Resolving interface structure and local internal mechanics of mitotic chromosomes

Domain: physics
Published: 2025-11-28
Editorial outcome: major_revision

Abstract
The interface of chromosomes enables them to interact with the cell environment and is crucial for their mechanical stability during mitosis. Here, we use Atomic Force Microscopy (AFM) to probe the interface and local micromechanics of the highly condensed and complex chromatin network of native human mitotic chromosomes. Our AFM images provide detailed snapshots of chromatin loops and Sister-Chromatids Intertwines. A scaling analysis of these images reveals that the chromatin surface has fractal nature. AFM-based Force Spectroscopy and microrheology further show that chromosomes can resist severe deformations, elastically recovering their initial shape following two characteristic timescales. Localized indentations over the chromatids reveal that the spatially varying micromechanics of the chromatin network is largely governed by chromatin density. Together, our AFM investigation provides insights into the structure and local mechanics of mitotic chromosomes, offering a toolbox for further characterization of complex biological structures, such as chromosomes, down to the nanoscale.

Introduction
During mitosis, the chromatin of eukaryotic cells undergoes a dramatic reorganization that results in the formation of highly condensed rod-like structures known as mitotic chromosomes1,2. In this process, chromosomes experience exogenous forces from the action of the mitotic spindle, which first pushes chromosomes towards the metaphase plate and then pulls the sister chromatids apart towards the spindle poles2–4. How chromosomes interact with their external environment and withstand these local forces is still unclear, even though this represents a key step for the successful completion of the mitotic cycle. In this context, AFM offers the unique possibility of accessing both the interface structure and the local micromechanics of mitotic chromosomes, thereby allowing a thorough characterization of their chromatin architecture.

Current structural models5 show that mitotic chromosomes consist of chromatin loops extruded from a central protein-rich scaffold where most of the condensin II protein complexes are located. Condensin I complexes further compact each condensin II-mediated loop into smaller loops. Numerous techniques were used to probe the whole structure of mitotic chromosomes6–10; however, the high susceptibility of chromosomes to external conditions11,12 and the soft yet very dense nature of their chromatin architecture8 still complicate access to smaller (sub-micron) features of the chromosome architecture and to their mechanics. Micropipette-based manipulation or optical tweezers (OTs) can be used to probe the macroscopic mechanical response of chromosomes to stretching deformations11,13–15 but are unable to access the local mechanics of the chromatin network at the nanoscale level. This knowledge gap can be bridged by exploiting the capabilities of Atomic Force Microscopy (AFM).

Indeed, AFM can be used to characterize the interface structure of mitotic chromosomes over the entire range of its length. In addition, AFM can be used to probe chromosome mechanics, via AFM-based Force Spectroscopy (AFM-FS)16, and microrheology, both with high spatial accuracy. All these measurements can be performed in a liquid environment, mimicking the native mitotic conditions. AFM has been applied to chromosomes before, though in most cases on fixed or dried samples. The few AFM studies characterizing chromosomes in liquid conditions have employed PBS17,18 or water as imaging solutions19, potentially altering both chromosome structure and mechanics due to the susceptibility of the chromatin network to variations in the physico-chemical properties of the environment11,12. Furthermore, most of these studies used chromosome spreads18–21, where chromosomes are covered by cytoplasmic debris that can only be removed via specific enzymatic treatments that could alter the delicate chromatin network22.

Here, we show how AFM can be used to resolve the interface structure and the internal mechanics of native human mitotic chromosomes in a liquid solution mimicking the physico-chemical properties of cells in mitosis. Our AFM images probe the chromosome interface over the entire range of scales of its structural hierarchy, revealing features like chromatin loops and Sister Chromatids Intertwines (SCIs). From the image analysis, we find that chromatin does not exhibit a specific higher-order organization and that the chromosome surface possesses a statistical fractal nature with self-similar scaling. Results from AFM-FS show that chromosomes can withstand and elastically recover from extremely high forces. Complementing these results with microrheology experiments, we find that the chromatin network relaxes on two prominent timescales. Finally, by performing localized indentations over the entire width of the chromatids we reveal that the local stiffness of the chromatin network is highly variable, both along a single chromosome and among different chromosomes. The variable micromechanics is mostly mediated by local chromatin density.

Results
AFM imaging of native mitotic chromosomes under physiological solution

The properties of the liquid environment in which mitotic chromosomes are probed play a crucial role on their structure and mechanics11,12. Here, isolated mitotic chromosomes were probed within a polyamine (PA) buffer that mimics the physico-chemical conditions of the mitotic environment in cells23 (“Methods”). In these conditions, mitotic chromosomes are fully hydrated and appear as a soft network of highly entangled chromatin fibers, displaying structural hierarchy at different length-scales (Fig. 1a–c). The sister chromatids of the chromosome reported in Fig. 1a, b, are in close contact (in agreement with previous studies24) and attached to each other at several points along their length by chromatin fibers (Fig. 1b). With our chromosomes, the sister chromatids are in close contact and attached to each other at several points along their length by chromatin fibers (Fig. 1b). Blue arrows in Fig. 1b point at several structures resembling chromatin loops stemming from inside the chromosome body; these features support the loop arrangement proposed by Gibcus et al.5, even though the overall chromatin network seems more disordered and randomly coiled than the predicted bottlebrush with a layered organization of loops. This is also supported by the presence of fibers spanning distances greater than the typical loop size (white arrows in Fig. 1b), some of them resembling inter-chromatids bridges25. Figure 1c highlights two different types of SCIs: those formed by individual chromatin fibers connected to both sister chromatids and those consisting of interlaced fibers forming loop structures (Fig. 1c and its schematic illustration in Fig. 1d). The presence of SCIs is in line with earlier studies demonstrating that also nocodazole-arrested yeast cells present these topological links26. Despite the high resolution of Fig. 1b, c, the displayed features appear not completely ‘sharp’; we interpret this observation as an indication that our imaging conditions still allow for some thermal motion of the soft chromatin architecture.

Chromatin organization and the fractal nature of chromosome surface

In the higher contrast, zoomed in image in the inset of Figs. 1e and S1, chromatin fibers and loops protruding outwards and forming a “halo” around the chromosome body can be observed. Similar structures have also been reported before19 and have been attributed to partial unraveling of the chromosome during the sample preparation and adsorption steps. In our images the “chromatin halo” appears approximately four times smaller, which might reflect the gentler sample preparation. In Fig. 1e a histogram describing the height distribution of the chromatin halo (Fig. S1) is shown. The histogram is fitted with a probability density function given by the sum of 5 Gaussian functions, where the first peak is located at approximately 8.5 nm; this height value is in agreement with previous studies6,27 and compatible with the “beads on a string” model for chromatin organization, describing irregularly arranged nucleosomes. The centers of the other Gaussian functions are located at 16.5, 31.3, 63.7, and 113.4 nm, all approximately multiples of the main 8.5 nm peak, probably describing the stacking of multiple chromatin fibers. The absence of prominent secondary peaks in the height distribution suggests that there is no preferential higher order organization for chromatin in the outer layers of the chromosome.

To gain more insights into the nature of the chromosome surface, we analyzed its roughness. A natural way to characterize the roughness of a random surface is to explore its fractal nature. This can be done by analyzing the roughness at different length-scales to test for self-affinity28,29. Briefly, we traced multiple height profiles, both parallel and perpendicular to the long axis of the chromosome (Fig. S2). Along each trace, we then measured the standard deviation of the height, σ, which characterizes the surface roughness of the chromosome (Fig. 1f). A regular non-fractal interface would exhibit a roughness that is independent of the length scale over which it is determined, provided this length scale exceeds the largest associated correlation length in the system. By contrast, for mitotic chromosomes plotting the standard deviation as a function of length-scale L (Figs. 1g, S3) reveals a power-law scaling regime σ ∝ Lα for length-scales from 40 to 250 nm, with α=0.6±0.1 (value from fit ± standard error) for longitudinal (Fig. 1g) and a similar α=0.6±0.1 (value from fit ± standard error) for perpendicular traces (Fig. S3). Such scaling relations indicate a non-trivial self-similarity that controls the roughness over a broad range of length scales.

Chromosomes elastically recover from repeated high-force deformations

Being able to access the native structure of mitotic chromosomes allows leveraging AFM-based Force Spectroscopy (AFM-FS) to probe local chromosome mechanics. AFM-FS employs higher forces than AFM imaging, potentially altering the chromatin architecture and its stability. To rule out such potential artifacts, we performed repeated indentations at the same location on the chromosome and monitored the variation of different structural and mechanical descriptors. Each AFM-FS measurement results in a force curve representing the force experienced by the tip as it penetrates and retracts from the sample16. In the representative force curve of Fig. 2a, the red cross indicates the contact point at which the AFM tip starts to push against the chromosome and can be interpreted as an indirect measure of the sample height. Figure 2b shows that after 30 repeated indentations (for 4 different chromosomes), the contact point (i.e., chromosome height) varies less than 5% from its initial value. This result rules out the introduction of any artifact from the probing process and highlights that chromosomes can withstand and elastically recover from local indentations extending deep through their entire body and reaching forces up to about 10 nN.

From the same force curves, it is possible to extract mechanical descriptors of the elastic and inelastic response of chromosomes to the applied deformations. The Young’s Modulus, which quantifies the relationship between the stress and the strain applied during the indentation process within the linear elastic regime, was obtained by fitting the initial part of the indenting force curve with a modified Hertz model30 (red dashed line in Fig. 2a). The inelastic mechanical response, on the other hand, is related to the energy dissipated during the indentation, due to viscous deformations. This can be described by the so-called viscoelasticity index, η31,32 (caption of Fig. 2a). A purely elastic response would result in η=0, while η=1 indicates a purely viscous response. The traces reported in Fig. 2c, d show that both the Young’s Modulus and viscoelasticity index displayed negligible variation during the four repeated-indentation series, indicating that chromosome mechanics is not affected by deformation history. Furthermore, the individual viscoelasticity index values (Fig. 2d), ranging from 0.5 to 0.8, indicate that, between the loading and unloading regimes, a fraction of the indentation energy is dissipated.

Chromatin is responsible for the fluid-like behavior of chromosomes

Further insights into the energy dissipation and time relaxation of the chromatin network can be obtained by leveraging AFM-based microrheology33. To this aim, we used a spherical probe with a radius of 870 nm (covering large portions of the chromosome body) to apply oscillatory deformations to the chromosomes. Figure 3a shows a representative microrheology force-time trace, where the tip is brought in contact with the chromosome and then oscillated at specific frequencies (“Methods”). Probing chromosome response to oscillatory deformations allowed for the calculation of the Shear Storage Modulus (G’) and the Shear Loss Modulus (G”), which give insights in the elastic and viscous responses of chromosomes at different deformation rates. Figure 3b, c report the values of G’ and G” obtained at different oscillation frequencies for 19 chromosomes. Although the values vary by almost an order of magnitude between different chromosomes, both G’ and G” show a highly consistent frequency dependence. Averaged data in Fig. 3d show that both G’ and G” increase with increasing frequency. G” has a stronger frequency dependence than G’, even though, within the probed frequency range, the elastic component (G’) remains dominant. This is also reflected by the frequency-dependent loss tangent (Fig. 3e), which is defined as the ratio between G” and G’ and quantifies the dissipative contributions in the sample (a loss tangent of zero indicates a completely elastic response). Furthermore, both G’ and G” exhibit a frequency dependence consistent with the high-frequency regime of the Rouse model for flexible chains34.

In the same figure, data obtained from the AFM (black trace) are compared with data obtained using OT15 (gray trace). For both methods, the loss tangent is dependent on the oscillation frequency; however, values obtained with AFM are almost one order of magnitude larger than those obtained with OT. This indicates that the chromosome behaves much more fluidly when deformed using AFM perpendicularly to its long axis than when stretched along the long axis using OT (Fig. 3f). Earlier, we have shown that the viscoelastic response of chromosomes along the long axis can be modeled to arise from an elastic central scaffold in parallel with an effective solution of entangled flexible fibers, representing chromatin loops dangling around the central scaffold11 (Fig. 3f). Based on this model, it is reasonable to assume that in our measurements the AFM tip is probing the same structures in series. Given the shallow indentation characterizing the Approach phase and the small amplitude of the oscillations (10 nm), the observed fluid-like behavior of the chromosome can then be understood as arising solely from the response of the entangled chromatin loops, with minimal contribution from the stiffer protein-rich scaffold2,5,35.

The chromatin network recovers from deformations following two characteristic timescales

Further analysis of the Force Relaxation Curves (FRCs, the Relaxation phase in Fig. 3a) can provide additional information on how the chromatin network relaxes from the applied deformations. As shown by previous studies36, FRCs can be fitted with a double exponential function:

F(t)=A1 e−t/τ1+A2 e−t/τ2+Fr (1)

Where A1, A2 are the amplitudes and τ1, τ2, the timescales of the fast and slow relaxation processes, respectively, and Fr=F(t→∞): Fitting the average FRCs of Fig. 3g yielded an average τ1 of 0.014±0.001 s and τ2 of 0.20±0.02 s (mean±SEM). Despite the variations in the values of the force drops (i.e., F(0)−Fr) among the probed (Table S1), neither τ1 nor τ2 showed any significant force-drop dependence (Fig. S4), suggesting that the initial deformation is within the linear elastic regime. Micropipette experiments have been used before to probe the force-relaxation response of mitotic chromosomes along their long axis (Fig. 3f), obtaining a characteristic timescale of 2 s13. The slower relaxation time during micropipette stretching could be indicative of larger structural rearrangements involving the entire chromosome body. On the other hand, the faster relaxation behavior observed with AFM most likely accounts for only minor re-arrangements of the chromatin architecture perpendicularly to the chromosome long axis. In the same micropipette study13 it was also found that the chromosome structure recovers the initial width (after being stretched) in approximately 0.05 s; this faster timescale is comparable with the value of τ1, which could hence describe the flow of liquid back into the chromatin network. The slower timescale, τ2, would then account for the structural reorganization of the chromatin network in the direction perpendicular to the long axis.

Chromatin density mediates chromatid’s mechanics

According to the nested loop model5 and electron-microscopy images37 the density of the chromatin network is not uniform, which could cause the micromechanical properties to vary within the chromatid cross-section. Figure 4a shows that the presence of both condensin I and II complexes in the central part of each chromatid9 results in a higher density of chromatin compared to the outer layers. The overlapping portions in between the two sister chromatids present additional chromatin entanglements and overlaps, likely produced by SCIs. To test whether these cross-chromatid structural heterogeneities translate into different mechanical responses, we performed indentations along lines perpendicular to the chromosome central axis (CCA, Methods), covering the entire chromosome, as indicated in Fig. 4b. For each indentation curve, we calculated the viscoelasticity index and Young’s Modulus, and plotted their value as a function of the distance between indentation point and CCA.

Figure 4c reports the values of viscoelasticity index recorded at different distances from the CCA (for 17 chromosomes). The dark green trace describes the average values of η when binned along the x-axis. The lower values of η in the outer parts of the chromatids suggest that chromatin-sparser areas respond more elastically to deformations. In contrast, the center of each chromatid is characterized by higher η values. This might be due to the higher chromatin and protein density of these areas, which generate a more entangled network that eventually takes longer to recover the initial structure. Finally, in the vicinity of the CCA, where the sister chromatids overlap, we recorded slightly lower η values, probably due to the presence of SCIs that increase the elasticity of the chromatin network. Taken together these results highlight that chromatin-dense regions take longer to relax from deformations compared to chromatin-sparse regions. Repeating the same characterization in selected regions along the chromatids (Fig. S5 and “Methods”) yielded very similar results (Figs. S6, S7a) for both centromeric and mid-arm regions (with only minor variations for the peripheral regions).

To investigate whether chromatin density also mediates chromosome stiffness, we calculated the local Young’s Modulus at different distances from the CCA. The scatterplot in Fig. 4d shows that regions located within 700 nm from the CCA (delimited by the dashed vertical gray lines and likely containing the central parts of the chromatids) are dramatically stiffer than the external regions, with local Young’s Moduli almost two orders of magnitude higher. The average Young’s Modulus recorded within 700 nm from the CCA is 290±10 kPa (mean±SEM), comparable with previous measurements on chromosome spreads rehydrated in PBS18. This result could reflect the higher density of chromatin and SCIs of these central regions. Selectively probing centromeric, mid-arm and peripheral regions yielded similar Young’s Moduli with the same two-orders of magnitude variation (Figs. S6, S7b).

The colored markers (blue and pink) in Fig. 4d identify the Young’s Modulus values for two representative chromosomes and highlight how the observed two-orders of magnitude variability can be mostly explained by inter-chromosome differences. In previous work, we found that the dissipation due to entangled chromatin loops scaled strongly with the chromosome density ρ11. Therefore, we wondered if these differences in Young’s Modulus could also be density dependent. As shown in Fig. 4e, we find that the Young’s Modulus scales with h (i.e., the contact point) as E ∝ h−2, and that differences in Young’s Modulus between chromosomes can mostly be attributed to their differences in height. Since differences in chromosome widths are relatively small and uncorrelated with differences in height (Fig. S8), we argue that h is mostly determined by how much each sample is pressed against the substrate and directly reflects the density: ρ ∝ h−1. Interestingly, our micromechanical results are consistent with the scaling E ∝ ρ2 for an entangled solution of flexible polymers38, providing further evidence that we are probing the response of entangled chromatin loops surrounding the central scaffold of the chromosome.

Methods
All the experiments were carried out in polyamine buffer (PA buffer) containing 15 mM TRIS (buffered at pH = 8), 2 mM EDTA, 80 mM KCl, 20 mM NaCl, 0.5 mM EGTA, 0.2% Tween 20, 0.5 mM spermidine and 0.2 mM spermine. The absence of magnesium-ions reduces enzymatic, which preserves the chromosomal structure and allows for storage of chromosomes40. The stabilizing function of the magnesium-ions is fulfilled by the similarly positively charged polyamines12. All buffers were made with ultrapure water (MilliQ, Millipore) and filtered (0.2 µm pore size, Whatman) before use.

Chromosome isolation

Cell culture and chromosome isolation were performed according to procedures already used and described by previous works15,41. Briefly, U2OS TRF1-BirA H2B-eGFP cells were cultured in high glucose GlutaMAX DMEM (Gibco), containing 10% FBS (Gibco) and penicillin-streptomycin (Gibco) at 37°C, 5% CO2. 48 h prior to chromosome isolation, the medium was supplemented with 12.2 µg/mL biotin. Metaphase arrest was achieved by incubating the cells with 50 ng/mL nocodazole for 4 h. Arrested cells were harvested via mitotic shake off, pelleted by centrifugation and washed twice with PBS. The mitotic cells were then incubated for 10 min at room temperature in a buffer consisting of 75 mM KCl and 5 mM Tris-HCl pH 8.0. Cells were then resuspended in polyamine buffer (see above) supplemented with Complete mini protease inhibitor and PhosSTOP phosphatase inhibitor (Roche). Using a dounce homogenizer with a tight-fitting pestle, the cells were lysed in ~25 strokes. The lysate was cleared by centrifugation and added to a glycerol step gradient (60% glycerol followed by 30% glycerol, both in polyamine buffer) and centrifuged at 1750 RCF for 30 minutes. The 60% glycerol fraction, containing the mitotic chromosomes, was stored at −20°C.

Surface and sample preparation for AFM experiments

All the AFM experiments were performed on poly-L-lysine (PLL)-coated glass coverslips. The coverslips were cleaned in a sonicator bath for 1 h in 2% (v/v) Hellmanex solution, followed by 30 minutes in milliQ water; after that, the coverslips were dried with an argon flow and stored in a sealed box. Before each experiment, the glass coverslips were activated with air plasma (Harrick Plasma Expanded Plasma Cleaner 115 V) for 3 min followed by immediate immersion in milliQ water for other 5 min. The activated slides were then incubated for 30 min in a 0.01% (w/v) PLL solution in 0.5 mM, pH 8.15 borate buffer (Na3BO3), at room temperature; at the end of the incubation the slides were thoroughly rinsed with milliQ water, dried with a gentle argon flow and placed in the AFM for the measurements. A 10 µl droplet of the chromosome preparation, diluted 1:10 in the PA buffer, was spotted on the glass coverslip and let to equilibrate for 20 min at room temperature before starting the experiments. All the AFM experiments (except for microrheology, see below) were performed on a Bruker Bioscope Catalyst coupled with an inverted optical microscope, at room temperature and using the PA buffer as imaging solution. The AFM was equipped with different probes depending on whether it was used for imaging or Force Spectroscopy (FS) experiments. Each chromosome was first located using the Nikon ECLIPSE Ti-U epi-fluorescence microscope, equipped with a 10x objective. After that, the AFM was used to scan the region of interest (ROI) and probe the chromosome with higher spatial accuracy and, in the case of AFM-FS, to run the mechanical characterization.

AFM imaging

Images were collected in Peak Force Tapping mode, using an Olympus BLAC40TS probe (spring constant 0.1 N/m, designed for high resolution measurements of biomolecules in liquid); the force setpoint and the other imaging parameters were tuned in order to accurately follow the topological features of the chromosome structure without introducing any deformation. In particular, the force setpoint was lowered until losing tip-sample contact; from this point, a small increase in the scanning force would guarantee the optimal imaging conditions, i.e., good tip-sample contact with minimal applied force. As also noted by Ushiki et al. in previous AFM works on chromosomes, the choice of a rectangular cantilever with a sharp tip having a high aspect ratio limited artifacts related to the shape of the chromosome42. Figures S9–S15 provide further evidence of the minimal influence of the scanning tip to the sample while imaging, in terms of both applied force and convolution effects. All the AFM images were processed using Gwyddion 2.5843.

Chromosome surface roughness analysis

The height profiles displayed in Fig. S2 were obtained using Gwyddion 2.58 and saved as .txt file for further analyses. For each profile, the average standard deviation is calculated over a rolling sampling window. The size of the sampling window converted in real units represents the length scale (L) under consideration. By repeating this procedure for different window sizes and by averaging the standard deviations obtained from all the profiles, we obtain the plot in Fig. 1f. A power-law fit (weighted on the variability of each value in the plot) is applied to the portion of the plots presenting 40 nm < L < 250 nm to yield the so-called scaling exponent. The entire procedure was executed using a custom Python script.

Chromatin halo height distribution

Gwyddion 2.5843 was used to extract the height data of the chromatin halo (masked area in Fig. S1). The height distribution was fitted with a sum of 5 Gaussian curves using a custom Matlab script.

AFM-force spectroscopy (AFM-FS)

DNPS10 probes (cantilever C, nominal tip radius 10 nm, nominal elastic constant 0.24 N/m, from Bruker, Camarillo, CA, USA), calibrated with the thermal noise method44 were used for all the AFM-FS measurements. Images at lower resolution were used to localize each chromosome in the center of the ROI; after that, multiple indentations (ramp size 1 µm and indentation rate 0.5 Hz) were performed on different parts of the chromosome. At the end of the indentations, a second image was performed to ensure that no permanent alterations were induced to the chromosome morphology. We note, however, that this happened very rarely. The recorded force curves were processed with custom made Python scripts to extract the information about the mechanical response of mitotic chromosomes. For each curve, the contact point was determined as the first data point along the indenting force curve (green trace in Fig. 2) at which a force greater than 50 pN was recorded (this threshold was chosen based on the signal-to-noise ratio that characterized the collected force curves). To ensure considering only indentations performed on the chromosome body, force curves presenting a contact point <150 nm were discarded from the analysis. For extracting the value of Young’s Modulus from each force curve, the modified Hertz fit from Dimitriadis et al.30 was applied to the indenting curve; more precisely, each curve was fitted from the contact point to an indentation depth equal to 30% of the chromosome thickness (estimated from the value of the contact point). The viscoelasticity index, η, is related to the energy dissipated during the indentation process and is defined as η=1−A2/A1, where A1 and A2 represent the areas under the indenting and retracting curves, calculated within the loading and unloading regimes (i.e., where the tip-sample interactions result in positive force values), respectively31,32. A custom Python script was used to calculate the areas under the indenting and retracting curves and to compute the values of η.

Cross-chromatid analysis

Series of indentations along lines distributed perpendicularly to the chromosome long axis and covering the entire chromosome, as described in Fig. 4b was performed on a total of 17 chromosomes. Indentations showing a contact point <150 nm were declared “non valid” and discarded from the analysis to ensure considering only indentations probing the actual chromosome body. For each indentation line, the coordinates of the middle point between the two most distal valid indentations were calculated; the chromosome central axis (CCA) was defined by all the middle points obtained via this procedure. The results shown in Fig. 4c, d report the results for all the valid indentations on all the probed chromosomes. The analysis was carried out using custom Python scripts.

Cross-chromatid analysis of centromeric, mid-arm and peripheral regions

We selected three different regions along the chromatids: the centromeric regions (referred as centromeric), the middle part of each arm (referred as mid-arm) and the most distal part of the arms (referred as peripheral), as shown in Fig. S5. Despite the length of the telomeres in chromosomes from U2OS cells can reach values >50 kb45, these values are still relatively small compared to the areas probed in the peripheral regions by the AFM tip; for this reason, we consider the indentations performed on the peripheral regions more representative of the subtelomeric structures46. For each region, we performed series of indentations along three lines running perpendicularly to the chromosome long axis (Fig. S5); a total of 21 chromosomes were probed. Mid-arm regions were selected at approximately half-distance between the centromere and each chromatid end. Due to diversity in chromosome arm’s length, it was possible to probe the arms on both sides of the centromere for only 6 chromosomes; for the other 15, only one side of the centromere presented arms that were long enough to enable a clear distinction between peripheral and centromeric regions. For all the 21 chromosomes it was possible to probe the peripheral regions (on both sides of the centromere) and the centromeric region. For each chromosome, the three different regions were located arbitrarily, since the instrumentation did not allow for a rigorous distance parametrization among the different chromosomes. Analysis of the indentation curves was performed using the same procedure described in the Cross-chromatids analysis, using custom Python scripts.

Comparing chromosome widths to heights

To determine whether there is any correlation between the width and the height of sampled chromosomes (the 17 chromosomes from the cross-chromatids analysis), we compared their mean widths and heights (Fig. S8). To obtain values for the mean width, three height profiles were traced for each chromosome in parts of the chromatids that exhibited a uniform width. For each profile, the width of the chromatids was defined as the width of the region higher than 150 nm, to discard the chromatin halo surrounding the chromosomes. The mean height values for each chromosome were obtained by averaging the contact point heights for all indentations performed within 200 nm from the CCA.

Young’s Modulus scaling with chromosome height

For determining whether the compaction of a chromosome affects its Young’s modulus, we used data from all measurement points within 200 nm from the CCA. For each measurement point, we plotted the Young’s modulus against the contact point height (Fig. 4e).

Microrheology measurements

Microrheology experiments were performed using Dynamic mechanical analysis (DMA) on a Bruker NanoWizard V BioAFM. Surface and sample preparations were identical to the ones used for the other AFM-FS experiments. A modified version of the PFQNM-LC-A-CAL probe, carrying a cylindrical tip with end radius of 870 nm and spring constant 0.137 N/m (Bruker, Camarillo, CA, USA) was used for the measurements. The commercially available version is now released as SAA-SPH−1UM (Bruker, Camarillo, CA, USA). The AFM setup was coupled with a fluorescence microscope that was used to locate the eGFP-labelled chromosomes on the surface. Once a single chromosome was localized within the ROI, multiple force curves were performed over the chromosome body. For these experiments, each measurement consisted of four different steps (see Fig. 3a): i) a shallow penetration of the sample until reaching a force of 150 pN, ii) a relaxation phase in which the tip was kept at a constant height and the relaxation force was monitored, iii) an oscillatory part where oscillatory deformations (with 10 nm amplitude) were applied to the sample and iv) a final retraction phase in which the tip was brought back to its initial position. For each chromosome, three different oscillatory frequencies were probed: 2, 20 and 200 Hz. Data were collected on 19 chromosomes. At least 15 force curves were collected from each chromosome for each oscillatory frequency. The FRC analysis was performed only on the dataset related to the 20 Hz oscillation frequency; this arbitrary decision is motivated by the fact that the parameters used in the “Approach” and “Relaxation” parts of the force curves (Fig. 3a) did not change when probing different oscillation frequencies. The data were analyzed combining the proprietary JPK-SPM Software (version 8.0.74) with custom Python scripts (for plotting and fitting purposes).

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "Authors use AFM technique to resolve the microstructure of the protein/DNA axis holding together two copies of mitotic chromosomes. The innovation of the work is in performing AFM in buffer that allows for keeping chromatin hydrated and closer to its condition inside the cell. With this approach, authors are able to resolve chromatin loops and intertwines. In addition, authors perform AFM force spectroscopy and microrheology studies. This part of the work confirms previously published studies and newly finds that Young’s modulus of the chromatin scales with its density. The manuscript is clearly written, all statements made by authors are well supported by experimental data. I recommend this paper for publication after minor corrections:\nLine 71: Authors state that polyamine buffer used for the chromosome preparation mimics intracellular conditions. However, I couldn’t find information on Mg content in the buffer despite the well-known fact that Mg2+ plays an important role in stabilizing DNA structure. Could authors clarify this detail?\nFig 1b: the features shown with white arrows may be an indication of the presence of “bridges” reported by Chu et al (Cell 2020). Though more accurate identification would require knowing the exact phase of mitosis.\nFig 1e: More details are needed to properly understand the figure. In particular, it is unclear what is plotted along y axis.\nLine 95: Could authors provide explanation for why the peak at 8.5 nm is compatible with beads-on-a-string model?\nFig 2a: The identing curve has a flat step. Is it reproducible? Could authors provide interpretation for this feature?\nFig 4: Could authors provide interpretation for the observed differences in height (and not width) of the chromosomes?", "reviewer_2": "In this paper, the authors employ a combination of atomic force microscopy (AFM) imaging and force spectroscopy to investigate the characteristics of chromosomes in a liquid environment that closely resembles in vivo conditions. Through high-resolution AFM characterization, the authors were able to identify several key structural features of chromosomes, including chromatin loops, different types of sister chromatid intertwine, and the surface roughness of chromosomes, as well as the presence of fractal structures. By employing AFM-based force spectroscopy and microrheology, the mechanical properties of chromosomes were quantified, and a correlation between these properties and the shape (density) of the chromosome was identified. These findings are highly beneficial for elucidating the mechanisms underlying chromosome folding, which represents a fundamental question in the field of life science. In conclusion, this paper is both interesting and meaningful. Nevertheless, there are several issues that require further attention before the manuscript can be considered for publication.\n1. As illustrated in figures 1a, b, and c, a network of highly entangled fibers can be observed on the surface of the chromosome, consistent with the description provided by the authors. However, the authors should provide a clear rationale for their consideration of these fibers as chromatin. Moreover, they should elucidate the criteria they employed to define the structure of loops and fibers not arranged in loops. As AFM is only capable of imaging the morphology of the sample surface, there is a possibility that the observed loop structure may in fact represent a hole within the fiber network. Similarly, the fibers not arranged in loops may represent one side of the loop exposed on the surface, with the loop vertically inserted into the chromosome (just as figure 1f shown). Additionally, it would be beneficial to understand the number of chromosomes imaged and whether the loop structures were observed in all the samples imaged, as well as the statistical size of the loops.\n2. It would be advantageous to ascertain the number of chromosomes investigated in order to determine the results of the fiber height measurement in the \"halo\" area. Furthermore, it would be beneficial to determine the data repeatability, as this could provide insight into the reliability of the results. It is important to note that during the process of AFM imaging, the imaging force always compresses the sample, which could potentially lead to a lower apparent height. This prompts the question of whether this effect may have influenced the results presented here.\n3. For the roughness measurements, how do the authors know that the loop is perpendicular to the surface? It is possible that the roughness is caused by the holes (loops) on the mesh surface. Another issue is that the imaging force from the AFM tip can deform the sample surface, which means that the imaging conditions will significantly affect the measured roughness. The authors must therefore explain how they have avoided this problem.\n4. As shown in Figure 4d, the author proposed that the central regions of the chromatids display greater stiffness than the external, lower regions. Figure 4e, however, depicts a contrasting trend between height and stiffness. In addition, the definition of the term \"h\" is unclear to me. It should also be noted that the gray dashed vertical lines in Figures 4c and 4d do not appear to align with ±700 nm, and the error bars appear to be incomplete."}, "author_response": "Response to Reviewers\nReviewer 1\nAuthors use AFM technique to resolve the microstructure of the protein/DNA axis holding\ntogether two copies of mitotic chromosomes. The innovation of the work is in performing\nAFM in buffer that allows for keeping chromatin hydrated and closer to its condition inside\nthe cell. With this approach, authors are able to resolve chromatin loops and intertwines. In\naddition, authors perform AFM force spectroscopy and microrheology studies. This part of the\nwork confirms previously published studies and newly finds that Young’s modulus of the\nchromatin scales with its density. The manuscript is clearly written, all statements made by\nauthors are well supported by experimental data. I recommend this paper for publication after\nminor corrections:\nWe thank the reviewer for their time and their supportive comments.\nComment 1\nLine 71: Authors state that polyamine buffer used for the chromosome preparation mimics\nintracellular conditions. However, I couldn’t find information on Mg content in the buffer\ndespite the well-known fact that Mg2+ plays an important role in stabilizing DNA structure.\nCould authors clarify this detail?\nResponse to comment 1\nIndeed, we used a buffer without any magnesium-ions, in order to reduce enzymantic activity,\nwhich preserves the chromosomal structure and allows for storage of chromosomes (Gasser,\nLaemmli, Exp. Cell. Res. 1987). The stabilizing function of the magnesium-ions is fullfilled by\nthe similarly positively charged polyamines.\nActions undertaken\nThe following sentence was added in the first section of Methods:\nThe absence of magnesium-ions reduces enzymatic, which preserves the chromosomal\nstructure and allows for storage of chromosomes (Gasser, Laemmli, Exp. Cell. Res. 1987). The\nstabilizing function of the magnesium-ions is fulfilled by the similarly positively charged\npolyamines (Beel, Andrew J., Pierre-Jean Matteï, and Roger D. Kornberg. \"Mitotic chromosome\ncondensation driven by a volume phase transition.\" bioRxiv (2021): 2021-07.).\nComment 2\nFig 1b: the features shown with white arrows may be an indication of the presence of\n“bridges” reported by Chu et al (Cell 2020). Though more accurate identification would require\nknowing the exact phase of mitosis.\nResponse to comment 2\nAs correctly noticed by the Reviewer, the linear strands observed in Figure 1b and Figure 1c\ncould indeed represent bridges; however, given the resolution of the image and the fact that\nthese non-looped structures were spotted not only in the connection site of the two\nchromatids, we cannot make any stronger statement about their identity. Nevertheless, we\nthank the Reviewer for pointing us to the work of Chu et al (Cell 2020), which we referenced\nat the end of the sentence as a possible interpretation for the non-looped structures located\nbetween sister chromatids.\nActions undertaken\nThe sentence:\nThis is also supported by the presence of fibers not arranged in loops and spanning distances\ngreater than the typical loop size (white arrows in Figure 1b).\nWas updated to:\nThis is also supported by the presence of fibers spanning distances greater than the typical\nloop size (white arrows in Figure 1b), some of them resembling inter-chromatids bridges (Chu\net al (Cell 2020)).\nComment 3\nFig 1e: More details are needed to properly understand the figure. In particular, it is unclear\nwhat is plotted along y axis.\nResponse to comment 3\nFigure 1e reports a histogram describing the height distribution of the chromatin halo\nhighlighted in Figure S1. The histogram is fitted with a probability density function, which is\nobtained by the sum of 5 Gaussian functions. The y axis of Figure 1e reports the values for the\nprobability density or “density” of the histogram. Using density as the statistical descriptor\nalong the histogram y-axis allows for fitting the histogram with a probability density function;\nthis wouldn’t be possible using counts or frequency as statistical descriptors for the y.-axis. We\napologize for not being specific enough about this result and thank the Reviewer for spotting\nthis.\nActions undertaken\nWe clarified the sentence describing the histogram and the related probability density\nfunction; in particular, the sentence:\nIn Figure 1e the height distribution of the chromatin halo (Figure S1) is shown. The\ndistribution is fitted with a sum of 5 Gaussian functions, where the first peak is located at\napproximately 8.5 nm\nWas changed to:\nIn Figure 1e a histogram describing the height distribution of the chromatin halo (Figure\nS1) is shown. The histogram is fitted with a probability density function given by the sum\nof 5 Gaussian functions, where the first peak is located at approximately 8.5 nm\nComment 4\nLine 95: Could authors provide explanation for why the peak at 8.5 nm is compatible with\nbeads-on-a-string model?\nResponse to comment 4\nOur claim is related to findings from Nishino et al. (already referenced in the sentence at line\n95), who performed a thorough X-rays scattering characterization of chromosomes and found\nregular chromatin structures associated with size values of 6 and 11 nm. They attribute these\nvalues to the irregular arrangement of nucleosomes into the so-called beads on string\nstructure. Given the lower accuracy (and statistics) of AFM compared to X-rays scattering, we\nbelieve that the observed 8.5 nm value is an average between the two populations found by\nNishino et al. but nevertheless compatible with describing irregularly arranged nucleosomes.\nWe thank the Reviewer for highlighting this issue and we edited the sentence to better convey\nour message.\nActions undertaken:\nThe sentence:\n… in agreement with previous studies6,25 and compatible with the “beads on a string” model\nfor chromatin organization.\nWas changed to:\n… this height value is in agreement with previous studies6,25 and compatible with the “beads\non a string” model for chromatin organization, describing irregularly arranged nucleosomes.\nComment 5\nFig 2a: The identing curve has a flat step. Is it reproducible? Could authors provide\ninterpretation for this feature?\nResponse to comment 5\nWe thank the Reviewer for pointing out this interesting aspect about the force curves.\nIn our indentation curves we often observe a flat region along the indenting curve, even though the\nextension and the force characterizing this plateau display some variability among the\nrecorded curves. A plateau along the indenting curve would describe an indentation at\nconstant force, meaning that the AFM tip is piercing through the chromatin network without\nexperiencing any stiffening of the underlying substrate. Given the variability in the extension\nand in the force at which we record these events, we are still not able to provide a thorough\nexplanation for their occurrence. A more systematic study is underway, where we aim to\nbetter understand what structures within the chromosome architecture might be responsible\nfor a specific mechanical response along the force curves.\nComment 6\nFig 4: Could authors provide interpretation for the observed differences in height (and not\nwidth) of the chromosomes?\nResponse to comment 6\nGiven that the chromosomes adsorb electrostatically to the substrate, their width is\nsomewhat bounded by the anchoring points. This explains why we see little variation of\nchromosome widths between samples. On the other hand, chromosome height can vary more\nfreely. We believe that the observed height differences can be explained by differences in the\ndensity of their chromatin network. As described by our mechanical characterization,\nchromosomes show a dramatic variation in their mechanical response (the Young’s Modulus\ncan vary up to two orders of magnitude between different chromosomes). The Young’s moduli\nscale with the inverse of the chromosome height, as expected if the chromosome density is\ninversely proportional to the chromosome height. It could be that chromosomes with a less\ndense chromatin network are able to relax more after adsorbing on the substrate, hence\nappearing thicker and softer than chromosomes with a denser and more entangled chromatin\nnetwork.\nReviewer 2\nIn this paper, the authors employ a combination of atomic force microscopy (AFM) imaging\nand force spectroscopy to investigate the characteristics of chromosomes in a liquid\nenvironment that closely resembles in vivo conditions. Through high-resolution AFM\ncharacterization, the authors were able to identify several key structural features of\nchromosomes, including chromatin loops, different types of sister chromatid intertwine, and\nthe surface roughness of chromosomes, as well as the presence of fractal structures. By\nemploying AFM-based force spectroscopy and microrheology, the mechanical properties of\nchromosomes were quantified, and a correlation between these properties and the shape\n(density) of the chromosome was identified. These findings are highly beneficial for\nelucidating the mechanisms underlying chromosome folding, which represents a fundamental\nquestion in the field of life science. In conclusion, this paper is both interesting and\nmeaningful. Nevertheless, there are several issues that require further attention before the\nmanuscript can be considered for publication.\nWe thank the reviewer for their efforts and for emphasizing the significance of our findings.\nComment 1\nAs illustrated in figures 1a, b, and c, a network of highly entangled fibers can be observed on\nthe surface of the chromosome, consistent with the description provided by the authors.\nHowever, the authors should provide a clear rationale for their consideration of these fibers\nas chromatin. Moreover, they should elucidate the criteria they employed to define the\nstructure of loops and fibers not arranged in loops. As AFM is only capable of imaging the\nmorphology of the sample surface, there is a possibility that the observed loop structure may\nin fact represent a hole within the fiber network. Similarly, the fibers not arranged in loops\nmay represent one side of the loop exposed on the surface, with the loop vertically inserted\ninto the chromosome (just as figure 1f shown). Additionally, it would be beneficial to\nunderstand the number of chromosomes imaged and whether the loop structures were\nobserved in all the samples imaged, as well as the statistical size of the loops.\nResponse to comment 1\nWe thank the Reviewer for the comment. We assumed the fibers to be chromatin on the basis\nof the height distribution in Fig. 1e. The main population of fibers has a peak at 8.5 nm which\nis compatible with regular chromatin structures as for example found by Nishino et al.\n(referenced in the sentence at line 95).\nNevertheless, as correctly pointed out by the Reviewer, AFM is only capable of imaging the\nmorphology of the sample surface; from this point of view, a fiber network presenting a hole\nin the plane perpendicular to the tip would appear as a hole surrounded by material in all\ndifferent directions. As indicated by the blue arrows in Figure 1b, we observe holes that\nappear to be surrounded by single fibers. Since the thickness of these fibers is consistent with\nthe thickness of chromatin (as we report in the height distribution of the halo of fibers, Figure\n1e), we interpret these holes surrounded by single fibers as chromatin loops. In addition,\nchromatin loops are expected to be the dominant structure in mitotic chromosomes and\nobservable by AFM. Hence, we are thus fairly confident about this interpretation.\nNonetheless, we have softened our language related to this conclusion to leave room for other\nexplanations.\nThe sentence:\nBlue arrows in Figure 1b point at several chromatin loops stemming from inside the\nchromosome body; these features support the loop arrangement proposed by Gibcus et\nal.\nWas changed to:\nBlue arrows in Figure 1b point at several structures resembling chromatin loops\nstemming from inside the chromosome body; these features support the loop\narrangement proposed by Gibcus et al.\nWe thank the Reviewer for the observation about the fibers “not arranged in loops”; after\nreflecting on the comment we agree that we cannot rule out the scenario of them still being\nparts of loops oriented perpendicular with respect to the ones identified by the blue arrows.\nWe still think that it is curious that these fibers span larger lengths compared to the other loop\nstructures. To account for this (and for the comment 2 made by Reviewer 1), we made the\nfollowing edits:\nThe sentence:\nThis is also supported by the presence of fibers not arranged in loops and spanning distances\ngreater than the typical loop size (white arrows in Figure 1b).\nWas changed to:\nThis is also supported by the presence of fibers spanning distances greater than the typical\nloop size (white arrows in Figure 1b), some of them resembling inter-chromatids bridges Chu\net al (Cell 2020).\nFinally, several other chromosome images also presented hints of an underlying organization\nwith fibers and loops. The image reported in the manuscript was the one presenting all the\nmentioned features in one image together with an exceptional resolution over the entire\nchromosome body. That said, statistics on loop size and structure would require more\nsystematic AFM imaging of many chromosomes. Thus, for now we consider the imaging\nprimarily as a demonstration of what can be seen using non-fixed chromosomes imaged in\nbuffer.\nActions undertaken\nWe further clarified in the main text that the descriptions of the features are referred to the\nchromosome displayed in Figure 1.\nThe following sentence:\nWith our chromosomes, the sister chromatids are in close contact and attached to each\nother at several points along their length by chromatin fibers (Figure 1b).\nWas changed to:\nThe sister chromatids of the chromosome reported in Figure 1 a and b, are in close contact (in\nagreement with previous studies, Wang, J. et al. ACS Nano 16, 8030–8039 (2022)) and\nattached to each other at several points along their length by chromatin fibers (Figure 1b).\nComment 2\nIt would be advantageous to ascertain the number of chromosomes investigated in order to\ndetermine the results of the fiber height measurement in the \"halo\" area. Furthermore, it\nwould be beneficial to determine the data repeatability, as this could provide insight into the\nreliability of the results. It is important to note that during the process of AFM imaging, the\nimaging force always compresses the sample, which could potentially lead to a lower apparent\nheight. This prompts the question of whether this effect may have influenced the results\npresented here.\nResponse to comment 2\nRegarding the results describing the halo surrounding the chromosome body, the analysis of\nother 3 separate AFM images of chromosomes yielded similar results, i.e., a height\ndistribution of the chromatin halo with a main peak around 10 nm. This further confirms\nfindings discussed in Figure 1e and note that the histogram reported in the main text accounts\nfor the chromosome showed in Figure 1a.\nThe Reviewer’s concern on the apparent lower height induced by the AFM tip to the scanned\nsamples is correct; however, this effect is limited by the recent AFM imaging technologies, like\nthe PeakForce tapping mode employed by our AFM. This imaging system allows scanning the\nsample while directly controlling the force exerted by the tip. Leveraging this feature, when\nimaging chromosomes we decreased the imaging force until losing the tip-sample contact;\nfrom this force value, a small increase in the scanning force would guarantee the optimal\nimaging conditions, i.e., good tip-sample contact with minimal applied force. This procedure\nalso minimizes lateral forces by the scanning tip on the sample. Evidence that the employed\nscanning force did not affect the chromosome structure (and hence height of the chromatin\nhalo) can be seen in the bottom part of Fig. 1a; here, the image looks blurrier because the tip\nlost the contact with the underlying sample, probably due to dangling chromatin fibers not\nfirmly attached to the chromosome body. In the case of the AFM tip actively pushing against\nthe sample this would not be possible.\nActions undertaken\nWe detailed the procedure for finding the optimal imaging force in the Methods section (AFM\nImaging), the following sentence was added to the text:\nIn particular, the force set point was lowered until losing tip-sample contact, from this point, a\nsmall increase in the scanning force would guarantee the optimal imaging conditions, i.e.,\ngood tip-sample contact with minimal applied force.\nComment 3\nFor the roughness measurements, how do the authors know that the loop is perpendicular to\nthe surface? It is possible that the roughness is caused by the holes (loops) on the mesh surface.\nAnother issue is that the imaging force from the AFM tip can deform the sample surface, which\nmeans that the imaging conditions will significantly affect the measured roughness. The authors\nmust therefore explain how they have avoided this problem.\nResponse to comment 3\nOur analysis of the chromosome roughness is independent of the structures that contribute\nto the surface heterogeneity. As correctly mentioned by the Reviewer both here and in\nComment 1, chromatin loops could be arranged along different planes, hence resulting in\nstructures of different shapes (from a topographical point of view). However, when calculating\nthe scaling behaviour of the height profiles in Figure 1g, we make no assumptions about the\norigins of the surface roughness. Rather than describing what the surface consists of, our\nanalysis shows that the chromosome surface is structured over many length scales. Our\nintention behind the cartoon in Figure 1f (defined in the caption as “illustrative”) was to\nschematically describe what the height profiles are measuring, but a real chromosome surface\nis of course more complex than a simple series of 2D loops all standing in the same plane.\nFor the last part of the Reviewer’s comment on potential deformations induced by the imaging\nconditions, we refer to our response for comment 2\nActions undertaken:\nThe caption of Figure 1f:\nf) Illustrative height profile (red trace) describing the height fluctuations of the chromosome\nsurface (gray structure).\nWas changed to:\nf) Illustrative height profile (red trace) describing the height fluctuations of the chromosome\nsurface (gray structure is a 2D simplified representation of loops making up the chromosome\nbody).\nComment 4\nAs shown in Figure 4d, the author proposed that the central regions of the chromatids display\ngreater stiffness than the external, lower regions. Figure 4e, however, depicts a contrasting\ntrend between height and stiffness. In addition, the definition of the term \"h\" is unclear to me.\nIt should also be noted that the gray dashed vertical lines in Figures 4c and 4d do not\nappear to align with ±700 nm, and the error bars appear to be incomplete.\nResponse to comment 4\nFigure 4 d shows how the Young’s Modulus (YM) of individual chromosomes varies as a\nfunction of the distance to the central chromosome axis (CCA). This plot shows that the YM\nvaries across the width of the chromosomes, potentially due to structural differences between\ncentral and peripheral regions of the chromosomes. However, we also find that near the\ncentral regions, there are large inter-chromosomal differences in the YM: the moduli can differ\nby two orders of magnitude.\nTo explain the inter-chromosomal YM variations near the central axis of the chromosomes,\nwe analyzed the YM as a function of the contact point height (Figure 4 e). To ensure we were\ncomparing data from structurally similar regions of the chromosomes, we only considered\npoints within 200 nm of the CCA (as mentioned in the “Young’s Modulus scaling with\nchromosome height” section of Methods). Figure 4 e shows that the large inter-chromosomal\ndifferences in the YM magnitudes (as already seen in Figure 4 d) can be largely attributed to\nthe height differences of the chromosomes.\nWith respect to the notation used in Figure 4 e, we point to the sentence in the main text\nstating: “As shown in Figure 4 e, we find that the Young’s Modulus scales with (i.e., the\ncontact point)”; the contact point is an indirect measure of chromosome height (as mentioned\nin the main text, when describing the results of Figure 3 b). We therefore think that the term\n(describing the chromosome height, measured from the contact point of the force curves)\nhas been already introduced thoroughly.\nFinally, we apologize for the mistake related to the position of the vertical grey lines and thank\nthe Reviewer for spotting this error. We have fixed the problem in the newest version of our\nmanuscript."}, {"round": 2, "reviewer_comments": {"reviewer_1": "The revised manuscript can be published as is.", "reviewer_2": "The authors have made significant efforts to address the majority of my previous concerns, and I appreciate their comprehensive responses and thoughtful revisions. The manuscript presents important insights into the structure and mechanics of mitotic chromosomes using AFM-based techniques, and the overall quality of the work is commendable. Nevertheless, I still have lingering concerns regarding the reproducibility of the key experimental observations, particularly those derived from AFM imaging.\nThe authors have noted that similar height distributions of chromatin halos have been observed in three additional chromosome samples (in response to Comment 2). However, no representative images or quantitative comparisons from these independent datasets have been presented. Given the inherently heterogeneous and soft nature of chromatin, and the fact that AFM imaging parameters (e.g., tip radius, applied force) can significantly influence the acquired data, it is critical to demonstrate that the structural features described—such as chromatin loops, intertwines, and fractal-like roughness—are robust across multiple conditions.\nIt is therefore strongly recommended that the authors include additional experimental data to validate the reproducibility of their findings. Specifically, the acquisition of AFM images of chromosomes using probes with varying tip radii is necessary to demonstrate that key structural features are not artifacts of tip convolution. In order to ensure the integrity of the observed surface features, it is imperative to provide AFM images acquired under varying imaging forces (within the limits of the PeakForce Tapping mode). This approach will confirm that the surface features remain unaltered or undamaged under different mechanical loads. If feasible, a quantitative comparison (e.g., height distribution, loop size statistics, or surface roughness metrics) across chromosomes imaged under these different conditions would enhance the rigor of the manuscript.\nAddressing these points would significantly strengthen the validity of the conclusions and alleviate concerns about the potential influence of AFM imaging parameters on the interpretation of chromosomal architecture."}, "author_response": "Response:\nAs requested, by the reviewer, we now report in the SI (and also below) 4 other\nindependent images (Figure S9 – S12) in which we recorded and measured the chromatin\nhalo around chromosomes. As can be seen from the height distributions of the chromatin\nhalos (Figure S13), the main peaks are located around 10 nm, which is in agreement with\nthe statements related to Fig.1. We do note that images from Figure S9 and Figure S10\nare referred to the same chromosome but were aquired from independent scans with\ndifferent scan size. Despite most imaging parameters remained unchanged, reducing the\nscanning size will inevitably cause the AFM tip to probe “more aggressively” since it will\ninteract more frequently with the sample; nevertheless, the two images showed\ncomparable height distribution, further confirming that the scanning force does not\nsignificantly altered the height of the chromatin halo.\nFigure S9: From top to bottom, i) an AFM image of a chromosome, ii) the high contrast version of the\noriginal image allows spotting a chromatin halo around the chromosome body; iii) the chromatin halo\nis masked for analysis of its height distribution.\nFigure S10: From top to bottom, i) AFM image of a chromosome, ii) the high contrast version of the\noriginal image allows spotting a chromatin halo around the chromosome body; iii) the chromatin halo\nis masked for analysis of its height distribution.\nFigure S11: From top to bottom, i) AFM image of a chromosome, ii) the high contrast version of the\noriginal image allows spotting a chromatin halo around the chromosome body; iii) the chromatin halo\nis masked for analysis of its height distribution.\nFigure S12: From top to bottom, i) AFM image of a chromosome, ii) the high contrast version of the\noriginal image allows spotting a chromatin halo around the chromosome body; iii) the chromatin halo\nis masked for analysis of its height distribution.\nFigure S13: Height distributions of the chromatin halos masked in Figures S9 – S12. For all the 4\nindependent images, the main peak of the chromatin distribution is located around 10nm.\nIn addition, to assure that the observed halo is not an effect of convolution from the tip\nscanning over the steep profile of the chromatids, we report both the “trace” and\n“retrace” channels for the high contrast image in Fig.1. When the AFM tip scans back and\nforth, two signals named “trace” and “retrace” are recorded, hence generating two\nindependent images, which will be affected by convolution in opposite directions (since\nthe scanning directions during retrace is opposite than the one during the trace). As can\nbe see from the comparison of the two images, the chromatin halo is very similar all\naround the chromosome, meaning that its xy shape does not derive nor it’s affected from\na convolution effect of the scanning tip.\nFigure S14: raw “trace” (left) and “retrace” (right) channels for the image of Figure 1. Despite the\nchromatids’ steep height profile being challenging for the AFM scanning tip, the chromatin halo of both\nimages remain fairly similar. Since the two images were recorded along opposite scanning directions,\nthis means that convolution (which affects only one scanning direction) has a minimal effect on the\nxy shape of the chromatin halo.\nThe same goes for the roughness exponent, we agree that changing imaging parameters,\nAFM tip etc.. will yield different exponent values, this for sure will need to be taken into\naccount in the case of a quantitative imaging analysis on multiple chromosomes. Here,\nwe just want to highlight that we obtained the same scaling exponent from both the\nroughness analyses on longitudinal and perpendicular traces (Fig. 1g, S2 and S3); this\ndoes not mean that the value of the exponent is universal for different chromsomes or in\nimages obtained with different parameters; however it supports the limited/absent effect\nof convolution and tip defects artifacts, since these effects would manifest only along a\npreferred direction.\nRelated to tip convolution effects, we now also report a comparison of the profile of one\nof the fibers visible in Fig.1 measured along the scanning direction and perpendicularly.\nSince the fiber makes an approx. 90° turn, it was possible to measure its width along both\ndirections. Tip convolution only affects features along the scanning direction, so the\nperpendicular direction is in principle convolution-free. As can be seen from the\nsimilarity between the two profiles, the convolution is minimal.\nThis was also one of the reasons that refrained us from measuring quantitative properties\nof fibers and loops on the xy plane. Since convolution is inevitably present (even with the\nsharpest tips), this would have to be taken into account in such analyses. It is also for this\nreason that the height distribution is a safer way to get information on chromatin\nproperties, since it is obtained from the z-direction which is not affected by convolution.\nFigure S15: Two perpendicular height profiles (left image) extracted from the same chromatin fiber\nmaking a roughly 90° turn. Profile 2 is traced parallel to the scanning direction and is in principle\naffected by convolution while profile 1 is perpendicular to the scanning direction and is not affected\nby convolution effects. On the right, the two height profiles are plotted and compared; despite being\nrecorded along perpendicular direction, their similarity supports confirms that convolution had\nminimal effect on the observed features.\nActions Undertaken:\nThe above images were also included in the SI of the manuscript to further support\nfindings from the AFM imaging. Moreover, we included more detailed information about\nthe used AFM tip in the “AFM Imaging” paragraph in the Methods, in particular:\nImages were collected in PeakForce Tapping mode, using an Olympus BLAC40TS probe\n(spring constant 0.1 N/m, designed for high resolution measurements of bio molecules\nin liquid)\n… As also noted by Ushiki et al. in previous AFM works on chromosomes, the choice of a\nrectangular cantilever with a sharp tip having a high aspect ratio limited artifacts related\nto the shape of the chromosome (Ushiki et al. Nanotechnology 19 (2008) 384022 (5pp)).\nFigures S9 to S15 provide further evidence of the minimal influence of the scanning tip to\nthe sample while imaging, in terms of both applied force and convolution effects. All the\nAFM images were processed using Gwyddion 2.58…\nAbout the point of using varying tip radii, we used high resolution tips with high aspect\nratio (i.e. among the highest quality tips currently commercially available); their\nsharpened top section has an higher aspect ratio than conventional pyramidal AFM tipsto\nreduce as much as possible convolution effects; moreover, we kept the imaging force as\nlow as possible in order not to introduce any damage to the chromosome. Imaging with\nblunter probes or with higher forces will inevitably result in worse image quality or in\nsample damages. Providing in the manuscript images affected by these issues seems to\nus not very suitable for supporting or contrasting our findings. We have now included in\nthe SI of the manuscript further checks to ensure that the presented data are not affected\nby any type of imaging artifacts, and we hope that this addresses the concerns of the\nreviewer.\nWith respect to the remark regarding the quantitative comparison across chromosomes,\nwe of course agree. However, for this manuscript we would like to note that we aimed to\ndemonstrate what can be obtained from imaging chromosomes in near native\nconditions. Thus, reporting the images and qualitatively describing features like fibers\nand loops was to provide a starting point for future quantitative and more thorough and\nsystematic imaging analyses on larger datasets. At the same time the main focus of this\nwork remains the quantitative mechanical characterization of chromosomes for which\nwe do provide ample statistics. We tried to make this even more clear in the manuscript\nnow.\nActions undertaken:\nIn the Discussion section, we added the following sentence:\nWe note that the data presented in this section are referred to the chromosome in Figure\n1. The aim is to provide a starting point for more thorough and systematic future imaging\nanalyses on larger chromosome datasets."}, {"round": 3, "reviewer_comments": {"reviewer_2": "It is evident that the authors have comprehensively addressed all of the concerns raised during the review process. The respondents' responses were both unequivocal and exhaustive, and the revised manuscript has been considerably enhanced as a consequence. It is my considered opinion that the current version of the paper is satisfactory and I have no further comments to make."}, "author_response": ""}]

Decision letter