR.
• Put each frog on its back on top of saran wrap-covered ice. Pull its skin up with a pair of
forceps and slice the skin with a scalpel. “Undress” the frog by making a longitudinal and a
transversal cut. On one side of the frog’s body: Pull up the muscles with a pair of forceps and
carefully cut a slit with a scalpel to get excess to the abdominal cavity. Pull out the fat body.
The white, oval-shaped testis is attached to its end. Cut off the testis with a pair of scissors,
leaving a small piece of fat body attached to it. Put it into the dish with cold 1xMMR. Do the
same on the other side and with the second frog.
• Put one testes in a 50 ml conical tube filled with 5 ml of 1xMMR containing 50 µg/ml
gentamycin. Store at 4°C for later in vitro fertilization (IVF) experiments.
• The other three testis are used to prepare permeabilized sperm nuclei according to the
following protocol:
• Transfer testes into petri dish filled with ice cold 1x XN buffer. Using fine forceps and
scissors (expensive tools! please take care!), carefully remove as much of fat body and blood
vessels from the testes as possible. It might help you to do this under a binoccular
microscope.
• Transfer “cleaned” testes into petri dish filled with 4 ml of fresh, ice cold 1x XN buffer. Using
two forceps like knifes, mascerate all three testes until all clumps are gone (be patient and
persistent).
• Carefully, pipette sperm suspension through a 100 µm mesh into a cut-off 15 ml conical
pre-chilled on ice. Rinse petri dish and mesh with more 1x XN buffer and into the same tube
to minimize loss.
• 1st spin: 5 minutes at 4.500 rpm in a JS-13.1 rotor (Beckman) at 4°C.
• Remove the supernatant.
• With a 1 ml pipette and 2 ml of 1x XN buffer: Resuspend the white sperms on top but not
the red blood cells at the bottom by carefully pipetting up and down. Then, transfer the
suspension into 2nd cut-off 15 ml conical pre-chilled on ice. Save the blue tip!
• 2nd spin: Add 8 ml 1x XN buffer and then spin 5 minutes at 4.500 rpm in a JS-13.1 rotor
(Beckman) at 4°C.
• During the centrifugation: Resuspend the blood cell pellet at the bottom of the 1st conical
with 1 ml XN-buffer and discard. The rinsed 1st conical will be re-used in the next step.
• After the centrifugation: Remove the supernatant. Then, using the same blue tip (see
above), carefully resuspend the white sperms in 2 ml 1x XN buffer leaving behind the red
blood cells at the bottom. Then, transfer the suspension into the rinsed 1st conical.
• 3rd spin: Add 8 ml 1x XN buffer and then spin 5 minutes at 4.500 rpm in a JS-13.1 rotor
(Beckman) at 4°C.
• During the centrifugation: Resuspend the blood cell pellet at the bottom of the 2nd conical
with 1 ml XN-buffer and discard. The rinsed 2nd conical will be re-used in the next step.
• After the 3rd centrifugation: Remove all supernatant. Then, using still the same blue tip (see
above), carefully resuspend the white sperms in 2 ml 1x XN buffer leaving behind the red
blood cells at the bottom. Transfer the suspension into the rinsed 2st conical.
• Add 0,4 ml of 2 mg/ml lysolecithin in 1x XN and incubate for 1 hour on ice. Using your good
old friend (the blue tip) mix occasionally by pipetting up and down.
• Quench the reaction by addition of 8 ml 1x XN buffer with 3% BSA.
• 4th spin: 5 minutes at 4.500 rpm in a JS-13.1 rotor (Beckman) at 4°C.
• Wash 3 more times as before (once with 1x XN buffer, 3% BSA and twice with just plain 1x
XN buffer).
• After the 7th and last spin: Remove the supernatant and thoroughly resuspend (guess with
who?) the permeabilized sperm in 0,5 ml of 1x XN buffer, 50% glycerol.
• Determine sperm density: Take out 1 µl and thoroughly mix with 99 µl of Hoechst 33342
solution (mutagenic!).
• Using a hemocytometer (see appendix, figure 12) and a fluorescence microscope: Count
the sperm and calculate the concentration of sperm in your undiluted preparation. Typical
densities are between 5x107 and 5x108 sperms per ml.
• Aliquot your sperm suspension and store at -80°C.
Solutions/reagents provided:
• 10% (w/v) benzocaine stock solution in ethanol (= 200x)
• 25x MMR (2,5 M NaCl; 50 mM KCl ; 25 mM MgCl2; 50 mM CaCl2, 2,5 mM EDTA/NaOH, pH
8,0; 125 mM HepesNaOH, pH 7,8; titrated to pH 7,8 with NaOH)
• 1xMMR containing 50 µg/ml gentamycin.
• 2x XN buffer (100 mM HepesKOH, pH 7,0; 500 mM sucrose; 150 mM NaCl; 1 mM
spermidine; 0,3 mM spermine)
• 1x XN buffer with 2 mg/ml lysolecithin
• 1x XN buffer with 3% BSA
• Hoechst 33342 solution (1µg/ml in 1x XN; mutagenic!)
• 100% glycerol
ad 4) PREPARATION OF CSF-EXTRACTS FROM XENOPUS EGGS AND THEIR USE TO
STUDY EXIT FROM MEIOSIS II/MITOSIS
A multitude of complex cellular processes, e.g. nuclear transport and replication, can be
studied in extracts from oocytes of the African clawed frog, Xenopus laevis. The strength of
this cell-free system is that it is amenable to biochemical manipulations. A protein can easily
be removed by immuno-depletion and replaced by recombinant versions to study the effect
of certain mutations on a given process, for example. These extracts are also capable of
autonomously undergoing several rounds of a rapid cell cycle (Figure 7), which consists just
of alternating S and M-phases and is typical for early embryos. Much of our current
understanding of the cell cycle was elucidated by using this powerful model system. In
response to an injection of chorionic gonadotropin on the previous day, female frogs lay
eggs, which are arrested in metaphase of the (mitosis-like) meiosis II due to presence of a
cytostatic factor. This CSF, which is also called XErp1 or Emi2, was recently shown to
represent an APC/C-inhibitor and to become degraded in response to a transient rise in
intracellular Ca2+ upon fertilization (Figure 2).
In this pratical course, we will prepare extract under conditions that preserve the CSFimposed arrest. We will then add non-degradable forms of cyclin B1, XErp1 or reference
buffer and study the effects of these supplements on APC/CCdc20 activity and mitotic/meiotic
exit. Before we mimic fertilization by Ca2+ addition, the CSF-extract will also be supplemented
with fluorescently labeled tubulin and sperm chromatin (which is later stained with the
intercalating, fluorescent dye DAPI). This allows us to study by fluorescence microscopy the
state of the microtubule cytoskeleton and of chromosome condensation, both reliable
indicators of mitotic/meiotic versus interphase state. As the haploid, one-chromatidchromosomes of the sperm are no suitable indicator of anaphase, we observe the
degradation of securin instead. To this end, we add to the egg extract in vitro-expressed,
fluorescently labeled securin, the APC/CCdc20-degradation of which can be followed by SDSPAGE and direct fluorescent imaging of the gel.
Figure 7: (A) Schematics of generation and uses of oocyte extracts (B) In vitro cell cycles (DNA in blue,
Microtubules in red).
Experiments:
• prepare 400 ml of CSF-XB:
1x XB-salts; (+ additional) 1 mM MgCl2; 5 mM EGTA/KOH, pH 8,0; 50 mM sucrose; 10 mM
HepesKOH, pH 7,7; check pH & adjust to 7.7 with KOH
• prepare 100 ml of dejellying solution:
2% (w/v) cysteine in 0.5x XB-salts; adjust pH to 7,8 with KOH; use within 1h
• Add 0,5 ml CSF-XB and 5 µl cytochalasin B to the bottom of an appropriate spinning tube.
Mix.
• Collect eggs and remove „bad“-looking ones according to your tutors instructions.
• Remove all liquid and add around 50 ml of dejellying solution. Immediately, remove liquid
and add remainder of dejellying solution. Swirl once, then let sit until dejellying process is
complete (eggs pack tightly due to loss of jelly coat). The dejellying process should
maximally take 10 minutes.
• Eggs are now sensitive against mechanical forces and should always be covered with
liquid!
• Wash eggs 4x in CSF-XB. (The cytotoxic dejellying solution must be removed completely.)
• Remove all activated (contracted, small, black animal pole), lysed, and odd looking eggs!
• Carefully transfer the eggs into the prepared spinning tube using a large-bore pipette (cutoff Pasteur, fire-polished). Pipette eggs directly into liquid at the bottom. Avoid buffer transfer
as much as possible.
• Remove all supernatant and add 100 µl of Nyosil M25 on top.
• For centrifugation: Use the provided adaptors and the JS13.1 rotor and centrifuge at 18°C.
• Packing spin: 1 minute at 200 g and 1 minute at 600 g.
• Carefully remove all supernatant!
• Crushing spin: 10 minutes at 13.000 g (equivalent to 9.000 rpm).
• Recovery of cytosolic extract (middle part of tube): Puncture the tupe with an 18-gauge
needle on a 1 ml syringe just above the dark cell debris (bottom phase). Suck out yellowish
cytosol avoiding the other layers. Take off the needle before ejection of the extract into a
fresh, pre-cooled 1,5 ml tube
• Estimate the volume of the extract and add sperm nuclei to 2000 per µl as well as
cytochalsinB and cycloheximide to 1x. Mix your extract carefully but thoroughly.
• To a fresh, pre-chilled tube: Add 15 µl of your fluorescently labeled securin and 1 µl
rhodamine-labeled tubulin. Then, add 184 µl CSF-extract and simultaneously mix by slowly
pipetting up and down several times. Take out 1 µl and combine with 20 µl SDS-sample
buffer in a tube labeled -15’. Also, on a microscope slide combine 1 µl of this extract with 2 µl
of DAPI/Fix and cover the sample with a cover slip.
Figure 8: Pipetting scheme for egg extract experiment. On each level (-15, 0, and 40 minutes) two samples are
taken, one for SDS-PAGE/fluoroimaging and one for fluorescence microscopy.
• To 3 fresh, pre-chilled tubes labeled „C“, „X“ and „R“, respectively: Add 1 µl of either nondegradable Cyclin B1 or non-degradable XErp1 or Reference (dialysis) buffer. Then, transfer
59 µl from the supplemented CSF-extract into each of these three tubes. Again, mix by
slowly pipetting up and down several times. Finally, transfer all three tubes from ice to room
temperature. (Do not put back on ice at any time for the remainder of the experiment!)
• Into 3 tubes labeled 40’C-, 40’X-, 40’R- pipette 1 µl each of sperm dilution buffer. Into
another 3 tubes labeled 40’C+, 40’X+, 40’R+ pipette 1 µl each of 25x CaCl2. (Do not cool
tubes.)
• 15 minutes after starting your extract incubations at room temperature:
From all three tubes „C“, „X“ and „R“, take out two 1 µl-samples for SDSPAGE and fluorescence microscopy as described above. Suggested
labeling: 0’C, 0’X, and 0’R.
• Release from CSF arrest by Ca2+ addition: Now, transfer 24 µl from „C“
into 40’C- and mix by pipetting. Start a timer. Start another sample every
minute, i.e. 24 µl from „C“ into 40’C+ after 1 minute, 24 µl from „C“ into
40’C- after 2 minutes, .... (6 samples in total).
• After 40 minutes: In the same order, in which you started your samples,
prepare two samples each as described above – one for SDS-PAGE and
one fluorescence microscopy.
• Heat your SDS-samples and resolve 10 µl each on a re-cast gradient SDS-PAG (see
scheme above) followed by fluoroimaging as before.
• Fluorescence microscopy: Inspect your slides with a 20x or 40x lens in DAPI and
rhodamine channels and take pictures for the protocol.
Figure 9: Examples of freshly added sperm, mitotic/meiotic chromatin and interphase nucleus (blue) associated
with microtubules (red) in egg extract.
By filling in (+/-) the above table, document for all extracts whether bright red microtubuleasters/spindles are associated with condensed chromatin or whether long, hair-like
interphase microtubules surround nuclei containing decondensed chromatin. (Please note:
Sometimes the release from mitosis into interphase is not homogenous, so that some mitotic
figures persist; however, the presence of any nuclei is always a good indication of a
successful release.)
solutions/proteins provided:
• 20x XB-salts (2 M KCl; 2 mM CaCl2; 20 mM MgCl2)
• 5 M KOH (-> wear protective glasses when handling bases!)
• 1 M HepesKOH, pH 7,7
• 1 M MgCl2
• 0,5 M EGTA-KOH, pH 8,0
• 2,5 M sucrose
• 1.000x Cytochalasin B (10 mg/ml in DMSO; expensive! toxic!)
• Nyosil M25
• rhodamine-labeled tubulin (expensive!)
• stable cyclin B1 (D-Box removed by N-terminal deletion)
• Stable, constitutively active XErp1-fragment (prepared by yourself)
• FluoroTect-labeled securin (prepared by yourself)
• 25x CaCl2 (15 mM in sperm dilution buffer)
• Sperm dilution buf. (5 mM HepesKOH pH 7,7; 150 mM sucrose; 100 mM KCl; 1mM MgCl2)
• DAPI/Fix (1x MMR; 48% glycerol; 11% formaldehyde; 1 mg/ml Hoechst 33342; toxic,
mutagenic!)
• 100x Cycloheximide (10 mg/ml in H2O; toxic!)
• 4x SDS-sample buffer
ad 5) ANALYSIS OF CELL CYCLE REGULATORS BY MICRO-INJECTION EXPERIMENTS
INTO XENOPUS EMBRYOS
Apart from the powerful egg extracts, the model system Xenopus has even more to offer.
Freshly spawned eggs can be fertilized in vitro and the resulting embryos can easily be
micro-injected due to their enormous size. Next to developmental studies, this enables
researchers to study the cell cycle during early embryogenesis. To this end, usually one half
of a 2-cell embryo is injected, with the other cell serving as an internal control. You will
conduct an IVF and then micro-inject either mRNA coding for non-degradable cyclin B1, your
self-made XErp1 protein, or a reference buffer into the resulting 2-cell embryos. You will
follow the subsequent cleavages by video microscopy and analyse the effects of your
manipulations.
Experiments:
• Injection of female frogs to induce ovulation: 20 h before the experiment inject female frogs
with 0,8 – 1,0 ml (= 500-900 I.U.) of human chorionic gonadotropin (hCG) into the dorsal
lymph sac using a 27-gauge needle. This will be done by your tutor.
• Your tutor will also squeeze eggs from a frog into a 5.3 cm petri dish filled with 1xMMR. To
this end he/she will hold the frog by grabbing it from above with your forefinger between the
legs and by covering the head/eyes between his/her palm and pinky finger. Most frogs will
squeeze out their eggs all by themselves if you just hold them. However, sometimes it might
be necessary to gently exert some pressure by moving your fingers laterally and ventrally
over the abdomen. Some frogs secrete copious quantities of a milky exudate from their skins:
Dripping of this slime into the dish should be avoided because eggs tend to lyse in its
presence. Do fertilization within 15 minutes after squeezing the eggs.
• Fertilization: Remove most of the 1x MMR. Then take the testes, which you have isolated,
and by grapping on to the attached partial fat body, bring it into the petri-dish. Using tweezers
like knives, macerate a little piece of the testes beginning from its tip. Thoroughly mix the
cloud of liberated sperm with the eggs by grapping on to the testes and vigorously moving it
around in the dish. Repeat this 3 more times. Then, fill dish all the way with 0.1x MMR and
leave for 20 minutes. This treatment ensures that the eggs get fully activated.
Activation/fertilization causes a contraction of the pigmented region of the cortex, and the
white spot at the center of the pigmented region becomes less distinct. Also, all the eggs will
turn so that the pigmented region is facing up. During this incubation…
• Prepare 50 ml of dejellying solution. To this end, a solution of 2% cysteine in H2O is adjust
with 5 M NaOH (wear eye protections!) to a pH of 7,8. Use within one hour.
• Following the 20 minute incubation of the fertilized eggs: Remove as much of the 0.1xMMR
as possible and replace with fresh dejellying solution. Repeat this procedure once more to
ensure complete exchange of buffers. Incubate about 5 minutes until the jelly coats are gone
and the embryos pack tightly together. Zygotes are now sensitive against mechanical forces
and should always be covered with liquid! Wash 6 times with 0.1x MMR to fully remove
cytotoxic cysteine. Pick lysed or strangely pigmented zygotes with a Pasteur pipette and
discard. Finally, transfer embryos into fresh petri dishes filled with 0.1x MMR by using a wide
diameter Pasteur pipette. Transfer as little buffer as possible at this step. Incubate aliquots of
embryos at different temperatures (e.g. 22, 18 and 14°C) for 80 to 120 min. This will later
allow you to inject two-cell stage embryos over a longer period of time. During this time...
• Pull your own needles for micro-injection on a capillary puller together with your tutor. Wear
gloves and do touch needles only at the ends to keep the middle part (which will become the
tips of two new needles when the capillary is pulled apart) RNAse-free. Carefully place the
needles onto a styrofoam support in a large petri dish and close the lid.
• Prepare a needle for injection: Carefully put a needle on top of a parafilm under the stereomicroscope. At 50x magnification, break off the tip of the needle using a scalpel. The outer
diameter of the needle should be below the distance between two pitch lines of the
graduation (= 15 µm at 50x magnification). Try to break the needle so that you will get a
pointy end, which is indicated by its bright reflection of light. Put the needle with its blunt end
into the tubing of the injector and screw the needle gently into the holder of the
micromanipulator. Wear gloves and work as RNAse-free as possible.
• Open on the “compressed air” outlet that connects to the PLI-100 injector. Put the “pressure
meter source”-knob in the Pclear position. The reading should be between 60 and 90 psi and
stay fairly constant. Put the “pressure meter source”-knob in the Pinject position and adjust the
reading to 10 psi as a starting point (between 5 - 15 psi is OK). Set the injection time to 300
msec (between 300 - 500 msec is OK).
• Before putting a petri dish under the microscope (or removing one) always remember to
retract the needle fully so that you will not brake the tip!
• Calibrate the needle: Through the tip, suck up RNAse-free H2O from a petri dish to fill the
needle by pressing the “fill” button. Put a petri dish filled with mineral oil under the
microscope. Move the tip of the needle into the oil while watching it at 50x magnification.
Inject H2O into the oil by pushing the “inject” button. Using the graduation of the eyepiece,
estimate the diameter of the droplet; it should be around 12 pitch lines at 50x magnification.
This size of drop corresponds to an injection volume of about 10 nl. If the droplet is too small
or too big increase or decrease, the injection time and/or pressure. If you need too long long
injection times and/or too high pressure, break off the tip a little more.
• Adjust the balance pressure: Put the “pressure meter source”-knob in the Pbalance position.
Adjust the balance pressure to the lowest value where you just see cords of water being
slowly ejected into the oil. This should be the case at – 0.5 to 1.0 psi. The balance pressure
avoids sucking of liquid into the needle due to capillary action. Wash needle free of oil. For
rapid ejection: Press the “clear” button.
• Each group should inject two groups of 2-cell embryos. You may choose between cyclin B1
mRNA and H2O or XErp1 protein and the corresponding dialysis buffer. One partner may
inject mRNA/protein while the other injects reference buffer. Alternatively, each person may
change the content of her/his needle. When you use the same needle for two types of
injections, then start with the negative control, i.e. the reference solution.
• Loading the needle with sample: Put 2 µl of your sample as a small droplet onto a piece of
parafilm. Set the magnification to 12x. Focus onto the top of the drop and center the drop in
the middle of your visual field by moving the parafilm. Using the micromanipulator, position
the tip of the needle to the top surface of the droplet by just using the forward screw.
Submerse the tip of the needle just below the surface of the liquid and press the fill pedal.
RNA is typically used at a concentration of 2 to 200 ng/µl (in H2O), which corresponds to
roughly 2 pg to 2 ng per injection. When injecting protein: Use as much as possible but do
not inject more than 20 nl per cell.
• Actual Injection: At 12x magnification,
transfer 2-cell stage embryos into the
center of a petri dish filled with 0.1x
MMR, 5% Ficoll. Using a blue pipette tip
(or forceps) in your left hand, move a
nice looking embryo in front of your
needle tip. Focus onto the embryo’s
surface. Move the needle carefully
forward and hold the embryo in place
with the blue pipette tip (or forceps) from
the opposite side (Fig. 10). Then, quickly
push the needle into the darkly
pigmented region at the upper third of
the embryo and immediately retract a Figure 10: Typical images illustrating the injection process
little bit to ensure injection close to the (bottom) and the result of microinjection of a cell cycle inhibitor
cell surface. Instantly, press the “inject” (top right).
foot pedal and then pull the needle out
of the embryo. All these actions should be coordinated into one quick maneuver. You should
see a whitening of the cell surface at the side of injection due to displacement of dark
pigment. Move the injected embryo to an empty area of the dish. Repeat this injection
procedure many times. Every now and then, transfer injected embryos into new petri dish
with 0.1x MMR (avoid transfer of Ficoll) and add new, uninjected embryos to your dish.
• After injection: Film the two groups of injected embryos next to each other over a time
period of three hours at 18°C. You may also fix interesting looking embryos by incubation in
DAPI/Fix (see 4).
ad 6) MICROSCOPY OF FLUORESCENTLY-LABELED MARKER PROTEINS IN LIVING
CELLS
Microscopy of living cells is crucial for the investigation of dynamic processes. Fusing marker
proteins with fluorescent proteins has revolutionized this field because it allows visualization
of specific regions of interest within the cell. In the lab course we will, for example, film
modified HeLa cells (cervix adeno-carcinoma), which have their chromosomes and
microtubule cytoskeleton labeled by stable expression of histone2B-RFP and α-tubulineGFP, respectively. To this end, we will use a BioStation IM (Nikon; Fig. 11), which combines
an incubator, a microscope and a cooled CCD camera in a compact body, allowing
multichannel (RFP and GFP fluorescence plus phase contrast), multipoint (thanks to a
motorized objective lens) time-lapse imaging.
Figure 11: Components (left) and incubation chamber on top of the optics (right) of the Biostation from Nikon.
• In the morning: Trypsinize the cells from an existing plate according to your tutor’s
instructions and seed them into a four-chambered slide for later imaging.
• In the evening: Replace the medium with one containing a) monastrol, b) staurosporine, c)
cytochalasin B, and d) DMSO (the solvent for the three small molecules = control).
• Your tutor will help you to then start your time-lapse imaging over night at 37°C and 5%
CO2.
• The next morning: Stop the imaging and transfer the aquired data to a DVD for statistical
analysis.
ad 7) CHARACTERIZATION OF ISOGENIC AND INDUCIBLE TRANSGENIC CELL LINES
BY FLOW CYTOMETRY, CHROMOSOME SPREADING, AND WESTERN BLOT
It has been a dogma in the field that the pathways that lead to anaphase and mitotic exit are
strictly separated downstream of the common trigger APC/CCdc20. However, recent
investigations have shown that there is cross-talk between Cdk1 and separase in
vertebrates. More specifically, separase can be phosphorylated and then stably bound and
inhibited by Cdk1. This alternative mechanism of separase inhibition nicely explains why
vertebrate securin is fully dispensable for timely segregation of chromosomes with high
fidelity.
In the lab course, you will analyze transgenic Hek293 (transformed human embryonal
kidney) cell lines, which inducibly over-express either wild type (WT) separase or a Cdk1-
resistant phosphorylation-site mutant (PM). In particular, you will apply the techniques of
chromosome spreading and flow cytometry to compare the effects that these two
recombinant separases might have on sister chromatid cohesion and the cell cycle profile.
• Cell culturing: Split transgenic Hek293 cells according to your tutor’s instructions to
generate four 10 cm plates per group, each of which should exhibit a density of ≈70% the
following day.
Groups 1 – 4: Prepare four 10 cm plates of „PM“ cells.
Groups 5 – 7: Prepare two 10 cm plates of „WT“ and two 10 cm plates of „PM“ cells.
• At the evening of the next day add tetracyclin and/or nocodazole according to the following
scheme:
Groups 1 – 4: - add 5 µl of tetracyclin to plate 1 („PM/I“ with „I“ for induced)
- add 5 µl of ethanol to plate 2 („PM/U“ with „U“ for uninduced)
- add 5 µl of tetracyclin and 0,5 µl of nocodazole to plate 3 („PM/I/N“)
- add 5 µl of ethanol and 0,5 µl of nocodazole to plate 4 („PM/U/N“)
Groups 5 – 7: - add 5 µl of tetracyclin to all plates („PM/I“ and „WT/I“)
then, also: - add 0,5 µl of nocodazole to plate 3 („PM/I/N“)
- add 0,5 µl of nocodazole to plate 4 („WT/I/N“)
• Prepare the following solutions: • 10 ml of hypotonic medium (40% D-MEM; 60% de-ionized
water; 500 ng/ml nocodazole); • 10 ml of Canoy’s Solution (methanol:acetic acid = 3:1,
toxic!); • 15 ml PBS-B (1x PBS; 0,2% BSA); • 50 ml of blocking buffer (5% milk powder in
PBS-Tw; store on ice).
• Flow cytometry: All groups harvest all their cells from plates 1 and 2 by trypsination and
centrifugation in 15 ml conicals according to the tutor’s instructions.
• Cells are washed once with 10 ml of 1x PBS (i.e. resuspended and re-pelleted).
• After aspiration of the supernatant: Resuspend cells in 200 µl of 1x PBS.
• For later Western analysis: Combine 20 µl cell suspension with 80 µl of 1x SDS sample
buffer.
• Fixation: To the residual ≈150 µl of cell suspension: Add 8 ml of 70% ethanol pre-cooled to
-20°C while vortexing. Store samples at -20°C as long as necessary.
• PI-Staining: Spin down ethanol-fixed cells at 1.500 rpm for 10 minutes in a clinical
centrifuge.
• Resuspend cells in 10 ml PBS-B and pass suspension through 70 µm cell strainer. Recentrifuge.
• Resuspend cells in 0,75 ml propidium iodide solution and incubate for 1h at 37°C.
• With the help of your tutor: Perform flow cytometry at the Beckman FC-500.
• Chromosome spreading: To harvest nocodazole treated cells, tab the corresponding plates
3 and 4 to dislodge even the last attached cells. Resuspend by pipetting up and down and
then transfer cell suspension into 15 ml conical. Rinse plate with 5 ml 1x PBS to minimize cell
loss. Pellet cells by centrifugation for 3 minutes at 300 x g in a clinical centrifuge.
• Wash cells once in 5 ml of 1x PBS and re-centrifuge as before.
• Swelling and fixation of cells for spreading: Resuspend the cells in 200 µl hypotonic
medium by carefully pipetting up and down.
• For later Western analysis: Combine 20 µl cell suspension with 80 µl of 1x SDS sample
buffer.
• After a 3 minute incubation, another 300 µl and then 2 ml of hypotonic medium are added to
the cell suspension. (To minimize cell loss, mixing in these two steps is achieved by zestful
addition rather than pipetting up and down.) Following a 5 minute-incubation at room
temperature, swollen cells are gently pelleted at 100 x g for 5 minutes and resuspended in 20
µl of hypotonic medium by flicking the tube.
• Then, 250 µl, 250 µl, and 2 ml of Canoy’s Solution are added stepwise at room
temperature. (Again: To minimize cell loss, mixing in these three steps is achieved by zestful
addition of the fixative rather than pipetting up and down. Do not flick or vortex tube because
cells will stick to walls.) After incubation for 15 - 30 minutes at room temperature, cells are
dehydrated further by washing two times with 1 ml Canoy’s Solution (centrifugations at 300 x
g for 4 minutes). Finally, the cell pellet is resuspended in 250 µl Canoy’s Solution. At this
stage, samples may be stored at -20°C until further use.
• Spreading: Drop 15 µl of cell suspension (in two aliquots, approx. 2 cm apart from each
other) onto a 24 x 60 mm cover slip, which has been cooled to 0°C on top of an icesubmersed metal block and moisturized by breath. Following proper spreading of the sample,
the cover slips are dried at 60°C on a metal block covered with a wet tissue.
• Chromosome staining by Hoechst (groups 1, 3, 5, and 7): After drying, specimen are
stained by incubating in Hoechst (1 µg/ml in 1x PBS) for 10 minutes. After 2 rounds of 1x
PBS-wash, salt is removed by submersion of the slide in de-ionized water. Finally, samples
are air-dried. Spread 20 µl of Mounting Medium on top of cover slip and carefully cover with
microscope slide, avoiding air bubbles.
• Chromosome staining by Giemsa (groups 2, 4, 6, and 8): After drying, lysed nuclei are
stained by incubating in Giemsa for 10 minutes at room temperature. Excessive Giemsa is
removed by dipping slide 3x into de-ionized water. Spread 20 µl of Mounting Medium on top
of air-dried slide and carefully cover with 24 x 60 mm cover slip, avoiding air bubbles.
• All groups: Determine the percentage of one- versus two-sister chromatid chromosomes in
each sample.
• Immunoblotting: Each group has 4 samples. Thus, two groups team up to run one pre-cast
gradient SDS-PAG. After electrophoresis, transfer onto PVDF membrane (remember to prewet with methanol!) by electro-transfer at at 12 V over night (or at 100 V for 1 hour and 4°C).
• Following transfer, Ponceau-stain for 3 minutes and then destain in water. Block unbound
binding sites on the membrane by rotating it in blocking buffer, cut the membrane in three
stripes. Incubate the top with anti-Myc, the middle with anti-tubulin, and the bottom with antiphosphoHistoneH3. Rotate for 3 hours at room temperature or over night at 4°C. Wash 3
times for 10 minutes with PBS-Tw, then incubate with anti-mouse-IgG-HRP for 1 hour. Wash
as before and develop while detecting with a chemiluminescence-sensitive CCD camera.
Solutions/proteins provided:
• tetracycline (5 mg/ml in ethanol; 2.000x; stored in dark at -20°C)
• nocodazole (5 mg/ml in water-free DMSO; 20.000x; toxic!)
• 10x PBS
• glacial acetic acid
• methanol
• 4x SDS sample buffer
• 70% ethanol (stored at -20°C)
• BSA (fraction V; stored in dry environment at 4°C)
• propidium iodide solution (69 µM in 38 mM sodium citrate; 100 µg/ml RNAse A; store in the
dark at 4°C; mutagenic!)
• Hoechst 33342 (1 µg/ml in 1x PBS; stored at -20°C; toxic!)
• Giemsa (To make, 380 mg Giemsa stain (Fluka 11700) were dissolved in 25 ml glycerol
and heated for 3 hours at 60°C. Then, 25 ml methanol were added. The solution was left to
stand for 5 days at room temperature and finally filtered.)
• Mounting Medium (0.5% p-Phenelynediamine (free base) in 90% glycerol, 20 mM Tris-HCl
pH 9.0; toxic!)
• Ponceau solution (1% PonceauS in 5% acetic acid)
• SDS running buffer (25 mM Tris base; 210 mM glycine; 0,1% SDS)
• 4x SDS sample buffer (240 mM TrisHCl pH 6,8; 40% glycerol; 8% SDS; 0.04%
bromophenol blue; 5% (0,72 M) β-mercaptoethanol)
• blot buffer (25 mM Tris base; 200 mM glycin; 20% methanol)
• PBS-TW (1x PBS; 0,1% Tween 20)
• various antibodies as ready-to-go solutions
• HRP chemiluminescence reagents
8) INSTRUCTIONS AND QUESTIONS FOR THE PROTOCOL
• Do not reproduce what is already written in the script, i.e. describe how you did something
only if it is missing in the above text (e.g. culturing of human cells) or if your actions deviated
from the script.
• Describe your observations and explain - especially, if they differ from your expectations.
• Show all your (properly labeled !) results, i.e. FACS profiles, images of gels, Xenopus
embryos, and fluorescence microscopy,…
Address the following questions (in some cases, this will require comparing your results to
those of other groups):
ad 1)
• Why is the binding to Ni2+-NTA done in presence of 5 mM imidazole and 0,55 M NaCl?
• Why is β-mercaptoethanol added during IMAC? Why only at this low concentration?
• Is one of the used E. coli strains more suitable for the expression of the short heterologous
His6-Sumo-XErp1(491-651) protein?
• Is the micro-fluidizer or the sonicator better for lysis?
• Was the tag successfully removed by SenP2 treatment? Did XErp1(491-651) remain
soluble? Why was imidazole and not His6-SenP2 used for elution?
• Suggest an additional purification step, by which one could remove the cleaved-off tag and
the Sumo protease from XErp1(491-651).
• Comparing your (His6-Sumo-)XErp1 signal in the final Coomassie gel to the SenP2 band of
known amount, estimate the overall yield of your purification.
ad 2)
• How do you think has the wheat germ extract been treated to ensure that only the mRNA
that is newly transcribed is translated?
• Why is it suboptimal to use fluorescently label Lys residues in our case? Why does
degradation nevertheless (usually) work?
ad 3)
• Why are spermine and spermidine in the XN buffer?
• What is lysolecithin and what does it do?
ad 4)
• Explain what you would expect for each type of extract in terms of securin degradation,
chromosome decondensation and spindle disassembly and why!
• Do your observations fit the theoretical expectations? - If not, discuss possible reasons!
• What is the purpose of cytochalasin B addition?
• What does cycloheximide do?
• Why does the CSF-XB buffer contain EGTA?
• Why is it important for the visualization of spindle-like structures to not put the extract on
ice?
• Why are the spindle-like structures stable even at 4ºC once you have prepared your slides?
• Sometimes two asters (microtubules associated with one sperm-genome) fuse to form
bipolar arrangements with the chromosomes aligned in their middle. Why do these structures
not represent real spindles but merely fusions of two half-spindles?
ad 5)
• Cysteine is a mild reductant. Can you guess what it reduces and why it can successfully be
used to remove the jelly coat?
• Categorize your injected embryos’ behavior into a) normal division, b) blocked division in
injected side only, and c) apoptosis/death. Do your statistics match with your expectations?
Explain!
ad 6)
• Describe the cultivation/splitting of human cells. What reagents/media did you use?
• In your movies: How many cells do you observe in each chamber, which a) divide normally,
b) arrest in mitosis with monoasters, c) undergo apoptosis, and d) fail to divide and become
tetraploid? Do your results/statistics match with your expectations? Explain!
ad 7)
• How many of which cells a) accumulate in G2/M (and why) and b) suffer from unscheduled
sister chromatid separation?
• For PM separase to prematurely resolve cohesion in Hek293 cells, it needs to be
overexpressed. Why?
• Why does overexpression of WT separase not trigger premature anaphase?
• Time-lapse movies of Tet-treated PM separase cells show that chromosomes still congress
in these cells but then loose cohesion (only) about 5 minutes too early, i.e. before anaphase
would usually commence. What do you conclude from this?
9) APPENDIX
Figure 12: (A) Prestained protein ladder (Fermentas). Do not boil! Use 5 µl per lane. (B) Hemocytometer.
Calculate your sperm density according to the following formula: Number of sperm per red square x 10.000 x
dilution factor = Numer of sperm per 1 ml.
Zellzyklus - Zeitplan, SS 2009, 27.04. - 08.05.
group
1-2
day
time
group
3-4
day
time
Mo
Tu
We
Th
Mo
Tu
We
splitting of
Hek293sep.
IMAC finish
dial.+ cleav.
snap freeze
cell fixation
for FACS
isolate
testis
egg extr.
experiments
egg extr.
microscopy
microscopy
of spreads
swell + fix
for spreads
isolate
sperm
SDS-Page
Fluoro, in-gel
Coomassie
Western finish
(tub.+ myc)
E.coli lysis
IMAC start
sec. in vitro
translation
sec. degrad.
RNAse + PI
(Fluoro, in-gel) for FACS
chromosome
spreading
Tet/Noc add.
to Hek293sep
SDS-Page
Western start
(tub.+ myc)
quantify
sperm
Mo
Tu
We
Th
Mo
E.coli lysis
IMAC start
IMAC finish
dial.+ cleav.
snap freeze
cell fixation
for FACS
isolate
testis
egg extr.
experiments
swell + fix
for spreads
isolate
sperm
sec. in vitro
translation
SDS-Page
Fluoro, in-gel
Coomassie
Western finish
(tub.+ myc)
Tet/Noc add.
to Hek293sep
SDS-Page
Western start
(tub.+ myc)
quantify
sperm
Tu
We
Th
splitting of
Hek293sep.
Mo
Th
Fr
splitting of
live-cell stop
HeLa-G/RFP movie analysis
flow
cytometry
in vitro
fertilization
evaluations
microinjection
drug add. to
HeLa-G/RFP
+
live-cell start
(BioStation)
discussion
Tu
We
Th
Fr
chromosome
spreading
in vitro
fertilization
microinjection
splitting of
live-cell stop
HeLa-G/RFP movie analysis
flow
cytometry
egg extr.
microscopy
evaluations
microscopy
of spreads
sec. degrad.
(Fluoro, in-gel) RNAse + PI
for FACS
Mo
Tu
We
drug add. to
HeLa-G/RFP
+
live-cell start
(BioStation)
discussion
Th
Fr