Molecular Biology of the Cell
Vol. 20, 3451–3458, August 1, 2009
Stathmin Regulates Centrosomal Nucleation of
Microtubules and Tubulin Dimer/Polymer Partitioning
Danielle N. Ringhoff* and Lynne Cassimeris†
Departments of *Chemistry and†Biological Sciences, Lehigh University, Bethlehem, PA 18015
Submitted February 17, 2009; Revised May 22, 2009; Accepted June 3, 2009
Monitoring Editor: Stephen Doxsey
Stathmin is a microtubule-destabilizing protein ubiquitously expressed in vertebrates and highly expressed in many
cancers. In several cell types, stathmin regulates the partitioning of tubulin between unassembled and polymer forms, but
the mechanism responsible for partitioning has not been determined. We examined stathmin function in two cell systems:
mouse embryonic fibroblasts (MEFs) isolated from embryos ?/?, ?/?, and ?/? for the stathmin gene and porcine kidney
epithelial (LLCPK) cells expressing stathmin-cyan fluorescent protein (CFP) or injected with stathmin protein. In MEFs,
the relative amount of stathmin corresponded to genotype, where cells heterozygous for stathmin expressed half as much
stathmin mRNA and protein as wild-type cells. Reduction or loss of stathmin resulted in increased microtubule polymer
but little change to microtubule dynamics at the cell periphery. Increased stathmin level in LLCPK cells, sufficient to
reduce microtubule density, but allowing microtubules to remain at the cell periphery, also did not have a major impact
on microtubule dynamics. In contrast, stathmin level had a significant effect on microtubule nucleation rate from
centrosomes, where lower stathmin levels increased nucleation and higher stathmin levels reduced nucleation. The
stathmin-dependent regulation of nucleation is only active in interphase; overexpression of stathmin-CFP did not impact
metaphase microtubule nucleation rate in LLCPK cells and the number of astral microtubules was similar in stathmin ?/?
and ?/? MEFs. These data support a model in which stathmin functions in interphase to control the partitioning of
tubulins between dimer and polymer pools by setting the number of microtubules per cell.
Microtubules (MTs) are dynamic polymers composed of
?/?-tubulin heterodimers that in cells exist at steady state
with a pool of unassembled tubulin dimers. In a nondiffer-
entiated cell, ?65% of tubulins are assembled into MTs and
35% are present as dimers (Zhai et al., 1996). This partition-
ing of tubulin dimers between polymer and dimer pools is
present in both interphase and metaphase of mitosis (Zhai et
al., 1996), although the lengths, number, and dynamic turn-
over rates of the MTs are very different between these two
cell cycle states (Desai and Mitchison, 1997). Cells are also
able to regulate the partitioning of tubulins between soluble
and polymer pools, for example, briefly shifting most tubu-
lins into the dimer pool at entry into mitosis (Zhai et al.,
1996) or shifting dimers into polymer form in T cells re-
sponding to signals downstream of the T cell receptor/CD3
complex (Holmfeldt et al., 2007).
Several potential mechanisms could regulate partitioning
of tubulin between dimer and polymer pools. First, shifts in
the plus end dynamic instability of MTs, favoring net as-
sembly or disassembly, could lead to changes in the parti-
tioning of tubulins. By this mechanism, changes in the level
or activity of microtubule-associated proteins regulating
various parameters of dynamic instability would alter tubu-
lin dimer partitioning. Such a shift in tubulins from polymer
to dimers was observed after depletion of microtubule-
associated protein (MAP) 4, an MT-stabilizing protein
(Nguyen et al., 1999). A second factor possibly regulating
dimer/polymer partitioning is the rate of MT nucleation
from centrosomes. Several computational models have sug-
gested that MT nucleation rate could have a significant
impact on tubulin partitioning within a cell (Mitchison and
Kirschner, 1987; Gregoretti et al., 2006), although this has not
been tested experimentally. Additional mechanisms that
could contribute to tubulin partitioning between dimer and
polymer pools include regulation of the total tubulin level or
sequestration of tubulin dimers to render them unable to
polymerize (Mitchison and Kirschner, 1987; Holmfeldt et al.,
2007; Sellin et al., 2008).
One MT regulatory protein recently demonstrated to con-
tribute to tubulin dimer/polymer partitioning is stathmin
(also named oncoprotein 18; gene name Stmn1), an 18-kDa
phosphorylation-regulated MT-destabilizing protein ex-
pressed ubiquitously in vertebrates (Steinmetz, 2007) and
overexpressed in a wide range of cancers (Brattsand et al.,
1993; Curmi et al., 2000; Mistry et al., 2005). The role of
stathmin in regulating tubulin dimer/polymer partitioning
was shown by overexpression or microinjection of stathmin
in vertebrate cells, which reduced MT polymer dramatically
(Larsson et al., 1999; Wittmann et al., 2004; Holmfeldt et al.,
2007), whereas microinjection of anti-stathmin antibodies
(Howell et al., 1999a) or short hairpin RNA-based depletion
of endogenous stathmin (Holmfeldt et al., 2006; Sellin et al.,
2008) led to increased MT polymer. Stathmin-dependent
regulation of MT polymer level is not confined to cells in
culture because stathmin knockout mice have more MT
polymer in the amygdala region of the brain compared with
wild-type mice (Shumyatsky et al., 2005).
This article was published online ahead of print in MBC in Press
on June 10, 2009.
Address correspondence to: Lynne Cassimeris (email@example.com).
Abbreviations used: MEF, mouse embryo fibroblast; MT, microtu-
bule; Stmn1, stathmin.
© 2009 by The American Society for Cell Biology3451
Stathmin could contribute to tubulin dimer/polymer
partitioning through several possible mechanisms. Each
stathmin molecule can bind and sequester two tubulin
dimers, thus preventing their polymerization (Howell et
al., 1999b; Steinmetz, 2007). A significant sequestering
function is possible in those cells where stathmin concen-
tration nears that of tubulin, such as Jurkat or K562 leu-
kemia cells (Sellin et al., 2008). In Jurkat cells (Sellin et al.,
2008) and Drosophila embryos (Fletcher and Rorth, 2007),
stathmin also contributes to regulation of the total tubulin
level, which may contribute to dimer/polymer partition-
ing. Stathmin also functions independently of tubulin
sequestration to stimulate MT dynamics by increasing
catastrophes at MT plus ends (Belmont and Mitchison,
1996; Howell et al., 1999b), which could also contribute to
tubulin dimer partitioning (Howell et al., 1999a).
Here, we use two cell systems derived from noncancerous
cells, which express relatively low levels of stathmin, to
explore how stathmin regulates tubulin dimer partitioning
in the absence of significant tubulin sequestering activity.
We isolated mouse embryonic fibroblasts (MEFs) from em-
bryos wild-type (stathmin?/?), heterozygous (stathmin?/?),
and null (stathmin?/?) for the stathmin gene. We also tran-
siently overexpressed stathmin in porcine kidney epithelial
(LLCPK) cells. For both cell types, we measured MT plus-end
dynamics and nucleation rates. Our results support a model in
which stathmin regulates tubulin partitioning by regulating
the rate of MT nucleation, rather than by regulating MT dy-
namics at the cell periphery.
MATERIALS AND METHODS
Isolation of MEFs
C57BL/6 Stmn1?/?male and female mice (a generous gift from G.
Shumyatsky, Rutgers University) were mated. Pregnant females were
killed by cervical dislocation 13.5 to 14.5 d after coitus, and fibroblasts
were isolated as described by Tessarollo (2001). Fibroblast cells from
individual embryos were plated and allowed to grow for 1–3 d before
storage of aliquots in liquid nitrogen.
Genotyping of MEFs was performed as described by Liedtke et al. (2002). In
brief, DNA was isolated from embryonic tissue by using isoamyl alcohol/
phenol extraction methods. Samples were amplified using polymerase chain
reaction (PCR) to identify embryos with intact Stmn1 intron III or the neo-
mycin cassette used to disrupt the Stmn1 gene (Schubart et al., 1996). The PCR
Jump Start REDTaq kit (Sigma-Aldrich, St. Louis, MO) was used for ampli-
fication; each sample contained 1.5 mM MgCl2, deoxynucleotide triphos-
phates (200 ?M each), primers (0.5 ?M each), and Taq polymerase (0.05 U/?l).
For the wild-type amplification, 35 temperature cycles were performed (95°C,
35 s; 66°C, 45 s; and 72°C, 45 s). Primers used for genotyping are listed in
Supplemental Table 1; these primers amplify either a portion of the region of
intron III deleted in the mutant allele or mutant-allele–specific primers that
recognize a portion of the neomycin insert (Schubart et al., 1996).
All cells were cultured at 37°C in a humidified atmosphere of 5% carbon
dioxide. Cells were grown in DMEM, pH 7.4, supplemented with 4.5 g/l
d-glucose, l-glutamine (Invitrogen, Carlsbad, CA), 44 mM sodium bicarbon-
ate, antibiotic/antimycotic (Sigma-Aldrich), 1 mM sodium pyruvate (Sigma-
Aldrich), and 10% fetal bovine serum (Invitrogen). LLCPK green fluorescent
protein (GFP)-tubulin (Rusan et al., 2001) and LLCPK GFP-EB1 (Piehl and
Cassimeris, 2003) cell lines were maintained as described previously (Piehl
and Cassimeris, 2003). MEF cultures were discarded after passage 7.
LLCPK GFP-?-tubulin cells were microinjected with bacterially expressed stath-
min-FLAG (130 ?M stock concentration; Holmfeldt et al., 2006) as described
of Alexa-Fluor 594-conjugated bovine serum albumin. Others have estimated
microinjectionvolumesof ?2.5–10%oftotalcellvolume(Graessmann etal.,1980;
Saxton et al., 1984; Howell et al., 1999a, 2000), indicating that intracellular con-
centration of stathmin-FLAG was ?3–13 ?M.
A plasmid for expression of stathmin-cyan fluorescent protein (CFP) was
constructed by amplifying the human stathmin cDNA from a pBS-STMN
plasmid (provided by Martin Gullberg) and introducing HindIII and BamH1
restriction sites by using primers listed in Supplemental Table 1. The reverse
primer also removed the stop codon. The HindIII/BamH1 fragment was
ligated into pECFP-N1 and pEGFP-N1 (Clonetech, Mountain View, CA). Cells
were transfected with plasmids encoding stathmin-GFP/CFP, GFP-tubulin,
or EB1-GFP as described previously (Piehl and Cassimeris, 2003; Warren et al.,
Fixed cells were analyzed by immunofluorescence as described previously
(Piehl and Cassimeris, 2003). Antibodies used included mouse monoclonal
antibody to ?-tubulin (B512; Sigma-Aldrich), rabbit anti-?-tubulin (Sigma-
Aldrich), mouse anti-EB1 (BD Biosciences, San Jose, CA), and goat-anti-mouse
or anti-rabbit Alexa Fluor 488 or 563 (Invitrogen). In some experiments, DNA
was also labeled with TO-PRO-3 iodide (Invitrogen).
Protein Isolation and Western Blot Analysis
Cell lysates or fractions from cytoskeletal and supernatant fractions were
prepared for SDS-polyacrylamide gel electrophoresis as described previously
(Graessmann et al., 1980; Saxton et al., 1984; Minotti et al., 1991; Howell et
al., 1999a). Protein concentrations of soluble fractions were determined by
Bradford assay (Bradford, 1976). Membranes were probed with anti-tubulin
antibodies DM1A or Tub2.1, or anti-stathmin (Sigma-Aldrich) followed by
goat anti-rabbit or mouse horseradish peroxidase-linked immunoglobulin G
(Sigma-Aldrich). Enhanced chemiluminescence (PerkinElmer Life and Ana-
lytical Sciences, Boston, MA) was used to develop immunoreactive bands
according to manufacturer’s specifications. For semiquantitative estimates of
stathmin and tubulin concentrations, band intensities from cell lysates were
compared with intensities for purified porcine brain tubulin (purified as
described in Vasquez et al., 1994) and probed with Tub2.1 anti-?-tubulin) or
bacterially expressed stathmin-FLAG (Holmfeldt et al., 2006; a generous gift
from Martin Gullberg).
Confocal microscopy was used to image fixed and living cells as described
previously (Warren et al., 2006). To image GFP-tubulin for MT dynamics or
EB1-GFP to measure MT nucleation rates, 30–35 images were acquired every
4 s by using two-line mean averaging, requiring a scan time of 3.5–4.0 s. To
count all EB1-GFP comets per cell, Z series were collected from live LLCPK-
EB1-GFP cells with or without stathmin-CFP expression. Z series also were
collected for fixed mitotic MEFs labeled with antibodies to EB1.
Image Analysis and Microtubule Tracking
MetaView imaging software (Molecular Devices, Sunnyvale, CA) was used to
measure polymer level in MEFs, as described previously (Howell et al., 1999a),
for cells fixed in ice-cold methanol and stained for ?-tubulin. MetaView
software also was used to track length changes of individual MTs in cells
expressing GFP-?-tubulin as described previously (Piehl and Cassimeris,
2003; Warren et al., 2006). Interphase MTs were selected for tracking if they
had clearly discernible plus-end tips at the cell periphery that persisted for at
least 30 frames, or 120 s. All clearly defined MTs within an image series were
tracked. Images acquired from LLCPK-GFP-tubulin cells expressing CFP or
stathmin-CFP were later used to measure MT density at the cell periphery by
counting the number of MTs within 5 ?m of the cell cortex. MetaMorph
imaging software (Molecular Devices) was used to manually count all EB1-
GFP comets per cell from Z-series image stacks.
Dynamic Instability Calculations
Parameters of dynamic instability were calculated as described previously
(Warren et al., 2006; Warren and Cassimeris, 2007). Rates of growth and
shortening, and total time spent in growth, pause, or shortening were calcu-
lated for each MT tracked. Dynamicity was calculated by summing the total
lengths of MT polymer gained and lost for each MT divided by the total time
observed (Warren et al., 2006). Dynamicity describes total gain and loss of
tubulins per unit time. Drift velocity, a measure of net growth or shortening,
was calculated as described previously (Vorobjev et al., 1999; Warren et al.,
2006). The frequencies of catastrophe (kcat) and rescue (kres), were calculated
as described by Rusan et al. (2001). The standard deviations for transition
frequencies were calculated from catastrophe frequencies, or rescue frequen-
cies, divided by the square root of the number of transitions observed (Walker
et al., 1988; Howell et al., 1999b). Statistical analyses were performed by
single-factor analysis of variance (ANOVA) in Excel (Microsoft, Redmond,
WA) or Student’s t test (Howell et al., 1999a).
Total RNA was isolated from cultured MEF cell lines by using TRIzol reagent
(Invitrogen) following manufacturer’s instructions, followed by DNase treat-
D. N. Ringhoff and L. Cassimeris
Molecular Biology of the Cell3452
ment. RNA was used to synthesize cDNA with SuperScript III First-Strand
Synthesis System (Invitrogen) for quantitative reverse transcription-PCR
Oligonucleotide probes were designed using Primer Express Software (Ap-
plied Biosystems, Foster City, CA; Supplemental Table 1) and cDNA se-
quences from the mouse genome (Benson et al., 2007). Stathmin message
levels were quantified for each genotype by using oligonucleotide probes
designed to amplify cDNA fragments containing stathmin (Supplemental
Table 1). Results were normalized to the signal from glyceraldehyde-3-phos-
phate dehydrogenase (GAPDH). Amplification was achieved using Power
SYBR Green PCR Master Mix (Applied Biosystems) and 7300 Real-Time PCR
System (Applied Biosystems) with SDS version 1.4 software. RT-PCR ampli-
con specificity was checked by electrophoresis of RT-PCR products on 2%
agarose gels (data not shown). Following manufacturer’s instructions (Ap-
plied Biosystems), the threshold for determining CTvalues was set to log scale
0.2, and internal normalization of genotype results to GAPDH was first
calculated (i.e., CTtarget mRNA ? CTGAPDH mRNA ? ?CT). ?CTvalues
were not calculated for target mRNA samples that exceeded 35 cycles before
crossing the threshold. The relative abundance of target mRNA between
genotypes was calculated as 2 raised to the negative of the difference in ?CT
values [i.e., 2?(?CT ?/? target mRNA ? ?CT ?/? or ?/? target mRNA)(Applied
All statistical analyses were done using Excel (Microsoft). RT-PCR data are
presented as -fold changes relative to MEF Stmn1?/?samples. SEs were
computed using ?CTvalues transformed to SE values of -fold change using
the formula 2?[(?CT ?/? target mRNA ? ?CT ?/? or ?/? target mRNA) ? SE ?CT].
Significance was determined by ANOVA of ?CTvalues (Gru ¨ndemann et al.,
2008; Westberry et al., 2008). Amplification efficiency was determined to be
100% for GAPDH for each genotype per manufacturer’s instructions (data not
shown), and it was assumed that efficiency of target genes was also 100% for
our statistical analysis.
Mouse Embryo Fibroblast Cell Lines Differing in Stathmin
To establish cell lines expressing varying levels of stathmin,
we isolated MEF lines from mouse embryos (?/?), (?/?),
and (?/?) for the Stmn1 gene. The genotypes of the em-
bryos and MEF lines were determined by PCR amplification
of either a region of intron 3 present in wild-type but not
in knockout lines, or a region of the neomycin cassette
that replaced a portion of the Stmn1 gene (Figure 1A and
Supplemental Table 1) (Schubart et al., 1996). Stmn1
mRNA levels, measured by qRT-PCR, corresponded to
the genotypes of the MEF lines: stathmin?/?MEFs con-
tained half as much stathmin mRNA as stathmin?/?
MEFs, whereas stathmin?/?MEFs were void of stathmin
mRNA (Figure 1B). At the protein level, stathmin?/?
MEFs expressed ?50% of the stathmin of wild-type cells.
Stathmin protein was undetectable in stathmin?/?MEFs
(Figure 1C). Measurements of stathmin mRNA and pro-
tein levels were consistent with those reported previously
for mouse neonatal brain and 2-mo-old testicular tissues
(Schubart et al., 1996) and demonstrate that stathmin
mRNA and protein expression are proportional to the
We also estimated the relative abundance of stathmin and
tubulin in MEFs based on Western blots by comparing the
band intensities from purified proteins or cell lysates. We
estimate that the stathmin level in wild-type MEFs is ?170
ng/mg total protein, or ?0.017% of total cell protein (Figure
1D), consistent with previous measurements from nontrans-
formed cells (120–330 ng/mg; Brattsand et al., 1993). Blots
also were probed with anti-? tubulin Tub2.1, a pan-tubulin
antibody that recognizes all ?-tubulin isotypes (Matthes et
al., 1988). The ?-tubulin concentration in MEFs was ?12
?g/mg total protein (data not shown) and was similar in all
three stathmin genotypes (Figure 2). Assuming that ?- and
?-tubulins are present at equal levels, we estimate that the
tubulin concentration in each cell line is ?24 ?g/mg total
protein, or 2.4% of total protein, which is also consistent
with previous estimates of tubulin concentration in cells
(?2–3% of total cell protein; Hiller and Weber, 1978; Sellin et
al., 2008). Based on our estimates for tubulin and stathmin
concentrations, tubulin is present at approximately a 25-fold
molar excess over stathmin in wild-type MEFs and, there-
fore, at a 50-fold molar excess over stathmin in MEFs het-
erozygous for stathmin.
Representative PCR amplicons are shown for each genotype. Geno-
types of embryos and cell lines were assigned based on the presence
or absence of the Stmn1 gene and the neomycin gene used to disrupt
the Stmn1 gene. (B) qRT-PCR of stathmin mRNA derived from each
cell line, shown as –fold change relative to stathmin?/?. Stathmin
mRNA is reduced by 50% in stathmin?/?cells and was not
detectable in stathmin?/?cells. (C) Stathmin protein level corre-
sponds to the copy number of the Stmn1 gene. (D) Semiquanti-
tative estimate of stathmin concentration in wild-type MEFs. The
left four lanes show decreasing loads of purified stathmin. The
detection limit was ?0.1 ng. Two stathmin?/?MEF lines (3 ?g of
total protein per lane) are shown relative to the stathmin stan-
dards. Based on immunoblot band intensities, these lines express
?0.5 ng/3 ?g total protein.
Stathmin genotypes and protein levels in MEF lines. (A)
Maximum intensity projections of MTs in MEF cell lines after fixa-
tion and staining for ?-tubulin. Bar, 25 ?m. (B) Mean MT polymer
content per cell ? SD averaged from three experiments. Both stath-
min?/?(n ? 22) and stathmin?/?(n ? 19) lines contain more MT
polymer than wild-type cells (**p ? 0.001; n ? 19). (C) Relative
?-tubulin levels in MEF lines detected by immunoblots probed with
a pan-?-tubulin antibody, Tub 2.1. Total tubulin remains nearly
constant between cell lines differing in STMN1 genotype. (D) Su-
pernatant (S) and cytoskeletal pellet (P) fractions from stathmin?/?
and stathmin?/?MEFs. Deletions of both copies of the stathmin
gene shift almost all tubulin into polymer.
MT polymer level increased with loss of stathmin. (A)
Stathmin Regulates Microtubule Nucleation
Vol. 20, August 1, 2009 3453
Stathmin Regulates Tubulin Dimer/Polymer Partitioning
and MT Nucleation with Little Impact on MT Plus-End
Stathmin regulation of tubulin dimer/polymer partition-
ing has been well documented in leukemia-derived lines
(Marklund et al., 1996; Holmfeldt et al., 2002; Sellin et al.,
2008), but mechanisms responsible for shifts in tubulin
dimer partitioning between soluble and polymer pools have
not been fully explored. Stathmin also regulates MT plus
end dynamics and polymer level, as shown in primary cul-
tures of newt lung cells (Howell et al., 1999a). Consistent
with these previous studies, we found that MT polymer
levels increased with stathmin level decrease, as shown by
fluorescence intensity measurements of ?-tubulin–stained
cells (Figure 2, A and B). Stathmin?/?cells had ?1.4 times
the MT polymer of wild-type MEFs. Polymer level increased
to 1.6 times the wild-type level in stathmin?/?MEFs (Figure
2B), similar to the increases observed after transient knock-
down or inhibition of stathmin (Holmfeldt et al., 2007). The
increase in MT polymer in stathmin?/?and stathmin?/?
MEFs was not associated with an overall increase in the
concentration of tubulin (Figure 2C). These data indicated
that the soluble tubulin pool should be reduced in stath-
min?/?MEFs. To test this prediction directly, we separated
soluble and cytoskeletal fractions from stathmin?/?and
stathmin?/?MEFs. As shown in Figure 2D, loss of stathmin
shifted most tubulin into the cytoskeletal fraction. A similar
increase in MT polymer and decreased tubulin dimer pool
was observed recently in K562 leukemia cells after transient
knockdown of stathmin (Sellin et al., 2008).
To determine the impact of stathmin on MT dynamics,
MEF cell lines were transiently transfected with GFP-?-
tubulin, and the dynamic instability of their individual
MTs was tracked by time-lapse fluorescence microscopy
at the cell periphery. In all three cell lines, MTs extended
to the cell periphery and were dynamic. In general, MT
dynamics were similar among cells from all three stath-
min genotypes. Growth and shortening rates were both
slightly reduced in stathmin?/?cells compared with
wild-type and stathmin?/?cells. Other parameters of
dynamic instability, including catastrophe and rescue fre-
quencies, were similar in all three cell lines (Table 1). The
similar parameters for dynamic instability in cells from all
three stathmin genotypes is also shown by the relatively
small differences in dynamicity (the total gain and loss of
subunits per unit time) and drift velocity (the net gain or
loss of subunits over time; Table 1). Although not signif-
icantly different, drift velocity was slightly negative in
stathmin?/?cells and shifted to slightly positive values in
stathmin?/?and stathmin?/?cells, indicating that MT
dynamics shift toward net growth in the absence of stath-
Differences in MT dynamics among MEFs of different
stathmin genotypes were small compared with the changes
in MT polymer level, indicating that stathmin could regulate
polymer level through additional mechanisms. To probe
whether stathmin contributes to MT nucleation rate from
centrosomes, we counted new MTs as they emerged from
the centrosome using GFP-EB1 comets to mark MT plus
ends (Piehl et al., 2004). Stathmin?/?MEFs nucleated 14 ? 3
MTs/min (Figure 3A). Nucleation rate was higher in stath-
min?/?MEFs, although not statistically significant, and
increased significantly to 20 ? 2 MTs/min in stathmin?/?
MEFs (Figure 3A).
MT Dynamics and Nucleation in LLCPK Cells with
Measurement of MT dynamics in MEF lines indicated that
stathmin knockout had little impact on plus-end dynamic
instability, whereas significantly increasing the amount of
MT polymer. To confirm these results, we also examined MT
dynamics and nucleation rate in LLCPK-GFP-tubulin epi-
thelial cells transiently overexpressing stathmin-CFP. Our
observations were confined to cells expressing moderate
levels of stathmin-CFP, in which MTs were detectable. In
Table 1. MT dynamic instability in MEFs of various stathmin ge-
Total MTs tracked
Total time tracked (s)
Growth rate (?m/min)
% time spent in growth
% time spent in
% time spent in pause
Drift velocity (?m/min) ?0.07 ? 1.32
Rescue frequency (s?1)
7.1 ? 4.0
9.0 ? 5.4
7.1 ? 3.3
8.3 ? 4.5
6.0 ? 2.3**
7.4 ? 5.2*
4.33 ? 3.063.07 ? 1.26
0.18 ? 0.73
3.09 ? 1.63
0.43 ? 0.80
0.042 ? 0.004 0.038 ? 0.004 0.035 ? 0.004
0.054 ? 0.005 0.050 ? 0.005 0.065 ? 0.006
Confocal microscopy and subsequent image analysis were used to
track changes in MT lengths and to quantify the various parameters
of dynamic instability as described in Materials and Methods. Data
are given as mean ? SD. ANOVA was used to determine significant
differences (growth rate, shortening rate, dynamicity, drift velocity,
nucleation rate). Two-tailed Student’s t test was applied to catastro-
phe and rescue frequencies. *p ? 0.05 and **p ? 0.01.
ation rate was measured by counting the number of EB1-GFP com-
ets emerging from the centrosome over time (Piehl et al., 2004). (A)
MT nucleation rate in MEFs increased in the absence of stathmin. (B)
Expression of stathmin-CFP in LLCPK epithelial cells decreased
nucleation rate. Data for A and B are means ? SD from five to 13
cells per condition. For each graph, *p ? 0.05 and **p ? 0.01.
MT nucleation rates in MEFs and LLCPK cells. Nucle-
D. N. Ringhoff and L. Cassimeris
Molecular Biology of the Cell 3454
these stathmin-CFP–expressing cells, MT dynamics were
similar to that measured in cells expressing CFP (Table 2).
The only significant change was a reduced rescue frequency.
All other parameters, including dynamicity and drift veloc-
ity, were not significantly different between cells expressing
CFP or stathmin-CFP.
The expression of stathmin-CFP in LLCPK cells showed
minor changes in MT dynamics compared with control cells,
whereas at higher expression, reduced MT density made it
difficult to image MTs. Attempts to estimate the level of
stathmin-CFP expression indicated a wide distribution of
expression levels (our unpublished observations), which
prevented qualitative estimates of the amount of stathmin-
CFP expression in those cells in which we measured MT
dynamics. As an alternative, we also microinjected LLCPK-
GFP-tubulin cells with purified stathmin-FLAG. Cells micro-
injected with stathmin-FLAG again showed MT dynamics
similar to control cells (expressing CFP; Table 2), although
growth and shortening rates were slightly slower and rescue
frequency was reduced. Cells injected with stathmin-FLAG
also showed reduced dynamicity. The small impact of stath-
min on MT dynamics at the cell periphery was therefore not
due to the tag on stathmin.
We next measured MT nucleation from the centrosome,
because nucleation rate was sensitive to stathmin level in
MEFs (Figure 3A). Expression of stathmin-CFP decreased
nucleation rate by ?40% compared with that measured in
cells expressing CFP (Figure 3B). The MT nucleation data are
consistent with the results in MEFs, pointing to a role of
stathmin in regulating MT nucleation from the centrosome.
We then asked whether stathmin-CFP or stathmin-FLAG,
at the levels expressed in those cells having sufficient MTs
for dynamics measurements, was able to modulate tubulin
dimer/polymer partitioning. We used images previously
collected for dynamics measurements and counted the num-
ber of plus ends extending to within 5 ?m of the plasma
membrane. By this measurement, MT density at the cell
periphery dropped by ?33% for cells expressing stathmin-
CFP compared with those expressing CFP and dropped by
50% in cells microinjected with stathmin-FLAG (Figure 4A).
We confirmed these results by counting the total number of
growing MTs per cell, marked by EB1-GFP comets, in either
untransfected cells or cells expressing moderate levels of
stathmin-CFP. As shown in Figure 4B, cells expressing stath-
min-CFP had significantly fewer growing MTs per cell.
Stathmin Does Not Regulate MT Nucleation at
The above-mentioned analyses focused on interphase cells
because stathmin is thought to be inactivated in mitosis by
phosphorylation of four serine residues (Larsson et al., 1995;
Tournebize et al., 1997). In contrast, others have suggested
that stathmin is required for mitotic progression (Rana et al.,
2008). Because our data pointed to stathmin functioning to
regulate interphase MT nucleation, rather than MT plus-end
dynamics, we used two measures to examine possible stath-
min function in mitosis. First, we compared the number of
astral MTs in stathmin?/?and stathmin?/?MEFs. As
shown in Figure 5A, stathmin genotype did not impact the
number of astral MTs. Second, we measured MT nucleation
rate in LLCPK cells at metaphase of mitosis. As shown in
Figure 5B, MT nucleation at metaphase was insensitive to
moderate expression of stathmin-CFP. Our data are consis-
Table 2. Dynamic instability in LLCPK cells
CFP controlStmn-CFP Stmn-Flag
Total MTs tracked
Total time tracked (s)
Growth rate (?m/min)
% time spent in growth
% time spent in
% time spent in pause
Rescue frequency (s?1) 0.086 ? 0.005 0.066 ? 0.004* 0.063 ? 0.004*
8.5 ? 5.8
11.3 ? 7.9
7.9 ? 4.5
10.5 ? 8.0
7.3 ? 3.3*
9.2 ? 6.3*
5.3 ? 2.7
0.7 ? 1.6
4.5 ? 3.2
0.2 ? 1.8
3.6 ? 1.5*
0.2 ? 1.0
0.053 ? 0.003 0.044 ? 0.003 0.044 ? 0.003
MT dynamics were measured in LLCPK cells stably expressing
GFP-?-tubulin and CFP or stathmin (stmn)-CFP. Additional cells
were microinjected with stathmin-FLAG as described in Materials
concentration. (A) MT density was measured by counting the num-
ber of MTs extending to within 5 ?m of the cell membrane and
shown as the number of MTs per 10 ?m of cell circumference.
Images used for measuring MT density were the same as those used
for dynamics measurements (see Table 2). Examples of images
(inverted gray scale) used to measure MT density are shown to the
right. These images were selected initially only for dynamics mea-
surements and the sole selection criterion was clarity of the MT
ends. Data shown are means ? SD from 19 to 24 cells. Bar, 5 ?m. (B)
The total number of growing MTs per cell was measured by count-
ing all EB1-GFP comets per cell. Data shown are means ? SD from
21 to 22 cells. Examples of images (inverted gray scale) are shown to
the right. Stathmin-CFP–expressing cell is marked by an arrow. The
insets show enlarged regions from the cell periphery. Bar, 20 ?m.
For both A and B, **p ? 0.01.
MT density is reduced in cells having increased stathmin
Stathmin Regulates Microtubule Nucleation
Vol. 20, August 1, 2009 3455
tent with previous results showing that stathmin is inacti-
vated during mitosis (Larsson et al., 1995; Tournebize et al.,
Stathmin Regulates Tubulin Dimer/Polymer Partitioning
and MT Nucleation from Centrosomes
Manipulating stathmin expression level in noncancerous
cells, either by isolation of MEFs differing in stathmin geno-
type or by transient overexpression of stathmin in LLCPK
cells, resulted in changes in tubulin partitioning between
dimer and polymer pools. Several different assays were used
to measure MT polymer, including measurement of MT
staining intensity, Western blot of soluble and cytoskeletal
fractions, and counting both the number of MTs within 5 ?m
of the cell periphery and the total number of growing MTs
(EB1-GFP comets) per cell. These data demonstrate that
lowering stathmin increases MT polymer, whereas raising
stathmin decreases MT polymer, consistent with previous
results in leukemia-derived cells (Holmfeldt et al., 2007;
Sellin et al., 2008), newt lung epithelial cells (Howell et al.,
1999a), and PtK1 cells (Wittmann et al., 2004). Given the low
concentration of stathmin relative to tubulin in MEFs (Figure
1) and LLCPK cells (Supplemental Figure 1), it is unlikely
that tubulin-sequestering activity plays a major role in reg-
ulating tubulin dimer/polymer partitioning.
Surprisingly, the changes in MT dimer/polymer partition-
ing were not accompanied by large changes in MT dynamics
at the cell periphery. We found that MTs remained dynamic
in both MEFs lacking stathmin and in LLCPK cells overex-
pressing stathmin. MTs at the cell periphery grew and short-
ened at similar rates regardless of stathmin level, although
these rates were reduced slightly in cells with either de-
creased or increased stathmin level (Tables 1 and 2). In
general, we observed relatively small changes in MT dy-
namics, as measured by MT dynamicity (the total net gain
and loss of tubulin dimers over time) and drift velocity (the
net gain or loss of dimers over time), for cells differing in
It is surprising that stathmin level had relatively mild
effects on MT dynamics, given its major effect on MT catas-
trophes and lengths in Xenopus egg extracts (Tournebize et
al., 1997) and on catastrophe frequency in newt lung epithe-
lial cells (Howell et al., 1999a). In contrast, our results agree
with recent results from leukemia-derived cell lines, in
which stathmin depletion did not have a detectable effect on
MT sensitivity to depolymerization by nocodazole (Sellin et
al., 2008). The different effects of stathmin on the MT system
in Xenopus extracts (Tournebize et al., 1997) or in several cell
types (Sellin et al., 2008; this study) may reflect different
responses of the MT system in unbounded (extract) or
bounded (cells) states, in which boundary effects from the
cell’s plasma membrane impact dynamics (Gregoretti et al.,
2006). It is not yet clear why stathmin regulated catastrophes
in newt lung cells but had no impact on catastrophes in the
two cell types studied here.
In most of our experiments, MT dynamics was measured
several days or longer after stathmin level manipulation
(transient overexpression or gene knockout), providing cells
time to adapt and reach a new steady state. In all cases, we
examined the MTs that remained at the cell periphery hours
to days after manipulating stathmin level and not immedi-
ately upon a change in stathmin concentration. It is possible
that up-regulation of stathmin initially increased MT catas-
trophes but that the system adapted to return the catastro-
phe rate to its baseline rate, for those MTs that remain. Such
an adaptation would have to occur in ?2 h, based on data
from cells injected with stathmin-FLAG. We cannot formally
rule out such an adaptation of the MT system but should
such adaptation occur, it is not sufficient to return the MT
polymer to its original level, based on MT density at the cell
periphery or the number of growing MT ends per cell (Fig-
How then does stathmin regulate tubulin partitioning
between soluble and polymer pools without major changes
to MT dynamics at the cell periphery? We suggest that
stathmin-dependent regulation of new MT formation at the
centrosome is the critical function regulating tubulin parti-
tioning. Nucleation rate was increased in stathmin?/?MEFs
(Figure 3A) and reduced in LLCPK-EB1-GFP cells overex-
pressing stathmin-CFP (Figure 3B). By regulating MT nu-
cleation, stathmin contributes to setting the number of
MTs per cell. We suggest that stathmin shifts tubulin
dimer/polymer partitioning through regulation of MT
number, rather than through regulation of MT dynamics
at the cell periphery. These data support a model origi-
nally proposed by Mitchison and Kirschner (1987), dem-
onstrating computationally that changes in the number of
nucleation sites can regulate tubulin dimer/polymer par-
titioning (Mitchison and Kirschner, 1987).
phase. (A) The number of astral MTs, measured by counting EB1
marked MT tips, was similar for metaphase spindles from stath-
min?/?and stathmin?/?MEFs. Data shown are means ? SD for
13–18 spindle poles. Images to the right are maximum intensity
projections from image stacks used to count EB1 comets. EB1
(green), ?-tubulin (red) and DNA (blue) are shown. (B) MT nucle-
ation rate was measured in LLCPK-EB1-GFP cells by counting the
number of EB1-GFP comets emerging from the centrosome over
time. Data shown are means ? SD from seven to 15 cells. Example
of EB1-GFP image from a mitotic cell expressing stathmin-CFP is
shown to the right. Nucleation rate was measured for the astral side
of the centrosome, as outlined by the white half-circle. Bar, 5 ?m.
Stathmin does not regulate MT nucleation during meta-
D. N. Ringhoff and L. Cassimeris
Molecular Biology of the Cell3456
Although our results point to a role for stathmin in regu-
lating MT nucleation, it is not clear where this regulation
occurs. Our nucleation assay uses EB1-GFP to detect new
MTs as they emerge from the centrosome; therefore, we
detect either nucleation or a step shortly thereafter. It is
possible that nascent MTs are more sensitive to stathmin
level than those MTs that reach the cell periphery, possibly
because nascent MTs have not yet accumulated sufficient
stabilizing MAPs to sustain growth. Alternatively, the nu-
cleation step(s) may be more sensitive to stathmin level than
is continued plus-end polymerization. In support of this
idea, stathmin regulates the number of MTs nucleated from
flagellar axoneme fragments in vitro in the absence of MAPs
(Howell et al., 1999b).
MT Growth Rate Seems Independent of Tubulin Dimer
Stathmin binds two tubulin dimers and can sequester those
dimers to prevent their polymerization (Steinmetz, 2007);
yet, we find that increased or decreased stathmin level did
not significantly impact MT growth rate (Tables 1 and 2). We
also estimate that stathmin is present at low concentrations
relative to tubulin in both MEFs (described above) and
LLCPKs (Supplemental Figure 1). These data indicate that
stathmin does not function as a sequestering protein in
noncancerous cells, consistent with our previous observa-
tions in newt lung cells (Howell et al., 1999a). Surprisingly,
we also found that the total tubulin pool was equal in all
MEF cell lines, whereas elimination of both copies of the
stathmin gene shifted most tubulin into polymer and dras-
tically reduced the amount of unpolymerized dimers (Fig-
ure 2, C and D). Given the low concentration of tubulin
dimers in stathmin?/?MEFs, it is surprising that MT
growth rate was only slightly slower in these cells (Table 1).
These data indicate that, in cells, MT growth rate shows little
variation across a surprisingly wide range of tubulin dimer
concentrations, as suggested several years ago by experi-
ments in Xenopus extracts and cytoplasts (Parsons and
Salmon, 1997; Rodionov et al., 1999).
Stathmin Regulation of the MT System
Stathmin regulates the MT system at several levels, includ-
ing dimer/polymer partitioning and in some cells, tubulin
expression (Fletcher and Rorth, 2007; Sellin et al., 2008). For
example, in Jurkat cells, stathmin depletion reduces both
total tubulin protein and mRNA levels, yet at the same time
shifts more tubulins into polymer (Sellin et al., 2008). In
Drosophila embryos, stathmin also regulates the total tubulin
pool (Fletcher and Rorth, 2007). In MEFs, we see approxi-
mately equal total tubulin expression, but shifts in dimer/
polymer partitioning, similar to previous reports for K562
cells (Holmfeldt et al., 2007; Sellin et al., 2008).
Stathmin is phosphorylated by a number of kinases,
which reduces stathmin’s MT destabilizing activity (re-
viewed in Cassimeris, 2002). It is likely that phosphorylation
also turns off stathmin’s ability to regulate MT nucleation,
suggesting that MT number can be modified rapidly in
response to various signal cascades. It is interesting to note
that stathmin is phosphorylated as cells enter mitosis (Larsson
et al., 1995), at a time when nucleation of MTs increases
dramatically (Kuriyama and Borisy, 1981; Piehl et al., 2004).
It is possible that stathmin inactivation contributes to the
greater MT nucleation during mitosis, although this has not
been tested directly. Here, we show that stathmin expression
level did not affect either metaphase MT nucleation rate or
the number of astral MTs, supporting the idea that stath-
min is inactivated during mitosis. Further experiments are
needed to understand how stathmin regulates MT nucle-
ation and the impact of this regulation on the MT system,
cell cycle progression, and other stathmin-dependent pro-
We thank Areeb Zamir for help collecting images of EB1-GFP in LLCPK cells.
We thank Ann Boulet (University of Utah) for advice on MEF isolation, Gleb
Shumyatsky for generously providing stathmin?/?mice, Jutta Marzillier
(Lehigh University) for assistance with qRT-PCR, and Cindy Spittle (Fox
Chase Cancer Center) for assistance with genotyping. We also thank Martin
Gullberg (University of Umea) for purified stathmin protein and for critically
reading the manuscript. This study was supported by National Institutes of
Health grant GM-058025 (to L.C.).
Belmont, L., and Mitchison, T. (1996). Identification of a protein that interacts
with tubulin dimers and increases the catastrophe rate of microtubules. Cell
Benson, D. A., Karsch-Mizrachi, I., Lipman, D. J., Ostell, J., and Wheeler, D. L.
(2007). GenBank. Nucleic Acids Res. 35, D21–25.
Bradford, M. M. (1976) A rapid and sensitive method for the quantitation of
microgram quantities of protein utilizing the principle of protein-dye binding.
Anal. Biochem. 72, 248–254.
Brattsand, G., Roos, G., Marklund, U., Ueda, H., Landberg, G., Nanberg, E.,
Sideras, P., and Gullberg, M. (1993). Quantitative-analysis of the expression
and regulation of an activation-regulated phosphoprotein (oncoprotein 18) in
normal and neoplastic-cells. Leukemia 7, 569–579.
Cassimeris, L. (2002). The oncoprotein 18/stathmin family of microtubule
destabilizers. Curr. Opin. Cell Biol. 14, 18–24.
Curmi, P., Nogue `s, C., Lachkar, S., Carelle, N., Gonthier, M., Sobel, A.,
Lidereau, R., and Bie `che, I. (2000). Overexpression of stathmin in breast
carcinomas points out to highly proliferative tumours. Br. J. Cancer 82,
Desai, A., and Mitchison, T. J. (1997). Microtubule polymerization dynamics.
Annu. Rev. Cell. Dev. Biol. 13, 83–117.
Fletcher, G., and Rorth, P. (2007). Drosophila stathmin is required to maintain
tubulin pools. Curr. Biol. 17, 1067–1071.
Graessmann, A., Graessmann, M., and Mueller, C. (1980). Microinjection of
early SV40 DNA fragments and T antigen. Methods Enzymol. 65, 816–825.
Gregoretti, I., Margolin, G., Alber, M., and Goodson, H. (2006). Insights into
cytoskeletal behavior from computational modeling of dynamic microtubules
in a cell-like environment. J. Cell Sci. 119, 4781–4788.
Gru ¨ndemann, J., Schlaudraff, F., Haeckel, O., and Liss, B. (2008). Elevated
alpha-synuclein mRNA levels in individual UV-laser-microdissected dopa-
minergic substantia nigra neurons in idiopathic Parkinson’s disease. Nucleic
Acids Res. 36, e38.
Hiller, G., and Weber, K. (1978). Radioimmunoassay for tubulin: a quantita-
tive comparison of the tubulin content of different established tissue culture
cells and tissues. Cell 14, 795–804.
Holmfeldt, P., Brattsand, G., and Gullberg, M. (2002). MAP4 counteracts
microtubule catastrophe promotion but not tubulin-sequestering activity in
intact cells. Curr. Biol. 12, 1034–1039.
Holmfeldt, P., Bra ¨nnstro ¨m, K., Stenmark, S., and Gullberg, M. (2006). Aneu-
genic activity of Op18/stathmin is potentiated by the somatic Q18–?e mu-
tation in leukemic cells. Mol. Biol. Cell 17, 2921–2930.
Holmfeldt, P., Stenmark, S., and Gullberg, M. (2007). Interphase-specific
phosphorylation-mediated regulation of tubulin dimer partitioning in human
cells. Mol. Biol. Cell 18, 1909–1917.
Howell, B., Deacon, H., and Cassimeris, L. (1999a). Decreasing oncoprotein
18/stathmin levels reduces microtubule catastrophes and increases microtu-
bule polymer in vivo. J. Cell Sci. 112, 3713–3722.
Howell, B., Hoffman, D., Fang, G., Murray, A., and Salmon, E. (2000). Visu-
alization of Mad2 dynamics at kinetochores, along spindle fibers, and at
spindle poles in living cells. J. Cell Biol. 150, 1233–1250.
Howell, B., Larsson, N., Gullberg, M., and Cassimeris, L. (1999b). Dissociation
of the tubulin-sequestering and microtubule catastrophe-promoting activities
of oncoprotein 18/stathmin. Mol. Biol. Cell 10, 105–118.
Stathmin Regulates Microtubule Nucleation
Vol. 20, August 1, 2009 3457
Kuriyama, R., and Borisy, G. (1981). Microtubule-nucleating activity of cen- Download full-text
trosomes in Chinese hamster ovary cells is independent of the centriole cycle
but coupled to the mitotic cycle. J. Cell Biol. 91, 822–826.
Larsson, N., Melander, H., Marklund, U., Osterman, O., and Gullberg, M.
(1995). G2/M transition requires multisite phosphorylation of oncoprotein 18
by two distinct protein kinase systems. J. Biol. Chem. 270, 14175–14183.
(1999). Op18/stathmin mediates multiple region-specific tubulin and microtu-
bule-regulating activities. J. Cell Biol. 146, 1289–1302.
Liedtke, W., Leman, E. E., Fyffe, R. E., Raine, C. S., and Schubart, U. K. (2002).
Stathmin-deficient mice develop an age-dependent axonopathy of the central
and peripheral nervous systems. Am. J. Pathol. 160, 469–480.
Marklund, U., Larsson, N., Gradin, H., Brattsand, G., and Gullberg, M. (1996).
Oncoprotein 18 is a phosphorylation-responsive regulator of microtubule
dynamics. EMBO J. 15, 5290–5298.
Matthes, T., Wolff, A., Soubiran, P., Gros, F., and Dighiero, G. (1988). Antitu-
bulin Antibodies. 2. Natural autoantibodies and induced antibodies recognize
different epitopes on the tubulin molecule. J. Immunol. 141, 3135–3141.
Minotti, A., Barlow, S., and Cabral, F. (1991). Resistance to antimitotic drugs
in Chinese hamster ovary cells correlates with changes in the level of poly-
merized tubulin. J. Biol. Chem. 266, 3987–3994.
Mistry, S., Bank, A., and Atweh, G. (2005). Targeting stathmin in prostate
cancer. Mol Cancer Ther. 4, 1821–1829.
Mitchison, T., and Kirschner, M. (1987). Dynamic instability of microtubule
growth. Nature 312, 237–242.
Nguyen, H., Gruber, D., and Bulinski, J. (1999). Microtubule-associated pro-
tein 4 (MAP4) regulates assembly, protomer-polymer partitioning and syn-
thesis of tubulin in cultured cells. J. Cell Sci. 112, 1813–1824.
Parsons, S., and Salmon, E. (1997). Microtubule assembly in clarified Xenopus
egg extracts. Cell Motil. Cytoskeleton 36, 1–11.
Piehl, M., and Cassimeris, L. (2003). Organization and dynamics of growing
microtubule plus ends during early mitosis. Mol. Biol. Cell 14, 916–925.
Piehl, M., Tulu, U., Wadsworth, P., and Cassimeris, L. (2004). Centrosome
maturation: measurement of microtubule nucleation throughout the cell cycle
by using GFP-tagged EB1. Proc. Natl. Acad. Sci. USA 101, 1584–1588.
Rana, S., Maples, P., Senzer, N., and Nemunaitis, J. (2008). Stathmin 1, a novel
therapeutic target for anticancer activity. Expert Rev. Anticancer Ther. 8,
Rodionov, V., Nadezhdina, E., and Borisy, G. (1999). Centrosomal control of
microtubule dynamics. Proc. Natl. Acad. Sci. USA 96, 115–120.
Rusan, N. M., Fagerstrom, C. J., Yvon, A. M., and Wadsworth, P. (2001). Cell
cycle-dependent changes in microtubule dynamics in living cells expressing
green fluorescent protein-alpha tubulin. Mol. Biol. Cell 12, 971–980.
Saxton, W., Stemple, D., Leslie, R., Salmon, E., Zavortink, M., and McIntosh,
J. (1984). Tubulin dynamics in cultured mammalian cells. J. Cell Biol. 99,
Schubart, U. K., Yu, J., Amat, J. A., Wang, Z., Hoffmann, M. K., and Edelmann,
W. (1996). Normal development of mice lacking metablastin (P19), a phos-
phoprotein implicated in cell cycle regulation. J. Biol. Chem. 271, 14062–14066.
Sellin, M. E., Holmfeldt, P., Stenmark, S., and Gullberg, M. (2008). Global
regulation of the interphase microtubule system by abundantly expressed
Op18/Strathmin. Mol. Biol. Cell 19, 2897–2906.
Shumyatsky, G. P., et al. (2005). Stathmin, a gene enriched in the amygdala,
controls both learned and innate fear. Cell 123, 697–709.
Steinmetz, M. (2007). Structure and thermodynamics of the tubulin-stathmin
interaction. J. Struct .Biol. 158, 137–147.
Tessarollo, L. (2001). Manipulating mouse embryonic stem cells. Methods
Mol. Biol. 158, 47–63.
Tournebize, R., Andersen, S., Verde, F., Dore ´e, M., Karsenti, E., and Hyman,
A. (1997). Distinct roles of PP1 and PP2A-like phosphatases in control of
microtubule dynamics during mitosis. EMBO J. 16, 5537–5549.
Vasquez, R. J., Gard, D. L., and Cassimeris, L. (1994). XMAP from Xenopus
eggs promotes rapid plus end assembly of microtubules and rapid microtu-
bule polymer turnover. J. Cell Biol. 127, 985–993.
Vorobjev, I. A., Rodionov, V. I., Maly, I. V., and Borisy, G. G. (1999). Contri-
bution of plus and minus end pathways to microtubule turnover. J. Cell Sci.
Walker, R. A., O’Brien, E. T., Pryer, N. K., Soboeiro, M. F., Voter, W. A.,
Erickson, H. P., and Salmon, E. D. (1988). Dynamic instability of individual
microtubules analyzed by video light microscopy: rate constants and transi-
tion frequencies. J. Cell Biol. 107, 1437–1448.
Warren, J. C., and Cassimeris, L. (2007). The contributions of microtubule
stability and dynamic instability to adenovirus nuclear localization efficiency.
Cell Motil. Cytoskeleton 64, 675–689.
Warren, J. C., Rutkowski, A., and Cassimeris, L. (2006). Infection with repli-
cation-deficient adenovirus induces changes in the dynamic instability of host
cell microtubules. Mol. Biol. Cell 17, 3557–3568.
Westberry, J., Prewitt, A., and Wilson, M. (2008). Epigenetic regulation of the
estrogen receptor alpha promoter in the cerebral cortex following ischemia in
male and female rats. Neuroscience 152, 982–989.
Wittmann, T., Bokoch, G., and Waterman-Storer, C. (2004). Regulation of
microtubule destabilizing activity of Op18/stathmin downstream of Rac1.
J. Biol. Chem. 279, 6196–6203.
Zhai, Y., Kronebusch, P., Simon, P., and Borisy, G. (1996). Microtubule dy-
namics at the G2/M transition: abrupt breakdown of cytoplasmic microtu-
bules at nuclear envelope breakdown and implications for spindle morpho-
genesis. J. Cell Biol. 135, 201–214.
D. N. Ringhoff and L. Cassimeris
Molecular Biology of the Cell 3458