Download Bone cell elasticity and morphology changes during the cell cycle

Journal of Biomechanics 44 (2011) 1484–1490
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Journal of Biomechanics
journal homepage: www.elsevier.com/locate/jbiomech
www.JBiomech.com
Bone cell elasticity and morphology changes during the cell cycle
Geraldine M. Kelly a,c,n, Jason I. Kilpatrick a, Maarten H. van Es a, Paul P. Weafer b, Patrick J. Prendergast c,
Suzanne P. Jarvis a
a
Nanoscale Function Group, Conway Institute of Biomedical and Biomolecular Research, University College Dublin, Belfield, Dublin 4, Ireland
Department of Mechanical and Biomedical Engineering, National University of Ireland, Galway, Ireland
c
Trinity Centre for Bioengineering, School of Engineering, Trinity College Dublin, Ireland
b
a r t i c l e i n f o
a b s t r a c t
Article history:
Accepted 8 March 2011
The mechanical properties of cells are reported to be regulated by a range of factors including interactions
with the extracellular environment and other cells, differentiation status, the onset of pathological states,
as well as the intracellular factors, for example, the cytoskeleton. The cell cycle is considered to be a
well-ordered sequence of biochemical events. A number of processes reported to occur during its progression
are inherently mechanical and, as such, require mechanical regulation. In spite of this, few attempts have
been made to investigate the putative regulatory role of the cell cycle in mechanobiology. In the present
study, Atomic Force Microscopy (AFM) was employed to investigate the elastic modulus of synchronised
osteoblasts. The data obtained confirm that osteoblast elasticity is regulated by cell cycle phase; specifically,
cells in S phase were found to have a modulus approximately 1.7 times that of G1 phase cells. Confocal
microscopy studies revealed that aspects of osteoblast morphology, namely F-actin expression, were also
modulated by the cell cycle, and tended to increase with phase progression from G0 onwards. The data
obtained in this study are likely to have implications for the fields of tissue- and bio-engineering, where prior
knowledge of cell mechanobiology is essential for the effective replacement and repair of tissue. Furthermore,
studies focused on biomechanics and the biophysical properties of cells are important in the understanding of
the onset and progression of disease states, for example cancer at the cellular level. Our study demonstrates
the importance of the combined use of traditional and relatively novel microscopy techniques in understanding mechanical regulation by crucial cellular processes, such as the cell cycle.
& 2011 Elsevier Ltd. All rights reserved.
Keywords:
Osteoblast
Atomic force microscopy
Cell cycle
Elastic modulus
Cytoskeleton
1. Introduction
The cell cycle is a well-ordered sequence of biochemical events,
occurring in eukaryotic cells between one cell division and the next
(Cooper, 2000). However, a number of processes that occur during
cell cycle progression, for example cytokinesis, are inherently
mechanical and thus also require mechanical regulation. This has
been demonstrated in a number of studies, where functionalised
substrates were used to restrict cell spreading resulting in late G1
phase arrest, decreased chromatin condensation and a reduction in
nuclear swelling and DNA synthesis (Huang et al., 1998; Ingber
et al., 1995; Roca-Cusachs et al., 2008). These irregularities coincided
with disruption of the cytoskeleton, coupled with de-regulated
protein expression. Furthermore, application of mechanical loads
to cells can result in initiation of specific cell cycle phases (Liao et al.,
2004; Sedding et al., 2003; Sun et al., 2004). These data demonstrate
the importance of mechanical forces, cell shape, as well as interplay
between the extracellular matrix (ECM) and cytoskeleton in
regulating cell cycle progression. However, few studies have investigated the influence of cell cycle phase on mechanical properties of
cells. Progression through the cell cycle involves significant remodelling of cytoskeletal systems (Foisner, 1997; Skalli et al., 1992).
Thus, changes in cytoskeletal organisation may influence mechanical properties associated with the cell, for example, cell elasticity.
In this study, we tested the hypothesis that the elasticity and
morphology of bone cells change during the cell cycle. This hypothesis was tested using Atomic Force Microscopy (AFM) to measure the
mean elastic modulus and height of individual phase synchronised
osteoblasts. Confocal measurements were also made to investigate
the effects of cell cycle progression on the nature of the actin
cytoskeleton. Corroboration of this hypothesis will provide useful
insights into the relationship between the cell cycle and cell
mechanobiology.
2. Methods
2.1. MC3T3-E1 osteoblast cell culture and synchronisation
n
Corresponding author at: Nanoscale Function Group, Conway Institute
of Biomedical and Biomolecular Research, University College Dublin, Belfield,
Dublin 4, Ireland. Tel.: þ 353 1 716 6780; fax: þ 353 1 716 6777.
E-mail address: [email protected] (G.M. Kelly).
0021-9290/$ - see front matter & 2011 Elsevier Ltd. All rights reserved.
doi:10.1016/j.jbiomech.2011.03.011
MC3T3-E1 osteoblasts (ATCC-LGC Standards, Middlesex, UK) were cultured in
alpha minimum essential medium (a-MEM; Sigma, Dorset, UK), as described by
McGarry et al. (2007). Cell synchrony was achieved via serum starvation, as described
G.M. Kelly et al. / Journal of Biomechanics 44 (2011) 1484–1490
previously (Han et al., 2003; Schorl and Sedivy, 2007). Briefly, osteoblasts were seeded
at a density of 3500 cells/cm2 in standard growth medium in tissue culture dishes.
After 24 h, medium was removed and cells were washed in phosphate buffered saline
(PBS). Cells were then cultured for 72 h in medium supplemented normally but
containing only 0.1% FBS to induce G0 arrest.
1485
for G1 and t¼ 25–27 h after release for S. Cell height measurements were then
obtained by comparing the POC for each of the curves taken on the cell relative to
the POC of the substrate curves. Data were corrected for instrument drift by
assuming a linear drift rate. Data are presented as mean7standard deviation.
2.6. Statistical analysis
2.2. Analysis of cell cycle progression using flow cytometry
Synchronised cells were released from arrest in standard growth medium and
pulse-labelled with 10 mm bromodeoxyuridine (BrdU; Sigma) for the final 60 min of
culture. BrdU staining was carried out as described previously (Gandarillas and Watt,
1997), in order to identify S phase cells. Cellular DNA content was investigated using
propidium iodide (PI; Sigma) staining solution (50 mg/ml PI, 200 mg/ml RNase A in
PBS), added for 30 min in the dark. Samples were analysed using a Cyan flow
cytometer (DAKO, Stockport, UK). Fluorochromes were excited using a 488 nm laser.
FITC fluorescence was obtained between 515 and 545 nm and PI fluorescence was
obtained above 580 nm. At least 10,000 events per sample were collected.
2.3. Measurement of cytoskeletal dynamics
3. Results
3.1. Osteoblast synchronisation and cell cycle progression
2.4. AFM measurement of cell elasticity
Cells were seeded at a density of 3500 cells/cm2 on sterile glass slides, placed
in tissue culture dishes and synchronised as described in Section 2.1. Prior to
experiments, culture medium was removed and cells were washed in Krebs
bicarbonate buffer (KBB) (Murphy et al., 2009). The slide to which the cells were
adhered was positioned on the AFM stage and cells remained immersed in KBB for
the duration of experiments.
An Asylum Research (Santa Barbra, CA, USA) MFP-3D AFM combined with a
Nikon Eclipse Ti fluorescent microscope was used here. The spring constant and
Inverse Optical Lever Sensitivity (InvOLS) of each cantilever (Lever D, DNP, Veeco,
USA, kE0.03 N/m) was calibrated prior to data collection (Hutter and Bechhoefer,
1993; Meyer and Amer, 1988). Force maps arrays (4 8) were collected in a region
of cytoplasm over a period of approximately 90 s, with 3 s between indentations.
A constant velocity of 600 nm/s and a sample rate of 500 Hz were used. The
maximum force applied was 500 nN, resulting in indentation depths typically less
than 700 nm. Elastic modulus (E) values were determined using the Hertzian
analysis (Eq. (1)) modified for the use of a conical indenter (Sneddon, 1965).
Indenter parameters were calculated from the specifications of the manufacturer.
An empirical algorithm was developed, in house, to facilitate robust automated determination of the point of contact (POC) as this has been shown to be a
critical parameter in determining the value of E (Dimitriadis et al., 2002). Use of
this algorithm minimised subjectivity in pre-processing and allows for improved
consistency and minimised user intervention (Jaasma et al., 2006; Lin et al., 2007).
E values were expressed as mean 7 standard error:
ð1Þ
The Hertz formula (Eq. (1)) (Hertz, 1881) and modifications made to it
(Sneddon, 1965) relates E to the force applied (F) and the indentation depth (d),
determined from force versus distance curves. Poisson’s ratio used was 0.37, as
assumed previously for osteoblasts (McGarry and Prendergast, 2004), while the
half angle of the conical indenter was calculated to be 23.75 7 2.81.
2.5. AFM measurement of cell height
Cell samples were prepared as in Section 2.4 with PBS substituted for KBB and
then height measurements were made using a tipless cantilever (Lever F, NSC12,
MikroMesch, Spain, kE0.65 N/m) in order to minimise the influence of the local
cell topography. Calibration was performed as in Section 2.4. Force curves were
collected at a velocity of 1 mm/s with a peak force of 20 nN to the glass substrate
beside a cell of interest, 2 separate regions of cell cytoplasm, the nucleus and finally
repeated on the glass substrate. Measurements were made at t¼ 1–3 h after release
Synchronisation of cells resulted in accumulation of 490% of
cells in G0/G1. Cells then moved synchronously through G1, with
490% on average remaining in this phase up to 18 h post-release
from arrest (Fig. 1). The number of cells in both S and G2/M during
this period was significantly lower (F¼29,842.9; po0.001). The
percentage of cells in G1 began to decrease 18 h after release from
G0, while the percentage of cells in S was observed to increase,
indicating cells were approaching the G1/S border (Fig. 1).
Cells began to enter S 24 h post-release, when a significantly
larger percentage of cells were recorded in this phase, compared
with G1 and G2/M (F¼96.40, p o0.001) (Fig. 1). The proportion of
cells in S continued to increase until 30 h after release, when
approximately 60% of cells were observed in this phase. It was
expected that cells would now move, relatively synchronously,
into G2/M. However, cells appeared to lose their synchronicity
once they exited S and samples analysed 36 h post-release were
characteristically similar to asynchronised populations (Fig. 1).
3.2. Quantifiable difference in osteoblast morphology during the cell
cycle
Analysis of AFM height data revealed that total height of cells
synchronised in S (n¼ 39) was significantly larger than that of G1
100
% Cells per Cell Cycle Phase
Synchronised cell samples were harvested every 3 h post-release from arrest
and fixed in 4% paraformaldehyde (PFA). Samples were stained using an actin
cytoskeleton and Focal Adhesion Staining Kit (Chemicon, Cork, IE). In brief, an
anti-vinculin primary antibody conjugated to a Gt Ms FITC-conjugated secondary antibody and rhodamine-conjugated phalloidin were used to label focal
adhesions and F-actin, respectively. Cells were counterstained using DAPI. DAPI,
FITC, and rhodamine were excited at 402, 488, and 561 nm, respectively. The
staining protocol was standardised across all samples. Imaging was conducted
immediately after staining using optimal imaging settings, as determined at the
beginning of the first experiment. Emitted fluorescence was obtained at 470, 525,
and 590 nm for each fluorophore. Images of 50 cells were acquired at each time
point (n¼ 3). The Nikon NIS-Elements AR 3.0 software package was used to
measure mean rhodamine–phalloidin fluorescent intensity, from which F-actin
concentrations were inferred and expressed as mean 7standard error.
2E
d2
F¼
p tan að1n2 Þ
One-way analyis of variance (ANOVA) tests were employed to determine
statistical differences between the means of three or more independent groups.
Statistical differences between the means of two independent experimental
groups were determined using unpaired Student’s t-tests. The null hypothesis
was rejected for p40.05. For modulus values, outliers for each cell were identified
as those being beyond the inner fence limits (Q1 1.5 IQR and Q3 þ1.5 IQR)
and were excluded from analysis.
90
G1 Phase
S Phase
G2/M Phase
80
70
60
50
40
30
20
10
0
0
4
8
12
18
20
24
30
36
Hours Released from G0 (h)
Fig. 1. Flow cytometric analysis of osteoblast progression through G1 and S phase.
Statistical analysis of data obtained from flow cytometry experiments (n ¼3).
Between 0 and 12 h post-release, the percentage of cells in G1 was 490%.
However, by 18 h, the average percentage in G1 had decreased to 85%,
indicating that cells were approaching the G1/S transition. 20 h after release from
G0 cells were still in the G1/S transition. Cells entered S phase between 24 and
30 h post-release, when approximately 60% of cells were observed to be in this
phase, on average. Cells did not synchronously enter G2/M as expected. Instead
synchronicity appeared to be lost at 36 h post-release (***p o 0.001).
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G.M. Kelly et al. / Journal of Biomechanics 44 (2011) 1484–1490
(n¼24) cells (po0.001; Fig. 2). It was found that the nucleus
contributed significantly to total cell height. The average measured cytoplasmic height in S cells (n ¼73) was also significantly
larger than that of G1 cells (n¼ 46) (p o0.01; Fig. 2).
3.3. S phase cells express significantly more F-actin than G1 phase
cells
At 0 h post-release actin protein appeared quite diffused
throughout the cell, with very few polymerised filaments apparent (Fig. 3). As cells progressed through G1, actin became
organised into long, linear, well defined filaments, arranged in
6
Total Cell Height
Height of Cytoplasm
Height (µm)
5
large bundles tending to run parallel to each other. This arrangement persisted as cells progressed through G1 into S. However
during S, actin filaments tended to transverse the nucleus more
frequently than in G1 cells (Fig. 3, 24 and 33 h). Quantification of
rhodamine–phalloidin fluorescence revealed that actin expression
was significantly higher in S than in G1 (p o0.01; Fig. 4a). Fig. 4b
shows that actin expression tended to increase for the first 12 h
after release from G0, with an abnominally high value recorded
3 h post-release. Interestingly, expression levels decreased after
12 h with a minima at 24 h, corresponding to the period during
cell cycle progression associated with G1/S transition. Actin
expression levels then increased with progression through S and
beyond and were greatest during this period.
Focal adhesions were observed as intense spots of fluorescence
at terminal ends of actin bundles. Neither the quantity nor
distribution of focal adhesions appeared to change significantly
as cells progressed through the cell cycle, indicating that the cells
were well adhered to the substrate at all times (Fig. 3).
4
3.4. Osteoblast elasticity is modulated by the cell cycle
3
G1 cells had an average elastic modulus of 3.5 70.3 kPa
(n ¼63), compared with an average of 5.9 70.5 kPa (n ¼59) for S
cells (Fig. 5). Thus, S cells were significantly stiffer than G1 cells
(p o0.001) and osteoblast elasticity is correlated with the cell
cycle. Representative force map data, obtained from the cytoplasm of individual cells (Fig. 5), illustrates large intracellular
variability in elasticity, indicative of intracellular heterogeneity.
2
1
0
G1 Phase
S Phase
4. Discussion
Fig. 2. Osteoblast cell height is mediated by the cell cycle. The total height
(including the nucleus) of S phase cells was significantly larger than that of G1
phase cells (***p o0.001). The height of the cytoplasm, the region indented in this
study, was also significantly larger in S phase than in G1 phase (**p o0.01). The
cell heights were found to be G1 cytoplasm height (m) ¼ 1.18 10 6 7 7.39 10 7
(n¼ 46), G1 nucleus height (m) ¼ 3.51 10 6 7 7.45 10 7 (n ¼24), S cytoplasm
height (m) ¼ 1.69 10 6 7 9.51 10 7 (n¼ 73), and S nucleus height (m) ¼4.45 10 6 71.17 10 6 (n¼ 39).
The data obtained during this study has corroborated the
hypothesis that bone cell elasticity and morphology are correlated with cell cycle progression.
6h
18h
G1 Phase
0h
4.1. Modulation of osteoblast elasticity and morphology by the cell
cycle
20 µm
50 µm
21h
20 µm
30h
S Phase
24h
50 µm
20 µm
20 µm
Fig. 3. Investigation of actin cytoskeleton arrangement and focal adhesion distribution during osteoblast cell cycle progression. Polymerised actin (red fluorescence) was
generally visible as long, linear bundles of filaments that transversed the cytoplasm of the cell surrounding the nucleus (blue fluorescence). As shown in the images above,
this arrangement did not change drastically as cells progressed through G1 phase. However, in S phase (24–30 h) it was found that filaments tended to actually cross the
nucleus more frequently. Focal adhesions were observed as intense spots of green fluorescence at the terminal ends of actin filaments. The number and distribution of focal
adhesions did not appear to change due to cell cycle progression. However, they did not appear well formed in G0/early G1, and a large amount of non-specific vinculin
staining in the perinuclear region was visible here (0 h). (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this
article.)
μm
35
30
25
20
0
15
8
6
4
2
0
10 20
μm
30 S
25
20
15
10
5
0
30
0
25
20
15
10
5
0
-5
10 20 30
μm
10
5
0
Cell Cycle Phase
50
45
Mean Fluorescent Intensity (AFU)
30 G1
25
20
15
10
5
0
kPa
**
G1 phase
S phase
Average Elastic Modulus (kPa)
Mean Fluorescent Intensity (AFU)
40
1487
kPa
μm
G.M. Kelly et al. / Journal of Biomechanics 44 (2011) 1484–1490
40
35
7
6
G1 phase
S phase
5
4
3
2
1
0
Cell Cycle Phase
30
Fig. 5. Osteoblast elasticity is modulated by the cell cycle. The force maps are
representative of raw data from single cells in G1 and S phases. The minimum and
maximum elastic modulus values obtained from the G1 map were 0.2 and
12.6 kPa, respectively, while the average value was 2.3 7 3.3 kPa. The minimum
and maximum elastic moduli obtained from the S phase map were 1.5 and
19.5 kPa, respectively, while the average was 5.27 0.6 kPa. The bar chart shows
the average of 63 and 59 G1 and S cells, respectively. S phase cells had a
significantly higher elastic modulus than G1 cells, confirming that osteoblast
elasticity is modulated by the cell cycle (***p o0.001).
25
20
15
10
5
0
0
3
6
9
12 18 21 24 27
Release from G0 (h)
30
33
36
Fig. 4. Comparison of the mean actin expression of G1 and S phase cells. The sum
fluorescent intensity was calculated for each cell and then normalised to cell area
to give a mean intensity value. Cells in S phase had an average mean intensity of
34.5 7 3.5 compared with 25.3 73.9 for G1 phase cells (a). Thus cells in S phase
expressed more actin than those in G1. (b) Mean fluorescent intensity at each time
point assessed. This tended to increase with cell cycle progression, although a
decrease was recorded that coincided with the G1/S transition period (**p o 0.01).
Primary somatic cells are sometimes considered a better
in vitro model than immortalised cell lines. The principle difference between these cell types is the finite proliferative capacity of
primary cells, compared to unlimited replicative ability of immortalised cells (Oh et al., 2007). Techniques used to confer immortality to cells can raise concerns in relation to their functionality;
however in most cases this is unperturbed (May et al., 2005;
Ouyang et al., 2000). Use of immortalised cells was considered
appropriate here, as cell elasticity is reportedly modulated by a
number of intra- and extracellular factors, including differentiation (Ng et al., 2006; Yourek et al., 2007). Use of immortalised
cells, with limited capacity for differentiation, removes this
variable ensuring more accurate results.
Synchronization procedures used here yielded relatively pure
populations of G1 cells. However, failure of approximately 25% of
cells to progress to S meant these populations were not homogenous. It was demonstrated from a relatively pure population of
G1 cells that both elastic modulus and cell height are lower in this
phase than in S. Thus, it is likely that the values obtained for cells
in S will be an underestimate of the true value, due to the
unavoidable influence of G1 cells within the sample. Failure of
some cells to re-enter the cell cycle is likely to be due to cellular
damage during synchronisation; serum starvation has been
reported to result in DNA damage and eventual apoptosis in
some cases (Hayes et al., 2005; Huang et al., 1997). Nonetheless,
serum starvation was deemed to be a more appropriate technique
to synchronise cells than other pharmacological methods commonly used, for example treatment with colchine or nocodazole,
which results in rapid microtubule depolymerisation and eventual cell cycle arrest at G2/M phase (Blajeski et al., 2002; Ng et al.,
1998). Microtubule depolymerisation would significantly impact
the mechanical attributes of cells, which would adversely affect
our ability to isolate the effects of cell cycle progression.
Our study has demonstrated that both cytoplasmic and overall
cell height varied according to cell cycle phase. Cell heights were
larger in S than G1 and were comparable to those measured in
previous studies (Andersen et al., 2005; Darling et al., 2008;
Lehenkari et al., 2000). Indentation depth was determined by
the peak loading force of 500 nN and was found to be in the range
of 300–700 nm dependant on the local heterogeneity of the cell
mechanics. This value is similar to that used in previous AFM
studies of osteoblast cell lines (Jaasma et al., 2006; Charras and
Horton, 2002; Takai et al., 2005). It should be noted that although
the indentation depth was a significant proportion of the cytoplasm height it is thought to be consistent with the magnitude of
the in vivo cell deformations (Jaasma et al., 2006).
Whilst previous studies have adopted more advanced models
for the analysis of AFM indentation data (Jaasma et al., 2006;
Mahaffy et al., 2004) it should be noted that the values obtained
in this study are in good agreement with those in the previous
studies (Darling et al., 2008; Jaasma et al., 2006; Domke et al.,
2000; Hansen et al., 2007). Indeed, according to Kuznetsova et al.
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G.M. Kelly et al. / Journal of Biomechanics 44 (2011) 1484–1490
(2007), although indentation depth in AFM experiments often
exceeds the limits of the Hertz model it remains an adequate
analysis technique, which sufficiently represents the data. If the
model were to be grossly inaccurate then it is unlikely that the
results contained herein would be in such an agreement with
studies devoid of such model limitations. Indentation depths
greater than 10 % of the cell height are anticipated to result in
modulus values higher than the true cell modulus due to the
influence of the stiff substrate (Saha and Nix, 2002). Thus, we
would assume that for a system where the substrate dominates
the interactions that lower cell heights would result in a higher
stiffness value. In fact we have found that the opposite is true;
height is negatively correlated with elastic modulus. This observation, also reported by Darling et al. (2008), indicates that the
increase in modulus from G1 to S is both real and statistically
significant (po0.001). The removal of any influence from the
substrate due to large indentation depths is likely to increase the
effect measured in this study.
Standard errors associated with average modulus values measured are relatively small, indicating that intra-phase variations
were not significant. However, intracellular variations recorded in
osteoblast elasticity were relatively large, with data obtained from
a single cell often spanning an order of magnitude. It has been
reported that application of external forces to cells results in
cytoskeletal reorganisation and cell stiffening (Deng et al., 2004;
Wang et al., 1993). Strain induced stiffening was not evident on
the timescale of force maps obtained here and variability in
elasticity within a force map is considered to reflect intracellular
heterogeneity (Hofmann et al., 1997). Avoiding the nucleus during
collection of force map data may have implications for results, as
the nucleus is reportedly the stiffest and the most mechanically
responsive part of cells (Walker et al., 1999; Tseng et al., 2004).
Although the nucleus was excluded from this study it was
considered that the relatively large area of each cell mapped
(32 points over 90 mm2) was sufficiently representative of samples.
Our results are consistent with previous reports, documenting
that mechanical properties of the cells are cell cycle dependant
(Anderson et al., 1991; Needham et al., 1991). Tsai et al. (1996)
utilised micropipette aspiration to investigate cell cycle dependence of HL-60 cell deformability, and also found that cells in S
were less deformable than those in G1. The above three studies
also reported proliferating populations of cells to be heterogeneous in size and morphology, an observation also made here. In
this study, the average height, measured for cells in S phase, was
seen to increase by a factor of 1.3, with respect to that of cells in
G1. Increases in cell size during cell cycle progression have been
reported previously in relation to many other cell types (Gazitt
et al., 1978; Pasternak et al., 1971; Rubin et al., 1989; Skog and
Tribukait, 1986).
4.2. Role of the cytoskeleton in determining cell cycle dependence of
osteoblast elasticity
It was found that, on average, cells in S expressed 1.36 times
more actin than those in G1. Again, this may be an underestimation of the true ratio, due to unavoidable contribution made from
cells remaining in G1. Leger et al. (1990) reported that the amount
of actin and tubulin in fibroblasts doubled from G1 to G2 phase.
Here, comparison of actin expression levels at the start of G1 with
those at the end of S revealed expression levels to be 1.6 times
higher at the latter time point. Thus, our results are consistent
with those reported by Leger et al. (1990). The role played by the
actin cytoskeleton in determining cell elasticity is well documented (Pourati et al., 1998; Stamenovic and Coughlin, 1999). However, increases in actin protein content alone cannot account for
increases in cell rigidity observed here, as the concurrent increase
in cell height observed is an indication that cell volume has also
increased. Thus, it may be inferred that protein concentrations
remain relatively constant throughout cell cycle progression.
F-actin assemblies utilise actin-binding proteins (ABPs) to construct networks with specific morphologies and mechanical
properties. ABPs regulate network microstructure, governing local
deformation mode and micro- and macroscopic elastic response
of cross-linked actin networks (Lieleg et al., 2010). Appreciable
changes in the organisation of actin stress fibres or focal adhesion
formation, with cell cycle progression were not obvious in this
study. However, ABP mediated changes in F-actin microstructures
might account for observed cell cycle dependence of osteoblast
elasticity. Similarly, reorganisation of the nucleus and its association with the cytoskeleton during DNA synthesis might be a
contributing factor; it was observed that actin filaments transversed the nucleus more frequently in S than in G1 cells, lending
support to this idea.
4.3. Implications
This study utilised traditonal and novel microsocpy techniques
to illustrate that both morphological and mechanical properties of
cells are correlated with the cell cycle. The data obtained
demonstrate the importance of measuring mechanical properties
of cells at subcellular (as well as whole cell) level and confirms
the importance of AFM as a tool for investigating cellular
processes due to its ability to make localised measurements.
Our results are pertinent to the field of tissue-engineering, where
cell-seeded 3D scaffolds are commonly used for effective replacement/repair of tissue (Hutmacher, 2000). Better understanding of
the factors that regulate elastic and viscoelastic properties of cells
will facilitate tighter control over mechanical interplay between
compliant scaffolds and cells and may reduce negative cellinduced mechanical effects commonly reported, such as scaffold
deformation, shrinkage, tearing, and resultant redefinition of
scaffolds properties (Bell et al., 1979; Levy-Mishali et al., 2008;
Pins et al., 2000). In addition, changes in elastic properties of cells
have been linked to the onset/progression of diseases and is
involved in pathogenesis (Dulinska et al., 2006; Miller et al.,
2002). Understanding the mechanisms and resultant changes in
the cell’s mechanical properties is therefore critical to understand
the developmental processes underlying disease progression
(Suresh et al., 2005; Suresh, 2007).
Conflict of interest statement
The authors of this article wish to declare that they have no
personnal relationships with other people or organisations that
could inappropriately influence or compromise the data
presented here.
Acknowledgements
This work was supported by Science Foundation Ireland (Grant
no. 07/IN1/B931). The study sponsors had no involvement in the
study design, in the collection, analysis and interpretation of data,
in writing the article, and in the decision to submit the article for
publication.
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