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Back to Journal »International Journal of Nanomedicine» Volume 16

Lipids in Chips: A brief review of microfluidic formation of liposomes

Author Zhang Geng, Sun Jie

Published on November 3, 2021, the 2021 volume: 16 pages 7391-7416

DOI https://doi.org/10.2147/IJN.S331639

Single anonymous peer review

Editor who approved for publication: Dr. Mian Wang

Sun Jiaming Plastic Surgery, Union Hospital, Tongji Medical College, Huazhong University of Science and Technology, Wuhan, China Zhang Guo Communication: Sun Jiaming Email[email protected] Abstract: Liposomes are ubiquitous tools in biomedical applications, such as drug delivery, membrane science And artificial cells. Micro- and nano-manufacturing technologies have revolutionized the preparation of micro-scale liposomes. This review introduces the most advanced liposome formation on microfluidic chips and its related applications. We try to provide references for liposome researchers by comparing various microfluidic technologies used for liposome formation. Keywords: liposomes, lipid vesicles, microfluidics, chips

Liposomes or lipid vesicles are artificially synthesized from one or several phospholipid bilayers, enclosing a water core, ranging in size from nanometers to hundreds of microns. First discovered in the 1960s, 1,2 liposomes have become practical tools for drug delivery, 3-6 membrane science 7-9 or artificial cell synthesis 10-12 because they are similar to cells and natural vesicles and are easy to form and manipulate And modify.

The overwhelming success of lipid-based mRNA vaccines has brought a glimmer of hope in response to the COVID-19 pandemic. 13,14 As a safe and effective carrier, liposomes can protect the loaded materials from external degradation, so they are rapidly developing as a multifunctional vaccine adjuvant delivery system in inducing the immune response required for cancer treatment and vaccines. Has a strong ability. 15-18 In addition, recently reported breakthroughs in lipid-mediated gene delivery technologies (such as targeted genome editing), 19,20 a promising era for the treatment of genetic diseases. At the same time, liposomes have become an indispensable tool for membrane protein research and synthetic biology, helping to explore new drugs for the treatment of cancer and other diseases, and to clarify complex cell functions. 21-23

The popularity of liposomes in research has facilitated the search for preparation methods in an economically viable way. In addition to anti-evaporation 24 and ethanol injection 25, common production methods include film hydration, 26, 27 extrusion, 28, 29 electroforming, 30, 31 freeze-drying 32, 33, and double emulsion. 34,35 Although commonly used, because of their availability, these macroscopic techniques have been criticized for their inability to precisely control the size and polydispersity of liposomes and the insufficient use of reagents and materials. For example, artificial cell chambers require precise shape and size control to simulate living cells, but it is extremely challenging (nearly impossible) to achieve such accuracy using macroscopic techniques. It is well known that smaller liposome size (≤100 nm) may be suitable for enhancing the activity of anti-tumor drugs encapsulated in liposomes. 36,37 The production of "limit-size" liposomes with a diameter of less than 50 nm is only suitable for the use of macroscopic techniques such as sonication and homogenization; however, microfluidic technology can easily achieve the scalability of SUVs with a size of 20-50 nm Production. 38 In addition, encapsulation materials in liposomes dissolved in aqueous solutions are sometimes very expensive (such as mRNA), and batch methods inevitably tend to have poor reproducibility and waste of materials. The emergence of microfluidic technology has solved this dilemma, reduced the formation of liposomes to centimeter-level chips, greatly reduced the related costs, and improved the production efficiency and operability of liposomes.

As an updated version of a review covering similar topics published earlier, 39-41 this review focuses on the current state-of-the-art liposome preparation methods in microfluidic chips and related applications. A comprehensive comparative analysis of various methods classified according to design principles is carried out to highlight their advantages and disadvantages. We intend to provide insightful guidance for each method and its specific applications by detailing the advantages and limitations.

Lipid vesicles can be divided into unilamellar vesicles (UVs), oligolamellar vesicles (OLVs), multilamellar vesicles (MLVs) and multi-vesicles (MVVs) according to their lamellar structure. 42 In addition, UVs can be divided into small unilamellar vesicles (SUV) with vesicles with a particle size of <100 nm, large unilamellar vesicles (LUV) with a particle size between 100 and 1000 nm, and those with a particle size> 1 μm. Giant unilamellar vesicles (GUV) (according to their size) (Figure 1). Generally, dynamic light scattering (DLS) and nanoparticle tracking analysis (NTA) techniques are used to assess the size distribution of vesicles. 43,44 Common methods used to determine the laminarity of liposomes include nuclear magnetic resonance (NMR), 45 small-angle X-ray scattering (SAXS) 46, and transmission electron microscopy (TEM), 47 to name a few. Figure 1 Classification of vesicle size and layered structure. The diameter of small unilamellar vesicles (SUV) is less than 100 nm; large unilamellar vesicles (LUV) are between 100 and 1000 nm; and giant unilamellar vesicles (GUV) are larger than 1 μm. Oligolamellar vesicles (OLV) are similar to multilamellar vesicles (MLV) and consist of two to five concentric bilayers, while MLV contains more than five concentric bilayers. Multivesicular vesicles (MVV) encapsulate multiple non-concentric bilayer liposomes.

Figure 1 Classification of vesicle size and layered structure. The diameter of small unilamellar vesicles (SUV) is less than 100 nm; large unilamellar vesicles (LUV) are between 100 and 1000 nm; and giant unilamellar vesicles (GUV) are larger than 1 μm. Oligolamellar vesicles (OLV) are similar to multilamellar vesicles (MLV) and consist of two to five concentric bilayers, while MLV contains more than five concentric bilayers. Multivesicular vesicles (MVV) encapsulate multiple non-concentric bilayer liposomes.

As a drug delivery system, MLV and MVV are more commonly used to achieve long-term drug delivery effects. 48,49 In order to function effectively and efficiently, liposomes of a certain size need to be prepared to allow absorption into cells. Size is a key parameter that determines liposome drug encapsulation, circulating half-life, and smaller liposome diameter, which may help optimize drug release in vivo. 50 Therefore, SUV as a vaccine drug carrier is the first choice and anti-tumor drug, to name a few. 36,51 Because they are similar in size to cells, GUVs are more commonly used in membrane science and artificial cell synthesis. 52 The difference in size leads to differences in properties and applications, but for monodispersity, 1 the main motivation for preparing liposomes using microfluidic chips must be ensured to maintain reproducibility.

The stability of vesicles is not only affected by size and lipid composition, but also by the physical and chemical environment (such as pH and temperature) in the vesicles. 53,54 For example, the double-layer cross-linked MLV made by Moon et al. retains about 95% of the amount of protein they are encapsulated when stored in PBS at 4°C for more than 30 days. 55 Merging ppylation has also become a popular method for enhancing stability. 56,57 Compared with polyethylene glycol (PEG) modified liposomes that can basically remain unchanged. After 2 months of storage, conventional liposomes lacking PEG surfactants showed gradual aggregation and precipitation, indicating that PEG The modification improves the stability of the vesicles. 57 The surface electrostatic potential of liposomes has been shown to play a key role in the binding constant, and in general, charged liposomes are not prone to aggregation due to electrostatic repulsion. 59 Zeta potential of liposomes, used to absorb charges close to the surface, can be measured by laser Doppler velocimetry (LDV). 60

Encapsulation efficiency (EE) is defined as the ratio of the encapsulated drug to the initial amount of drug in the system. It is affected by many factors, such as drug properties, liposome size, lamellar structure and preparation method. Effective separation of liposomes from samples is the most critical step in determining EE. Common separation methods used to determine EE include centrifugation, dialysis, chromatography, and ultrafiltration. It can be quantified after liposome separation and destruction. However, NMR and fluorescence methods can directly determine EE without a separation step and avoid errors caused by the process (Table 1). Table 1 Comparison of methods to determine liposome encapsulation efficiency

Table 1 Comparison of methods to determine liposome encapsulation efficiency

Yamamoto et al. developed nanoparticle size exclusion chromatography, which can determine EE without pretreatment. Only 5 μL of liposome suspension and 3 minutes of analysis time are required to quantify the amount of unencapsulated drug. 67 Differences in separation methods may lead to different results. Therefore, it is necessary to choose a suitable method to obtain reliable EE. It is worth noting that high EE does not mean better performance. Bioactive molecules such as proteins may be affected by various physical and chemical factors during the encapsulation process, resulting in only a few liposomes eventually functioning normally. 39

This method was first proposed by Jahn et al. to prepare nano-scale lipid vesicles in a microfluidic chip (Figure 2). 68 The central channel of lipids dissolved in alcohol is vertically connected to the two side channels by the aqueous solution. When the alcohol is mixed and diluted to a critical concentration by the aqueous solution, the lipids spontaneously self-assemble into liposomes. Hydrodynamic flow focusing (HFF) can prepare monodisperse liposomes, and their size can be adjusted by adjusting the flow rate ratio (FRR) of the water phase to the lipid phase. Subsequent studies confirmed that liposomes with similar size distribution can be prepared by adjusting FRR with different chip geometries, and the reduction of FRR will increase the size of vesicles. 69-71 Lipid composition and concentration have also been identified as the main factors affecting liposome size and polydispersity index. 72,73 In order to further study the self-assembly process of liposomes, Jahn et al. further combined the HFF device with propane jet freezing. The disc-like intermediate structure formed at the interface of lipid-alcohol and buffer reveals more details about the formation of non-equilibrium liposomes. 74 Figure 2 Schematic diagram of an example of the hydrodynamic flow focusing (HFF) method. Schematic diagram of HFF liposome formation process. The color outline indicates the concentration ratio of alcohol to aqueous buffer. Reprinted with permission from Jahn A, Lucas F, Wepf RA, Dittrich PS. Freezing continuous flow self-assembly in a microfluidic device: imaging of liposome formation. Langmuir. 2013;29(5):1717–1723. doi: 10.1021/la303675g. Copyright 2013 American Chemical Society. 74

Figure 2 Schematic diagram of an example of the hydrodynamic flow focusing (HFF) method. Schematic diagram of HFF liposome formation process. The color outline indicates the concentration ratio of alcohol to aqueous buffer. Reprinted with permission from Jahn A, Lucas F, Wepf RA, Dittrich PS. Freezing continuous flow self-assembly in a microfluidic device: imaging of liposome formation. Langmuir. 2013;29(5):1717–1723. doi: 10.1021/la303675g. Copyright 2013 American Chemical Society. 74

Balbino et al. designed a dual-hydrodynamic focusing (DHF) device to produce unilamellar cationic liposomes by adding a pair of additional HFF-based channels. 75 DHF doubles the production rate compared to HFF, and can also adjust FRR to form liposomes with a smaller diameter, while the polydispersity index (PDI) is no different from HFF. 71,76

In order to further improve productivity, Hood et al. used a concentric capillary array to create a three-dimensional microfluidic hydrodynamic focusing (3D-MHF) device. The array consists of seven identical glass capillaries fused together into a circle. 77 Poly(ether-ether-ketone) (PEEK) tubing was used as the lipid feed line and passed through the center of the array (Figure 3A). Due to the complete radial symmetric diffusion of the ethanol-lipid solution, the 3D-MHF device can synthesize nano-scale liposomes with low PDI at a productivity four orders of magnitude higher than the previous HFF method. FRR and the size of capillary pores in the ring play an important role in the size control of liposomes prepared by 3D-MHF. However, due to the complicated assembly process of the capillary array, the usability is affected. Figure 3 Schematic diagram of an improved HFF device. (A) An example of 3D microfluidic hydrodynamic focusing (3D-MHF). The device consists of a narrow-bore capillary surrounded by a glass multi-capillary array. The liposome is prepared as an alcohol-soluble lipid solution continuously injected into a ring capillary, and is hydrodynamically focused in a three-dimensional space through the flow of an aqueous buffer. Not to scale. Reprinted with permission of the Royal Society of Chemistry from Hood RR, DeVoe DL, Atencia J, Vreeland WN, Omiatek DM. An easy way to synthesize monodisperse nano-scale liposomes using 3D microfluidic hydrodynamics in concentric capillary arrays. Laboratory chip. 2014;14(14):2403–2409. License communicated through the Copyright Licensing Center, Inc.77 (B) Vertical Flow Focus (VFF) technology. In the VFF design (not to scale), the focus axis is perpendicular to the focus axis (top) of the traditional MHF device. Picture of a VFF device with a focus channel aspect ratio of 100:1 (bottom). Reprinted from Hood RR, DeVoe DL. The vertical flow of microfluidics focuses on high-throughput continuous flow to produce nano-scale liposomes. small. 2015; 11(43): 5790-5799. doi:10.1002/smll.201501345. Licensed by © 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim. 78

Figure 3 Schematic diagram of an improved HFF device. (A) An example of 3D microfluidic hydrodynamic focusing (3D-MHF). The device consists of a narrow-bore capillary surrounded by a glass multi-capillary array. The liposome is prepared as an alcohol-soluble lipid solution continuously injected into a ring capillary, and is hydrodynamically focused in a three-dimensional space through the flow of an aqueous buffer. Not to scale. Reprinted with permission of the Royal Society of Chemistry from Hood RR, DeVoe DL, Atencia J, Vreeland WN, Omiatek DM. An easy way to synthesize monodisperse nano-scale liposomes using 3D microfluidic hydrodynamics in concentric capillary arrays. Laboratory chip. 2014;14(14):2403–2409. License communicated through the Copyright Licensing Center, Inc.77 (B) Vertical Flow Focus (VFF) technology. In the VFF design (not to scale), the focus axis is perpendicular to the focus axis (top) of the traditional MHF device. Picture of a VFF device with a focus channel aspect ratio of 100:1 (bottom). Reprinted in Hood RR, DeVoe DL. The vertical flow of microfluidics focuses on high-throughput continuous flow to produce nano-scale liposomes. small. 2015; 11(43): 5790-5799. doi:10.1002/smll.201501345. Licensed by © 2015 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim. 78

Soon after, the same team launched the Vertical Flow Focus (VFF) device. The device is made of precisely aligned multilayer thermoplastic devices with an aspect ratio as low as 1:100 (Figure 3B). Use this method to prepare monodisperse liposomes (80 to 200 nm in diameter) at a productivity of up to 95 mg/hour. 78 Based on the VFF design, Michelon et al. and Chen et al. used soft lithography or 3D printing to fabricate similar microfluidic chips for high-throughput synthesis of liposomes, respectively. 79,80 Although the use of PMDS to manufacture VFF is an ingenious alternative to improve the throughput of liposome synthesis, as the aspect ratio increases, the patterning of micro-scale features becomes more and more difficult, such as soft lithography. Therefore, in the study of Michelon et al., the throughput of liposomes is limited because the maximum aspect ratio is limited to 6:1. 79 The VFF device manufactured by Hood et al. overcomes these limitations and is more impressive and practical. . At the same time, Chen et al. successfully manufactured VFF chips using stereolithography technology, but due to the limited resolution of commercial printers, the minimum channel width is only 200 μm. 80 Nevertheless, it is believed that 3D printing technology provides great potential for the realization of future improvements in the tunable and high-throughput synthesis of nanoliposomes.

Considerable effort has been made to utilize HFF design to generate liposomes for drug delivery. For example, liposomes encapsulating plasmid DNA synthesized by HFF can effectively deliver genes into HeLa cells. 81 Ran and others successfully prepared liposomes of various formulations through HFF, and then further studied their physical and chemical properties. 82 Although high-throughput-based HFF methods such as VFF can quickly and robustly prepare liposomes on a large scale, they are inevitably affected by low EE. According to reports, the EE of methotrexate-loaded liposomes prepared by DHF or β-carotene-loaded liposomes prepared by VFF is about 60%. 79,83 Hood et al. integrated purification and remote drug loading components into the HFF device. Although the device has unprecedentedly reduced the total on-chip residence time of drug-loaded liposomes to 3 minutes, the EE is about 72%, which is less than traditional long-term remote loading (>95%). 84,85 Since the formation of vesicles, it has been determined by mixing a dilution solvent in an aqueous solution. Solvents such as ethanol cannot be completely removed from the liposomes, which may affect the stability of the membrane. Last but not least, HFF-based devices, while practical, have limitations in the size and quantity of materials produced. For example, liposomes with a diameter of less than 50 nm produced by HFF can only be achieved at a flow rate ratio of 30 or higher, resulting in a large amount of material dilution. 70 The emergence of the microfluidic interleaved herringbone mixer (SHM) can solve the above problems.

Similar to HFF, liposomes can also be formed by using a microfluidic micromixer. As a powerful and versatile method for generating smaller-sized liposomes, the SHM designed by Stroock et al. is used to efficiently mix flows in microchannels. The special interlaced chevron pattern on the channel bottom plate greatly improves the mixing efficiency, which can be changed by the asymmetry of the chevron and the number of chevrons per half cycle. 86 Zhigaltsev et al. pioneered SHM (Figure 4A) as a scalable manufacturing method to produce lipid nanoparticles (LNP) with a size as small as 20 nm by mixing lipids dissolved in ethanol with water streams for milliseconds . They demonstrated the excellent ability of nano-scale liposomes synthesized by SHM to load and retain doxorubicin, and confirmed the double-layer structure of SUV through 31P NMR and cryo-TEM (cryo-TEM). 87 They further reported that by using SHM, it is possible to limit the SUV with a diameter to produce a wavelength of about 20 nm when the FRR is only 3 (the flow rate of the water channel is 1.5 mL/min). Figure 4 Lipid nanoparticle formulation process using a micromixer. (A) A schematic diagram of the preparation process of lipid nanoparticles using a staggered chevron microfluidic mixing device. Reprinted with permission from Zhigaltsev IV, Belliveau N, Hafez I, et al. Using millisecond microfluidic mixing bottom-up design and synthesis of extreme size lipid nanoparticle systems with water and triglyceride cores. Langmuir. 2012;28(7):3633–3640. doi: 10.1021/la204833h. Copyright 2012 American Chemical Society. 87 (B) Invasive lipid nanoparticle production (iLiNP) equipment. The three-dimensional and top view of iLiNP device, the basic structure is 20 baffle mixer structure group. Reproduced from Kimura N, Maeki M, Sato Y, etc. with permission of the American Chemical Society. Development of iLiNP device: Fine-tune lipid nanoparticle size within 10 nm for drug delivery. ACS Omega. 2018;3(5):5044-5051. https://pubs.acs.org/doi/10.1021/acsomega.8b00341. Further permission related to the excerpted material should be directed to the ACS.95 (C) acoustically driven microfluidic micromixer. Fluorescence indicates a mixture of lipid solvent and water in the presence of a sound field. Republished with permission from the Royal Society of Chemistry of Rasouli MR, Tabrizian M. An ultra-fast acoustic micromixer for the synthesis of organic nanoparticles. Laboratory chip. 2019;19(19):3316–3325. Schematic diagram of the electrofluidic power micromixer, Inc.96 (D) through the license conveyed by the Copyright Licensing Center. When the voltage is turned off and on, the fluorescence indicates laminar flow of water and ethanol. Reprinted with permission from Modarres P, Tabrizian M. Electrohydrodynamics-driven micro-mixing for the synthesis of highly monodisperse nano-scale liposomes. ACS applies nanomaterials. 2020;3(5):4000-4013. doi:10.1021/acsanm.9b02407. Copyright 2020 American Chemical Society. 97

Figure 4 Lipid nanoparticle formulation process using a micromixer. (A) A schematic diagram of the preparation process of lipid nanoparticles using a staggered chevron microfluidic mixing device. Reprinted with permission from Zhigaltsev IV, Belliveau N, Hafez I, et al. Using millisecond microfluidic mixing bottom-up design and synthesis of extreme size lipid nanoparticle systems with water and triglyceride cores. Langmuir. 2012;28(7):3633–3640. doi: 10.1021/la204833h. Copyright 2012 American Chemical Society. 87 (B) Invasive lipid nanoparticle production (iLiNP) equipment. The three-dimensional and top view of iLiNP device, the basic structure is 20 baffle mixer structure group. Reproduced from Kimura N, Maeki M, Sato Y, etc. with permission of the American Chemical Society. Development of iLiNP device: Fine-tune lipid nanoparticle size within 10 nm for drug delivery. ACS Omega. 2018;3(5):5044-5051. https://pubs.acs.org/doi/10.1021/acsomega.8b00341. Further permission related to the excerpted material should be directed to the ACS.95 (C) acoustically driven microfluidic micromixer. Fluorescence indicates a mixture of lipid solvent and water in the presence of a sound field. Republished with permission from the Royal Society of Chemistry of Rasouli MR, Tabrizian M. An ultra-fast acoustic micro-mixer for synthesizing organic nanoparticles. Laboratory chip. 2019;19(19):3316–3325. Schematic diagram of the electrofluidic power micromixer, Inc.96 (D) through the license conveyed by the Copyright Licensing Center. When the voltage is turned off and on, the fluorescence indicates laminar flow of water and ethanol. Reprinted with permission from Modarres P, Tabrizian M. Electrohydrodynamics-driven micro-mixing for the synthesis of highly monodisperse nano-scale liposomes. ACS applies nanomaterials. 2020;3(5):4000-4013. doi:10.1021/acsanm.9b02407. Copyright 2020 American Chemical Society. 97

Kastner et al. found that the increase in FRR leads to an increase in PDI, but the size of liposomes prepared by SHM decreases. 88 However, the results of Maeki et al. showed that the size and PDI of liposomes prepared by SHM simultaneously decreased with the increase of FRR. 89 In addition, the number of SHM cycles and the location of the first SHM are also important factors in the production of vesicles. For example, the size of liposomes gradually increases as the number of SHM cycles decreases and the distance from the location of the first SHM increases. Due to the insufficient mixing of aqueous and lipid solutions observed under high FRR conditions, Maeki et al. hypothesized that instead of a completely mixed solution, rapid dilution of ethanol to a critical concentration is a necessary condition for the formation of vesicles. 89 In addition to the parameters of SHM itself, lipid composition, 90 concentration 91 and even solvent selection 92 are considered to be the key parameters for liposome formation. It should be pointed out that not all LNPs prepared from SHM constitute a continuous bilayer, making them liposomes; some LNPs are micellar-like structures (micelles) with a non-aqueous core. 93 Unfortunately, in some studies, techniques such as low-temperature TEM are unaffordable or just not reported to confirm the structure of LNP prepared by micromixers. According to the results of these reports, the formation of liposomes or micelles in SHM may mainly depend on lipid composition and synthesis parameters. 87,90,92,94

Kimura et al. designed an invasive lipid nanoparticle production (iLiNP) device composed of 20 groups of zigzag mixer structures instead of SHM (Figure 4B). The iLiNP device can accurately adjust the size of the LNP at 10 nm intervals through various baffle parameters, flow rates and FRR, ranging from 20 to 100 nm, which cannot be achieved by traditional SHM. 95 It is worth noting that dissolving aqueous solutions of packaging materials such as mRNA and siRNA are expensive, so LNP must be produced under low and constant FRR conditions. The iLiNP device can produce LNP with a size range of 20-40 nm at flow rates of 50, 100 and 500 μL/min and an FRR of 9.

In order to further improve the mixing speed and uniformity, Rasouli et al. introduced an acoustically driven microfluidic micromixer (Figure 4C). 96 Acoustic microflow is enhanced by integrating sharp edges and bubbles in this device, which leads to higher throughput of PDI liposomes. Compared with passive micromixers, the strong and universal acoustic microflow in the device prevents the formation of large nanoparticle aggregates and blockage of channels. However, as the flow rate increases, the vibration amplitude of sharp edges decreases rapidly, which limits throughput. In addition, for non-professionals, the manufacturing of the entire device is somewhat complicated; therefore, the usability is compromised.

Subsequently, the same team used an electrohydrodynamic micromixer to prepare liposomes with a narrower size distribution at a throughput of 1∼2 × 1010 min-1 (Figure 4D). 97 The size of liposomes is mainly determined by the initial lipid concentration and FRR. This device. Compared with traditional micro-mixers that require soft lithography to make grooves, this novel device is easier to implement, requiring only one additional metal deposition step. It is worth mentioning that although the electric field can prevent contaminants from clogging or liquid leakage, liposome electroporation and lipid oxidation may occur, affecting the size and stability of liposomes, and even the lipid bilayer. Passive permeability. 98,99

SHM obviously can produce a large number of liposomes/LNPs of limited size at a lower FRR, avoiding a large amount of material dilution and waste. However, the problem of relatively low EE has been reported. In the study of Joshi et al., the EE of liposomes prepared by SHM loaded with hydrophilic drugs (metformin) or lipophilic drugs (glipizide) was about 20% or 40%, respectively. When used to encapsulate highly lipophilic propofol, the EE is about 40%, which is significantly higher than the EE (15%) in liposomes prepared by ultrasound. 100 It is worth mentioning that the LNP with micellar structure produced by SHM has higher EE when encapsulating biomolecules. For example, 101 Belliveau et al. used a device with six parallel SHM elements to produce 1-palmitoyl-2 -Oleoyl-sn-glycerol-3-phosphocholine (POPC)/cholesterol composition of restricted-size LNP, encapsulating siRNA at very high throughput (72 mL/min or 580 mg LNPs/min), while the EE of siRNA More than 95% and PDI below 0.1.102. These LNPs exhibit an electronically dense core in cryo-TEM, indicating a micellar structure. This morphology is significantly different from the double-layer LNP, which exhibits an electronically dense ring and a low-density interior, which is related to a single-layered vesicle system with an aqueous interior. Leung et al. further proved that the high siRNA encapsulation level of siRNA in the LNP system is inconsistent with the double-layer structure, because the encapsulation depends on the presence of cationic lipids. 103

Generally, in order to prepare liposomes/LNPs for drug delivery, micromixers may be a better choice because of their relatively high EE, ease of use, and full utilization of packaging materials. The throughput of the micromixer increases exponentially by integrating multiple arrays of mixing channels operating simultaneously. 104 In fact, commercial devices based on micromixers (such as the NanoAssemblr™ platform) have been commonly used in many studies to prepare liposomes/LNPs for different applications, such as CRISPR-Cas9 genome editing for cancer treatment 105 and Intrauterine mRNA delivery for monogenic fetal diseases. 106 It is worth noting that the COVID-19 vaccine nanoparticles manufactured by Pfizer are reported to be prepared by a microfluidic mixer. 93

Unlike HFF and micromixers, discrete droplets of definite size produced by immiscible fluids can be used as templates for liposome formation in this method. Generally, these liposomes with a diameter in micrometers are more used in membrane science and artificial cell research than in drug delivery.

Weiss et al. developed a method to combine vesicles and water-in-oil (W/O) droplets to create cell-like compartments. 107 In this method, the LUV prepared by extrusion is first encapsulated in copolymer-stabilized W/O droplets. By introducing Mg2 solution during the droplet production process, due to the transformation of the encapsulated LUVs, a continuous lipid bilayer was formed at the internal interface of the droplet, which they named droplet stable GUVs (dsGUVs). The assembled lipid compartment can be released from the droplet into the aqueous phase by another microfluidic device (Figure 5). Before demulsification, pico injection technology (inspired by Weitz laboratory design) 108 can be used to sequentially load various proteins and molecules into dsGUV to form synthetic cells. By combining different microfluidic technologies, copolymer-stabilized W/O droplets are used as templates for high-throughput preparation of GUVs (103/s) with unique and precise composition. This elegant technology ensures high EE, and no trace of residual oil is detected in the released GUV under the Raman microscope. This technology successfully obtained the bottom-up remodeling of actin cytoskeleton, microtubules and even FoF1-ATP synthase in dsGUVs, overcoming the basic limitations of complex synthetic cell design. Figure 5 Formation of stable GUV droplets. (A) LUVs or GUVs are encapsulated in W/O copolymer stabilized droplets by microfluidics. In order to convert the encapsulated vesicles into a supporting lipid bilayer at the internal interface of the copolymer stabilized droplets, 10mM Mg2 was applied during the production of the droplets. (B) Representative combined image of green fluorescence from lipids and brightfield microscope of encapsulated LUV and GUV and dsGUV. Scale bar, 10 microns. (C) The microfluidic device is designed to release the assembled lipid compartment from the surrounding stable polymer droplets into the aqueous phase. The image on the left shows the fluorescence image of monodisperse dsGUV in the oil phase before release. After injection, the droplets are separated from each other at the T-joint, where the tributary oil stream containing 20 vol% of unstable surfactant merges with the droplet stream. The passive trapping structure can drain the continuous oil phase and decelerate the droplets before they enter the water phase. Comparison of the dsGUVs and the released lipid compartments shows that they are comparable in size. Scale bar, 20 microns. Reprinted with permission from Springer Nature Customer Service Center Co., Ltd.: [Springer Nature] [Nature material] Weiss M, Frohnmayer JP, Benk LT, etc. Continuous bottom-up assembly of mechanically stable synthetic cells through microfluidics. Nat. 2018;17(1):89-96. Copyright 2018. https://www.nature.com/articles/nmat5005.107

Figure 5 Formation of stable GUV droplets. (A) LUVs or GUVs are encapsulated in W/O copolymer stabilized droplets by microfluidics. In order to convert the encapsulated vesicles into a supporting lipid bilayer at the internal interface of the copolymer stabilized droplets, 10mM Mg2 was applied during the production of the droplets. (B) Representative combined image of green fluorescence from lipids and brightfield microscope of encapsulated LUV and GUV and dsGUV. Scale bar, 10 microns. (C) The microfluidic device is designed to release the assembled lipid compartment from the surrounding stable polymer droplets into the aqueous phase. The image on the left shows the fluorescence image of monodisperse dsGUV in the oil phase before release. After injection, the droplets are separated from each other at the T-joint, where the tributary oil stream containing 20 vol% of unstable surfactant merges with the droplet stream. The passive trapping structure can drain the continuous oil phase and decelerate the droplets before they enter the water phase. Comparison of the dsGUVs and the released lipid compartments shows that they are comparable in size. Scale bar, 20 microns. Reprinted with permission from Springer Nature Customer Service Center Co., Ltd.: [Springer Nature] [Nature material] Weiss M, Frohnmayer JP, Benk LT, etc. Continuous bottom-up assembly of mechanically stable synthetic cells through microfluidics. Nat. 2018;17(1):89-96. Copyright 2018. https://www.nature.com/articles/nmat5005.107

The same team further reduced the diameter of dsGUV to ~2 μm through the microfluidic droplet separator, paving the way for the targeted delivery of advanced cargo such as microparticles, viruses or macromolecular DNA robots. 109 They studied the interaction between GUVs of different charges and cells and proved that small GUVs are a practical alternative to transport and deliver very large and complex goods such as baculovirus to specific cells. This is a traditional SUV-based solution. The delivery cannot be achieved. The problem is that the mechanical stability of the droplet splitting GUV is insufficient, resulting in low yield of the sub-GUV. Nevertheless, their strict control over the composition and functionalization of GUVs allows the construction of GUVs with functional diversity and high targeting specificity. However, the disadvantages of a single emulsion template include the challenging requirements for soft lithography (the minimum width of the droplet separator microchannel is less than 2 μm) and the special requirements of pico injection devices.

First proposed by Sugiura et al., the 110 lipid-coated ice drop hydration method is used to prepare monodisperse W/O droplets with a diameter between 4 and 20 µm as a template. After freezing, the droplets precipitate and separate, and then the hexane solution containing lipids is replaced. Next, the solvent is evaporated while the water droplets are still frozen, and the aqueous solution is added, resulting in the formation of oligolamellar or multilayered giant bubbles. However, this method must generate UV through extrusion, and the EE after extrusion drops from 35% to 12%, which limits the practicality. 110 This method has been successfully applied to the entrapment of enzyme (α-chymotrypsin), but the problem is that the problem of low EE has not been solved because it cannot prevent the leakage of part of the initially formed droplets. 111

Pautot et al. first described the droplet emulsion transfer method. The 34,112 W/O emulsion prepared by stirring the mixture of the aqueous solution and the lipid suspension was transferred to the water phase again. The droplets in the emulsion pull out the second monolayer from the oil-water interface to complete the double layer, thereby forming UV (Figure 6A). They successfully encapsulated various macromolecules such as G-action into GUV as a microbial reactor with up to 98% EE. Using this preparation technology, it is also possible to independently prepare asymmetric GUVs of each leaflet with a controllable composition. However, these GUVs are polydisperse because the emulsion is prepared by stirring. Further improvements using microfluidic devices have solved this problem (Figure 6B). Microfluidic droplet formation techniques (such as T-junctions and flow focusing) can be used to prepare monodisperse GUVs from lipid-stabilized W/O droplets. However, the droplets must be recollected to pass through the oil-water interface. 113-115 Importantly, Pautot et al. report that only a small fraction of the larger emulsion droplets will pass through the interface to form liposomes, because most of them may rupture.112 They also observed that asymmetric vesicles require several Hours of equilibration time can achieve complete lipid coverage of the second unilamellar, thereby transforming droplets into unilamellar vesicles. Unfortunately, a shorter equilibration time (<2 hours) leads to insufficient coverage and uncontrolled lipid composition, while a longer equilibration time (> 3 hours) causes lipids to accumulate in the multilayer structure, thereby This is a key limitation of the technology. Figure 6 The method of preparing liposomes using droplet emulsion transfer. (A) Droplet emulsion transfer method for vesicle formation. Reprinted from Pautot S, Frisken BJ, Weitz DA. Engineering asymmetric vesicles. Proc Natl Acad Sci. 2003;100(19):10718. doi: 10.1073/pnas.1931005100. Approved by (2003) National Academy of Sciences 34 (B) Giant liposome formation device using W/O emulsion. Reprinted with permission from Hu PC, Li S, Malmstadt N. Microfluidic manufacturing of asymmetric giant lipid vesicles. ACS application program interface. 2011; 3(5): 1434–1440. doi:10.1021/am101191d. Copyright 2011 American Chemical Society. 114 (C) An integrated microfluidic device that forms vesicles by the droplet transfer method. Reprinted from Matosevic S, Paegel BM. The huge unilamellar vesicles are gradually synthesized on the microfluidic assembly line. J Am Chem Soc. 2011;133(9):2798-2800. doi:10.1021/ja109137s. https://pubs.acs.org/doi/10.1021/ja109137s with permission from the American Chemical Society. 116 (D) Asymmetric giant liposome formation device. Reprinted from Karamdad K, Law RV, Seddon JM, Brooks NJ, Ces O. Investigate the influence of asymmetry on the flexural stiffness of the lipid membrane formed by microfluidics. Chemical Commune. 2016;52(30):5277–5280. doi:10.1039/C5CC10307J.118 (E) Schematic diagram of cell membrane module layer by layer. Phospholipid stabilized water-in-oil droplets are first formed and trapped in a static droplet array. When the oil/water phase boundary passes through the droplets, the lipid monolayer deposition proceeds, resulting in the formation of double bilayer vesicles after the three-phase exchange. Reprinted with permission from Springer Nature Customer Service Center Co., Ltd.: Springer Nature, Nature Chemistry, Matosevic S, Paegel BM. Assemble the cell membrane layer by layer. National Chemistry 2013; 5(11): 958-963. Copyright 2013. 119

Figure 6 The method of preparing liposomes using droplet emulsion transfer. (A) Droplet emulsion transfer method for vesicle formation. Reprinted from Pautot S, Frisken BJ, Weitz DA. Engineering asymmetric vesicles. Proc Natl Acad Sci. 2003;100(19):10718. doi: 10.1073/pnas.1931005100. Approved by (2003) National Academy of Sciences 34 (B) Giant liposome formation device using W/O emulsion. Reprinted with permission from Hu PC, Li S, Malmstadt N. Microfluidic manufacturing of asymmetric giant lipid vesicles. ACS application program interface. 2011; 3(5): 1434–1440. doi:10.1021/am101191d. Copyright 2011 American Chemical Society. 114 (C) An integrated microfluidic device that forms vesicles by the droplet transfer method. Reprinted from Matosevic S, Paegel BM. The huge unilamellar vesicles are gradually synthesized on the microfluidic assembly line. J Am Chem Soc. 2011;133(9):2798-2800. doi:10.1021/ja109137s. https://pubs.acs.org/doi/10.1021/ja109137s with permission from the American Chemical Society. 116 (D) Asymmetric giant liposome formation device. Reprinted from Karamdad K, Law RV, Seddon JM, Brooks NJ, Ces O. Investigate the influence of asymmetry on the flexural stiffness of the lipid membrane formed by microfluidics. Chemical Commune. 2016;52(30):5277–5280. doi:10.1039/C5CC10307J.118 (E) Schematic diagram of cell membrane module layer by layer. Phospholipid stabilized water-in-oil droplets are first formed and trapped in a static droplet array. When the oil/water phase boundary passes through the droplets, the lipid monolayer deposition proceeds, resulting in the formation of double bilayer vesicles after the three-phase exchange. Reprinted with permission from Springer Nature Customer Service Center Co., Ltd.: Springer Nature, Nature Chemistry, Matosevic S, Paegel BM. Assemble the cell membrane layer by layer. National Chemistry 2013; 5(11): 958-963. Copyright 2013. 119

Matosevic et al. successfully integrated the generation and transfer of droplets into a microfluidic chip (Figure 6C). 116 They used triangular columns to drive lipid-stabilized droplets to transfer across the interface, leading to the deposition of a second lipid monolayer. However, due to premature contact with the water stream, the droplets were easily lost, so the assembly yield was only 5% . EE also dropped to 83%. Karamdad et al. further optimized the device to increase liposome production and EE. 117,118 The device contains a microstep junction fabricated by double-layer lithography, in which W/O droplets are transferred through the oil-water interface to construct monodisperse asymmetric vesicles. Throughput (Figure 6D). It is worth noting that double-layer lithography requires high professional skills.

In addition, Paegel's team cleverly integrated a static array on the microfluidic chip to capture lipid-stabilized W/O droplets. 119 By alternately introducing oil and water, each time the oil/water boundary crosses, a new monolayer of lipid is deposited on the fixed droplets, indicating that the platform can be used to assemble phospholipid membranes layer by layer on these droplets (Figure 6E) . Asymmetric multilamellar vesicles were successfully assembled on this chip, which is considered to be a new tool for studying lipid biosynthesis enzymes and transporters.

Based on droplet emulsion transfer, Morita et al. developed a droplet ejection and size filtration (DSSF) method to prepare monodisperse GUV (10-20 μm) with controlled asymmetric lobules in the lipid bilayer. 120 Polydisperse W/O droplets are drawn from the tip of the glass capillary. Only 10-20% of the smallest diameter droplets pass through the oil-water interface selectively through kinetic size filtration (Figure 7), as previously observed by Pautot et al. . 112 1 μL sample solution is enough to generate hundreds of liposomes through DSSF without wasting any material. However, the limited amount of solution in the capillary makes it difficult to efficiently prepare liposomes. In addition, the residual oil between the leaflets was confirmed, although the author claimed that it was low enough to not affect the physical, chemical and biochemical properties. 120,121 Figure 7 The formation of cell-sized liposomes by means of droplet ejection and size filtration (DSSF). (A) Capillary-based microfluidic device. (B) Production of cell-sized liposomes (inside the rectangle shown in (A)). Reprinted from Morita M, Onoe H, Yanagisawa M, etc. Droplet ejection and size filtration (DSSF) method for synthesizing cell-sized liposomes with controlled lipid composition. Chemistry Biochemistry. 2015;16(14):2029-2035. doi:10.1002/cbic.201500354 with permission from Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim120.

Figure 7 The formation of cell-sized liposomes by droplet ejection and size filtration (DSSF) method. (A) Capillary-based microfluidic device. (B) Production of cell-sized liposomes (inside the rectangle shown in (A)). Reprinted from Morita M, Onoe H, Yanagisawa M, etc. Droplet ejection and size filtration (DSSF) method for synthesizing cell-sized liposomes with controlled lipid composition. Chemistry Biochemistry. 2015;16(14):2029-2035. doi:10.1002/cbic.201500354 with permission from Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim120.

The main advantage of the droplet emulsion transfer method based on the microfluidic device is the preparation of a monodisperse GUV with asymmetry and high EE, making it suitable for membrane science and the encapsulation of a variety of biologically active macromolecules. Adir et al. prepared GUVs by emulsion transfer method, and constructed a cell-free expression system by wrapping bacterial lysate in GUVs. The obtained synthetic cells are then used to study protein/RNA production and activity in an isolated environment. 122 However, as mentioned above, the issue of equilibration time is still unresolved, and there is a dispute about the presence of oil residues between liposome leaflets because of the transfer of droplets from the oil phase to the water phase. Kubatta et al. demonstrated that an oil layer accumulated on the top of giant vesicles (3-5 mm) prepared by the batch transfer method, and Hu et al. also observed oil residues in the double layer 114,123. However, the results of the study were Matosevic et al. and Pautot et al. Disagree, because they did not detect double-layer oil traces through thin-layer chromatography or quantitative chemical detection respectively. No inhibition of membrane protein insertion and function was observed. 112,119 Pautot et al. recommended the use of squalene because it has a good emulsion continuous phase and is not miscible in the lipid bilayer. 112 However, due to sensitivity, the possibility of oil residue cannot be ruled out due to the limitations of the technology they use. It seems that different agreements may lead to conflicting conclusions. Imaging based on cryo-TEM or high-sensitivity techniques (such as Raman microscopy) is required to verify the membrane morphology.

First proposed by Shum et al., they used a glass microcapillary device as a template to prepare lipid-stabilized water-in-oil-in-water (W/O/W) double emulsion droplets. The internal phase consists of an aqueous solution containing materials for encapsulation. The middle phase is a phospholipid dissolved in a mixture of toluene and chloroform, while the outer phase is a solution of polyvinyl alcohol (PVA) and glycerin (Figure 8A). 124 As the solvent layer thins during the evaporation process, the phospholipids are concentrated and then forced to rearrange on the double emulsion template to form a GUV. Although the monodisperse GUV with a certain size and high EE can be prepared by this method, a small amount of organic solvent still remains between the lipid leaflets after dialysis. In order to eliminate the existence of poor solvents, the same team used another glass capillary microfluidic device to produce W/O/W droplets with ultra-thin oil shell (<1 μm) as a template to prepare the membrane with the least residual solvent GUV (Figure 8B). 125,126 This time, the internal phase was replaced by a solution of PEG and PVA, and the oil phase was a mixture of chloroform and hexane. Dehumidification occurs because chloroform evaporates faster than hexane. The hexane floats on top of the droplets as an oil bag and eventually evaporates to form a GUV with little residual oil. The 126 GUV prepared by this device was used as a cell-free platform for the synthesis of green fluorescent protein, indicating that the internal PVA aqueous phase may not affect the biological activity of liposomes. 127 Using this technique, a therapeutic enzyme (Cu, Zn-superoxide dismutase) was successfully encapsulated in liposomes to treat mouse ear edema model. An EE of 59 ± 6% and an enzyme activity of 82 ± 3% were obtained. 128 Figure 8 Double emulsion template method. (A) Formation of a phospholipid-stabilized W/O/W double emulsion in a glass microcapillary device. The double emulsion droplets have a water core surrounded by a solvent shell containing phospholipids (top). As the solvent layer thins during evaporation, the phospholipids are concentrated to form GUVs (bottom). Reprinted with permission from Shum HC, Lee D, Yoon I, Kodger T, Weitz DA. Double emulsion templated monodisperse phospholipid vesicles. Langmuir. 2008;24(15):7651–7653. doi: 10.1021/la801833a. Copyright 2008 American Chemical Society. 124 (B) Preparation of water-in-oil-in-water double emulsion drops with ultra-thin shell (top). The hexane floats as an oil pocket on the top of the oil droplets, and then finally evaporates to form a GUV (bottom) with little residual oil. Reprinted with permission from Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim from Arriaga LR, Datta SS, Kim SH, etc. The ultra-thin shell double emulsion templated giant unilamellar lipid vesicles with controllable microdomain formation. small. 2014; 10(5): 950–956. doi:10.1002/smll.201301904.126 (C) Schematic and microscope image of the formation of a double emulsion in a PDMS chip used to prepare liposomes. Reprinted from Teh SY, Khnouf R, Fan H, Lee AP. Stable and biocompatible lipid vesicles are generated through the droplet microfluidic technology based on solvent extraction. Biological microfluidics. 2011; 5(4): 44113–4411312. doi: 10.1063/1.3665221, licensed by AIP. Figure 129 (D) Schematic diagram of the device for producing oil-in-water (W/O/O/W) triple emulsion droplets. The cross-sectional picture corresponds to the position of the dashed line in the horizontal view. Reprinted with permission of the Royal Society of Chemistry from Arriaga LR, Huang Y, Kim SH, etc. One-step assembly of asymmetric vesicles. Laboratory chip. 2019;19(5):749–756. doi:10.1039/C8LC00882E. With permission from the Copyright Licensing Center, Inc.132 (E) an image of the process of forming asymmetric vesicles in a PDMS microfluidic device. (a) Once W/O droplets are formed in the first flow focus area, the emulsion will pass through the serpentine channel. (b and c) The images were taken on the first and last triangular pillars, with fluorescent images of lipid 1 (red) and lipid 2 (green). They confirmed the "double squeeze" separation strategy of replacing lipid 1 (red) with lipid 2 (green) in the oil solution surrounding the droplet. The excess oil/lipid 1 solution flows to the waste outlet. (d) W/O/W double emulsion (indicated by the dashed red arrow) and excess oil droplets (indicated by the solid blue arrow) are formed in the second flow focus area. Reprinted with permission from Lu L, Schertzer JW, Chiarot PR. Continuous microfluidic fabrication of synthetic asymmetric vesicles. Laboratory chip. 2015; 15(17): 3591-3599. doi:10.1039/C5LC00520E. Copyright 2015 American Chemical Society. 133

Figure 8 Double emulsion template method. (A) Formation of a phospholipid-stabilized W/O/W double emulsion in a glass microcapillary device. The double emulsion droplets have a water core surrounded by a solvent shell containing phospholipids (top). As the solvent layer thins during evaporation, the phospholipids are concentrated to form GUVs (bottom). Reprinted with permission from Shum HC, Lee D, Yoon I, Kodger T, Weitz DA. Double emulsion templated monodisperse phospholipid vesicles. Langmuir. 2008;24(15):7651–7653. doi: 10.1021/la801833a. Copyright 2008 American Chemical Society. 124 (B) Preparation of water-in-oil-in-water double emulsion drops with ultra-thin shell (top). The hexane floats as an oil pocket on the top of the oil droplets, and then finally evaporates to form a GUV (bottom) with little residual oil. Reprinted with permission from Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim from Arriaga LR, Datta SS, Kim SH, etc. The ultra-thin shell double emulsion templated giant unilamellar lipid vesicles with controllable microdomain formation. small. 2014; 10(5): 950–956. doi:10.1002/smll.201301904.126 (C) Schematic and microscope image of the formation of a double emulsion in a PDMS chip used to prepare liposomes. Reprinted from Teh SY, Khnouf R, Fan H, Lee AP. Stable and biocompatible lipid vesicles are generated through the droplet microfluidic technology based on solvent extraction. Biological microfluidics. 2011; 5(4): 44113–4411312. doi: 10.1063/1.3665221, licensed by AIP. Figure 129 (D) Schematic diagram of the device for producing oil-in-water (W/O/O/W) triple emulsion droplets. The cross-sectional picture corresponds to the position of the dashed line in the horizontal view. Reprinted with permission of the Royal Society of Chemistry from Arriaga LR, Huang Y, Kim SH, etc. One-step assembly of asymmetric vesicles. Laboratory chip. 2019;19(5):749–756. doi:10.1039/C8LC00882E. With permission from the Copyright Licensing Center, Inc.132 (E) an image of the process of forming asymmetric vesicles in a PDMS microfluidic device. (a) Once W/O droplets are formed in the first flow focus area, the emulsion will pass through the serpentine channel. (b and c) The images were taken on the first and last triangular pillars, with fluorescent images of lipid 1 (red) and lipid 2 (green). They confirmed the "double squeeze" separation strategy of replacing lipid 1 (red) with lipid 2 (green) in the oil solution surrounding the droplet. The excess oil/lipid 1 solution flows to the waste outlet. (d) W/O/W double emulsion (indicated by the dashed red arrow) and excess oil droplets (indicated by the solid blue arrow) are formed in the second flow focus area. Reprinted with permission from Lu L, Schertzer JW, Chiarot PR. Continuous microfluidic fabrication of synthetic asymmetric vesicles. Laboratory chip. 2015; 15(17): 3591-3599. doi:10.1039/C5LC00520E. Copyright 2015 American Chemical Society. 133

Double emulsion droplets were also produced in the PDMS chip. 129-131 Unlike capillary microfluidic devices, these W/O/W droplets are composed of an internal phase of aqueous solution, an intermediate phase of oleic acid, and an external phase containing ethanol (Figure 8C). It is worth noting that ethanol is used to extract oleic acid from the middle layer and then finally evaporates. GUVs are formed by self-assembly of phospholipids at the oil/water interface. Most GUVs prepared by this method can remain stable for a long time (> 3 months) and can be used as synthetic cells for cell-free protein expression. 129 Oleic acid or excess phospholipids may remain in the lipid shell, but this method does not use toxic solvents such as chloroform, so it is more biocompatible than those mentioned above. In addition, the device does not require any complicated design principles and processing except for the extra step of hydrophilization of the external channel.

The double emulsion template method can also be used to prepare asymmetric UV. Arriaga et al. further upgraded the glass microcapillary device. Complex water-in-oil-in-water (W/O/O/W) triple emulsion droplets are used as templates to continuously high-throughput vesicles with asymmetric membranes in one step (Figure 8D). 132 At the same time, Lu et al. An integrated PDMS chip was designed to produce asymmetrical ultraviolet rays. Through their ingenious design, lipid-stable W/O droplets are first formed in the first flow focus area, and then the oil/inner leaf lipid solution surrounding the droplets is replaced by the oil/outer leaf lipid solution. After the W/O/W droplets are formed in the second flow focus area, the asymmetric vesicles are collected outside the chip, and then the oil is extracted between the leaflets of the double emulsion (Figure 8E). 133 The oil phase is also extracted with ethanol. This technique indicates that residual oleic acid or excess phospholipids may remain in the lipid shell. It is reported that more than 80% of liposomes remain stable for at least 6 weeks, and membrane asymmetry remains for more than 1 day. Compared with W/O/O/W emulsion templates, this method seems to be easier to handle and produces asymmetric vesicles with monodisperse size, high EE, and high throughput. However, the choice of solvent is limited to the PDMS chip. For example, chloroform, a solvent commonly used in glass capillary devices, can significantly swell PDMS and cause blockages in the microchannel network. 134

Although these results may increase the utility of vesicles with high EE as model biofilms or drug delivery applications, there are still some limitations in terms of usability. For example, the preparation of liposomes by this technique requires careful handling and assembly of multiple glass capillaries or PDMS chips, and precise control of several microfluidic parameters at the same time. In the study of Arriaga et al., five pumps were used to operate the glass capillary device at the same time, not to mention the strict selection of lipid solvents. 132 Secondly, a variety of ingredients have been introduced, such as PEG, PVA and Pluronic F-68. During the formation of the aqueous solution, this may affect the biological properties of the liposomes. It is important that residual solvent still exists after liposome formation.

In order to solve the problem of oil residue in liposomes once and for all, Deshpande et al. developed a microfluidic technology, namely octanol-assisted liposome assembly (OLA), to form a monolayer, monolayer with efficient, autonomous and fast solvents. Scattered, cell-sized GUV. Extraction mechanism (Figure 9). 135 They replaced the traditional oil phase with 1-octanol. Within a few seconds after its generation, the W/O/W droplet develops into an internal water core, surrounded by a lipid bilayer and a 1-octanol bag containing excess lipids. It is completely combined with the lipid within a few minutes. Plastid separation. By inserting alpha-hemolysin across the bilayer protein pore, the unilaminarity of liposomes formed by OLA was confirmed. The leak-free separation of the oil phase also ensures excellent EE. In addition, the biocompatibility of OLA-based liposomes was verified by encapsulating bacterial cleavage proteins. 135,136 Figure 9 Schematic diagram of octanol-assisted liposome assembly (OLA). (A) The schematic diagram shows the working principle of using OLA to produce liposomes on a chip. (B) Display the corresponding fluorescence image of each of the above steps. (C) Time-resolution sequence showing the separation of 1-octanol droplets from liposomes. The first frame of the sequence in c is obtained about 1 minute after the formation of the double emulsion droplets. Reprinted from Deshpande S, Caspi Y, Meijering AE, Dekker C. Octanol-assisted liposome chip assembly. Nat community. 2016; 7: 10447. doi:10.1038/ncomms10447.135

Figure 9 Schematic diagram of octanol assisted liposome assembly (OLA). (A) The schematic diagram shows the working principle of using OLA to produce liposomes on a chip. (B) Display the corresponding fluorescence image of each of the above steps. (C) Time-resolution sequence showing the separation of 1-octanol droplets from liposomes. The first frame of the sequence in c is obtained about 1 minute after the formation of the double emulsion droplets. Reprinted from Deshpande S, Caspi Y, Meijering AE, Dekker C. Octanol-assisted liposome chip assembly. Nat community. 2016; 7: 10447. doi:10.1038/ncomms10447.135

Similarly, Deng et al. used a surfactant-assisted microfluidic strategy with coaxial glass capillaries to induce spontaneous and complete dewetting of W/O/W droplets to prepare oil-free monodisperse UV. 21,137 The triblock copolymer surfactant Pluronic F-68 is added to the interface energy of the external phase to control the development, which promotes the gradual dewetting of the oil shell from the inside; therefore, the liposomes finally pass through the excess lipids and oil pockets like OLA Separate. After the mixture is poured into the sucrose solution, floating oil droplets can be easily removed by pipetting or evaporation. Multilamellar liposomes can be formed by changing the flow rate using an additional droplet generator (Figure 10). This work realized the controllable production of multilamellar liposomes in microfluidics for the first time. However, the lipids here are dissolved in a toxic organic solvent mixture of chloroform and hexane. Figure 10 Formation of liposomes with different multichambers. (A and B) Schematic and snapshot of the manufacture of a double emulsion with two different droplets. (C) Confocal image of monodisperse liposomes with two different compartments. (D) Prepared liposomes with controlled structures and various configurations. Reprinted with permission from Deng-NN, Yelleswarapu M, Huck WTS. Monodisperse unilamellar and multilamellar liposomes. J Am Chem Soc. 2016;138(24):7584–7591. doi:10.1021/jacs.6b02107. Copyright 2016 American Chemical Society. 137

Figure 10 Formation of liposomes with different multichambers. (A and B) Schematic and snapshot of the manufacture of a double emulsion with two different droplets. (C) Confocal image of monodisperse liposomes with two different compartments. (D) Prepared liposomes with controlled structures and various configurations. Reprinted with permission from Deng-NN, Yelleswarapu M, Huck WTS. Monodisperse unilamellar and multilamellar liposomes. J Am Chem Soc. 2016;138(24):7584–7591. doi:10.1021/jacs.6b02107. Copyright 2016 American Chemical Society. 137

Although there is concern that the external water environment where the liposomes are collected may be contaminated by the rupture of the oil bag, and some 1-octanol may remain in the double layer, OLA solves the limitations of other methods to a considerable extent, such as time -Consuming solvent extraction and residual organic solvents in the lipid bilayer. Since the organic solvents used are bio-friendly, OLA is becoming a multifunctional platform for drug delivery, microreactors, and synthetic cells. For example, the OLA platform is combined with optical fluid transmission analysis to create a complete microfluidic total analysis system for quantifying drug permeability. The transport of norfloxacin and ciprofloxacin through liposomes is measured at physiological pH and salt concentration. 138 Vaezi et al. used OLA to produce liposomes to encapsulate Taxotere. Combined with a microfluidic cell analysis system, the efficacy of encapsulated liposomes is analyzed in a single cell in the capture chip, which is used to quantify drug permeability and cell apoptosis determination. 139 The EE of liposomes loaded with Taxotere was 65.49 ± 3.08%, according to reports. 139 Niederholtmeyer et al. developed a porous cell mimic that can use OLA for gene expression and communication through diffusion protein signals. 140 In addition, OLA has proven to be a powerful tool for controlling and studying the formation of membraneless organelles in liposomes, helping to promote the creation of bottom-synthetic cells. 141

Funakoshi et al. first reported that 142 this method mimics the process of blowing soap bubbles. The pulsed jet generated by the micro-nozzle is applied to the planar lipid bilayer formed by the droplet contact method, because the bilayer is then stretched and finally forms a free vesicle (Figure 11A). 143 The size of the generated vesicles can be adjusted for different spray dispensing time in the following ways. Repeating this process can prepare a large number of monodisperse GUVs (300 μm -600 μm), while forming a small number of smaller "satellite vesicles". However, residual organic solvent (n-decane) was observed inside the film. Subsequently, the same team proved that due to the uneven breakdown of deformed lipid microtubules, smaller "satellite vesicles" would be produced. Based on "satellite vesicles", they prepared an asymmetric GUV with a diameter of 3-20 μm by modifying parameters such as the area of ​​the planar lipid bilayer and jet distribution time (Figure 11B). 144 They confirmed the use of Raman scattering microscopes in these GUVs. Observation of the trigger movement of lipid molecules in the asymmetric GUV indicates that the presence of n-decane between the lobules has little effect on membrane dynamics. 144 Recently, the team successfully produced cell-sized vesicles containing vesicles, namely MVV, which was produced by applying pulsed jets to two parallel plane asymmetric lipid bilayers formed in a three-hole device. The above is similar to the formation of W/O/W droplets. 145 In addition, by applying pulse jets with longer duration and higher pressure to the asymmetric planar lipid bilayers, nano-scale asymmetric lipid vesicles are formed. However, the resulting vesicles are polydisperse, about 30% are multilamellar. 146 Figure 11 Example of pulse jet flow method. (A) Conceptual diagram of the "blowing" method. The green area represents organic solvents. Continuous images formed by vesicles captured by a high-speed CCD camera. The flat membrane is stretched to form a columnar shape and decomposes into spherical vesicles within 10 milliseconds. Reprinted with permission from Funakoshi K, Suzuki H, Takeuchi S. Large lipid vesicle-like compartments are formed from the planar lipid membrane by pulsed jets. J Am Chem Soc. 2007;129(42):12608-12609. doi: 10.1021/ja074029f. Copyright 2007 American Chemical Society. 142 (B) Formation of lipid microtubules from asymmetric planar lipid bilayers by jet flow method. Reprinted from Kamiya K, Kawano R, Osaki T, Akiyoshi K, Takeuchi S. Cell-sized asymmetric lipid vesicles are helpful for the study of asymmetric membranes. National Chemistry 2016; 8(9): 881–889. doi:10.1038/nchem.2537 is approved by Springer Nature.144 (C) Acoustic Jet Method. The focused flow generated by the ultrasonic transducer deforms the planar lipid bilayer, forming a time-series image of GUV and GUV. Reprinted from Armstrong M, Vahey MD, Hunt TP, Fletcher DA. Acoustic jets are used to form and load huge unilamellar vesicles. Biological microfluidics. 2020;14(6):064105. doi: 10.1063/5.0021742, with the permission of AIP publishing. 151

Figure 11 Example of pulse jet method. (A) Conceptual diagram of the "blowing" method. The green area represents organic solvents. Continuous images formed by vesicles captured by a high-speed CCD camera. The flat membrane is stretched to form a columnar shape and decomposes into spherical vesicles within 10 milliseconds. Reprinted with permission from Funakoshi K, Suzuki H, Takeuchi S. Large lipid vesicle-like compartments are formed from the planar lipid membrane by pulsed jets. J Am Chem Soc. 2007;129(42):12608-12609. doi: 10.1021/ja074029f. Copyright 2007 American Chemical Society. 142 (B) Formation of lipid microtubules from asymmetric planar lipid bilayers by jet flow method. Reprinted from Kamiya K, Kawano R, Osaki T, Akiyoshi K, Takeuchi S. Cell-sized asymmetric lipid vesicles are helpful for the study of asymmetric membranes. National Chemistry 2016; 8(9): 881–889. doi:10.1038/nchem.2537 is approved by Springer Nature.144 (C) Acoustic Jet Method. The focused flow generated by the ultrasonic transducer deforms the planar lipid bilayer, forming a time-series image of GUV and GUV. Reprinted from Armstrong M, Vahey MD, Hunt TP, Fletcher DA. Acoustic jets are used to form and load huge unilamellar vesicles. Biological microfluidics. 2020;14(6):064105. doi: 10.1063/5.0021742, with the permission of AIP publishing. 151

Unlike the microdispenser used by Funakoshi et al., Stachowiak et al. use piezoelectric inkjet nozzles for microfluid ejection. 147,148 GUVs with a diameter of 10-400 μm are formed by changing the pulse amplitude, pulse number and fluid viscosity. Using this technology, they synthesized an asymmetric GUV with directional membrane protein incorporation and constructed a synthetic system in which the membrane protein was delivered to the outside of the GUV to simulate all aspects of exocytosis. 149

The problem with this method is that the formation of the double layer produced by manual pipetting requires a certain incubation time, and the rate of membrane regeneration limits the rate of vesicle generation. In addition, the difficulty of repositioning the micro-nozzles relative to the planar lipid bilayer can hinder repeatability and throughput. Therefore, rapid and continuous liposomes cannot be prepared in large quantities by this method. Piezo inkjet technology can increase the productivity by 200 Hz to some extent, but the lipid bilayer needs to pause for 2-4 seconds to re-grow. 148 In order to solve this problem, Gotanda et al. arranged six holes on a circular rotating platform through an automated system to precisely control the rotation speed parameters and ejection time, and sequentially prepared various asymmetric liposomes. Approximately 300 asymmetric cell-sized GUVs (average diameter of 8.9 ± 6.7 μm) were collected in 2 minutes. 150 Armstrong et al. pioneered the use of focused high-intensity ultrasound instead of micro-nozzles to induce focused flow jets to prepare GUVs (Figure 11C), eliminating stability challenges. By controlling the numerical aperture of the acoustic lens and the wavelength of the acoustic wave, a GUV with a diameter of 100-300 μm is formed. 151 It is worth noting that ultrasound itself will affect the stability of liposomes. It is very important to ensure the stability of the planar lipid bilayer during vesicle formation.

In this method, materials such as DNA and cells, regardless of their size, concentration, or chemical properties, can be directly encapsulated into liposomes by spraying, similar to pico injection, which brings high EE (~90%) 144 However, organic solvent residues were found between the leaflets of this method. 142,144,152 Therefore, complex equipment requiring operating conditions and low liposome production limits its further applications beyond membrane science and synthetic cells. Nevertheless, Kamiya et al. used pulsed stream jets to stably produce asymmetric GUVs. 144 These asymmetric GUVs successfully mimicked the lipid triggers corresponding to apoptotic cells and showed the trigger dynamics affected by antibiotic peptides.

Membrane hydration is a common method used to form vesicles. The process involves depositing a lipid film on the substrate after the organic solvent has evaporated. Then the film swells, peels off, and finally self-assembles into liposomes without an aqueous solution. Since the hydration process cannot be controlled, most liposomes prepared are polydisperse MLVs. Need to carry out subsequent extrusion or other treatments to obtain smaller monodisperse UV. 153-155

To solve this problem, Osaki et al. focused on the control of the lipid deposition process. Array micro-holes were fabricated on indium tin oxide (ITO)-glass slides by soft lithography. After using the electrospray deposition (ESD) method, a selectively patterned lipid film was formed on the conductive surface (Figure 12). 156 Through a simple hydration process of dried lipids, they succeeded in forming huge liposomes on top of the pattern. Although ESD can produce monodisperse giant bubbles, the area of ​​the sprayed film is too small to achieve high yields. Figure 12 Electrospray deposition method. (A) An image of a lipid patterned substrate for gentle hydration. (B) A cross-sectional illustration of lipid hydration. The aqueous solution (MilliQ) is simply injected into the hole. ©2011 IEEE. Reprinted with permission, from Osaki T, Kuribayashi-Shigetomi K, Kawano R, Sasaki H, Takeuchi S. Giant liposomes of uniform size are formed and are gently hydrated. In: 2011 IEEE 24th International Conference on Microelectromechanical Systems; 2011; IEEE.156.

Figure 12 Electrospray deposition method. (A) An image of a lipid patterned substrate for gentle hydration. (B) A cross-sectional illustration of lipid hydration. The aqueous solution (MilliQ) is simply injected into the hole. ©2011 IEEE. Reprinted with permission, from Osaki T, Kuribayashi-Shigetomi K, Kawano R, Sasaki H, Takeuchi S. Giant liposomes of uniform size are formed and are gently hydrated. In: 2011 IEEE 24th International Conference on Microelectromechanical Systems; 2011; IEEE.156.

One disadvantage of hydration is low EE, because vesicles cannot be separated from materials in the external environment during the formation process. 157 In addition, this process requires the deposition of a lipid film on the substrate followed by time-consuming evaporation, which prevents continuous preparation of vesicles because lipids cannot be replenished during the operation. Other microfluidic methods used to prepare liposomes, such as micromixers, are more practical when applied to drug delivery.

In 1986, Angelova and Dimitrov first described the electroforming method for preparing giant liposomes. 30 Similar to hydration, a dry lipid film is prepared on the surface of the electrode. An electric field is then applied to the hydrated membrane immersed in the aqueous solution, resulting in the formation of huge vesicles. However, the yield of monodisperse vesicles by electroforming is low, and an effective collection process for preparing liposomes is lacking. 30,154,158

Kuribayashi et al. successfully integrated the electroforming method into the microfluidic chip. 159 Two parallel microfluidic channels formed in a silica gel sheet are sandwiched between ITO glass plates to prepare giant vesicles. The liposomes with different lipid components prepared by this method can encapsulate different types of materials. Because they are polydisperse, micro-contact printing is used to control the size distribution of vesicles. 160-162 Use curtain polydimethylsiloxane (PDMS) stamps to deposit lipids on ITO glass to prepare giant liposomes with a narrow size distribution. In addition, Diguet et al. prepared giant liposomes with controllable size and narrow size distribution by dragging a lipid solution on a microstructured silicon substrate for electroforming. 163,164 Recently, Wang et al. integrated the preparation, separation, and collection of liposomes on a PDMS chip. After electroforming, the liposomes are driven by fresh buffer and captured by the filter membrane in a special collection chamber, thereby increasing the liposome collection rate to 40%. 165

In 2013, Bi et al. introduced the use of coplanar interdigital electrodes for liposome electroformation (Figure 13), breaking the tradition of using two opposite electrodes. 166 In this method, a smaller interdigital electrode produces a larger GUV, which provides a reliable solution. The size distribution of giant liposomes is controlled by electroformation. Figure 13 Schematic diagram of the GUV electroforming setup of the coplanar interdigital electrode. (A) The interdigital ITO electrode (bottom) and the glass cover glass (top) are separated by a PTFE gasket. (B) Side view of GUV electroforming setup. (C) The actual size of the interdigital ITO electrode (top view). Not to scale. Republished by Bi H, Yang B, Wang L, Cao W, Han X with permission from the Royal Society of Chemistry. Electroforming of giant monolayer vesicles using interdigital ITO electrodes. J Mater Chem A. 2013;1(24):7125-7130. Permission communicated through the Copyright Licensing Center, Inc.166

Figure 13 Schematic diagram of the GUV electroforming setup of the coplanar interdigital electrode. (A) The interdigital ITO electrode (bottom) and the glass cover glass (top) are separated by a PTFE gasket. (B) Side view of GUV electroforming setup. (C) The actual size of the interdigital ITO electrode (top view). Not to scale. Republished by Bi H, Yang B, Wang L, Cao W, Han X with permission from the Royal Society of Chemistry. Electroforming of giant monolayer vesicles using interdigital ITO electrodes. J Mater Chem A. 2013;1(24):7125-7130. Permission communicated through the Copyright Licensing Center, Inc.166

In addition to those defects discussed in the hydration method, irregular and asymmetric lipid distribution was observed in the GUV formed by electroforming. 167 The electric field itself may adversely affect the liposomes described in the micromixer. 98,99 Recent studies are mainly focused on exploring new microfluidics for liposome preparation based on electroforming methods, and there are few reports on related applications. 167,168 means that due to the above shortcomings, electroforming The application of the method is limited.

With the successful development of micro- and nano-processing, the preparation of 169 liposomes has changed from macro-scale to micro-scale. Microfluidic technology can not only accurately control the size and lamellar structure of liposomes, but also reduce the volume and cost of reagents required. Table 2 briefly summarizes the comparison between the various microfluidic methods used to prepare liposomes. Different methods have their own advantages and disadvantages. Film hydration and electroforming are more biocompatible, avoiding the presence of organic solvents. When applied to membrane science, the problem of low EE is trivial. Although pulse jets and single emulsion templates with higher EE are more difficult to achieve, they are almost perfect techniques for synthesizing cells. Double emulsion templates, especially the OLA method, are also useful tools for preparing GUVs to explore new drugs and elucidate cell functions. For large-scale production of liposomes/LNPs for drug delivery, micromixers may be the first choice due to their high throughput. Table 2 Comparison between microfluidic methods for liposome production

Table 2 Comparison between microfluidic methods for liposome production

In the case of a single emulsion template, combining dsGUV with pico injection is one of the few ways to directly encapsulate any required material into liposomes with the help of pico injection technology, thereby avoiding dilution or contamination of the encapsulation material . Obviously, LNP is not suitable for larger macromolecules with a radius of gyration as large as 1 μm, which may be necessary for potential applications such as drug delivery and gene therapy. In addition, their various strategies for GUV biofunctionalization (charge, bioligand coupling or PEG coupling) broaden the scope of GUV applications, such as targeted cell delivery. Once the carrier reaches its destination, the potentially immunogenic cargo can be eliminated from the cell through the lysosomal escape mechanism without triggering an immune response. The work of Spatz's group proposed a refined model combining liposome and microfluidic technology, and 107,109 promoted further basic and applied liposome research. Coincidentally, some research has been using other external energies, such as sound waves and electromagnetics, to promote current microfluidic methods by improving production efficiency96,97 or simplifying device manufacturing. 151 These initiatives support recent efforts to research and improve nanotechnology for liposome formation, but future research must critically assess whether the properties of liposomes are easily destroyed by physical methods such as ultrasound and electric fields.

Regardless of the method, bulk or microfluidics, liposomes need to be separated and purified after preparation before further application. Some microfluidic methods, such as single emulsion and double emulsion templates, may require additional post-processing steps (Table 2). For example, floating oil droplets formed by rapid dewetting of double emulsion droplets must be removed by additional pipetting or evaporation steps. 137 OLA also has 1-octanol droplets, which is an inevitable by-product. 135 In order to separate 1-octanol droplets from liposomes formed by OLA, the same team developed a one-step density-based separation technology integrated on the same chip. 170 High separation efficiency (~93%) is achieved compared to unseparated samples (~45%). Similarly, Dimov et al. introduced a microfluidic system composed of SHM and a tangential flow filtration device for continuous purification. This on-chip platform can remove most non-encapsulated materials and organic solvents (>90%) in less than 4 minutes, while providing liposome production and purification. 171 Although the purification step (described in Encapsulation Efficiency) can be performed off-chip, it is important to retain liposomes in the microfluidic chip. It reduces purification time, eliminates possible material loss during off-chip processing, and more importantly, enables downstream on-chip experiments. In fact, efforts have been made to establish a one-step microfluidic platform for the subsequent observation, purification or manipulation of liposomes, such as capture and immobilization, 116,172 separation and purification, 165 dynamic monitoring 173 and even functional evaluation. 138,174 Efforts on these chips will undoubtedly broaden the research horizon of liposomes/LNPs.

LNPs are an upgraded version of liposomes and are now used as a multifunctional nano-drug delivery platform. Although LNP is currently receiving attention due to the success of the COVID-19 mRNA vaccine, viral vectors are more commonly used than LNP because they are more effective in delivering DNA. RNA only needs to enter the cytoplasm to function, while gene therapy requires DNA to be transported through the nuclear membrane to the nucleus. The use of LNP to effectively deliver DNA molecules in vitro and in vivo requires more in-depth research on lipid nanoparticle-DNA formulations.

Most liposomes/LNPs are undoubtedly used in medical science, but scholars who usually use these nanoparticles for drug delivery prefer traditional batch methods, such as ethanol dilution, solvent injection, or homogenization to prepare liposomes. 19,175,176 They focus on designing and optimizing the nanocarriers themselves. Nanocarrier preparation new technologies. Since the production technology cannot solve the key clinical issues in liposome applications, such as delivery efficiency, organ-specific and cell-specific delivery, and toxicity, to name a few, it is not surprising to choose these batch methods because of their simplicity and high cost- Saving and safety. However, researchers who promote the development of preparation technology pay more attention to the basic design of microfluidic devices, such as geometry or the basic physics behind microfluidic-assisted nanoparticle production, such as process dynamics. 177 The vigorous development of liposomes/LNPs requires the coordinated development of medicine, physics, chemistry, engineering and other disciplines. In view of this review, we encourage researchers not only to choose the most suitable technology for the required research, but also to bravely explore the possible applications of new technologies. With the popularity of commercial micromixers (such as the NanoAssemblr™ platform), this situation may change in the near future. 178,179 Based on the current progress and success, we firmly believe that microfluidics technology will soon completely change the production of complex artificial cells and lipids. Based on nano-encapsulation.

The author thanks Dr. Zhenxing Wang for his constructive comments during the preparation of this manuscript. This work was supported by the National Key Research and Development Program (2019YFA0110500) and the National Natural Science Foundation of China (8201001114).

The author has not declared a conflict of interest related to the content of this article.

1. Bangham AD, Horne RW. The negative staining of phospholipids and their structural modification by surfactants were observed under an electron microscope. J Molecular Biology. 1964; 8(5): 660-IN610. doi:10.1016/S0022-2836(64)80115-7

2. Bangham AD, Standish MM, Watkins JC. The monovalent ions diffuse through the expanded phospholipid layer. J Molecular Biology. 1965;13(1):238–IN227. doi:10.1016/S0022-2836(65)80093-6

3. Nakamura T, Miyabe H, Hyodo M, Sato Y, Hayakawa Y, Harashima H. Liposomes loaded with STING pathway ligands and cyclic diGMP enhance cancer immunotherapy for metastatic melanoma. J Control release. 2015;216:149-157. doi:10.1016/j.jconrel.2015.08.026

4. Kulkarni JA, Myhre JL, Chen S, etc. Design of lipid nanoparticles for delivery of plasmid DNA in vitro and in vivo. Nanomed Nanotechnol Biol Med. 2017;13(4):1377–1387. doi:10.1016/j.nano.2016.12.014

5. Deng Hua, Song Ke, Zhao X, etc. Tumor microenvironment activated membrane fusion liposomes with rapid antibody and doxorubicin delivery are used for the collaborative treatment of metastatic tumors. ACS application program interface. 2017; 9(11): 9315-9326. doi:10.1021/acsami.6b14683

6. Lechanteur A, Sanna V, Duchemin A, Evrard B, Mottet D, Piel G. Cationic liposomes carrying siRNA: the influence of lipid components on physical and chemical properties, cytotoxicity and endosomal escape. nanomaterials. 2018;8(5):270. doi:10.3390/nano8050270

7. Elani Y, Law RV, Ces O. Artificial cells based on vesicles are used as chemical microreactors with spatially isolated reaction pathways. Nat community. 2014;5(1):5305. doi:10.1038/ncomms6305

8. Kaddah S, Khreich N, Kaddah F, Charcosset C, Greige-Gerges H. Cholesterol regulates the fluidity and permeability of liposome membranes of hydrophilic molecules. Food Chemical Toxicology. 2018; 113:40-48. doi:10.1016/j.fct.2018.01.017

9. Crone NSA, Minnee D, Kros A, Boyle AL. Peptide-mediated liposome fusion: the influence of anchor positioning. International J Molecular Science. 2018;19(1):211. doi:10.3390/ijms19010211

10. Elani Y, Law RV, Ces O. Protein synthesis in artificial cells: the use of partitions for spatial organization in vesicle bioreactors. Physical Chemistry Chemistry Physics. 2015;17(24):15534–15537. doi:10.1039/C4CP05933F

11. Löffler PMG, Ries O, Rabe A, etc. DNA-programmed liposome fusion cascade. Angewandte Chemie Int Edn. 2017;56(43):13228–13231. doi:10.1002/anie.201703243

12. van Nies P, Westerlaken I, Blanken D, Salas M, Mencía M, Danelon C. DNA replicates itself in liposome-based synthetic cells through its encoded proteins. Nat community. 2018;9(1):1583. doi:10.1038/s41467-018-03926-1

13. Polack FP, Thomas SJ, Kitchin N, etc. Safety and effectiveness of BNT162b2 mRNA Covid-19 vaccine. N Engl J Med. 2020;383(27):2603-2615. doi:10.1056/NEJMoa2034577

14. Khurana A, Allawadhi P, Khurana I, etc. The role of nanotechnology behind the success of the COVID-19 mRNA vaccine. Nano today. 2021;38:101142. doi:10.1016/j.nantod.2021.101142

15. Butts C, Maksymiuk A, Goss G, etc. An updated survival analysis of patients with stage IIIB or IV non-small cell lung cancer receiving the BLP25 liposome vaccine (L-BLP25): a phase IIB randomized, multicenter, open-label trial. J Cancer Res Clin Oncol. 2011;137(9):1337–1342. doi:10.1007/s00432-011-1003-3

16. Yang-ZZ, Li Jiaqing, Wang-ZZ, Dong Dewei, Qi XR. Tumor targeting dipeptide modified cationic liposomes for delivery of siRNA and docetaxel to gliomas. biomaterials. 2014;35(19):5226-5239. doi:10.1016/j.biomaterials.2014.03.017

17. Huang Q, Ji Ke, Tian Sheng, etc. A single dose of mRNA vaccine provides long-term protection for hACE2 transgenic mice from SARS-CoV-2. Nat community. 2021;12(1):776. doi:10.1038/s41467-021-21037-2

18. Huang WC, Zhou S, He X, et al. Particle inoculation can enhance the induction of SARS-CoV-2 RBD neutralizing antibodies. Adv Mater. 2020;32(50):2005637. doi:10.1002/adma.202005637

19. Cheng Q, Wei T, Farbiak L, Johnson LT, Dilliard SA, Siegwart DJ. Selective organ targeting (SORT) nanoparticles for tissue-specific mRNA delivery and CRISPR-Cas gene editing. Nat Nanotechnology. 2020;15(4):313–320. doi:10.1038/s41565-020-0669-6

20. Liu S, Cheng Qi, Wei Tao, et al. Membrane destabilization of ionizable phospholipids for organ selective mRNA delivery and CRISPR-Cas gene editing. Nat. 2021;20(5):701–710. doi:10.1038/s41563-020-00886-0

21. Deng-NN, Yelleswarapu M, Zheng L, Hack WTS. The microfluidic assembly of monodisperse vesicles serves as an artificial cell model. J Am Chem Soc. 2017;139(2):587–590. doi:10.1021/jacs.6b10977

22. Deng-NN, Hack WTS. Microfluidic formation of monodisperse coacervate organelles in liposomes. Angewandte Chemie Int Edn. 2017; 56(33): 9736-9740. doi:10.1002/anie.201703145

23. Langton MJ, Keymeulen F, Ciaccia M, Williams NH, Hunter CA. Controlled membrane translocation provides a mechanism for signal transduction and amplification. National Chemistry 2017; 9(5): 426–430. doi:10.1038/nchem.2678

24. Otake K, Shimomura T, Goto T, etc. A modified supercritical reverse phase evaporation method was used to prepare liposomes. Langmuir. 2006;22(6):2543-2550. doi:10.1021/la051654u

25. Gouda A, Sakr OS, Nasr M, Sammour O. Ethanol injection technology for liposome formulations: Insights into development, influencing factors, challenges and applications. J Drug Deliv Sci Technol. 2021; 61:102174. doi:10.1016/j.jddst.2020.102174

26. Nomura SM, Tsumoto K, Hamada T, Akiyoshi K, Nakatani Y, Yoshikawa K. Gene expression in cell-sized lipid vesicles. Chemistry Biochemistry. 2003;4(11):1172-1175. doi:10.1002/cbic.200300630

27. Kučerka N, Liu Y, Chu N, Petrache HI, Tristram-Nagle S, Nagle JF. The structure of DMPC and DLPC lipid bilayers in a fully hydrated fluid phase using X-ray scattering from oriented multilayer arrays and unilamellar vesicles. Journal of Biophysics 2005;88(4):2626-2637. doi:10.1529/biophysj.104.056606

28. Berger N, Sachse A, Bender J, Schubert R, Brandl M. Using different equipment to filter and extrude liposomes: comparison of liposome size, encapsulation efficiency and process characteristics. Int J Pharm. 2001;223(1):55–68. doi:10.1016/S0378-5173(01)00721-9

29. Jousma H, Talsma H, Spies F, Joosten JGH, Junginger HE, Crommelin DJA. Characterization of liposomes. The effect of multi-layer vesicle extrusion through polycarbonate film on particle size, particle size distribution and number of double layers. Int J Pharm. 1987;35(3):263-274. doi:10.1016/0378-5173(87)90139-6

30. Angelova MI, Dimitrov DS. Liposomes are formed electrically. Faraday discusses the chemical society. 1986; 81:303-311.

31. Estes DJ, Mayer M. Electroformation of giant liposomes from spin-coated lipid membranes. Colloidal surfing B Biological interface. 2005;42(2):115–123. doi:10.1016/j.colsurfb.2005.01.016

32. Li C, Deng Y. A new method for preparing liposomes: freeze-drying of single-phase solutions. J Pharm Sci. 2004;93(6):1403-1414. doi:10.1002/jps.20055

33. Cui J, Li C, Deng Y, Wang Y, Wang W. Lyophilize liposomes using a tert-butanol/water co-solvent system. Int J Pharm. 2006;312(1):131–136. doi:10.1016/j.ijpharm.2006.01.004

34. Pautot S, Frisken BJ, Weitz DA. Engineering asymmetric vesicles. Proc Natl Acad Sci. 2003;100(19):10718. doi:10.1073/pnas.1931005100

35. Liau JJ, Hook S, Prestidge CA, Barnes TJ. Lipid-based multi-compartment system: Liposomal double emulsion for oral vaccine delivery. Eur J Pharma Biopharm. 2015; 97:15-21. doi:10.1016/j.ejpb.2015.09.018

36. Nagayasu A, Uchiyama K, Kiwada H. The size of liposomes: factors that affect their targeting efficiency on tumors and the therapeutic activity of liposome anti-tumor drugs. Adv Drug Deliv Rev. 1999; 40(1–2):75–87. doi:10.1016/S0169-409X(99)00041-1

37. Cabral H, Matsumoto Y, Mizuno K, etc. The accumulation of sub-100 nm polymer micelles in poorly permeable tumors depends on the size. Nat Nanotechnology. 2011; 6(12): 815–823. doi:10.1038/nnano.2011.166

38. Allen TM, Coolis PR. Liposome drug delivery system: from concept to clinical application. Adv Drug Deliv Rev. 2013;65(1):36-48.

39. van Swaay D, deMello A. Microfluidic method of liposome formation. Laboratory chip. 2013; 13(5): 752–767. doi:10.1039/c2lc41121k

40. Carugo D, Bottaro E, Owen J, Stride E, Nastruzzi C. Microfluidic liposome production: potential and limiting factors. Sci Rep. 2016;6(1):25876. doi:10.1038/srep25876

41. Has C, Sunthar P. A comprehensive review of recent liposome preparation techniques. J liposome research. 2020; 30(4): 336–365. doi:10.1080/08982104.2019.1668010

42. Akbarzadeh A, Rezaei-Sadabady R, Davaran S, etc. Liposomes: classification, preparation and application. Nano Res Lett. 2013;8(1):102. doi:10.1186/1556-276X-8-102

43. Filipe V, Hawe A, Jiskoot W. Critical evaluation of nanoparticle and protein aggregate measurement by nanosight with nanoparticle tracking analysis (NTA). Medical research. 2010;27(5):796-810. doi:10.1007/s11095-010-0073-2

44. Markones M, Drechsler C, Kaiser M, Kalie L, Heerklotz H, Fiedler S. Engineering asymmetric lipid vesicles: accurately and conveniently control the lipid composition of outer leaves. Langmuir. 2018;34(5):1999-2005. doi:10.1021/acs.langmuir.7b03189

45. Villasmil-Sánchez S, Rabasco AM, González-Rodríguez ML. Thermal and 31 P-NMR studies clarified the entrapment behavior of sumatriptan succinate in phosphatidylcholine/cholesterol liposomes. 31P-NMR comparative analysis of negatively charged and positively charged liposomes. Colloidal surfing B Biological interface. 2013; 105:14-23. doi:10.1016/j.colsurfb.2012.12.019

46. ​​Ghazal A, Gontsarik M, Kutter JP, etc. Microfluidic platform for the continuous production and characterization of multi-layered vesicles: synchrotron small-angle X-ray scattering (SAXS) research. J Phys Chem Lett. 2017;8(1):73-79. doi:10.1021/acs.jpclett.6b02468

47. Hoppe SM, Sasaki DY, Kinghorn AN, Hattar K. In situ transmission electron microscopy of liposomes in an aqueous environment. Langmuir. 2013;29(32):9958–9961. doi:10.1021/la401288g

48. Cohen SM. Extended pain relief trial using Exparel(®) (a long-acting multivesicular liposome preparation of bupivacaine) infiltration: a phase IV health economic trial for adult patients undergoing open colectomy. J Pain Research. 2012; 5: 567-572. doi:10.2147/JPR.S38621

49. Kao HW, Lin -YY, Gwathney WJ, Hong K. Formulation and evaluation of multilamellar ropivacaine in pain management. International J Nanomedicine. 2019; 14: 7891-7901. doi:10.2147/IJN.S215952

50. Liu D, Mori A, Huang L. The role of liposome size and RES blockade in controlling the biodistribution of GM1-containing liposomes and tumor uptake. Biochim Journal of Biophysics. 1992;1104(1):95-101. doi:10.1016/0005-2736(92)90136-A

51. Jason General Manager, Hodgson DF. Opportunities and challenges in the application of commercial drug liposomes. Adv Drug Del Rev. 2020; 154–155:2–12. doi:10.1016/j.addr.2020.07.016

52. Robinson T. Microfluidic processing and analysis of giant vesicles used as artificial cells: a review. Adv biological system. 2019;3(6):1800318. doi:10.1002/adbi.201800318

53. Briuglia ML, Rotella C, McFarlane A, Lamprou DA. The effect of cholesterol on liposome stability and drug release in vitro. Drug Deliv Transl Res. 2015; 5(3): 231–242. doi:10.1007/s13346-015-0220-8

54. Sato Y, Hatakeyama H, Sakurai Y, Hyodo M, Akita H, Harashima H. pH-sensitive cationic lipids promote liposomal siRNA delivery and gene silencing activity in vitro and in vivo. J Control release. 2012;163(3):267–276. doi:10.1016/j.jconrel.2012.09.009

55. Moon JJ, Suh H, Bershteyn A, etc. Double-layer cross-linked multilamellar vesicles are used as synthetic vaccines for effective humoral and cellular immune responses. Nat. 2011;10(3):243–251. doi:10.1038/nmat2960

56. Zhang Y, Mintzer E, Uhrich KE. Synthesis and characterization of pegylated bolaamphiphiles with enhanced retention in liposomes. J Colloid Interface Science. 2016; 482: 19-26. doi:10.1016/j.jcis.2016.07.013

57. Caddeo C, Pucci L, Gabriele M, etc. Stability, biocompatibility and antioxidant activity of PEG-modified liposomes containing resveratrol. Int J Pharm. 2018;538(1–2):40–47. doi:10.1016/j.ijpharm.2017.12.047

58. Lombardo D, Calandra P, Barreca D, Magazù S, Kiselev MA. Soft interactions in liposomal nanocarriers for therapeutic drug delivery. nanomaterials. 2016;6(7):125. doi:10.3390/nano6070125

59. Narenji M, Talaee MR, Moghimi HR. The effect of charge on liposome separation during stagnation. Iran J Pharm Res. 2017; 16(2): 423–431.

60. Clogston JD, Patri AK. Zeta potential measurement. In: Characterization of Nanoparticles for Drug Delivery. Humanities Press; 2011:63-70.

61. Gonzalez Gomez A, Syed S, Marshall K, Hosseinidoust Z. Liposome nanovesicles for effective encapsulation of staphylococcal antibiotics. ACS Omega. 2019; 4(6): 10866-10876. doi:10.1021/acsomega.9b00825

62. Ruysschaert T, Marque A, Duteyrat JL, Lesieur S, Winterhalter M, Fournier D. Liposome retention in size exclusion chromatography. BMC Biotechnology Company. 2005; 5:11. doi:10.1186/1472-6750-5-11

63. Ran C, Chen D, Xu M, Du C, Li Q, Jiang Y. Research on the characteristics of different sample pretreatment methods to evaluate the encapsulation efficiency of liposomes. J Chromatogr B. 2016; 1028:56-62. doi:10.1016/j.jchromb.2016.06.008

64. Zhang XM, Patel AB, de Graaf RA, Behar KL. Proton nuclear magnetic resonance spectroscopy was used to determine the encapsulation efficiency of liposomes. Chemical and physical lipids. 2004;127(1):113-120. doi:10.1016/j.chemphyslip.2003.09.013

65. Lohse B, Bolinger PY, Stamou D. Encapsulation efficiency measured on a single small unilamellar vesicle. J Am Chem Soc. 2008;130(44):14372-14373. doi:10.1021/ja805030w

66. Sun B, Chiu DT. Single-vesicle photolysis and confocal single-molecule detection were used to determine the encapsulation efficiency of a single vesicle. Anal chemistry. 2005;77(9):2770-2776. doi:10.1021/ac048439n

67. Yamamoto E, Miyazaki S, Aoyama C, Kato M. By directly injecting the liposome suspension into the liquid chromatography, a simple and rapid method for measuring the encapsulation efficiency of doxorubicin-loaded liposomes. Int J Pharm. 2018;536(1):21–28. doi:10.1016/j.ijpharm.2017.11.035

68. Jahn A, Vreeland WN, Gaitan M, Locascio LE. Controlled vesicle self-assembly in a microfluidic channel with hydrodynamic focusing. J Am Chem Soc. 2004;126(9):2674-2675. doi:10.1021/ja0318030

69. Jahn A, Vreeland WN, DeVoe DL, Locascio LE, Gaitan M. Microfluidics oriented to form liposomes of controlled size. Langmuir. 2007;23(11):6289–6293. doi:10.1021/la070051a

70. Jahn A, Stavis SM, Hong JS, Vreeland WN, DeVoe DL, Gaitan M. Microfluidic mixing and formation of nanoscale lipid vesicles. ACS nano. 2010;4(4):2077-2087. doi:10.1021/nn901676x

71. Tien Sing Young RV, Tabrizian M. Nano-scale 1,2-distearoyl-sn-glycerol-3-phosphocholine liposomes are quickly manufactured and loaded in a simple two-flow focused microfluidic device in one step. Biological microfluidics. 2015; 9(4): 046501. doi:10.1063/1.4926398

72. Mijajlovic M, Wright D, Zivkovic V, Bi JX, Biggs MJ. Microfluidic hydrodynamic focusing-based synthesis of POPC liposomes for model biological systems. Colloidal surfing B Biological interface. 2013; 104: 276-281. doi:10.1016/j.colsurfb.2012.12.020

73. Zizzari A, Bianco M, Carbone L, etc. Continuous production of injectable liposomes by microfluidic method. Material. 2017;10(12):1411. doi:10.3390/ma10121411

74. Jahn A, Lucas F, Wepf RA, Dittrich PS. Freezing continuous flow self-assembly in a microfluidic device: imaging of liposome formation. Langmuir. 2013;29(5):1717–1723. doi:10.1021/la303675g

75. Balbino TA, Aoki NT, Gasperini AAM, etc. Cationic liposomes are continuously produced at high lipid concentrations in microfluidic devices for gene delivery applications. Journal of Chemical Engineering 2013;226:423-433. doi:10.1016/j.cej.2013.04.053

76. Aghaei H, Solaimany Nazar AR. Continuous production of nano-scale liposomes in a dual-flow focused microfluidic device. Ind Eng Chem Res. 2019;58(51):23032–23045. doi:10.1021/acs.iecr.9b04079

77. Hood RR, DeVoe DL, Atencia J, Vreeland WN, Omiatek DM. An easy way to synthesize monodisperse nano-scale liposomes using 3D microfluidic hydrodynamics in concentric capillary arrays. Laboratory chip. 2014;14(14):2403–2409. doi:10.1039/C4LC00334A

78. Hood RR, DeVoe DL. The vertical flow of microfluidics focuses on high-throughput continuous flow to produce nano-scale liposomes. small. 2015; 11(43): 5790-5799. doi:10.1002/smll.201501345

79. Michelon M, Oliveira DRB, de Figueiredo Furtado G, Gaziola de la Torre L, Cunha RL. High-throughput continuous production of liposomes using hydrodynamic flow-focused microfluidic devices. Colloidal surfing B Biological interface. 2017; 156: 349-357. doi:10.1016/j.colsurfb.2017.05.033

80. Chen Z, Han JY, Shumate L, Fedak R, DeVoe DL. Formation of high-throughput nanoliposomes using 3D printed microfluidic flow focusing chips. Adv Mater Technol. 2019;4(6):1800511. doi:10.1002/admt.201800511

81. Balbino TA, Azzoni AR, de la Torre LG. Microfluidic device for continuous production of pDNA/cationic liposome complexes for gene delivery and vaccine therapy. Colloidal surfing B Biological interface. 2013; 111:203-210. doi:10.1016/j.colsurfb.2013.04.003

82. Ran R, Middelberg APJ, Zhao CX. Microfluidic synthesis of multifunctional liposomes for tumor targeting. Colloidal surfing B Biological interface. 2016; 148: 402-410. doi:10.1016/j.colsurfb.2016.09.016

83. Aghaei H, Nazar ARS, Varshosaz J. Preparation of methotrexate-loaded liposome nanoparticles based on dual-stream focused microfluidics: encapsulation efficacy, drug release and stability. Colloid surfing physical and chemical engineering. 2021;614:126166. doi:10.1016/j.colsurfa.2021.126166

84. Hood RR, Vreeland WN, DeVoe DL. Microfluidic remote loading for rapid single-step liposomal drug preparation. Laboratory chip. 2014;14(17):3359-3367. doi:10.1039/C4LC00390J

85. Sur S, Fries AC, Kinzler KW, Zhou S, Vogelstein B. Remote loading of pre-encapsulated drugs into invisible liposomes. Proc Natl Acad Sci. 2014;111(6):2283. doi:10.1073/pnas.1324135111

86. Stroock AD, Dertinger SKW, Ajdari A, Mezić I, Stone HA, Whitesides GM. Chaos mixer for microchannels. science. 2002;295(5555):647. doi:10.1126/science.1066238

87. Zhigaltsev IV, Belliveau N, Hafez I, etc. Using millisecond microfluidic mixing bottom-up design and synthesis of extreme size lipid nanoparticle systems with water and triglyceride cores. Langmuir. 2012;28(7):3633–3640. doi:10.1021/la204833h

88. Kastner E, Kaur R, Lowry D, Moghaddam B, Wilkinson A, Perrie Y. Use enhanced statistical tools for characterization and high-throughput production of size-regulated liposomes through a new microfluidic approach. Int J Pharm. 2014;477(1):361–368. doi:10.1016/j.ijpharm.2014.10.030

89. Maeki M, Saito T, Sato Y, etc. A strategy for synthesizing lipid nanoparticles using a microfluidic device with a mixer structure. RSC Advanced 2015; 5(57): 46181–46185. doi:10.1039/C5RA04690D

90. Lou G, Anderluzzi G, Woods S, Roberts CW, Perrie Y. A novel method based on microfluidics to formulate large unilamellar cationic liposomes with adjustable size: formulation, cellular uptake and biodistribution studies. Eur J Pharma Biopharm. 2019;143:51-60. doi:10.1016/j.ejpb.2019.08.013

91. Maeki M, Fujishima Y, Sato Y, etc. Understand the formation mechanism of lipid nanoparticles in microfluidic devices with chaotic micromixers. Public Science Library One. 2017;12(11):e0187962. doi:10.1371/journal.pone.0187962

92. Joshi S, Hussain MT, Roces CB, etc. Microfluidic-based liposome manufacturing simultaneously encapsulates hydrophilic and lipophilic drugs. Int J Pharm. 2016;514(1):160–168. doi:10.1016/j.ijpharm.2016.09.027

93. Tenchov R, Bird R, Curtze AE, Zhou Q. Lipid nanoparticles-from liposomes to mRNA vaccine delivery, to study the landscape of diversity and progress. ACS nano. 2021. doi:10.1021/acsnano.1c04996

94. Mitchell MJ, Billingsley MM, Haley RM, Wechsler ME, Peppas NA, Langer R. Engineered precision nanoparticles for drug delivery. Nat Rev drug discovery. 2021;20(2):101–124. doi:10.1038/s41573-020-0090-8

95. Kimura N, Maeki M, Sato Y, etc. Development of iLiNP device: Fine-tune lipid nanoparticle size within 10 nm for drug delivery. ACS Omega. https://pubs.acs.org/doi/10.1021/acsomega.8b00341. 2018; 3(5): 5044–5051.

96. Rasouli MR, Tabrizian M. An ultra-fast acoustic micromixer for the synthesis of organic nanoparticles. Laboratory chip. 2019;19(19):3316–3325. doi:10.1039/C9LC00637K

97. Modarres P, Tabrizian M. Electrohydrodynamics-driven micromixing for the synthesis of highly monodisperse nano-scale liposomes. ACS applies nanomaterials. 2020;3(5):4000-4013. doi:10.1021/acsanm.9b02407

98. Brittany M, Amirkavei M, Mir LM. Optimization of the electroformation of giant unilamellar vesicles (GUV) with unsaturated phospholipids. J Member Biology. 2015;248(5):827–835. doi:10.1007/s00232-015-9828-3

99. Runas KA, Malmstadt N. The low level of lipid oxidation fundamentally increases the passive permeability of the lipid bilayer. Soft matter. 2015; 11(3): 499–505. doi:10.1039/C4SM01478B

100. Kastner E, Verma V, Lowry D, Perrie Y. Manufacture of liposomes controlled by microfluidics, used to dissolve drugs that are difficult to dissolve in water. Int J Pharm. 2015;485(1):122–130. doi:10.1016/j.ijpharm.2015.02.063

101. Chen D, Love KT, Chen Y, etc. Quickly discover effective lipid nanoparticles containing siRNA through a controlled microfluidic formulation. J Am Chem Soc. 2012;134(16):6948-6951. doi:10.1021/ja301621z

102. Belliveau NM, Huft J, Lin PJ, etc. Microfluidic synthesis of high-efficiency extreme size lipid nanoparticles for in vivo delivery of siRNA. Mol Ther nucleic acid. 2012;1(8):e37. doi:10.1038/mtna.2012.28

103. Leung AKK, Hafez IM, Baoukina S, etc. Lipid nanoparticles containing siRNA synthesized by microfluidic mixing exhibit an electronically dense nanostructured core. J Phys Chem C. 2012;116(34):18440–18450. doi:10.1021/jp303267y

104. Shepherd SJ, Warzecha CC, Yadavali S, etc. Use parallel microfluidic devices to produce scalable mRNA and siRNA lipid nanoparticles. Nanolet. 2021; 21(13): 5671-5680. doi:10.1021/acs.nanolett.1c01353

105. Rosenblum D, Gutkin A, Kedmi R, etc. CRISPR-Cas9 genome editing for cancer therapy using targeted lipid nanoparticles. Advanced Science 2020; 6(47): eabc9450. doi:10.1126/sciadv.abc9450

106. Riley RS, Kashyap MV, Billingsley MM, etc. Ionizable lipid nanoparticles for in-utero mRNA delivery. Advanced Science 2021; 7(3). doi:10.1126/sciadv.aba1028.

107. Weiss M, Frohnmayer JP, Benk LT, etc. Continuous bottom-up assembly of mechanically stable synthetic cells through microfluidics. Nat. 2018;17(1):89-96. doi:10.1038/nmat5005

108. Abate AR, Hung T, Mary P, Agresti JJ, Weitz DA. Microfluidic high-throughput injection using picojectors. Proc Natl Acad Sci. 2010;107(45):19163. doi:10.1073/pnas.1006888107

109. Staufer O, Antona S, Zhang D, etc. Microfluidic production and characterization of bio-functionalized giant unilamellar vesicles for targeted intracellular cargo delivery. biomaterials. 2021;264:120203. doi:10.1016/j.biomaterials.2020.120203

110. Sugiura S, Kuroiwa T, Kagota T, etc. A new method for obtaining uniform giant vesicles from monodisperse water-in-oil emulsions prepared by microfluidic devices. Langmuir. 2008;24(9):4581–4588. doi:10.1021/la703509r

111. Kuroiwa T, Fujita R, Kobayashi I, etc. Efficient preparation of giant vesicles as a biomimetic compartment system has a high capture rate for biological macromolecules. Chemical biology diver. 2012; 9(11): 2453-2472. doi:10.1002/cbdv.201200274

112. Pautot S, Frisken BJ, Weitz DA. Use inverted emulsion to produce unilamellar vesicles. Langmuir. 2003;19(7):2870–2879. doi:10.1021/la026100v

113. Tan YC, Hettiarachchi K, Siu M, Pan YR, Lee AP. Controlled microfluidic encapsulation of cells, proteins and microbeads in lipid vesicles. J Am Chem Soc. 2006;128(17):5656-5658. doi:10.1021/ja056641h

114. Hu PC, Li S, Malmstadt N. Microfluidic manufacturing of asymmetric giant lipid vesicles. ACS application program interface. 2011; 3(5): 1434–1440. doi:10.1021/am101191d

115. Nishimura K, Suzuki H, Toyota T, Yomo T. Size control of giant unilamellar vesicles prepared from inverted emulsion droplets. J Colloid Interface Science. 2012;376(1):119-125. doi:10.1016/j.jcis.2012.02.029

116. Matosevic S, Paegel BM. The huge unilamellar vesicles are gradually synthesized on the microfluidic assembly line. J Am Chem Soc. 2011;133(9):2798-2800. doi:10.1021/ja109137s

117. Karamdad K, Law RV, Seddon JM, Brooks NJ, Ces O. Preparation and mechanical characterization of giant unilamellar vesicles by microfluidic methods. Laboratory chip. 2015; 15(2): 557–562. doi:10.1039/C4LC01277A

118. Karamdad K, Law RV, Seddon JM, Brooks NJ, Ces O. Investigate the influence of asymmetry on the flexural stiffness of the lipid membrane formed by microfluidics. Chemical Commune. 2016;52(30):5277–5280. doi:10.1039/C5CC10307J

119. Matosevic S, Paegel BM. Assemble the cell membrane layer by layer. National Chemistry 2013; 5(11): 958-963. doi:10.1038/nchem.1765

120. Morita M, Onoe H, Yanagisawa M, etc. Droplet ejection and size filtration (DSSF) method for synthesizing cell-sized liposomes with controlled lipid composition. Chemistry Biochemistry. 2015;16(14):2029-2035. doi:10.1002/cbic.201500354

121. Morita M, Katoh K, Noda N. Direct observation of bacterial growth in giant unilamellar vesicles: a new tool for bacterial culture. Chemistry opens. 2018; 7(11): 845–849. doi:10.1002/open.201800126

122. Adir O, Sharf-Pauker N, Chen G, etc. Use cell-free bacterial extracts, liposomes, and emulsion transfer to prepare protein-producing synthetic cells. Jupiter. 2020;158:e60829.

123. Kubatta EA, Rehage H. Characterization of giant vesicles formed during phase transfer. Colloidal Polymer Science. 2009;287(9):1117–1122. doi:10.1007/s00396-009-2083-3

124. Shum HC, Lee D, Yoon I, Kodger T, Weitz DA. Double emulsion templated monodisperse phospholipid vesicles. Langmuir. 2008;24(15):7651–7653. doi:10.1021/la801833a

125. Kim SH, Kim JW, Cho JC, Weitz DA. Double emulsion drops with ultra-thin shell for capsule template. Laboratory chip. 2011;11(18):3162–3166. doi:10.1039/C1LC20434C

126. Arriaga LR, Datta SS, Kim SH, etc. The ultra-thin shell double emulsion templated giant unilamellar lipid vesicles with controllable microdomain formation. small. 2014; 10(5): 950–956. doi:10.1002/smll.201301904

127. Caschera F, Lee JW, Ho KKY, Liu AP, Jewett MC. Cell-free compartmentalized protein synthesis is carried out in double emulsion template liposomes with in vitro synthesis and assembly of ribosomes. Chemical Commune. 2016;52(31):5467–5469. doi:10.1039/C6CC00223D

128. Costa C, Liu Z, Simões SI, etc. One-step microfluidic production of enzyme-loaded liposomes for the treatment of inflammatory diseases. Colloidal surfing B Biological interface. 2021; 199: 111556. doi:10.1016/j.colsurfb.2020.111556

129. Teh SY, Khnouf R, Fan H, Lee AP. Stable and biocompatible lipid vesicles are generated through the droplet microfluidic technology based on solvent extraction. Biological microfluidics. 2011; 5(4): 44113–4411312. doi:10.1063/1.3665221

130. Petit J, Polenz I, Baret JC, Herminghaus S, Bäumchen O. Vesicles-on-a-chip: a universal microfluidic platform for assembling liposomes and polymer bodies. Eur Phys J E. 2016; 39(6):59. doi:10.1140/epje/i2016-16059-8

131. Lu L, Doak WJ, Schertzer JW, Chiarot PR. Membrane mechanical properties of synthetic asymmetric phospholipid vesicles. Soft matter. 2016; 12(36): 7521-7528. doi:10.1039/C6SM01349J

132. Arriaga LR, Huang Y, Kim SH, etc. One-step assembly of asymmetric vesicles. Laboratory chip. 2019;19(5):749–756. doi:10.1039/C8LC00882E

133. Lu L, Schertzer JW, Chiarot PR. Continuous microfluidic fabrication of synthetic asymmetric vesicles. Laboratory chip. 2015; 15(17): 3591-3599. doi:10.1039/C5LC00520E

134. Lee JN, Park C, General Manager of Whitesides. Solvent compatibility of poly(dimethylsiloxane)-based microfluidic devices. Anal chemistry. 2003;75(23):6544-6554. doi:10.1021/ac0346712

135. Deshpande S, Caspi Y, Meijering AE, Dekker C. Captanol-assisted liposome chip assembly. Nat community. 2016; 7: 10447. doi:10.1038/ncomms10447

136. Deshpande S, Dekker C. On-chip microfluidic production of cell-sized liposomes. National agreement. 2018;13(5):856–874. doi:10.1038/nprot.2017.160

137. Deng-NN, Yelleswarapu M, Hack WTS. Monodisperse unilamellar and multilamellar liposomes. J Am Chem Soc. 2016;138(24):7584–7591. doi:10.1021/jacs.6b02107

138. Schaich M, Cama J, Al Nahas K, etc. An integrated microfluidic platform for quantifying drug penetration across biomimetic vesicle membranes. Moore Pharmaceuticals. 2019;16(6):2494–2501. doi:10.1021/acs.molpharmaceut.9b00086

139. Vaezi Z, Sedghi M, Ghorbani M, Shojaeilangari S, Allahverdi A, Naderi-Manesh H. Using a microfluidic platform to study programmed cell death caused by liposomes encapsulating cytoskeletal drugs. Microfluidic nanofluid. 2020;24(7):48. doi:10.1007/s10404-020-02353-3

140. Niederholtmeyer H, Chaggan C, Devaraj NK. Communication and quorum sensing in non-life simulants of eukaryotic cells. Nat community. 2018;9(1):5027. doi:10.1038/s41467-018-07473-7

141. Deshpande S, Brandenburg F, Lau A, etc. Temporal and spatial control of coacervate formation in liposomes. Nat community. 2019;10(1):1800. doi:10.1038/s41467-019-09855-x

142. Funakoshi K, Suzuki H, Takeuchi S. A giant lipid vesicle-like compartment is formed from a planar lipid membrane by pulsed jets. J Am Chem Soc. 2007;129(42):12608-12609. doi:10.1021/ja074029f

143. Funakoshi K, Suzuki H, Takeuchi S. Form a lipid bilayer by contacting a single layer in a membrane protein analysis microfluidic device. Anal chemistry. 2006;78(24):8169–8174. doi:10.1021/ac0613479

144. Kamiya K, Kawano R, Osaki T, Akiyoshi K, Takeuchi S. Cell-sized asymmetric lipid vesicles facilitate the study of asymmetric membranes. National Chemistry 2016; 8(9): 881–889. doi:10.1038/nchem.2537

145. Kamiya K, Osaki T, Takeuchi S. Use pulse jet method to form vesicles in vesicles with asymmetric lipid components. RSC Advanced 2019;9(52):30071–30075. doi:10.1039/C9RA04622D

146. Kamiya K, Osaki T, Takeuchi S. Use pulse jet method to form nano-sized lipid vesicles with asymmetric lipid composition. Sensor Actuator B: Chemistry. 2021;327:128917. doi:10.1016/j.snb.2020.128917

147. Stachowiak JC, Richmond DL, Li TH, Liu AP, Parekh SH, Fletcher DA. Unilamellar vesicle formation and microfluidic jet encapsulation. Proc Natl Acad Sci. 2008;105(12):4697. doi:10.1073/pnas.0710875105

148. Stachowiak JC, Richmond DL, Li TH, Brochard-Wyart F, Fletcher DA. Inkjet formation of unilamellar lipid vesicles for cell-like encapsulation. Laboratory chip. 2009;9(14):2003-2009. doi:10.1039/b904984c

149. Richmond DL, Schmid EM, Martens S, Stachowiak JC, Liska N, Fletcher DA. Form huge vesicles with controlled membrane composition, asymmetry and content. Proc Natl Acad Sci. 2011;108(23):9431. doi:10.1073/pnas.1016410108

150. Gotanda M, Kamiya K, Osaki T, Miki N, Takeuchi S. Automatic generation system for cell-sized liposomes. Sensor Actuator B: Chemistry. 2019;292:57-63. doi:10.1016/j.snb.2019.04.096

151. Armstrong M, Wach, MD, Hunter TP, Fletcher DA. Acoustic jets are used to form and load huge unilamellar vesicles. Biological microfluidics. 2020;14(6):064105. doi:10.1063/5.0021742

152. Kirchner SR, Ohlinger A, Pfeiffer T, etc. Membrane composition of jet lipid vesicles: Raman spectroscopy study. J Biophotonics. 2012; 5(1): 40–46. doi:10.1002/jbio.201100058

153. Reeves JP, Dowben RM. The formation and properties of thin-walled phospholipid vesicles. J Cell Physiology. 1969;73(1):49-60. doi:10.1002/jcp.1040730108

154. Rodriguez N, Pincet F, Cribier S. Giant vesicles formed by gentle hydration and electroformation: a fluorescence microscope comparison. Colloidal surfing B Biological interface. 2005;42(2):125–130. doi:10.1016/j.colsurfb.2005.01.010

155. Zhang H. Preparation of liposomes by extrusion after film hydration. In: liposomes. New York, New York: Humana Press; 2017:17-22.

156. Osaki T, Kuribayashi-Shigetomi K, Kawano R, Sasaki H, Takeuchi S. Giant liposomes of uniform size are formed and are gently hydrated. In: 2011 IEEE 24th International Conference on Microelectromechanical Systems; 2011; IEEE.

157. O'Neill CP, Suzuki T, Demurtas D, Finca A, Hubbel JA. A new method of encapsulating biomolecules into polymer vesicles through direct hydration. Langmuir. 2009;25(16):9025-9029. doi:10.1021/la900779t

158. Herold C, Chwastek G, Schwille P, Petrov EP. Efficient electroformation of super-giant unilamellar vesicles containing cationic lipids on ITO-coated electrodes. Langmuir. 2012;28(13):5518–5521. doi:10.1021/la3005807

159. Kuribayashi K, Tresset G, Coquet P, Fujita H, Takeuchi S. Electric formation of giant liposomes in microfluidic channels. Measurement science and technology. 2006;17(12):3121–3126. doi:10.1088/0957-0233/17/12/S01

160. Taylor P, Xu C, Fletcher PDI, Paunov VN. A new technology for preparing monodisperse giant liposomes. Chemical Commune. 2003; (14): 1732-1733. doi:10.1039/b304059c

161. Taylor P, Xu C, Fletcher PDI, Paunov VN. A two-dimensional array of giant liposomes was prepared by microcontact printing on a solid substrate. Physical Chemistry Chemistry Physics. 2003;5(21):4918-4922. doi:10.1039/b308082j

162. Nafday OA, Lowry TW, Lenhert S. Multifunctional lipid multilayer stamping. small. 2012;8(7):1021–1028. doi:10.1002/smll.201102096

163. Le Berre M, Yamada A, Reck L, Chen Y, Baigl D. Electroforming of giant phospholipid vesicles on silicon substrates: the advantages of controllable surface properties. Langmuir. 2008;24(6):2643–2649. doi:10.1021/la703391q

164. Diguet A, Le Berre M, Chen Y, Baigl D. Fabrication of phospholipid multilayer patterns with controlled size and thickness by capillary assembly on a microstructure substrate. small. 2009;5(14):1661–1666. doi:10.1002/smll.200900368

165. Wang Z, Wu C, Fan Tao, et al. The giant liposomes are formed and collected by electricity on the integrated microchip. China Chemical News. 2019;30(2):353–358. doi:10.1016/j.cclet.2018.12.001

166. Bi H, Yang B, Wang L, Cao W, Han X. Use interdigital ITO electrodes to electroform giant unilamellar vesicles. J Mater Chem A. 2013;1(24):7125-7130. doi:10.1039/c3ta10323d

167. Steinkühler J, De Tillieux P, Knorr RL, Lipowsky R, Dimova R. The charged giant unilamellar vesicles prepared by electroforming showed nanotubes and lipid asymmetry across the bilayer. Sci Rep. 2018;8(1):11838. doi:10.1038/s41598-018-30286-z

168. Witkowska A, Jablonski L, Jahn R. A convenient solution for using electroforming to generate giant unilamellar vesicles containing SNARE protein. Sci Rep. 2018;8(1):9422. doi:10.1038/s41598-018-27456-4

169. Cheng L, Wang Y, Sun G, etc. Hydration enhanced lubricating electrospun nanofiber membrane can prevent tissue adhesion. Research. 2020; 2020: 4907185. doi:10.34133/2020/4907185

170. Deshpande S, Birnie A, Dekker C. Liposome purification based on chip density. Biological microfluidics. 2017;11(3):034106. doi:10.1063/1.4983174

171. Dimov N, Kastner E, Hussain M, Perrie Y, Szita N. Formation and purification of custom liposomes for drug delivery using a module-based micro-continuous flow system. Scientific Reports 2017; 7(1): 12045. doi:10.1038/s41598-017-11533-1

172. Haller B, Göpfrich K, Schröter M, Janiesch JW, Platzman I, Spatz JP. Charge controlled microfluidic formation in lipid-based single- and multi-chamber systems. Laboratory chip. 2018;18(17):2665–2674. doi:10.1039/C8LC00582F

173. Birnbaumer G, Küpcü S, Jungreuthmayer C, etc. Use lab-on-a-chip for rapid liposome quality assessment. Laboratory chip. 2011;11(16):2753–2762. doi:10.1039/c0lc00589d

174. Ran R, Wang HF, Hou F, et al. Microfluidic tumor chip for evaluating tumor targeting and anticancer efficacy of multifunctional liposomes. Adv healthcare materials. 2019;8(8):1900015. doi:10.1002/adhm.201900015

175. Muthu MS, Kulkarni SA, Raju A, Feng SS. Therapeutic and diagnostic liposomes for the targeted co-delivery of docetaxel and quantum dots with TPGS coating. biomaterials. 2012;33(12):3494-3501. doi:10.1016/j.biomaterials.2012.01.036

176. Keck CM, Specht D, Brüßler J. The influence of lipid matrix composition on the biopharmaceutical properties of lipid nanoparticles. J Control release. 2021;338:149-163. doi:10.1016/j.jconrel.2021.08.016

177. Liu Z, Fontana F, Python A, Hirvonen JT, Santos HA. Microfluidics: Microfluidics for particle production: mechanisms, methods and applications (Small 9/2020). small. 2020;16(9):2070048. doi:10.1002/smll.202070048

178. Qiu M, Glass Z, Chen J, et al. The cooperative delivery of Cas9 mRNA and single guide RNA mediated by lipid nanoparticles realizes the liver-specific in vivo genome editing of Angptl3. Proc Natl Acad Sci US A. 2021;118(10):e2020401118. doi:10.1073/pnas.2020401118

179. Elia U, Ramishetti S, Rosenfeld R, etc. Design of SARS-CoV-2 hFc coupled receptor binding domain mRNA vaccine delivered by lipid nanoparticles. ACS nano. 2021; 15(6): 9627-9637. doi:10.1021/acsnano.0c10180

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