biology

Mechanism for Vipp 1 spiral formation, ring biogenesis, and membrane repair

The ESCRT-III-like protein Vipp1 couples filament polymerization with membrane remodeling. It assembles planar sheets as well as 3D rings and helical polymers, all implicated in mitigating plastid-associated membrane stress. The architecture of Vipp1 planar sheets and helical polymers remains unknown, as do the geometric changes required to transition between polymeric forms. *

In the article “Mechanism for Vipp1 spiral formation, ring biogenesis, and membrane repair” Souvik Naskar, Andrea Merino, Javier Espadas, Jayanti Singh, Aurelien Roux, Adai Colom and Harry H. Low show how cyanobacterial Vipp1 assembles into morphologically-related sheets and spirals on membranes in vitro.*

The spirals converge to form a central ring similar to those described in membrane budding. Cryo-EM structures of helical filaments reveal a close geometric relationship between Vipp1 helical and planar lattices. Moreover, the helical structures reveal how filaments twist—a process required for Vipp1, and likely other ESCRT-III filaments, to transition between planar and 3D architectures. *

Overall, the authors’ results provide a molecular model for Vipp1 ring biogenesis and a mechanism for Vipp1 membrane stabilization and repair, with implications for other ESCRT-III systems. *

NanoWorld Ultra-Short Cantilevers USC-F0.3-k0.3 for High-Speed AFM (HS-AFM) with a typical spring constant of 0.3 N nm−1 and a typical resonance frequency of about 300 kHz were used for image acquisition with fast scanning atomic force microscopy.*

Fig. 2 from Souvik Naskar et al. 2024 “Mechanism for Vipp1 spiral formation, ring biogenesis, and membrane repair”:Vipp1 assembles dynamic networks of spirals, rings and sheets on membrane a, F-AFM phase timecourse showing Vipp1 recruitment to the highly curved edge of membrane patches. Scan rate, 70 Hz; 256 × 256 pixels. The area in the dashed box is enlarged in b. b, Spiral and ring formation localized to the membrane edge. Scan rate, 70 Hz; 256 × 256 pixels. c, Left, phase timecourse showcasing a dense network of sheets, spirals, and rings that ultimately cover the entire membrane plane. Right, average of six F-AFM height images. Scan rate, 120 Hz; 256 × 256 pixels. d, Average F-AFM height image showing Vipp1 sheet, spiral, and ring detail. Red arrows mark the sheet branching into filaments ~13 nm wide. Scan rate, 20 Hz; 256 × 256 pixels. e, Vipp1 sheet and spiral filament height offset from the membrane. f–i, Quantification of Vipp1 filament and spiral characteristics. n = 124, 13, 278, and 278 independent measurements for panels f, g, h, and i, respectively. Error bars show one s.d. of the mean. NanoWorld Ultra-Short Cantilevers USC-F0.3-k0.3-10 with a typical spring constant of 0.3 N nm−1 and a typical resonance frequency of about 300 kHz were used for image acquisition. — Fig. 2 from Souvik Naskar et al. 2024 “Mechanism for Vipp1 spiral formation, ring biogenesis, and membrane repair”:
Vipp1 assembles dynamic networks of spirals, rings and sheets on membrane
a, F-AFM phase timecourse showing Vipp1 recruitment to the highly curved edge of membrane patches. Scan rate, 70 Hz; 256 × 256 pixels. The area in the dashed box is enlarged in b. b, Spiral and ring formation localized to the membrane edge. Scan rate, 70 Hz; 256 × 256 pixels. c, Left, phase timecourse showcasing a dense network of sheets, spirals, and rings that ultimately cover the entire membrane plane. Right, average of six F-AFM height images. Scan rate, 120 Hz; 256 × 256 pixels. d, Average F-AFM height image showing Vipp1 sheet, spiral, and ring detail. Red arrows mark the sheet branching into filaments ~13 nm wide. Scan rate, 20 Hz; 256 × 256 pixels. e, Vipp1 sheet and spiral filament height offset from the membrane. f–i, Quantification of Vipp1 filament and spiral characteristics. n = 124, 13, 278, and 278 independent measurements for panels f, g, h, and i, respectively. Error bars show one s.d. of the mean.

*Souvik Naskar, Andrea Merino, Javier Espadas, Jayanti Singh, Aurelien Roux, Adai Colom and Harry H. Low
Mechanism for Vipp1 spiral formation, ring biogenesis, and membrane repair
Nature Structural & Molecular Biology (2024)
DOI: https://doi.org/10.1038/s41594-024-01401-8

Open Access The article “Mechanism for Vipp1 spiral formation, ring biogenesis, and membrane repair” by Souvik Naskar, Andrea Merino, Javier Espadas, Jayanti Singh, Aurelien Roux, Adai Colom and Harry H. Low is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this license, visit http://creativecommons.org/licenses/by/4.0/.

Real-time multistep asymmetrical disassembly of nucleosomes and chromatosomes visualized by high-speed atomic force microscopy

During replication, expression, and repair of the eukaryotic genome, cellular machinery must access the DNA wrapped around histone proteins forming nucleosomes. These octameric protein·DNA complexes are modular, dynamic, and flexible and unwrap or disassemble either spontaneously or by the action of molecular motors. Thus, the mechanism of formation and regulation of subnucleosomal intermediates has gained attention genome-wide because it controls DNA accessibility.*

In the article “Real-Time Multistep Asymmetrical Disassembly of Nucleosomes and Chromatosomes Visualized by High-Speed Atomic Force Microscopy” Bibiana Onoa, César Díaz-Celis, Cristhian Cañari-Chumpitaz, Antony Lee and Carlos Bustamante describe how they imaged nucleosomes and their more compacted structure with the linker histone H1 (chromatosomes) using high-speed atomic force microscopy to visualize simultaneously the changes in the DNA and the histone core during their disassembly when deposited on mica.*

Furthermore, Bibiana Onoa et al. trained a neural network and developed an automatic algorithm to track molecular structural changes in real time. *

The authors’ results show that nucleosome disassembly is a sequential process involving asymmetrical stepwise dimer ejection events. The presence of H1 restricts DNA unwrapping, significantly increases the nucleosomal lifetime, and affects the pathway in which heterodimer asymmetrical dissociation occurs. *

Bibiana Onoa et al. observe that tetrasomes are resilient to disassembly and that the tetramer core (H3·H4)2 can diffuse along the nucleosome positioning sequence. Tetrasome mobility might be critical to the proper assembly of nucleosomes and can be relevant during nucleosomal transcription, as tetrasomes survive RNA polymerase passage. These findings are relevant to understanding nucleosome intrinsic dynamics and their modification by DNA-processing enzymes. *

To characterize the nucleosomes dynamics in 2D, individual molecules were observed in buffer using an Ando-type high speed atomic force microscope together with NanoWorld Ultra-Short Cantilevers for HS-AFM of the USC-F1.2-K0.15 AFM probe type ( typical spring constant 0.15 N/m, typical resonance frequency in air 1200 kHz, resonance frequency 500–600 kHz in liquid). *

The AFM data presented in the article allow the authors to directly visualize the dynamics of DNA and histones during nucleosome and chromatosome disassembly, providing a simultaneous observation of DNA unwrapping and histone dissociation. *

The experimental and analytical strategy presented shows that real-time HS-AFM is a robust and powerful tool for studying single nucleosomes and chromatin dynamics. *

graphical abstract from Bibiana Onoa et al 2024 "Real-Time Multistep Asymmetrical Disassembly of Nucleosomes and Chromatosomes Visualized by High-Speed Atomic Force Microscopy" - NanoWorld Ultra-Short Cantilevers of the USC-F1.2-k0.15 AFM probe type were used for the high-speed atomic force microscopy — graphical abstract from Bibiana Onoa et al 2024 “Real-Time Multistep Asymmetrical Disassembly of Nucleosomes and Chromatosomes Visualized by High-Speed Atomic Force Microscopy”

*Bibiana Onoa, César Díaz-Celis, Cristhian Cañari-Chumpitaz, Antony Lee and Carlos Bustamante
Real-Time Multistep Asymmetrical Disassembly of Nucleosomes and Chromatosomes Visualized by High-Speed Atomic Force Microscopy
ACS Central Science 2024, 10, 1, 122–137
DOI: https://doi.org/10.1021/acscentsci.3c00735

Open Access The article “Real-Time Multistep Asymmetrical Disassembly of Nucleosomes and Chromatosomes Visualized by High-Speed Atomic Force Microscopy” by Bibiana Onoa, César Díaz-Celis, Cristhian Cañari-Chumpitaz, Antony Lee and Carlos Bustamante is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this license, visit http://creativecommons.org/licenses/by/4.0/.

A beginner’s guide to the Characterization of Hydrogel Microarchitecture for Cellular Applications

Hydrogel materials show a number of properties which make them interesting candidates to be utilized to mimic the extracellular matrix (ECM). Therefore, these materials are attractive for use in biological applications such as tissue engineering, cell culture 3D bioprinting and more.

Are you planning to use hydrogels for the first time in your research?

Then have a look at the insightful article “A beginner’s guide to the Characterization of Hydrogel Microarchitecture for Cellular Applications” by Francisco Drusso Martinez-Garcia, Tony Fischer, Alexander Hayn, Claudia Tanja Mierke, Janette Kay Burgess and Martin Conrad Harmsen.

In their article the authors describe and evaluate the different technologies that are most commonly used to assess hydrogel microarchitecture.

Francisco Drusso Martinez-Garcia et al. explain the working principle of the various methods and also discuss the merits and limitations of each of them in view of their usefulness for the characterization of hydrogels.

They introduce and explore the pros and cons of the following methods: Scanning Electron Microscopy (SEM), Cryogenic Scanning Electron Microscopy (Cryo-SEM), Environmental Scanning Electron Microscopy (ESEM), Micro-Computed Tomography (µ-CT), Confocal Laser Scanning Microscopy (CLSM), Second Harmonic Generation and Atomic Force Microscopy (AFM).*

Atomic force microscopy (AFM) can be used to investigate the hydrogel surface topology as well as a hydrogel’s mechanical properties. The latter can be achieved through mathematical modelling of force-distance curves.

When using the AFM to characterize the elasticity of a hydrogel sample it is essential to take the stiffness of the investigated material into account when choosing what kind of AFM probe to use for these experiments.

If an AFM cantilever used for probing a soft sample is too stiff (if the force constant/spring constant is too high) this might result in a poor signal-to-noise ratio.

If a soft AFM probe (an AFM probe with an AFM cantilever with a low force constant) is chosen to investigate a soft material this should lead to a better signal-to-noise ratio. On the other hand, if an AFM cantilever is too soft (if the force constant is too low) then it might not be stiff enough to indent the investigated material.

Another critical factor is the shape and the size of the AFM tip.

Spheroidal AFM probes might stick to the material, resulting in artefacts, disrupted force–distance curves, or even damaged AFM cantilevers. If the AFM tip is much smaller than the pore size of the hydrogel, it might get stuck in the fibrous network microarchitecture.

On the other hand, if the spherical AFM tip, e.g. as in colloidal AFM probes (a sphere glued to end of a tipless AFM cantilever), is too large, the weight of the sphere can have a negative influence on the spring characteristics of the AFM cantilever.

All these factors and more as described in the cited article have to be carefully weighed before deciding on the settings of the atomic force microscope and choosing an AFM probe for the investigation of a specific hydrogel.

NanoWorld tipless ArrowTL2 cantilever arrays with polystyrene beads glued to them were used by the authors of this beginner’s guide to achieve the AFM data presented in the article.*

Figure 6. from Francisco Drusso Martinez-Garcia et al. 2022: Atomic force microscopy. (A) Equipment. (B) Schematic of an AFM setup with a four-quadrant photodiode (1), in which the four-quadrant photodiode (1) receives a laser (2) reflected from a cantilever (3), in this case positioned over a hydrogel (4) mounted in a piezo stage (5). For example, the height differences in a sample (4) are measured by adjusting the stage using piezo elements (5) to counter the cantilever bending on a nanometer scale. (C) The AFM can then generate a surface heightmap of the hydrogels such as a GelMA hydrogel (shown). AFM can also be used to determine the mechanical properties of hydrogels. (D) Schematic of the AFM technique to determine the elastic moduli of hydrogels with a tipless cantilever (1), spheroidal probe (2, red), hydrogel (3), and stiff substrate (4). As the cantilever represents a spring with a known spring constant, the cantilever bending due to elastic counterforces exerted by the soft material is correlated with the piezo stage height (4). (E) The so-called force–distance curves are recorded. Data from a collagen type-I hydrogel (3.0 g/L) are shown. (F) Young’s moduli of a 1.5 g/L and 3.0 g/L collagen type-I hydrogel. Outliers indicated by ◆. AFM equipment detailed in Appendix A of the cited article. NanoWorld tipless ArrowTL2 cantilever arrays with polystyrene beads glued to them were used by the authors of this beginner’s guide to achieve the AFM data presented in the article. — Figure 6. from Francisco Drusso Martinez-Garcia et al. 2022:
Atomic force microscopy. (A) Equipment. (B) Schematic of an AFM setup with a four-quadrant photodiode (1), in which the four-quadrant photodiode (1) receives a laser (2) reflected from a cantilever (3), in this case positioned over a hydrogel (4) mounted in a piezo stage (5). For example, the height differences in a sample (4) are measured by adjusting the stage using piezo elements (5) to counter the cantilever bending on a nanometer scale. (C) The AFM can then generate a surface heightmap of the hydrogels such as a GelMA hydrogel (shown). AFM can also be used to determine the mechanical properties of hydrogels. (D) Schematic of the AFM technique to determine the elastic moduli of hydrogels with a tipless cantilever (1), spheroidal probe (2, red), hydrogel (3), and stiff substrate (4). As the cantilever represents a spring with a known spring constant, the cantilever bending due to elastic counterforces exerted by the soft material is correlated with the piezo stage height (4). (E) The so-called force–distance curves are recorded. Data from a collagen type-I hydrogel (3.0 g/L) are shown. (F) Young’s moduli of a 1.5 g/L and 3.0 g/L collagen type-I hydrogel. Outliers indicated by ◆. AFM equipment detailed in Appendix A of the cited article.

*Francisco Drusso Martinez-Garcia, Tony Fischer, Alexander Hayn, Claudia Tanja Mierke, Janette Kay Burgess and Martin Conrad Harmsen
A Beginner’s Guide to the Characterization of Hydrogel Microarchitecture for Cellular Applications
Gels 2022, 8(9), 535
DOI: https://doi.org/10.3390/gels8090535

The article “A Beginner’s Guide to the Characterization of Hydrogel Microarchitecture for Cellular Applications” by Francisco Drusso Martinez-Garcia, Tony Fischer, Alexander Hayn, Claudia Tanja Mierke, Janette Kay Burgess and Martin Conrad Harmsen is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The images or other third-party material in this article are included in the article’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this license, visit https://creativecommons.org/licenses/by/4.0/.