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\begin{document}

\title{Piezo1-mediated microtubule destabilisation promotes extracellular
matrix rigidity induced smooth muscle cell hypertrophy}



\author[1]{Robert Johnson}%
\author[1]{Sultan Ahmed}%
\author[1]{Finn Wostear}%
\author[1]{Reesha Solanki}%
\author[1]{Christopher Morris}%
\author[1]{Stefan Bidula}%
\author[1]{Derek Warren}%
\affil[1]{University of East Anglia}%


\vspace{-1em}



  
  \date{May 26, 2024}


\begingroup
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\selectlanguage{english}
\begin{abstract}
Decreased aortic compliance is a precursor to numerous cardiovascular
diseases. Compliance is regulated by the rigidity of the aortic wall and
the vascular smooth muscle cells (VSMCs) within it. Extracellular matrix
stiffening, observed during ageing, reduces compliance and contributes
to hypertension. In response to increased rigidity, VSMCs generate
enhanced contractile forces that result in VSMC stiffening and a further
reduction in compliance. Due to a lack of suitable in vitro models, the
mechanisms driving VSMC response to matrix rigidity remain poorly
defined. Human aortic-VSMCs were seeded onto polyacrylamide hydrogels
whose rigidity mimicked either healthy or aged/diseased aortae. VSMC
response to contractile agonist stimulation was measured through changes
in cell area and volume. VSMCs were pre-treated with pharmacological
agents prior to agonist stimulation to identify regulators of VSMC
hypertrophy. VSMCs undergo a differential response to contractile
agonist stimulation based on matrix rigidity. On pliable matrices, VSMCs
contract, decreasing in cell area. Meanwhile, on rigid matrices VSMCs
undergo a hypertrophic response, increasing in area and volume. Piezo1
mediated calcium influx drives VSMC hypertrophy by promoting microtubule
destabilisation. Pharmacological stabilisation of microtubules or
blocking calcium influx prevented VSMC hypertrophy on rigid matrices
whilst maintaining contractility on pliable matrices. In response to
extracellular matrix rigidity, VSMCs undergo a hypertrophic response
driven by piezo1-mediated microtubule destabilisation. Pharmacological
targeting of this response blocks matrix rigidity induced VSMC
hypertrophy whilst VSMC contractility on healthy mimicking matrices is
unimpeded. Through delineating this rigidity-induced mechanism, we
identify novel targets whose pharmacological inhibition may prove
beneficial against VSMC-driven cardiovascular disease.%
\end{abstract}%



\sloppy


\textbf{Piezo1-mediated microtubule destabilisation promotes
extracellular matrix rigidity induced smooth muscle cell hypertrophy}

\textbf{Robert T. Johnson\textsuperscript{1+}, Sultan
Ahmed\textsuperscript{1}, Finn Wostear\textsuperscript{1}, Reesha
Solanki\textsuperscript{1}, Chris J. Morris\textsuperscript{1,2}, Stefan
Bidula\textsuperscript{1} and Derek T. Warren\textsuperscript{1+}}

\textsuperscript{1} School of Pharmacy, University of East Anglia,
Norwich Research Park, Norwich, United Kingdom.

\textsuperscript{2} School of Pharmacy, University College London,
London, United Kingdom.

\textsuperscript{+} Corresponding Author(s): Dr. Robert Johnson
(robert.johnson@uea.ac.uk) \&

Dr. Derek Warren (derek.warren@uea.ac.uk) School of Pharmacy, University
of East Anglia, Norwich Research Park, Norwich, Norfolk, UK, NR4 7TJ

\emph{ORCiD IDs:} RTJ (https://orcid.org/0000-0003-3618-238X)

FW (https://orcid.org/0009-0003-0703-8917)

CJM (https://orcid.org/0000-0002-7703-4474)

SB (https://orcid.org/0000-0002-3790-7138)

DTW (https://orcid.org/0000-0003-0346-7450)

\emph{Running Title:} Microtubules regulate smooth muscle cell
hypertrophy

\emph{Word Count:} 3,683

\emph{Author Contributions:} RTJ, SA, FW and RS were responsible for
performing experiments and analysing data. DTW analysed data. RTJ, CJM,
SB and DTW were involved in the conceptualisation of the study. CJM and
SB reviewed and edited this manuscript. RTJ and DTW were responsible for
the design, writing and editing of this manuscript.

\emph{Acknowledgements:} This work was funded by a Biotechnology and
Biological Sciences Research Council Research Grant (BB/T007699/1) and a
British Heart Foundation PhD Studentship (FS/17/32/32916) awarded to DTW
and a UKRI Biotechnology and Biological Sciences Research Council
Norwich Research Park Doctoral Training Partnership PhD Studentship
awarded to FW.

\emph{Conflict of Interests:} The authors declare that the research was
conducted in the absence of any commercial or financial relationships
that could be construed as a potential conflict of interest.

\emph{Data Availability:} The data that support the findings of this
study are available from the corresponding author upon reasonable
request.

\emph{Abstract}

\textbf{Background and Purpose:}Decreased aortic compliance is a
precursor to numerous cardiovascular diseases. Compliance is regulated
by the rigidity of the aortic wall and the vascular smooth muscle cells
(VSMCs) within it. Extracellular matrix stiffening, observed during
ageing, reduces compliance and contributes to hypertension. In response
to increased rigidity, VSMCs generate enhanced contractile forces that
result in VSMC stiffening and a further reduction in compliance. Due to
a lack of suitable \emph{in vitro} models, the mechanisms driving VSMC
response to matrix rigidity remain poorly defined.

\textbf{Experimental Approach:} Human aortic-VSMCs were seeded onto
polyacrylamide hydrogels whose rigidity mimicked either healthy or
aged/diseased aortae. VSMC response to contractile agonist stimulation
was measured through changes in cell area and volume. VSMCs were
pre-treated with pharmacological agents prior to agonist stimulation to
identify regulators of VSMC hypertrophy.

\textbf{Key Results:} VSMCs undergo a differential response to
contractile agonist stimulation based on matrix rigidity. On pliable
matrices, VSMCs contract, decreasing in cell area. Meanwhile, on rigid
matrices VSMCs undergo a hypertrophic response, increasing in area and
volume. Piezo1 mediated calcium influx drives VSMC hypertrophy by
promoting microtubule destabilisation. Pharmacological stabilisation of
microtubules or blocking calcium influx prevented VSMC hypertrophy on
rigid matrices whilst maintaining contractility on pliable matrices.

\textbf{Conclusions and Implications:}In response to extracellular
matrix rigidity, VSMCs undergo a hypertrophic response driven by
piezo1-mediated microtubule destabilisation. Pharmacological targeting
of this response blocks matrix rigidity induced VSMC hypertrophy whilst
VSMC contractility on healthy mimicking matrices is unimpeded. Through
delineating this rigidity-induced mechanism, we identify novel targets
whose pharmacological inhibition may prove beneficial against
VSMC-driven cardiovascular disease.

\emph{Introduction}

Aortic compliance describes the ability of the aorta to change shape in
response to changes in blood pressure. Maintaining aortic compliance is
essential for cardiovascular (CV) health and decreased aortic compliance
is a major risk factor associated with a variety of age-related CV
diseases (Glasser et al., 1997; Mitchell et al., 2010; Lacolley et al.,
2020). Clinical measurements of aortic compliance, using pulse wave
velocity, identify that reduced compliance is associated with increased
CV mortality (Safar et al., 2002; Zhong et al., 2018; Sequ\selectlanguage{ngerman}í-Domínguez et
al., 2020). The rigidity of the aortic wall is a major contributor to
aortic compliance. In the healthy aortic wall, rigidity and compliance
are determined by the balance between extracellular elastic fibres,
including elastin, which provides pliability, and non-elastic
extracellular matrix (ECM) components, including collagen-I, that
provides tensile strength to the aortic wall (Tsamis et al., 2013).
However, during ageing and CV disease, elastic-fibres degrade, and
collagen-I accumulates. These ECM changes increase the rigidity of the
aortic wall and decrease aortic compliance (Ahmed and Warren, 2018).

Vascular tone is regulated by the contraction of vascular smooth muscle
cells (VSMCs), the predominant cell type within the aortic wall (Leloup
et al., 2019). These mechanosensitive cells generate actomyosin-derived
forces, with force production increasing as ECM rigidity increases
(Petit et al., 2019; Sanyour et al., 2019; Johnson et al., 2021).
Enhanced actomyosin force generation increases VSMC stiffness and in
turn contributes to increased aortic wall rigidity and reduced aortic
compliance observed in ageing and CV disease (Qiu et al., 2010; Sehgel
et al., 2015b; Lacolley et al., 2017). In healthy aortae, wall rigidity
and compliance are a balance between ECM rigidity and VSMC stiffness
(Johnson et al., 2021). However, this balance is disrupted in ageing and
CV disease, resulting in VSMC dysfunction (Sazonova et al., 2011;
Lacolley et al., 2017). For example, in hypertension, aortic stiffness
increases as a result of enhanced ECM rigidity and VSMC hypertrophy, a
process through which cell mass increases without a corresponding
increase in cell number (Owens and Schwartz, 1983; Rizzoni et al., 2000;
Zhang et al., 2005; Schiffrin, 2012; Sehgel et al., 2013). VSMC
hypertrophy increases aortic wall thickness and rigidity, resulting in
reducing aortic compliance (Zieman et al., 2005; Hayashi and Naiki,
2009; Sehgel et al., 2015a). However, mechanisms driving VSMC
hypertrophy remain poorly defined.

In response to matrix rigidity, ECM adhesions activate the Rho/ROCK
signalling pathway, resulting in actin polymerisation and myosin light
chain phosphorylation, that in turn enhance actomyosin activity (Ahmed
and Warren, 2018). In other cell types, actomyosin activity and actin
cytoskeleton reorganisation in response to ECM rigidity have been shown
to activate stretch activated ion channels (SACs) (Kobayashi and Sokabe,
2010; Nourse and Pathak, 2017). SAC activation enables mechanical
stimuli to be converted into biochemical responses through mediating the
diffusion of ions, including calcium, across the cell membrane
(Kobayashi and Sokabe, 2010). Whether VSMCs possess a similar response
to ECM rigidity remains unknown, however, VSMCs possess numerous stretch
activated ion channels including piezo1 and members of the TRP
(transient receptor potential) family (Lowis et al., 2023). In normal
physiology, these channels are activated transiently by blood flow
derived stretching of the aortic wall (Liu and Lin, 2022). This
stimulates VSMC contraction as a result of calcium ion influx. These
SACs are also reported to contribute to VSMC dysfunction, with piezo1
implicated in atherosclerosis and abdominal aortic aneurysm induced
vascular remodelling (Qian et al., 2022; Yin et al., 2022). Whether
SACs, including piezo1, are involved in VSMC hypertrophy and the
mechanisms by which they drive VSMC dysfunction remain unknown.

Investigating the cellular response to matrix rigidity requires us to
move away from the use of tissue culture plastic and glass, materials
whose stiffness is around a thousand times greater than that of a
healthy aorta (Minaisah et al., 2016). The rigidity of the aortic wall,
known as its Young's modulus (measured in kilopascals (kPa)), has been
experimentally determined using atomic force microscopy (Hayenga et al.,
2011; Tracqui et al., 2011). In this study, we utilised polyacrylamide
hydrogels, substrates that can be fabricated to the same rigidity as
that of a healthy or diseased aorta (Hayenga et al., 2011; Tracqui et
al., 2011; Minaisah et al., 2016; Rezvani-Sharif et al., 2019). We
identify that enhanced ECM rigidity promotes VSMCs to undergo
hypertrophy following contractile agonist stimulation. VSMC hypertrophy
is driven by piezo1-mediated microtubule instability. Finally,
pharmacological stabilisation of microtubules, or inhibition of piezo1
prevented ECM rigidity induced VSMC hypertrophy whilst leaving VSMC
contractility on healthy mimicking substrates unimpeded.

\emph{Methods}

\textbf{2.1 Polyacrylamide Hydrogel Preparation}

Hydrogels were prepared as described previously (Minaisah et al., 2016).
Glass coverslips were activated by treating with
(3-Aminopropyl)triethoxysilane for 2 minutes, washed 3x in
dH\textsubscript{2}O, then fixed in 0.5\% glutaraldehyde for 40 minutes.
After fixation, coverslips were washed and left to air dry overnight.
Polyacrylamide hydrogel buffer was comprised as follows: 12 kPa -- 7.5\%
acrylamide, 0.15\% bis-acrylamide in dH\textsubscript{2}O; 72 kPa --
10\% acrylamide, 0.5\% bis-acrylamide in dH\textsubscript{2}O. To
prepare hydrogels for fabrication, the appropriate volume of buffer was
supplemented with 10\% APS (1:100) and TEMED (1:1,000) then placed on a
standard microscopy slide and covered by an activated coverslip (13 mm
coverslips required 30 \selectlanguage{greek}µ\selectlanguage{english}l of supplemented buffer; 33 mm coverslips used
50 \selectlanguage{greek}µ\selectlanguage{english}l). Once set, the hydrogels were washed 3x in dH\textsubscript{2}O
to remove any unpolymerized acrylamide, crosslinked with sulfo-SANPAH
(1:3,000) under UV illumination (365 nm) for 5 minutes, then
functionalised with collagen I (0.1 mg/ml) for 10 minutes at room
temperature. Hydrogel stiffnesses have previously been confirmed using a
JPK Nanowizrd-3 atomic force microscope (Porter et al., 2020).

\textbf{2.2 Vascular Smooth Muscle Cell Culture}

Human adult aortic VSMCs were purchased from Cell Applications Inc.
(354-05a). Standard VSMC culture (passages 3-9) was performed as
previously described (Ragnauth et al., 2010; Warren et al., 2015). VSMCs
were seeded onto polyacrylamide hydrogels in basal media (Cell
Applications Inc Cat\# 310-500), 18 hours prior to the beginning of the
experiment. Briefly, VSMCs were pre-treated with pharmacological agents
for 30 minutes, prior to co-treatment with a contractile agonist for an
additional 30 minutes. Experimental specific concentrations are provided
in the corresponding figure legends. Please see \textbf{Supplementary
Table} \textbf{S1} for details of compounds used in this study.
Keratinocyte calcium-free basal medium (Cell Applications Inc
Cat\#132-500) was used for experiments where VSMCs were cultured in the
absence of extracellular calcium.

\textbf{2.3 Immunofluorescence and VSMC Area/Volume Analysis}

Cells were fixed in 4\% paraformaldehyde for 10 minutes, permeabilised
with 0.5\% NP40 for 5 minutes, then blocked with 3\% BSA/PBS for 1 hour.
Primary staining against lamin A/C (1:200) (Sigma-Aldrich Cat\#
SAB4200236, RRID:AB\_10743057) was performed overnight at 4 ?\selectlanguage{english}C in 3\%
BSA/PBS. Secondary staining was performed using the appropriate Alexa
Fluor 488 antibody (1:400) (Thermo Fisher Scientific Cat\# A-11001,
RRID:AB\_2534069) in the dark for 2 hours. F-actin was visualised using
Rhodamine Phalloidin (1:400) (Thermo Fisher Scientific Cat\# R145).
Images were captured at 20x magnification using a Zeiss LSM980-Airyscan
confocal microscope. Cell area and volume was measured using FIJI,
open-source software (Schindelin et al., 2012; Ahmed et al., 2022)

\textbf{2.4 Traction Force Microscopy}

VSMCs were seeded onto polyacrylamide hydrogels containing 0.5 \selectlanguage{greek}µ\selectlanguage{english}m red
fluorescent (580/605) FluoSpheres (1:1000) (Invitrogen). Following
angiotensin II stimulation (30 minutes), cell lysis was achieved by the
addition of 0.5\% Triton X-100. Images were captured at 20x
magnification before and after lysis at 2-minute intervals using a Zeiss
Axio Observer live cell imaging system. Drift was corrected using the
ImageJ StackReg plugin and traction force was calculated using the
ImageJ FTTC plugin that measured FluoSphere displacement (Tseng et al.,
2012). Briefly, bead displacement was measured using the first and last
image of the movie sequence. The cell region was determined by
overlaying the traction map with the phase image, selecting the cell
traction region with an ROI and extracting the traction forces in each
pixel using the XY coordinate function in FIJI (Porter et al., 2020;
Ahmed et al., 2022).

\textbf{2.5 Cold-Stable Microtubule Stability Assay}

Cold-stable microtubules were identified as per previous studies
(Atkinson et al., 2018). Following treatment, cells were placed on ice
for 15 minutes before being washed once with PBS and twice with PEM
buffer (80 \selectlanguage{greek}μ\selectlanguage{english}M PIPES pH 6.8, 1 mM EGTA, 1 mM MgCl\textsubscript{2}, 0.5\%
Triton X-100 and 25\% (w/v) glycerol) for 3 minutes. Cells were fixed in
ice-cold methanol for 20 minutes then blocked with 3\% BSA/PBS for 1
hour. Microtubules were visualised by staining for \selectlanguage{greek}α-\selectlanguage{english}tubulin (1:200)
(Cell Signalling Technology Cat\# 3873, RRID:AB\_1904178) whilst cell
nuclei were visualised using a DAPI containing mounting media. Images
were captured at 40x magnification using a Zeiss AxioPlan 2ie
microscope.

\textbf{2.6 Cell Viability Assay}

Cell viability was determined using a RealTime-Glo MT Cell Viability
Assay, as per manufacturers instructions. Briefly, 5,000 cells per well
seeded in a 96-well plate and exposed to a range of drug concentrations
for 1 hour. Luminescence was subsequently measured using a Wallac
EnVision 2103 Multilabel Reader (PerkinElmer).

\textbf{2.7 siRNA Knockdown}

VSMC siRNA transfection was performed using HiPerFect (Qiagen) as per
manufacturer's instructions, the day before cells were seeded onto
hydrogels. The next afternoon, cells were seeded onto hydrogels as
above, serum was withdrawn overnight to induce quiescence and the next
morning VSMCs were stimulated with Angiotensin II (10 \selectlanguage{greek}µ\selectlanguage{english}M) for 30 minutes
prior to fixation and downstream histochemical analysis.

siRNA \#1 -- CCGCGTCTTCCTTAGCCATTA

siRNA \#2 - CGGCCGCCTCGTGGTCTACAA

\textbf{2.8 Western Blotting}

Western blotting was performed as previously described (Ragnauth et al.,
2010). When looking for Piezo1 expression specifically, lysates were run
on a TruPAGE precast 4-20\% gradient gel (Sigma-Aldrich) at 120 V for 2
hours. Protein was transferred onto PVDF membrane at 30 V for 3 hours
prior to the membrane being blocked in 5\% milk/TBST. The following
antibodies were used: anti-Piezo1 (1:500) (Novus Cat\# NBP1-78537,
RRID:AB\_11003149), anti-GAPDH (1:4000) (Cell Signalling Technology
Cat\# 2118, RRID:AB\_561053) and anti-Rabbit-HRP (1:2000) (Sigma-Aldrich
Cat\# GENA934, RRID:AB\_2722659).

\textbf{2.9 Statistical Analysis}

The data and statistical analysis in this study complies with the
recommendations on experimental design and analysis in pharmacology
(Curtis et al., 2018). Experiments were performed by one researcher,
with a second researcher then performing the microscopy and downstream
analysis. Statistical analysis was performed using GraphPad Prism 9.5.
Results are presented as mean \selectlanguage{ngerman}± SEM, with individual data points shown.
The number of independent repeats performed, and total number of cells
analysed per experiment are detailed in the corresponding figure legend.
Unpaired Student's t-tests were used for the comparison of two
conditions. To compare more than two conditions a one-way ANOVA was
performed, with either a Tukey's or Sidak's multiple comparison post-hoc
test being performed as appropriate. Concentration-response curves are
presented as mean ± SEM plotted on a logarithmic scale. Log(agonist) vs
response curves were generated using non-linear regression. Comparisons
between concentration ranges on different hydrogel stiffness were
performed using a two-way ANOVA followed by Sidak's post-hoc test.
Differences between conditions were considered statistically significant
when \emph{P} \textless{} 0.05.

\emph{Results}

\textbf{3.1 Matrix rigidity alters isolated smooth muscle cell response
to contractile agonist stimulation.}

We set out to determine how enhanced matrix rigidity, akin to that of an
aged/diseased aortic wall, would affect VSMC response to contractile
agonists. Quiescent VSMCs grown on pliable (12 kPa) or rigid (72 kPa)
hydrogels were stimulated with increasing concentrations of the
contractile agonist angiotensin II. Changes in VSMC area were used as a
measure of contractile response (Ahmed et al., 2022). As previously
observed, VSMCs on pliable hydrogels contracted, indicated by a decrease
in cell area as angiotensin II concentration increased (\textbf{Figure
1a, b \& d} ) (Ahmed et al., 2022). In contrast, VSMCs seeded on rigid
hydrogels failed to contract (\textbf{Figure 1a, c \& d} ). VSMCs seeded
on rigid hydrogels were initially smaller than those on pliable
hydrogels, yet when exposed to increasing concentrations of angiotensin
II, cell area increased (\textbf{Figure 1a \& d} ). Stimulation of VSMCs
with increasing concentrations of an alternative contractile agonist,
carbachol, again resulted in a differential response whereby VSMC area
was reduced on pliable hydrogels but increased on rigid hydrogels
(\textbf{Figure 1e-h} ). Subsequent experiments were performed by
stimulating VSMCs with 10 \selectlanguage{greek}μ\selectlanguage{english}M of either angiotensin II or carbachol, a
concentration which induced maximal area changes on both rigidities of
hydrogel.

To confirm that the above changes were specific for receptor activation,
we utilised receptor antagonists irbesartan and atropine. Irbesartan
antagonises the angiotensin II type 1 receptor, AT1, whilst atropine
antagonises acetylcholine receptors thereby blocking the effects of
carbachol. Quiescent VSMCs grown on pliable or rigid hydrogels were
stimulated with either angiotensin II or carbachol in the presence of an
increasing concentration of their respective antagonist. On pliable
hydrogels, increasing concentrations of irbesartan or atropine prevented
VSMCs from undergoing a contractile response (\textbf{Figure 2a, b, e \&
f} ). Likewise, treatment with irbesartan or atropine prevented
contractile agonist induced enlargement of VSMCs on rigid hydrogels
(\textbf{Figure 2a, c, e \& g} ).

\textbf{3.2 Isolated smooth muscle cells undergo a hypertrophic response
on rigid substrates following contractile agonist stimulation.}

The above data demonstrates that VSMC response to contractile agonist
stimulation is regulated by matrix rigidity. We next sought to determine
whether VSMC volume, as well as area was enlarged following contractile
agonist stimulation on rigid substrates. Quiescent VSMCs were seeded on
pliable and rigid hydrogels and stimulated with angiotensin II. Confocal
microscopy was used to measure VSMC volume. As previously observed
(Ahmed et al., 2022), VSMCs on pliable hydrogels underwent a contractile
response following angiotensin II stimulation, decreasing in cell area
but displaying no change in volume (\textbf{Figure 3a-c} ). In contrast,
angiotensin II stimulation of VSMCs on rigid hydrogels resulted in both
cell area and volume enlargement, indicative of a hypertrophic response
(\textbf{Figure 3a-c} ). On both rigidities of hydrogel, angiotensin II
stimulated VSMCs displayed a positive correlation between cell area and
volume (\textbf{Figure 3d} ). Linear regression revealed that the
correlation between VSMC area and volume was similar on both matrices
(12 kPa + AngII R\textsuperscript{2} = 0.6186, 72 kPa + AngII
R\textsuperscript{2} = 0.6023) (\textbf{Figure 3d} ).

\textbf{3.3 Extracellular matrix rigidity regulates microtubule
stability within isolated smooth muscle cells.}

Based on the morphological changes observed above, contractile agonist
stimulation promotes VSMC contraction on pliable hydrogels and VSMC
hypertrophy on rigid hydrogels. Previous studies have shown that matrix
rigidity promotes increased VSMC traction stress generation (Sazonova et
al., 2015; Petit et al., 2019). To confirm that angiotensin II
stimulated VSMCs seeded on rigid hydrogels generated enhanced traction
stresses, we next performed traction force microscopy. Analysis revealed
that VSMCs seeded on rigid hydrogels generated greater maximal and total
traction stress following angiotensin II stimulation, compared to their
counterparts on pliable hydrogels (\textbf{Supplementary Figure S1} ).
As cells generate actomyosin derived traction forces, deformational
stresses are also placed upon the cell membrane (Johnson et al., 2021).
Microtubules exist in a mechanical balance with actomyosin activity,
serving as compression bearing struts capable of resisting
actomyosin-generated deformational stresses (Stamenovi\selectlanguage{polish}ć, \selectlanguage{english}2005;
Brangwynne et al., 2006; Johnson et al., 2021). This relationship is
defined by the tensegrity model which predicts that microtubule
destabilisation will lead to greater actomyosin derived force generation
(Stamenovi\selectlanguage{polish}ć, \selectlanguage{english}2005). As VSMCs on rigid hydrogels generate enhanced
traction stresses, we predicted that decreased microtubule stability may
contribute to increased traction stress generation. To test this, we
performed a cold-stable microtubule assay on isolated VSMCs in the
presence or absence of angiotensin II stimulation. Upon exposure to cold
temperatures, microtubules readily undergo catastrophe, with only
stabilised microtubule filaments remaining. After clearing the tubulin
monomers, the number of stabilised microtubules can be counted (Ochoa et
al., 2011). On pliable hydrogels, angiotensin II stimulation had no
effect on microtubule stability (\textbf{Figure 3e \& f} ). Microtubule
stability was found to increase in unstimulated VSMCs that were seeded
on rigid compared to pliable substrates (\textbf{Figure 3e \& f} ).
However, following angiotensin II stimulation this enhanced stability
was lost, with angiotensin II stimulated VSMCs on rigid hydrogels
displaying similar levels of microtubule stability as cells on pliable
hydrogels (\textbf{Figure 3e \& f} ).

\textbf{3.4 Microtubule destabilisation promotes isolated smooth muscle
cell hypertrophy following contractile agonist stimulation.}

The above data shows that on rigid hydrogels, VSMCs stimulated with
angiotensin II undergo a hypertrophic response that is accompanied by an
increase in traction force generation and a decrease in microtubule
stability. We hypothesised that the loss of microtubule stability was
driving VSMC hypertrophy on rigid substrates. Quiescent VSMCs were
seeded onto pliable and rigid hydrogels and pre-treated with increasing
concentrations of microtubule stabilisers prior to angiotensin II
stimulation. Treatment with two microtubule stabilisers, paclitaxel or
epothilone B, had no effect on the contractile response of VSMCs seeded
on pliable hydrogels (\textbf{Figure 4a, b, d-f \& h} ). Meanwhile,
increasing concentrations of either microtubule stabiliser was
sufficient to prevent the increase in VSMC area observed following
angiotensin II stimulation on rigid hydrogels (\textbf{Figure 4a, c-e, g
\& h} ).

Given that microtubule stabilisation prevented the enlargement of VSMCs
on rigid substrates following angiotensin II stimulation, we then sought
to determine whether microtubule destabilisation would have the opposite
effect. VSMCs were pre-treated with increasing concentrations of the
microtubule destabilisers colchicine or nocodazole, and then stimulated
with angiotensin II. On pliable hydrogels, VSMCs treated with either
colchicine or nocodazole displayed increased cell area following
angiotensin II stimulation (\textbf{Figure 5a, b, d-f \& h} ). In
contrast, treatment with the microtubule destabilisers had no effect on
VSMCs seeded on rigid hydrogels (\textbf{Figure 5a, c-e, g \& h} ). All
microtubule targeting agents were used at concentrations that did not
cause cell death, as confirmed through a viability assay for
concentrations of epothilone B and nocodazole (\textbf{Supplementary
Figure S2} ) or previously for paclitaxel and colchicine (Ahmed et al.,
2022).

Having determined that microtubule stability regulated changes in VSMC
area, we next sought to measure changes in VSMC volume and subsequently
confirm if microtubule instability was driving the hypertrophic
response. Quiescent VSMCs were pre-treated with concentrations of
paclitaxel or colchicine that altered both VSMC area (\textbf{Figure
4a-d, 5a-d} ) and caused a detectable change in the number of
cold-stable microtubules (\textbf{Supplementary Figure S3} ). Following
pre-treatment, VSMCs were stimulated with angiotensin II and confocal
microscopy was used to assess changes in cell volume. Treatment with the
microtubule stabiliser paclitaxel had no effect on the volume of VSMCs
seeded on pliable hydrogels (\textbf{Figure 6a-c} ). In contrast,
microtubule stabilisation prevented the angiotensin II induced increase
in VSMC volume on rigid hydrogels (\textbf{Figure 6a-c} ). Meanwhile,
microtubule destabilisation, via colchicine pre-treatment, increased
VSMC volume on pliable hydrogels following angiotensin II stimulation,
indicative of a hypertrophic response. No further increase in VSMC
volume following microtubule destabilisation was observed in VSMCs
seeded on rigid hydrogels (\textbf{Figure 6d-f} ).

\textbf{3.5 Extracellular calcium ion influx promotes VSMC hypertrophy
on rigid hydrogels}

The above data shows that in response to angiotensin II stimulation,
VSMCs undergo contraction on pliable hydrogels, but display a
hypertrophic response on rigid hydrogels. Angiotensin II mediated
activation of the AT1 receptor drives calcium ion
(Ca\textsuperscript{2+}) release from the sarcoplasmic reticulum,
increasing cytosolic Ca\textsuperscript{2+} levels (Woodrum and Brophy,
2001). Therefore, we next investigated whether ECM rigidity promoted
differences in intracellular calcium handling. Quiescent VSMCs were
pre-treated with an increasing concentration of either the ryanodine
receptor antagonist, dantrolene, or the IP\textsubscript{3} channel
blocker, xestospongin C, prior to angiotensin II stimulation. On pliable
hydrogels, increasing concentrations of dantrolene or xestospongin C
prevented VSMCs from undergoing a contractile response and decreasing in
cell area (\textbf{Supplementary Figure S4a, b, d-f \& h} ). Likewise,
treatment with dantrolene or xestospongin C prevented the angiotensin II
induced enlargement of VSMCs on rigid hydrogels (\textbf{Supplementary
Figure S4a, c-e, g \& h} ). Neither compound had any effect on VSMC
viability (\textbf{Supplementary Figure S5a \& b} ). This suggests that
sarcoplasmic Ca\textsuperscript{2+} release is a common component of
both the contractile and hypertrophic response.

ECM rigidity is known to promote the activation of SACs, enabling the
influx of extracellular Ca\textsuperscript{2+} in cardiac fibroblasts
(Stewart and Turner, 2021). To test whether this was also true in VSMCs,
we next utilised the SAC blocker, GsMTx-4. Quiescent VSMCs were
pre-treated with an increasing concentration of GsMTx-4 prior to
angiotensin II treatment on pliable and rigid hydrogels. Analysis
revealed that GsMTx-4 pre-treatment had no effect on VSMC area on
pliable hydrogels (\textbf{Figure 7a, b \& d} ). In contrast, GsMTx-4
pre-treatment blocked the increase in VSMC area on rigid hydrogels in a
concentration dependent manner (\textbf{Figure 7a, c \& d} ). GsMTx-4
pre-treatment had no effect on the viability of VSMCs
(\textbf{Supplementary Figure S5c} ). To confirm that changes in VSMC
area correlated with changes in volume, we next performed confocal
microscopy on GsMTx-4 pre-treated VSMCs. SAC blockade had no effect on
angiotensin II treated VSMC area or volume on pliable hydrogels
(\textbf{Figure 7e-f} ). However, on rigid hydrogels, GsMTx-4
pre-treatment blocked the angiotensin II induced increases in VSMC area
and volume (\textbf{Figure 7e-f} ).

\textbf{3.6 Piezo1-mediated calcium influx destabilises microtubules and
drives VSMC hypertrophy}

The above data shows that VSMC hypertrophy on rigid substrates is
mediated by SAC activation. Piezo1 is a SAC known to be involved in
atherosclerosis and abdominal aortic aneurysm mediated VSMC dysfunction
(Qian et al., 2022; Yin et al., 2022). Whether piezo1 is involved in
VSMC hypertrophy remains unknown. We therefore investigated its
potential role in regulating VSMC response to ECM rigidity by utilising
an siRNA mediated knockdown approach. Western blotting confirmed that
two independent siRNAs efficiently depleted piezo1 in VSMCs
(\textbf{Figure 8a \& b} ). Analysis revealed that piezo1 depletion had
no effect on angiotensin II induced VSMC contractility on pliable
hydrogels, with no change in VSMC area or volume detected.
(\textbf{Figure 8c-e} ). In contrast, piezo1 depletion blocked the
angiotensin II mediated increase in VSMC area and volume on rigid
hydrogels (\textbf{Figure 8c-e} ). These data confirm that piezo1 is a
novel mediator of VSMC hypertrophy. Finally, we sought to determine if
piezo1 activity was driving VSMC hypertrophy through the induction of
microtubule instability. To test this idea, we performed cold-stable
microtubule assays on VSMCs that were either pre-treated with GsMTx-4 or
depleted for piezo1. Analysis revealed that GsMTx-4 mediated SAC
blockade (\textbf{Figure 9a \& b} ) or piezo1 depletion (\textbf{Figure
9c \& d} ) restored microtubule stability on VSMCs seeded on rigid
hydrogels whilst having no effect on those seeded on pliable hydrogels.

To confirm that Ca\textsuperscript{2+} influx was driving microtubule
destabilisation, we performed a cold-stable microtubule assay in the
presence or absence of extracellular Ca\textsuperscript{2+}. The absence
of extracellular Ca\textsuperscript{2+} had no effect on the number of
cold-stable microtubules detected in angiotensin II stimulated VSMCs on
pliable hydrogels (\textbf{Figure 9e \& f} ). In contrast, the number of
cold-stable microtubules increased within angiotensin II stimulated
VSMCs on rigid hydrogels when extracellular Ca\textsuperscript{2+} was
absent (\textbf{Figure 9e \& f} ). This suggests that piezo1-mediated
Ca\textsuperscript{2+} influx decreases microtubule stability and
induces VSMC hypertrophy on rigid hydrogels following angiotensin II
stimulation.

\emph{Discussion}

Our understanding of the mechanisms driving VSMC dysfunction and its
contribution to decreased aortic compliance in ageing and CV disease
remains limited. Progress has been hindered by a lack of easy to
use,\emph{in vitro} tools through which pathological mechanisms can be
investigated. In this study, we utilised polyacrylamide hydrogels whose
rigidity (pliable 12 kPa, rigid 72 kPa) mimicked the respective
stiffness of healthy and aged/diseased aortic walls (Hayenga et al.,
2011; Tracqui et al., 2011; Minaisah et al., 2016; Rezvani-Sharif et
al., 2019). Polyacrylamide hydrogels are biologically inert and can be
easily fabricated to a specific rigidity in-house, using generic
research equipment and skills (Kandow et al., 2007; Caliari and Burdick,
2016; Minaisah et al., 2016; Mohammed et al., 2019). This makes them
ideal substrates for investigating how ECM rigidity regulates cellular
function. Previous studies have shown that ECM rigidity promotes the
dedifferentiation of VSMCs, downregulating contractile markers whilst
increasing the expression of proliferative genes (Brown et al., 2010;
Sazonova et al., 2015; Nagayama and Nishimiya, 2020). Increased VSMC
migration, adhesion and proliferation have also been reported (Wong et
al., 2003; Brown et al., 2010; Sazonova et al., 2015; Nagayama and
Nishimiya, 2020; Rickel et al., 2020). Furthermore, in response to
matrix rigidity, VSMC reorganise their actin cytoskeleton and generate
enhanced traction stresses, a finding we recapitulate in this study
(Brown et al., 2010; Sazonova et al., 2015; Petit et al., 2019; Sanyour
et al., 2019). We now show that contractile agonist stimulation of
quiescent VSMCs on pliable (healthy) or rigid (aged/diseased) matrices
results in a differential response; VSMCs on pliable matrices decreased
in area as they undergo contraction, whereas increased cell area was
observed in VSMCs on rigid matrices. Furthermore, on rigid matrices,
contractile agonist stimulation also promoted an increase in VSMC
volume. These findings suggest that changes in VSMC morphology following
contractile agonist stimulation are differentially regulated on pliable
and rigid matrices. On pliable matrices, actomyosin activity drives
contraction and VSMC area subsequently decreases but volume remains
unchanged. In contrast, on rigid matrices, changes in VSMC morphology
are driven by enhanced volume. Importantly, we identify for the first
time that enhanced matrix rigidity initiates a hypertrophic response
within VSMCs, akin to that seen \emph{in vivo} in response to
hypertension (Owens and Schwartz, 1983; Rizzoni et al., 2000; Zhang et
al., 2005; Schiffrin, 2012).

Hypertension is regarded as one of the strongest risk factors for the
progression of CV disease (Kjeldsen, 2018). VSMCs dedifferentiate as a
result of hypertension, becoming dysfunctional and hypertrophic (Touyz
et al., 2018). VSMC hypertrophy contributes to aortic wall thickening,
increased aortic stiffness and reduced aortic compliance, which in turn
leads to hypertension (Brown et al., 2018). The causality between VSMC
hypertrophy and hypertension is cyclic in nature, with the development
of one promoting the onset of the other. Current methods of
investigating VSMC hypertrophy generally utilise animal models,
including rat models of hypertension and genetic depletion in mice
(Owens and Schwartz, 1983; Hixon et al., 2000; Choi et al., 2019; Bai et
al., 2021). We now show that the \emph{in vitro} induction of VSMC
hypertrophy can be achieved through culturing cells on rigid matrices.
This enables the mechanisms regulating VSMC hypertrophy and the
potential of anti-hypertrophic compounds to be investigated at a greater
throughput whilst using a less invasive and more ethical method.
Importantly, we correlated increased VSMC area following angiotensin II
stimulation with an increase in VSMC volume. This enabled us to predict
potential beneficial or deleterious effects of pharmacological agents on
VSMC hypertrophy, prior to confirming those predictions through
3-dimensional confocal microscopy. Through this method, we identified
novel regulators of VSMC hypertrophy using physiologically or
pathologically relevant ECM rigidities.

In hypertension, VSMC Ca\textsuperscript{2+} handling is known to become
dysregulated, with VSMCs entering a hypercontractile state that further
increases aortic rigidity whilst simultaneously reducing aortic
compliance (Touyz et al., 2018). Our findings suggest that the internal
release of Ca\textsuperscript{2+} from the sarcoplasmic reticulum,
following angiotensin II stimulation, is unperturbed by increased ECM
rigidity. Instead, VSMC hypertrophy is driven by the extracellular
influx of Ca\textsuperscript{2+} through mechanosensitive SACs.
Specifically, we identify piezo1 as a driver of ECM rigidity induced
VSMC hypertrophy. VSMC piezo1 expression is upregulated in mouse models
of atherosclerosis and abdominal aortic aneurysms, with dysregulation of
VSMC mechanosensation shown to drive these disease states (Qian et al.,
2022; Yin et al., 2022). Furthermore, reduced aortic compliance is
observed in both atherosclerosis and abdominal aortic aneurysms (Glasser
et al., 1997; Mitchell et al., 2010; Lacolley et al., 2017). Enhanced
VSMC actomyosin force generation is known to contribute to reduced
aortic compliance with inhibition of actomyosin activity resulting in
reduced aortic wall rigidity (Qiu et al., 2010). Our study now suggests
that VSMC hypertrophy may also contribute to the increased aortic wall
rigidity observed in ageing and CV disease. Furthermore, we identify
piezo1 as a critical driver of VSMC hypertrophy in response to increased
ECM rigidity. Pharmacological blockade of piezo1, may prove beneficial
and could form part of a potential therapeutic strategy to reduce VSMC
hypertrophy, decrease aortic wall rigidity and restore aortic
compliance. Further research to test these findings in \emph{ex
vivo}settings is an essential next step in confirming the validity of
our findings.

VSMC response to increased actomyosin-derived force generation on rigid
hydrogels is predicted by the tensegrity model. In this model,
microtubules serve as compression bearing struts that resist actomyosin
generated strain (Johnson et al., 2021). Healthy VSMC behaviour is
therefore a balance between microtubule stability and actomyosin
activity; actomyosin activity drives VSMC contraction while the
microtubules maintain VSMC morphology and protect against strain induced
cellular damage (Stamenovi\selectlanguage{polish}ć, \selectlanguage{english}2005; Brangwynne et al., 2006). The
tensegrity model also predicts that microtubule destabilisation results
in increased traction stress generation, a hypothesis previously
confirmed by both wire myography and traction force microscopy (Paul et
al., 2000; Zhang et al., 2000; Platts et al., 2002; Ahmed et al., 2022).
We show that the tensegrity model drives VSMC hypertrophy on rigid
matrices, where angiotensin II stimulation triggers microtubule
destabilisation. Importantly, treatment with microtubule stabilising
agents blocked VSMC hypertrophy on rigid matrices whilst maintaining
VSMC contractility on pliable matrices. This suggests that therapeutic
pathways that promote microtubule stabilisation may be a potential novel
approach to target VSMC hypertrophy specifically in areas of aortic
stiffness. Current antihypertensive therapies target the
renin-angiotensin system and either prevent the production of
angiotensin II or block the activation of the AT1 receptor
(Hern\selectlanguage{ngerman}ández-Hernández et al., 2002). Whilst these therapies are effective
and well tolerated, they target healthy and dysfunctional VSMCs alike.

Whilst microtubule stabilisation blocked VSMC hypertrophy, treatment
with microtubule destabilising agents had the opposite effect and
induced VSMC hypertrophy on pliable matrices. Colchicine, a microtubule
destabilising agent, is currently being researched for its CV benefits.
Colchicine treatment has been experimentally shown to protect against
endothelial cell dysfunction (Huang et al., 2014; Kajikawa et al., 2019;
Z\selectlanguage{polish}ă\selectlanguage{english}lar et al., 2022), reduce VSMC proliferation and migration \emph{in
vitro} (Zhang et al., 2022), and supress atherosclerotic plaque
development in some animal models (Mart\selectlanguage{ngerman}ínez et al., 2018). However, wire
myography and traction force microscopy both show that colchicine
treatment also increases VSMC isomeric force generation (Zhang et al.,
2000; Platts et al., 2002; Ahmed et al., 2022), suggesting that
colchicine treatment will increase aortic wall rigidity and decrease
aortic compliance. VSMC hypertrophy may subsequently be an unwanted side
effect of any potential colchicine therapy.

Our data suggest that microtubule stability serves as a hypertrophic
switch, in VSMCs destabilisation promotes hypertrophy, whereas
stabilisation blocks hypertrophy. Previous studies have shown that
microtubule catastrophe can be driven by increased levels of cytosolic
Ca\textsuperscript{2+} (O'Brien et al., 1997; Thompson et al., 2014). In
this study, we identify that on rigid matrices, angiotensin II
stimulation of VSMCs leads to an influx of Ca\textsuperscript{2+}through
mechanosensitive piezo1 channels. Blocking the influx of
Ca\textsuperscript{2+} maintains microtubule stability and in turn
prevents VSMC hypertrophy. These findings pave the wave for confirmatory
studies in \emph{ex vivo} and \emph{in vivo} models. Furthermore,
through comparing the response of VSMCs on pliable and rigid matrices we
not only identify potential anti-hypertrophic targets, but also identify
side effects of proposed therapies for the treatment of CV and other
diseases. Although we have established a role for piezo1 and microtubule
stability in regulating VSMC hypertrophy, further research is required
to elucidate the molecular mechanisms through which ECM rigidity drives
this response and how Ca\textsuperscript{2+} regulates microtubule
stability. Developing a better understanding of the connection between
Ca\textsuperscript{2+} and microtubule stability will enhance our
mechanistic understanding of VSMC hypertrophy and will likely identify
further targets that have the therapeutic potential to restore aortic
compliance in ageing and CV disease.

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\emph{Figure Legends}

\textbf{Figure 1. VSMC response to contractile agonist stimulation is
matrix stiffness dependent.} Representative images of isolated VSMCs
cultured on 12 or 72 kPa polyacrylamide hydrogels. Actin cytoskeleton
(purple) and Lamin A/C labelled nuclei (green). Scale bar = 100 \selectlanguage{greek}μ\selectlanguage{english}m. (a)
VSMCs were treated with increasing concentrations of angiotensin II
(0.01 -- 100 \selectlanguage{greek}µ\selectlanguage{english}M) for 30 minutes. VSMC area on (b) 12 kPa and (c) 72 kPa
hydrogels representative of 5 independent experiments with [?]158 cells
analysed per condition. Significance determined using a one-way ANOVA
followed by Tukey's test. (d) Comparison of VSMC response to angiotensin
II on 12 and 72 kPa hydrogels. Data is expressed as the mean of the
means calculated from 5 independent experiments; significance determined
using a two-way ANOVA followed by Sidak's test. (e) VSMCs were treated
with increasing concentrations of carbachol (0.01 -- 100 \selectlanguage{greek}µ\selectlanguage{english}M) for 30
minutes. VSMC area on (f) 12 kPa and (g) 72 kPa hydrogels representative
of 5 independent experiments with [?]149 cells analysed per condition.
Significance determined using a one-way ANOVA followed by Tukey's test.
(h) Comparison of VSMC response to carbachol on 12 and 72 kPa hydrogels.
Data is expressed as the mean of the means calculated from 5 independent
experiments; significance determined using a two-way ANOVA followed by
Sidak's test. (* = \emph{p \textless{} 0.05} , error bars represent \selectlanguage{ngerman}±
SEM).

\textbf{Figure 2. VSMC response to contractile agonist stimulation is
blocked by receptor antagonism.} Actin cytoskeleton (purple) and Lamin
A/C labelled nuclei (green). Scale bar = 100 \selectlanguage{greek}μ\selectlanguage{english}m. (a) Representative
images of isolated VSMCs cultured on 12 or 72 kPa polyacrylamide
hydrogels treated with angiotensin II (10 \selectlanguage{greek}μ\selectlanguage{english}M) for 30 minutes in the
presence of increasing concentrations of irbesartan (0.023 -- 230 nM).
VSMC area on (b) 12 kPa and (c) 72 kPa hydrogels representative of 5
independent experiments with [?]97 cells analysed per condition.
Significance determined using a one-way ANOVA followed by Tukey's test.
(d) Comparison of VSMC response to angiotensin II in the presence of
irbesartan on 12 and 72 kPa hydrogels. Data is expressed as the mean of
the means calculated from 5 independent experiments; significance
determined using a two-way ANOVA followed by Sidak's test. (e)
Representative images of isolated VSMCs cultured on 12 or 72 kPa
polyacrylamide hydrogels treated with carbachol (10 \selectlanguage{greek}μ\selectlanguage{english}M) for 30 minutes
in the presence of increasing concentrations of atropine (0.038 -- 380
nM). VSMC area on (f) 12 kPa and (g) 72 kPa hydrogels representative of
5 independent experiments with [?]93 cells analysed per condition.
Significance determined using a one-way ANOVA followed by Tukey's test.
(h) Comparison of VSMC response to carbachol in the presence of atropine
on 12 and 72 kPa hydrogels. Data is expressed as the mean of the means
calculated from 5 independent experiments; significance determined using
a two-way ANOVA followed by Sidak's test. (* = \emph{p \textless{} 0.05}
, error bars represent \selectlanguage{ngerman}± SEM).

\textbf{Figure 3. VSMC undergo a hypertrophic response on rigid
substrates following angiotensin II stimulation.} (a) Representative
images of isolated VSMCs cultured on 12 or 72 kPa polyacrylamide
hydrogels treated with angiotensin II (AngII) (10 \selectlanguage{greek}µ\selectlanguage{english}M). Actin
cytoskeleton (purple) and Lamin A/C labelled nuclei (green). Top --
Representative XY images of VSMC area, scale bar = 100 \selectlanguage{greek}μ\selectlanguage{english}m. Bottom --
Representative XZ images of VSMC height, scale bar = 20 \selectlanguage{greek}μ\selectlanguage{english}m. (b) VSMC
area and (c) volume, representative of 5 independent experiments with
[?]82 cells analysed per condition. Significance determined using a
one-way ANOVA followed by Sidak's test. (d) Correlation between area and
volume in angiotensin II stimulated VSMCs on 12 and 72 kPa hydrogels.
(e) Representative images of isolated VSMCs seeded on 12 or 72 kPa
polyacrylamide hydrogels in the presence or absence of AngII (10 \selectlanguage{greek}µ\selectlanguage{english}M)
stimulation. Cold-stable microtubules, (\selectlanguage{greek}α-\selectlanguage{english}tubulin, aqua) and nuclei
(DAPI, blue). Scale bar = 50 \selectlanguage{greek}μ\selectlanguage{english}m. (f) number of cold-stable microtubules
per cell, representative of 5 independent experiments, with [?]81 cells
analysed per condition; significance determined using a two-way ANOVA
followed by Sidak's test. (* = \emph{p \textless{} 0.05} , error bars
represent \selectlanguage{ngerman}± SEM).

\textbf{Figure 4. Microtubule stabilisation prevents VSMC enlargement on
rigid substrates following angiotensin II stimulation.} Representative
images of isolated VSMCs cultured on 12 or 72 kPa polyacrylamide
hydrogels stimulated with angiotensin II (10 \selectlanguage{greek}µ\selectlanguage{english}M). Actin cytoskeleton
(purple) and Lamin A/C labelled nuclei (green). Scale bar = 100 \selectlanguage{greek}μ\selectlanguage{english}m. (a)
VSMCs were pre-treated with increasing concentrations of paclitaxel
(0.001 -- 10 nM). VSMC area on (b) 12 kPa and (c) 72 kPa hydrogels
representative of 5 independent experiments with [?]100 cells analysed per
condition. Significance determined using a one-way ANOVA followed by
Tukey's test. (d) Comparison of VSMC response to paclitaxel
pre-treatment on 12 and 72 kPa hydrogels. Data is expressed as the mean
of the means calculated from 5 independent experiments; significance
determined using a two-way ANOVA followed by Sidak's test. (e) VSMCs
were pre-treated with increasing concentrations of epothilone B (0.001
-- 10 nM). VSMC area on (f) 12 kPa and (g) 72 kPa hydrogels
representative of 5 independent experiments with [?]53 cells analysed per
condition. Significance determined using a one-way ANOVA followed by
Tukey's test. (h) Comparison of VSMC response to epothilone B
pre-treatment on 12 and 72 kPa hydrogels. Data is expressed as the mean
of the means calculated from 5 independent experiments; significance
determined using a two-way ANOVA followed by Sidak's test. (n.s.
=\emph{non-significant} , * = \emph{p \textless{} 0.05} , error bars
represent \selectlanguage{ngerman}± SEM).

\textbf{Figure 5. Microtubule destabilisation prevents VSMC contraction
on pliable substrates following angiotensin II
stimulation.}Representative images of isolated VSMCs cultured on 12 or
72 kPa polyacrylamide hydrogels stimulated with angiotensin II (10 \selectlanguage{greek}µ\selectlanguage{english}M).
Actin cytoskeleton (purple) and Lamin A/C labelled nuclei (green). Scale
bar = 100 \selectlanguage{greek}μ\selectlanguage{english}m. (a) VSMCs were pre-treated with increasing concentrations
of colchicine (0.1 -- 1000 nM). VSMC area on (b) 12 kPa and (c) 72 kPa
hydrogels representative of 5 independent experiments with [?]79 cells
analysed per condition. Significance determined using a one-way ANOVA
followed by Tukey's test. (d) Comparison of VSMC response to paclitaxel
pre-treatment on 12 and 72 kPa hydrogels. Data is expressed as the mean
of the means calculated from 5 independent experiments; significance
determined using a two-way ANOVA followed by Sidak's test. (e) VSMCs
were pre-treated with increasing concentrations of nocodazole (0.001 --
10 nM). VSMC area on (f) 12 kPa and (g) 72 kPa hydrogels representative
of 5 independent experiments with [?]58 cells analysed per condition.
Significance determined using a one-way ANOVA followed by Tukey's test.
(h) Comparison of VSMC response to nocodazole pre-treatment on 12 and 72
kPa hydrogels. Data is expressed as the mean of the means calculated
from 5 independent experiments; significance determined using a two-way
ANOVA followed by Sidak's test. (n.s. = \emph{non-significant} , *
=\emph{p \textless{} 0.05} , error bars represent \selectlanguage{ngerman}± SEM).

\textbf{Figure 6. VSMC hypertrophy is regulated by microtubule
stability.} Representative images of isolated VSMCs cultured on 12 or 72
kPa polyacrylamide hydrogels stimulated with angiotensin II (AngII) (10
\selectlanguage{greek}µ\selectlanguage{english}M). Actin cytoskeleton (purple) and Lamin A/C labelled nuclei (green).
Top -- Representative XY images of VSMC area, scale bar = 100 \selectlanguage{greek}μ\selectlanguage{english}m. Bottom
-- Representative XZ images of VSMC height, scale bar = 20 \selectlanguage{greek}μ\selectlanguage{english}m. (a)
Paclitaxel (1 nM) pre-treated VSMCs. (b) VSMC area and (c) volume,
representative of 5 independent experiments with [?]114 cells analysed per
condition. (d) Colchicine (100 nM) pre-treated VSMCs. (e) VSMC area and
(f) volume, representative of 5 independent experiments with [?]113 cells
analysed per condition. Significance determined using a one-way ANOVA
followed by Sidak's test. (* = \emph{p \textless{} 0.05} , error bars
represent \selectlanguage{ngerman}± SEM).

\textbf{Figure 7. Extracellular calcium influx promotes VSMC hypertrophy
on rigid substrates.} Representative images of isolated VSMCs cultured
on 12 or 72 kPa polyacrylamide hydrogels stimulated with angiotensin II
(AngII) (10 \selectlanguage{greek}µ\selectlanguage{english}M). Actin cytoskeleton (purple) and Lamin A/C labelled
nuclei (green). (a) VSMCs were pre-treated with increasing
concentrations of GsMTx-4 (0.5 -- 5000 nM). Scale bar = 100 \selectlanguage{greek}μ\selectlanguage{english}m. VSMC
area on (b) 12 kPa and (c) 72 kPa hydrogels representative of 5
independent experiments with [?]72 cells analysed per condition.
Significance determined using a one-way ANOVA followed by Tukey's test.
(d) Comparison of VSMC response to GsMTx-4 pre-treatment on 12 and 72
kPa hydrogels. Data is expressed as the mean of the means calculated
from 5 independent experiments; significance determined using a two-way
ANOVA followed by Sidak's test. (e) GsMTx-4 (500 nM) pre-treated VSMCs.
Top -- Representative XY images of VSMC area, scale bar = 100 \selectlanguage{greek}μ\selectlanguage{english}m. Bottom
-- Representative XZ images of VSMC height, scale bar = 20 \selectlanguage{greek}μ\selectlanguage{english}m. (f) VSMC
area and (g) volume, representative of 5 independent experiments with
[?]116 cells analysed per condition. Significance determined using a
one-way ANOVA followed by Sidak's test. (* = \emph{p \textless{} 0.05} ,
error bars represent \selectlanguage{ngerman}± SEM).

\textbf{Figure 8. Piezo1 regulates VSMC hypertrophy in response to ECM
rigidity.} (a) Representative Western blot and (b) densitometric
analysis of siRNA mediated piezo1 depletion. Data representative of 5
independent experiments. Significance determined using a one-way ANOVA
followed by Dunnett's test. (c) Representative images of piezo1-depleted
VSMCs cultured on 12 or 72 kPa hydrogels following angiotensin II (10
\selectlanguage{greek}µ\selectlanguage{english}M) stimulation. N.T. = non-targeting siRNA. Actin cytoskeleton (purple)
and Lamin A/C labelled nuclei (green). Top -- Representative XY images
of VSMC area, scale bar = 100 \selectlanguage{greek}μ\selectlanguage{english}m. Bottom -- Representative XZ images of
VSMC height, scale bar = 20 \selectlanguage{greek}μ\selectlanguage{english}m. (d) VSMC area and (e) volume,
representative of 5 independent experiments with [?]120 cells analysed per
condition. Significance determined using a one-way ANOVA followed by
Sidak's test. (* = \emph{p \textless{} 0.05} , error bars represent \selectlanguage{ngerman}±
SEM).

\textbf{Figure 9. Extracellular calcium influx via Piezo1 ion channels
triggers microtubule destabilisation.} Representative images of isolated
VSMCs cultured on 12 or 72 kPa polyacrylamide hydrogels treated with
angiotensin II (10 \selectlanguage{greek}µ\selectlanguage{english}M). Cold-stable microtubules, (\selectlanguage{greek}α-\selectlanguage{english}tubulin, aqua) and
nuclei (DAPI, blue). Scale bar = 50 \selectlanguage{greek}μ\selectlanguage{english}m. (a) GsMTx-4 (500 nM) pre-treated
VSMCs. (b) number of cold-stable microtubules per cell, representative
of 5 independent experiments, with [?]100 cells analysed per condition.
(c) siRNA-mediated piezo1 depleted VSMCs. N.T. = non-targeting siRNA.
(d) number of cold-stable microtubules per cell, representative of 5
independent experiments, with [?]109 cells analysed per condition. (e)
VSMCs were cultured in the presence or absence of extracellular calcium
(f) number of cold-stable microtubules per cell, representative of 5
independent experiments, with [?]98 cells analysed per condition.
Significance determined using a one-way ANOVA followed by Sidak's test.
(* = \emph{p \textless{} 0.05} , error bars represent \selectlanguage{ngerman}± SEM).\selectlanguage{english}
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