*2.1.* μ*Ps Fabrication by Advanced Processes*

Several conventional methods are used to prepare μPs, such as phase separation, spray drying, and batch emulsion techniques. The resulting polymeric material is often characterized by heterogeneous size distribution and limited control over their shape [49,50]. To overcome these limitations and achieve even highly complex μPs morphology and composition, new advanced techniques were implemented for μPs' fabrication in the past decades, as highlighted in Figure 1. Microfluidics is a revolutionary technique that manipulates fluids into microscale channels for fluid mixing, merging, splitting, and reaction [51]. Microfluidic emulsion is one of the most investigated techniques for the high-throughput production of monodisperse modular microstructures variable in size, shape, and composition. Microfluidic devices are generally made of transparent and chemically strong

devices obtained by assembling glass capillaries or patterning channels in a silicone elastomer, e.g., polydimethylsiloxane (PDMS), through soft lithography methods (Figure 1a,b) [52]. Oil-in-water single emulsions generated with the assistance of the co-flow devices schematized in Figure 1a,b enabled the production of uniform and size-controlled droplets by simple modulation of flow rate of continuous and disperse phases. These droplets can be conveniently converted into uniform beads (Figure 1c) or beads with core-shell, patchy, and Janus architectures (Figure 1d) by the adequate choice of solutions' compositions and controlling the solvent evaporation and phase separation mechanism [45,53]. For instance, using PLGA/PCL as model materials, the authors showed that the core-shell, patchy, and Janus types of particles can be produced with a high yield and a narrow size distribution by precisely controlling interfacial tensions and spreading coefficients between immiscible phases of the generated droplets. As a direct consequence, μPs' hydrophilicity, degradation rate, and drug delivery properties can be tuned depending on the specific application [45].

The formation of multiple emulsions within these microfluidic devices may enable the fabrication of μPs with multiples cores and drug delivery capability (Figure 1e). More importantly, these methods are able to improve loading efficiency of hydrophobic polymeric μPs by changing their polarity [54]. Microfluidic flow-focusing devices were fabricated to generate droplets of different sizes and shapes and narrow size distribution in either PDMS or glass capillary devices [55]. Indicating the diameter of an undeformed (regular) spherical droplet as dS = (6V/π)1/3, non-spherical droplets are obtained when dS is larger than at least one of the dimensions of the outlet channel, as the confinement hinder shape relaxation of droplets into spheres after breakup. As a direct consequence, in wide channels, when w > dS (w = width) while the height h < dS, the drops assume a discoid shape with rounded borders (Figure 1f). For channels with both h and w smaller than dS, the droplet makes contact with all channel walls and assumes a rod-like morphology (Figure 1g). As shown in Figure 1h, microfluidic approaches were also used for preparing porous μPs with a large surface area, good mechanical strength, and high interconnectivity to be suitable as μ-scaffolds for cells' culture [53]. This was achieved by injecting an unstable water-in-oil emulsion, made of gelatin and poly (vinyl alcohol) (PVA) as discontinuous phase in a PLGA solution in dichloromethane that served as continuous phase. The resultant water-oil-in-water droplets were subsequently solidified by solvent extraction and evaporation in a collection phase (water), generating porous μPs.

Lithography-based processes, such as those reported in Figure 1, are the subsequent example to manufacture precisely shaped polymeric μPs. Flow-lithography processes (e.g., continuous or stop-flow-lithography) enable for continuously synthesizing a variety of different shapes and sizes using several oligomers and produce multifunctional Janus particles. These approaches use ultraviolet (UV) light combined with light-transparent PDMS devices for the selective photopolymerization of a fluidic bead with the aid of proper masks (Figure 1i). Particles' shape in the x–y plane is determined by the transparency masks pattern (Figure 1j–l); whereas the z-plane projection is dependent on the height of the channel used and the thickness of the oxygen inhibition layer [47]. For instance, polyethylene glycol diacrylate (PEGDA) microgels were synthesized with tunable shapes such as triangles, squares, and hexagons, showing good fidelity to the original mask features (Figure 1j–l) [47]. The fundamental limitations of the flow-lithography technique are mainly dependent on the optical resolution and the depth-of-field of the microscope objective used as well, as it requires a short polymerization time or slow flow rate to avoid smearing of the patterned feature in the hydrogels. The main limitation of this technique is the use of prepolymer solutions with high concentrations of monomer and/or photoinitiator, necessary for reducing μPs' setting time that may induce a possible cytotoxic effect. A stop-flow-lithography (SFL) process was recently proposed to overcome this limitation. This process involves stopping the liquid flow, polymerizing the patterned solution, and the flowing of the particles out of the device. This workflow proved suitable for fabricating cell-laden PEGDA particles with controlled shapes and size for TE (Figure 1m,n) [56]. Nevertheless, flow-lithography processes are mainly limited to materials that can polymerize under UV light (hydrogels) and therefore, cannot be used for synthetic materials such as thermoplastic polymers.

Recent advances in micro/nanotechnology have allowed fabrication of μPs made of thermoplastic polymers with uniform sizes and well-defined shapes and composition, which are otherwise impossible to fabricate using conventional μPs' manufacturing methods, providing new building blocks libraries for modular TE. In particular, soft-lithography techniques involve the use of elastomeric PDMS stamps with topological microfeatures to fabricate μPs with precise control over size and geometry in a simple, versatile, and cost-effective modality (Figure 1o) [57–59]. After solvent evaporation, the dried polymer is deposited on selective portions of the mold in the form of particles, and it is removed from the PDMS mold by stamping it onto a PVA sacrificial layer at temperatures and pressures in the range of 80–120 ◦C and 30–90 KPa, respectively [58]. The μPs are released from the mold by dissolving the PVA layer in water. The versatility of these fabrication methods has been demonstrated using materials of biomedical interest including thermoplastic polymers such as PCL and PLGA (Figure 1p–r) [59], polyethylene glycol dimethacrylate (PEGDMA) hydrogels, and chitosan. An advancement in this fabrication technique was reported recently by McHugh and co-workers that developed a microfabrication method, termed StampEd Assembly of polymer Layers (SEAL), for fabricating modular micrometric structures, such as injectable pulsatile drug-delivery PLGA μPs with complex geometry at a high resolution (Figure 1s,t) [60]. In another study, de Alteriis and co-workers used microspheres to obtain shaped μPs by a soft-lithography approach [46]. This was achieved by positioning PLGA microspheres into PDMS mold cavities with different shapes and deforming them under gentle process conditions, i.e., at room temperature using a solvent/non-solvent vapor mixture. By this approach, it was also possible to preserve the microstructure and bioactivity of molecules loaded inside the μPs (Figure 1u). In conclusion, all of the discussed advanced μPs' fabrication methods may open new avenues for the fabrication of multifunctional building blocks for modular TE applications.

**Figure 1.** Microfluidic emulsion: Fabrication of microparticles (μPs) with advanced processes. (**a**) Co-flow and (**b**) flow-focusing pictures of fluidic emulsion devices. Effect of processing conditions on μPs morphology, composition and structure: (**c**) spherical monodisperse μPs, (**d**) Janus μPs, (**e**) core-shell μPs with dual and triple cores, (**f**) disks and (**g**) rods μPs obtained by controlling the dimension of the outlet channel, (**h**) highly porous polylactic-co-glycolic acid (PLGA) spherical μPs prepared by double emulsion. Flow-lithography: (**i**) Picture of the flow-lithography continuous process for making shape-controlled μPs by exposure of precursor solution to patterned ultraviolet (UV) light. Morphology of (**j**) triangles, (**k**) squares and (**l**) hexagons μPs prepared by the continuous flow-lithography process. Single-cell encapsulated within (**m**) square and (**n**) triangular μPs prepared by the stop-flow-lithography (SFL) process. Soft-lithography: (**o**) Schematic drawing of the soft-lithography and lift-out molding fabrication protocol of μPs: (**p**,**q**,**r**) effect of mold type on μPs shape. Morphology of μPs obtained by the StampEd Assembly of polymer Layers (SEAL) process before (**s**) and (**t**) after sealing. (**u**) Morphology of vascular endothelial growth factor (VEGF)-loaded PLGA microsphere after solvent vapor shaping

process. **c**, **f**, **g** Reproduced with permission from Reference [55] (Xu, Angewandte Chemie International Edition; Published by John Wiley and Sons, 2005); **d** Reproduced with permission from Reference [45] (Cao, RCS. Advances; published by Royal Society of Chemistry, 2015); **e**, **i** Reproduced with permission from Reference [54] (Baah, Microfluid Nanofluid; published by Springer Nature, 2014); **h** Reproduced with permission from Reference [53] (Choi, Small; published by John Wiley and Sons, 2010); **j**, **l** Reproduced with permission from Reference [47] (Dendukuri, Nature Materials; published by Springer Nature, 2006); **m**, **n** Reproduced with permission from Reference [56] (Panda, Lab Chip; published by Royal Society of Chemistry, 2008); **o** Reproduced with permission from Reference [57] (Canelas, Nanomed Nanobiotechnol; published by John Wiley and Sons, 2009); **p**, **q**, **r** Reproduced with permission from Reference [59] (Guan, Biomaterials; published by Elsevier Ltd, 2006); **s**, **t** Reproduced with permission from Reference [60] (Kevin J. McHugh, Science; published by American Association for the Advancement of Science, 2017); **u** Reproduced with permission from Reference [46] (Renato de Alteriis, Scientific Reports; published by Springer Nature, 2015).
