Scientific statement
Passed research interest
2013-2018
My PhD project led me to investigate on the ultrastructure of nanoscopic membrane-lined channels called plasmodesmata, which are embedded in the cell walls between plant cells, allowing them to communicate with each other. They establish a very special kind of communication, proper to the plant kingdom, called symplastic communication, by establishing a cytosolic and membrane continuum of the plasma membrane (PM) and the endoplasmic reticulum (ER) between the cells. While they are central to plants and actively studied, the relation between ultrastructure and function is not fully understood. When I started my PhD, no information was available on the 3-dimensional organization of these pores and the dynamics of their membranes.
The ambition of my PhD project was to pioneer a new perspective on plasmodesmata pores by providing a detailed understanding of how they are organized within the narrow space between the ER and the PM and how this was related to the control of cell-to-cell connectivity. The ultrastructural analysis of plasmodesmata was a formidable challenge given their nanoscopic size (they can be as narrow as 20 nm in diameter), location (embedded in the plant cell wall) and their dynamic nature. And because electron tomography had not been developed yet on the Bordeaux Imaging Centre (BIC), I took a great part in establishing this technic from the ground up. Using electron tomography of high pressure-frozen, near-native plasmodesmata, I provided for the first-time data on their 3D membrane architecture at an unprecedented resolution, forcing a reconsideration of the current model.
2018-2022
As a young microscopist experienced in 3D Electron microscopy on plastic section and given the lack of work pushing forward cryo-electron tomography (cryo-ET) in the plant biology field, the next relevant step towards near-native imaging was cryo-ET. Hence, I was appointed at Caltech (Pasadena, USA) in January 2018 as a joint post-doctoral scholar in two labs, the Meyerowitz lab, specialized in apical meristem development (plant biology), and the Jensen lab, specialized in cryo-ET method development. At the interface between these two labs, I started a couple projects on the structural characterization cellulose synthesis.
The study of the arrangement of the cellulose fibers and their synthesis by the Cellulose Synthase Complexes in the plant cell wall by ECT
Because plant tissues are notoriously thick, cryo-EM requires thinning technics. Cryo-ET is only compatible with samples less than a micron thick at the most. Anything above requires thinning. The white onion (Allium cepa) abaxial epidermal cell wall peels biological system was chosen for its thickness (~10um) compatible with plunge freezing and FIB milling. It is also a biological model for the study of mechanical and structural properties of the plant cell wall. I generated ~100-200nm thick lamellae by FIB milling, then transferred in the cryo-transmission electron microscope (cryo-TEM). I then had access to the whole depth of the cell wall. Such process allowed the high-resolution imaging, in near-native conditions, of the cellulose fibrils and other components within the cell wall, revealing a striking bimodal angular distribution (see panel below) and a mysterious meshing bathing the fibers that gradually disappear at deeper depths of the cell wall.
This work won the 3rd place poster-award at the Southern California Cryo-EM symposium 2020 and the postdoctoral work award of the Microscopy and Microanalysis conference in July 2021. It was published in Current biology (https://www.sciencedirect.com/science/article/pii/S0960982222005930?via%3Dihub) and the protocol was published in Bioprotocol (https://en.bio-protocol.org/en/bpdetail?id=4559&type=0). The protocol paper made the cover of the issue.
The study of lipid droplet biogenesis in whole plant cells by cryo-ET
Despite the successful attempt at milling through onion cell walls, we did not have access to the full cells because the preparation methods involved stripping the cells away from the cell wall. My holy grail being looking inside whole plant cells, I started to investigate the type of cells that held the best chance to be compatible with plunge freezing and Gallium FIB milling. Pollen tube cells happen to be cylindrical cells 10 microns in diameter which is within the confines of plunge freezing and FIB milling. These cells emanate from the pollen grain and shuttle the male haploid nucleus from to the ovule deep in the carpel. I therefore set out to grow Arabidopsis pollen tubes from pollen grains, freeze them and FIB mill them. This allowed me to open 10-20um wide windows in different sections of the pollen tube showing intense vesicular trafficking. A subset of the data clearly displayed beautiful views of lipid droplet biogenesis so I set out to characterize the life cycle of lipid droplets in pollen tubes. This project relied on extensive use of convolutional neural networks to exhaustively segment the tomograms in 3D in order to gather as much structural/morphological information as possible on vesicles, lipid droplets and other organelles in the cell. This work has yet to be published but a snapshot of it is available in a poster presented at the Microscopy and Microanalysis conference in August 2021 (https://www.researchgate.net/publication/373449087_Cryo-electron_tomography_methods_applied_to_plant_science).
The study of the cellulose synthesis and crystallization process in prokaryotes
It is thought that plants acquired the ability to make cellulose through early endosymbiosis events with cyanobacteria. Therefore, understanding the mechanism of cellulose synthesis in bacteria is a good proxy to gain insights into understanding this process in plants. Among several bacteria that produce cellulose as a component of their biofilms, only one genus can make crystalline cellulose just like in plants, Gluconacetobacter. I thus started to study the structure of this bacterium by cryo-ET to better understand the cellular determinants of the cellulose crystallization process. This also includes elucidating the structure of its cellulose synthase machinery, namely the Acetobacter Cellulose Synthase (ACS), a multimeric complex that spans the double bacterial membrane. This aspect of my work has been granted funds from the Center for Environmental Microbial Interactions of Caltech in October 2018. This project has allowed the discovery of a novel, unknown prokaryotic cytoskeletal element we named the cortical belt.
The later piece of work has been published in Journal of Bacteriology in February 2021 and made the cover of volume 203, issue 3 (https://jb.asm.org/content/203/3.cover-expansion).
The study of the life cycle of predatory bacterium Bdellovibrio bacteriovorus
My extensive experience in tomography, tomogram analysis and segmentation led me to collaborate heavily in this very fun project spearheaded by Mohammed Kapan, also postdoc in the Jensen lab at the time (now Assistant Professor at University of Chicago (https://profiles.uchicago.edu/profiles/display/36292832). Several members of the Jensen lab, including Mohammed, amassed several hundreds of tomograms of Bdellovibrio bacteriovorus for multiple projects, all archived on the internal database of the Jensen lab (a subset of it was made public: https://etdb.caltech.edu/). This peculiar bacterium latches on its bacterial victims, enters their periplasm and eats them alive from the inside is still unclear. Its life cycle is not well understood and the massive database we had allowed us to piece back together the life cycle, in a still image style. This led to several unique observations, updating the molecular mechanisms involved in different steps of the life cycle:
· Hosts (victims) are detected by the use of pilli. Upon identification, the Bdellovibrio cell is brought closer to the host
· Upon contact with host, the bacterium creates a polar attachment plaque.
· Bdellovibrio’s flagellum is “reeled” inside the periplasm
· The outer-membrane of the host is digested, an air-tight seal is created between Bdellovibrio and host cell and the former penetrates the periplasm of the latter
· It will feed on the cytoplasm of the host cell until all cytoplasm is gone, divide and break loose, completing the life cycle.
This extensive work was published in Nature microbiology and made the cover of volume 8 issue 7 (https://www.nature.com/articles/s41564-023-01401-2).
Current research interest
2022-present
I took on a cryo-EM lab manager position in the Howard Hughes Medical Institute (HHMI) Gonen lab at UCLA (Los Angeles). My main duties are maintaining the facility, troubleshooting/fixing the equipment, coordinating with vendors and customer service for all consummables and training lab members and exterior users. I also have side projects, among which developing accurate fluorescence-guided FIB milling methods that I then apply to difficult to mill membrane protein crystals that grow in a tooth-paste like matrix called Lipidic-Cubic Phase (LCP). Because of this new interest in crystals and micro-ED, combined with my interest in cell-biology, I have a couple of projects that have to do with biogenic-crystals (crystals naturally made by cells that hold a biological function). More to come on this...
Development of efficient fluorescence-guided FIB milling methodologies ("FIBucial" method)
In the Gonen lab, a lot of focus is put on solving structures of membrane protein crystals that require special growth solvants and conditions. Namely, several require the use of a tooth-paste like substance called lipidic cubic phase (LCP). When LCP matrix loaded with crystals is loaded on a grid, it looks like amorphous piles ressembling hills or mountains (sometimes several hundreds or nanometers high). It is impossible to see where the crystals are, and it can take a lot of trial and error to make a lamella right through the crystal.
We therefore use fluorescence guided plasma-FIB milling to target as best we can the crystals. While the X-Y correlation is quite straightforward and could be done by eye, the Z correlation, taking into account the different imaging perspectives of the SEM, FIB and our in-chamber fluorescence light microscope (iFLM), is impossible to do by eye.
When I arrived into the lab, the process of x,y,z correlation was not streamlined at all and the success rate was low. Additionally, the process took time, not because of milling time but correlation time. I thus setout to establish the best correlation pipeline to do the correlation on-the-fly, allowing for rapid targeting. Correlation methods often require landmarks, or fiducial markers that are electron-dense fluorescent spheres that can be spotted in all imaging modalities. Unfortunately, LCP is very hard to mix with aqueous fiducial marker mixes. I developed a method where the fiducial markers are directly “drilled” into the sample. I call them “FIBucials” and it allows quick x,y,z registration of the targets of interests allowing to focus the session time on the milling.
Although the streamlining of this method is still ongoing, it was used in two published manuscripts in Nature Communications (https://www.nature.com/articles/s41467-023-36733-4) and IUCrJ (https://journals.iucr.org/m/issues/2023/04/00/fq5021/index.html).
A focused paper on the FIBucial method is on its way.
De-novo structure of Vasopressin-1 receptor (V1B) by micro-ED
Building on the pipeline I setup and am still optimizing, I am actively collaborating in solving the structure of the human Vasopressin-1 B (V1b) receptor. Michael Martynowicz and Anna Shiriaeva provide me with the grids with the LCP loaded with V1b crystals. V1b is a G-Protein Coupled Receptor (GPCR) central in the regulation of vasoconstriction, which controls blood pressure and osmotic homeostasis, which balances the body's water and electrolyte levels. This work has been published as a preprint (https://www.biorxiv.org/content/10.1101/2023.07.05.547888v1.abstract) and has yet to be published.
