Differential cellulose distribution drives polarized growth of cotton fibers

Domain: biology
Published: 2026-04-02
Editorial outcome: accept

Abstract
Plant cell morphogenesis relies on the mechanical properties of the primary cell wall, yet it remains unclear which components predominantly regulate wall extensibility. Cotton fibers, highly elongated single cells, offer a unique system to investigate polarized cell expansion. Here, by combining atomic force microscopy and cellulose labeling, we find a basipetal gradient of cellulose microfibril density from the apex to the shank that underlies cell wall heterogeneity and directed cotton fiber elongation. Live-cell imaging shows that cellulose synthase complexes accumulate more densely toward the shank which is guided by specific microtubule organization and is supported by genetic disruption and microtubule perturbation. A mechanical model further demonstrates that a cellulose gradient is sufficient to reshape axial strain for directional growth. Collectively, our findings provide single-cell evidence for a cellulose-dependent mechanism of directional growth, expanding our understanding of primary cell wall extensibility in plant morphogenesis and offering potential strategies to improve cotton fiber quality.

Introduction
Plant primary cell walls (PCWs), mainly composed of three distinct polysaccharides (cellulose, hemicelluloses and pectins), are strong but extensible extracellular structures that critically underpin cell growth and environmental adaptation 1,2. From a biomechanical perspective, plant cell expansion is driven by the prevalent turgor pressure within the cells. And the heterogeneity of plant cell walls, including the spatial distribution of cell wall components and their structural organizations, results in anisotropic mechanical responses to turgor pressure. Such behavior would convert the isotropic internal pressure into directionally biased deformation, contributing to the formation of diverse cell shapes. Therefore, the mechanical properties that determine dynamic extensibility of PCW are central to plant cell growth and morphogenesis 3,4. However, how different cell wall components determine anisotropic expansion of plant cells remain controversial. Homogalacturonans (HGs), the most abundant form of pectins, are often considered as major determinants of plant cell wall heterogeneity 5-7. However, new computational models support a prominent role of sliding cellulose microfibrils in cell wall extensibility during constant stress 3. Due to substantial differences in cell types and contexts that these observations have been made, we argue that a single-cell system that combines diffuse and tip-related growth might be ideal to address cell wall organization and the driving force for plant cell growth.

Cotton fibers are highly elongated and unbranched single cells differentiated from the ovule epidermis, and are one of the longest plant cells characterized 8. Elongating fiber cell walls typically contain 40% cellulose, 25-30% hemicelluloses, 30-35% pectins, and a variety of wall proteins 9. Notably, the length of cotton fibers can reach 3-6 cm, which is 1000-3000 times greater than the diameter during the elongating stage 10. After the PCW-dominated rapid elongation stage, cotton fibers gradually stop elongating and enter a secondary cell wall thickening stage. In this phase, cellulose accumulates to become the most prevalent wall components, accounting for > 95% dry weight of cotton fibers 11. Owing to the highly polarized unicellular feature and abundant cellulose content, cotton fibers have long been regarded as an excellent model for investigating plant cell elongation and cellulose biosynthesis 12 . Elongating cotton fiber cells may therefore be a suitable single-cell system to decipher how different cell wall components contribute to heterogenous wall organization and dynamic extensibility during plant cell growth 9.

Qin and Zhu proposed a linear growth mode of cotton fibers that combines tip and diffuse growth, based on cytoskeletal organization using immunostaining 13. Ten years later, our previous studies provided the first live-cell observations on the cytoskeletal organization in elongating fiber cells, and further proposed a unique, tip-biased diffuse growth mode for cotton fibers 14,15. However, direct exploration of wall structures and mechanics that drive cotton fiber elongation is still lacking. Here, using atomic force microscopy (AFM) alongside cellulose labeling, we show that differential cellulose deposition underpins the mechanical properties of cotton fiber cell walls. By integrating live-cell imaging with computational modeling, we propose that a basipetal cellulose gradient, tightly associated with the microtubule-mediated spatial distribution of cellulose synthase complexes (CSCs), drives cotton fiber elongation. This unique cellulose deposition pattern not only supports tip-biased diffuse growth in cotton fibers but further highlights the indispensable role of cellulose in regulating plant cell expansion at the single-cell level.

Results
Cotton fiber cell walls are soft at the tip, with stiff shanks
After anthesis, cotton fibers initiate and proceed into a rapid elongating stage (Fig.1a). This stage relies on the heterogeneous extensibility of the PCWs. To assess mechanical characteristics, as well as morphological surface differences along the PCWs, we examined the apparent elastic modulus and apparent stiffness via AFM scanning (Fig. 1b and c). Cell wall elastic modulus describes the intrinsic elasticity of the cell wall material, while cell wall stiffness reflects the overall resistance of the wall to deformation under an applied force. In our AFM measurements, the reported modulus is best interpreted as an apparent elastic modulus, as the fiber walls show adhesion and nonlinear force-displacement behavior. Although AFM indentation primarily measures the mechanical response perpendicular to the cell wall surface, it provides valuable insight into the local mechanical properties that may influence wall extensibility and morphogenesis in growing tissues 16-19. Therefore, in this study AFM-derived apparent modulus and stiffness are used here as comparative indicators of the local mechanical state, which could be interpreted as a relative mechanical landscape and is informative for understanding cotton fiber elongation. We found a gradual increase in apparent modulus of cotton fiber cell walls at 7 DPA (corresponding to rapid elongation) as compared to 3 DPA (early elongating stages/late initiating stage), which may underlie an enhanced capacity to bear turgor-generated stress during fiber expansion (Fig. 1c and Supplementary Fig.1). By comparing AFM-derived mechanical parameters across two major cotton cultivars, i.e. the upland cotton (Gossypium hirsutum) TM-1 and the sea-island cotton (Gossypium barbadense) Hai7124 harboring longer fibers, we found that Hai7124 exhibited lower apparent elastic modulus and stiffness than TM-1 (Fig. 1c). This association between a more mechanically compliant primary wall and greater final fiber length suggests a possible link between early wall mechanics and fiber elongation capacity. If confirmed in broader cotton populations, such features could potentially serve as mechanical indicator for fiber-quality breeding.

We next analyzed mechanical properties at different regions of individual fiber cells. Strikingly, the cell walls near the tip exhibited lower apparent modulus and stiffness, suggesting that the PCWs of cotton fibers are softer and mechanically more compliant at the tip that we interpret as conferring a higher local expansion potential (Fig. 1d), most likely related to fiber tip-biased diffuse growth mode. To further test whether this mechanical heterogeneity correlates with regional differences in wall deformation, we performed osmotic shrinkage assays. Treatment of 3 DPA fibers with 0.8 M mannitol caused a significantly larger reduction in width at the tip than at the shank, whereas fibers under water conditions showed minimal changes (Fig. 1e and f). These results indicate that apical walls are more deformable under turgor-related perturbation, in agreement with their lower apparent stiffness and modulus.

To further confirm the tip-biased elongation of fiber cells, we attempted to visualize newly forming cell walls in cotton fibers from the cultured ovules via a click-chemistry-based strategy 20. We used a glucose that contains a clickable group (2-Azido-2-deoxy-D-glucose, aGlu), which was applied in the growth medium. This molecule was readily incorporated into the fiber cell walls (Fig. 2a and b), thus reflecting newly formed cell walls that we could track by a click reaction to the dibenzocyclooctyl-linked cyanin 5 (DBCO-Cy5) (Supplementary Fig. 2a-c). While we are unsure what cell wall polymer that is labelled by this approach, we could therefore monitor the incorporation of glucose into the extracellular matrix (Supplementary Fig. 2d). These analyses confirmed a tip-biased growth pattern of cotton fibers (Fig. 2c and Supplementary Fig. 2a-c).

A gradient of cellulose accumulation determines heterogeneous cell wall properties of cotton fibers
Local modifications of pectins play a crucial role in cell wall extensibility 6,7,21,22. To see whether the ratio of de-methyl- / methyl-esterification of pectins in cotton fibers could explain the cell wall heterogeneity we observed in our AFM assays, we performed immunolabelling assays with two monoclonal antibodies, JIM5 and JIM7, that specifically recognize the lowly-esterified HGs (JIM5) and highly methyl-esterified HGs (JIM7), respectively 21. Notably, both JIM5- and JIM7-labeling were uniformly distributed along cotton fibers (Supplementary Fig. 3a and b). To further investigate the role of pectins in fiber extension, we treated the fibers with the pectinase. However, no obvious impact was detected on cotton fiber growth and development (Supplementary Fig. 3c). We therefore inferred that pectin modifications if involved, are unlikely to have major effects on cotton fiber development or on the heterogeneous cell wall formation that we observed.

Cellulose is the principal load-bearing cell wall component, and we therefore speculated that differential distribution of cellulose may contribute to the heterogeneity of cotton fiber PCWs 9,23. To test this hypothesis, we assessed the thickness of cellulosic cell walls of cotton fibers between the apex and shanks and performed functional perturbations. Here, we used calcofluor-white (CFW) that preferentially binds cellulose in cell walls of land plants and quantified the full width at half maximum (FWHM) of the cross-sectional CFW intensity profile as an indicator of the apparent width of cellulosic PCWs (Fig. 3a and b). These results revealed a clear difference of CFW-derived signal width between fiber shank and apex, indicating higher cellulose accumulation in the shank than in the apex. In parallel, staining with the cellulose-specific dye S4B, we found transverse banded labeling patterns that are in line with previous observation of cellulose microfibrils by transmission electron microscopy24, and likewise showed stronger cellulose accumulation in the fiber shanks (Fig. 3c and d). Interestingly, we also found that within apex cross-sections, the CFW-derived apparent wall signal width positively correlated with fiber diameter (Fig. 3e). Together with the differential cellulose accumulation between fiber shank and apex, these data infer a progressive increase in cellulose deposition along the fiber axis, which may contribute to heterogeneous wall mechanics and underpin anisotropic expansion of the fibers. To functionally test whether cellulose deposition is required for fiber growth and cell wall heterogeneity, we used either the cellulose biosynthesis inhibitor (CBI) isoxaben25 or cellulase R-10 to reduce cellulose deposition in the fibers at 2 DPA. Both treatments led to loss of the anisotropic growth behavior of the cotton fibers, indicating that cellulose deposition is indispensable for cotton fiber development (Supplementary Fig. 4). Taken together, these results support differential cellulose deposition along the fiber axis, which is consistent with and may contribute to the observed cell wall heterogeneity and directed cotton fiber elongation.

The CMF deposition gradient is associated with differential CSC distribution across the cotton fiber surface
PCW cellulose is presumed to be synthesized by GhCESA1, GhCESA3 and GhCESA6 in cotton fiber cells 26,27. To directly observe the cellulose biosynthesis and the deposition of CMFs in elongating cotton fibers, we generated mNeoGreen-tagged GhCESA3 single, or mNeoGreen-CESA3 and mCherry-TUB9 dual, fluorescent markers that label CSCs and microtubules, respectively. Using these lines, we performed live-cell imaging to track the spatial distribution and dynamics of CSCs. As shown in Supplementary Fig. 5, we observed PM-localized mNeoGreen-CESA3 particles in various cell types, such as pavement and petiole cells. These particles moved along linear trajectories, similar to what has been reported in Arabidopsis, with movement that coincided with underlying MTs (Fig. 4a). In addition, we also observed CESA3 fluorescence from cytoplasmic foci that moved erratically, and likely resemble that of small CSC compartments (SmaCCs) or MT-associated CESA compartments (MASCs) that have been reported on in Arabidopsis 28,29.

To further explore the CSC dynamics in elongating cotton fibers, we imaged both initiating and elongating fibers. Similar to other cell types, fluorescent CESA3 particles localized to the PM in fiber cells across these developmental stages (Fig. 4b and Supplementary Fig. 5a-d). The speed of the PM-located CSCs is thought to correspond to the catalytic activity of the CSCs 29. Therefore, we quantified the speed of the PM-localized CSCs across various cotton cell types. Surprisingly, while the CSC moved with about 150 nm/min in pavement (mean ± SD = 148.3 ± 63.6, five cells) and petiole cells (mean ± SD = 147.1 ± 68.6, five cells), the CSCs moved significantly faster in the two developmental stages (initiation, mean ± SD = 224.5 ± 84.2, five cells; elongation, mean ± SD = 365.2 ± 110.4, five cells ) of cotton fibers (Fig. 4c and d). Taken together, these data indicate a higher catalytic activity of CSCs in cotton fiber cells than in pavement cells or petiole cells, likely due to high demands of cellulose in the rapidly expanding fiber cells.

As the shanks and the apex regions of cotton fiber cells exhibit heterogeneous cell wall mechanics and differences in cellulose levels, we speculated that the catalytic activity of CSCs might vary correspondingly. To test this, we imaged mNeoGreen-CESA3 at the different regions of elongating fibers. However, only mild changes in CSC speed at the different regions were detected (Fig. 4e and f, and Supplementary Movie 1). Alternatively, the density of PM-localized mNeoGreen-CESA3 particles appeared to be higher in fiber shanks than at the apex (Fig. 4e and g, Supplementary Fig 5e, and Supplementary Movie 2). We confirmed this observation by quantifying the number of fluorescent CSCs per area unit, where shanks had 0.84 CSCs per μm2 and apex had only 0.59 CSCs per μm2. Therefore, the increase in cellulosic PCWs along the cotton fiber is likely driven by a biased PM-localized CSC distribution.

The microtubule arrangement defines the differential CSC distribution for cotton fiber elongation
CSCs typically track along MTs, which may define CSC trajectories in plant cells 29,30. To corroborate that such relationship also determines the spatial patterning of CSCs in cotton fibers, we examined the cortical MT organization in cotton fiber cells at both the initiation and the elongation stages (Fig 5a, Supplementary Fig. 5f, and Supplementary Movie 3, 4). We observed a mesh-like MT array in fibers at the initiation stage. By contrast, the MT array was well-aligned and arranged perpendicular to the growth axis of elongating fiber cells. Correspondingly, we observed fluorescent CSCs moving along the MTs, with trajectories aligning with the MTs (Fig. 4c). We next assessed the MT organization in the shanks and the apex regions of elongating cotton fiber cells. As expected, we observed reduced MT density at the apex as compared to the shanks, which corresponded well with the observed CSC distribution (Fig. 5a-c). Additionally, we detected a MT depleted zone (MDZ) in the fiber cell apex, similar to our previous observations in cotton fibers and in Arabidopsis trichomes 14,31. Then, we further validated the functional relationship between MTs and CSCs by pharmacological treatments in cotton fiber cells. Here, the MT stabilizing drug Taxol or MT nucleation inhibiting drug Propyzamide reduced the differences in MT density between the shanks and the apex of fiber cells (Fig. 5d and e). Surprisingly, these treatments led to an even distribution of CSCs across the fiber cell PM (Supplementary Fig. 5g and h). These fibers did not display tip-biased fiber elongation as illustrated by the chase-labelling experiments (Fig. 5f).

The CSCs are guided by MTs via the protein CSI1 in Arabidopsis 30,32. Thus, we first validated the biochemical interaction between GhCSI1 and GhCESA3 (Supplementary Fig. 6a and b). To further explore the relationship of CSC and MTs in cotton, we used the CRISPR-Cas9 technology to mutate the GhCSI1 paralogs (Gh_A11G212800, Gh_D11G239500) (Supplementary Fig. 6c), which are predominantly expressed in development fiber cells, of both the At and Dt cotton subgenomes as reported in CottonGVD database (https://db.cngb.org/cottonGVD/). Using advanced fiber information system (AFIS) measurements of cotton fiber lengths, we found that the CSI1-mutated lines had significantly shorter fibers compared to wild type (Supplementary Fig. 6d-f). These results indicate an important role of MT-guided CSC distribution to control tip-biased growth of cotton fibers.

Computational simulation of how CMF deposition patterns underpin cotton fiber growth
To explore how CMFs support both directional expansion and differential growth of cotton fibers, we performed computational simulations. Some of the key modeling parameters are referenced from a previously reported cotton fiber model 33 (see also Supplementary Note 1). In our model the cell wall matrix and embedded CMFs are simplified to longitudinally arranged matrix spring segments and numerous transversely arranged CMF springs (Supplementary Fig. 7a), similar to what we observed with the S4B staining. This method allowed us to simulate and analyze the effects of different CMF deposition patterns on the polarized elongation of cotton fiber cells. According to our simulations, the uniformly distributed CMF springs along the cells led to equivalent strains along cell strains along cell walls, which correlated with a diffuse expansion mode of plant cells (Supplementary Fig. 7b and c). By contrast, a gradient of CMF springs led to a gradually increasing strain at the apex (Fig. 5g and h, and Supplementary Movie 5), which is in line with our observations in cotton fiber cells.

Additionally, we noticed that the growth rate of the apex would reach saturation with the increase in shank-to-apex CMF density ratio in the model (Supplementary Fig. 7d). Similarly, at higher shank-to-apex CMF density ratios (e.g., greater than 6), the reduction in applied force will be less pronounced. (Supplementary Fig. 7e). These results indicated that maintaining a range of shank-to-apex CMF density ratio (i.e. 2 to 6) is sufficient to generate tip-biased cotton fiber elongation. Excessive CMF deposition differences between cotton fiber apexes and shanks would perhaps not be an efficient way to drive cell expansion as this may lead to unnecessary amounts of cellulose at the shank. Our modelling data thus supports our live-cell imaging, where we observed a two-fold shank-to-apex ratio of CMF density. Collectively, our simulation results, together with the observed cellulose and mechanical heterogeneity, are consistent with a tip-biased diffuse growth mode of cotton fibers, in which apical regions exhibit higher elongation potential than the shank (Fig. 5i). In this model, elongation remains distributed along the fiber but is biased towards the apex.

Methods
Plant materials and growth conditions
The cotton cultivars TM-1 and Hai7124 were used as the wild type control for the upland cotton (Gossypium hirsutum) and the sea-island cotton (Gossypium barbadense), respectively. The TM-1 was also used as recipient of the proCESA3-mNeoGreen-CESA3 and the proCESA3-mNeoGreen-CESA3-proTUB1-mCherry-TUB9 constructs. Cotton transformation was performed as described previously 14. All the cotton lines were grown in the field under normal farming practices, or in a greenhouse with long-day (16 h light/8 h dark) conditions at 28 ℃. Flowers were tagged on the day of anthesis, and fibers were dissected from ovules in bolls at various DPA for further analyses. Bolls from similar positions (primarily the middle boll positions) were uniformly harvested for fiber analyses.

Plasmids Construction
The constructs of proCESA3-mNeoGreen-CESA3 and proCESA3-mNeoGreen-CESA3-proTUB1-mCherry-TUB9 were generated to obtain stable transgenic cotton lines expressing the fluorescent marker of the mNeoGreen-tagged CESA3, or cotton lines simultaneously expressing the double fluorescent marks of mNeoGreenCESA3 and mCherry-TUB9. The Phanta DNA polymerase with high fidelity (Vazyme Biotech Co., Ltd, AWS1207) was used to amplify all the required gene products as described in the previous studies 55,56. In detail, for the construct of proCESA3-mNeoGreen-CESA3, the promoter (1,930 bp upstream from the initiation ATG) of CESA3 gene (Gh_A08G167200), the coding sequence of CESA3 genes and the 1,125 bp downstream untranslated region was amplified from the genomic DNA, respectively. And these fragments fused with the fragment of mNeoGreen were used to obtain the final expression constructs through the in-fusion cloning using the DNA Assembly Mix PLUS KIT (LABLEAD, D0204P). For the construct of proCESA3-mNeoGreen-CESA3-proTUB1-mCherry-TUB9, the promoter (1,608 bp upstream from the initiation ATG) of TUB1 gene (Gh_A08G207900), the coding sequence of TUB9 gene (Gh_A08G20660N0) and the 931bp downstream untranslated region was amplified from the genomic DNA, respectively. These fragments fused with the DNA fragment encoding mCherry were insert to the construct of proCESA3-mNeoGreen-CESA3 through the in-fusion cloning using the DNA Assembly Mix PLUS KIT (LABLEAD, D02A04P) to obtain the final expression construct.

Atomic Force Microscopy
Ovules at different developmental stages were collected, and fiber bundles were gently cut from the ovules and mounted on clean glass slides, then allowed to air-dry at room temperature. Atomic force microscopy (AFM) imaging and nano-indentation were performed using a Dimension Icon AFM (Bruker, Santa Barbara, CA, USA) operating in PeakForce Tapping mode with ScanAsyst and Quantitative Nanomechanical Property Mapping (QNM) as described previously 57. All images were acquired at 512 × 512 pixels over 2 µm × 2 µm scan areas positioned on visually flat regions of the fiber surface near the tip or the shank.

ScanAsyst-Air silicon-nitride probes (Bruker; nominal spring constant 0.2-0.7 N/m; tip radius ~10 nm) were used. The actual spring constant was calibrated by the thermal-tune method and the deflection sensitivity was determined on a hard glass surface using the NanoScope software. During PeakForce imaging, the force setpoint was adjusted to keep the maximum load below ~5 nN, which resulted in an indentation depths on the range of ~100 nm. At every pixel a force curve was recorded and later analyzed in NanoScope Analysis 1.8 (Bruker). Regions of interest selected on relatively smooth areas of the fiber surface were used for statistical analysis. Modulus and stiffness are derived from force maps, with a built-in Hertz contact model with adhesion enabled, as implemented in the software. Because the fiber cell walls exhibited pronounced adhesive minima and nonlinear force-displacement behavior, the AFM-derived moduli should be regarded as apparent elastic modulus. Pixels affected by edges, dust, obvious baseline drift or poor fits were excluded from the analysis.

Ovule culture in vitro culture and chase-labelling of newly formed cell walls
The fresh flowers at 2 DPA were harvested from the cotton plants. The ovaries were then sterilized with 70% ethanol after removing petals and the sepals. The cotton ovules (2 DPA) were disserted from the bolls and cultured at 28 °C in BT medium in dark. To access the influence of cellulose for cotton fiber development and morphology, either 0.1 μM of Isoxaben (DR. EHRENSTORFER, DRE-C14480000) or the 0.1% (w/v) of cellulase R-10 (Yakult, L0112) was applied into the medium. The growth of cultured cotton fibers were observed either by a stereoscope or by a confocal microscope.

To determine the newly formed cell walls in cultured cotton fibers, 2.5 μM of 2-Azido-2-deoxy-D-glucose (aGlu; HARVEYBIO, A33673) were applied or washed out to the medium in two stages of 4-hour periods. The ovules were washed three times with PBS (pH=7.4) and then labelled with 1 μM DBCO-Cy5 (AAT Bioquest, Lot: 923) for 20 min. Fluorescence of labeled cotton fibers was visualized via a confocal microscope (Leica SP8, excited at 633 nm, EM 650–670 nm) under a 40X objective.

Immunolabelling of pectins in cotton fiber cells
The cotton ovules were harvested from fresh flowers at 2-3 DPA and then fixed for 30 min with 4% (w/v) paraformaldehyde in PBS buffer (pH=7). After three times washes with PBS, the ovules were then incubated with 3% (w/v) milk protein in PBS buffer to block non-specific binding sites. The primary antibodies (JIM5 or JIM7; PlantProbe) were diluted in 1:10 with PBS and incubated with the ovules overnight at 4 ℃. After three washes with PBS, the ovules were then incubated with the secondary antibody (FITC conjugated goat anti-rat IgG; ABclonal, AS019) diluted in 1:100 with PBS for 1 h in dark at room temperature. Fibers were carefully cut off the ovules and placed onto Poly-Prep slides (Sigma-Aldrich) after three washes with PBS and examined via confocal microscope excited at 488 nm.

To access the influence of pectins for cotton fiber development and morphology, fresh cotton ovules at 2 DPA were harvested and cultured in at 28 °C in BT medium in dark. 0.1% (w/v) of pectinase (Biotopped, P6280C) was applied into the medium. After 12h incubation, the pectinase treated ovules were performed immunolabelling with JIM5 to test the degradation of pectins.

Assessing the thickness of cellulose in cotton fiber cell walls
The cotton ovules were harvested from fresh flowers at 10 DPA. and then fixed overnight in 4 % (w/v) paraformaldehyde in PBS buffer (pH=7) at 4 °C. Then the ovules were dehydrated in ethanol and embedded in Technovit 7100. The cotton fibers were cut into sections (thickness, 5 μm) with a Leica RM2265 microtome, and stained with 0.05 % (w/v) Calcofluor White (CFW; Sigma-Aldrich, BCCNF8014) for 5 min at room temperature in dark. The stained fiber cross sections were examined under 40X objective of the ZEISS Axio observer Z1 at excitation of 445 nm. The fluorescent of CFW intensity profile was measured using Line Tools of ImageJ (https://imagej.nih.gov/ij/). The FWHM was calculated through a self-written macro after the Gaussian fitting.

Scanning Electron Microscopy and S4B staining
Cotton ovules at various developmental stages (0 DPA, 3 DPA and 7 DPA) were harvested from the fresh flowers, and frozen in the liquid nitrogen via frozen transmission system (Quorum PP3010T). The samples were broken and sublimated in -70 °C for 10 min. After sputtering with platinum (10 mA, 60 s), the imaging was performed in a high-resolution SEM 7401F (JEOL, Japan).

To display the cellulose deposition by S4B staining, the ovules were collected from fresh flowers at 3 DPA and incubated with 0.1% (v/v) Tween-20 in PBS buffer (pH=7) for 5 min. Then the samples were stained with 0.01% (w/v) S4B in PBS buffer for 15 min in dark. After three times washes, the fiber cells were cut off the ovules and placed onto slides. The fluorescent signals of stained celluloses were examined under 60X oil objective of confocal microscope with excitation at 561 nm.

CRISPR/Cas9-mediated genome editing
The CRISPR/Cas9-mediated genome editing was implied to knockout the GhCSI1 genes in both At (Gh_A11G212800) and Dt (Gh_D11G239500) of cotton genome. Dual guide RNA (sgRNA) 5’-ATCAGGATCTCTTGGAGTGA-3’ and 5’-GTGCTTTGGAAGCTGCTAAA-3’ was designed from both of the coding region. CRISPR-Cas9 experiments were conducted following previously reported experimental protocols 58 and performed by WIMI Biotechnology with TM-1. The edited type of GhCSI1-A and GhCSI1-D were identified via Sanger sequencing (BGI Tech Solutions Co.) to obtain homozygous gene mutants with specific primers. All the cotton lines were grown in the field under normal farming practices, or in a greenhouse with long-day (16 h light/8 h dark) conditions at 28 ℃. Mature fibers (~10 g) were collected from the bolls of similar positions (primarily the middle boll positions) to determine the fiber length parameters by Advanced Fiber Information System (AFIS).

Spinning-disc confocal microscopy and live-cell imaging
Live-cell imaging was carried out under a spinning-disk confocal microscope (Ultraview VoX, PerkinElmer, Beaconsfield, Buckinghamshire, UK) equipped with the Yokogawa Nipkow CSU-X1 spinning disk scanner, Hamamatsu EMCCD 9100-13, Nikon TiE inverted microscopy with the Perfect Focus System. mNeoGreen was excited at 488 nm, and mCherry was excited at 561 nm. A 100X Plan Apo oil immersion TIRF objective (NA=1.49) was used for most image sequences acquisition. Each time-lapsed image stack was acquired over five minutes with 30s intervals. Acquired images were processed and analysed using Volocity (Perkin Elmer), ImageJ (http://rsbweb.nih.gov/ij), as described previously 29,59.

Computational simulation
The detailed description of cotton fiber growth model is available in Supplementary Note 1.

Quantification and statistical analysis
All statistical details (statistical test used, number of samples, and p values) for each experiment can be found in the Figure labels or legends. The statistical analyses were performed using GraphPad Prism 9 software. Data were considered statistically significant when p < 0.05 by statistical test performed as indicated in the Figure labels or legends. Two-tailed unpaired Student’s t test was used to compare two different groups of samples when indicated.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_2": "The paper investigates the mechanism of polarized growth in cotton fiber cells, focusing on how the differential distribution of cellulose microfibrils (CMFs) drives the heterogeneity of the cell wall and directional expansion. The study found that the cellulose gradient is a key factor driving polarized growth, while pectin modification has a minimal impact on fiber elongation. These findings deepen the understanding of the mechanisms underlying polarized growth in plant cells, and provide new insights for improving crop fiber quality.\nAll my concerns have been addressed in this revision.\nThere are some minor comments need to be revised:", "reviewer_4": "The authors examine the biomechanics of cotton fibres in the context of cotton fibre morphogenesis. I like the idea of using the cotton fibre to look at growth, especially because appears to be a blend of tip and diffuse growth, and has a relatively simple and highly anisotropic shape.\nThe authors have shown there is a gradient of cellulose, which they suggest translates into a gradient of stiffness that in turn results in growth gradient driving elongation. Unfortunately, they have no quantification showing the presence of growth gradients, and their method for measuring stiffness is not really suitable for the diffuse growing section, as it can only measure stiffness normal to the surface.\nWithout any growth measurements and stiffness measurements limited to the normal direction, it is difficult to be convinced of their main hypothesis.\nGeneral comments\nI notice not much data is presented showing cotton fibres do indeed display both tip and diffuse growth, it seems to be just taken as given. It would be nice if the authors could point to some growth tracking data that does indeed show this. The authors wrote a Current Opinion in 2011 on this subject, but did not present any tracking data there either. Does this mean it is not possible to track growth in cotton fibres? like people have done for trichomes and pollen tubes? In any case, if this is the state of the art, then statements of how they grow need to be presented more carefully as hypotheses, not fact. In addition, it would be good to know what the relative percentages are believed to be. Is it mostly at the tip? the flank? 50-50?\nIt is great that you have been able to track the cellulose synthase in this system with live imaging, so is there a reason you cannot track the growth as well? These things grow pretty fast, so one does not need to keep it happy that long.\nMost of the results in relation to mechanical properties of the wall depend on AFM measurements. However, I am not sure that this technique is so useful to determine stiffness in diffuse growing areas, at least not in a way that is relevant for morphogenesis. In tip growing cells the wall at the apex has no cellulose, and thus growth there is mostly isotropic. In this case the AFM can likely be used to measure the stiffness that could be relevant for growth. In the diffuse part the wall is anisotropic because of the cellulose. Since the AFM can only measure in the direction normal to the surface, it cannot capture the directions that are important for growth. What is needed is an in-plane measurement of the stiffness, and in this case the longitudinal direction would be most important.\nOne could use potentially use the AFM to “see” cellulose bundles, as was done in Sampathkumar et al. 2014 using similar equipment. In that work the images look pretty convincing. Here the images appear very different, and I am not so convinced that these images visualize cellulose fibres. In order to show they do, the authors would need to show they are not just showing for irregularities in the surface shape, for example wrinkling from being plasmolysed. If a smooth patch of surface is found, with stiffer and softer sections, then this could indicate the presence of cellulose bundles. So it would be good to see both the surface topography and the stiffness map side by side.\nIn any case, the cellulose in this system is highly transversely oriented, so we have no way of knowing if more cellulose means more stiffness in the longitudinal direction. To access this, one would need a different method, such at stretching with an extensometer, or osmotic treatments. The system is too small for an extensometer, but what about osmotic treatments? One should be able to image the fibre, plasmolyse it, and then image again and observe the shrinkage. One could put florescent beads to make landmarks to track the deformation, again as has been done in trichomes and pollen tubes.\nAnother related issue throughout is the authors are stressing the importance of a cellulose gradient as a main driver for shape and morphogenesis in the cotton fiber. Since the cellulose is predominately transverse, would that not imply it primarily stop the cells from widening? And how is it that they don’t get too wide towards the tip, where there is much less cellulose? especially since the stress is higher in the transverse direction. One has to wonder if this system mimics a bit pollen tube growth. In pollen tubes there is pure tip growth, but the tip has no cellulose, it appears displaced from the tip later on, mostly in a transverse pattern as here. Thus in pollen tubes, you get a very elongated shape from isotropic growth, that it limited to a small area at the tip. Cellulose in pollen tubes does not appear to be involved in creating the shape, but only appears in areas after the growth has stopped. The more tapered shape in cotton fibres that enters the region with cellulose suggests that diffuse growth is required, and that it cannot be solely tip growth\nThe authors frequently conflate the concepts of elasticity and extensibility. They are not the same, and just because a wall is more elastic (softer) does not mean it is more extensible. A wall needs to be elastic (have stretch) and have extensibility factors (ie expansins, etc) in order to grow. Without the latter the wall will not grow. It could be that there is a gradient of extensibility factors, rather than differences in elasticity, that are the primary drivers of growth gradients. Although expansin appears as the predominant extensibility factor in cell wall with cellulose, it is not clear what the extensibility actors are at the tip where growth happens without cellulose, since the usual pectin suspects have been apparently ruled out.\nNot heard of the Turkey test (brings strange images to mind), perhaps the authors mean Tukey test?\nSpecific comments:\nLine 40: GhCSI1 is mentioned in the abstract without saying what it is.\nLine 52: What about callose?\nLine 58: Technically speaking, CMFs create anisotropy in the wall that governs elastic behaviour, how the wall extends to make shape is a different matter, although they are usually, but not always, correlated.\nLine 76: Deposition stops, but secondary cell wall is produced? Is that not still cell wall (ie cellulose) deposition?\nLine 110: It is not clear what the authors mean by modulus and stiffness here. The authors only talk about stiffness in the methods. The fibres were dried, so I don’t understand how they measure modulus if they say it relates turgor pressure. If modulus is being calculated from the stiffness with some sort of Herzian contact model, I am not sure why they should be so qualitatively different. This section needs substantial clean up, and needs to be made much more precise.\nThe max force for the AFM and indentation depth should also be reported.\nLine 124: Again, you are conflating elasticity and extensibility. (also Lines 130,\nLine 312: What existing paradigms about cell wall extensibility are being challenged?\nLine 339: Do you really know what the gradient of extensibility looks like? If you have now time-lapse tracking, how do you know there is a gradient of growth. It could bethe tip grows fast, and the whole shank grows more slowly, but the same. I don’t think you can say this with any certainty.\nLine 342: It is not known that microtubules respond to mechanical cues, it is a hypothesis, which so far has little direct experimental support.\nLine 375: The idea that cellulose is the most important player for elongation, and that pectin is not important, is in obvious contrast with rapid growth at the tip. Again this stresses the necessity to have growth data in this system to support this hypothesis.", "reviewer_5": "The key messages of the study is a basipetal gradient of cellulose microfibril density from the apex to the shank. Apical regions are reported to be softer, while the shank is stiffer to support overall cell integrity. It make sense and is verified by computational simulation that CMF distribution influences strain patterns along the fibers. Primary wall during cotton fiber elongation is reported with 22% semi-crystalline cellulose fibrils, which are surrounded by matrix polymers including xyloglucan and pectin. So far, there has been a lot of evidence that presence and modification of the matrix polymer may regulate fiber elongation.\nIn this study, the focus is solely on cellulose accumulation and the main evidence by Raman spectroscopy lacks integrity of data and spectral interpretation. Only 3 spectra are shown from 10DPA and analysis is only based on the 380cm-1 band, which is taken and interpreted as cellulose accumulation. Raman spectra of cellulose reflect also changes in cellulose orientation and crystallinity, which is not analysed and taken into account at all. Further, pectin and hemicellulose bands should not be overseen and discussed. Unfortunately only 10DPA is analysed with Raman, while data from AFM are from 3DPA and 7DPA and staining from 3DPA and 2DPA, cell wall synthesis 3DPA. The study would benefit from a more conclusive experimental design including Raman at different developmental stages (same as AFM) and mutants added. Additional the study will benefit from a more comprehensive Raman spectra interpretation, instead of plotting the intensity of one peak for cellulose content. That can only be done, if changes in orientation and crystallinity can be ruled out based on the spectral signatures.\nIn the Material and Method part of Raman experiments no information is given, but only the link to a paper which has nothing to do with Raman: 51) Liu, W. et al. Towards a better recording of microtubule cytoskeletal spatial organization and dynamics in plant cells. J Integr Plant Biol 61, 388-393 (2019). https://doi.org/10.1111/jipb.12721.\nPlease provide all necessary information or at least correct citation. And it would be interesting for the reader to comment why laser tweezers Raman microspectroscopy was chosen and not conventional Raman imaging using a Confocal Raman microscopy, which might give more detailed information including spatial changes in content and orientation.\nSome questions also remain regarding AFM results: Why are peak force error images shown instead of stiffness and modulus images. They might not reflect changes in mechanical properties, but where the feedback system struggles to maintain the setpoint force. Stiffness and modulus are defined in a strange way, as “modulus refers to elasticity of the cell walls under turgor pressure, while stiffness reflects the overall resistance of the cell wall against deformation”(Line110-112). Why is modulus related to turgor pressure? Normally modulus is referred to the “intrinsic” material properties (taking geometry into account), while stiffness just refers to materials resistance to deformation under the applied force. In line 116 suddenly a conclusion is made on toughness, which is a mechanical parameter usually not gained by AFM data as no full strain-stress curve is acquired. Please take care about basic mechanical definitions.\nIn line 120 “ a remarkable correlation between mechanical parameters and cotton fiber growth” is reported, but just due to the fact that the two cultivars showed differences in modulus. Based on the presented data it is not easy to follow and verify such correlation and statement! It could be a bit too far fetched to conclude from a lower modulus and stiffness in Hai fibers to a connection to greater final length (Line 123-126)! In Fig. 1, the modulus is similar in both cotton variants at 3DPA, but only the stiffness different. This raises again the question on how these values were derived. It would be more trustable to show at least in the supplementary the raw data – the acquired force curves of the two species, developmental stages and position. It would also be helpful to comment why the DMT model was used, as this is usually more recommended when adhesion forces are significant, like in hydrated conditions. Have you compared results to the Hertz model? How was the quality of the fit to the force-distance data? Was the decision for DMT model based on quality of the fit or other considerations? Have you measured wet cellulose fibrils and therefore chosen DMT? Please add this information in M&M.\nSome detailed comments:\nLine 39-42: conclusion from microtubule perturbation to CSC cellulose distribution and cellulose gradient not clear and not shown. Or have there been any Raman and AFM measurements on the mutant? Figure1: b) why do peak force error images show mechanical features and which ones? Why not pictures of stiffness and modulus are shown? Line 774: what means differed developmental stages? 3 and 7? And are in c) the different positions mixed or is it the base or the shank or the tip? It is misleading that in c the different colors refer to different cotton species and in d) to different developmental stages. Are in the different regions (d) the species mixed or which species?\nSupplementary Figure 1:what are “represent images”? Based on the pictures it is difficult to derive how height and width was measured….please replace “represent images” by which images these are and mark the width and height of the measured fibre in the image.\nFigure 2: a) Not necessary to show a cellulose standard. What is shown in the last? I guess it is the difference between all spectra, but at this size nothing is really visible. B) you talk about relative, so this means it is normalized? If so on what? Why only at 10DPA? c and d relevant for proofing a basipetal gradient of cellulose?"}, "author_response": "Point-by-point responses:\nReviewer #2 (Remarks to the Author):\nReviewer Comments:\nThe paper investigates the mechanism of polarized growth in cotton fiber cells, focusing\non how the differential distribution of cellulose microfibrils (CMFs) drives the\nheterogeneity of the cell wall and directional expansion. The study found that the\ncellulose gradient is a key factor driving polarized growth, while pectin modification\nhas a minimal impact on fiber elongation. These findings deepen the understanding of\nthe mechanisms underlying polarized growth in plant cells, and provide new insights\nfor improving crop fiber quality.\nAll my concerns have been addressed in this revision.\nResponse: We sincerely thank the reviewer for your thoughtful evaluation and kind\nwords regarding our manuscript. We are very pleased to know that the revised version\nhas fully addressed your previous concerns. We are grateful for your supportive\ncomments on our work.\nThere are some minor comments need to be revised:\nResponse: We thank the reviewer for the careful inspection of the figures. For all\nconventional 2D images, we have checked and ensured that scale bars are now properly\nshown. For the 3D reconstruction panels, the images are rendered with an overlaid 3D\ngrid, where each grid unit corresponds to the underlying voxel size. Adding an\nadditional 2D scale bar would obscure the 3D rendering and is not recommended for\nthis representation. To avoid confusion, we have now clarified the spatial scale of these\n3D views explicitly in the figure legends.\nnot indicate which groups were compared, and the asterisks are missing in the figure.\nResponse: We appreciate the reviewer’s comment. The asterisks have been replaced\nwith exact p-values and group comparisons are now clearly stated in the figures.\nReviewer #4 (Remarks to the Author):\nThe authors examine the biomechanics of cotton fibres in the context of cotton fibre\n1\nmorphogenesis. I like the idea of using the cotton fibre to look at growth, especially\nbecause appears to be a blend of tip and diffuse growth, and has a relatively simple and\nhighly anisotropic shape.\nThe authors have shown there is a gradient of cellulose, which they suggest translates\ninto a gradient of stiffness that in turn results in growth gradient driving elongation.\nUnfortunately, they have no quantification showing the presence of growth gradients,\nand their method for measuring stiffness is not really suitable for the diffuse growing\nsection, as it can only measure stiffness normal to the surface.\nWithout any growth measurements and stiffness measurements limited to the normal\ndirection, it is difficult to be convinced of their main hypothesis.\nResponse: We thank the reviewer for the thoughtful and constructive evaluation of our\nwork and for highlighting these two important limitations. We fully agree that direct,\nspatially resolved quantification of growth and in-plane measurements of wall stiffness\nwould provide the most stringent test of a growth gradient model. In the present study,\nour experimental evidence and modelling are intended to support a tip-biased diffuse\ngrowth mode rather than to claim that we have directly measured a continuous growth\ngradient along the fiber. We have carefully revised the text to avoid overstating this\npoint and to clarify the scope of our conclusions.\nTechnically, precise tracking of local growth rates in individual cotton fibres is\nchallenging due to their small diameter, curvature, and movement on the ovule surface,\nand would likely require dedicated microfluidic or long-term immobilization setups that\nare beyond the scope of the current work. Nonetheless, we have tried our best to address\nthe concerns by performing a simple tracking assay, which qualitatively supports faster\nelongation near the apex. Given the limited spatial resolution, these pilot data are\nsummarized in this response letter but not included as a formal figure. In addition, we\ncarried out osmotic deformation experiments to functionally probe regional differences\nin wall deformability. We refer to these new data in the Results and Discussion as\nsupportive evidence for regional differences in growth potential, while still avoiding\nover-interpretation as a fully quantified growth gradient.\nRegarding the stiffness measurements limitation, we agree that AFM does not directly\nmeasure the longitudinal stiffness that is most relevant for axial elongation. In our study,\nAFM-derived apparent modulus and stiffness are therefore used as comparative\nindicators of the local mechanical state of the cell wall, as is common in plant\nbiomechanics studies, rather than as absolute elasticity parameters to directly evaluate\nplant cell growth rate. The new osmotic deformation data help bridge this gap by\nshowing that regions with lower apparent stiffness and modulus are also more\ndeformable under turgor-related perturbation. We have moderated the wording\nthroughout to better reflect these limitations and the scope of our conclusions.\nGeneral comments\ndiffuse growth, it seems to be just taken as given. It would be nice if the authors could\npoint to some growth tracking data that does indeed show this. The authors wrote a\nCurrent Opinion in 2011 on this subject, but did not present any tracking data there\neither. Does this mean it is not possible to track growth in cotton fibres? like people\nhave done for trichomes and pollen tubes? In any case, if this is the state of the art, then\nstatements of how they grow need to be presented more carefully as hypotheses, not\nfact. In addition, it would be good to know what the relative percentages are believed\nto be. Is it mostly at the tip? the flank? 50-50?\nIt is great that you have been able to track the cellulose synthase in this system with\nlive imaging, so is there a reason you cannot track the growth as well? These things\ngrow pretty fast, so one does not need to keep it happy that long.\nResponse: We appreciate this insightful comment and fully agree that direct tracking\nof growth dynamics would strengthen the support for the growth model of cotton fiber.\nIn fact, we had considered and attempted growth-tracking experiments during the initial\nphase of this work. However, precise live tracking of cotton fiber growth remains\ntechnically challenging. At the rapid elongation stage, fibers are long, curved and\nfrequently intertwined on the ovule surface, which makes it difficult to unambiguously\nidentify and follow the apical regions over time. At earlier stages when fibers are shorter\nand less entangled, the instantaneous growth rate is relatively low, so that detectable\nlength changes over experimentally accessible time windows are close to the resolution\nlimit. In addition, we had to partially excise fiber bundles from the ovule and mount\nthem in ovule culture medium under a coverslip; under these conditions, fiber growth\nis usually perturbed and the fibers tend to drift in the liquid, further compromising the\nstability required for accurate long-term tracking. By contrast, tracking the dynamic of\nCESAs and MT organization in live cotton fiber cells is more technically accessible.\nBecause CESA particles provide bright, membrane-localized signals that can be\nfollowed without the need for long-term mechanical stability.\nWe have tried our best to perform a short-term tracking assay using fluorescent beads\nas fiduciary markers on the fiber surface. Although the beads did not allow\nhomogeneous labelling along the entire fiber and their adhesion was not sufficiently\nstable for long-term recording, we followed several extension events from the apical\nand shank regions. A representative example is shown below, together with a plot of\nrelative extension (ΔL/L ) over time (please see Responsive Figure 1).\n0\npreliminary data qualitatively support faster elongation near the apex compared with\nthe shank, consistent with a tip-biased diffuse growth mode. As we felt that the spatial\nresolution and the measurement precision of these attempts were not yet sufficient for\na robust quantitative analysis, we have chosen not to include them in the main text.\nfluorescent carboxylate-modified polystyrene beads (circles) used as fiduciary markers at apical\n(dark red) and shank (cyan) regions. Dashed lines indicate the tracked fiber segments. Scale bar 10\nμm. Right, relative extension (ΔL/L ) over time for the apical (Tip, red) and shank (green) segments;\n0\nlines indicate the mean trend and shaded areas show the confidence interval across tracked fibers.\nRegarding the question about the relative contributions of the tip versus the shank (e.g.\n“50–50”), our current data are not sufficient to provide a robust quantitative partitioning\nof growth along the fiber. While the apical region clearly shows higher instantaneous\nextension potential, the shank spans a much larger fraction of the cell length, and its\ncumulative contribution to final length is likely non-negligible. The precise spatial\nprofile of growth and the relative contributions of apex and shank remain to be\nquantified in future work. Such quantification will likely require dedicated microfluidic\nor immobilization setups that allow stable, long-term tracking of individual fibers. We\nhave now clarified these technical limitations and moderated our statements in the\nDiscussion accordingly. Please see Line 473-479.\nMost of the results in relation to mechanical properties of the wall depend on AFM\nmeasurements. However, I am not sure that this technique is so useful to determine\nstiffness in diffuse growing areas, at least not in a way that is relevant for\nmorphogenesis. In tip growing cells the wall at the apex has no cellulose, and thus\ngrowth there is mostly isotropic. In this case the AFM can likely be used to measure the\nstiffness that could be relevant for growth. In the diffuse part the wall is\nanisotropic because of the cellulose. Since the AFM can only measure in the direction normal to the surface, it cannot\ncapture the directions that are important for growth. What is needed is an in-plane measurement of the stiffness, and in this\ncase the longitudinal direction would be most important.\nResponse: We appreciate this important comment and fully agree that AFM indentation\nprimarily probes the out-of-plane mechanical response of the wall and does not directly\nprovide in-plane elastic constants, particularly in the longitudinal direction. Direct in-\nplane measurements at cellular resolution remain technically challenging in most plant\nsystems, including cotton fibers.\nNevertheless, AFM-based stiffness mapping reflects the composite mechanical state of\n4\nthe wall and has been widely used to reveal spatial heterogeneity of wall mechanics\nrelevant to morphogenesis in plant systems such as guard cells and the shoot apical\nmeristem (Peaucelle, Braybrook et al. 2011, Sampathkumar, Krupinski et al. 2014,\nCarter, Woolfenden et al. 2017). In these contexts, regions with reduced stiffness are\nassociated with higher wall deformation or growth, suggesting that local wall softening\ncan accompany increased expansion potential. In this study, we follow the same logic\nand use AFM-derived apparent modulus and stiffness to provide a relative mechanical\nlandscape that reports local differences in wall mechanical state, rather than as absolute\nin-plane stiffness values or direct measurements of growth rate.\nTo further address this concern, we have now performed osmotic deformation assays.\nUnder hyperosmotic treatment, the apical regions exhibit greater transverse shrinkage\nthan the shank, consistent with a softer and more deformable wall at the apex. These\nresults are in line with the AFM measurements and support the idea that regions with\nlower apparent stiffness have higher deformation potential under changes in turgor.\nWe have incorporated these new data into the revised Fig1. We have also moderated our\nwording to emphasize that our AFM measurements and osmotic assays provide a\nrelative mechanical context for tip-biased diffuse growth, rather than a direct\nquantification of longitudinal stiffness or a fully resolved growth field. Please see Line\n114-123 and Line 142-148.\nOne could use potentially use the AFM to “see” cellulose bundles, as was done in\nSampathkumar et al. 2014 using similar equipment. In that work the images look pretty\nconvincing. Here the images appear very different, and I am not so convinced that these\nimages visualize cellulose fibres. In order to show they do, the authors would need to\nshow they are not just showing for irregularities in the surface shape, for example\nwrinkling from being plasmolysed. If a smooth patch of surface is found, with stiffer\nand softer sections, then this could indicate the presence of cellulose bundles. So it\nwould be good to see both the surface topography and the stiffness map side by side.\nResponse: We appreciate this valuable comment and the reference to Sampathkumar\net al. (2014) (Sampathkumar, Krupinski et al. 2014). The system used in that study was\nthe Arabidopsis cotyledon epidermis, where cellulose microfibrils form a reticulate\nnetwork on a relatively flat surface that is particularly well suited for AFM imaging. By\ncontrast, our own data indicate that the primary walls of cotton fibers contain densely\npacked cellulose microfibrils that are highly aligned in a predominantly transverse\norientation, rather than forming a crossed mesh. Importantly, scanning electron\nmicroscopy (please see Responsive Figure 2) and S4B staining both reveal densely\npacked, transversely oriented cellulose microfibrils on the fiber surface, consistent with\nthe banded patterns in the AFM maps. Thus, the more parallel bands observed in our\nimages most likely reflect a genuine biological difference in CMF organization.\nshowing the inner and outer faces of the cell wall. Transverse banding is observed on both the inner\nand outer faces. (Bottom) Higher-magnification view of the wall surface, revealing densely packed\ncellulose microfibrils aligned and transverse to the fiber axis.\nFrom a technical standpoint, cotton fibers are slender cylindrical protrusions on the\novule surface, which makes it challenging to obtain large, perfectly planar imaging\nareas as in leaf epidermis. To minimize geometric artefacts, we therefore restricted our\nAFM scans to visually smooth segments of the fiber surface and used relatively small\nscan windows to reduce curvature effects. And all measurements acquired and fitted\nunder identical conditions to ensure that relative differences are meaningful. Under\nthese conditions, the apparent modulus maps provide a local readout of wall mechanics,\nwhich are used as comparative indicators of local wall mechanical state between\nregions and genotypes rather than as absolute in vivo stiffness values.\nFor the datasets shown, scan regions were selected only when the height images showed\nno obvious folds or detachment and the force–distance curves were stable and\nreproducible across the field. Under these conditions, the apparent modulus maps\nprovide a robust readout of local wall mechanics, which is the quantity analyzed in this\nstudy. Representative topography images are provided in Supplementary Fig. S1, and\nSEM together with S4B staining (Responsive Fig. 2) independently confirm that the\nbanded patterns reflect transversely aligned CMFs rather than surface wrinkling.\nFollowing suggestions from another reviewer, we now show apparent modulus maps in\nthe revised Fig. 1b, which more directly represent the mechanical contrast used in our\nanalysis. The surface topography remain provided as 3D AFM height renderings in\nExtended Data Fig. S1, which were used for quantifying fiber morphology. Together\nwith the SEM and S4B results, these maps support the interpretation that the AFM\ncontrast reflects the underlying cellulose microfibril organization and wall mechanics,\nrather than artefacts from surface irregularities or plasmolysis.\nway of knowing if more cellulose means more stiffness in the longitudinal direction. To\naccess this, one would need a different method, such at stretching with an extensometer,\nor osmotic treatments. The system is too small for an extensometer, but what about\nosmotic treatments? One should be able to image the fibre, plasmolyse it, and then\nimage again and observe the shrinkage. One could put florescent beads to make\nlandmarks to track the deformation, again as has been done in trichomes and pollen\ntubes.\nResponse: We appreciate this insightful suggestion. In cotton fibers, cellulose\nmicrofibrils are predominantly oriented in the transverse direction, so their primary\ngeometric role is indeed to resist circumferential expansion. At the same time, as the\nmajor load-bearing component of the wall, local cellulose accumulation is expected to\ninfluence the overall stiffness of the wall in both transverse and longitudinal directions\nthrough its interactions with matrix polymers. This view is consistent with recent\ntheoretical treatments of the cell wall as an anisotropic composite material, in which\ncellulose reinforcement influences the overall stiffness of the wall in multiple directions\n(Coen and Cosgrove 2023). In our study, we therefore combine information on cellulose\ndistribution, wall mechanics and modelling to infer how cellulose accumulation may\nshape deformation patterns along the fiber.\nTo provide the in-plane deformation readout suggested by the reviewer, we have now\nperformed osmotic treatments on living 3-DPA fibers. Fibers were imaged after\ntreatment with either 0.8 M mannitol or pure water, and changes in fiber width were\nquantified at the tip and the shank. Mannitol-induced plasmolysis caused a significantly\nlarger transverse shrinkage at the tip than at the shank, whereas mock-treated fibers\nshowed minimal changes (revised Fig. 1e,f). These data indicate that apical walls are\nmore deformable under turgor-related perturbation, consistent with the softer wall\nmechanics at the tip and with the higher elongation potential predicted by our\nmechanical modelling. Although in the current experiments we quantified transverse\nshrinkage rather than axial shortening, the osmotic assay nevertheless provides an in-\nplane, turgor-driven measure of local wall compliance that complements our AFM and\nRaman analyses. We agree that fully resolving how transverse cellulose reinforcement\ntranslates into effective longitudinal stiffness will require more refined in-plane\nmechanical measurements. We have now discussed this point in the revised Discussion\nas an important direction for future work. Please see Lines 476-479.\nAnother related issue throughout is the authors are stressing the importance of a\ncellulose gradient as a main driver for shape and morphogenesis in the cotton fiber.\nSince the cellulose is predominately transverse, would that not imply it primarily stop\nthe cells from widening? And how is it that they don’t get too wide towards the tip,\nwhere there is much less cellulose? especially since the stress is higher in the transverse\ndirection. One has to wonder if this system mimics a bit pollen tube growth. In pollen\ntubes there is pure tip growth, but the tip has no cellulose, it appears displaced from the\ntip later on, mostly in a transverse pattern as here. Thus in pollen tubes, you get a very\nelongated shape from isotropic growth, that it limited to a small area at the tip. Cellulose\nin pollen tubes does not appear to be involved in creating the shape, but only appears\nin areas after the growth has stopped. The more tapered shape in cotton fibres that enters\nthe region with cellulose suggests that diffuse growth is required, and that it cannot be\nsolely tip growth\nResponse: We thank the reviewer for raising this important point. We fully agree that\ntransversely oriented cellulose microfibrils primarily act to restrict radial expansion and\nthereby help maintain the anisotropic geometry of fiber cells during elongation. In the\npresent study, however, our main focus is on how differential cellulose deposition and\nwall mechanics relate to axial elongation, rather than on providing a full mechanical\ndescription of fiber width and tip shape. To avoid overstating this aspect, we have now\nclarified this point more explicitly in our description of the growth model. Please see\nLines 406-411.\nWe also wish to clarify that we do not propose purely tip growth in cotton fibers; instead,\nour data support a tip-biased diffuse growth mode in which both the apex and the shank\nelongate, but with higher elongation potential near the tip. Cotton fiber growth is\ntherefore distinct from pollen tubes, which rely on highly localized, isotropic tip growth\nsupported by actin-rich vesicle delivery. In our interpretation, the apical region with\nlower CMF density allows faster longitudinal wall expansion under turgor, while the\nremaining cellulose and matrix polymers still provide sufficient anisotropic\nreinforcement to limit excessive radial swelling. As a consequence, axial expansion at\nthe tip outpaces lateral expansion, producing a narrow apex, and differential expansion\nbetween apical and subapical regions naturally generates a tapered shape. This differs\nfrom pollen tubes, where cellulose predominantly appears behind the growth zone and\nfunctions mainly as a stabilizer after expansion has ceased.\nThe detailed establishment of tip shape is likely to involve further biochemical and\nbiomechanical mechanisms beyond the cellulose gradient itself. It has been implicated\nfeatures such as local cytoskeletal organization and regional differences in wall\npolymers in shaping cotton fiber tips. And wall-modifying or structural proteins are also\nexpected to fine-tune tip morphology through more complex mechanisms. These\nprocesses lie beyond the scope of the present work and will require future targeted\nstudies. We now state this explicitly in the Discussion. Please see Line 406-411\nThe authors frequently conflate the concepts of elasticity and extensibility. They are not\nthe same, and just because a wall is more elastic (softer) does not mean it is more\nextensible. A wall needs to be elastic (have stretch) and have extensibility factors (ie\nexpansins, etc) in order to grow. Without the latter the wall will not grow. It could be\nthat there is a gradient of extensibility factors, rather than differences in elasticity,\nthat are the primary drivers of growth gradients. Although expansin appears as the\npredominant extensibility factor in cell wall with cellulose, it is not clear what the\nextensibility actors are at the tip where growth happens without cellulose, since the\n8\nusual pectin suspects have been apparently ruled out.\nResponse: We appreciate this important clarification and fully agree that wall elasticity\nand extensibility are conceptually distinct. Our AFM measurements quantify apparent\nmodulus and stiffness to describe the mechanical state of the fiber cell wall, rather than\nto directly measure growth rate or extensibility. At the same time, previous studies have\nreported that regions with lower apparent wall stiffness are spatially associated with\nzones of higher wall deformation or growth, suggesting that local wall softening can\naccompany increased expansion potential (Peaucelle, Braybrook et al. 2011, Carter,\nWoolfenden et al. 2017, Cosgrove 2018, Traas 2023). In this sense, elasticity provides\na necessary physical background that can facilitate extension. We apologize that our\nearlier wording may have conflated these concepts. In the revised manuscript, we now\nmake it explicit that the mechanical differences we observe between tip and shank are\ninterpreted as differences in local wall compliance that may underlie differential\ndeformation capacity, rather than as direct measurements of extensibility. Please see\nLines 112-123\nWe also agree that biochemical wall-loosening factors are likely to play an important\nrole in establishing local extensibility patterns. Factors such as expansins and XTHs\nmay form spatial gradients that act in concert with cellulose-based mechanical\nheterogeneity to generate growth heterogeneity. However, the spatial distribution of\nthese proteins in cotton fibers remains largely unexplored. We have now expanded the\nDiscussion to acknowledge this possibility and emphasize that mapping wall-loosening\nfactors will be an important direction for future work. Please see Lines 439-444.\nSpecific comments:\nNot heard of the Turkey test (brings strange images to mind), perhaps the authors mean\nTukey test?\nResponse: Thank you for pointing this out. We apologize for the typographical error.\nWe have corrected it to Tukey test throughout the manuscript.\nLine 40: GhCSI1 is mentioned in the abstract without saying what it is.\nResponse: Thank you for this comment. To avoid introducing an undefined gene name\nwithin the strict word limit of the abstract, we have removed GhCSI1 from the abstract.\nThe full name and description of GhCSI1 are retained and defined in the main text.\nLine 52: What about callose?\nResponse: Thank you for this helpful comment. Callose can transiently accumulate in\nspecific developmental or stress-related contexts, but it is generally not considered a\nmajor, ubiquitous structural component of primary walls. To avoid confusion, we now\nexplicitly state that primary cell walls are “mainly” composed of cellulose,\nhemicelluloses and pectins. Please see the revised sentence at Line 51.\nbehaviour, how the wall extends to make shape is a different matter, although they are\nusually, but not always, correlated.\nResponse: Thanks for this insightful remark. We agree that cellulose microfibrils\nprimarily create anisotropy in the wall that governs its elastic behavior, and that how\nthe wall extends to generate shape is a related but distinct issue. We have revised the\nsentence to clarify that wall heterogeneity leads to anisotropic mechanical responses to\nturgor pressure, which in turn bias deformation direction, rather than implying a direct\ncontrol of growth. Please see Line 57-60.\nLine 76: Deposition stops, but secondary cell wall is produced? Is that not still cell wall\n(ie cellulose) deposition?\nResponse: Thank you for pointing out this ambiguity. Our intention was not to imply\nthat all wall deposition stops, but rather to refer to the end of the PCW dominated\nelongation stage. We have revised the sentence to avoid the misleading wording. Please\nsee Line 77-79.\nLine 110: It is not clear what the authors mean by modulus and stiffness here. The\nauthors only talk about stiffness in the methods. The fibres were dried, so I don’t\nunderstand how they measure modulus if they say it relates turgor pressure. If modulus\nis being calculated from the stiffness with some sort of Herzian contact model, I am not\nsure why they should be so qualitatively different. This section needs substantial clean\nup, and needs to be made much more precise.\nThe max force for the AFM and indentation depth should also be reported.\nResponse: Thank you for this helpful comment. We agree that our previous wording\nwas not sufficiently precise and may have caused confusion. In the revised manuscript,\nwe now state more clearly that both modulus and stiffness are AFM-derived descriptors\nof the elastic properties of the fiber cell wall surface, and that we use them as\ncomparative measures of local wall mechanics rather than as direct readouts of turgor-\nrelated elasticity or growth. Please see Lines 112-123.\nAFM measurements on intact cotton fibers are technically challenging due to curvature\nand surface roughness and, to our knowledge, comparable datasets are still scarce.\nCotton fiber walls are biological composites that are not perfectly linearly elastic, and\nAFM measurements on fiber surfaces can be influenced by factors such as adhesion.\nOur initial analysis pipeline for modulus may not have fully accounted for these\ncomplexities. Because the fiber cell walls exhibit strong adhesion and nonlinear force-\ndisplacement behavior, In the revision, we have therefore re-analyzed the AFM data by\ndirectly using the stiffness and modulus values provided in the QNM maps (Hertz\ncontact model with adhesion enabled). The overall trends, including softer apex versus\nshank and differences between cultivars, remain unchanged. Please see revised Fig.1.\nWe thus treat AFM-derived modulus and stiffness as comparative indicators rather than\nabsolute measurements of in vivo wall properties, and interpret these parameters as\nproviding a relative mechanical landscape that reports local differences in wall\nmechanical state.\nThe force setpoint and the typical indentation depth used in the AFM measurements are\nnow reported in the Methods section.\nLine 124: Again, you are conflating elasticity and extensibility. (also Lines 130,\nResponse: Thank you for pointing this out. We agree that our original wording at these\nlines could be read as conflating elasticity with extensibility. In line with our revised\ndescription of AFM-derived parameters as comparative indicators of wall mechanics\nrather than direct readouts of extensibility, we have now rephrased these sentences in\nterms of mechanical compliance and expansion potential. Please see Lines 123-128 and\n131-134 in the revised manuscript.\nLine 312: What existing paradigms about cell wall extensibility are being challenged?\nResponse: Thank you for pointing this out. We agree that our original wording was too\nstrong and too vague. What we intended to convey is that the traditional view of wall\nextensibility has largely emphasized pectin modifications and matrix polysaccharides\nas the primary regulators of wall yielding, with cellulose often considered mainly as a\nstructural scaffold. In our system, however, we find that differential cellulose deposition\ncreates marked mechanical heterogeneity along the fiber and is closely associated with\npolarized elongation, whereas pectin modification appears to play a more limited role\nduring the early rapid-elongation phase.\nTo avoid overstatement, we have revised the sentence to specify this point and to frame\nour results as “refining and extending”, rather than challenging the conventional view.\nPlease see the revised text around Line 359-363\nLine 339: Do you really know what the gradient of extensibility looks like? If you have\nnow time-lapse tracking, how do you know there is a gradient of growth. It could be\nthe tip grows fast, and the whole shank grows more slowly, but the same. I don’t think\nyou can say this with any certainty.\nResponse: Thank you for raising this important point. We agree that we do not directly\nmeasure a gradient of wall extensibility or growth, and that our data are insufficient to\nmap a continuous growth field along the fiber. Our observations of CSC density and\nCMF distribution, together with the computational modelling and a short-term tracking\nassay that qualitatively supports faster elongation near the apex (see Responsive Figure\n1), are consistent with a tip-biased diffuse growth mode, but they do not allow us to\ndefine a precise gradient of extensibility.\nTo avoid over-interpretation, we have revised the text accordingly. We now refer to a\ncellulose gradient to spatial variation in wall mechanical state and deformation potential,\nas inferred from AFM and osmotic assays. Please see Line 388-391. We have also added\na discussion of future directions for directly resolving growth gradients in cotton fibers\nPlease see Line 469-476.\nhypothesis, which so far has little direct experimental support.\nResponse: Thank you for this comment. We agree that the idea that microtubules\nrespond to mechanical cues is still best regarded as a working hypothesis, rather than\nan established universal principle, and that the strength of experimental support varies\nbetween systems. Our intention was not to claim this as a fully proven mechanism, but\nto refer to a possible link between wall mechanics and MT organization. Please see Line\n393-397.\nLine 375: The idea that cellulose is the most important player for elongation, and that\npectin is not important, is in obvious contrast with rapid growth at the tip. Again this\nstresses the necessity to have growth data in this system to support this hypothesis.\nResponse: Thank you for this comment. We would like to clarify that our data and\ninterpretation do not imply that cellulose content is positively correlated with local\ngrowth rate. Rather, our conclusion is that differential cellulose deposition plays a key\nmechanical role in shaping the wall properties and anisotropy that underlie fiber\nelongation in our system. In our model, the apical region with lower cellulose density\nis mechanically softer and more compliant, which allows greater deformation under\nturgor provided that wall-loosening factors are present, whereas the gradual increase in\ncellulose toward the shank reinforces the wall and limits further extension. Thus, the\nobservation of rapid growth at tip is not in contradiction with a central mechanical role\nfor cellulose.\nTo avoid overstatement, we have softened the wording in the Discussion to emphasize\nthat compared to pectins, cellulose emerges as a major mechanical contributor to fiber\ncell wall extensibility in our system. Please see Lines 435-436 in the revised manuscript.\nRegarding the need for growth data, we fully agree that direct measurements of growth\ndistribution provide an important test of any mechanistic model. As detailed in our\nresponses to earlier comments, we have now added osmotic deformation assays and a\nshort-term tracking experiment, and we have revised the Discussion to frame our\nconclusions as being consistent with a tip-biased diffuse growth mode. Future work will\nbe required to quantitatively map growth gradients along the fiber. Line 469-479.\nReviewer #4 (Remarks on code availability):\nThe model seems pretty straightforward, more springs = slower growth, so the outcome\nis intuitive and I am sure it can be easily reproduced.\nResponse: We thank the reviewer for this positive assessment.\nReviewer #5 (Remarks to the Author):\nfrom the apex to the shank. Apical regions are reported to be softer, while the shank is\nstiffer to support overall cell integrity. It make sense and is verified by computational\nsimulation that CMF distribution influences strain patterns along the fibers. Primary\nwall during cotton fiber elongation is reported with 22% semi-crystalline cellulose\nfibrils, which are surrounded by matrix polymers including xyloglucan and pectin. So\nfar, there has been a lot of evidence that presence and modification of the matrix\npolymer may regulate fiber elongation.\nResponse: We thank the reviewer for the clear and positive summary of our study and\nare pleased that the overall framework, which is differential CMF distribution is\nassociated with wall mechanical heterogeneity and polarized elongation in cotton fibers,\nis considered mechanistically sensible. We fully agree that previous work has\nestablished important roles for matrix polymers such as pectins and hemicelluloses in\nregulating primary wall extensibility. In the present work, our focus is specifically on\nhow differential cellulose deposition contributes to wall mechanical heterogeneity and\npolarized elongation during the early elongation of cotton fiber development. We have\nclarified this scope more explicitly in the revised manuscript. And in the following\nresponses we address the detailed comments on the Raman analysis and AFM-based\nmechanical measurements.\nIn this study, the focus is solely on cellulose accumulation and the main evidence by\nRaman spectroscopy lacks integrity of data and spectral interpretation. Only 3 spectra\nare shown from 10DPA and analysis is only based on the 380cm-1 band, which is taken\nand interpreted as cellulose accumulation. Raman spectra of cellulose reflect also\nchanges in cellulose orientation and crystallinity, which is not analyzed and taken into\naccount at all. Further, pectin and hemicellulose bands should not be overseen and\ndiscussed. Unfortunately only 10DPA is analyzed with Raman, while data from AFM\nare from 3DPA and 7DPA and staining from 3DPA and 2DPA, cell wall synthesis 3DPA.\nThe study would benefit from a more conclusive experimental design including Raman\nat different developmental stages (same as AFM) and mutants added. Additional the\nstudy will benefit from a more comprehensive Raman spectra interpretation, instead of\nplotting the intensity of one peak for cellulose content. That can only be done, if\nchanges in orientation and crystallinity can be ruled out based on the spectral signatures.\nResponse: We thank the reviewer for these detailed and very helpful comments on the\nRaman analysis. In the revision, we have extended our laser-tweezers Raman\nmeasurements to additional developmental stages and to wall-polymer standards, and\nwe have clarified the scope of what we infer from the spectra.\nIn the revised manuscript we now show spectra from 3, 7 and 10 DPA fibers together\nwith spectra of cellulose, pectins and hemicellulose (Xylan) standards. Across\ndevelopment, the overall spectral shape of the fibers progressively converges towards\nthat of crystalline cellulose, consistent with an increasing contribution of cellulose to\nthe wall signal. However, the spectra at 3 and 7 DPA differ from all three standards:\nthey show broader features and additional bands that likely reflect a mixture of wall\n13\nand intracellular components, especially given the thin walls and relatively large\ncytoplasmic volume at early stages. Although spectra within each stage are reproducible,\ntheir interpretation in terms of individual wall polymers is not straightforward. For this\nreason, we now explicitly state that the 3/7 DPA Raman data are used only qualitatively\nto illustrate the developmental trend towards more cellulose-like spectra, and not for\nquantitative comparison of cell wall composition (please see Line 179-193). We also\napologize that our sampling strategy was not made sufficiently clear in the original\nfigure. The three 10-DPA traces represent averaged spectra, not single measurements:\nfor each position, multiple spectra were collected from several fibers, background-\ncorrected and normalized to their maximum, and then averaged. We now state these\nnumbers explicitly in the figure legends.\nTo address the concern that our original analysis relied on a single band at ~380 cm-1,\nwe have added a second cellulose-associated window around ~1120 cm-1. After\nbackground correction and normalization of each spectrum to its maximum intensity,\nwe quantify the relative peak intensities at ~380 and ~1120 cm-1 along 10 DPA fibers.\nBoth bands report a consistent increase in cellulose-related signal from tip to sub-tip to\nshank (revised Fig. 3 and Supplementary Fig. 4), supporting our conclusion of a\nbasipetal enrichment of cellulose. In the text we now emphasize that the 380 and 120\ncm-1 band is used as a convenient representative peak for this trend, rather than as the\nsole basis of our interpretation. Please see Line 193-198\nWe also have explicitly considered the contributions of pectins and hemicelluloses.\nPectin and xylan standards display characteristic bands that are clearly distinct from the\ncellulose standard. The fiber spectra do not show strong changes in the pectins or xylan-\nassociated bands. On this basis we infer that the dominant Raman differences along the\nfiber axis most likely reflect changes in cellulose content. At the same time, we now\nclearly state in the Results that the absence of distinct pectin/xylan bands in our spectra\ndoes not imply that these polymers are absent from the wall; rather, their signals are\nprobably masked by the stronger cellulose deposition or fall below the sensitivity of\nour current laser-tweezers setup. Please see Line 199-208.\nWe agree that Raman spectra of cellulose also contain information about crystallinity\nand molecular orientation. A rigorous analysis of these aspects would require\npolarization-resolved measurements on well-defined wall geometries and a more\nextensive spectral deconvolution than is feasible within the scope of this study. Our\nprimary goal was to use Raman spectroscopy as a strength tool to assess relative\ncellulose enrichment along the fiber axis, rather than to perform an exhaustive,\nspectroscopy-driven analysis of all structural parameters. Moreover, how changes in\ncrystallinity and microfibril orientation quantitatively influence cotton fiber\ndevelopment is still not well understood. We therefore use the Raman data to inferring\nrelative cellulose enrichment along the fiber axis, and we avoid making claims about\nchanges in cellulose orientation or crystallinity. In addition, we note that Raman\nanalyses specifically focused on cotton fibers are still very scarce. A more exhaustive\n14\nspectral interpretation, including systematic analyses of mutants and a detailed changes\nof crystallinity and molecular orientation, would be an interesting follow-up direction\nbuilding on the current work. Please see Line 208-216.\nTogether with the CFW and S4B staining and the distribution of cellulose synthase\ncomplexes, the revised Raman analysis provides a more coherent basis for our\nconclusion that cellulose content increases basipetally along elongating fibers.\nCorresponding changes and clarifications have been made in the Results.\nIn the Material and Method part of Raman experiments no information is given, but\nonly the link to a paper which has nothing to do with Raman: 51) Liu, W. et al. Towards\na better recording of microtubule cytoskeletal spatial organization and dynamics in\nplant cells. J Integr Plant Biol 61, 388-393 (2019). https://doi.org/10.1111/jipb.12721.\nPlease provide all necessary information or at least correct citation. And it would be\ninteresting for the reader to comment why laser tweezers Raman microspectroscopy\nwas chosen and not conventional Raman imaging using a Confocal Raman microscopy,\nwhich might give more detailed information including spatial changes in content and\norientation.\nResponse: We thank the reviewer for pointing out this issue. The previous citation in\nthe Raman Methods section was indeed incorrect and could be resulted from a reference\nmismatch during manuscript merging. We have now replaced it with two appropriate\nRaman references describing a similar laser-tweezers Raman micro-spectroscopy\napproach (Lu, Chen et al. 2020, Lu, Li et al. 2022) and we have substantially expanded\nthe description of our experimental conditions.\nRegarding the choice of laser-tweezers Raman versus confocal Raman imaging, our\nlaboratory is currently only equipped with an LTRS system but not with an imaging\nmodule. In combination with CFW and S4B staining, which independently reveal the\nspatial distribution of cellulose in the fiber wall, these point spectra from LTRS. have\nprovided sufficient information to support our conclusion of a basipetal increase in\n\ncellulose content. We now note that future work using confocal Raman imaging on fiber\nsamples would be valuable for more detailed mapping of spatial changes in wall\ncomposition and orientation. Please see Line 211-214.\nSome questions also remain regarding AFM results: Why are peak force error images\nshown instead of stiffness and modulus images. They might not reflect changes in\nmechanical properties, but where the feedback system struggles to maintain the setpoint\nforce. Stiffness and modulus are defined in a strange way, as “modulus refers to\nelasticity of the cell walls under turgor pressure, while stiffness reflects the overall\nresistance of the cell wall against deformation” (Line110-112). Why is modulus related\nto turgor pressure? Normally modulus is referred to the “intrinsic” material properties\n(taking geometry into account), while stiffness just refers to materials resistance to\ndeformation under the applied force. In line 116 suddenly a conclusion is made on\ntoughness, which is a mechanical parameter usually not gained by AFM data as no full\n15\nstrain-stress curve is acquired. Please take care about basic mechanical definitions.\nResponse: We thank the reviewer for these careful comments on the AFM analysis and\nfor drawing attention to the mechanical terminology. We fully agree that peak force\nerror images mainly reflect feedback performance and are not the most direct\nrepresentation of mechanical properties. We apologies for this inappropriate choice in\nthe initial version. In the revised Fig. 1b we now show apparent modulus maps, which\ndirectly correspond to the parameter used for our quantitative analysis.\nWe also agree that our earlier wording about modulus and stiffness was not sufficiently\nprecise and could be misleading. In the revision, we explicitly state that both quantities\nare AFM-derived descriptors report the intrinsic elastic properties of the wall material,\nrather than as direct measures of in vivo elasticity under turgor or of growth rate.\nPrevious studies have reported that regions with lower apparent wall stiffness are\nspatially associated with zones of higher wall deformation or growth, suggesting that\nlocal wall softening can accompany increased expansion potential (Peaucelle,\nBraybrook et al. 2011, Carter, Woolfenden et al. 2017, Cosgrove 2018, Traas 2023).\nAlthough our AFM measurements cannot be taken as absolute in vivo stiffness values,\nthey provide a relative mechanical landscape that is informative for understanding\ncotton fiber elongation. We have rewritten the relevant sentences accordingly. Please\nsee Lines 116-123.\nWe further agree that it is not appropriate to infer “toughness” from our AFM data, as\na full stress-strain curve is not obtained. We have therefore removed the previous\nstatement referring to toughness and no longer use this term in the revised manuscript.\nThe stress-strain curve will require future in-plane mechanical measurements and high-\nresolution strain mapping, which is currently not applicable. Instead, we have\nperformed osmotic deformation assays. The apical regions exhibit significantly greater\ntransverse shrinkage than the shank under hyperosmotic treatment, in contrast to\nminimal changes in water-treated controls. These results are consistent with the AFM\nmeasurements and support the idea that regions with lower apparent stiffness have\nhigher deformation potential under changes in turgor. We have incorporated these new\ndata into the revised Fig. 1e,f. Therefore we interpret AFM-derived modulus and\nstiffness in combination with the osmotic shrinkage assays for inferring cell wall are\nmechanically compliant, rather than as standalone, absolute elasticity values. Please see\nLine 131-146 and Line 142-148.\nIn line 120 “ a remarkable correlation between mechanical parameters and cotton fiber\ngrowth” is reported, but just due to the fact that the two cultivars showed differences in\nmodulus. Based on the presented data it is not easy to follow and verify such correlation\nand statement! It could be a bit too far fetched to conclude from a lower modulus and\nstiffness in Hai fibers to a connection to greater final length (Line 123-126)! In Fig. 1,\nthe modulus is similar in both cotton variants at 3DPA, but only the stiffness different.\nThis raises again the question on how these values were derived. It would be more\ntrustable to show at least in the supplementary the raw data – the acquired force curves\n16\nof the two species, developmental stages and position. It would also be helpful to\ncomment why the DMT model was used, as this is usually more recommended when\nadhesion forces are significant, like in hydrated conditions. Have you compared results\nto the Hertz model? How was the quality of the fit to the force-distance data? Was the\ndecision for DMT model based on quality of the fit or other considerations? Have you\nmeasured wet cellulose fibrils and therefore chosen DMT? Please add this information\nin M&M.\nResponse: We thank the reviewer for these thoughtful comments on the AFM analysis\nand the interpretation of the mechanical parameters. We agree that our previous\nwording was too strong. In the revised manuscript, we now describe this only as a\npossibility: we note that the cultivar with lower apparent wall modulus and stiffness\nalso develops longer fibers. This co-occurrence raises the hypothesis that early wall\nmechanics may influence final fiber length. If this potential link could be tested in future\nstudies using larger cotton populations, it might provide a useful mechanical indicator\nfor fiber-quality breeding. Please see Line 132-136.\nAFM measurements on intact cotton fibers are technically challenging due to curvature\nand surface roughness, and cotton fiber walls are biological composites that are not\nperfectly linearly elastic; in addition, adhesion at the wall interface can influence the\nfitted parameters. Therefore, we re-analyzed the AFM datasets by re-selecting regions\nof interest and directly using the stiffness and modulus values provided by the\ninstrument software. The overall trends, including softer apex versus shank and\ndifferences between cultivars, remain unchanged. Please see revised Fig.1. We thus\ntreat AFM-derived modulus and stiffness as comparative indicators rather than absolute\nmeasurements of in vivo wall properties, and interpret these parameters as providing a\nrelative mechanical landscape that reports local differences in wall mechanical state.\nWe now provide representative averaged force-indentation curves for both cultivars,\nand different positions. Please see Responsive Figure 3. These curves display smooth,\nphysically reasonable force-distance behavior and are consistent with the lower\napparent stiffness of fiber apex regions as reported in Fig 1. If the editor and reviewers\nconsider it useful for readers, we would be happy to include this figure as a\nsupplementary figure.\nforce-indentation responses (lines with error bars) are shown for different cultivars, TM-1 (G.h) and\nHai7124 (G.b), developmental stages (3 and 7 DPA) and positions along the fibre (tip, shank and\nbase), acquired under identical probe and imaging conditions. These curves are consistent with the\nrelative stiffness differences between fiber tip and shanks.\nBecause the fiber cell walls exhibit strong adhesion and nonlinear force-displacement behavior,\nin our re-analysis, all quantitative stiffness and modulus values reported in\nFig. 1 are taken directly from the QNM maps, as calculated by NanoScope Analysis 1.8\nusing the built-in Hertz contact model with adhesion enabled. As our analysis focuses\non relative differences between regions or cultivars, rather than on absolute modulus\nvalues, we did not perform a systematic comparison with alternative models. The\ndescription has been corrected accordingly in the Methods.\nSome detailed comments:\nLine 39-42: conclusion from microtubule perturbation to CSC cellulose distribution\nand cellulose gradient not clear and not shown. Or have there been any Raman and\nAFM measurements on the mutant? Figure1: b) why do peak force error images show\nmechanical features and which ones? Why not pictures of stiffness and modulus are\nshown? Line 774: what means differed developmental stages? 3 and 7? And are in c) the different positions mixed or is it the base or the shank or the tip? It is misleading that in c the different colors refer to different cotton species and in d) to different developmental stages. Are in the different regions (d) the species mixed or which species?\nSupplementary Figure 1:what are “represent images”? Based on the pictures it is\ndifficult to derive how height and width was measured….please replace “represent\n18\nimages” by which images these are and mark the width and height of the measured fibre\nin the image.\nResponse: We thank the reviewer for these careful comments. In this study, the\nmicrotubule-perturbation experiments as well as the csi1-cr lines were designed to test\nthe coupling between cortical microtubules and CSC distribution, rather than to directly\nre-measure the cellulose gradient or wall mechanics in these backgrounds. We have\ntherefore revised the Abstract to make this clear. We did not perform AFM or Raman\nmeasurements on drug-treated ovules or mutants, because short-term microtubule\nperturbations in cultured ovules are unlikely to produce robust, spatially resolved\nchanges in cellulose deposition that can be captured by these techniques. Moreover,\nperform AFM on fiber samples as well as interpretation of Raman spectra remain\ntechnically challenging. A rigorous characterization of cellulose gradients and wall\nmechanics under such perturbations would therefore require a dedicated, technically\noptimized study in future.\nWe agree that peak force error images mainly report feedback performance and are not\nthe most direct representation of mechanical properties. We apologize for this earlier\nchoice. In the revised Fig. 1b, we now show the apparent modulus maps, which\ncorrespond to the parameter used for analysis.\nRegarding the legend for Fig. 1c, d, we apologize for our previous wording could be\nconfusing. In Fig. 1c, only apical regions of fibers at 3 and 7 DPA are analyzed, and the\ntwo colors represent the two cultivars (TM-1 and Hai7124). This is now explicitly\nstated in the legend. In Fig. 1d, the analysis of different positions (tip vs shank) is\nperformed on TM-1 fibers at 3 and 7 DPA. Here the colors indicate developmental\nstages (3 and 7 DPA), not cultivars. We have clarified in the legend that TM-1 is used\nin this panel.\nIn Supplementary Figure 1, we have revised the legend with a more precise description\nof the samples shown. On each 3D AFM height renderings, we have added a double-\nhead arrow indicating the line along which the height profile was taken. The CMF width\nand height were obtained by extracting the height profile along this line and fitting it\nwith a Gaussian function. Full width and peak height from the fit were taken as the\napparent width and height of the CMF.\nFigure 2: a) Not necessary to show a cellulose standard. What is shown in the last? I\nguess it is the difference between all spectra, but at this size nothing is really visible. B)\nyou talk about relative, so this means it is normalized? If so on what? Why only at\n10DPA? c and d relevant for proofing a basipetal gradient of cellulose?\nResponse: We thank the reviewer for these helpful comments on the Raman analysis\nand presentation. In the revised version, the cellulose, pectins and xylan standards have\nbeen moved to a Supplementary Figure 4.\nAnd we now display the mean spectra from tip, sub-tip and shank of 10 DPA fibers in\nFig 3 together with the mean spectra from 3 and 7 DPA. Because the measurements\nfrom 3 and 7 DPA fibers show more complex and deviate from the cellulose standard,\nlikely reflecting a mixture of thin primary wall and intracellular contributions, we now\nuse these spectra only to illustrate the developmental evolution of the fiber Raman\nsignature. The description has been corrected accordingly in the Main text. Please see\nLine 185-193.\nAll Raman spectra were processed by Savitzky-Golay smoothing, polynomial baseline\nsubtraction, and min–max normalization. On this basis, the peak heights at ~380 cm-1\nand ~1120 cm-1 are used as representative cellulose-associated bands to evaluate\nrelative cellulose enrichment. We now explicitly state it in the Methods and figure\nlegend. This is now explicitly stated in the Methods and figure legend.\nThe CFW stainings are intended as independent imaging evidence for a differential\nenrichment of cellulose, complementary to the Raman data. In the same figure, the\nisoxaben inhibition results are included as functional assays showing that cellulose\ndeposition is essential for normal wall formation and fiber elongation. Although these\nperturbations are not used to reveal the differential cellulose deposition, they support\nthe important role of cellulose in cotton fiber growing. So we consider it appropriate to\npresent them together with the Raman results.\nReferences\nCarter, R., H. Woolfenden, A. Baillie, S. Amsbury, S. Carroll, E. Healicon, S. Sovatzoglou, S.\nBraybrook, J. E. Gray, J. Hobbs, R. J. Morris and A. J. Fleming (2017). \"Stomatal Opening\nInvolves Polar, Not Radial, Stiffening Of Guard Cells.\" Curr Biol 27(19): 2974-2983 e2972.\nCoen, E. and D. J. Cosgrove (2023). \"The mechanics of plant morphogenesis.\" Science\n379(6631): eade8055.\nCosgrove, D. J. (2018). \"Diffuse Growth of Plant Cell Walls.\" Plant Physiol 176(1): 16-27.\nLu, W., X. Chen, L. Wang, H. Li and Y. V. Fu (2020). \"Combination of an Artificial Intelligence\nApproach and Laser Tweezers Raman Spectroscopy for Microbial Identification.\" Anal Chem\n92(9): 6288-6296.\nLu, W., H. Li, H. Qiu, L. Wang, J. Feng and Y. V. Fu (2022). \"Identification of pathogens and\ndetection of antibiotic susceptibility at single-cell resolution by Raman spectroscopy combined\nwith machine learning.\" Front Microbiol 13: 1076965.\nPeaucelle, A., S. A. Braybrook, L. Le Guillou, E. Bron, C. Kuhlemeier and H. Hofte (2011).\n\"Pectin-induced changes in cell wall mechanics underlie organ initiation in Arabidopsis.\" Curr\nBiol 21(20): 1720-1726.\nSampathkumar, A., P. Krupinski, R. Wightman, P. Milani, A. Berquand, A. Boudaoud, O.\nHamant, H. Jonsson and E. M. Meyerowitz (2014). \"Subcellular and supracellular mechanical\nstress prescribes cytoskeleton behavior in Arabidopsis cotyledon pavement cells.\" Elife 3:\ne01967.\nTraas, J. (2023). \"Morphogenesis at the shoot meristem.\" C R Biol 345(4): 129-148.\nThe author has addressed all my comments and it seems to be accepted.\nResponse: We thank Reviewer #2 for the positive evaluation of our revised\nmanuscript. We are pleased that the revisions have addressed all of the\nreviewer’s concerns.\nReviewer #4 (Remarks to the Author):\nI am satisfied with the changes made by the authors.\nResponse: We thank Reviewer #4 for the positive evaluation and for\nrecognizing the improvements made in this revision.\nThe comments related to imprecise terminology in places have been resolved,\nalong with the tempering of some statement where required to avoid\noverstatement. The addition of osmotic treatments is very nice, although it is a\nbit a pity they could not spatially resolve elongation with beads or similar,\nalthough I appreciate the technical challenges there.\nMy other comments were mostly related to growth tracking, which appears to\nbe too technically challenging in this system, although I appreciate the effort\nthere. I guess it is the same as longitudinal deformation tracking in osmotic\ntreatments. It is too bad, because knowing where the growth, and the relative\ncontribution between shank and tip, would strengthen their story, but it is what\nit is.\nResponse: We are pleased that the imprecise terminology has been corrected\nand that statements have been appropriately tempered to avoid overstatement.\nWe also appreciate the reviewer’s comments on the osmotic treatments. We\nagree that spatially resolved elongation tracking would further strengthen the\nmanuscript. However, as the reviewer notes, implementing robust high\nresolution tracking in this system is technically challenging. We are also looking\nforward to emerging approaches those may enable spatially resolved\ndeformation and elongation mapping in cotton fibers.\nA few small points:\nLine 112-116: This is OK, bit I would go with “cell wall stiffness” or “cell wall\nelastic modulus” for the cell wall, and “apparent stiffness” or “apparent elastic\nmodulus” for the AFM stiffness.\nResponse: Thanks for pointing this out. We agree and have revised the\nmanuscript to clearly distinguish intrinsic cell wall elastic modulus/stiffness from\nAFM indentation-derived apparent elastic modulus/apparent stiffness, and we\nnow consistently use the “apparent” terminology when referring to AFM-derived\nvalues throughout the text.\nLine 360-363: The way this is worded, it sounds at first like you are talking about\ncell walls in general, maybe start with “In cotton fibres, our findings …”\nResponse: Thanks for pointing this out. We agree and have revised the\nsentence to explicitly anchor the statement in cotton fibers and to clarify that\nthe “conventional view” is largely based on evidence from other plant systems.\nThe revised text:\n”In cotton fibers, our findings refine and extend the conventional view, largely\nderived from other plant systems, that cell wall extensibility is primarily\nregulated by pectin modifications ...”\nLine 908: I not sure what “mock” means here? You treated some fibres and not\nothers? Normally this would be presented as a before/after experiment. A\npicture of the measurement before treatment (in water) and then after in\nMannitol, ideally with the figure panel showing the same fibre.\nResponse: We thank the reviewer for pointing this out. We agree that “mock”\nwas unclear and have replaced it with “water condition”. The medium was\nexchanged from water to mannitol while imaging/measuring the same ovule\nregion. Because fibers are densely packed, individual fiber pairing cannot be\nreliably ensured, so we report a within-region before/after comparison. We have\nadded this statement in the legend.\nReviewer #4 (Remarks on code availability):\nI looked at the code, but did not verify it in any way. The model is pretty simple,\nand does what one would expect. I don't think it would be hard to reproduce.\nResponse: We thank Reviewer #4 for checking the code availability. We are\npleased that the model implementation is clear and should be straightforward\nto reproduce.\nReviewer #5 (Remarks to the Author):\nThank you for taking into account the mistakes in terminology and for correcting\naccordingly. Instead of additional analysis and/or measurement of the sample\nspectra, non relevant reference spectra for hemicellulose and pectin have been\nadded as Supplementary data as well four spectra of developmental stages\nwithout spectral interpretation. Supplementary reference spectra from my point\nof view can be removed as published elsewhere and the xylan spectrum as well\nas the pectin looks very strange and no clear material and methods from where\nthese references were purchased. Same for 3 and 7DPA as long as they are\nnot analysed as the author state them as “complex and deviate from the\ncellulose standard”. Does this mean at this stage cellulose plays no role? In the\ncurrent form adding the spectra is not beneficial as no conclusions are drawn\nand the idea to have data from the same developing stages from Raman and\nAFM still not fulfilled.\nBased on the new spectra shown in Fig. 3a in the analysis of the 10DPA spectra\nit seems problematic when doing a min-max normalisation over the whole\nwavenumber region as in some spectra 380 will be max and in some 1095,\nwhich of course influences the analysis of peak heights. I do not understand\nwhat the Pearson correlation with the cellulose standard should tell you? As it\nis higher in the tip and sup-tip, you have more cellulose there than in the shank?\nAnd no cellulose in 3DPA and 7DPA?\nBased on the plotted data in Fig. 3b) it is difficult to follow that differences are\nreally statistically significant! Data contradict the authors assumption that these\nbands can be interpreted as cellulose content as they give not the same results.\nFor 1120band the shanke seems not really different anymore (statistical test\ncorrect?) and for 380 probalby some spectra were normalised on the 380band\n(see red spectra in a) and therefore high!.\nFig. 3a,c: I do not understand what the pearson correlation vs. cellulose should\nbring in 3DPA and 7DPA? Important in spectral analysis is not only correlation,\nbut interpreting the peaks. It is logical that cellulose dominates as soon as\npresent in crystalline form. I therefore still think that cellulose in such Raman\ndata can not be analysed only by cellulose content, as the influence of\ncrystallinity and orientation is tremendous on the height of the Raman signal!\nAnd even if contributions of hemicellulose and pectin are naturally less reflected\nin the spectra, they are obviously there and need to be taken into account. If\nthis is not possible for the authors they have to use another method.\nFig. 3d): does not bring any information, it is evident from the spectra that they\nare different and will group accordingly. Interpreting bands of loading 1 and 2\nwould be informative. But please do not show a PCA without discussing\nloadings..than it is better not to show.\nFig. 3e and f: has nothing to do with the Raman results.\nResponse: We sincerely thank Reviewer #5 for the careful and technically\ninformed assessment of our Raman spectroscopy experiments. We apologize\nthat our presentation of the Raman-related data caused confusion.\nWe fully understand the reviewer’s expectation for a more detailed\ninterpretation of Raman features across developmental stages. We also agree\nthat Raman band intensities and spectral shapes can be influenced by factors\nsuch as crystallinity, microfibril orientation, and cellular/background\ncontributions. We have tried our best to collect Raman spectra from 3 and 7\nDPA fibers. However, we noted that the spectra were complex and likely\ninfluenced by cellular/background contributions, making robust interpretation\nchallenging in our hands. Assigning and interpreting cellulose-related Raman\nfeatures across stages would require reference datasets about these factors,\nwhich are not yet well established for cotton fiber cell walls. In the previous\nversion, we therefore used Pearson correlation primarily to describe broad\nsimilarity trends across stages rather than as a direct measure of cellulose\ncontent.\nIn view of these complexities, and following the editor’s guidance, we have\nremoved the Raman dataset in its entirety from the manuscript and\nSupplementary Information, together with all Raman-related analyses and\nclaims. We have accordingly refocused the study on AFM-based mechanics,\ncellulose labeling, live-cell imaging and mathematical modeling. Importantly,\nthe central conclusions are unchanged and are now supported solely by these\nindependent lines of evidence."}]

Decision letter