DNA serves as the carrier of genomic information, interacting with a wide variety of proteins to form chromatin during interphase. Before cell division, chromatin is compacted into thick, noodle-like chromosome structures, a process which plays a critical role in ensuring the equitable distribution of genetic material (Vagnarelli, 2013). This process is essential for maintaining genome integrity, as any errors in chromosome behavior during meiosis or mitosis can lead to conditions such as cancer, infertility, miscarriage, or congenital diseases (Potapova and Gorbsky, 2017; Theisen and Shaffer, 2010). Despite many decades of research, chromosome structure and organization are not yet understood.

Chromosome compaction during mitosis and meiosis is not a random process, but a tightly regulated sequence of events. In mammals, centimeters-long segments of DNA are compacted into chromosomes less than 10 μm long, via a multi-level process (Kschonsak and Haering, 2015). This process begins with the formation of nucleosomes, wherein DNA wraps around histone octamers, following the ‘beads on a string’ model (Alberts et al., 2002). Linker histone H1 binds to the entry and exit points of DNA on the nucleosome and interacts with the linker DNA region between nucleosomes.

However, the interaction between histones and DNA does not come close to explaining the full extent of chromosome compaction, indicating that other factors must be involved in this process (Wood and Earnshaw, 1990; Paulson et al., 2021; Batty and Gerlich, 2019). There are different models for chromosome organization, with the most popular being the ‘Scaffold/Radial-Loop’ model (Laemmli et al., 1978; Maeshima and Laemmli, 2003; Paulson and Laemmli, 1977; Earnshaw and Laemmli, 1983) and the ‘Chromatin Network’ model (Biggs et al., 2019). The Scaffold/Radial-Loop model proposes a continuous central protein core with chromatin loops stacked around it, based on electron microscopy observations. The Chromatin Network model suggests that structure proteins link chromatin segments without forming a continuous central core. MNase, an enzyme that degrades DNA/chromatin, ablated chromosome stiffness, in discord with a pure Scaffold/Radial-Loop model and instead indicating a key role for chromatin cross-bridges (Poirier and Marko, 2002). Experiments with DNases, which affect chromosome stiffness depending on their cutting frequencies, lend support to the Chromatin Network model (or equivalently, a chromosome radial looping model without a continuous proteinaceous scaffold and with some degree of crossbridging between adjacent loops).

Chromosome compaction and decompaction occur in different phases of the cell cycle (Antonin and Neumann, 2016), and the shape and size of chromosomes vary accordingly (Zickler and Kleckner, 2023). It is reasonable to hypothesize that chromosome stiffness also differs between stages, and between mitotic and meiotic chromosomes, which have similar shapes but experience different molecular events (Marko and Siggia, 1997; Koshland and Strunnikov, 1996). Chromosome stiffness measurements have shown that prophase I spermatocyte chromosomes are approximately 10 times stiffer than those in mitosis, indicating differences in chromosome organization between meiosis I and mitosis (Biggs et al., 2020).

During meiosis I, a unique railroad-track-like protein structure, known as the synaptonemal complex (SC), forms between the two homologous chromosomes (Heyting, 1996; Hollingsworth, 2020). Although the SC was thought to be a potential factor increasing stiffness of meiotic chromosomes, it has been observed that SYCP1, a key SC component, does not contribute to chromosome stiffness (Biggs et al., 2020). SYCP1 laterally connects the two ‘rails’ of the SC, axial elements (AEs), but may not impact the longitudinal stiffness. It is more likely AEs provide mechanical strength to the meiotic chromosomes longitudinally. Cohesin proteins, which connect sister chromatids in both mitosis and meiosis, are fundamental components of AEs (Peters et al., 2008; Ishiguro et al., 2011). In meiosis, meiosis-specific cohesin proteins, such as SMC1β, RAD21L, REC8, and STAG3, may play a role in chromosome structure and stiffness (Zheng and Xie, 2019; Rong et al., 2016; Gyuricza et al., 2016).

Aging can also induce significant changes in chromosomes, potentially leading to apoptosis, senescence, or cancer, all of which affect the lifespan and well-being of both animals and humans (Shammas, 2011; Faggioli et al., 2011). Moreover, chromosome-associated protein levels change with age, with some increasing while others decreasing (Thakur, 1983; Oliviero et al., 2022). For instance, cohesin proteins along chromosome axes decrease with age, potentially contributing to increased rates of aneuploidy and unsuccessful gamete production (Lister et al., 2010; Nakagawa and FitzHarris, 2017). Given the elevated aneuploidy rates in oocytes from older individuals, it is essential to explore how aging impacts chromosome mechanics and to gain a better understanding of the molecular mechanisms driving these changes.

In this study, we aspired to identify how meiotic cell cycle stage and aging influence meiotic chromosome stiffness. Prior studies found a large difference between mitotic and spermatocyte meiotic chromosome stiffness (Biggs et al., 2020). Spermatocyte meiotic prophase chromosomes were found to be 10 times stiffer than somatic chromosomes, which may be due to chromosome compaction or folding mechanisms specific to meiosis. During prophase I of meiosis, the SC zips up the two homologous chromosomes with components that are meiosis specific (Gordon et al., 2021). Previously, we found that SYCP1, an essential component of the SC that connects the lateral and central elements, does not contribute to the high prophase I chromosome stiffness in spermatocytes (Biggs et al., 2020), consistent with a ‘liquid crystal’ organizational scheme of the SC (Rog et al., 2017).

Here, we found that high chromosome stiffness also occurs in MI oocytes, which have a Young’s modulus approximately 10 times larger than that of mitotic chromosomes, five times larger than that of MII chromosomes, and approximately the same as that of prophase I chromosomes (Figure 2). We note that the Young’s modulus corrects for the varying thickness of chromosomes and indicates the elasticity in a volume- and geometry-independent way. However, the same trend of a large increase in chromosome stiffness during progression from somatic metaphase to meiotic prophase I and persisting through MI, followed by a reduction in chromosome stiffness upon progression to MII, is reflected in the force needed to double the length of chromosomes (Figure 2—figure supplement 1). This robust trend of a strong stiffening of chromosomes during prophase I and MI is in general accord with expectations from the chromosome compaction/relaxation and mechanics progression proposed by Kleckner et al., 2004. To further test Kleckner et al.’s model it would be desirable to measure the elasticity of somatic mitotic prophase chromosomes, but this has proven to be technically problematic in our attempts to date.

The SC central element dissociates from chromosomes before MI, in accord with our prior result that SYCP1 does not impart high meiotic chromosome stiffness (Biggs et al., 2020). We also measured chromosome stiffness in MII oocytes and found that it is significantly lower than in MI oocytes (in quantitative agreement with prior work Hornick et al., 2015, which measured a doubling force of MII chromosomes approximately double that of somatic mitotic chromosomes). Given the high and comparable stiffnesses of prophase I and MI chromosomes, we focused on factors specific to meiosis I, noting that AEs of the SC can still load onto chromosomes even in Sycp1^−/− spermatocytes which display WT prophase I chromosome elasticity (Biggs et al., 2020).

Cohesin proteins are components of the meiotic prophase chromosome axis, and load onto chromosomes during DNA replication. Some meiosis-specific cohesins, such as REC8, persist on chromosome arms until anaphase I. We hypothesized that cohesins rather than the central element might contribute to meiotic chromosome stiffness, at least through MI. However, comparing results from experiments on WT and cohesin-deficient (Rec8^−/−, Rad21l^−/−, and Stag3^−/−) spermatocytes, we found no difference in chromosome stiffness. We conclude that the meiosis-specific cohesins do not account for high stiffness in meiotic chromosomes. While we do not yet understand the molecular origin of the high stiffness of meiotic chromosomes, we hypothesize that factors present during prophase I and persisting through MI, drive high meiotic chromosome stiffness.

A limitation of the current experiments is that we were restricted to analyzing homozygous meiotic cohesin mutants in prophase I in spermatocytes. While it would be valuable to compare these results with prophase I oocytes, this is currently not a practical option since oocytes reach this stage in utero, making genotyping at that time highly challenging. It would have been preferable to carry out control experiments on heterozygous spermatocytes, but the quantitative lack of any difference in prophase I mechanics across the four genetic cases studied (Figure 3) strongly suggests that cohesin mutations do not substantially affect chromosome mechanics at prophase I.

Another limitation is that our mechanics experiments are extracellular, following removal of chromosomes from MI and MII oocytes, prophase I spermatocytes and cultured somatic cells. The extracellular environment (phosphate-buffered saline [PBS], or cell culture buffer for somatic cells) is different from the environment inside the cell, and one might hypothesize that this changes chromosome mechanics, for example, via loss of chromosome-folding proteins after their isolation. In addition to past experiments indicating that mitotic chromosomes are stable for long periods after their isolation (Pope et al., 2006), we carried out control experiments on mouse oocyte chromosomes where we incubated them for 1 hr in PBS, or exposed them to a flow of Triton X-100 solution for 10 min; there was no change in chromosome stiffness in either case (Methods and Figure 1—figure supplement 2). Across all our experiments we found quantitative agreement across trials under corresponding conditions, arguing against there being large uncontrolled changes in chromosome structure resulting from our isolation method.

Previous research on mitotic chromosome stiffness revealed that chromosomes have a chromatin network organization (Poirier and Marko, 2002). Several factors, including condensin, have been found to affect chromosome stiffness (Sun et al., 2018). Condensin exists in two distinct complexes, condensin I and condensin II, and both are active during meiosis. Published studies indicate that condensin II is more sharply defined and more closely associated with the chromosome axis from anaphase I to metaphase II (Lee et al., 2011). Additionally, condensin II appears to play a more significant role in mitotic chromosome mechanics compared to condensin I (Sun et al., 2018). Thus, condensin II likely contributes more significantly to meiotic chromosome stiffness than condensin I. It would be interesting to determine to what extent condensin defects affect meiotic chromosome structure.

Age also plays a role in altering chromosome stiffness. We found that chromosomes from aged MI oocytes had higher stiffness compared to those from younger oocytes, indicating that aging influences chromosome stiffness, in accord with a corresponding result for MII chromosomes (Hornick et al., 2015). This also provided further support for our conclusion that cohesins are not the primary factor making meiotic chromosomes stiffer as cohesin-protein levels reduce with age. The question of the molecular-level cause of oocyte meiotic chromosome stiffening with age remains open.

Hundreds of DNA double-stranded breaks are spontaneously introduced by the SPO11 protein at the onset of meiosis and aging also induces DNA damage (Carofiglio et al., 2013; Schumacher et al., 2021). How can meiotic chromosomes still become stiffer even in the presence of hundreds of DNA breaks in vivo? The answer may lie in the DNA repair proteins that are recruited to or near the DNA damage sites to repair the DNA damage and maintain genome integrity. Therefore, we hypothesized that DNA repair proteins contribute to meiotic chromosome stiffness. To test this hypothesis, we used a chemotherapy drug-etoposide to induce DNA damage in MI oocytes. Etoposide can increase the levels of TOP2–DNA covalent complexes, which generate DNA damage (Menendez et al., 2022). We found that etoposide treatment caused chromosome stiffness to decrease. This result is consistent with our previous study that DNA breaks can reduce chromosome stiffness in vitro (Poirier and Marko, 2002), reaffirming that meiotic chromosomes integrity relies on DNA continuity rather than the linkage of chromosome axis components (Biggs et al., 2019).

Since DNA repair proteins do not increase chromosome stiffness, other factors must be responsible for the high level of meiotic chromosome stiffness. During senescence, the amount of nuclear protein increases by around twofold, even though the cohesin level is decreased (Tsutsumi et al., 2014; De Cecco et al., 2011). Thus, it is possible that some nuclear proteins increase chromosome stiffness with age. However, further investigation is needed to determine which nuclear proteins contribute to chromosome stiffness. Additionally, histone methylation might influence chromosome stiffness because the level of histone methylation is proportional to chromosome stiffness (Biggs et al., 2019). Given that histone methylation levels are altered during aging, it is also plausible that some histone methyltransferases and demethylases regulate chromosome stiffness (Soriano-Tárraga et al., 2019; Wang et al., 2022). Moreover, histone methylation, especially H3K4, H3K9, and H3K36, plays a very important role in synapsis and recombination and has been found on meiotic chromatin from leptotene stage onward (Wang et al., 2017; Hochwagen and Marais, 2010; Powers et al., 2016). Further experiments could test the hypothesis that histone methylation regulates meiotic chromosome stiffness across different cell types.

No matter what factors are involved in stiffening meiotic chromosomes, they must alter chromosome organization to regulate chromosome stiffness. This indicates that chromosome stiffness is an important parameter for understanding chromosome structure. Defective chromosome organization is often related to various diseases, such as cancer, infertility, and senescence (Thompson and Compton, 2011; Harton and Tempest, 2012; He et al., 2018), and can be expected to cause changes in chromosome mechanics. By using micromanipulation experiments to study chromosome stiffness, we can better understand the mechanisms underlying these chromosome defects, potentially leading to new treatments and therapies.

A well-documented characteristic of aged oocytes is their higher rate of aneuploidy compared to that of younger oocytes (Selesniemi et al., 2011; Ma et al., 2020). The majority of aneuploidies can be traced to meiosis I, especially anaphase I, because it is an error-prone process (Kolano et al., 2012). It would be intriguing to investigate whether the increased chromosome stiffness contributes to this issue. A common cause of aneuploidy is lagging chromosomes during anaphase I (Godek and Compton, 2018). Chromosome stiffness may be related to the occurrence of lagging chromosomes, as chromosome separation relies on the pulling forces exerted by microtubules (Duro and Marston, 2015; Cohen-Fix, 2000). During anaphase I, homologous chromosomes, rather than sister chromatids, segregate. Thus, the chromosome region between the centromere and crossover must resist the tension applied by the spindle (Duro and Marston, 2015). Chromosomes must be stiff enough to prevent breakage under the pulling force. If the chromosomes are excessively stiff, the tightly connected homologous chromosomes may not properly separate, potentially causing aneuploidy. This suggests that chromosome stiffness change may be related to the high rate of aneuploidy observed in aged oocytes. Modulating factors altering chromosome stiffness may help reduce the incidence of aneuploidy. Moreover, further investigation into the relationship between chromosome stiffness and lagging chromosomes could provide insight into the mechanisms underlying aneuploidy in oocytes.

All data generated or analyzed during this study are included in the manuscript and supporting files.

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Author details

1. Ning Liu

   Department of Comparative Biosciences, University of Illinois at Urbana-Champaign, Urbana, United States

   Contribution

   Investigation, Methodology, Validation, Visualization, Writing – original draft

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0009-0002-3738-4004

2. Wenan Qiang

   1. The Chemistry of Life Processes Institute, Northwestern University, Evanston, United States
   2. Division of Reproductive Science in Medicine, Department of Obstetrics and Gynecology, Feinberg School of Medicine, Northwestern University, Chicago, United States

   Contribution

   Investigation, Methodology, Resources

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0001-5068-7128

3. Philip W Jordan

   1. Biochemistry and Molecular Biology Departments, Johns Hopkins University Bloomberg School of Public Health, Baltimore, United States
   2. Biochemistry and Molecular Biology Department, School of Medicine, Uniformed Services University of the Health Sciences, Bethesda, United States

   Contribution

   Methodology, Resources, Writing – review and editing

   Competing interests

   is on the scientific advisory board of Gameto, Inc. The opinions and assertions expressed herein are those of the author(s) and do not reflect the official policy or position of the Uniformed Services University of the Health Sciences or the Department of Defense

   "This ORCID iD identifies the author of this article:" 0000-0003-4890-2647

4. John F Marko

   1. Department of Molecular Biosciences, Northwestern University, Evanston, United States
   2. Department of Physics and Astronomy, Northwestern University, Evanston, United States

   Contribution

   Funding acquisition, Investigation, Methodology, Resources, Supervision, Validation, Writing – review and editing

   For correspondence

   john-marko@northwestern.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-4151-9530

5. Huanyu Qiao

   Department of Comparative Biosciences, University of Illinois at Urbana-Champaign, Urbana, United States

   Contribution

   Conceptualization, Funding acquisition, Investigation, Methodology, Project administration, Resources, Supervision, Validation, Visualization, Writing – original draft, Writing – review and editing

   For correspondence

   hqiao@illinois.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-0966-8077

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