Coronal Complexation Induces Snowman‐Shaped Janus Polymersome Formation

Abstract

Engineering polymersome shape is a keen area of interest because minimal variations in polymersome properties can greatly affect the performance of nanoreactors, drug delivery vehicles, and more. Although methods to transform polymersome shapes have been evolving rapidly, limitations remain in the scope of applicable polymersome compositions and adaptability to engineer a desired shape. In this work, we developed a method utilizing cationic polymer additives and coronal complexation to transform charged spherical polymersomes into asymmetric Janus polymersomes. The resulting morphologies feature a hollow polymersome compartment conjoined to a protruding bleb consisting of the complexed polymer additive, which we propose forms via a dewetting process. This methodology provides enhanced shape control through adjustments in additive composition or plasticizer content, offering additional routes to engineering novel polymersome structures.

Snowman‐like Janus polymersomes were produced in a simple, yet versatile two‐step approach involving a plasticizing agent and a cationic polymer additive to complex with the charged polymersome. Shape transformation was driven by coronal complexation altering membrane structure, followed by segregation of the polymer additive to form a protruding bleb of the Janus polymersome.

Introduction

To perform vital functions (e.g., endo/exocytosis, migration, or division), cells require the ability to alter their shape.^{[ 1 ]} This is often triggered by environmental fluctuations to create imbalances across the membrane either by osmotic pressure or changes in membrane composition (i.e., lipid or protein adsorption/insertion) to generate curvature.^{[ 1} , ² , ^{3 ]} For mammalian cells to be able to undergo shape changes or to allow lateral diffusion of lipids or protein, high membrane fluidity is a prerequisite. This can be achieved by changing the temperature or altering the membrane composition.^{[ 4 ]} Material scientists have long tried to replicate the ability of nature to reorganize the lipid membrane structure to achieve organized surfaces or shape transformations. Vesicles or polymersomes often serve as a platform as their bilayer structures are reminiscent of biological systems.^{[ 5} , ⁶ , ^{7 ]} Traditionally, polymersome shape transformation has been achieved via osmotic pressure; however, the transformations were reliant on the glass transition temperature (T _g) and final shapes varied with membrane and solvent compositions.^{[ 8} , ⁹ , ¹⁰ , ^{11 ]}

Recently, a so‐called “additive approach” toward polymersome shape transformation has emerged due to its simplicity and applicability to a wide range of polymersome chemistries. The approach has notably opened up access to novel polymersome shapes ^{[ 5} , ^{6 ]} (e.g., multi‐armed discs, starfish, hairy polymersomes) that were previously unattainable using conventional methods, involving osmotic pressure and liquid crystallinity, bringing us closer to replicating the true complexity of non‐spherical cell/organelle shapes such as the mitochondria.

A pioneering example of the additive approach was developed by Sun et al.^{[ 12} , ^{13 ]} (Figure 1a). The authors relied on a lower critical solution temperature (LCST) polymer additive that inserts itself into the outer leaflet of a polymersome membrane. Upon plasticization, the LCST polymer undergoes a hydrophilic‐to‐hydrophobic transition and subsequent conformational change. This effectively changes the surface area of the outer leaflet and induces bilayer asymmetry, thus initiating the onset of shape transformation into structures with protruding “arms” comprising a segregated form of the polymer additive.

Figure 1.

Schematics of shape transformation techniques utilizing bilayer asymmetry via a) LCST‐active membrane insertion^{[ 13 ]} b) Nucleobase pairing membrane insertion,^{[ 14 ]} and this work c) coronal complexation of polymeric additive.

Another method (Figure 1b) developed by Thomas et al.^{[ 14 ]} is conceptually similar, utilizing a polymer additive to achieve asymmetric membrane insertion. The difference, however, lies in the type of polymer used. Rather than using an LCST polymer, Thomas et al.^{[ 14 ]} used a nucleotide‐containing polymer additive that can complex with the membrane structure via nucleobase pairing. The accumulation of the additive within the outer membrane leaflet once again induces bilayer asymmetry, resulting in the formation and elongation of “nodes” on the polymersome surface. Although the two methods discussed involve the insertion of polymeric additives into the membrane, the mechanics for transformation differ greatly by virtue of random insertion and segregation behaviors to produce unusual structures. However, due to specific requirements for LCST and base‐pairing, the scope of applicable polymers for shape transformation is limited. Furthermore, these studies utilize neutral polymer coronas, leaving charged species unexplored despite the advantages of leveraging electrostatic interactions to easily manipulate membrane composition with endless possible charged polymer combinations to explore.

In this work, we introduce a new additive approach that provides additional levels of control, allowing coronal manipulation at the molecular level by combining polyelectrolyte interactions with membrane plasticity. Our approach to polymersome transformation consists of three steps: i) plasticization of negatively charged polyacrylic acid‐block‐polystyrene (PAA‐b‐PS) polymersomes, ii) coronal complexation of a cationic additive polymer via electrostatic interactions, and iii) segregation of the complex to form snowman‐shaped Janus polymersomes featuring a hollow polymersome compartment conjoined to a cationic polymer complex. This method offers versatility to simultaneously manipulate factors (e.g., degree of plasticization and properties of the cationic additive) to steer transformation into a variety of shapes, as we will demonstrate.

Results and Discussion

We used PAA₃₈‐b‐PS₈₄ (Figure 2a) as building blocks for the polymersomes (see Figure S1 and the Supporting Information for synthetic procedures and characterization data). We previously showed that this anionic block copolymer can form into polymersomes in a relatively narrow phase window.^{[ 15 ]} Given that the block length of the block copolymer used in this work differs from our earlier work, we had to first screen through a series of solvents to identify suitable self‐assembly conditions to target the polymersome morphology. Briefly, the self‐assembly process was conducted by mixing equivolumes of a 1 mg mL⁻¹ THF solution of PAA₃₈‐b‐PS₈₄ with 100–500 µM aqueous hydrochloric acid (HCl) containing 1 mM of aqueous sodium chloride (NaCl). The purpose of HCl addition is to modulate the charged state of the PAA block, which has a pKa of ∼4.5,^{[ 16 ]} while NaCl addition serves to screen some PAA charges to ensure dynamic equilibrium between charged groups.

Figure 2.

a) Chemical structure of PAA₃₈‐b‐PS₈₄. b) DLS intensity‐weighted distributions of polymersomes assembled at varying HCl concentrations (100–500 µM) in the presence of 1 mM NaCl as the aqueous phase. Respective TEM micrographs were prepared using c) 100 µM, d) 300 µM, and e) 500 µM HCl in the presence of 1 mM NaCl as the aqueous phase.

We show in Figure 2b dynamic light scattering (DLS) data obtained from the resulting particles in their organic/water mixtures (further provided inset Figure 2b are hydrodynamic diameters (D _h,intensity) and polydispersity values (PDI) for each sample). As can be seen from Figure 2b, the self‐assembly of PAA₃₈‐b‐PS₈₄ is sensitive to changes in HCl concentration due to the varying degrees of PAA chain protonation. Larger particles appear to form with increasing HCl concentration, though it is worth noting that the lowest HCl concentration tested (100 µM) also shows a small shoulder peak centered around ∼20 nm, likely indicative of micelle formation. To probe particle morphology, we analyzed the samples using transmission electron microscopy (TEM) (Figure 2c–e). Indeed, as we hypothesized, the sample prepared at 100 µM HCl is composed of a mixture of micelles and polymersomes, while both the samples prepared at 300 and 500 µM HCl contained pure phases of polymersomes. We decided to proceed with subsequent experiments using polymersomes formed with 300 µM HCl instead of 500 µM HCl due to a slight improvement in latter's polydispersity index (PDI) (refer back to Figure 2b(ii,iii)). As a purification step, we removed the organic solvent (THF) from the polymersome sample by extensive dialysis against the same aqueous solution.

To induce shape transformation, we used the block copolymer, polyethylene oxide‐block‐poly(4‐vinylpyridine) (PEO₄₄‐b‐P4VP₃₈), whose chemical structure is shown in Figure 3a. Briefly, PEG is a neutral hydrophilic polymer, while P4VP is a cationic polymer with pyridyl groups that becomes protonated below its pK _a of ∼5.2.^{[ 17 ]} Under acidic conditions, the pyridyl groups become protonated (P4VP⁺) to render PEO₄₄‐b‐P4VP⁺ ₃₈ doubly hydrophilic, thus enabling it to form polyelectrolyte complexes with the oppositely charged PAA chains of PAA₃₈‐b‐PS₈₄ polymersomes. In the case of our experiments, we chose to acidify PEO₄₄‐b‐P4VP⁺ ₃₈ with aqueous HCl to pH 3.0 to ensure the polymer was well‐dissolved.

Figure 3.

a) Schematic of the complexation‐driven shape transformation method using PEO₄₄‐b‐P4VP_n. Plasticization allows the rigid PS polymersome membrane to become flexible, which can then deform upon complexation. b) TEM micrographs of Janus polymersomes by variation of 4VP molar ratio (i) 0.2 eq, (ii) 0.5 eq, (iii) 1.0 eq, and (iv) 2.0 eq. c) TEM micrographs of polymersomes transformed using varying P4VP block length polymers, where n denotes the number of repeating units (i) 10, (ii) 23, (iii) 38, and (iv) 73 units.

The procedure for the shape transformation process is illustrated in Figure 3a. We typically begin by plasticizing the membrane structure of PAA₃₈‐b‐PS₈₄ polymersomes by doping 1 mL of 0.2 mg mL⁻¹ aqueous polymersome solution with 20% (v/v) of THF. A slow THF addition rate of 1 drop of THF per 30 s under stirring was implemented to prevent abrupt changes in solvent quality that could lead to polymersome dissolution. After incubation for 1 h to ensure that THF is homogenized across the entire polymersome sample, we added different amounts of 10 mg mL⁻¹ of acidified PEO₄₄‐b‐P4VP⁺ ₃₈ to achieve complexation at different P4VP:PAA mol ratios ranging from 0.2 to 2.0 eq. For context, this was calculated to be the addition of between 0.021–0.214 mg of PEO₄₄‐b‐P4VP⁺ ₃₈ per 1 mL of 0.2 mg mL⁻¹ of PAA₃₈‐b‐PS₈₄ polymersomes. This was followed by mild evaporation under stirring, which helps to remove the plasticizer (THF) from the sample, allowing PS to regain its glassy state and prevent further structural changes. Following THF evaporation, the resulting samples were subjected to purification by repeated centrifugation/redispersion cycles to remove any excess (uncomplexed) PEO₄₄‐b‐P4VP⁺ ₃₈.

We then used TEM to probe how complexation at different P4VP:PAA mol ratios influenced particle morphology (Figure 3b). Beginning from the lowest equivalency of 0.2 mol eq. of P4VP used, we observed significant clustering of the polymersomes into ill‐defined structures (Figure 3b(i)). Interestingly, at P4VP mol ratios above 0.5 eq., we find in all samples a population of Janus polymersomes—hollow structures tethered conjointly to a single protruding bleb with relatively dark electron contrast (Figure 3b(ii–iv)). Given how most Janus polymersomes reported to date rely on complex chemical processes to produce (e.g., metal deposition, crosslinking, seeded polymerization),^{[ 18} , ¹⁹ , ²⁰ , ^{21 ]} we were motivated to study these structures in more detail, as will be discussed below.

In addition to Janus polymersomes, we also observe a population of particles that do not have the typical vesicular (hollow) feature of polymersomes but share similarities to the dark protruding blebs of the Janus polymersomes. These particles were noticeably smaller in size (<100 nm) compared to the Janus polymersomes (>150 nm), which suggests that the size of the parent PAA₃₈‐b‐PS₈₄ polymersomes may have some influence on the shape transformation.

Under neutral conditions, the PAA corona is sufficiently deprotonated to give a negative surface charge, which leads to electrostatic repulsion between polymersomes. However, the presence of only 0.2 eq. of PEO₄₄‐b‐P4VP⁺ ₃₈ appears to cause significant agglomeration between the polymersome structure (Figure 3b(i)). We presume that here the additive complexes at the interface of multiple PAA coronae to give an effect akin to crosslinking (see schematic in Figure S2a for clarity). Interestingly, a further increase to 0.5 eq. of PEO₄₄‐b‐P4VP⁺ ₃₈ eliminates agglomeration (Figure 3b(ii)). We note that while one may assume that 0.5 eq. of cationic additive is only half the required amount to complex all available PAA in the sample, it is important to recognize that most of the PAA chains on the inner leaflet of the polymersome membrane are inaccessible to the cationic additive due to its inability to transverse the PS membrane. Based on the relative ratio of surface area between the external and internal polymersome leaflets (Figure S3), we estimate that only ∼65% of PAA chains are available for complexation. Thus, the use of 0.5 eq. of cationic additive should nearly saturate the surface of the polymersomes. This is asserted by the fact that no further morphological transformations were observed when the amount of cationic additive added was increased to 1.0 and 2.0 eq. (Figure 3b(iii,iv)). We postulate that at higher eq. of cationic additive added, the oppositely charged P4VP and PAA chains complex to form a new interdigitated corona with PEO chains present on the outermost surface, as depicted in Figure S2b.

Next, we explored how the properties of the cationic additive influence shape transformation. More specifically, we varied the block length of P4VP in the cationic additive to alter: i) the overall hydrophobicity of the polymer and its resulting complex, ii) the physical mass loaded onto the outer leaflet of the polymersome membrane, and iii) the conformation of the cationic additive upon complexation. Three additional PEO₄₄‐b‐P4VP _n polymers with n = 10, 23, and 73 units were synthesized (see Figure S4 and the Supporting Information for synthetic procedures and characterization data). All of the polymers were acidified to pH 3.0 to ensure that the P4VP was protonated and completely solubilized. This helps to eliminate the complexity of the experiment to only a single experimental variable (negating pH effects), allowing us to solely explore the effect that P4VP block length has on shape transformation. We performed shape transformation using acidified PEO₄₄‐b‐P4VP⁺ _10/23/73 following the procedure already described in Figure 3a. In these experiments, we chose to use an excess of cationic additive (>1.0 eq.) to ensure complete saturation/complexation of the polymersome surface. After removal of THF, the resulting structures were purified by centrifugation to remove excess cationic additive and subsequently probed with TEM. As shown in Figure 3c, the different block lengths of cationic additive resulted in a range of morphologically distinct structures.

The shortest block length of n = 10 units gave structures that strongly resemble the original (parent) polymersome structures (Figure 2d). Upon increasing the block length to n = 23 units, all polymersomes appear to have transformed into structures featuring a single and pronounced indentation resembling stomatocytes—a classic non‐spherical form of polymersomes that have been widely reported in the literature.^{[ 10} , ^{11 ]} A further increase to n = 38 units leads to Janus polymersomes featuring a single protruding bleb, as already discussed around Figure 3b. At n = 73, Janus polymersomes are also produced, although there is a greater prevalence of ill‐defined and solid particles compared with the n = 38 additive (Figure 3c(iv)).

Unlike the cationic additive with block length n = 10, which gave no morphological changes, the n = 23, 38, and 73 additives all led to (shape‐)asymmetric polymersomes. We therefore presume that the complex formed between PAA and P4VP chains must possess some level of hydrophobicity to allow the resulting PAA/P4VP complex to form a segregated domain on the parent polymersome structure. However, the n = 23 additive forms a comparatively lesser hydrophobic complex, which is insufficient for the formation of a protrusion, such as in the case of the Janus polymersomes. The n = 23 additive still permits complex segregation on the molecular level, as evidenced by the formation of single indentations on the polymersomes (Figure 3d(ii)). Although not visually obvious, the location of these indentations is presumably where most of the hydrophobic complex lies, as the polymersome can only deform in this region only if there exists some degree of structural asymmetry in the bilayer structure.^{[ 13} , ²² , ^{23 ]} Finally, with the n = 38 and 73 additives, the complex is sufficiently hydrophobic enough to form a phase‐separated domain in the form of a protrusion. Given its dark electron contrast, the domain must consist of electron‐dense components such as PS or P4VP. We will discuss the origins of the Janus structure in more detail in a later section.

Next, we continued our efforts to optimize the formation of the Janus polymersomes. We were curious to understand how THF, which was used as a plasticizer to lower the energetic barrier to membrane deformation (refer back to Figure 3a), affects the shape transformation process. Here, we chose to explore the effect of THF content from 0 to 30% (v/v) while maintaining complexation using 1.0 eq. of PEO₄₄‐b‐P4VP⁺ ₃₈. Size changes before and after transformation were monitored by DLS (Figure S6). The resulting particles were purified as already discussed and then imaged by TEM (Figure 4).

Figure 4.

a)–d) TEM micrographs of polymersomes complexed with PEO₄₄‐b‐P4VP⁺ ₃₈ (1.0 eq) at THF contents from 0 to 30% (v/v). e) Column graphs summarizing the population (n > 200 per sample) of the different structures present at the specified THF contents. Figure legend (dark green = spherical, pale green = indented, grey = ill‐defined, pale purple = Janus, and dark purple = solid) contains representative images of the structures classified under the specified categories.

From Figure 4a, it is apparent that no shape transformation is observed at 0% (v/v) THF content, and no noticeable size changes were observed (D _h ∼157 nm). This is expected since there is insufficient THF for the PS membrane to transition out of its glassy state. Note that the minor indentations seen in Figure 4a should not be confused with shape transformation, as they are a common imaging artifact caused by buckling of the polymersome membrane during TEM sample preparation/drying but are not apparent in cryoTEM (see Figure S5). An increase in THF content to around 10% (v/v) promotes membrane flexibility and facilitates the onset of shape transformation, yielding indented polymersomes (Figure 4b). A slight increase to 20% (v/v) THF permits membrane components to reorganize laterally to give rise to Janus polymersomes (Figure 4c), with an increase in particle size to D _h ∼164 nm. Further increases beyond 30% (v/v) THF saw significant size increase (D _h ∼171 nm) and broadening distribution (PDI ∼0.155) evident of a complete transformation and loss in polymersome morphology. This occurs as the internal leaflet is now accessible for complexation, resulting in dissociation of the membrane structure.

To ensure that the observed transformations were indeed attributed to complexation, we also prepared a set of negative controls at the same THF contents, but without the addition of the cationic additive (Figure S7).

Visual inspections aside, we also quantified the population of each morphology seen in Figure 4e to gain insights into the shape transformation progression. At each %THF content tested, particle populations (n > 200 particles) were assessed by image analysis and categorized according to their shape and morphology. For simplicity, the particles were divided into four categories (Figure 4e): “Spherical” refers to untransformed polymersomes that are (quasi‐)spherical in shape (Figure 4e). “Indented” comprises of polymersomes with a “mouth” present (Figure 4e). “Janus” refers to polymersomes with a single protruding cap (Figure 4e). “Solid” consists of particles with no apparent membrane structure but a consistently dark contrast across a large portion of their structure (Figure 4e). Lastly, “Ill‐defined” refers to polymersomes that appear to be transformed but cannot be classified under the aforementioned categories.

According to Figure 4e, 20% (v/v) THF is the optimal condition for Janus polymersome formation with an apparent yield of only ∼43%. We therefore explored the possibility of isolating the Janus polymersomes by centrifugation. We anticipated that this was possible because the other ∼57% of (unwanted) ill‐defined polymersomes appear to have smaller sizes and hence overall lower density. Indeed, centrifugation at 7000 rpm for 5 min leads to the Janus polymersomes being pelleted, while the indented polymersomes remain in the supernatant. Careful removal of the supernatant and subsequent resuspension of the pellet eventually yielded a pure phase of Janus polymersomes (Figure 5a).

Figure 5.

a) TEM micrograph of purified Janus polymersomes prepared with PEO‐b‐P4VP⁺ ₃₈. b) Cryo‐TEM micrograph of Janus polymersomes in multiple orientations as indicated by the arrows on the adjacent 3D representations, where purple = polymersome region, and yellow = protruding bleb.

Although various types of Janus polymersomes have been reported in the literature, there is, to the best of our knowledge, no report on polymersomes with a protruding bleb such as in this work. We were therefore motivated to study their structure in more detail. In pursuit of this, we first examined the n = 38 particles under cryogenic‐transmission electron microscopy (cryo‐TEM) to visualize them in a (native) frozen‐hydrated state. In the resulting cryo‐TEM image shown in Figure 5b, we find three Janus polymersomes coincidently oriented in three different ways (side‐on, top–down, and off‐axis), making structural investigations straightforward (see also Figure S8 for n = 73 Janus polymersomes). To aid visualization, we superimpose 3D representations of the three particles in the same cryo‐TEM image, with purple being used to indicate the polymersome component and yellow to indicate the protruding bleb. Arrows in blue are also shown to indicate the orientation of the Janus polymersomes relative to the cryo‐TEM grid (background).

What was most peculiar from the image was the resolvement of the internal structure of the protruding bleb. The porosity of the bleb is clearly visible without any signs of short‐ or long‐range ordering. Also interesting was the lack of any clear boundary between the protruding bleb and the polymersome segment. This indicated that the PAA/P4VP complex does not simply segregate on the polymersome surface but rather reorganizes a portion of the membrane structure, notably at the interface where the bleb and the polymersome join.

To further analyze the structure, we performed cryogenic electron tomography (cryo‐ET), collecting a series of cryo‐TEM images at multiple tilt angles (60 images at 2° increments) and reconstructing a 3D projection using the collected images. Figure 6 shows the resulting 3D projection and a series of left‐to‐right images sliced along the z‐axis of the 3D projection. The images were assembled to create a movie that can be found in the associated content.

Figure 6.

a) 3D projection of a Janus polymersome obtained from cryo‐ET. b) Graphical representation depicting sliced regions, where purple = polymersome region and yellow = protruding bleb. c) Cryo‐ET micrographs of the sliced images along the z‐axis of the 3D projection. Scale bar = 50 nm. d) (i) Magnified image of the protruding bleb, (ii) cartoon schematic of the composition, and (iii) overlay image with the cryo‐ET micrograph. e) Schematic of the interactions of the polymers at the molecular level, where navy = PS, yellow = PAA/P4VP complex, and blue = PEO.

In our system, we know that only two polymer components (PS and P4VP) can contribute to the dark electron contrast seen in Figure 6. Intuitively, this informs us that the structure seen in Figure 6 must be composed of either PS or P4VP. The vesicular portion of the Janus polymersome reflects the original features of the parent polymersome (refer back to Figure 2), while the protruding bleb likely comprises a newly formed polymer domain arising from the complexation between the cationic additive and the anionic polymersome corona. To assist in visualizing the composition of the bleb, a magnified image and cartoon overlays are shown in Figure 6d(i–iii).

Upon complexation, both P4VP and PAA lose their ability to disperse in water, as the complexed domain of P4VP/PAA is hydrophobic in nature. Since the domain is tethered to the parent polymersome structure, frustration arises, leading to their segregation into a compartment as seen in all our Janus polymersome structures. This segregation behavior of P4VP/PAA is reminiscent of domain formation by interpolyelectrolyte complexes,^{[ 24} , ^{25 ]} with the presence of salts and plasticizers to encourage further structural rearrangements. Simultaneously, with the insertion of PEO₄₄‐b‐P4VP⁺ ₃₈ into the polymersome shell, the PAA layer becomes denser, and the polymer chains start to stretch in response to it. The membrane thickness increases from 18.6 to 23.2 ± 1.8 nm  (n = 50) at 0.5 eq PEO₄₄‐b‐P4VP⁺ ₃₈ to 27.6 ± 2.3 nm (n = 56) at 1.0 eq cationic polymer. The denser brushes might have prevented further complexation, which could explain why there are no significant morphology changes at 1.0 eq PEO₄₄‐b‐P4VP⁺ ₃₈. The apparent light contrast “holes” can only be attributed to PEO, given its poor electron contrast nature. Recall that PEO is covalently bound to the cationic additive (Figure 6e) and must therefore also be present somewhere within the newly formed domain. Since PEO is hydrophilic and generally incompatible with hydrophobic polymers (hence its use as a hydrophilic component in many amphiphilic block copolymer systems), it is perhaps unsurprising that PEO would constitute holes within the bleb.

We propose that the protruding bleb forms via a dewetting mechanism.^{[ 26 ]} To elaborate, we can simplify the system by considering the Janus polymersomes as an immiscible triphasic system: (Phase A) the parent polymersome component, (Phase B) the protruding bleb (or complexed polymer domain), and (Phase C) the non‐solvent (water) wherein the polymersomes are dispersed in. As illustrated in Figure 7b, there are three possible configurations for the interaction of the three immiscible phases, namely, complete engulfing, partial engulfing, and non‐engulfing.^{[ 27} , ^{28 ]} These are governed by the spreading coefficients (S_i = γ_jk – (γ_ij + γ_ik ) for the ith phase, where γ = interfacial tension) dictate whether a fluid would spread (S_i  > 0) or retract (i.e., “dewet”) (S_i  < 0) from an interface (Figure 7a).^{[ 29 ]}

Figure 7.

a) Wetting of a liquid (blue) on a surface given its spreading coefficient (S). b) (i) Complete, (ii) partial, and (iii) non‐engulfing configurations of an immiscible triphasic system where Phases A = polymersome (purple), B = cationic polymer complex (yellow), and C = solvent (grey).

We consider the fact that when complexation occurs between the cationic additive and PAA corona, charge neutralization must render the complexed polymer (Phase B) hydrophobic. Intuitively, this should promote phase separation since an excess of the interface between Phase B and C (i.e., the BC interface) is highly unfavorable due to the hydrophobic effect. Interestingly, however, despite the complex being hydrophobic in nature, coronal complexation can proceed at low THF content (0% (v/v), Figure 4a) without any changes to the original polymersome morphology—this is consistent with the complete engulfing scenario in Figure 7b(i). This can be further explained because Phase A (i.e., the polymersome) does not have any membrane flexibility to allow morphing or any further minimizations in interfacial energies.

However, at a high THF content (c.a., 20% (v/v), Figure 4c), the membrane is now afforded with flexibility, which enables Phases A and B to morph into their equilibrium configurations. Phase B is now able to dewet (S _B <0) from Phase A to reduce the energetic penalty associated with an excessively large BC interface, forming a protruding bleb. Since the association of Phases A (predominantly PAA) and B (predominantly P4VP) is considered a thermodynamically favorable interaction due to electrostatics, Phase B dewets to a partial engulfing configuration (Figure 7b(ii)) rather than a non‐engulfing configuration (i.e., non‐interacting) (Figure 7b(iii)). Furthermore, this partial engulfing configuration (Figure 7b(ii)) correctly describes the shape of the Janus polymersomes (Figure 6) produced.

Lastly, to address the tendency to form a single bleb rather than multiple blebs, we constructed model polymersomes containing a single or two blebs of the same total volume (Figure S9). Given that the change in interfacial energy dE = γ*dA (where A = interfacial area)^{[ 30 ]} and the relative magnitudes of interfacial tension are γ _AC < γ _AB ≈ γ _BC; we determined that the two bleb configuration results in a net energetic penalty due to a ∼21.6% increase in higher energy Phase B‐containing interfacial areas when compared with the single bleb configuration. Hence, the single bleb configuration is preferred.

Conclusion

In summary, this work reports on a new approach to shape transformation utilizing charged polymersomes as a platform to synthesize Janus polymersomes. Our approach offers control to vary properties of the additive or membrane plasticity to affect the final polymersome shape with a relatively simple process, involving only three key steps: i) plasticization of the negatively charged PAA‐b‐PS polymersomes, ii) coronal complexation of a cationic PEO‐b‐P4VP additive, followed by iii) segregation or dewetting. For Janus polymersome formation, we found that it was crucial for the membrane to be sufficiently plasticized (20% (v/v) THF) for structural rearrangement to occur and that ≥ 0.5 eq of additive was present to ensure complete saturation of the corona (and thus prevention of agglomeration). The result is a singular protruding bleb consisting of segregated complexed additives, which forms via dewetting into a partial engulfing configuration representative of the Janus polymersome structure. This complexation‐driven approach offers new opportunities to leverage charged polymers for polymersome shape transformations.

Supporting Information

Materials, characterization, synthesis, self‐assembly procedure, and supplementary figures can be found in the supporting information. A movie, obtained from the stacked cryo‐TEM images, shows cross‐sections of the 3D structure.

Conflict of Interests

The authors declare no conflict of interest.

Supporting information

Supporting Information

Supporting Information

Lai R. Y., Wong C. K., Stenzel M. H., Angew. Chem. Int. Ed.. 2025, 64, e202507756. 10.1002/anie.202507756

Data Availability Statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.