Biomechanical Characterization of Endothelial Cells Exposed to Shear

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bioRxiv preprint doi: https://doi.org/10.1101/2020.09.29.319772; this version posted September 30, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-ND 4.0 International license.

1 Biomechanical characterization of endothelial cells exposed to 2 shear stress using acoustic force spectroscopy

3 Giulia Silvani1#, Valentin Romanov1#, Charles D. Cox1,2 & Boris Martinac1,2,*

4 1Victor Chang Cardiac Research Institute, Darlinghurst, Sydney, NSW, Australia

5 2 St Vincent’s Clinical School, University of New South Wales, Sydney, NSW, Australia

6 #These authors contributed equally

7 * Correspondence: 8 Corresponding Author 9 [email protected]

10 Keywords: cell mechanics, microrheology, junction, cytoskeleton, contractile prestress, stress- 11 stiffening. (Min.5-Max. 8)

12 Abstract

13 Characterizing mechanical properties of cells is important for understanding many cellular processes, 14 such as cell movement, shape, and growth, as well as adaptation to changing environments. In this 15 study, we explore mechanical properties of endothelial cells that form the biological barrier lining 16 blood vessels, whose dysfunction leads to development of many cardiovascular disorders. Stiffness 17 and contractile prestress of living endothelial cells were determined by Acoustic Force Spectroscopy 18 (AFS) focusing on the displacement of functionalized microspheres located at the cell-cell periphery. 19 The specific configuration of the acoustic microfluidic channel allowed us to develop a long-term 20 dynamic culture protocol exposing cells to laminar flow, reaching shear stresses in the physiological 21 range (i.e. 8 dyne cm-2) within 48 hours of barrier function maturation. A staircase-like sequence of 22 increasing force steps, ranging from 186 pN to 3.5 nN, was applied in a single measurement revealing 23 a force-dependent apparent stiffness in the kPa range. Moreover, our results show that different degrees 24 of stiffening, defining the elastic behavior of the cell under different experimental conditions, i.e. static 25 and dynamic, are caused by different levels of contractile prestress in the cytoskeleton, and are 26 modulated by shear stress-mediated junction development and stabilization at cell borders. These 27 results demonstrate that the AFS is capable of fast and high-throughput force measurements of 28 adherent cells under conditions mimicking their native microenvironment, and thus revealing the shear 29 stress dependence of mechanical properties of neighbouring endothelial cells.

31 1 Introduction

32 Cells in vivo, whether isolated or part of a larger collective, are constantly exposed to physical forces 33 such as extensional, compressive, and shear stresses, all playing a critical role in regulating 34 physiological or pathological conditions [1, 2]. Cellular mechanics and rheological properties (e.g. 35 viscosity and stiffness) determine the ability of cells to respond to mechanical cues, and are important 36 for many physiological processes, including growth, division and migration[3]. bioRxiv preprint doi: https://doi.org/10.1101/2020.09.29.319772; this version posted September 30, 2020. The copyright holder for this preprint (which was not certified by peer review) is the author/funder, who has granted bioRxiv a license to display the preprint in perpetuity. It is made available under aCC-BY-ND 4.0 International license.

37 A remarkable example of cells under potentially high mechanical stimuli are endothelial cells (ECs) 38 that line the inner surface of blood vessels and form the endothelial barrier. ECs form a tight tessellated 39 monolayer which is constantly exposed to shear force generated by blood flowing along their apical 40 surface [4, 5]. The formation and maintenance of EC contacts, which provides the functional integrity 41 of blood vessels, requires a complex interplay of plasma membrane proteins, cytoskeletal components 42 and associated signaling molecules [6]. ECs have evolved to form a size-selective barrier, consisting 43 of specialized transmembrane proteins, called inter-endothelial adherens junctions, located at the cell- 44 cell border which are ubiquitously expressed in endothelia of all vascular beds [7]. Among them, 45 vascular endothelial cadherin (VE-cadherin), is one of the main structural and regulatory proteins 46 controlling endothelial barrier function and permeability [8-10]. Not only do these junctions link the 47 cells together, they also work as surface receptors generating a cascade of intracellular signaling upon 48 external applied stimuli (e.g. shear stress), triggering dynamic interactions with cytoskeletal elements. 49 The shear stress exerted by blood flow, typically in the range of ~1-15 dyne/cm2 [11-16], is known to 50 be an important regulator of mechanotransduction in vascular physiology [17, 18], including 51 mechanosensory processes of actin-mediated stabilization of junctions [13, 19-21], followed by 52 morphological remodeling [6, 22-24] and tension redistribution [25] such that cells can maintain or 53 change their shape.

54 According to the tensegrity model of living cells [26], the shape and stability of neighboring cells are 55 dictated by the internal framework of contractile actomyosin filaments, better known as actin stress 56 fibres. These active filaments are interconnected with each other and with cell-cell or cell-extracellular 57 matrix adhesion complexes, generating a tensile prestress through a balance of complementary forces 58 within the network [26-28]. Any variation of this structure, whether spatially or temporally, would 59 affect the cell’s mechanical properties, e.g. cell stiffness and deformability, as well as shape stability 60 [29-32]. In other words, the cytoskeletal prestress defines the constitutive elastic behavior of the cell 61 and is modulated by shear stress-mediated junction development and stabilization at cell borders which 62 in turn is strictly coordinated with actin stress fibre reorganization within the cell [32, 33]. Although 63 the precise role of VE-Cadherin as a sensor of external stimuli remains unclear, several studies have 64 demonstrated the importance of shear stress in in vitro experiments to achieve a functional monolayer 65 with barrier properties [15, 34-39]. Besides, it is widely accepted that any alteration of the pattern of 66 blood flow can lead to a wide range of vascular pathologies, including atherosclerosis and pulmonary 67 arterial hypertension [40, 41]. As a result, understanding how the mechanical behavior of collective 68 ECs may vary when exposed to fluidic shear stress is of critical importance in elucidating the cellular 69 malfunction.

70 A number of micro-rheological techniques have been developed aiming at characterization of the 71 viscoelasticity of adherent cells. These techniques include Atomic Force Microscopy (AFM), Magnetic 72 or Optical tweezers, Micropipette Aspiration and Particle Tracking Microrheology (PTMR) [42]. Two 73 major limitations of these techniques are measurements of reproducibility and throughput [43]. More 74 importantly, most of the techniques require an open chamber configuration, making it difficult to mimic 75 the physiological shear stresses necessary for EC maturation during culture. An alternative method, the 76 recently introduced Acoustic Force Spectroscopy (AFS) technology, has the potential to overcome 77 these drawbacks and greatly improve investigation of adherent cell mechanics. Originally designed for 78 single-molecule rheology, this method uses controlled acoustic forces (in the range of pNs to nNs ) 79 over a microfluidic channel to stretch multiple molecules in parallel, individually tethered to 80 functionalized microspheres [44, 45]. Later on, it has been used for characterization of mechanical 81 properties of red blood cells upon different chemical treatments [46]. Recently, AFS methodology has 82 been applied to Human Embryonic Kidney (HEK) cells capturing their inherent heterogeneity and

83 showing the impact of temperature and pharmacological treatments on the mechanical properties at the 84 membrane level [47].

85 In this study, we exploit the unique configuration of the closed AFS channel to probe the evolution of 86 ECs mechanical properties during maturation of a confluent monolayer. By doing so, we build on the 87 work of Nguyen et al. [48], while overcoming some reported limitations. As pointed out in their paper, 88 viscoelastic measurements with AFS require rigorous calibration and data analysis, nevertheless a 89 proper physiological shear stress is also needed to achieve the development of a phenotypic 90 endothelium. First, we developed a protocol for long term, dynamic cell culture under physiological 91 shear stress, i.e. 8 dyne/cm2. Second, after monolayer maturation, which was evaluated by 92 immunofluorescence (IF) microscopy directly in situ at different steps, we performed creep tests by 93 locally pulling the periphery of the cellular membrane with functionalized silica particles. The 94 viscoelastic response of the cells was then modeled by a Power law model, to estimate the two 95 parameters characteristic of the model, i.e. stiffness and the power-law exponent, both related to the 96 active contractile prestress in the cytoskeleton [49]. Two different EC lines, Human Umbilical Vein 97 Endothelial Cells (HUVECs) and Human Aortic Endothelial Cells (HAECs), were investigated to show 98 the potential of this tool to capture heterotypic properties of endothelial barriers. Each experiment 99 allows up to 80 measurements with forces ranging through the physiologically relevant range of pNs- 100 nNs. Under these conditions, we demonstrated that the AFS is capable of high-throughput force 101 measurements allowing for direct comparison between actin stress fibres reorganization, VE-Cadherin 102 formation, and shear stress induced stiffness modulation.

103

104 2 Material and methods

105 Experimental set-up.

106 The AFS chip, based on a lab-on-chip device, custom fabricated by LUMICKS B.V. [44], is described 107 in detail in SI.

108 Cell culture and seeding into the AFS channel.

109 Human Umbilical Vein Endothelial cells (HUVECs) and Human Aortic Endothelial Cells (HAECs) 110 were purchased from Lonza (Cat. No. CC-2517, Cat. No. CC-2535). The culture medium was the 111 endothelial basal medium-2 (EBM-2) supplemented with endothelial growth medium (EGM-2) 112 BulletKit from Lonza (Cat. No. CC-3162). Cells were grown in tissue culture flasks and maintained in 113 humidified atmosphere at 37°C and 5% CO2. The culture medium was changed every 2 days and cells 114 were used up to the 5th passage to ensure the expression of key endothelial protein components. Further 115 details on functionalization of the AFS chip for cell seeding are given in SI.

116 Immunofluorescence staining and Image analysis

117 Junction morphology and cytoskeleton organization was tested under different experimental 118 conditions. VE-cadherin protein and actin stress fibres were imaged by fluorescence microscopy and 119 ImageJ was used for scanning and analysis of the changes in morphology of both cytoskeletal proteins, 120 as described in SI.

121 Microsphere functionalization and tracking

122 Microspheres were functionalized with fibronectin and tracked as described in SI.

123 Force steps measurement

124 To determine the acoustic radiation force, 퐹푟푎푑, we performed a force-balance on acoustically driven 125 beads [47]. The method is explained in Supplementary material together with the experimental protocol 126 of force application. The quadratic dependence of the force with the voltage is also shown (Fig.S1).

127 Data fitting

128 To quantify the force dependence of the creep response we used a superposition of power-law models. 129 Details of data fitting are given in SI. 130 131

132 3 Results

133 The main advantage of the AFS technology is the confined fluidic circuit, shown in Fig.1A. The device 134 is also engineered and optimized to simultaneously allow for acoustic resonance and optical imaging 135 thanks to a transparent piezoelectric element glued on top [45]. Given the small dimension of the 136 microfluidic channels, the flow rate and the properties of the fluid, the flow is fully laminar, meaning 137 that the flow pattern is completely predictable and does not reach turbulence (Fig.1Ai-ii). Moreover, 138 its solid structure allowed for high perfusion rates, mimicking the shear stress magnitudes exerted on 139 ECs in vivo, during monolayer maturation. Here, we exploited the unique geometry of the AFS chip to 140 perform long term culture of ECs by connecting the microfluidic channel to a peristaltic pump and 141 maintaining the flow of growth media for 48 hours at a shear stress of 8 dyne cm-2. A sketch of the set- 142 up is shown in Fig.1B. The chip and the medium reservoir were placed in a dry incubator at 37 °C and 143 5% CO2 in order to avoid damaging the piezo-glass connection. During maturation, the endothelial 144 morphology changed considerably, from an initial disorganized configuration of rounded spread cells 145 to a compact endothelial monolayer, characterized by cell elongation in the stream direction (Fig 1C). 146 Brightfield images of HAECs, after 4, 24, and 48 hours from seeding procedure are shown in Fig1 D- 147 F. The elongation of cells over time in the direction of the flow is indicative for a phenotypic 148 endothelium [50]. 149 150

151

152 Fig 1. Long-term cell culture in the AFS chip. (A) Cartoon representation of the AFS chip (i-ii) Fluid dynamic simulation 153 of fluid streamlines. (B) Sketch of the set up with the AFS fluidic channel connected to a peristaltic pump. (C) Distribution 154 of EC orientations after 4 h (green), 24h (red) and 48h (blue) from seeding obtained with “orientationJ distribution” of 155 ImageJ software. (D) Crop section of the channel imaged in transillumination mode, showing ECs 4 hours from seeding 0 156 procedure and (E,F) cells after 24 and 48 hours, respectively, under flow culture condition at 37 C with 5% CO2 and no 157 humidity. The flow was applied from left to right. 158 159 160 Toward EC monolayer and junction maturation

161 It is generally proposed that the control of ECs adhesion, migration and barrier function, critically 162 depends on the mutual interaction of VE-cadherin with actin filaments via receptor protein complexes 163 [51, 52], as both structures are remodelled under physiological shear stress [22, 24]. Here, we 164 performed IF microscopy, to assess the phenotype of these key protein components, for both EC lines, 165 grown in the AFS device after 4 hours from seeding and at the end of the flow culture protocol, to 166 demonstrate the significance of shear stress and timing in promoting cell barrier formation. In this 167 section, we refer to HAECs staining and evaluation, although the same IF results were obtained with 168 HUVECs (Fig.S2). When subjected to shear stresses, HAECs showed a varied response. The most 169 striking one was the change in morphology, as they went from sub-confluence to complete confluence. 170 Without any shear applied, i.e. CTRL, cells appeared larger and grew in a polygonal shape without 171 directionality and with VE-cadherin proteins forming an irregular intermittent network (Fig.2A, upper 172 left panel) at the cell periphery. After 48 hours of flow, cell contact was achieved and VE-cadherin 173 exhibited linear oriented junctional staining (Fig.2A, lower left panel) suggesting a well-established, 174 mature confluent endothelium, as expected of a tight cobblestone monolayer [50]. Fig 2.B shows the 175 fluorescence intensity profile for VE-Cadherin protein concentrated at the cell border for both 176 experimental conditions. Under flow (blue curve), cells exhibited higher junctional localization of VE-

177 Cadherin staining, as opposed to those grown under static conditions (red curve), where the junction 178 was yet to form. Regarding rearrangement of actin stress fibres within the cellular cytoplasm, we 179 performed fast and automated image analysis by monitoring changes in IF signal under both culture 180 conditions. Phalloidin staining revealed a strict cortical organization of actin filaments, referred to as 181 circumferential actin bundles [33] in cells subjected to shear stress (Fig. 2A, lower middle panel), 182 whereas cells under static conditions exhibited more stress fibres across the cell body (Fig. 2A, lower 183 middle panel). Therefore, at the end of the cell culture protocol, actin filaments aligned along the cell 184 periphery sustaining and reinforcing junction proteins, as shown in the lower right panel of Fig.2A 185 by a linear, overlayed pattern of VE-Cadherin/Phalloidin. This result was also confirmed by 186 quantification of actin stress fibres along the cells’ smaller axis, i.e. number of fluorescence peaks per 187 μm (Fig.2 C). We found that, for both HUVECs and HAECs, the mid-range of fibre densities for cells 188 under flow conditions decreased compared to control cells, suggesting substantial remodelling of actin 189 filaments is necessary for junction stabilization (Fig. 2D,E).

190

191 Fig 2. Reorganization of VE-Cadherin and actin filaments under flow culture condition. A) Fluorescence images of 192 HAECs monolayer stained for junction VE-Cadherin (red, left panel) and Actin Filaments (green, middle panel) under 193 static (CTRL) or dynamic (Flow culture) condition. B,C) Representative graphs showing fluorescence intensity (a.u.) 194 profile for VE-Cadherin signal across cell-cell junctions and actin filaments, respectively. Insets: examples of the line along 195 which fluorescence intensity profile was obtained). D,E) Box plots of F-actin stress fibres density for both HAECs and 196 HUVECs. 197 198 Manipulating endothelium monolayer with acoustic forces

199 In our AFS experiments, the “endothelialized” microfluidic channel was resonantly excited at 14.50 200 MHz, the frequency of which determines the location of the nodal plane for the acoustic standing wave. 201 When acoustic force is applied, silica microspheres attached to the cell surface stretch the membrane 202 of multiple adherent cells as they are pulled toward the node, capturing the cell-dependent viscoelastic 203 creep response. The principle of an AFS experiment is shown in Fig. 3A. Only beads in the near 204 proximity of cell-cell borders have been chosen for these experiments (Fig. 3B). In order to study the 205 dependence of contractile prestress in the cytoskeleton with shear stress-mediated mechanical 206 properties of neighbouring ECs, a staircase-like sequence of increasing force steps (Fig.3C), ranging 207 from 186 pN to 3.5 nN, was applied in a single measurement (i.e. differential creep test), for cells under 208 static or dynamic conditions. Indeed, from a single-step creep experiment, no reliable conclusion about 209 contractile prestress and shear-dependence can be made. Under the application of single stretch force, 210 the cytoskeletal structure may not return quickly enough to its original undisturbed state in between 211 measurements, and a large number of independent measurements is required. Besides, at higher forces, 212 cells show a stress-stiffening response, which is consistent with adherent cell rheological behaviour in 213 the physiologically relevant regime of large external forces and deformation [53]. We utilized the 푡 훽 214 Power-Law model, 퐽(푡) = 퐽표 ( ) , to capture the viscoelastic parameters of cells at each force step, 휏0 215 where E0, the inverse of the prefactor 퐽표, is the apparent stiffness (units of Pascal) and β, the power- 216 law exponent, defining the solid- or liquid-like behaviour of the cell. The displacement of beads at each 217 force step and for all cells measured, always follow a weak power-law response (Fig. 3C,D). In Fig. 218 3E, the cell compliance 퐽표, decreases with increasing forces, indicating stress-stiffening behaviour.

219 220 Fig 3. Rheology creep test for measuring nonlinear viscoelastic properties of ECs. A) Sketch showing the physical 221 principle of the experiment. B) Bright field image of endothelium monolayer seeded in the AFS channel together with a 222 Silica bead attached to the cellular surface. The dashed line represents the boundary of multiple cells. C) Creep test made 223 of a staircase-like sequence of increasing force steps, from 180pN to 3.5nN. The bead displacement is fit to a superposition 224 of creep responses (blue dashed line). D) Compliance curves at a specific force step taken from beads located all over the 225 AFS channel and collected into a single experiment over 45 minutes (퐹푟푎푑= 2.8 nN, n=80, HAEC cells, EGM, 9.2 um Silica 226 beads). E) Histogram showing the decrease of the factor J0 with increasing force, indicating stress stiffening, i.e. non-linear 227 viscoelastic behaviour. Data are presented as means ± SEM. 228 229 Local mechanical properties measured by Acoustic Force Spectroscopy 230 Measurements of local mechanical properties using AFS in this study indicate that different degrees of 231 stiffening, characteristic of cells under different experimental conditions, were caused by different 232 levels of contractile prestress in the cell, revealing the shear stress-dependence of endothelial 233 mechanical properties.

234 When subjected to the staircase-like sequences of forces, both HUVECs and HAECs behaved like a −1 235 viscoelastic material, in the non-linear regime, where the stiffness, 퐽표 , increased with increasing force −1 3 236 (Fig.4A,C). We found that 퐽표 is in the order of 10 Pa, consistent with other rheology measurements 237 of adherent cells using other methods [43, 53-57]. However, variation of the power law exponent, β, 238 was not observed (Fig.4B,D). Significantly, the same stress-stiffening was found for both experimental 239 conditions, with the only difference that the stiffness for cells under static conditions (CTRL, red 240 curves) was higher compared to cells subjected to shear stress for 48 hours (Flow culture, blue curves). 241 As confirmed by the stress-fibre density analysis, during monolayer maturation, actin filaments

242 rearranged to the cell periphery by minimizing radial tension forces in the cell’s body, and thus 243 apparently causing the cells to become softer. To further confirm this hypothesis and to find the 244 relationship between the degree of stiffening and the prestress in the cytoskeleton, we fitted equation 245 (S.5) to all force-stiffening curves. As shown in Fig.4E, for both cell types, prestress was higher for 246 measurements under static conditions, indicating that the tension inside the cells was not yet relaxed 247 by the formation and maturation of protein junctions and was rather carried by contractile actomyosin 248 filaments. These results seem to be in line with previous findings where stiffer cells exhibited higher 249 prestress [53]. Interestingly, the prestress of HAECs was lower compared to HUVECs, while for the 250 latter, the magnitude of the prestress under different conditions remained almost the same. This could 251 be due to the heterotypic resistant properties of endothelium barriers from different vascular beds [58], 252 however more experiments need to be performed in order to further confirm such hypothesis.

253

254

255 Fig 4. Effect of shear stress on the viscoelastic properties of ECs. (A,C) Stiffness, 1/J0, versus applied external stress 256 for HAECs and HUVECs respectively, under static and dynamic condition (n=100 each). (B,D) The power law exponent, 257 β, does not show any notable variation (E) Final stiffness versus prestress for all data grouped by cell type and experimental 258 conditions. Data are presented as means ± SEM.

259 4 Discussion

260 The mechanisms by which ECs respond and transduce signals, such as mechanical cues, are of great 261 importance in vascular physiology, as they are often altered during disease, driving multifunctional

262 behavior and pathology. It is broadly accepted that the cytoskeleton of adherent cells is critical to 263 mechanotransduction by redistributing external force across the cytoplasm e.g. shear stress from the 264 blood. In particular, actin fibres are known to have a central role in force transmission, regulating many 265 cellular functions, including morphological stability, adhesion, and motility [59, 60]. Although it is 266 known that any changes in spatial distribution of actin fibres are shear dependent and occur in 267 correlation with cadherin junction localization at cell borders, there is little quantitative knowledge 268 about how stress fibre density and organization modulate cellular stiffness. Given that, it is important 269 to characterize the mechanical properties of ECs under relevant physiological conditions in order to 270 understand resulting cellular malfunction.

271 Much of the pioneering work characterizing adherent cells are performed using methods lacking in 272 physiological reproduction of the cell’s native microenvironment [42]. Experimental limitations, such 273 as an open chamber configuration, makes it difficult to assess ECs monolayer maturation under 274 physiological shear stress conditions during cell culture. Moreover, the thickness of the cell may alter 275 indentation measurements as it will determine the contribution from the substrate to the overall stiffness 276 [61, 62]. Refer to [42] for a more comprehensive review on factors that may influence rheological 277 measurements. In this study, we utilized a new technique, called Acoustic Force Spectroscopy, which 278 operates in similar fashion to Optical Tweezers but overcomes the low throughput. Moreover, AFS 279 technology exploits acoustic mechanical waves to manipulate a particle position, which requires 280 simpler experimental equipment and sample preparation compared to laser set-up and usage.

281 After establishing a mature endothelial monolayer, assessed by IF microscopy at different growing 282 steps, several beads at once have been pushed toward the acoustic node, stretching different cellular 283 membranes at the surface receptors/cytoskeleton linkage site in a high-throughput fashion. By fitting 284 a power law model to the bead’s displacement using a custom-written MATLAB program, we could 285 determine the stiffness of ECs upon application of a range of forces and under different experimental 286 conditions, revealing the shear stress dependence of mechanical properties of neighbouring ECs. We 287 showed that shear stress-driven actin filament reorganization in the cytoplasm is essential for junction 288 formation and stabilization, confirming the significance of the interplay between actin filaments and 289 VE-Cadherin junction during barrier function maturation. Importantly, the shear stress experienced by 290 ECs in our experiments was within the range of shear forces encountered by the cells in their native 291 environment. This could help us to demonstrate how shear stress-driven morphology remodeling 292 modulates cell stiffness while determining contractile prestress in the cellular body, going further with 293 the fundamental viscoelastic measurements than Nguyen et al. [48]. Furthermore, we have extended 294 the potential of the AFS by performing IF microscopy in situ, proving the applicability of AFS for 295 further studies using live imaging. In summary, our results demonstrate the potential of AFS as a novel 296 tool for shedding light on mechanobiology of endothelial cells, meeting several requirements during 297 the cell culture and allowing for fast and reproducible high-throughput measurements of the 298 viscoelastic properties at the single cell level.

299

300 5 Author Contributions

301 G.S. designed, ran, processed, and analysed the experiment and wrote the manuscript. V.R. designed, 302 ran, processed, and analysed the experiment and edited the manuscript. C.C. helped with formulating 303 experiments and editing of manuscript. B.M. formulated the experiments and editing the manuscript.

304 6 Supplementary Material

305 Supplementary material is included and contains methods for Experimental set up, Cell culture and 306 seeding into the AFS channel, Immunofluorescence staining and Image analysis, Microsphere 307 functionalization and tracking, Force steps measurement, Acoustic Force calibration and Data fitting.

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