Cell loss disrupts mechanical homeostasis to drive retinal pigment epithelium ageing-like phenotype in vitro

Domain: neuroscience
Published: 2026-04-08
Editorial outcome: major_revision

Abstract
Tissue homeostasis emerges from mechanical feedback loops balanced by cell loss and proliferation, a balance that in postmitotic tissues must be maintained without compensatory proliferation. Yet how these tissues preserve mechanical homeostasis and how this challenges function in ageing remains unclear. To establish the relationship between cell density, mechanical homeostasis, and function, we induced age-mimicking cell loss in a postmitotic retinal pigment epithelium (RPE) in vitro. This model recapitulates key structural hallmarks of RPE ageing, including reduced cell height, shortened microvilli and cytoskeletal reorganisation. The density-reduced RPE establishes a new mechanical equilibrium characterised by tissue stiffening and increased junctional contractility. Functionally, these monolayers exhibit impaired phagocytosis of photoreceptor outer segments due to compromised apico-lateral plasticity, which is mechanistically linked to the modulation of actin nucleators, Arp2/3 and formins. Altogether, our findings show that a cell loss-induced shift in mechanical homeostasis drives age-related RPE dysfunction, demonstrating that structural remodelling and mechanics alone can compromise tissue function in ageing.

Introduction
The structure and form of a biological entity determine its function, as described by the form-function paradigm1,2. This paradigm applies across various length scales, from the molecular3,4 and sub-cellular levels5,6 to the tissue and organ levels, where proper architecture and cellular connectivity are essential for functionality7–9. In recent decades, alongside form and organisation, increasing recognition as key parameters influencing function and homeostasis has also been given to the mechanical properties of tissues10–12. These properties are determined by the cellular organisation and environment13–15, and their regulation within a preferred range engages mechanosensitive feedback loops that sustain physiological function16–18, a concept referred to as mechanical homeostasis19–21. Despite its importance, mechanical homeostasis remains poorly understood in postmitotic tissues such as the central nervous system and the retina, which must adapt structurally to naturally occurring cell death to maintain the tissue’s integrity and functional state22–25.

An emblematic example of this adaptation is the age-related changes occurring in the retinal pigment epithelium (RPE)26. In the retina, the RPE is located between the light-detecting photoreceptor cells and the choroid capillaries and provides vital metabolic support to photoreceptor cells27. In ageing, this support is challenged by the steady rate of apoptosis in RPE28–30, which needs to be actively compensated by the remaining cells, leading to major structural changes of the tissue. These include monolayer thinning, with the average RPE cell height reported to decrease, while cell area increases29,31,32. Furthermore, the RPE actin cytoskeleton remodels, leading to local fibres thickening or thinning with the formation of transcellular stress fibres33. Finally, ageing has been shown to be associated with changes in RPE apical microvilli, which become shorter and sparser31,34. In temporal correlation to these structural changes, the functional ability of the RPE to support photoreceptor cells changes towards a reduced ability to phagocyte and process photoreceptor outer segments (POS) fragments, with important implications for retinal degenerative pathologies35,36.

While it has been proposed that the age-related changes in RPE organisation stem from a time-associated reduction in cell density, direct evidence supporting this hypothesis remains elusive. Moreover, whether such alterations drive a shift in tissue mechanical homeostasis that implies functional decline remains a critical unresolved question. Notably, we have recently shown that the strength of both cell-substrate and cell–cell adhesion forces in RPE cells directly relates to their ability to phagocytose POS, with higher actomyosin contractility leading to reduced phagocytic activity21. Finally, because rodent models and human post-mortem tissue studies cannot reveal the precise effects of reduced cell density on monolayer structure from other ageing-related processes, such as oxidative stress or extracellular matrix (ECM) remodelling, it is necessary to establish a reductionist, human-relevant in vitro model.

To address this knowledge and technical gap, we present an optimised in vitro model that employs large-scale apoptosis on demand to promote a reduction in cell density in postmitotic human induced pluripotent stem cell-derived RPE (iRPE). This density-reduced iRPE (dr-iRPE) displays structural changes such as enlarged and thinner cells with shortened apical microvilli and cytoskeleton reorganisation, alongside increased tissue stiffness, enhanced junctional contractility and a remodelled apical cortex. The mechanical changes coincide with impaired POS phagocytosis, reflected by reduced apicolateral deformation, decreased number of internalised particles and increased fragment size. Pharmacological modulation of actin nucleation allowed us to partially rescue function, suggesting that functional alteration may result from a shift in force balances within the RPE that prioritises lateral structural reinforcement instead of apical plasticity needed for POS phagocytosis.

Results
Cell density reduction in vitro mimics ageing RPE structure
Inspired by the natural reduction in cell density in RPE in vivo28,30, we aimed to replicate cell density reduction in vitro by inducing large-scale apoptosis and to study the adaptation of the epithelium to the cell loss. To model a native RPE monolayer, iRPE was employed as a recognised and reliable model for the native tissue37–40. Importantly, to investigate cellular force balance, iRPE were cultured on 4kPa soft polyacrylamide hydrogels with basement membrane-like coating, thereby recapitulating the physiological mechanical and biochemical cues of the native substrate as previously characterised21 (Supplementary Fig. 1a). In these conditions, iRPE assumes a characteristic honeycomb-like organisation, apicobasal polarisation (Supplementary Fig. 1b), and arrest of proliferation (0.01 to 0.41% within 24h) (Fig. 1a and Supplementary Fig. 1c). To facilitate on-demand apoptosis, on the 10th day of culture we transduced the cells with an adeno-associated virus (AAV) carrying a construct designed to promote the expression of FKBP-caspase8 (Fig. 1b, Supplementary Fig. 1d). FKBP-caspase8 is pro-apoptotic fusion protein that can be activated on demand through AP20187-induced dimerisation, initiating the cellular suicide programme41. Therefore, after reaching a stable expression, on the 15th day, we incubated the monolayer with the compound AP20187, inducing monolayer-scale apoptosis that led to the extrusion of 2% of the cells within the observation time (Fig. 1b, c, Supplementary Fig. 1e and Supplementary Movie 1). By the 20th day (96h after apoptosis completion), the iRPE monolayer showed a reduction of cell density of approximately 8% (Fig. 1d), which was not compensated by increased proliferation. In fact, we quantified significantly lower proliferation rates compared to control monolayers (Fig. 1a). As a result, the obtained monolayer was referred to as density-reduced iRPE or dr-iRPE.

Morphologically, dr-iRPE presents cells with a larger area on average (Fig. 1e, Supplementary Fig. 1f and g) without experiencing any fluidification or unjamming, as evident by cellular area distribution and shape factor (Supplementary Fig. 1h and i), recapitulating morphological changes observed in human RPE ageing29. Furthermore, we noticed that one-tenth of cells within dr-iRPE presented transcellular stress fibres (Fig. 1f), which were also reported to occur frequently in human RPE ageing33. Next, we quantified the cell height in fixed and mounted samples (Fig. 1g, h), showing that dr-iRPE is significantly thinner than the controls (9.1 vs 11.6 µm), in agreement with observations from the ageing RPE31,42. Lastly, motivated by the documented changes of microvilli size and number in the ageing RPE in vivo31,34, we evaluated ezrin as a polarity marker and microvilli organiser43–45. Qualitative analyses of ezrin staining revealed a distinct morphology of the cellular apices between dr-iRPE and control (Fig. 1i), which can also be quantitatively captured by the reduced protein level detected in the cells by western blot (Fig. 1j). Moreover, the examination of the microvilli length by confocal microscopy showed approximately one-third shorter microvilli in the dr-iRPE compared to the control (Fig. 1k). Based on the studies performed on Ezrin-/- mice43, these observed differences strongly suggest that in the dr-iRPE conditions, the microvilli are less elaborated and more disorganised. Finally, the quantification of the active form of ezrin (phosphorylated ezrin), which connects the actin cortex to the membrane46, revealed a significant decrease in the dr-iRPE (Fig. 1l).

To exclude the influence of the viral delivery on the observed structural changes, we included a matched viral control, which promoted the expression of GFP (GFP-iRPE) (Supplementary Fig. 2a). Compared to this control, the dr-iRPE was still showing a reduced cell density and cell height (Supplementary Fig. 2b–d), as well as significant differences in both organisation and length of ezrin-positive microvilli (Supplementary Fig. 2e). Next, we addressed the observation that despite most of the cells within the monolayer expressed the reporter after transduction, only a few of the cells ultimately underwent apoptosis during the AP20187 treatment (Supplementary Movie 1). To exclude any possible effects of the leftover recombinant protein in the remaining cells, we included an additional mixed control in the analyses, consisting of an iRPE monolayer made by a mixed population of transduced (10%) and naïve cells. This condition allowed us to target the caspase activation only on a subpopulation of cells (Supplementary Fig. 2f, g). Interestingly, with this approach, apoptosis still caused a significant cell density and height reduction, confirming that these features are independent of the possible presence of inactive recombinant protein in the cells (Supplementary Fig. 2h, i). Lastly, we asked whether the changes were induced by secondary factors released in the medium after the large-scale apoptosis induction47. Hence, we transferred a medium pre-conditioned by the dr-iRPE sample during and after apoptosis onto control monolayers (Supplementary Fig. 2j). Nonetheless, the quantification still showed no effect of the “conditioned medium” on cell morphology, density, or height (Supplementary Fig. 2k–m).

Altogether, we present an in vitro model based on cell density reduction via on-demand apoptosis, and we demonstrate that the RPE monolayer responds to large-scale apoptosis with structural adaptation. This includes the formation of transcellular actin fibres, reduced cell height, larger cell area and shorter and remodelled microvilli. Interestingly, these cellular responses appear to result from intrinsic tissue stretching and preservation of cell volume rather than from cellular hypertrophy. Finally, our model recapitulates key morphological features of the aged RPE in vivo, suggesting a causal relationship between cell loss and the structural changes observed in ageing.

Cell density reduction compromises RPE phagocytotic activity
One of RPE’s main functions in vivo is the daily engulfment and internalisation of oxidised fragments of photoreceptor outer segments (POS), often referred to in the literature as POS phagocytosis48 or, due to the active nature of nibbling involved, as trogocytosis49. Along the lines of the form-function paradigm, we asked whether the structural adaptation observed in response to cell density reduction would alter iRPE’s ability to internalise POS. Thus, following an established assay21,40, we incubated control and dr-iRPE with fluorescently labelled porcine POS fragments in the presence of bridging molecules Protein-S and MFG-E8 for 4h (Fig. 2a). During this time, some of the fragments bound to the apical surface of the iRPE and some got internalised by the cells (Fig. 2a, b). After 3D segmentation, we used the apical actin signal to differentiate between bound and internalised fragments (Supplementary Fig. 3a). Quantification revealed that the dr-iRPE exhibited significantly fewer internalised particles, both within the field of view or when normalised by cell number (Fig. 2c, Supplementary Fig. 3b). Next, motivated by a reported trend of larger POS phagosomes in the RPE of aged animals36, we further quantified the volume of the internalised photoreceptor fragments. Surprisingly, we found the average volume of internalised fragments to be larger in dr-iRPE monolayers (Fig. 2b, d). Regarding the bound particles, neither their number nor their average volume was found to be significantly different (Supplementary Fig. 3c). Similar differences in internalised particle number and volume were observed in the mixed cell population subjected to cell density reduction, with fewer internalised particles and a larger average particle volume (Supplementary Fig. 3e, g). Finally, the medium transfer from the apoptotic-induced samples showed no significant impact on either the internalised particle count or on the fragment volume (Supplementary Fig. 3f, h).

Altogether, these data demonstrate that structurally remodelled dr-iRPE exhibit deficient internalisation of POS fragments, internalising fewer and larger fragments. By phenocopying the phagocytic deficits observed in ageing RPE, this model suggests that reduced cell density alone can drive functional decline in the remaining cells.

Density reduction alters actomyosin plasticity and mechanics
POS phagocytosis relies on the active remodelling of the apical actin network, which plays a crucial role in fragment internalisation and phagosome size21,40,49. Actin dynamics are essential throughout the process, guiding key steps from initial binding and ensheathment to the final scission of the phagosome48,50. Thus, we examined the morphology of the apical actin during internalisation, on a few-cell scale in our 3D segmentation and on a larger monolayer scale. In the side view, we noticed the apical actin of control iRPE monolayers experienced a substantial deformation during the internalisation, such as bulging and enlargement of the protruding cell portion above the cell junctions and clear formation of myosin IIB-positive cups (Fig. 2e–h). In contrast, dr-iRPE did not experience much evident deformation, and phagocytotic cups were less resolved (Fig. 2f, g). Next, we noticed that cells in the dr-iRPE displayed a different apical actin topography compared to the control, having an overall smoother surface, as quantified by area per field of view (Fig. 2h). Finally, on the monolayer scale, we noticed that the dr-iRPE frequently presented large actomyosin-rich apical structures (Fig. 2g, i, Supplementary Fig. 4a, b), which were absent in iRPE samples.

These observations suggest that iRPE cells in the density-reduced condition exhibit altered ability in remodelling their actomyosin cytoskeleton when internalising POS fragments. Consequently, we wondered whether the differences stemmed from distinct patterns of actin-cytoskeleton-related signalling. Hence, we examined the myocardin-related transcription factor A (MRTFA), which is known to regulate actin-associated signalling51. We observed a short-term translocation of MRTFA into the nucleus at day 17, shortly after the large-scale cell loss, which went back to levels comparable to the control at day 20, or the day of the read-out (Fig. 3a, b). This demonstrates that the cell density reduction leads to a transient transcriptional adaptation, possibly conveying the observed structural remodelling of dr-iRPE. We then examined the mRNA expression of actin-associated proteins after bulk RNA sequencing (Fig. 3c, Supplementary Fig. 5a). Examining more precisely the expression of these proteins showed an array of significantly differentially expressed actin-related genes in the dr-iRPE. Upregulated genes in dr-iRPE contained genes related to actin branching, polymerisation, depolymerisation, and crosslinking (ARPC2, PFN1, FHOD3, INF2, CFL2, ACTN1, FSCN1, FLNA). Downregulated genes included ezrin and the formins DIAPH1, DIAPH2, DAAM1 and DAAM2 (Fig. 3d, e). Next, to further assess the parallel between our model and ageing RPE, we compared global gene expression changes in iRPE and dr-iRPE with those reported in ageing RPE from two independent studies52,53 (Supplementary Fig. 5b–d). Considering the significant differences from our reductionist approach and in vivo human data, as well as the differences between the results of the studies, it is worth noting that our model may mimic further aspects of RPE ageing besides the actomyosin remodelling discussed in this work.

Altogether, these insights point to a distinct signature of actin-associated proteins in dr-iRPE that implies altered actomyosin network organisation, consistent with the observed changes in subapical F-actin topography. Such reorganisation may prevent the formation of the prominent apical deformations or bulges required for efficient phagocytosis, instead promoting the emergence of localised actin-rich structures.

As we previously showed that RPE mechanical equilibrium influences the epithelium phagocytotic activity21, we sought to determine whether the observed actin network remodelling would alter the mechanical properties of dr-iRPE. To characterise the viscoelastic properties of the monolayer as a multicellular system, we conducted nanoindentation using a spherical tip with a diameter of 20µm, allowing to probe a contact area of approximately 10µm in diameter (Fig. 4a, Supplementary Fig. 6a, Supplementary Movie 2). This approach allowed us to measure the Effective Young’s modulus of the uppermost 2.5µm of the monolayer, which approximately corresponds to the apical-most 9% of the cell height (average height of live iRPE is 27.8 ± 0.6µm). The Effective Young’s modulus of dr-iRPE was roughly one-third higher (694 Pa vs 513 Pa), pointing to a stiffer tissue (Fig. 4b). Next, using the same nanoindentation setup, we quantified both storage and loss moduli at different probing frequencies to characterise the monolayer viscoelasticity54. With this approach, we found a significant increase in the storage modulus of the dr-iRPE while the loss modulus remained unchanged (Fig. 4c, Supplementary Fig. 6b). Similar differences were observed when dr-iRPE was compared to the virus control or when the condition with mixed cell population was analysed without or with cell extrusion (Supplementary Fig 6d, e). In the latter case, the differences in stiffness and storage modulus were smaller and only apparent at the lower probed frequencies (1 and 2Hz). Lastly, medium transfer controls seemed to experience softening rather than stiffening, excluding an effect of apoptotic debris on the observed stiffening in dr-iRPE (Supplementary Fig. 6f). Collectively, these measurements revealed a reinforced cellular interconnectivity that results in a less deformable monolayer, i.e. stiffer, as well as more elastic, suggesting an actomyosin-based reinforcement55.

As mechanical properties and contractility are often linked56,57, we examined the levels of phosphorylated myosin light chain 2 (pMLC2) to obtain a general idea about actomyosin contractility58. Immunoblotting revealed general increased levels of pMLC2 in dr-iRPE (Supplementary Fig. 6c), pointing to overall higher actomyosin contractility, especially localised at the cell-cell junctions as shown by immunofluorescent staining (Fig. 4d and dI). This observation was further supported by enrichment of vinculin at the cell-cell and at the tricellular junctions (Fig. 4e, eI), pointing to higher contractile force at cell-cell junctions59.

Nanoindentation enabled us to characterise the biomechanical properties of the RPE from the apical side. Our indentation setup was designed to probe multicellular length scale deformation, where the indentation of the 20µm tip at the selected depth is counteracted by intercellular junctions60 (Fig. 4a, Supplementary Fig. 6a, Supplementary Movie 2). To characterise subcellular mechanical properties of the RPE monolayer non-invasively, i.e., independently from intercellular adhesion, we applied Brillouin micro-spectroscopy. Here, the detection of photons, inelastically scattered by thermally-induced acoustic phonons, enables the probing of viscoelastic properties within the sample at microscale resolution61. Using Brillouin microscopy, we first performed low-resolution z-scans through iRPE and dr-iRPE monolayers, including the surrounding hydrogel and medium (Fig. 4f). These line profiles revealed the presence of a peak in the Brillouin shift, corresponding to the cellular basal side, which gradually decreased towards the apical cell region (Fig. 4g). The quantification of fitted peak values of these line profiles revealed significantly lower Brillouin shift in dr-iRPE compared to iRPE (Fig. 4h). Moreover, based on the peaks shown in Fig. 4g and confirmed by live-cell brightfield imaging, we identified the apical and basal z-levels for conducting 2D high resolution raster scans (Fig. 4i). The 2D raster scans (50 x 50µm) revealed mechanical compartmentalisation into areas with different Brillouin shifts, which may correspond to cellular and intercellular regions (Fig. 4j). Notably, this compartmentalisation was much more evident at the basal sides. The quantification of the mean Brillouin shifts revealed a difference between the conditions, with dr-iRPE cells showing lower average values at the apical (6.30 vs 6.36 GHz) and basal (6.35 vs 6.37 GHz) sides of the cells (Fig. 4k). Interestingly, when comparing apical and basal shifts within the same conditions, dr-iRPE cells showed significantly different apical shifts compared to basal, which was not the case in iRPE (Fig. 4l). As the Brillouin shift reflects cytoskeletal organisation and water content61–63, this supports the idea that the apical sides of the cells undergo the most pronounced reorganisation in dr-iRPE. This is consistent with microvilli and apical actin remodelling observed in dr-iRPE, i.e., transcellular stress fibres and F-actin surface topography.

As the intercellular junctions showed an increased pMLC2, thereby an increased contractility and stiffness, we sought to isolate the shifts of “cell edges” from those of “cell bodies”. As this distinction was not clear in the apical scans, we performed the analysis on the basal sides, where the distribution of shift values revealed two distinct modes (Supplementary Fig. 7a, b). Quantification based on segmented Brillouin shift plots shows that, regardless of the experimental condition, the interstitial regions of the monolayer (“cell edges”) consistently exhibit lower shifts than the cell cores (“cell bodies”). This also confirms a trend toward reduced core shifts in dr-iRPE independently of the interstitium (Supplementary Fig. 7c, d). Notably, the shift quantified at the interstitium was unexpectedly lower than at the cell bodies, despite enrichment of F-actin. This likely reflects the absence of well-defined intercellular junctions at the basal side, and instead, the presence of loosely interdigitated basal membranes visible with F-actin staining (Supplementary Fig. 7e). These looser basal intercellular interactions likely explain why structural and mechanical remodelling following cell density reduction is predominantly apical, affecting load-bearing apical junctions rather than the more compliant basal domain.

Altogether, biomechanical characterisation at the two different length scales showed that, in addition to the structural and functional adaptation, the reduction of cell density causes RPE to adopt a different mechanical state. This mechanical state is associated with increased contractility at the apical intercellular junctions, which confers a stiffer and more elastic tissue, as evident by the nanoindentation. Brillouin shift analyses at the subcellular level also confirmed distinct mechanical states between the two conditions, most pronounced at the apical surface. Taken together, this apical prevalence of the mechanical changes may result from the intimate link between the epithelial junctional complex and the organisation of the apical domains, which in the case of the RPE may create a mechanical feedback loop between tissue integrity and function.

Actin modulation differentially impacts POS phagocytosis
Motivated by our observations on cortical actin topography, mechanics, and the differential expression of actin-related regulators, we aimed to directly investigate the cause–effect relationship between POS internalisation and actin remodelling. To this end, we targeted actin nucleation, a key process influencing cortical thickness and membrane tension64. For the experimental design, we considered the upregulation of Arp2/3 subunit mRNA expression and the downregulation of several formins in dr-iRPE (Fig. 3d), along with the distinct actin structures observed during internalisation. Dr-iRPE exhibited a larger, flatter actin surface topography (Fig. 2h), suggesting an abundance of heavily branched actin, similar to that found in lamellipodia. In contrast, control samples displayed more numerous, smaller, and protruding actin structures (Fig. 2h), resembling filopodia, which are characterised by linear actin cores and highly dependent on formin activity65. Based on these findings, we inhibited Arp2/3 activity using the compound CK-66666 and inhibited formin activity using the compound SMIFH267.

Formin inhibition in iRPE did not affect the cell height but significantly stiffened the monolayer on a tissue level (Supplementary Fig. 8a, b). In dr-iRPE, the inhibition did not produce a change in cell height, but led to a softer monolayer (Supplementary Fig. 8c, d). When tested for the POS fragments internalisation, the quantification showed a highly variable effect on the number of internalised particles, which nevertheless showed an overall significant reduction in the number of both internalised and bound POS fragments (Fig. 5a, b; Supplementary Fig. 8e and f). In contrast, no difference was detected in SMIFH2-treated dr-iRPE (Supplementary Fig. 8g). Next, we quantified the fragments’ size, which was significantly affected by SMIFH2 only in iRPE, resulting in larger fragments on average (Fig. 5a, b, Supplementary Fig. 8h). Since SMIFH2 affected internalisation of POS only in iRPE, we further analysed 3D-segmented actin and monolayer deformation during internalisation. SMIFH2-treated iRPE monolayers displayed a smoother apical actin topography; however, this was not captured by the quantification of the total apical actin area (Fig. 5c; Supplementary Fig. 8i). Furthermore, SMIFH2-treated iRPE monolayers also bulged out upwards similarly to untreated controls, but showed denser F-actin signal at the junction, suggesting a lateral reinforcement comparable to dr-iRPE (Fig. 5d). In addition, the bulging cells showed a more irregular apical morphology (Fig. 5d).

Although SMIFH2 is widely accepted as a reliable formin inhibitor, it has been reported to have a concentration-dependent off-target effect on Myosin II activity68. To evaluate whether the effects observed with 20µM SMIFH2 were not due to partial Myosin II inhibition, we conducted SMIFH2 treatments across a range of concentrations (5–100µM). We then assessed iRPE and dr-iRPE monolayers for changes in stiffness and phagocytic activity (Supplementary Fig. 8j, l, n, p, r, t). As a positive control for Myosin II inhibition, we used blebbistatin (Supplementary Fig. 8k, m, o, q, s, u). In iRPE, stiffness quantification confirmed a significant monolayer stiffening with 20µM SMIFH2, whereas blebbistatin treatment induced an expected softening as cellular contractility is reduced (Supplementary Fig. 8j, k). Differently, dr-iRPE treated with SMIFH2 displayed an overall softening effect, most prominent at 100µM, consistent with the trend observed following blebbistatin treatment (Supplementary Fig. 8l, m). Finally, analyses of the POS internalisation process revealed that both SMIFH2 at all concentrations and blebbistatin reduced the number of internalised POS particles in iRPE. Still, only the low concentrations of SMIFH2 (5 and 20µM) led to increased average phagosome size, while blebbistatin did not affect fragment size (Supplementary Fig. 8n–q).

In dr-iRPE, SMIFH2 significantly reduced the number of internalised particles only at 5µM with no effect at higher concentrations, whereas blebbistatin reduced it (Supplementary Fig. 8r, s). Average phagosome size was clearly reduced by SMIFH2 treatment in dr-iRPE, in opposite direction to iRPE (Supplementary Fig. 8t), while blebbistatin did not affect the phagosome size (Supplementary Fig. 8u). Overall, while we cannot fully exclude the possibility of minor off-target inhibition of Myosin II by 20µM SMIFH2, we are confident that Myosin II inhibition does not drive our results. This confidence stems from the clear differences in trends for iRPE and dr-iRPE regarding both stiffness and internalised POS number and volume, as well as the contrasting effects seen with blebbistatin treatments.

The inhibition of Arp2/3 using CK-666 in iRPE did not change the cell height or stiffness, whereas in dr-iRPE, the treatment led to a slightly increased cell height (Supplementary Fig. 9a, b), but no significant changes in monolayer stiffness (Supplementary Fig. 9c, d). During phagocytosis, no effect was detected when iRPE were treated, but in dr-iRPE, the treatment led to an increased number of internalised particles (Fig. 5e, f; Supplementary Fig. 9e, g), while the average internalised particle size and the number of bound particles remained comparable (Fig. 5e, g, Supplementary Fig. 9f, h). As the phagocytic process was affected only in dr-iRPE, we proceeded to analyse the morphology of the 3D-segmented cortical actin during internalisation. The analyses showed the occasional presence of thinner and longer ruffles (Fig. 5g), which resulted in a significant increase in the quantified apical actin area (Supplementary Fig. 9i). Furthermore, orthogonally, the treated dr-iRPE showed some degree of bulging during internalisation, which was more pronounced than in the untreated dr-iRPE (Fig. 5h).

Overall, these results demonstrate that modulating actin cytoskeleton remodelling by interfering with actin nucleation effectively altered cortical actin responsiveness during internalisation. These manipulations were sufficient to replicate or rescue differences in POS fragments internalisation observed in dr-iRPE. Particularly, Arp2/3 activity appears to be related to cell bulging, modulating apical actin area and cell height, which impacts the number of internalised particles, while formins activity modulates tissue-level stiffness, affecting both the number and size of the internalised particles.

These insights strongly indicate that dr-iRPE cells establish a new mechanical equilibrium, in which the actin cytoskeleton acts as a critical link between tissue structure, mechanics, and the execution of phagocytosis. The altered organisation of the actomyosin network enhances contractility at cell–cell junctions, and simultaneously reduces the capacity for apical remodelling required for the formation of phagosomes. We therefore conclude that, in response to reduced cell density, the RPE monolayer reroutes actin-related processes toward reinforcing tissue integrity. This shift diminishes the plasticity of the actin-myosin network, which is essential for efficient POS phagocytosis (Fig. 6).

Methods
Preparation of cell hydrogel substrates
The surface of 20mm glass-bottom dishes (Cellvis, D35-20-0-N) or 6-well glass bottom plates (Cellvis, P06-1.5H-N) was activated to allow hydrogel attachment by etching with 0.1M NaOH (Merck, 106498), followed by reaction with 4% (3-Aminopropyl)triethoxysilane (Sigma Aldrich, 440140) in isopropanol (VWR, 20842) and final incubation with 1% Glutaraldehyde (Merck, 354400) in double distilled water (ddH2O) (B. Braun, 0082479E).

Polyacrylamide hydrogels of 4kPa stiffness were prepared from an 5%/ 0.1% Acrylamide/Bis-Acrylamide (Bio-Rad 1610140, Bio-Rad 1610142) mixture in ddH2O by radical polymerisation upon addition of the initiators ammonium persulfate (Sigma Aldrich, 248614) at 0.5% (vol/vol) and N’-Tetramethylethylenediamine (Sigma Aldrich, 411019) at 0.05% (vol/vol)104.

Hydrogels were trimmed to size of 4mm before functionalisation with Laminin-511 (Biolamina, LN511) and Laminin-332 (Biolamina, LN332) at a concentration of 15µg/ml respectively. Briefly, a functionalisation solution was prepared from ddH2O, 0.5 M HEPES (Carl Roth, 9105) pH 6, 100% ethanol (Sigma Aldrich, 32205) and 0.2% (vol/vol) Bis-Acrylamide. The solution was degassed for 20 min and was completed by adding 0.2% Di(trimethylopropane)tetraacrylate (Sigma Aldrich, 408360), 3% (wt/vol) Hydroxy-4′-(2-hydroxyethoxy)-2-methylpropiophenone (Sigma, 41089 and 0.03% (wt/vol) Acrylic acid N-hydroxysuccinimide ester (Sigma, A8060)105. Hydrogels were dehydrated with 70% ethanol for 5 min, incubated under UV for 10 min with the functionalisation solution and washed with 25 mM HEPES and PBS for a total of 20 min before adding the protein solution. The gels were incubated with the protein solution overnight at 4°C.

Culture of human-induced pluripotent stem-cell-derived RPE
HiPSCs-derived RPE cells (Fujifilm Cellular Dynamics, R1102) were seeded in 24-well plates coated with 2.5μg/ml vitronectin XF (StemCell Technologies, 07180). Cells were cultured in MEM-alpha with nucleosides (Thermo Fisher, 12571063), supplemented with 1x Glutamax (Thermo Fisher, 35050061), 5% (vol/vol) Knockout Serum replacement (Gibco, 10828028), 1% (vol/vol) N2-Supplement (Gibco, 17502048), 55nM Hydrocortisone (Sigma Aldrich, H6009), 250µg/ml Taurine (Sigma Aldrich, T8691), 14pg/ml Triiodo-L-thyronine (Sigma-Aldrich, T5516) and 25µg/ml gentamycin (Gibco, 15750060) following the manufacturer’s instructions. The medium was exchanged every other day. For seeding on hydrogels, cells were detached from the 24-well plate by 30min incubation with 5mM PBS/EDTA (Sigma-Aldrich, E6511), followed by 5min incubation with TrypLE Express (Gibco, 126050). Cells were seeded on the UV-sterilised functionalised hydrogels at 5000–6000 cells/mm2. The medium of cells on hydrogels was exchanged halfway twice a week.

On-demand apoptosis for generation of density-reduced iRPE (dr-iRPE)
Apoptosis was induced by expression and on-demand dimerisation of recombinant FKBP-caspase8 (FKBP-casp8) fusion protein41. The FKBP-casp8 containing plasmid was delivered packaged in AAV5 (VectorBuilder, VB230127-1043eaq). Briefly, on day 10 after seeding on the hydrogel, cells were transduced with the virus at 7×106 infectious units per cell for 24h. On day 15, 120h after the transduction, FKBP-casp8 was dimerised by addition of 100nM AP20187 in medium (Sigma Aldrich, SML2838) for 24h. Paired control monolayers were also subjected to the AP20187 treatment. The read-out was performed on day 20, or 96h after apoptosis treatment finalisation.

Virus, mixed and medium-transfer controls
To exclude effects of the AAV as a delivery system, we used an AAV5-packaged construct that promotes the expression of green fluorescent protein (GFP) (VectorBuilder, VB10000-9287ffw) as a control. Cells were transduced and treated the same way as described for FKBP-casp8 expressing cells, including same timing of viral transduction, AP20187 treatment and read-out.

To obtain a mixed population of transfected and naïve cells, 2 days prior to seeding on hydrogels, we transduced hiPSCs-derived RPE in a 24-well plate at 7×106 infectious units per cell for 24h with the FKBP-casp8 vector (VectorBuilder, VB230127-1043eaq). The virus solution was replaced with fresh medium after 24h. After another 24h, we seeded a 10%/90% mixture of the transduced hiPSCs-derived RPE cells and naïve hiPSCs-derived RPE on the hydrogels as described above. On day 15, corresponding to the same apoptosis timing of dr-iRPE generation, apoptosis was induced by addition of 100nM AP20187 (Sigma Aldrich, SML2838) for 24h. The read-out was performed at 96h after apoptosis treatment finalisation.

To exclude possible effects of the release of apoptotic debris during the apoptosis, 16h after AP20187 addition, the medium from dr-iRPE samples was transferred onto control conditions. The medium was then replaced after apoptosis treatment finalisation. 24h after apoptosis finalisation, the medium from dr-iRPE samples was again transferred to control samples and the samples were maintained in this conditioned medium until read-out.

EdU incorporation-based proliferation assay
Proliferation was assessed by evaluating the incorporation of the base 5-Ethinyl-2’-Desoxyuridin (EdU) with the Click-iT EdU kit (Thermo Fisher, C10337). Cells were incubated with 1:1 mixture of 20µg/ml EdU and cell culture supernatant for 24h. The subsequent EdU development reaction was performed following the manufacturer’s instructions. Monolayers were imaged completely as overviews of stitched tiles at inverted epi-fluorescent microscope Zeiss Axio Observer 7. Positive cells were identified by using the particle analysis tool in Fiji and their number was normalised to the total number of cells, as detected by DNA labelling by DAPI (1μg/ml, Invitrogen D1306).

Photoreceptor fragment phagocytosis assay
Photoreceptor outer segment fragments were isolated from porcine eyes and labelled with FITC (Thermo Fisher, F1906), following an established protocol106. For the assay, the vial was thawed and washed with medium at 2300g for 5min. The pellet was resuspended in complete hiPSCs-derived RPE medium together with the bridging molecules human Protein-S (Coachrom, pp012A) at 2µg/ml and human MGF-8 (R&D Systems, 2767-MF-050) at 2.4µg/ml. An approximate ratio of ten photoreceptor outer segment fragments per RPE cell seeded was used. Cells were incubated for 4h, washed four times with DPBS with Ca2+ and Mg2+ (Gibco, 14040117) and fixed with 2% paraformaldehyde (Merck, 818715) for 8min at room temperature (RT). Samples were then permeabilised with 0.3% Triton-X-100 (Sigma Aldrich, T8787) for 2min and labelled for actin with Phalloidin AF568 (Molecular probes, A12380). Three z-stacks per monolayer were recorded at Zeiss LSM710 Confocal Microscope with ZEN Black 2.1 SP3 software with 40× water immersion objective (ZEISS, 420862-9970-000) in Airyscan superresolution mode. The subsequent analysis was performed in Imaris v10.2.0 (Oxford Instruments).

Nanoindentation and dynamic mechanical analysis
Nanoindentation was performed on a Chiaro Nanoindentor (Optics11 Life) mounted on an inverted epi-fluorescence microscope Zeiss Axio Observer 7 with environmental control. Measurements were performed in the Piuma V3.5.0 software on living cells in displacement control mode using a spherical probe of 20µm diameter and 0.025 N/m stiffness. Three matrix scans of 16 indentations were recorded per monolayer, indenting 6µm into the cells. Dynamic mechanical measurements were performed at 1, 2, 4 and 10Hz to allow for viscoelastic characterisation. For the analysis, indentation portions between the contact point and 2.5µm were fitted using the Hertz model in the Optics11 Analysis Software Data Viewer v2.6.0, allowing a range for the contact point fit of 30% of the maximum load. Effective Young’s modulus values of fit R2 better than 0.95 were used for further statistics. Loss and storage moduli were determined for the different frequencies (1, 2, 4 and 10Hz) using the software default methods.

Immunofluorescent labelling and imaging
Monolayers on hydrogels were fixed with 2% paraformaldehyde (Merck, 818715) for 8min at RT and permeabilised with 0.3% Triton-X-100 (Sigma Aldrich, T8787) for 2min at RT. Samples were then blocked with 1% BSA (Serva, 11930) in PBS for 30min at RT, followed by incubation with primary antibodies in blocking solution overnight at 4°C. The following primary antibodies were used: phospho-myosin light chain 2 (Ser19) (Cell Signalling Technology, Cat# 3671) at 1:50, vinculin (Sigma-Aldrich, Cat# V4139) at 15µg/ml, ezrin (Abcam, Cat# ab4069) at 10µg/ml and non-muscle myosin heavy chain IIB (BioLegend, Cat# 909901) at 4µg/ml. After washing, secondary antibodies diluted in PBS were added for 2h at RT. Secondary antibodies included: Alexa Fluor 488 anti-mouse IgG (H+L) (Invitrogen, Cat# A11001) at 8µg/ml, Alexa Fluor 594 anti-mouse IgG (H+L) (Invitrogen, Cat# A11005) at 8µg/ml, Alexa Fluor 594 anti-rabbit IgG (H+L) (Invitrogen, Cat# A11012) at 8µg/ml, Alexa Fluor 647 anti-rabbit IgG (H+L) (Jackson ImmunoResearch/Dianova, Cat# 111-605-144) at 7.5µg/ml, Alexa Fluor 488 anti-rabbit IgG (H+L) (Jackson ImmunoResearch/Dianova, Cat# 711-546-152) at 8µg/ml. Samples were washed and mounted in Mowiol mounting medium (Carl Roth, 9002-89-5) before imaging. Unless started otherwise, samples were imaged at Zeiss LSM710 Confocal Microscope with 40× water immersion objective (ZEISS, 420862-9970-000) in Airyscan superresolution mode as z-stacks of 0.5µm optical slicing. Uniform imaging settings were used between samples within the same experiment. Images were then inspected and further processed for different analyses using Fiji107.

Bulk RNA sequencing
RNA isolation for RNA sequencing was performed using the RNeasy Micro RNA isolation kit (Qiagen, 74004) with DNase treatment following the manufacturers protocol. Bulk RNA sequencing was performed by the Life & Brain NGS Core Facility (Bonn, Germany) after QuantSeq 3’-mRNA Library Prep/ Lexogen enrichment at NovaSeq 6000 with a depth of 20M reads. The raw data was processed using nf-core/rnaseq v3.12.0 (https://doi.org/10.5281/zenodo.1400710) of the nf-core collection of workflows108. The pipeline was executed with Nextflow v23.04.1109. The complete dataset is available on the Gene Expression Omnibus repository (GEO accession number GSE297557).

Differential expression analysis was performed using DESeq2 package110 in R v4.4.3, applying an alpha of 0.05. Genes were considered differentially expressed at an adjusted p-value lower than 0.1.

Heatmaps were generated from the DESeq data set after z-normalisation using the package pheatmap111 and a custom-curated gene list. Functional enrichment of terms related to differentially expressed genes was performed using the g:Profiler online tool112 using the terms GO molecular function, GO cellular component and GO biological process as well as the pathways KEGG, Reactome and WikiPathways. For comparison, publicly available data on the ageing human RPE was obtained from Butler et al.53 and Cai et al.52.

Immunoblotting
Samples were harvested using RIPA buffer113 with phosphatase inhibitors PhosSTOP™ (Roche, 4906845001) and were subjected to DNA digestion for 15min by Pierce Universal Nuclease for cell lysis (Thermo Fischer, 88700) prior to addition of Lämmli buffer114. Samples were then heated to 95°C for 7min and ca. 10µg were loaded onto a 10 to 14% (depending on target protein molecular weight) polyacrylamide gel115 for SDS-PAGE together with a ProSieve Quad color Protein Marker (Lonza, 00193837). Proteins were blotted onto Immobilion-P PFDV membrane (Thermo Fischer 88518) and labelled with primary and secondary antibodies for different targets. The following primary antibodies were used: phospho-myosin light chain 2 (Ser19) (Cell Signalling Technology, Cat# 3671) at 1:1000, ezrin (Abcam, Cat# ab4069) at 1µg/ml and phospho-ezrin (Thr567) (Invitrogen, Cat# PA5-37763) at 1µg/ml. Secondary antibodies included: HRP anti-rabbit IgG (Abcam, Cat# ab99696) at 1:5000 and HRP anti-mouse IgG (H+L) (Jackson ImmunoResearch/Dianova, Cat# 115-035-003) at 1:5000. Bands were detected at the Fusion Solo (Vilber Lourmat) with the FusionCapt Advance Software after detection with the Pierce™ ECL Western blotting substrate (Thermo Fischer, 32106). The signal was quantified in Fiji107 by using equal-sized region of interest (ROI) for all bands and measuring the mean grey value. The measurements for the proteins of interest were normalised to the housekeeping protein GAPDH.

Modulation of actin nucleator and myosin II activity
Actin nucleation by Arp2/3 was modulated by using CK-666 (Tocris Bioscience, #3950) at 500nM in medium. After 1h incubation, the monolayers were characterised by nanoindentation. For phagocytosis studies, the monolayers were incubated for 30min with 500nM CK-666, followed by incubation with the photoreceptor fragments and the bridging molecules in CK-666 containing medium for 4h. For modulating formin activity, 20µM SMIFH2 (Tocris Bioscience, #4401) was used instead keeping the same timing and procedure. DMSO at a paired concentration was used as carrier control. For the incubation with different SMIFH2 concentrations, a series of dilutions was prepared starting from a 100µM solution, resulting in a dilution series of 100, 40, 20 and 5µM concentrations. For nanoindentation and POS functional studies, the same timing and procedure were used as described above with DMSO as a carrier control. Myosin II inhibition was performed using 5µM blebbistatin (Abcam, ab120425) for 1h before nanoindentation and for 30min before POS addition in blebbistatin-containing medium (see above).

Analysis of extrusion events
Extrusion events were quantified from live cell imaging of apoptosis induction on SiR-Actin (Spirochrome, SC001) labelled cells. The last 15 hours of apoptosis induction were used for the quantification which relies on extrusion events being visible as very bright spots. The video was aligned to correct for stage shifts, smoothed using a Gaussian filter and overlayed in a maximum intensity z-projection in Fiji. The resulting image was thresholded to generate a binary mask and extruded cells were counted using the particle analysis tool in Fiji.

Analysis of cell density
Cell density was quantified from snaps of nuclear staining obtained on epi-fluorescence microscope at 20x magnification. Three to four snaps were used per condition per experiment. Nuclear signal was smoothed using a Gaussian filter, then detected by thresholding. The binary mask was subjected to watersheding and the number of nuclei was counted by using the particle analysis tool in Fiji. Density was calculated by dividing the obtained nuclei count to the area of the field of view. Data originates from at least three independent experiments.

Analysis of cell height
Cell height was determined from super-resolution z-stacks of F-actin staining in cross-sectional view. Cells were imaged fixed after mounting in Mowiol, which leads to lower absolute height compared to the height detectable in live-cell imaging. Ten cells per stack were measured with the line tool in three stacks per condition from three independent experiments.

Analysis of cell outline segmentation and shape factor calculation
Cellular outlines from F-actin labelling were segmented using Cellpose v.3.0.11116. The generated regions of interest (ROI) were imported in Fiji and the cell area and perimeter were measured to calculate the dimensionless shape factor117,118.

Analysis of stress fibres
Cells with stress fibres were manually counted using the multipoint tool in Fiji and normalised to the cell number as calculated after segmentation of cells via Cellpose v.3.0.11.

Analysis of microvilli length
Microvilli length was characterised from cross-sectional views of ezrin labelling in Fiji. Briefly, at least three cross-sections from at least three stacks per condition from three independent experiments were blurred with a Gaussian blur and a threshold was applied to capture the structure. The resulting masks were selected via the magic wand tool and an eclipse fit was performed. The minor axis was used as an approximate measure of the microvilli length.

Photoreceptor fragment phagocytosis analysis
The photoreceptor fragment phagocytosis was characterised using the Software Imaris v10.2.0 (Oxford Instruments). Briefly, the actin labelling was used to create a surface capturing only the apical actin portion of the monolayer. The average z-position of the surface was determined as a cuff-off height to distinguish internal vs external particles. Separately, photoreceptor fragments were segmented as a surface and the cut-off height was used to distinguish between internalised and external fragments. The count and volume of the fragments of both populations was recorded and used for further analysis.

Quantification of apical actin area
The apical actin area of 300×300 pixel field of view was segmented from z-stacks of phalloidin staining during photoreceptor internalisation as a surface in Imaris v10.2.0 (Oxford Instruments). For the region of interest, only a set number of slices including the same upper portion of the cell monolayer were used. The covering area, as internally determined by the software, was then noted.

Quantification of vinculin junctional to cytoplasmic ratio
Junctional signal of vinculin was quantified as a ratio of junctional to cytoplasmic mean grey value in single optical sections. For this, paired F-actin signal was used to segment cell outlines via Cellpose v.3.0.11 as described above. The generated ROIs were imported in Fiji and a custom macro was used to slim-down the ROIs, invert the mask, watershed the signal and generate ROIs that encompass the junctional regions around cells. The generated junctional ROIs were used to measure grey value as an average per image. The slimmed-down cell ROIs were used to measure the cytoplasmic signal. The average values of junctional and cytoplasmic signals per image were used to generate the junctional to cytoplasmic vinculin measurements.

Quantification of junctional pMLC2
Paired samples from the same experiment were stained with the same antibody solution and imaged with the same imaging settings. In single optical sections at the junctional level, pMLC2 signal was thresholded in Fiji making sure the junctional signal is captured. The mean grey value was measured only within the thresholded region. At least three images were measured per condition per experiment.

Quantification of MRTFA
MRTFA was quantified as nuclear to cytoplasmic ratio of the mean grey value. For this, paired cell nuclei signal (DAPI staining) was segmented via Cellpose v.3.0.11 to generate nuclear ROIs. Nuclear MRTFA was measured as average of the mean grey value of all nuclei ROIs within the image. Cytoplasmic MRTFA was measured after setting nuclear ROIs pixels as zero and thresholding the rest of the image. Then, the mean grey value of the thresholded, non-zero region was measured.

Brillouin microscopy
Brillouin imaging was performed on a commercial setup based on an inverted Nikon Ti-2 Eclipse microscope using a vertically polarised, continuous-wave laser (Cobolt Flamenco, λ=660nm, ~100mW) beam with a temperature-tuned air-spaced etalon (LightMachinery) attenuated by 0.6 optical density (OD) filter, resulting in approximately 9mW incident power at the sample. Light was coupled into a confocal circulator (LightMachinery, HF-13311) and directed to the microscope through a polarising beam splitter and a quarter wave plate (QWP) yielding circularly polarised light. Measurements were performed on a motorised translation stage (Prior Scientific, H117E2NN) using an air objective (Nikon S Plan Fluor 20X NA 0.45), achieving an estimated 0.73µm lateral and 6.52µm axial resolution. Brightfield and fluorescence imaging were performed sequentially using a Teledyne sCMOS camera and epi-fluorescence illumination (Nikon D-LEDi).

Backscattered Rayleigh and Brillouin signals were collected with the same objective and routed back through the confocal circulator. The Brillouin signal was acquired using a LightMachinery HyperFine spectrometer (HF-8999-PK-660), equipped with a “pump killer” (PK) module with an air-spaced, pressure-tuned etalon to suppress the elastically scattered Rayleigh light and laser line reflection (~60dB attenuation). Frequency-shifted Brillouin Stokes and anti-Stokes components were directed into the spectrometer, comprised of a virtually imaged phase array (VIPA) and a reflective echelle diffraction grating for spectral dispersion. The cross-dispersed orders were focused onto the CMOS camera sensor (Hamamatsu ORCA-Fusion, C14440-20UP) simultaneously acquiring the full spectrum. The typical acquisition time of a single spectrum is 500ms, with a spectral resolution <1pm.

Spectral data acquisition and raster scanning was facilitated with LightMachinery LabView-based software. The x-y pixel coordinates on the CMOS sensor are converted to wavelength (or frequency) and a Lorentzian fit is applied over each Brillouin peak as described by equation (1).

I(ω) = I0 * (ΓB/2)^2 / ((ω - ωB)^2 + (ΓB/2)^2) (1)

The final Brillouin shift ωB and linewidth Γ values for a given spectrum were computed by averaging the corresponding values from the Stokes and anti-Stokes peaks forming a single voxel in the raster scan. Z-scans were acquired as a 3×3 matrix in a step size of 20μm and z-step of 2μm, z-range of 80μm, while apical and basal scans were acquired over the region of 50×50μm with a lateral step size of 1.5μm. The z-position of apical and basal Brillouin scans within the monolayer was selected based on the appearance of the cells in brightfield and SiR-actin fluorescence images. The scans were limited to 15 minutes to obtain an optimal raster scan resolution while maintaining the cell viability. It should be noted that although the average laser power at sample is within the range used in biological samples, by comparing brightfield images taken before and after the Brillouin scan, some deformation of the sample was observed. This could have occurred due to the heat of the laser affecting the gel or the cells as the RPE cells’ pigmentation contributes to the linear laser light absorption.

Brillouin shift data analysis
For segmentation, the Brillouin shift maps from basal layers were linearly scaled and converted into 8-bit grayscale images, proportionally reflecting the original Brillouin shift values. The grayscale images were segmented using Cellpose v.3.0.11 to identify the individual structures within the images. Following segmentation, two populations were defined: “core”, corresponding to segmented regions, and “interstitium”, corresponding to areas between them. These modified images were then imported into a custom-written MATLAB script, where appropriate binary masks were created and applied to the original Brillouin shift maps to separate the “core” and the “interstitium”. For each raster scan, the mean Brillouin shift and the standard deviation were calculated separately for the two regions. The 3×3 raster z-scans resulted in nine representative z-axis line profiles per region, capturing mechanical transition across the sample. Each profile typically displayed a stiffness change from the hydrogel to the monolayer and then from the monolayer to the surrounding cell media. For each sample, 5 of these scans were obtained.

The Brillouin Shift data were imported into a custom-written MATLAB script to identify the local peaks or, where absent, plateaus in the Brillouin shift profiles. Each line profile was smoothed with a Savitzky-Golay filter, the peak was detected using MATLAB’s findpeaks function, with a minimum width imposed to prevent false peak identification. A gradient of the filtered z-line was taken. To refine peak localisation, a Gaussian fit was applied across a range of filtered Brillouin shift values. The fitting range was bound by the z-positions at which the local maximum and the minimum of the derivative of the z-line was within an appropriate range from the previously identified peak. The fitted peak value and position were retrieved. In the case of no peak detection, the code identifies the onset of a drop in the plateau by considering the change in the derivative over a moving average window. The corresponding plateau value was then recorded. The performance of the script was checked for individual z-profiles to exclude outliers.

Statistics and reproducibility
Statistical analysis was performed using GraphPad Prism 10 on paired averaged values originating from at least three independent biological experiments. Datapoints from at least three technical replicates or from matrix measurements within the samples, epithelial monolayers, were averaged into representative values from a biological replicate. Assuming normal distribution, the data was tested with a paired, two-sided test depending on experimental design with statistical tests and number of biological replicates for each graph given in the captions of the figures. Samples were randomly assigned to experimental conditions. Data were excluded only if obvious sample damage occurred or if technical failures prevented reliable analysis. No other data were excluded. The investigators were not blinded to allocation during experiments or outcome assessment.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "Reviewer report on “Tissue organization alone drives age-like changes in a postmitotic epithelium” by Piskova et al. (2025)\nTissue mechanics (including elasticity, viscosity, pressure, compliance, among others) play a critical role in cancer\ndevelopment, influencing how tissue grows, functions, and responds to external mechanical cues. This study explores how\ncellular organization alone—without genetic mutations or external stressors—can mimic ageing effects in the retinal pigment\nepithelium (RPE), a crucial postmitotic tissue in the eye. The authors used an in vitro model system based on human\ninduced pluripotent stem cell-derived RPE (iRPE) cultured on soft hydrogels. The authors showed that the described AgeD-\niRPE model in vitro model recapitulates structural ageing features previously shown in human RPE tissues including (1)\nlarger cell area and reduced cell height, (b) shortened and disorganizes microvilli, and (c) transcellular actin stress fibers.\nThen, the authors proceeded to investigate if the model recapitulates impaired internalization of outer segment fragments,\nand indeed they showed that AgeD-iRPE monolayers exhibit deficient internalization. Then, the authors hypothesized that\ninternalization deficiency could be due to active remodeling of the actomyosin network. The authors performed RNA\nsequencing to investigate changes in mRNA expression of actin regulatory proteins and observed a vast amount of\nupregulated and downregulated actin associated genes. Next, the authors investigate changes in mechanical properties by\nnanoindentation and Brillouin microscopy. The authors observed a significant increase in elasticity by Young’s modulus and\nstorage modulus that seem to correlate with an apparent increase in phosphorylated myosin (pMLC2) and vinculin\nexpression. Surprisingly, with Brillouin microscopy the authors observed an opposite mechanical behavior with decrease\nBrillouin shift indicating softening. Last, the authors performed small molecule pharmacological perturbations to inhibit actin-\nbased nucleation either inhibition of branched actin nucleation (ARP2/3) by CK-666 or inhibition of linear actin nucleation\n(formins) by SMIFH2. The authors observed that by modulating actin nucleators phagocytosis is impacted (a) Arp2/3\ninhibition (via CK-666) improved internalization in AgeD-RPE whereas (b) formins inhibition (via SMIFH2) increase\nphagosome size in control iRPE.\nAltogether, this manuscript seems to describe a new in vitro model to investigate RPE ageing, however there are technical\nconcerns in some conducted experiments and some controls are missing. The current manuscript version is rather\nsuperficial and of incremental nature. I believe a revised version of this manuscript will be more suitable to be submitted for\npublication consideration in a specialized cell biology journal. Therefore, I do not recommend the publication of this\nmanuscript in Nature Communications.\nComments:\n1. Figure 1: The transcellular stress fibers the authors describe in the manuscript are not clearly visible. Even trying to\ndistinguish the transcellular stress fibers of zoomed-in images from the AgeD-iRPE that has been highlighted with arrows\nare hard to see (Fig. 1F).\n2. Figure 1: In addition, to this reviewer looking at the F-actin images from both control iRPE and AgeD-iRPE it seems that\nthere is the same amount of transcellular stress fibers on both conditions (Fig. 1E). Looking at these images alone is hard to\ncompare both conditions and judge if there is indeed an increase of 10 folds more percentage of transcellular stress fibers. I\nsuggest the authors add zoomed-in images from the control iRPE in this figure (Fig. 1F) next to the zoomed-in images of\nAged-iRPE so readers can compare both conditions and judge the changes by themselves.\n3. Figure 4: Can the authors provide more detailed explanations of why the Brillouin shift is significantly lower in AgeD-iRPE\nwhen compared to control iRPE and how this is contradictory to the obtained microindentations results. There is published\nwork correlating changes in microindentation based mechanical properties changes with Brillouin shift and have shown that\nwhen it is observed an increase in stiffness measured by microindentation measurements this corresponds to a significant\nincrease in measured Brillouin shift (Weber et al. Physical Biol. 14, 2017; Scarcelli et al. Nat. Methods 12, 2015). Therefore,\nthe results shown in this work are contradictory with published work and need to be clearly clarified and discussed.\n4. As briefly discussed in the discussion section of this manuscript, SMIFH2 has been shown to have nonspecific off target\neffects and on many occasions strongly inhibits non muscle myosin II myosin ATPase activity (Nishimura et al. J. Cell Sci.\n134, 0221). In addition, it seems to inhibit several other non-muscle myosin types. Therefore, without a targeted perturbation\nof formins protein expression to validate the SMIFH2 results or by performing SMIFH2 experiments at different\nconcentrations it is hard to judge if these results are valid or artefactual.\n5. Continuing on this line about the formin inhibition results, a lot of published work has shown an opposite effect to what the\nauthor describes in this manuscript. Previous work (Mittal et al. Commun. Mater. 5, 2024; Efremov et al. BBA-Mol. Cell Res.\n1853, 2015) inhibiting formins in many different cell types including (3T3 fibroblasts, DU145 cells, and Vero cells) has\nshown a drastic softening effect. Therefore, can the authors comment on this surprising contradiction.\n6. Control experiments are missing. The authors must perform experiments on AgeD-iRPE treated with SMIFH2 and control\niRPE treated with CK-666 to contrast the effects of inhibition in both conditions.\n7. Minor comment. On several occasions throughout the paper the authors make statements such as “first-of-a-kind”,\n“for the first time”, and others. I suggest the authors tone down the manuscript and avoid using such statements.", "reviewer_2": "In this study, Piskova and colleagues developed and characterized an in vitro aging model of retinal pigmented epithelium\n(RPE) to investigate how actin contributes to the reduced phagocytosis of photoreceptor outer segments during aging. This\nwork presents novel mechanisms that will be of interest to a broad audience.\nThe experimental design is sound with appropriate controls, and the methodological approaches are well-suited\nto address the research questions. The data analyses appear rigorous, and the conclusions are supported by the presented\nevidence.\nHowever, the manuscript has several presentation issues that limit its accessibility. The abstract lacks clarity due to ill-\ndefined terminology, and the results section focuses heavily on data presentation but provides insufficient interpretation of\nthe biological significance of the observations.\nOverall, I support this manuscript for publication, but the authors must address the clarity and presentation issues outlined\nbelow to make their important findings accessible to the broad readership of Nature Communications.\nMajor Comments\n1) The novelty and impact of this study are difficult to understand from the abstract, primarily due to the use of numerous ill-\ndefined and unconventional terms. For example, what do the authors mean by \"mechanical homeostasis\" and \"structural\nageing\"? Similarly, the term \"impaired-plasticity\" requires clarification. More critically, the main research question addressed\nin this study is unclear. The authors state they are investigating \"the connection between age-related structural changes and\nmechanical homeostasis,\" but it is not evident how this relates to their proposed answer: \"a mechanistic link between\ncytoskeletal adaptation and phagocytic dysfunction in ageing retinal pigment epithelium.\" I recommend rephrasing the\nabstract to clearly define key terms, explicitly state the research question, and demonstrate how the findings address that\nquestion.\n2) The novelty of the findings is not sufficiently emphasized throughout the manuscript. This may be because much of the\ndata focuses on characterizing the AgeD-iRPE model, showing results consistent with in vivo data such as reduced cell\nheight, shortened microvilli, actin cytoskeletal alterations, and decreased phagocytic ability. While the cellular and\nsubcellular mechanical characterization of aging RPE is novel, the authors should better highlight the importance of these\nmechanical measurements and their meaning.\nA few key findings can be emphasized more clearly: first, the establishment of an in vitro aging RPE model, which\nrepresents a significant methodological advance for the field. Second, the challenge to the current notion of cellular\n'hypertrophy' is particularly noteworthy. The results demonstrate that aging involves not cell volume enlargement but\nrather 'tissue stretching from within,' pointing to individual cell volume conservation. This novel concept, which challenges\ntraditional views of cellular aging, should be highlighted more prominently in the results section rather than being buried in\nthe discussion.\n3) Related to point #1, the authors frequently use poorly defined terminology that obscures their interesting findings. Several\nexamples require clarification:\n- What do the authors mean by \"a new mechanical equilibrium and homeostasis\" and \"actin-myosin network plasticity\" (line\n368)?\n- How should readers interpret \"a shift in the range of mechanical equilibrium\" (line 431)?\n- What is meant by \"intercellular reinforcement\" (line 481)?\n4) The results section is difficult to follow because while data are presented and quantifications are explained, there is\ninsufficient interpretation of what the results mean and why these observations are important. The authors need to better\nexplain the significance of their findings. Several examples illustrate this issue:\n- What do the nanoindentation data mean (lines 242-252)?\n- Why does the difference in Brillouin shift at different sections (apical to basal) lead to the conclusion that \"the apical sides\nof the cells undergo the most pronounced reorganisation\" (lines 293-306)? Moreover, what specific type of 'reorganisation'\nare the authors referring to, and why is this important?\n- What is the significance of the segmented Brillouin shift data (lines 311-317)? Why should readers care about \"the\npresence of loosely interdigitated basal membranes\"?\n- Why is the observation that \"age-associated RPE phenotype has a different biomechanical state of the tissue\"\nimportant?\n- What does the finding that \"bulging cells showed a more irregular apical morphology (Fig. 5D)\" imply for RPE function?\nWhy do the authors highlight this observation? A similar question applies to the statement that \"the treated AgeD-iRPE\nshowed some degree of 'bulging' during phagocytosis, which was more noticeable compared to the untreated\nAgeD-iRPE (Fig. 5H).\"\nMinor Comments\n5) Fig. 6: Why does the middle panel showing top views highlight the observed characteristics only in the central cell and not\nin the neighboring cells as well?\n6) Supp. Fig. 3A(2): The screenshot from an application is not appropriate for a scientific publication.\n7) The statement that \"a balance between basal cell-ECM adhesion and apical cell-cell adhesion exists and is normally well\nconserved (78–80)\" (line 456) requires more detailed discussion.", "reviewer_3": "This is a well executed study on the physiological changes of the retinal pigmented epithelial monolayer in response to 2%\napoptosis in vitro. The quality of the experimental design, the chosen assays and the execution is all excellent. The framing\nof the experiments is where this reviewer had much issue. There are flaws in the data interpretation and conclusions which\nprohibit publication or require revision. They make bodacious claims as to what their results signify. To so without\nqualifications of which there are many. They also use duplicitous naming of their model attempting to more closely align\nwhat they are studying (epithelial monolayer response to apoptosis) to what they claim to be studying (epithelial monolayer\naging). Moreover there is no qualifications with regards to the differences between in vivo and in vitro conditions nor the\nknown differences between iPSC-RPE compared to native RPE.\nThe experiments detailed in this manuscript does not support the title. “Tissue alone” is too vague for any understanding\n“alone” claim has not been supported by the evidence of this MS. The in vitro model is lacking many of the conditions in a\nlive aging organism and therefore the conclusions can only be made in that model. Therefore, the title should qualify the\nstatement. “within an in vitro model…\nThe abstract is too general and does not discuss what is done, what results were found and what conclusions from those\nresults can be made.\nThe in vitro aging may exhibit similar morphological changes for very different reasons. Lack of vascular support, lack of the\nability to maintain ionic balance of the media apically and basolaterally, no differential in nutrient access apically to\nbasolaterally. These over time can also lead to a decrease in physiological function and have nothing to do with the native\nenvironment.\nHydrogels do not have a composition resembling native RPE environment. Water content, diffusion gradient,\nThe induced apoptosis is too synthetic to model native RPE aging. Therefore what is being modeled is the cellular response\nto apoptosis.\nThere is no age dependent condition and therefore, the way the experiments are described is a misnomer.\nReplace AgeD RPE with apoptosis responsive ARD-RPE and then the manuscript is on a more accurate foundation from\nwhich to draw conclusions. To mix apoptosis with age is duplicitous.\nThe lack of proliferation in response to apoptosis is an interesting response that warrants further study. What is missing is a\nspectrum of apoptosis responses from 0 to 50% at regular intervals. Is there an apoptosis % that proliferation can be\ntriggered? If so, what are the mechanisms governing this? Is there another more natural apoptosis approach that can support\nthese findings or must it be the manipulation of caspase to lead to these changes within this timeframe?\nThe results are informative so long as the authors are more honest with their claims.\nThere is sufficient detail in the methods section to reproduce the experiments.\nCommend them on the proper protocol for assaying phagocytosis.\nIn general experimental execution and data collection quality is high."}, "author_response": "We thank the reviewer for the constructive feedback. We respectfully disagree that the\nwork is incremental, as the manuscript identifies a previously unrecognised mechanism\nof homeostasis in the RPE and more broadly in postmitotic tissues, and demonstrates\nits consequences for ageing-related dysfunction.\nWe have carefully addressed all specific comments and revised the manuscript to clarify\nthe conceptual advances, strengthen the methodological description, and improve the\nintegration of findings across disciplines. We believe these revisions now more clearly\narticulate the impact of our work within mechanobiology, cell biology, and retinal\nphysiology.\nComments:\n1. Figure 1: The transcellular stress fibers the authors describe in the manuscript are not\nclearly visible. Even trying to distinguish the transcellular stress fibers of zoomed-in\nimages from the AgeD-iRPE that has been highlighted with arrows are hard to see (Fig.\n1F).\nFigure 1f was improved by enlarging the images and adding a zoom-in of a single cell and\nstress fiber to better highlight the phenomenon.\n2. Figure 1: In addition, to this reviewer looking at the F-actin images from both control\niRPE and AgeD-iRPE it seems that there is the same amount of transcellular stress fibers\non both conditions (Fig. 1E). Looking at these images alone is hard to compare both\nconditions and judge if there is indeed an increase of 10 folds more percentage of\ntranscellular stress fibers. I suggest the authors add zoomed-in images from the control\niRPE in this figure (Fig. 1F) next to the zoomed-in images of Aged-iRPE so readers can\ncompare both conditions and judge the changes by themselves.\nFigure 1f now displays both conditions side by side, which indeed allows for better\ncomparison.\n3. Figure 4: Can the authors provide more detailed explanations of why the Brillouin shift\nis significantly lower in AgeD-iRPE when compared to control iRPE and how this is\ncontradictory to the obtained microindentations results. There is published work\ncorrelating changes in microindentation based mechanical properties changes with\nBrillouin shift and have shown that when it is observed an increase in stiffness measured\nby microindentation measurements this corresponds to a significant increase in\nmeasured Brillouin shift (Weber et al. Physical Biol. 14, 2017; Scarcelli et al. Nat.\nMethods 12, 2015). Therefore, the results shown in this work are contradictory with\npublished work and need to be clearly clarified and discussed.\nWe understand the reviewer’s concern, and we are aware of the apparent contradiction,\nbut believe the results are reconcilable because of two major factors:\n1. Indentation-based and optical elastography-based methods differ fundamentally in\nhow they assess mechanical properties.\nSpecifically, they differ in probing frequency, invasiveness (mechanical vs. optical),\nand hydration influence. Notably, a recent study demonstrated that hydration levels\nin heterogeneous materials, such as cells, greatly affect the longitudinal elastic\nmodulus measured by Brillouin microscopy. However, indentation-based\napproaches like AFM and nanoindentation are not impacted by hydration because\nthey allow water to expand transversely during measurement (ref:\n10.1021/acsphotonics.5c00808).\n2. In the nanoindentation vs Brillouin microscopy, we probe the tissue at different length\nscales.\nDuring nanoindentation, we use a 20 µm in diameter sphere that indents into a few\ncells at a time, often impacting on cell-cell contacts. As we were interested in probing\ntissue-level mechanics, so important for epithelia, we did not segregate between\nmeasurements hitting purely on a cell vs measurements hitting on cell-cell junctions.\nThus, our nanoindentation results heavily reflect the stiffness of the cell-cell\njunctions, counteracting the deformation. On the contrary, as our Brillouin\nmicroscopy measurements relied on point-by-point laser scanning within the apical\nand basal optical planes of the monolayers, the cell cortices may have had a higher\ncontribution to the obtained average values rather than the cell-cell contacts, as the\ncortical, non-junctional portion of the images occupies more pixels.\nLastly, direct comparisons between new methods like Brillouin microscopy and\nestablished ones such as nanoindentation are important to reveal discrepancies and\nencourage careful interpretation, since these techniques rely on fundamentally different\nprinciples. As also recently highlighted in a work on retinal tissue (DOI: 10.1088/2515-\n7647/ad5ae3), as complementary, both techniques are necessary for a comprehensive\ncharacterisation of a system.\nTo avoid diverting attention from the main message of our work, we did not emphasise\nthe apparent contradiction with other biological systems (e.g., Scarcelli et al. Nat.\nMethods 12, 2015) or length scales (Weber et al. Physical Biol. 14, 2017) when comparing\nBrillouin microscopy and nanoindentation. Nonetheless, we explicitly mentioned that a\ndirect comparison cannot be made (lines 429-430) and that any difference may be\nattributed to changes in microvilli density and apical actin remodelling (lines 430-432\nand 254-255).\n4. As briefly discussed in the discussion section of this manuscript, SMIFH2 has been\nshown to have nonspecific off target effects and on many occasions strongly inhibits non\nmuscle myosin II myosin ATPase activity (Nishimura et al. J. Cell Sci. 134, 0221). In\naddition, it seems to inhibit several other non-muscle myosin types. Therefore, without a\ntargeted perturbation of formins protein expression to validate the SMIFH2 results or by\nperforming SMIFH2 experiments at different concentrations it is hard to judge if these\nresults are valid or artefactual.\nWe thank the reviewer for the valuable comment and agree that additional perturbation\nexperiments were necessary. We have now included treatments with SMIFH2 at multiple\nconcentrations, as well as the Myosin II inhibitor blebbistatin as a positive control. The\ndata are now shown in Suppl. Fig. 8j–u and presented in lines 314-338 and discussed in\nlines 491-493.\nInterestingly, the effects quantified following SMIFH2 treatment were not consistent with\nthose observed using blebbistatin, particularly at 20 µM SMIFH2, which was the initial\nconcentration used. Notably, SMIFH2 treatment increased iRPE stiffness only at 20 µM,\nwhich rules out a possible inhibitory effect on Myosin function, an effect that would\ninstead be expected to soften the monolayer, as demonstrated by blebbistatin and\nreported in the literature (e.g., Figure 2 in DOI: 10.3389/fcell.2022.946190).\n5. Continuing on this line about the formin inhibition results, a lot of published work has\nshown an opposite effect to what the author describes in this manuscript. Previous work\n(Mittal et al. Commun. Mater. 5, 2024; Efremov et al. BBA-Mol. Cell Res. 1853, 2015)\ninhibiting formins in many different cell types including (3T3 fibroblasts, DU145 cells, and\nVero cells) has shown a drastic softening effect. Therefore, can the authors comment on\nthis surprising contradiction.\nWe thank the reviewer for raising this important point. The studies mentioned (e.g. Mittal\net al. 2024; Efremov et al. 2015) primarily examined single cells. Our work focuses on\npostmitotic, static epithelium where cell-cell adhesions are in mechanical equilibrium.\nTo our knowledge, the mechanical response to formin inhibition has not been previously\ncharacterised in this context. In our system, we believe that the increased stiffness\nreflects a redistribution of tension within mechanically integrated monolayers, rather\nthan a cell-autonomous softening response.\n6. Control experiments are missing. The authors must perform experiments on AgeD-\niRPE treated with SMIFH2 and control iRPE treated with CK-666 to contrast the effects of\ninhibition in both conditions.\nWe have now included the missing controls after performing a new series of experiments,\nand the results are presented in the Sup. Fig. 8 c, d, g and h and in Sup. Fig. 9 c, d, g and\nh. In the text, we have described the results in the last chapter, starting from line 297.\n7. Minor comment. On several occasions throughout the paper the authors make\nstatements such as “first-of-a-kind”, “for the first time”, and others. I suggest the authors\ntone down the manuscript and avoid using such statements.\nWe have now changed our wording and toned down the narrative.\nReviewer #2 (Remarks to the Author):\nIn this study, Piskova and colleagues developed and characterized an in vitro aging\nmodel of retinal pigmented epithelium (RPE) to investigate how actin contributes to the\nreduced phagocytosis of photoreceptor outer segments during aging. This work presents\nnovel mechanisms that will be of interest to a broad audience. The experimental design\nis sound with appropriate controls, and the methodological approaches are well-suited\nto address the research questions. The data analyses appear rigorous, and the\nconclusions are supported by the presented evidence.\nHowever, the manuscript has several presentation issues that limit its accessibility. The\nabstract lacks clarity due to ill-defined terminology, and the results section focuses\nheavily on data presentation but provides insufficient interpretation of the biological\nsignificance of the observations. Overall, I support this manuscript for publication, but\nthe authors must address the clarity and presentation issues outlined below to make\ntheir important findings accessible to the broad readership of Nature Communications.\nWe thank the reviewer for recognising the value of our work and conclusions. We agree\nthat the text greatly benefited from the substantial revisions to clarify the significance of\nour findings. As detailed below, we have now made the suggested changes to the text.\nMajor Comments\n1) The novelty and impact of this study are difficult to understand from the abstract,\nprimarily due to the use of numerous ill-defined and unconventional terms. For example,\nwhat do the authors mean by \"mechanical homeostasis\" and \"structural ageing\"?\nSimilarly, the term \"impaired-plasticity\" requires clarification. More critically, the main\nresearch question addressed in this study is unclear. The authors state they are\ninvestigating \"the connection between age-related structural changes and mechanical\nhomeostasis,\" but it is not evident how this relates to their proposed answer: \"a\nmechanistic link between cytoskeletal adaptation and phagocytic dysfunction in ageing\nretinal pigment epithelium.\" I recommend rephrasing the abstract to clearly define key\nterms, explicitly state the research question, and demonstrate how the findings address\nthat question.\nWe completely rewrote the abstract, clarifying the reviewer's points and avoiding niche\nterms.\n• Now the term “Mechanical homeostasis” is understandable from the context of\nthe first three sentences.\n• The terms plasticity and structural ageing were removed or rephrased.\n• The abstract contains the main question:\n[… Yet, how these adaptations relate to epithelial mechanical homeostasis and\nage-associated functional decline remains poorly understood…]\n• How the findings address the question:\n[…To establish the relationship between structural changes, mechanical\nhomeostasis and function, we developed an in vitro reductionistic model\nmimicking age-related reduction in RPE cell density. Inducing large-scale\napoptosis in postmitotic stem cell-derived RPE monolayers recapitulates\nstructural hallmarks of aged tissue, such as reduced cell height, shortened\nmicrovilli and cytoskeletal reorganisation. This new structure acquires a new\nmechanical equilibrium, evidenced by tissue stiffening and enhanced junctional\ncontractility. Functionally, the monolayers display impaired vision-supporting\nphagocytosis of photoreceptor outer segments. Mechanistically, modulation of\nactin nucleators, Arp2/3 and formins, demonstrates that apicolateral monolayer\ndeformation is critical for phagocytosis and may be compromised in aged RPE…]\n2) The novelty of the findings is not sufficiently emphasized throughout the manuscript.\nThis may be because much of the data focuses on characterizing the AgeD-iRPE model,\nshowing results consistent with in vivo data such as reduced cell height, shortened\nmicrovilli, actin cytoskeletal alterations, and decreased phagocytic ability. While the\ncellular and subcellular mechanical characterization of aging RPE is novel, the authors\nshould better highlight the importance of these mechanical measurements and their\nmeaning.\nA few key findings can be emphasized more clearly: first, the establishment of an in vitro\naging RPE model, which represents a significant methodological advance for the field.\nSecond, the challenge to the current notion of cellular 'hypertrophy' is particularly\nnoteworthy. The results demonstrate that aging involves not cell volume enlargement but\nrather 'tissue stretching from within,' pointing to individual cell volume conservation. This\nnovel concept, which challenges traditional views of cellular aging, should be highlighted\nmore prominently in the results section rather than being buried in the discussion.\nWe appreciate the reviewer’s feedback, which helped us sharpen the focus of our\nnarrative. We have revised the manuscript accordingly to address the points raised.\n• On the importance of the mechanical measurements:\nWe have added explanations in the introduction, lines 31-36, results, lines 271-\n281 and discussion, lines 453-468.\n• On the in vitro model:\nAlthough we align with the reviewer's enthusiasm to define this model as a\npossible ageing model, we want to be careful to consider this reductionistic in\nvitro system representative of the complex in vivo ageing process. We aimed to\ndissect only one aspect of ageing RPE, focusing more on the structural and\nmechanical perspectives. As correctly pointed out by reviewer #3, with our model,\nwe study the direct response to cell-density reduction, which helps us to\nspeculate on RPE ageing in the native environment. More studies are necessary\nto understand how far this approach can be considered as an ageing model.\nWe included more clarification about our model and how it helps to make age-\nrelated conclusions in the introduction, lines 36-40, results, lines 124-126, and\nabstract: [… To establish the relationship between structural changes,\nmechanical homeostasis and function, we developed an in vitro reductionistic\nmodel mimicking age-related reduction in RPE cell density…]\n• On the cellular hypertrophy:\nOn the same line regarding the validity of the ageing model, we are careful to\nconclude that the observations are directly transferable to RPE ageing. In fact, in\nthe discussion, we want to explain what is happening in our model, which will be\na stretch to compare directly with ageing RPE. However, we added a line in the\ndiscussion (lines 399 - 402) highlighting the significance for postmitotic epithelia\nhomeostasis and emphasised the findings in the results (lines 123-124).\n3) Related to point #1, the authors frequently use poorly defined terminology that\nobscures their interesting findings. Several examples require clarification:\n- What do the authors mean by \"a new mechanical equilibrium and homeostasis\" and\n\"actin-myosin network plasticity\" (line 368)?\n- How should readers interpret \"a shift in the range of mechanical equilibrium\"\n(line 431)?\n- What is meant by \"intercellular reinforcement\" (line 481)?\nWe thank again the reviewer for their valuable inputs regarding the text. We have\naddressed the raised points in the revised manuscript version as follows:\n• Regarding “mechanical equilibrium and homeostasis” and “actin-myosin\nnetwork plasticity”:\nWe clarify the conclusions with an expanded paragraph in lines 359-367\n• Regarding “a shift in the range of mechanical equilibrium”:\nIn the revised version, we introduce the concept of mechanical feedback loops\nand the interconnection with mechanical equilibrium and homeostasis early on\nin the introduction (lines 8-14), we highlight the change of mechanical state at the\nend of the corresponding result section (line 271-281), and we explain its\nimplications for phagocytosis in the discussion (lines 494-497).\n• Regarding “intercellular reinforcement\":\nThe sentence has been optimised for clarity, specifying that reinforcement is\nassociated with an increase in contractility at the cell-cell junction (lines: 475 -\n477).\n4) The results section is difficult to follow because while data are presented and\nquantifications are explained, there is insufficient interpretation of what the results mean\nand why these observations are important.\nWe thank the reviewer for this helpful feedback on the clarity of our exposition of the\nwork. Now we have added several more explanations throughout the results section.\nThe authors need to better explain the significance of their findings. Several examples\nillustrate this issue:\n- What do the nanoindentation data mean (lines 242-252)?\nWe added a short explanation in lines 219-221.\n- Why does the difference in Brillouin shift at different sections (apical to basal) lead to\nthe conclusion that \"the apical sides of the cells undergo the most pronounced\nreorganisation\" (lines 293-306)?\nMoreover, what specific type of 'reorganisation' are the authors referring to, and why is\nthis important?\nWe expanded the data explanation at the end of the results section in lines 254-258.\n- What is the significance of the segmented Brillouin shift data (lines 311-317)? Why\nshould readers care about \"the presence of loosely interdigitated basal membranes\"?\nWe improved the clarification of why we did the segmentation of the interstitium vs the\ncore, and the unexpected lower shift measured at the interstitium in lines 259-261 and\n267-270.\nWe thought mentioning the loosely interdigitated basal membrane may provide a likely\nexplanation for any questions readers may have regarding the data.\n- Why is the observation that \"age-associated RPE phenotype has a different\nbiomechanical state of the tissue\" important?\nWe expanded the explanation in a few points of the manuscript. In the results in lines\n277-281 and lines 359-367, and in the discussion in lines 458-465.\n- What does the finding that \"bulging cells showed a more irregular apical morphology\n(Fig. 5D)\" imply for RPE function? Why do the authors highlight this observation? A similar\nquestion applies to the statement that \"the treated AgeD-iRPE showed some degree of\n'bulging' during phagocytosis, which was more noticeable compared to the untreated\nAgeD-iRPE (Fig. 5H).\"\nThe rationale for investigating formins vs. Arp2/3 function in iRPE and dr-iRPE stems from\nthe apical F-actin topography observed during phagocytosis (Fig. 2h). We explain this in\nlines 287-295. With the expanded explanations in the results, lines 359-367, and in the\ndiscussion, lines 458-465, we believe this is now clearer.\nMinor Comments\n5) Fig. 6: Why does the middle panel showing top views highlight the observed\ncharacteristics only in the central cell and not in the neighboring cells as well?\nWe thank the reviewer for the observation and have now improved Figure 6 accordingly.\n6) Supp. Fig. 3A(2): The screenshot from an application is not appropriate for a scientific\npublication.\nWe have now substituted the low-resolution exported panel with a better-resolved graph.\n7) The statement that \"a balance between basal cell-ECM adhesion and apical cell-cell\nadhesion exists and is normally well conserved (78–80)\" (line 456) requires more detailed\ndiscussion.\nWe added some more detailed discussion in lines 441-446.\nReviewer #3 (Remarks to the Author)\nThis is a well executed study on the physiological changes of the retinal pigmented\nepithelial monolayer in response to 2% apoptosis in vitro. The quality of the experimental\ndesign, the chosen assays and the execution is all excellent. The framing of the\nexperiments is where this reviewer had much issue. There are flaws in the data\ninterpretation and conclusions which prohibit publication or require revision. They make\nbodacious claims as to what their results signify. To so without qualifications of which\nthere are many. They also use duplicitous naming of their model attempting to more\nclosely align what they are studying (epithelial monolayer response to apoptosis) to what\nthey claim to be studying (epithelial monolayer aging). Moreover there is no qualifications\nwith regards to the differences between in vivo and in vitro conditions nor the known\ndifferences between iPSC-RPE compared to native RPE.\nWe thank the reviewer for appreciating our technical execution and experimental design\nand for pointing out how we could refine the interpretation of the data without\noverstretching their significance. We have now significantly changed the manuscript as\nspecified below.\nImportantly, we replaced the abbreviation with a more appropriate one that focused on\nthe studied phenomenon. AgeD-RPE is now density-reduced iRPE or dr-iRPE.\nAdditionally, we now explain how iPSC-derived RPE are an accepted model for in vivo\nRPE in lines 58-59 and discussed the limitation of our in vitro system for comparison with\nthe in vivo native tissue in lines 415-418.\nThe experiments detailed in this manuscript does not support the title. “Tissue alone” is\ntoo vague for any understanding “alone” claim has not been supported by the evidence\nof this MS. The in vitro model is lacking many of the conditions in a live aging organism\nand therefore the conclusions can only be made in that model. Therefore, the title should\nqualify the statement. “within an in vitro model…\nThanks to the reviewer, we believe that we now have a much more precise and impactful\ntitle. The new title is: “Cell loss disrupts mechanical homeostasis to drive retinal pigment\nepithelium ageing-like phenotype in vitro”\nThe abstract is too general and does not discuss what is done, what results were found\nand what conclusions from those results can be made.\nAs also pointed out from reviewer #2, we now have rewritten the abstract.\nThe in vitro aging may exhibit similar morphological changes for very different reasons.\nLack of vascular support, lack of the ability to maintain ionic balance of the media\napically and basolaterally, no differential in nutrient access apically to basolaterally.\nThese over time can also lead to a decrease in physiological function and have nothing\nto do with the native environment.\nHydrogels do not have a composition resembling native RPE environment. Water\ncontent, diffusion gradient,\nWe agree with the reviewer that our model is highly reductionistic; therefore, conclusions\ncannot be drawn directly for a complex in vivo system. This was not sufficiently clear in\nthe previous version of the manuscript. We have now rewritten many parts of the\nmanuscript, clarifying that the conclusions stem from cell-density reduction in the\nmodel and not ageing. This is, for example, in lines: 409-410, 371-374, 471-473 or 494-\n495.\nFurthermore, the difference between our model and in vivo, including the hydrogel\nsubstrate, is clarified in the discussion in lines 415-418 and 447-450.\nThe induced apoptosis is too synthetic to model native RPE aging. Therefore what is being\nmodeled is the cellular response to apoptosis.\nWe agree with the reviewer and have clarified our conclusions accordingly. The revised\nmanuscript explicitly states that our in vitro model does not directly replicate ageing.\nHowever, as the model is inspired by the cell loss seen in RPE ageing in vivo, and because\nwe can reproduce key features of aged RPE, we believe it is reasonable to speculate that\nthe observed adaptation to apoptosis may highlight a previously unrecognised aspect of\nageing.\nThere is no age dependent condition and therefore, the way the experiments are\ndescribed is a misnomer.\nReplace AgeD RPE with apoptosis responsive ARD-RPE and then the manuscript is on a\nmore accurate foundation from which to draw conclusions. To mix apoptosis with age is\nduplicitous.\nAs mentioned earlier, the condition is now called density-reduced iRPE or dr-RPE,\ntherefore avoiding any bias of interpretation.\nThe lack of proliferation in response to apoptosis is an interesting response that warrants\nfurther study. What is missing is a spectrum of apoptosis responses from 0 to 50% at\nregular intervals. Is there an apoptosis % that proliferation can be triggered? If so, what\nare the mechanisms governing this? Is there another more natural apoptosis approach\nthat can support these findings or must it be the manipulation of caspase to lead to these\nchanges within this timeframe?\nThese are all relevant and interesting questions, which, we think, exceed the scope of the\ncurrent study and may remove the focus from our main message of a\nstructure/mechanics-driven functional decay. Furthermore, it is technically challenging\nwith the current method to finely modulate the apoptosis rate, and we made sure to\ninclude a control for the presence of the recombinant caspase (sup. fig. 2 and 3). Other\nmeans of cell death (e.g., staurosporin) have been initially explored with mixed success\ndue to off-target effects.\nRegarding the question of the proliferative response to varying levels of cell death, we\nnow highlight this as a relevant aspect for future investigation in the context of important\nretinal pathologies (lines 404-408).\nThe results are informative so long as the authors are more honest with their claims.\nThere is sufficient detail in the methods section to reproduce the experiments.\nCommend them on the proper protocol for assaying phagocytosis.\nIn general experimental execution and data collection quality is high.\nWe thank the reviewer for the positive assessment."}, {"round": 2, "reviewer_comments": {"reviewer_2": "In this revised manuscript, the authors have addressed most of the points I raised in my original review. I appreciate the\nauthors' thorough responses.\nOne minor comment relates to my original point #4: Why should readers care about 'the presence of loosely interdigitated basal membranes'? The authors responded that 'We thought mentioning the loosely interdigitated basal membrane may provide a likely explanation for any questions readers may have regarding the data.' However, without context, readers (including myself) remain confused about what this observation means. To clarify this point, I suggest providing at least one specific example of a question that readers might have about your data that would be answered by this basal membrane observation and explain how this structural feature addresses that question in the text. This will help readers understand the relevance of this finding to your overall results. Since this observation is featured in both the text and figures, readers need a clear explanation of its relevance.\nOnce this issue is addressed, I support this manuscript for publication.", "reviewer_3": "The authors satisfied all of the major issues in my previous review. I support acceptance."}, "author_response": "We thank the reviewer for appreciating our efforts in providing an exhaustive revised\nmanuscript. Regarding the comment about the remaining confusion on the interdigitated\nbasal membrane, we apologise if this remained unclear. We have now explicitly added a\nsentence explaining the significance of this observation for the overall interpretation of\nthe data. This has been included in lines 274–277."}]

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