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Cell lipid membranes are the site of vital biological processes, such as motility, trafficking, and sensing, many of which involve mechanical forces. Elucidating the interplay between such bioprocesses and mechanical forces requires the use of tools that apply and measure piconewton-level forces, e.g., optical tweezers. Here, we introduce the combination of optical tweezers with free-standing lipid bilayers, which are fully accessible on both sides of the membrane. In the vicinity of the lipid bilayer, optical trapping would normally be impossible due to optical distortions caused by pockets of the solvent trapped within the membrane. We solve this by drastically reducing the size of these pockets via tuning of the solvent and flow cell material. In the resulting flow cells, lipid nanotubes are straightforwardly pushed or pulled and reach lengths above half a millimeter. Moreover, the controlled pushing of a lipid nanotube with an optically trapped bead provides an accurate and direct measurement of important mechanical properties. In particular, we measure the membrane tension of a free-standing membrane composed of a mixture of dioleoylphosphatidylcholine (DOPC) and dipalmitoylphosphatidylcholine (DPPC) to be 4.6 × 10-6 N/m. We demonstrate the potential of the platform for biophysical studies by inserting the cell-penetrating trans-activator of transcription (TAT) peptide in the lipid membrane. The interactions between the TAT peptide and the membrane are found to decrease the value of the membrane tension to 2.1 × 10-6 N/m. This method is also fully compatible with electrophysiological measurements and presents new possibilities for the study of membrane mechanics and the creation of artificial lipid tube networks of great importance in intra- and intercellular communication.
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Cell lipid membranes are the site of vital biological processes, such as motility, trafficking, and sensing, many of which involve mechanical forces. Elucidating the interplay between such bioprocesses and mechanical forces requires the use of tools that apply and measure piconewton-level forces, e.g., optical tweezers. Here, we introduce the combination of optical tweezers with free-standing lipid bilayers, which are fully accessible on both sides of the membrane. In the vicinity of the lipid bilayer, optical trapping would normally be impossible due to optical distortions caused by pockets of the solvent trapped within the membrane. We solve this by drastically reducing the size of these pockets via tuning of the solvent and flow cell material. In the resulting flow cells, lipid nanotubes are straightforwardly pushed or pulled and reach lengths above half a millimeter. Moreover, the controlled pushing of a lipid nanotube with an optically trapped bead provides an accurate and direct measurement of important mechanical properties. In particular, we measure the membrane tension of a free-standing membrane composed of a mixture of dioleoylphosphatidylcholine (DOPC) and dipalmitoylphosphatidylcholine (DPPC) to be 4.6 × 10-6 N/m. We demonstrate the potential of the platform for biophysical studies by inserting the cell-penetrating trans-activator of transcription (TAT) peptide in the lipid membrane. The interactions between the TAT peptide and the membrane are found to decrease the value of the membrane tension to 2.1 × 10-6 N/m. This method is also fully compatible with electrophysiological measurements and presents new possibilities for the study of membrane mechanics and the creation of artificial lipid tube networks of great importance in intra- and intercellular communication.
This thesis presents a reliable technology to assemble free standing lipid membranes using microfabricated devices. A microfluidic cartridge consisting of parallel channels connected with a rectangular aperture was designed and characterized to assemble artificial membranes. This methodology resulted on a system capable of assembling lipid bilayer membranes of different lipid composition. Using decane as organic solvent, ~70% of the aperture was covered by the lipid bilayer, while the remaining are was occupied a pocket of solvent (annulus). An almost complete depletion of the annulus can be achieved by choosing a solvent (chloroform) capable of being absorbed by the flowcell material. In comparison with others methods, this approach is an important contribution to the field as it is allows real-time control over conditions (voltage, molecules in solution, pH) over both leaflets of the membrane. Furthermore, the lipid bilayer plane is perpendicular to the microscope focal plane, allowing observation of morphological changes in the lipid membrane and straightforward combination with optical techniques. This work shows the first successful operation of optical tweezers combined with planar lipid membranes accessible from both sides. Direct manipulation of the membrane is demonstrated with membranes with a reduced annulus. One of the microfluidic devices designed in this thesis can host several membranes simultaneously, which are all accessible with an optical tweezers. This device facilitates optical tweezers studies of by allowing to work with different membranes of same lipid composition in a same device with access to both sides of the membranes. Direct mechanical manipulation and adjustable buffer conditions both simultaneously are highly desired features that this technique offers, enabling the study of biological processes that depend on asymmetric conditions on each membrane sides. In addition, the easy access facilitates the study of the formation of lipid nanotube via the intrusion of objects on a flat membrane. The methodology developed in the context of this thesis can be used for combined electrophysiology and force spectroscopy of lipid membranes. To reach the full potential of this technique, a more complete descriptive model of the membrane is needed. More complex lipid membranes could also be implemented as future work.
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This thesis presents a reliable technology to assemble free standing lipid membranes using microfabricated devices. A microfluidic cartridge consisting of parallel channels connected with a rectangular aperture was designed and characterized to assemble artificial membranes. This methodology resulted on a system capable of assembling lipid bilayer membranes of different lipid composition. Using decane as organic solvent, ~70% of the aperture was covered by the lipid bilayer, while the remaining are was occupied a pocket of solvent (annulus). An almost complete depletion of the annulus can be achieved by choosing a solvent (chloroform) capable of being absorbed by the flowcell material. In comparison with others methods, this approach is an important contribution to the field as it is allows real-time control over conditions (voltage, molecules in solution, pH) over both leaflets of the membrane. Furthermore, the lipid bilayer plane is perpendicular to the microscope focal plane, allowing observation of morphological changes in the lipid membrane and straightforward combination with optical techniques. This work shows the first successful operation of optical tweezers combined with planar lipid membranes accessible from both sides. Direct manipulation of the membrane is demonstrated with membranes with a reduced annulus. One of the microfluidic devices designed in this thesis can host several membranes simultaneously, which are all accessible with an optical tweezers. This device facilitates optical tweezers studies of by allowing to work with different membranes of same lipid composition in a same device with access to both sides of the membranes. Direct mechanical manipulation and adjustable buffer conditions both simultaneously are highly desired features that this technique offers, enabling the study of biological processes that depend on asymmetric conditions on each membrane sides. In addition, the easy access facilitates the study of the formation of lipid nanotube via the intrusion of objects on a flat membrane. The methodology developed in the context of this thesis can be used for combined electrophysiology and force spectroscopy of lipid membranes. To reach the full potential of this technique, a more complete descriptive model of the membrane is needed. More complex lipid membranes could also be implemented as future work.
