Fragmentation and aggregation of cyanobacterial colonies

Reviewer #1 (Public review):

Sinzato et. al. investigated how shear flow in a rheological chamber affects the assembly and fragmentation of cyanobacterial aggregates, with the goal of understanding how such aggregates might form naturally, and/or be destroyed industrially. The authors used a combination of experiments and models to show that cyanobacterial colonies can be difficult to fragment with fluid flows. Additionally, they provide biophysical support for the idea that such aggregates likely form primarily when cells stay together after cell division, rather than coming together from disparate paths.

This work has significant relevance to the field, both practically and naturally. Combatting or preventing toxic cyanobacterial blooms is an active area of environmental research that offers a practical backbone for this manuscript's ideas. Additionally, the formation and behavior of cellular aggregates in general is of widespread interest in many fields, including marine and freshwater ecology, healthcare and antibiotic resistance research, biophysics, and microbial evolution. In this field, there are still outstanding questions regarding how microbial aggregates form into communities, including if and how they come together from separate places. Therefore, I believe that researchers from many distinct fields would find interest in the topic of this paper, and particularly Figure 5, in which a phase space that is meant to represent the different modes of aggregate formation and destruction is suggested, dependent on properties of the fluid flow and particle concentration.

Altogether, the authors were successful in their investigation, and I find their claims to be justified. In particular, the authors achieve strong results from their experiments. Below, I outline key claims of the paper and indicate the level to which they were supported by their data.

• Their first major claim is that fluid flows alone must be quite strong in order to fragment the cyanobacterial aggregates they have studied. With their rheological chamber, they explicitly show that energy dissipation rates must exceed "natural" conditions by multiple orders of magnitude in order to fragment lab strain colonies, and even higher to disrupt natural strains sampled from a nearby freshwater lake. This claim is well-supported by their experiments and data.

• The authors then claim that the fragmentation of aggregates due to fluid flows occurs primarily through erosion of small pieces from larger aggregates. Because their experimental setup does not allow them to directly observe this process (for example, by watching one aggregate break into pieces), they rely on indirect methods to support the claim. Overall, the experimental evidence is generally supportive, but the models leave some gaps. I describe this conclusion in more detail below.

• The strongest evidence for the erosion-dominated process comes from the authors' measurements of transfer of biomass between large and small size classes, as in Figure 2E and Figure 2D. The authors claim that only the erosion model can reproduce this kind of biomass transfer. However, it also seems that the idealized erosion model alone is not fully sufficient to capture the observed behavior. In Figure 2D, there remains a gap between their experiment and the prediction of the erosion model, which grows larger over time (Supplemental Figure S9). While the authors suggest that the erosion model is better than the equal-fragmentation model, it is also true that tracking the mean size (Figure 2B) or small size distribution (Figure S6) cannot distinguish between these models.

• Taken altogether, the experimental evidence favors an erosion-dominated process. However, a few minor questions remain regarding the models. Why does the equal-fragmentation model predict no biomass transfer between size classes? To what extent, quantitatively, does the erosion model outperform the equal fragments model at capturing the biomass size distributions? Finally, why does the idealized erosion fail to capture the size distribution at late stages in Supplemental Figure S9 - would this discrepancy be resolved if the authors considered individual colony variances in cell adhesion (for instance, as hypothesized by the authors in lines 133-137)? I do not believe these questions curb the other results of the paper.

• Their third major claim is that fluid flows only weakly cause cells to collide and adhere in a "coming together" process of aggregate formation. They test this claim in Figure 3, where they suspend single cells in their test chamber and stir them at moderate intensity, monitoring their size histogram. They show that the size histogram changes only slightly, indicating that aggregation is, by-and-large, not occurring at a high rate. Therefore, they lend support to the idea that cell aggregation likely does not initiate group formation in toxic cyanobacterial blooms. Additionally, they show that the median size of large colonies also does not change at moderate turbulent intensities. These results agree with previous studies (their own citation 25) indicating that aggregates in toxic blooms are clonal in nature. This is an important result, and well-supported by their data, but only for this specific particle concentration and stirring intensity. Later, in Figure 5 they show a much broader range of particle concentrations and energy dissipation rates that they leave untested. However, they refer to other literature that does test these regions of the phase map.

• The fourth major result of the manuscript is displayed in Equation 8 and Figure 5, where the authors derive an expression for the ratio between the rate of increase of a colony due to aggregation vs. the rate due to cell division. They then plot this line on a phase map, altering two physical parameters (concentration and fluid turbulence) to show under what conditions aggregation vs. cell division are more important for group formation. Because these results are derived from relatively simple biophysical considerations, they have the potential to be quite powerful and useful, and represent a significant conceptual advance. By combining their experiments with discussions of other experimental investigations of scum formation in cyanobacterial blooms, the authors have investigated the two most relevant zones of this map for the present study (Zones II and III), and have made a strong contribution to the literature in regards to artificial mixing to disrupt cyanobacterial blooms.

Other notes:

The authors rely heavily on size distributions to make the claims of their paper. I was pleased to find the calibration histograms in Supplemental Figure S8, which provide information as to how and why they made corrections to the histograms they observed. From these calibration histograms, it seems that larger colonies are more accurately measured in the cone-and-plate shear setup, while smaller colonies can be missed, presumably due to resolution issues.

Reviewer #2 (Public review):

Summary:

In this work, the authors investigate the role of fluid flow in shaping the colony size of a freshwater cyanobacterium Microcystis. To do so, they have created a novel assay by combining a rheometer with a bright field microscope. This allows them to exert precise shear forces on cyanobacterial cultures and field samples, and then quantify the effect of these shear forces on the colony size distribution. Shear force can affect the colony size in two ways: reducing size by fragmentation and increasing size by aggregation. They find limited aggregation at low shear rates, but high shear forces can create erosion-type fragmentation: colonies do not break in large pieces, but many small colonies are sheared off the large colonies. Overall, bacterial colonies from field samples seem to be more inert to shear than laboratory cultures, which the authors explain in terms of enhanced intercellular adhesion mediated by secreted polysaccharides.

Strengths:

• This study is timely, as cyanobacterial blooms are an increasing problem in freshwater lakes. They are expected to increase in frequency and severeness because of rising temperatures, and it is worthwhile learning how these blooms are formed. More generally, how physical aspects such as flow and shear influence colony formation is often overlooked, at least in part because of experimental challenges. Therefore, the method developed by the authors is useful and innovative, and I expect applications beyond the presented system here.

• A strong feature of this paper is the highly quantitative approach, combining theory with experiments, and the combination of laboratory experiments and field samples.

Weaknesses:

• Especially the introduction seems to imply that shear force is a very important parameter controlling colony formation. However, if one looks at the results this effect is overall rather modest, especially considering the shear forces that these bacterial colonies may experience in lakes. The main conclusion seems that not shear but bacterial adhesion is the most important factor in determining colony size. The writing could have done more justice to the fact that the importance of adhesion had been described elsewhere. This being said, the same method can be used to investigate systems where shear forces are biologically more relevant.

Author response:

The following is the authors’ response to the original reviews

Reviewer #1 (Public review):

(1) Their first major claim is that fluid flows alone must be quite strong in order to fragment the cyanobacterial aggregates they have studied. With their rheological chamber, they explicitly show that energy dissipation rates must exceed "natural" conditions by multiple orders of magnitude in order to fragment lab strain colonies, and even higher to disrupt natural strains sampled from a nearby freshwater lake. This claim is well-supported by their experiments and data.

We thank the reviewer for this positive comment. We fully agree, as our fragmentation experiments on division-formed colonies clearly demonstrate their strong mechanical resistance in naturally occurring flows.

(2) The authors then claim that the fragmentation of aggregates due to fluid flows occurs through erosion of small pieces. Because their experimental setup does not allow them to explicitly observe this process (for example, by watching one aggregate break into pieces), they implement an idealized model to show that the nature of the changes to the size histogram agrees with an erosion process. However, in Figure 2C there is a noticeable gap between their experiment and the prediction of their model. Additionally, in a similar experiment shown in Figure S6, the experiment cannot distinguish between an idealized erosion model and an alternative, an idealized binary fission model where aggregates split into equal halves. For these reasons, this claim is weakened.

The two idealized models of colony fragmentation, namely erosion of single cells and fragmentation into equal sizes (or binary fission), lead to distinguishable final size distributions. We believe that our experiments for division-formed colonies support the hypothesis of the erosion mechanism. Specifically, Figure 2E shows that colony fragmentation resulted in a decrease of large colonies and a strong increase of single cells and dimers (two cells). In our view, the strong increase of single cells and dimers provides quite convincing (but indirect) evidence supporting the erosion mechanism. This is described on lines 112-121. To further address the reviewer’s concern, we have included in the revised version of Figure 2 (panels B and D) a direct comparison between these two fragmentation models for large division-formed colonies fragmented at a high dissipation rate of ε = 5.8 m²/s³. Furthermore, we have included the new Supplementary Figure S9, which details the model predictions for the colony size distribution at various time points.

The ideal equal fragments model (i.e., where every fracture event produces two identical fragments with half the original biovolume) does not capture the biovolume transfer from large colonies to single cells, as observed for the experimental results in panel D of Figure 2 and panel E of Figure S9. In contrast, the erosion model, in panel D of Figure 2 and panel D of Figure S9, provides a good prediction of the experimental results within the experimental uncertainty. The different fragmentation models are discussed in lines 226-228 of the revised manuscript and lines 865-873 of the SI.

(3) Their third major claim is that fluid flows only weakly cause cells to collide and adhere in a "coming together" process of aggregate formation. They test this claim in Figure 3, where they suspend single cells in their test chamber and stir them at moderate intensity, monitoring their size histogram. They show that the size histogram changes only slightly, indicating that aggregation is, by and large, not occurring at a high rate. Therefore, they lend support to the idea that cell aggregation likely does not initiate group formation in toxic cyanobacterial blooms. Additionally, they show that the median size of large colonies also does not change at moderate turbulent intensities. These results agree with previous studies (their own citation 25) indicating that aggregates in toxic blooms are clonal in nature. This is an important result and well-supported by their data, but only for this specific particle concentration and stirring intensity. Later, in Figure 5 they show a much broader range of particle concentrations and energy dissipation rates that they leave untested.

We thank the reviewer for this positive comment. We agree that our experimental results show clear evidence that aggregated colonies have a weaker structure in comparison to division-formed colonies, thus supporting the hypothesis that clonal expansion is the main mechanism for colony formation under most natural settings. The range of energy dissipation rates of our experimental setup covers almost entirely the region for which aggregated and division-formed colonies differ in their fragmentation behavior (Zone III of Figure 5). Within this zone, aggregated colonies are fragmented and only the division-formed colonies are able to withstand the hydrodynamic stresses. Furthermore, we show that this fragmentation behavior has a low sensitivity to the total biovolume fraction, as displayed in the Supplementary Figures S2 and S4 and discussed in lines 151-154 and 160-163. We agree that our cone-and-plate setup covers a limited parameter range, and we have added a detailed discussion of these limitations in the revised manuscript, under section Materials and Methods in lines 462-473.

(4) The fourth major result of the manuscript is displayed in Equation 8 and Figure 5, where the authors derive an expression for the ratio between the rate of increase of a colony due to aggregation vs. the rate due to cell division. They then plot this line on a phase map, altering two physical parameters (concentration and fluid turbulence) to show under what conditions aggregation vs. cell division are more important for group formation. Because these results are derived from relatively simple biophysical considerations, they have the potential to be quite powerful and useful and represent a significant conceptual advance. However, there is a region of this phase map that the authors have left untested experimentally. The lowest energy dissipation rate that the authors tested in their experiment seemed to be \dot{epsilon}~1e-2 [m^2/s^3], and the highest particle concentration they tested was 5e-4, which means that the authors never tested Zone II of their phase map. Since this seems to be an important zone for toxic blooms (i.e. the "scum formation" zone), it seems the authors have missed an important opportunity to investigate this regime of high particle concentrations and relatively weak turbulent mixing.

We agree with the reviewer that Zone (II) of Figure 5 is of great importance to dense bloom formation under wind mixing and that this parameter range was not covered by our experiments using a cone-and-plate shear flow. The measuring range of our device was motivated by engineering applications such as artificial mixing of eutrophic lakes using bubble plumes, as well as preliminary experiments which demonstrated that high levels of dissipation rate were required to achieve fragmentation. The range of dissipation rates that can be achieved by the cone-and-plate setup is limited at the lower end by the accumulation of colonies near the stagnation point at the conical tip and at the upper end by the spillage of fluid out of the chamber. We now discuss this measuring range in lines 462-473 of the revised manuscript.

Although our setup does not cover Zone (II), we now refer to recent results in the literature for evidence of aggregation-dominance at Zone (II). The experimental study of Wu et al. (2024) (reference number 64 of the revised manuscript) investigated the formation of Microcystis surface scum layers in wind-mixed mesocosms. Their study identified aggregation of colonies in the scum layer, resulting in increases of colony size at rates faster than cell division. These results agree with our model, and the parameters range investigated fall within the Zone II. We have included in the revised version, lines 328-337, a detailed discussion elucidating the parameter range covered in our experiments and the findings of Wu et al. (2024).

Other items that could use more clarity:

(5) The authors rely heavily on size distributions to make the claims of their paper. Yet, how they generated those size distributions is not clearly shown in the text. Of primary concern, the authors used a correction function (Equation S1) to estimate the counts of different size classes in their image analysis pipeline. Yet, it is unclear how well this correction function actually performs, what kinds of errors it might produce, and how well it mapped to the calibration dataset the authors used to find the fit parameters.

We agree with the reviewer that more details of the correction function should be included. We have included in the revised version of the Supporting Information, in lines 785-796, a more detailed explanation of the correction function. Furthermore, a direct comparison of raw and corrected histograms of the size distribution and its associated uncertainty is presented in the new Supplementary Figure S8.

(6) Second, in their models they use a fractal dimension to estimate the number of cells in the group from the group radius, but the agreement between this fractal dimension fit and the data is not shown, so it is not clear how good an approximation this fractal dimension provides. This is especially important for their later derivation of the "aggregation-to-cell division" ratio (Equation 8)

We agree with the reviewer that more details on the estimation of fractal dimension are needed. The revised version, under Materials and Methods in lines 508-515, now includes the detailed estimation procedure, the number of colonies analysed, and the associated uncertainty.

Reviewer #1 (Recommendations For The Authors):

In light of the weak evidence for claim #2 outlined above, I believe the paper would benefit from a more explicit comparison in Figure 2C of the two models - idealized erosion, and idealized binary fission. With such a comparison, the authors would have stronger footing to claim that one process is more important than the other.

As mentioned in our answer above to comment #2 of public review, we have included in the revised version of Figure 2 (panels B and D) a direct comparison between the erosion and equal fragments (binary fission) models for large division-formed colonies fragmented under ε = 5.8 m²/s³. The comparison is further detailed in the new Supplementary Figure S9 for representative time points. Only the erosion models can recover the biovolume transfer from large colonies to single cells, as observed for the experimental results in Figure 2D and further detailed in Figure S9D. We believe that the revised version of Figure 2 and the new Supplementary Figure S9 provide strong evidence in support of the erosion fragmentation model.

Would the authors comment on their chosen range of experimental dissipation rates? For instance, was their goal more to investigate industrial/engineering applications where the goal is to disrupt the cyanobacteria, but not really typical natural conditions under which the groups might form?

The choice of experimental dissipation rates in our experiment was such that it covers engineering applications such as artificial mixing of eutrophic lakes using bubble plumes. We have now clarified in the Introduction, on lines 37-39, that artificial mixing has been successfully applied in several lakes to suppress cyanobacterial blooms. Furthermore, we have now clarified in the caption of Figure 5 that the bars on the right side indicate typical values of dissipation rates induced by natural wind-mixing, bubble plumes in artificially mixed lakes, and laboratory-scale experiments such as cone-and-plate systems and stirred tanks. The dissipation rates induced by the bubble plumes in artificially mixed lakes could potentially fragment aggregated cyanobacterial colonies and thus disrupt bloom formation. However, our preliminary experiments demonstrated that high levels of dissipation rate were required to achieve fragmentation, therefore we’ve focused on the upper range of values (0.01 to 10 m²/s³).

The dissipation rates generated by the cone-and-plate approach are indeed higher than the dissipation rates under typical natural conditions in lakes. We have now added a detailed discussion of the range of dissipation rates generated by the cone-and-plate approach in the revised manuscript, under section Materials and Methods in lines 462-473, where we also explain that these values are higher than the natural dissipation rates generated by wind action in lakes. However, the more generic insights obtained by our study, shown in Figure 5, are relevant for dissipation rates of natural lakes (e.g., Zone II). Therefore, in our discussion of Figure 5 we have now included the recent findings of Wu et al. (2024) (reference number [64] of the revised manuscript), who studied bloom formation of Microcystis in mesocosm experiments at dissipation rates representative of natural conditions; see also our reply to the next comment.

The authors should consider testing the space of Zone II on their phase map, for instance at very high particle concentrations and even lower rotational speeds, in order to show that their derivations match experiments.

Good point. As mentioned in our answer above to comment #4 of the public review, Zone II lies beyond the measuring range of our experimental setup. Instead, we refer to the recent study of Wu et al. (2024) (reference number [64] of the revised manuscript) which demonstrated that dense scum layers of Microcystis colonies are aggregation-dominated. These mesocosm experiments agree with our model predictions and their parameter range falls within Zone II. We have included in the revised version, lines 328-337, a detailed discussion where we elucidate the parameter range covered in our experiments and compare our predictions for Zone II with the recent findings of Wu et al. (2024).

The authors should show their calibration data and fit for the correction function of equation S1. Additionally, you may consider showing "raw" and "corrected" histograms of the size distribution, to demonstrate exactly what corrections are made.

As mentioned in our answer above to comment #5 of the public review, we have included in the revised version of the Supporting Information the new Supplementary Figure S8, which shows the raw and adjusted histograms of the size distribution, including the associated uncertainties. Furthermore, the correction function is now explained in detail in the new Supporting Information Text in lines 785-796.

The authors might consider commenting on Figure S3 a bit more in the main text. Even at very high dissipation rates, the cyanobacterial groups don't plummet to size 1, but stay in an equilibrium around 10-20x the diameter of a single cell. What might this mean for industrial applications trying to break up the groups?

We agree with the reviewer that further discussion of Figure S3, panels E and F, is warranted. In the revised version of the manuscript, under section Fragmentation of Microcystis colonies occurs through erosion in lines 133-137, we have now included a discussion of this figure. Figure S3F shows that more than 90% of the total biovolume ends up in the category “small colonies” (mostly single cells and dimers); hence, most of the initially large colonies do fragment to single cells or dimers. Only about 5-10% of the biovolume remains as “large colonies” of 10-20 cells. Although it is challenging to draw definitive conclusions about the behavior of these remaining large colonies, as they account for only a minor fraction of the suspension, one hypothesis is that variability in mechanical properties between colonies results in a subset of colonies exhibiting exceptional resistance even to very high dissipation rates (see lines 133-137).

Minor comments:

Typo Caption of Figure 2: Should read [m^2/s^3] for units

Thanks for catching this typo. The units in the caption of Figure 2 has been corrected to [m^2/s^3].

There is no Equation 10 in Materials and Methods as indicated in the rheology section.

We thank the reviewer for pointing out the lack of clarity in this algebraic manipulation. In fact, the yield stress has to be substituted in the current Equation 11 (previously Eq.10), from which the critical dissipation rate must be substituted in Equation 3. The result is the critical colony size (l* = 2.8) mentioned in line 243 of the revised manuscript. The correct equation numbers and algebraic substitutions are now indicated in lines 241-243 of the revised version of the manuscript.

<Reviewer #2 (Public review):

Especially the introduction seems to imply that shear force is a very important parameter controlling colony formation. However, if one looks at the results this effect is overall rather modest, especially considering the shear forces that these bacterial colonies may experience in lakes. The main conclusion seems that not shear but bacterial adhesion is the most important factor in determining colony size. As the importance of adhesion had been described elsewhere, it is not clear what this study reveals about cyanobacterial colonies that was not known before.

We would like to emphasize several key findings that our study reveals about the impacts of fluid flow on cyanobacterial colonies:

(I) Quantification of mechanical strength in cyanobacterial colonies: Our results demonstrate the high mechanical strength of cyanobacterial colonies, as evidenced by the requirement of high shear rates to achieve fragmentation. This is new knowledge, that was not known before for cyanobacterial colonies. To this end, our study highlights the resilience of these colonies against naturally occurring flows and bridges the gap between theoretical assumptions about colony strength and experimentally measured mechanical properties.

(II) The discovery that the mechanical strength of colonies differs between colonies formed by cell division and colonies formed by aggregation. This is again new knowledge, that was not known before for cyanobacterial colonies.

(III) Validation of a hypothesis regarding colony formation: Using a fluid-mechanical approach, we confirm the findings of recent genetic studies (references 25 and 67 of the revised version of the manuscript) which indicated that colony formation occurs predominantly via cell division rather than cell aggregation under natural conditions (except in very dense blooms).

(IV) Practical guidelines for cyanobacterial bloom control: Our findings provide valuable insights into the design of artificial mixing systems applied in several lakes. Artificial mixing of lakes is based on fundamentals of fluid flow, aiming at preventing aggregation of buoyant cyanobacteria in scum layers at the water surface. Our results show that the dissipation rates generated by bubble blumes in artificially mixed lakes can fragment cyanobacterial colonies formed by aggregation, but are not intense enough to cause fragmentation of division-formed colonies (see Figure 5 and lines 348-360).

The agreement between model and experiments is impressive, but the role of the fit parameters in achieving this agreement needs to be further clarified.

The influence of the fit parameters (namely the stickiness α1 and the pairs of colony strength parameters S1,q1,S2,q2) is discussed in the sections Dynamical changes in colony size modelled by a two-category distribution in lines 247-253 and Materials and Methods in lines 559-565. We kept the discussion concise to maintain readability. However, we agree with the reviewer that additional details about the importance of the fit parameters and the sensitivity of the results to these parameters could be beneficial. In the revised version of the section Materials and Methods in lines 560-563, we have included a detailed discussion of the fit parameters.

The article may not be very accessible for readers with a biology background. Overall, the presentation of the material can be improved by better describing their new method.

We apologize for the limited readability of the description of the experimental setup and model used. In the revised version of the manuscript and the SI, we have detailed further the new methods presented here. The modifications include a detailed description of the operating range of the cone-and-plate shear setup (subsection Cone-and-plate shear of the section Materials and Methods, in lines 462-473). Furthermore, we think that incorporation of the recent experimental results of Wu et al. (2024), on lines 331-337 of the manuscript, will appeal to readers with a biology background. Their mesocosm experiments support our model prediction that aggregation is the dominant mechanism for colony formation in region (II) of Figure 5.

Reviewer #2 (Recommendations For The Authors):

(1) The authors seem too modest in claiming technological advance. They should describe the technological advance of combining microscopy with rheometry, in such a way that this invites others to apply this or similar approaches on biological samples. Even though I feel that the advancement of knowledge of this system by their method is relatively modest, there may be more advances in other systems.

We appreciate the positive view of the reviewer towards the importance of this technology and we agree that its advantages should be advertised to researchers investigating similar systems. We have now given more attention to the technological advance of combining microscopic imaging with rheometry in the final paragraph of the Conclusions (lines 386400), where we now also briefly discuss an interesting recent study of marine snow (Song et al. 2023, Song and Rau 2022, reference numbers 70 and 71 of the revised manuscript), which used a similar combination of microscopy and rheometry as in our study. Furthermore, in the Methods section, we now briefly explain how the rheometry can be adjusted to investigate other systems (lines 474-480).

(2) It seems reasonable -also based on what we already know about these aggregates - to assume that the main difference in shear sensitivity between field samples and cultures lies in the production of extracellular polysaccharide substance (EPS). To go beyond what is already known, the study could try to provide more direct and quantitative evidence for EPS involvement. For example, using a chemical quantification of EPS levels, or perturbing EPS levels using digestive enzymes.

We agree with the reviewer that further characterization of the EPS is highly relevant to understand the mechanical strength of colonies. However, we believe that chemical quantification and/or degradation of EPS lies beyond the scope of our article and should be addressed by future studies.

(3) Assuming EPS is indeed the reason for the differences in shear resistance: the authors speculate the reason why the field samples have more EPS lies in chemical composition (Calcium/nitrogen levels). In addition, there could be grazing that is known to promote aggregation (possibly increasing EPS), or just inherent genetic differences between strains. I am not necessarily expecting the authors to explore this direction experimentally, but it seems certainly feasible and would make the final result less speculative.

We agree with the reviewer that there are more biotic and abiotic factors that can influence EPS amount and composition. The influence of grazing and other relevant factors on cell adhesion is discussed in references [26-29], cited in our introduction in lines 50-53. As discussed in our answer to recommendation #2, we believe that a quantitative investigation of these various factors is beyond the scope of this work and should be addressed in future studies.

(4) A cool finding seems to be the critical relative diameter (Fig 2E), a colony size that seems invariant under shear. I was slightly surprised that the authors seem to take little effort to understand this critical diameter mechanistically (for example by predicting it, or experimentally perturbing it). Again, not a necessary requirement, but this is where the study could harness its technological advantage to provide a more quantitative understanding of something that goes beyond the existing knowledge of the system.

We apologize to the reviewer if our descriptions and discussions of Figure 2 were unclear. One of the key conclusions from our experiments is that the critical relative diameter depends on the dissipation rate, as shown in Figure 2F. This dependence is also incorporated into the model through the constitutive equation (2). Furthermore, we expect the mechanical resistance of colonies, quantified by the critical relative diameter, to be affected by other biotic and abiotic factors that influence EPS amount and composition.

(5) The jump from 0.019 to 1.1 m²/s³ seems large. What was the reason for not exploring intermediate values? The authors should also define low, modest and intense dissipation rates more clearly. Currently, they seem somewhat arbitrarily defined, i.e. 0.019 m²/s³ is described as low (methods) and moderate (results). In Fig 2, the authors further talk about low dissipation rates without a quantitative description.

We thank the reviewer for pointing out the lack of clarity in the choice of parameter range and the nomenclature. Regarding the former, the suspension of division-formed colonies of Microcystis strain V163 displayed negligible fragmentation for dissipation rates between 0.019 to 1.1 m²/s³, as seen in Figures S2A and S3A. Due to the low sensitivity of the fragmentation results in this region, we don’t expect change in behavior for intermediate values. Regarding the nomenclature, we have corrected the inconsistencies throughout the text. We have chosen to name the dissipation rate values as: low for values typical of windmixing, moderate for values typical of the core of bubble plumes, and intense for values typical of propellers. Whenever mentioned in the text, the numerical value of dissipation rate is also included to avoid doubt.

(6.) The structure and narrative of the paper can be improved. The article first describes all lab culture experiments and then the model, while the first figure already shows model fits. Perhaps it would be better to first describe the aggregation experiments, to constrain the appropriate terms of the model, and then move to fragmentation.

We appreciate the recommendation of the reviewer regarding the structure. We have chosen to describe first the fragmentation experiments (Fig. 2), as these can be understood without introducing the aggregation effects. In contrast, the steady state results in the aggregation experiments (Fig. 3) come from the balance between aggregation and fragmentation. Therefore, we judged the current order to be more appropriate. The model fits are combined with the experimental results in Figures 2 and 3 to have a concise display. We have ensured that all the concepts required to understand each figure panel are explained prior to their discussion.

(7) The number of data points that go into the histogram needs to be indicated. The main reason is that the authors report the distribution in terms of the biovolume fraction, suggesting the numerical counts are converted into volume. This to me seems like the most sensible parameter, but I could not find how this conversion is calculated (my apologies if I missed it). This seems especially relevant because a single large colony can impact this histogram quite considerably.

We apologize for the lack of clarity in the calibration and conversion steps of the size distribution. As discussed above in the answer to comment #5 of the reviewer #1, more details of the calibration process have been added to the revised version of the Supporting Information Text in lines 785-796. Furthermore, the new Supplementary Figure S8 presents examples of the raw and adjusted size distribution, including the total number of counted colonies per histogram and the associated uncertainties in the concentration and biovolume distributions.

(8) Over the timescales measured here, colonies could start sinking (or floating), possibly in a size-dependent manner, that could lead to a bias due to boundary effects. Did the authors consider this potential artifact?

The sinking or floating of colonies is a relevant process which was taken into account in the choice of our parameter range for the dissipation rate. The minimum dissipation rate used in our experiments ensures that the upward inertial velocity near stagnation is sufficient to counteract the sedimentation of colonies. A detailed discussion of the choice of the parameter range is now included in the revised version of the Materials and Methods in lines 462-473.

(9) "On the one hand, sequencing of the genetic diversity within Microcystis colonies supports the hypothesis that colony formation undernatural conditions is primarily driven by cell division [25]. On the other hand, cell aggregation can occur on a shorter time scale and may offer improved protection against high grazing pressure [26]." This appears somewhat constructed, as what is described as "on the other hand" is not evidence against the genetic diversity.

We agree that the suggested dichotomy in this text appeared somewhat constructed, and we have now removed the wording “on the one hand” and “on the other hand”. The studies from reference [25] demonstrated that the genetic diversity between independent Microcystis colonies is much greater than the diversity within colonies. If cell aggregation was the dominant mechanism, a similar genetic diversity would be observed between and within colonies, which contrasts the findings from reference [25]. We have adjusted the text in the revised manuscript, in lines 46-54, to clarify this point.

(10) The phase diagram seems largely based on extrapolations that are made outside of the measurement regime (e.g. dark red bars indicating the dissipation rate, Fig 5 - by the way 1 this color scheme could use some better contrast, by the way 2 Fig S7 suggests a wider dissipation rate range as indicated in Fig 5, why?). Hence there seems to be the need to more clearly lineate experimental results, simulations, and extrapolations in the phase diagram.

We agree with the reviewer that further clarifications should be given about the parameter range covered in our experiments and apologize for the lack of readability in the color scheme of Fig 5. In lines 329-337, 346-347, 353-355, we have highlighted the parameters range covered by our experiments as well as the range covered by previous studies of windmixed mesocosm (namely reference [64] of the revised manuscript). Regarding the color scheme of Figure 5, we have modified the legend of the figure to improve readability. The color contrast was increased and leader lines were added to connect the colored bars with the respective label.

(11) Unfortunately, the manuscript did not contain line numbers.

We apologize to the reviewer for the lack of line numbers in our initial version. The revised version of the manuscript now contains line numbers, both in the main text and the supporting information.

(12) Fig 2D. Caption is too minimal. Y-axis could better be named "Fraction of colonies" as both small and large colonies are plotted.

The caption for Figure 2D was extended to better describe the plot. We have kept the y-axis label as “Fraction of small colonies”, since this is the quantity displayed by the three curves in the plot.

(13) An inset should have axis labels.

All the insets in our plots display the same variables as their respective plots. In order to keep the plots light and preserve readability, we therefore prefer to present the axis labels only along the x-axis and y-axis of the main plots, which implies by convention that the same axis labels also apply to the insets. To the best of our knowledge, this is a common approach.

(14) Page 5, first words. Likely Fig 3A, not 2A was meant.

We thank the reviewer for pointing out this readability issue. We intend to compare both Figures 2A and 3A. The text of the revised manuscript, in lines 146-148, has been adjusted with the correct figure numbers.

(15) Introduction, second last paragraph, third last line. "suspension leaded to a broad distribution" I assume you meant "... led to a ..."

We thank the reviewer for pointing out this typo. It has been corrected (line 122).