**2.4 Polymer as a stabilizing agent in the formation of microbubbles**

The term, "polymer microbubble" typically refers to a special class of microbubbles that are stabilized by a thick shell comprising cross-linked or entangled polymeric species. Polymer shells are more resistant to expansion and compression; therefore during drug delivery, it reduces echogenicity. During insonification polymer microbubbles release gas core which was unstable and rapidly dissolved [8, 20]. In 1990 a new polymer-shelled microbubble was reported by Wheatley et al. [21] in which the shell was formed by the ionotropic gelation of alginate. By using concentric jells of air and alginate solution, the microbubbles were prepared that was sprayed into a reservoir. On plunging into the calcium solution, the alginate was absorbed by the gas/liquid interface and was hardened. To increase the microbubble yield, sonicate the solution prior to spraying. By using the flow rate of air around the syringe needle, microbubble size was primarily determined. The diameters of microbubbles ranged between 30 and 40 μm and were therefore too large for intravenous administration. In 1997, Bjerknes et al. [22] introduced a method for making microbubbles using an emulsification—solvent evaporation method—encapsulated by a proprietary double-ester polymer with ethylidene units. The polymer microbubbles had a diameter ranging from 1 to 20 μm diameter. Optical microscopy and cryogenic transmission electron microscopy (cryo-TEM) were used for the determination of elongated, crumpled shapes of the microbubbles. The polymer shell was typically 150–200 nm thick. Acoustic tests determine a dose-dependent increase in acoustic attenuation. In 1999, Nayaran and Wheatley describe the preparation of microbubbles by using the biodegradable copolymer poly(D,L-lactide-co-glycolide) (PLGA). By using a volatile solid core, the microspheres were made hollow which could be sublimed. Manipulation of the solution viscosity, polydispersity, and shearing rate microbubble size was controlled. The size distribution ranged from 2 to 20 μm diameter. After incubation in serum the zeta potential of the microbubbles became less negative. In 2005, Cui et al. [23] reported the fabrication of PLGA microbubbles by using a double emulsion, solvent evaporation method. Coulter counter determines the size ranges between 1 and 2 μm diameter. Scanning electron microscopy (SEM) is used to study surface of particles so that the smooth surfaces, visible pores, or cavities can be explained. Confocal scanning microscopy explains internal morphology so that a single hollow core to a more honeycomb structure could be explained depending on the emulsification conditions. In 2005, Cavalieri et al. [24] determined a method of coating microbubbles by using PVA. In this case chemical cross-linking of PVA with microbubbles occurs at the air/water interface with a speed of 8000 rpm, so the mean diameter was approximately 6 ± 1 μm. By decreasing the operating temperature from room conditions to 4°C, the shell thickness could be decreased from 0.9 to 0.7 μm. PVA microbubbles enhance the shelf life of microbubbles by several months. This also increases the incorporation capability of hydrophobic drug and targeting ligand in microbubbles. Bohmer et al. [25] in 2006 used inkjet printing and developed a new technique for the preparation of polymer microbubbles. In this method they injected copolymer polyperfluorooctyl oxycaronyl-poly (lactic acid) (PLA-PFO) having a diameter of 4–5 μm as an organic phase into the aqueous solution (**Figure 2C**).
