Regulation of cell cycle progression by cell-cell and cell-matrix forces

Prior research has shown that cells respond to mechanical cues from their environment, including forces at cell-cell and cell-ECM interfaces. However, the specific temporal evolution of these forces during the cell cycle remains unclear. While it was already known that mechanical forces influence cell behavior, no prior work had resolved how these forces change dynamically throughout the cycle. This gap motivated the need to measure forces in a growing epithelium. Existing studies lacked direct quantification of tension and traction across the full cell cycle. No prior work had demonstrated how these forces might regulate transitions like G1 to S or mitotic rounding. That uncertainty drove the development of new tools to track mechanical states in real time. This study addresses the unresolved question of how forces evolve and whether they guide cell cycle progression.

The aim of this study was to investigate how mechanical forces at cell-cell and cell-ECM interfaces evolve during the cell cycle and whether these forces influence cycle progression. The researchers sought to determine if tension patterns correlate with key transitions like G1 to S and mitotic rounding. They also wanted to compare the predictive power of tension versus geometric properties for G1 duration. The study aimed to quantify forces in a growing epithelium to test the hypothesis that tension regulates cycle timing. The motivation stemmed from the lack of direct measurements linking mechanical states to cell cycle stages. The researchers hypothesized that tension changes could be a primary regulator of cycle progression. They aimed to test this using optogenetic tools to manipulate contractility. The study sought to establish a causal link between tension and cell cycle events.

The researchers used a growing epithelium to measure cell-cell tension and cell-ECM traction throughout the complete cell cycle. They employed high-resolution imaging to track mechanical states in a large cell population. Optogenetic tools were used to control contractility and test the effects of tension changes. The study combined live-cell imaging with mechanical force quantification. They tracked temporal patterns of tension across the cell cycle phases. The researchers compared tension levels with geometric properties like cell area. They analyzed how tension correlates with transitions such as G1 to S and mitotic rounding. The methods included statistical analysis to determine the predictive power of tension versus geometry for G1 duration.

Cells with higher intercellular tension showed a higher probability of transitioning from G1 to S and shorter G1 and S-G2-M phases. Tension levels increased during the cell cycle but dropped 3 hours before mitosis. The tension drop was linked to mitotic rounding through optogenetic manipulation. Tension and mechanical energy predicted G1 duration better than geometric properties. The study found that tension patterns span the entire cell cycle and regulate its timing. Mechanical forces at interfaces influence key transitions like G1 to S. The results show that tension is a better predictor of cycle progression than cell shape. The findings suggest that tension and contractility regulate mitotic rounding.

The authors propose that cell cycle progression is regulated by forces between dividing cells and their neighbors. They suggest that tension patterns influence cycle timing and key transitions. The study supports the idea that mechanical forces at interfaces regulate G1 to S and mitotic rounding. The results indicate that tension is a better predictor of G1 duration than geometric properties. The authors suggest that tension increases during the cycle but drops before mitosis. This drop facilitates mitotic rounding through contractility changes. The findings establish a causal link between tension and cycle progression. The study demonstrates that mechanical forces regulate the cell cycle cooperatively.