Spontaneous Formation and Rearrangement of Artificial Lipid Nanotube Networks as a Bottom-Up Model for Endoplasmic Reticulum

1. Preparation of Phospholipid Vesicle Suspension

NOTE: For all materials referred to as "clean" in this protocol, thoroughly wash them with isopropanol followed by deionized water and blow-dry them with nitrogen. Note that a treatment of glass substrates with strongly oxidizing acidic agents (Piranha solution), which is typically applied in preparation protocols for supported lipid films on solid substrates, should not be performed on Al₂O₃-coated carriers.

1. Place in a clean 10 mL round-bottom or inverted pear-shaped glass flask: Soy L-α phosphatidyl choline (PC, 69% w/w), 1,2-dioleoyl-sn-glycero-3-phosphoethanolamine (DOPE, 30% w/w), and a lipid-conjugated fluorophore of choice [e.g., Texas Red 1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine triethylammonium salt (TR-DHPE, 1% w/w)] in chloroform; for a total amount of 3000 µg of lipids in 300 µL of chloroform, resulting in a final concentration of 10 mg/mL.
   NOTE: Use clean, glass, gas-tight syringes with polytetrafluoroethylene plungers when handling compounds containing chloroform.
   CAUTION: Chloroform is toxic and highly volatile and should always be handled under a fume hood with associated personal protective equipment.
2. Connect the flask to a rotary evaporator, position with a tilt of 45°, and rotate at 24 rpm inside a water bath at 23 °C for 6 h with reduced air pressure to slowly and completely remove the chloroform. Start reducing the pressure right after initiating the rotation by steps of 20 kPa every 2 min until it reaches 20 kPa (150 Torr, 80% vacuum).
   NOTE: The formation of a homogenous lipid film of uniform thickness in the preparation vessel is the most important requirement for the rotavap procedure. Lipid preparations are sensitive to rotation velocity, rapid pressure changes, and final pressure value; therefore, strictly follow the slow reduction steps as well as the end pressure and rotation velocity. Position the flask with a tilt of 45° to guarantee that the dehydrated lipid cake is formed evenly as a film on the wall of the flask. Too fast of a rotation leads to turbulences and too slow of a rotation leads (due to gravity) to the accumulation of a thick layer of fluid at the bottom of the flask. During the subsequent overnight swelling process, a very inhomogenous lipid mass is produced that does not respond well to the final sonication step, and the resulting fractions are of different compositions. Pressure and time within the range specified in the method ensures slow desolvation. With chloroform as the solvent, too rapid of a drop in pressure cools down the mixture, resulting in increased viscosity and aggregate and uneven film formation. The long time period of 6 h is recommended in order to remove solvent to the greatest extent possible, as the organic solvent partitions into the lipid material upon rehydration.
3. After 6 h, stop the rotation and increase the air pressure again, gradually, by steps of 20 kPa every 2 min until reaching 100 kPa. Remove the flask from the rotary evaporator and add 3 mL of PBS and 30 µL of glycerol. Gently swirl the flask to dissolve the glycerol. Use an air-tight glass stopper to seal the flask containing the lipids.
   NOTE: The glycerol is used to prevent the complete dehydration of the lipid film and enables bilayer separation¹². It should be heated before use to decrease its viscosity, which facilitates the handling of this compound. The warmed glycerol does still not immediately mix with the PBS buffer. Gentle swirling is required until the glycerol is completely dissolved.
4. Store the flask in the refrigerator at 4 °C overnight for rehydration and swelling of the lipid films.
5. On the following day, sonicate the lipids with an ultrasonic water bath at room temperature (RT, ~21 °C) and at 35 kHz frequency until achieving a uniform, slightly turbid lipid suspension.
   NOTE: Sonication can take around 10-30 s. Prolonged sonication (~1 min) produces heat and is detrimental to vesicle formation.
6. Steps 1.1-1.5 yield a suspension containing two types of vesicular structures: multilamellar vesicles (MLV) and giant unilamellar vesicles (GUV) (Figure 1A-1F).
7. For storage, divide the lipid suspension into 100 µL aliquots, using a total of 30 microcentrifuge tubes, and store them in a freezer at -20 °C.
   NOTE: Flash freezing with liquid nitrogen is not necessary and not used before storage. The protocol can be paused here. Leaving lipid suspensions in the 4 °C refrigerator for prolonged times causes lipid lysis, which affects the membrane composition.

2. Preparation of Substrates

NOTE: The following protocol is performed at a cleanroom classified as ISO 8 in the ISO 14644-1 standard specification. Atomic layer deposition (ALD) is used to fabricate Al₂O₃ substrates. The specified process parameters are instrument-dependent and may vary between different models of equipment. They can be used as initial parameters to develop the process.

1. Set the temperature of the ALD reactor to 200 °C.
2. Load the glass surfaces (e.g., glass coverslips) into the sample chamber together with a silicon wafer, which will be used later as a reference surface to determine thickness of the deposition by ellipsometry.
   NOTE: Glass substrates were used out-of-the-box and were not solvent-cleaned before deposition. They were only flushed with nitrogen gas to remove particles.
3. Evacuate the loading chamber to 400 Pa (3 torr) and transfer the samples into the main reaction chamber, and evacuate it to 200 Pa.
   NOTE: The temperature of the reactor must be maintained at 200 °C for proper deposition. Temperature fluctuations after sample loading must therefore be equilibrated before initiating the deposition. Chamber pressure is set to be higher than reactor pressure to avoid any precursors to spread outside the chamber.
4. Start depositing the atomic film. One cycle consists of a 150 ms pulse trimethyl aluminum exposure, followed by a 1 s purge, and subsequently, a H₂O exposure of 200 ms duration followed by a 1 s purge.
   NOTE: All settings including chamber and reactor pressure, length of the cycles, and purges are automated to achieve a defined rate of deposition. These parameters may vary among the various models of equipment. The pre-configured recipes are often prepared by the vendor or tool responsible in the clean room and communicated to the user as deposited film thickness per unit of time.
5. To reach 10 nm of Al₂O₃ on the substrate, repeat the process for 100 cycles. The number of cycles depends on the deposition rate, which may vary among different recipes or equipment.
6. To remove the samples from the reactor, first vent the chamber until its pressure reaches atmospheric pressure, then remove the samples.
7. Store the samples in air tight containers at RT until use.
   NOTE: No further cleaning is recommended before use. The protocol can be paused here.
   NOTE: The samples should ideally be used immediately after deposition. The optimal storage requires positioning the surfaces inside polypropylene wafer carriers, followed by enveloping the carriers in clean room-compatible plastic bags, which are to be nitrogen-flushed before vacuum sealing. The purpose is to avoid exposing the surface to air-borne contaminants. If necessary, the surfaces can be kept in air tight containers at RT at maximum for 5 days. Longer storage is not recommended. For users who do not have easy access to a clean room nearby, and purchase or obtain the surfaces from abroad, re-oxidizing the substrates by means of oxygen plasma or ozone treatment may be an alternative solution¹³.

3. Transformation of Molecular Phospholipid Films to Tubular Networks

1. Thaw the lipid suspension and transfer a 4 µL droplet of suspension onto a clean glass microscope slide/coverslip.
2. Desiccate the droplet for 20 min. The droplet will collapse into a flat circular film of lipids after desiccation, which is visible to the eye.
3. Rehydrate the lipid film with 1 mL of HEPES buffer (see Table of Materials) for 3 min.
   NOTE: The volume of rehydration buffer affects the density of vesicle suspension (the number of vesicles per unit volume), which is subsequently transferred to the observation chamber. Depending on the volume of the observation chamber and the desired vesicle density, the rehydration volume can be tuned to 0.5-1 mL. Clean borosilicate slides tend to support droplets of several hundred microliters up to 1.5 mL without problems. Since the coverslip does not need to be moved, this does not lead to a technical problem. On more hydrophobic surfaces, such as SU-8 polymer-coated slides, even 1.5 mL can be deposited¹².
   NOTE: Lipids should be freshly prepared, as exposure of the rehydrated lipid film for more than 20 min at RT leads to evaporation of the buffer and partial dehydration of the previously rehydrated vesicles, which leads to a poorly defined composition.
4. Prepare the observation chamber: to allow for buffer exchange by means of an automatic pipette, which is required to initiate the ER transformation, an observation chamber with an open top was used. This chamber consists of a polydimethylsiloxane (PDMS) frame with dimensions 1.5 x 1.5 x 0.5 cm, adhered onto the Al₂O₃ deposited coverslip. A scheme of the mounted observation chamber is represented in Figure 1G. The following steps were performed to fabricate the PDMS frame and assemble the observation chamber:
   1. Prepare a KOH solution by mixing 100 g of KOH with 100 mL of isopropanol in a beaker in an ice bath. Stir for 10 h or longer until the KOH is completely dissolved using a magnetic stirrer and magnetic stir plate.
      CAUTION: KOH solution is corrosive and can lead to skin burns, so always handle with proper personal protective equipment.
      NOTE: The solubility of KOH in isopropanol is not as high as in water. The dissolution is exothermic. Crushing the KOH pellets prior to dissolving and continuous stirring is advisable.
   2. Submerge a glass Petri dish (d = 6 cm) into the KOH solution at RT and keep it overnight.
   3. The following day, remove the glass dish from the solution, immerse it in a container with deionized water for 5 min, rinse it several times with water, and place it inside a drying oven at 80 °C for 1 h. Blow the surface briefly with a stream of nitrogen to ensure that particles are removed.
   4. To passivate the surface and prevent bonding to PDMS, transfer 200 µL of dimethyldichlorosilane with a plastic syringe into a clean plastic container such as a weighting boat.
   5. Store the glass Petri dish together with the silane for 1 h in an evacuated desiccator (low vacuum, ~20 kPa).
      CAUTION: Dimethyldichlorosilane is toxic and should always be handled under a fume hood with associated personal protective equipment.
   6. Wait 15 min before collecting the Petri dish in order for the remaining dimethyldichlorosilane vapor to dissipate. The Petri dish is now silanized, and the surface is hydrophobic.
      NOTE: A quick way to test the success of this step is to place a water droplet onto the silanized Petri dish. The contact angle of the droplet with the surface must visibly increase compared to untreated glass.
   7. In a 250 mL plastic container (transparent plastic cup fresh from the package), mix 10 g of silicone elastomer base with 1 g of silicone elastomer curing agent (10:1). Stir using a plastic stirrer/plastic spatula for 5 min.
      NOTE: Air bubbles are formed upon stirring, and the PDMS will look pale-white.
   8. Desiccate the mixture to degas at < 20 kPa until all expanding air bubbles have collapsed (higher vacuum accelerates the process). Pour the degassed mixture into the silanized Petri dish.
   9. Cure at 65 °C for 2 h in an oven.
      NOTE: It is possible to double the curing speed by increasing the temperature to > 95 °C. The increasing curing temperature results in an increase in stiffness of the material.
   10. Cool down the Petri dish filled with the cured PDMS to RT and remove the PDMS slab with a spatula.
   11. With a scalpel, cut the frame into the dimensions and geometry appropriate for the available opening in the microscope stage. Dimensions of 1.5 (length) x 1.5 (width) x 0.5 (height) cm are suitable for most setups.
   12. Bring the smooth side (the bottom side that was in contact with the Petri dish) of the PDMS frame into contact with the active side of the surface where the Al₂O₃ film resides, and gently apply pressure to push the frame and surface against each other to make them adhere.
       NOTE: The adhesion between the PDMS frame and Al₂O₃ substrate is weak. The presence of air bubbles at the contact interface may cause de-attachment and, consequently, the leakage of the buffer and related content. The PDMS frame can be used several times if immediately after and before each use it is rinsed with isopropanol, followed by rinsing with DI water and blow-drying with nitrogen. The silanized Petri dish can also be reused.
5. Fill the observation chamber with Ca²⁺-HEPES buffer (see Table of Materials).
   NOTE: The surface should be used immediately after unsealing the package. Contact with air leads to adsorption of contaminants, which decreases the activity of the surface gradually. The chamber should be filled with buffer immediately after assembly. Do not fill the entire chamber volume to allow for addition of the rehydrated lipids in the subsequent step.
6. Place the chamber onto the confocal microscope stage. Transfer the rehydrated lipid material, now a suspension containing giant vesicles, into the chamber with a plastic Pasteur pipette (Figure 1A-1G).
7. Wait 10-20 min to let the vesicles adhere onto the substrate and spread across the surface (Figure 1H-1J).
   NOTE: The spreading starts immediately after deposition of lipids on the surface. The rate of spreading might slightly vary depending on the lipid composition, Al₂O₃ deposition technique (ADL, RF-sputtering, chemical vapor deposition, etc.), freshness of the substrate, and divalent cation concentration in the buffer. Ensure that buffer exchange is performed before rupturing of the spreading patches¹⁴.
8. After observing multiple lipid spreads, slowly remove the ambient buffer via an automatic pipette, such that only a thin buffer film remains on the bottom.
   NOTE: Rapid removal of buffer perturbs the lipid structures on the surface.
9. Proceed to the ambient buffer exchange by slowly filling the observation chamber with chelator-HEPES buffer (see Table of Materials) using an automatic pipette (Figure 1K).
   NOTE: Abrupt addition of buffer perturbs the lipid structures on the surface.
10. This final step yields dynamic nanotubular networks, formed as a result of the chelator-induced depinning and retraction of the DLBM to the MLV⁴ (Figure 1L-1Y).

4. Microscopy Observation

1. Acquire the images with an inverted laser scanning confocal microscope using a 40X oil (1.3 NA) immersion objective with a scanning frequency of 400 Hz. Employ a white light laser source to excite the Texas Red DHPE at 595 nm. Collect the emission from 605 to 700 nm by using a hybrid photon detector.
   NOTE: Alternatively, an epi-fluorescence microscope can be used for imaging. Depending on the light sources available, select an appropriate lipid-dye conjugate.