* NPs can be self-assembled at the surface of emulsion droplets prior to the formation of polymer shell via classical emulsion-based processes (especially interfacial polymerization). This strategy has been used to produce composite polymer capsules that incorporate TiO₂, gold, or silica nanoparticles.²⁸¹⁻²⁸³

As an alternative to these conventional methods, we showed that our strategy of thermo-enhanced interfacial complexation of PLL-g-PNIPAM offers a straightforward way to incorporate NPs in the polymer shell. We demonstrated that, upon adsorption of PLL derivatives on the surface of NPs, the latter can participate to the thermo-induced accumulation of matter at the interface, resulting in the formation of polymer-NPs mixed shells simply by gentle heating. We evidenced this process using fluorescent NPs whose distribution in the droplets can be directly visualized by microscopy, in particular upon thermal response of the polymer.

Then, gold NPs were used as a promising additive to trigger the collapse transition of PLL-g-PNIPAM by thermoplasmonic excitation, extending the stimuli-responsiveness of our system towards light stimulation. For this photothermal application, being able to measure temperature at microscales with high spatial resolution is a major issue and will be discussed before precisely focusing on a particular microscopy method enabling to retrieve the temperature rise generated *in situ* by thermoplasmonic excitation. The power of this thermometry method will be demonstrated on experimental examples of temperature measurements in emulsions containing both polymer and gold NPs.
## 4.2 Interactions between PLL derivatives and nanoparticles

### 4.2.1 Fluorescent polystyrene nanoparticles

We used two types of commercial fluorescent polystyrene NPs with different surface modifications: NeutrAvidin-modified polystyrene NPs (NeutravNPs) and carboxylate-modified polystyrene NPs (CarboxyNPs). At physiological pH, the surface of NeutravNPs is expected to be almost neutral (the pI of NeutrAvidin protein being of 6.3) and the surface of CarboxyNPs is expected be highly anionic (the pKa of carboxylic acids being around 4).

Zeta potential measurements were performed to assess the modification of the surface charge of the NPs after incubation with PLL chains at pH = 7.3. For these experiments, NPs with a diameter of 200 nm were used in order to facilitate the removal of excess polymer chains by filtration after the incubation step (see **Appendix A** for details). The zeta potential distributions of the NPs before (commercial solutions) and after incubation with PLL (PLL-g-PNIPAM-coated NPs) are shown in **Figure 4.1**, both for CarboxyNPs and NeutravNPs.

<figure><img src="image_76.png" /><figcaption>**Figure 4.1.** Zeta potential distributions of CarboxyNPs (left) and NeutravNPs (right) from commercial solutions (dashed lines) and after incubation with PLL solutions (solid lines). The distributions are the average of three measurements by sample.</figcaption></figure>

Both commercial NeutravNPs and CarboxyNPs were highly anionic with a distribution of the zeta potential centered around –42 mV and –64 mV respectively. This value is not surprising for CarboxyNPs since
surface carboxylic acids are deprotonated at pH = 7.3. The anionic character of NeutravNPs was less pronounced since Neutravidin is only slightly negatively charged at pH = 7.3 (pI = 6.3). A fraction of the sample even exhibited near neutral surface charge as suggested by a small shoulder observed around -11 mV in the zeta potential distribution.

After incubation of the NPs with PLL and filtration of the polymer excess, a shift of the zeta distribution towards “less negative” values was observed for both types of NPs. This effect can be attributed to a significant adsorption of cationic PLL chains at the surface of the NPs, hence increasing their surface charge by partial compensation of the initial anionic charges. For CarboxyNPs, the distribution shifted to around -37 mV (with a small shoulder reaching neutral values). For NeutravNPs, two populations of same magnitude appeared, one still negatively charged around -38 mV and one near neutral around -5 mV. This partially efficient adsorption of PLL suggests that the Neutravidin surface coverage is non-uniform within the NPs sample. In any case, PLL apparently develops electrostatic interactions with our two types of negatively-charged surface-modified polystyrene NPs. This propensity of PLL to bind to anionic surface has already been reported in the literature for various types of materials including silica, ceramics, plastics such as polystyrene, protein particles, etc.<sup>285-288</sup>

The adsorption of PLL on NPs is not necessarily prejudicial *per se* (as long as it does not destabilize the NPs solution). It implies that *functional* PLL derivatives such as PLL-g-PNIPAM would also be able to coat anionic NPs. In that case, since PNIPAM chains become hydrophobic at high temperature, PLL-g-PNIPAM-coated NPs may undergo aggregation upon heating via inter-coating hydrophobic interactions, as already demonstrated in previous works.<sup>288</sup>

This process was assessed for “naked” and PLL-g-PNIPAM-coated NeutravNPs or CarboxyNPs by monitoring the evolution of their hydrodynamic diameter by dynamic light scattering (DLS) upon heating. To evidence any aggregation process occurring upon the LCST transition of PNIPAM, the mean hydrodynamic diameter measured at each temperature was normalized by the initial diameter value of ~ 200 nm measured at 20 °C (see **Figure 4.2**).
<figure><img src="image_77.png" /><figcaption>**Figure 4.2.** Evolution of the mean effective diameter measured by DLS in NPs solution upon heating. The effective diameter has been normalized by its initial value of 200 nm (at 20 °C) to evidence the progressive aggregation of PLL-g-PNIPAM-coated NPs compared to non-coated NPs.</figcaption></figure>

For commercial NPs solutions, the apparent diameter did not change between 20 °C and 50 °C, indicating that the colloidal suspensions were stable upon heating. For PLL-g-PNIPAM-coated NPs, the mean diameter measured in each sample increased continuously as temperature increased from 35 °C to 50 °C. As this trend started at a temperature which fairly corresponds to the cloud point of PLL-g-PNIPAM (34 °C, see Chapter 1), this process can be reasonably attributed to a progressive agglomeration of NPs driven by hydrophobic interactions between PLL-g-PNIPAM layers which decorate their surface. The amplitude of this phenomenon was markedly higher for CarboxyNPs than for NeutravNPs. Indeed, the effective diameter of the aggregates at 50 °C was 6 times the diameter of the individual NPs in the case of CarboxyNPs, corresponding to an aggregation number of ~ 160 NPs per aggregate (calculated as $0.74 \times (\frac{d_{agg}(T=50°C)}{d_{NP}(T=20°C)})^3$ in a model of compact stacking of hard spheres). Conversely, in the case of NeutravNPs, the effective diameter was only 2 times the initial one, corresponding to ~ 6 NPs per aggregate.

These results confirm that (i) PLL-g-PNIPAM can adsorb at the surface of anionic nanoparticles without affecting their stability below the LCST of PNIPAM, (ii) the hydrophilic-to-hydrophobic transition of PLL-g-PNIPAM coating induces the aggregation of the NPs above the LCST, and (iii) the adsorption of PLL-g-PNIPAM is more efficient on CarboxyNPs than on NeutravNPs since the aggregation process is more pronounced. This last point is consistent with a partial adsorption of PLL on NeutravNPs as previously suggested by the presence of two populations of NPs in the zeta potential (see Figure 4.1).
Actually, regarding the initial goal of incorporating NPs in polymer capsules formed by interfacial
complexation, the possibility to coat NPs with PLL-g-PNIPAM can be a major advantage. As shown in
Chapter 1, hydrophobic PNIPAM can be accumulated in pre-formed PLL-g-PNIPAM shells upon heating.
As PLL-g-PNIPAM-coated NPs tend to form hydrophobic aggregates, we could also expect their
accumulation in PLL-g-PNIPAM shells. This phenomenon will be investigated by confocal microscopy on
fluorescent NPs in section 4.3.

4.2.2 Gold nanoparticles (AuNPs)

We used two types of commercial gold NPs (AuNPs): 1) 40-nm bare AuNPs (their surface was actually
occupied by residual citrate groups that are highly labile complexing agents), and 2) 50-nm AuNPs coated
with poly(vinyl pyrrolidone) (PVP) that favors steric stabilization of the colloidal suspension.

The aggregation of AuNPs in presence of additives can be readily revealed by UV-Vis
spectrophotometry. Indeed, non-aggregated spherical AuNPs exhibit a sharp absorbance peak at 532 nm,
corresponding to the plasmon resonance. Upon aggregation, this peak in the green range progressively
disappears in favor of a red-shifted peak which is characteristic of plasmon resonance in elongated
AuNPs.²⁸⁴ UV-Vis spectra of AuNPs solutions in presence and absence of PLL were thus compared (see
**Figure 4.3**, and **Appendix A** for details). This simple qualitative experiment was performed for both bare
and PVP-coated AuNPs commercial solutions, supplemented with PLL at concentrations in the wt% range
(typically used in the experiments of capsule formation).

<figure><img src="image_78.png" /><figcaption>**Figure 4.3.** UV-Vis spectra of bare AuNPs (left) and PVP-coated AuNPs (right) solutions containing various concentrations in PLL.</figcaption></figure>
For commercial solutions of bare and PVP-coated NPs, the plasmon peak at 530 nm was well defined (in dark red, corresponding to the real color of the commercial AuNPs solutions). When PLL was added in the solution of bare AuNPs, the intensity of the 530-nm peak drastically decreased (for any PLL concentration tested), and a broad bump appeared at higher wavelengths. This red-shift is typical of anisotropic aggregates of AuNPs and was also evidenced by a color switch of the solution from dark red to blue. This phenomenon was expected since bare AuNPs have a zeta potential of -52 mV (according to the datasheet), favoring the adsorption of cationic PLL at their surface, eventually inducing a massive aggregation by interparticle bridging.

For PVP-coated AuNPs, the addition of PLL did not drastically change the shape of the initial spectrum, except that the resonance peak was slightly shifted to 538 nm and lower in intensity. The preservation of the plasmon resonance characteristic of spherical AuNPs suggests that the presence of PLL did not induce any significant interparticle aggregation. This high colloidal stability can be attributed to the presence of the PVP coating which probably limits the adsorption of PLL at the surface of the AuNPs. To confirm this result, the zeta potential of PVP-coated AuNPs was measured before and after incubation with PLL (see Figure 4.4). The zeta potential shifted from -32 mV in the absence of PLL (in agreement with the commercial datasheet) to +6 mV after incubation with PLL, betraying a high coverage and charge reversion of the AuNPs. This result actually indicates that PLL can strongly bind to the surface of PVP-coated NPs, but obviously without altering their colloidal stability according to UV-Vis data.

<figure><img src="image_79.png" /><figcaption>**Figure 4.4.** Zeta potential distributions of PVP-coated AuNPs before and after incubation with PLL.</figcaption></figure>
To confirm UV-Vis and zeta results, the effective size of the AuNPs aggregates was measured by DLS for both bare and PVP-coated AuNPs solutions containing various amounts of PLL (see **Figure 4.5**).

<figure><img src="image_80.png" /><figcaption>**Figure 4.5.** Effective hydrodynamic diameters of objects measured by DLS in solution of AuNPs containing various amounts of PLL.</figcaption></figure>

For the initial solutions of AuNPs (in absence of PLL), the hydrodynamic diameters measured by DLS were consistent with the commercial datasheet: ~ 40 nm for bare AuNPs and ~ 70 nm for PVP-coated AuNPs (the Au core in itself has a diameter of 50 nm according to the datasheet TEM data, but the presence of the PVP coating increases the effective hydrodynamic diameter which is measured by DLS). The addition of PLL induced a slight increase of the effective diameter for PVP-coated AuNPs up to 100-150 nm, suggesting limited colloidal aggregation. For bare AuNPs solutions, the increase of PLL concentration induced a continuous increase of the effective mean diameter (up to ~ 650 nm for 5 wt% of PLL) and a concomitant broadening of the size distribution, betraying a massive clustering of the AuNPs in presence of PLL. These results are consistent with the UV-Vis measurements, and tend to validate the propensity of PLL to trigger a spontaneous and uncontrolled colloidal aggregation of bare NPs. On the contrary, PVP-coated AuNPs undergo with very limited aggregation, despite an effective coating of their surface with PLL. This behavior is analogous to the one of the fluorescent polystyrene NPs introduced in the previous section. For all these reasons, only AuNPs initially coated with PVP were used throughout this PhD.
## 4.3 Design of “composite” polymer shells by thermo-enhanced interfacial complexation

### 4.3.1 Proof of concept on NeutrAvidin-modified polystyrene NPs

For the proof of concept, the formation of polymer-NPs mixed shells was investigated in detail with fluorescent NeutravNPs using confocal microscopy. In these experiments, the PLL-g-PNIPAM used to form the capsules was not dye-labeled so that the fluorescence signal was only due to the NPs, enabling to directly visualize their distribution in the emulsion droplets.

**Figure 4.6.**A shows confocal micrographs of an emulsion containing 2 wt% of PLL-g-PNIPAM and 0.025 wt% of NeutravNPs in the aqueous phase at 18 °C and 45 °C, i.e. below and above the LCST of PNIPAM respectively. At 18 °C, the NeutravNPs were homogeneously distributed within the droplets. At 45 °C, the NeutravNPs were essentially located at the droplet edge, indicating that they had been captured at the water/oil interface concomitantly with the collapse transition of PNIPAM (occurring at 34 °C). This process of thermo-induced co-segregation of polymer chains and NeutravNPs at the interface is depicted in Figure 4.6.B (Krytox molecules and PLL positive charges have been removed for the sake of readability). It may be reasonably attributed to the combination of two results previously described: (i) the adsorption of PLL-g-PNIPAM chains at the surface of NeutravNPs (see section 4.2.1) and (ii) the thermo-induced accumulation of PNIPAM derivatives onto the PLL-g-PNIPAM shell at high temperature (see section 3.5.3 in Chapter 1). In other words, PLL-g-PNIPAM-coated NPs become hydrophobic and sticky at high temperature and agglomerate onto the polymer shell which is hydrophobic and sticky as well.
<figure><img src="image_81.png" /></figure>

<figure><img src="image_82.png" /></figure>

<figure><img src="image_83.png" /></figure>

<figure><img src="image_84.png" /><figcaption>**Figure 4.6.** Thermo-induced capture of NeutravNPs in PLL-g-PNIPAM shell upon collapse transition of PLL-g-PNIPAM. A) Confocal micrographs of an emulsion containing 2 wt% of PLL-g-PNIPAM (non-fluorescent) and 0.025 wt% of 50-nm fluorescent NeutravNPs in the aqueous phase and 0.05 wt% of Krytox in the oil phase, at 18 °C and 45 °C (scale bars = 50 µm). B) Schematic representation of the thermo-induced co-segregation of PLL-g-PNIPAM and NeutravNPs at the interface upon heating. For the sake of readability, positive charges of PLL and Krytox molecules are not represented. C) Radial kymograph and fluorescence profiles of a representative droplet upon heating from 27 °C to 45 °C (scale bars in the micrographs = 10 µm). D) Confocal micrographs and fluorescence radial profiles of a representative droplet during temperature switches between 18 °C and 45 °C.</figcaption></figure>

The evolution of the emulsion was recorded over time while temperature was increased (from 27 °C to 45 °C) to more precisely assess the dynamics of NeutravNPs within the droplets. For the representative droplet showed in Figure 4.6.C, the evolution of the radial profile and the radial kymograph are represented along the temperature sweep. In the radial kymograph, the projections of the radial fluorescence image at increasing temperatures have been put side by side in a continuous image. It illustrates the progressive disappearance of fluorescence in the droplet core concomitantly with an accumulation of the fluorescence at the droplet edge. This process is also highlighted by the evolution of the radial profile as temperature rises: the uniform profile of fluorescence at low temperature (below the cloud point) was progressively replaced by a sharp peak of fluorescence localized at the droplet edge (above the cloud point).
The consistency of the so-formed polymer-NPs mixed layer was assessed by two qualitative observations: (i) micron-size aggregates of NeutravNPs that had reached the interface were totally immobilized at 45 °C as shown in **Figure 4.7**, (ii) the corona of NeutravNPs formed at 45 °C did not redisperse as the emulsion was cooled below 34 °C (see Figure 4.6.D), suggesting that the co-segregation of NeutravNPs and PLL-g-PNIPAM at the interface was irreversible. Together, these data suggest that the polymer-NPs mixed layers formed upon heating behave like highly viscous or gel-like shells, probably due to PLL-NPs interactions that play the role of physical cross-links.

<figure><img src="image_85.png" /><figcaption>**Figure 4.7.** Temporal evolution at 45 °C of a representative water droplet containing 2 wt% of PLL-g-PNIPAM and 0.025 wt% of NeutravNPs, dispersed in oil phase containing 0.05 wt% of Krytox (scale bars = 10 µm). The white circle highlights the immobilization of a µm-size aggregate of NeutravNPs within the polymer shell.</figcaption></figure>

To confirm the role of hydrophobic PNIPAM strands in the thermo-induced capture of NeutravNPs at the interface, a control emulsion was made with non-thermoresponsive PLL-g-PEG shells (PLL-g-PNIPAM was replaced by PLL-g-PEG in the formulation). The NeutravNPs distribution was uniform in the droplets at 18 °C and 45 °C (see **Figure 4.8**), without any noticeable aggregation of the NeutravNPs at the interface upon heating. This result suggests that the interfacial capture of NeutravNPs observed with PLL-g-PNIPAM was due to the thermoresponsiveness of PNIPAM strands, and was not inherent neither to the nature of the NPs nor to the heating process in itself.

<figure><img src="image_86.png" /><figcaption>**Figure 4.8.** Control experiment with non-thermoresponsive PLL-g-PEG. A) Confocal micrographs of an emulsion containing 2 wt% of (non-fluorescent) PLL-g-PEG and 0.025 wt% of (fluorescent) NeutravNPs in the aqueous phase and 0.05 wt% of Krytox in the oil phase, at 18 °C and 45 °C (scale bars = 50 µm). B) Radial fluorescence profile of the droplet encircled in white in the micrographs, at 18 °C and 45 °C.</figcaption></figure>
### 4.3.2 Case of anionic carboxylate-modified polystyrene NPs

As a complement to the experiments on NeutravNPs, the thermally-induced capture of NPs in the polymer shell was also demonstrated with CarboxyNPs in identical conditions (see **Figure 4.9**).

<figure><img src="image_87.png" /><figcaption>**Figure 4.9.** Thermo-induced capture of Carboxy-NPs in PLL-g-PNIPAM shells. A) Confocal micrographs of an emulsion containing 2 wt% of PLL-g-PNIPAM (non-fluorescent) and 0.025 wt% of 50-nm CarboxyNPs fluorescent nanoparticles in the aqueous phase and 0.05 wt% of Krytox in the oil phase, at T = 18°C and T = 45°C (scale bars = 30 µm). B) Fluorescence radial profiles of the droplet indicated with a white arrow in the micrographs.</figcaption></figure>

This new approach of formation of polymer-NPs (or “composite”) shells has the advantage to rely on spontaneous deposition of PLL on NPs and spontaneous aggregation of the polymer-coated NPs upon gentle heating. Hence, it does not require any pre-treatment of the NPs of interest or post-modification of the polymer capsules. Here, the irreversible capture of NPs in PLL-g-PNIPAM shells has been demonstrated in the case of two types of anionic NPs, but this approach is likely to be generic as PLL can interact with a great variety of materials (silica, oxides, plastics, gold).

### 4.3.3 Incorporation of AuNPs in PLL-g-PNIPAM shells

Because AuNPs are not fluorescent, their precise localization in PLL-g-PNIPAM capsules formed by interfacial complexation could not be probed by fluorescence microscopy. However, as AuNPs are metallic, their incorporation in PLL-g-PNIPAM capsules may be evidenced on SEM images by contrast with the polymer matrix.

The capsules were formed and observed by SEM following the protocol described in Chapter 1 (see also **Appendix A**). Briefly, an aqueous solution was prepared by mixing 1 wt% of PLL-g-PNIPAM
with 0.05 wt% of 50-nm PVP-coated AuNPs in phosphate buffer, and then emulsified in FC-70 oil phase containing 0.05 wt% of Krytox to form aqueous-core polymer capsules by interfacial complexation. SEM imaging of dried capsules are shown in **Figure 4.10**.

<figure><img src="image_88.png" /><figcaption>**Figure 4.10.** SEM micrographs of dried emulsion containing 1 %wt of PLL-g-PNIPAM and 0.05% AuNPs in the aqueous phase, and 0.05 wt% of Krytox in the oil phase. The small white dots correspond to AuNPs aggregates embedded in the polymer shell (as indicated by the white arrow on the second micrograph).</figcaption></figure>

The presence of AuNPs in the formulation did not prevent the formation of PLL-g-PNIPAM capsules (showing the same structural features than the one presented in section 3.4.1 in Chapter 3 in absence of NPs). In addition, a multitude of nanometric white spots were clearly visible at the outer surface of the capsule shells, which was not the case in pure PLL-g-PNIPAM capsules. The typical size and high SEM contrast of these spots undoubtedly indicate that they are AuNPs aggregates.

SEM micrographs only indicate that AuNPs were incorporated in the polymer shells for capsules in a dried state but did not give any information on the AuNPs distribution in the initial state of the emulsion (before drying). However, since PLL can adsorb at the surface of AuNPs (see 4.2.2) as for NeutravNPs, we can reasonably assume that AuNPs would exhibit the same behavior than NeutravNPs in presence of PLL-g-PNIPAM in emulsion, i.e. a thermo-induced accumulation in the polymer shell upon heating.
## 4.4 Thermoplasmonic effects in AuNPs-loaded droplets

### 4.4.1 Photothermal transition of PLL-g-PNIPAM in emulsion: a qualitative approach

In section 4.3.3, we showed that AuNPs can be incorporated in the shell of PLL-g-PNIPAM capsules. As mentioned in Chapter 2 (section 2.3.4), AuNPs are interesting for their thermoplasmonic properties. The incorporation of AuNPs in PLL-g-PNIPAM capsules should in principle enable to trigger the thermal collapse transition of the polymer upon light irradiation in individual droplets, instead of heating the whole sample. First, qualitative experiments will be presented as a proof of concept to show that photothermal effects can be generated in AuNPs-loaded emulsion droplets and to identify the main parameters that may influence the efficiency of the process. Then a quantitative approach will be presented to measure *in situ* the temperature variations that can be generated by light excitation of AuNPs.

#### 4.4.1.1 Evidence for collapse transition of PLL-g-PNIPAM induced by light irradiation of AuNPs

The possibility to use thermoplasmonic effects of AuNPs to trigger the aggregation of PLL-g-PNIPAM was directly assessed in *situ*, i.e. in emulsion droplets. An aqueous solutions containing 10 wt% of PLL-g-PNIPAM and 0.05 wt% of AuNPs (PVP-coated) was prepared in phosphate buffer and emulsified in an oil phase containing 0.05 wt% of Krytox. The use of a high concentration in PLL-g-PNIPAM should saturate the droplet interface and facilitate the visualization of the collapse transition of the polymer upon heating, through the formation of a large amount of micrometric aggregates in the droplet cores.

A laser source emitting at 532 nm with a nominal power of 40 mW was used to excite the AuNPs. The laser beam was focalized on the sample through the side port of the microscope used to image the sample. At the level of the sample, the laser power has been reduced to 25 mW due to power losses within the microscope set-up. The diameter of the laser spot was typically 10 µm in the focal plane of the objective.

**Figure 4.11** shows an irradiation experiment on an individual droplet with a diameter of 30 µm. The droplet was exposed to different light powers (5, 10, 15 mW) for a few seconds (see Figure 4.11.A). At 5 mW, some granules appeared in the irradiated part of the droplet after 10 s of irradiation, suggesting that enough heat had been generated to locally reach 34 °C and trigger the collapse transition of PLL-g-
PNIPAM. This phenomenon was better evidenced at 15 mW since the droplet core got entirely filled with polymer granules after only 1 s of irradiation. At 25 mW, the collapse transition of PLL-g-PNIPAM even turned into a macro-phase separation of the polymer within the droplet core within 10 s of irradiation.

<figure><img src="image_89.png" /><figcaption>**Figure 4.11.** Collapse transition of PLL-g-PNIPAM triggered by AuNPs thermoplasmonic excitation. A) Phase contrast micrograph of an emulsion containing 10 wt% of PLL-g-PNIPAM and 0.05 wt% of AuNPs in the aqueous phase, and 0.05 wt% of Krytox in the oil phase. A 532-nm laser was focused on an individual 30-µm droplet to induce the collapse of PLL-g-PNIPAM in its core. The collapse transition of the polymer was clearly identified upon 1 s of irradiation with a light power P of 15 and 25 mW. B) Representation of the light-induced thermal collapse transition of PLL-g-PNIPAM occurring in the water droplet via thermoplasmonic effects of encapsulated AuNPs.</figcaption></figure>

These preliminary results suggest that the presence of AuNPs in the water droplets allows to generate enough heat upon light irradiation to induce the collapse transition of PLL-g-PNIPAM.
4.4.1.2 Parameters influencing the efficiency of light-induced PLL-g-PNIPAM collapse

The possibility to induce the collapse of PLL-g-PNIPAM with light is intrinsically related to the maximum
temperature rise that can be generated by AuNPs under irradiation. In a given droplet, the temperature rise
essentially depends on two parameters: the number of encapsulated AuNPs (itself depending on the AuNPs
concentration and the droplet size) and the laser power. The simple experiment presented in the previous
section does not give access to a quantitative measurement of the temperature in the droplet. However, we
can affirm that the temperature has exceeded the cloud point of PLL-g-PNIPAM (34 °C) as soon as the first
polymer aggregates have become visible in the droplet core. This qualitative approach can be used to
perform a first screening of the crucial parameters that govern this process.

Concretely, we prepared different emulsions containing 10 wt% of PLL-g-PNIPAM and different AuNPs
concentration (from 0.020 wt% to 0.100 wt%). The emulsions were successively thermalized at 20 °C,
24 °C and 28 °C, i.e. at temperatures closer and closer to the theoretical cloud point of 34 °C. Accordingly,
a smaller and smaller increase of temperature should be required to trigger the collapse transition of PLL-
g-PNIPAM entrapped in the droplets. At each temperature, we focused the 532-nm laser on different
droplets (with diameters ranging from 20 to 50 µm) and determined the light power which was required to
observe the formation of polymer granules. The results are shown in **Figure 4.12**.

<figure><img src="image_90.png" /><figcaption>**Figure 4.12.** Laser power required to trigger the collapse transition of PLL-g-PNIPAM in emulsion droplets containing various concentrations in AuNPs and thermalized at different temperatures (error bars = standard deviation on 5-8 droplets).</figcaption></figure>
The laser power which was required to trigger the collapse of PLL-g-PNIPAM decreased as the AuNPs concentration increased and/or the temperature of the sample increased. These results are consistent with the fact that (i) the highest AuNPs concentrations generate the highest temperature rises at a given laser power and (ii) decreasing the temperature gap that separates the actual temperature from the cloud point enables to decrease the laser power required to trigger the collapse.

The high dispersity which was observed for some experimental conditions may be associated to the influence of the droplet size on the required laser power. However, we did not identify any consistent evolution of the laser power with the droplet size, probably because the droplets were not uniformly irradiated (the 10-µm laser spot was smaller than the droplet diameters). At this stage, these experiments only provide qualitative arguments for choosing the suitable set of experimental parameters that may enable to trigger the collapse transition of PLL-g-PNIPAM with light. However, this method requires highly concentrated polymer solutions to visualize the collapse phenomenon, what is contradictory with the design of hollow capsules. In addition, this approach does not give access to quantitative values of temperature. The only quantitative information provided by this approach is if the critical temperature of 34 °C has been reached or not. Along this line, if we wanted to quantitatively screen the influence of the irradiation parameters, we should reproduce the previous collapse experiment with many thermoresponsive polymers exhibiting different collapse temperature, which could be tedious.

In the next section, we present an alternative microscopy approach which enables a quantitative *in situ* measurement of the temperature in the droplet, independently of the presence of the polymer.

### 4.4.2 Quantitative analysis of thermoplasmonic effects in emulsion based on quantitative phase microscopy

#### 4.4.2.1 The measurement of temperature at microscales

As demonstrated in the previous section, generating heat by thermoplasmonic effects is easy as it occurs as soon as AuNPs are irradiating with a sufficiently high laser power. The most challenging issue in any thermoplasmonic application is rather to be able to measure the temperature which is reached upon light excitation. Indeed, temperature gradients are generated at nano- and microscales that are obviously not accessible to standard thermometry techniques. The development of techniques for *in situ* measurement of temperature generated by thermoplasmonic systems is quite recent as it started in the last 2000s-early 2010s.²¹⁰ The first thermometry techniques implemented in thermoplasmonics were essentially based on
fluorescence properties of extrinsic reporters dispersed in the vicinity of the plasmonic structures. Indeed, many fluorescence features such as intensity, life-time, spectrum or anisotropy are temperature-dependent.²⁸⁹ Thus, measuring changes of these properties upon heating may enable to retrieve local temperature values, provided a suitable calibration. However, these approaches based on fluorescence may be limited in terms of spatial resolution, and may suffer from measurement biases related to photo- or thermo-bleaching of the fluorescent reporters.²¹⁰ As an alternative, some label-free and non-invasive imaging techniques – that may have already been used for “conventional” microscopy – have been adapted for temperature mapping of thermoplasmonic nanostructures. It was the case of quantitative phase microscopy techniques. Before detailing how they can be used in the context of thermoplasmonics, these techniques will be presented in more detail.

### 4.4.2.2 Quantitative phase microscopy

When a sample is subjected to a uniform light illumination, a distortion of the incident planar wavefront is observed as the light passes through refractive features (see **Figure 4.13.A**). Quantitative phase microscopy (QPM) techniques are based on the detection of this wavefront distortion to retrieve structural information on the sample. More precisely, QPM measures the optical path difference (OPD), also termed optical thickness in the literature, imprinted to the light wavefront upon sample imaging.²⁹⁰ The OPD, usually denoted $\delta$, is defined as the product of the thickness of the sample by the mismatch of refractive index between the sample and its environment (see Figure 4.13.B). It contains information on the geometrical profile and the refractive index of the distorting objects crossed by the illumination light. Since the OPD is directly proportional to the phase delay undergone by light during its propagation, the OPD image of the sample can be trivially converted into a phase image, hence the term of quantitative phase microscopy.

<figure><img src="image_91.png" /><figcaption>**Figure 4.13.** Physical values measured and imaged by quantitative phase microscopy techniques. A) Distortion of a planar light wavefront passing through a refractive object characterized by a geometrical profile $h(x, y)$ and a refractive index $n$. B) The wavefront distortion can be equivalently quantified either by the optical path difference $\delta(x, y)$ or by the phase delay $\Delta\phi(x, y)$, which are both directly proportional to the geometrical profile of the illuminated object.²⁹⁰</figcaption></figure>
Classical microscopy techniques are based on the measurement of light intensity. They are sensitive to diffusive or reflecting objects, and are accordingly less suitable for imaging samples which are mostly transparent. QPM techniques have the advantage to be highly sensitive to very small variations of refractive index, enabling the imaging of multi-media transparent samples, such as living cells,²⁹¹ with a high spatial resolution. For this reason, QPM techniques have been extensively used for the observation of biological samples, and has actually been for long restricted to this application field.²⁹²

The extension of QPM to the fields of physics and material science is only a recent trend, that has been instigated by Guillaume Baffou at the Fresnel Institute in Marseille. In this context, QPM turned out to be particularly efficient for the characterization of thermoplasmonic systems and for the precise mapping of temperature at microscales. Indeed, upon excitation, plasmonic nanostructures generate temperature gradients in their vicinity, which result in variations of refractive index in the surrounding medium. These local variations of refractivity may induce wavefront distortion of planar light used to image the sample (see **Figure 4.14**). By mapping the OPD upon plasmonic excitation, QPM techniques may thus enable to retrieve the temperature distribution around the plasmonic nanostructure.²⁹³

<figure><img src="image_92.png" /><figcaption>**Figure 4.14.** Distortion of a planar light wavefront passing through a temperature gradient locally generated by the photo-excitation of a plasmonic gold nanoparticle.</figcaption></figure>

The most common QPM techniques are based on the generation of interferences as it is a simple way to convert (invisible) phase information into (visible) intensity modulations. These techniques, such as digital holographic microscopy, rely on the splitting of the source light along two paths, one passing through the sample and the other not (reference path), to create an interference pattern which contains the phase information of the sample.²⁹⁴ These techniques benefit from high spatial resolution but suffer from instability issues. Indeed, extended propagation paths are required for the interfering light beams. Thus, the interference-based optical set-ups are highly sensitive to environmental perturbations (vibrations, temperature variations), resulting in ersatz fringes or speckle features which deteriorate the quality of the final phase image.
Alternative QPM techniques, that do not involve extended set-up, can be used for phase imaging of thermoplasmonic systems. Grating-shadow phase microscopy is one of these techniques and will be presented in the next section.

### 4.4.2.3 The technique of grating-shadow phase microscopy

Grating-shadow phase microscopy (GSPM) is a QPM technique based on wavefront sensing as described in the previous section. This technique follows on from precursor works of Primot in the mid-90s and has been commonly termed quadriwave lateral shearing interferometry (QLSI) in the literature.²⁹⁰ The name of grating-shadow phase microscopy has been recently proposed as a more simple and more relevant alternative to better understand and disseminate this microscopy technique.²⁹⁰

GSPM relies on an imaging system made of the association of a diffraction grating and a common CCD camera. The grating is a bidimensional periodic arrangement of transmitting glass squares (windows) separated by opaque horizontal and vertical lines. A chessboard-like phase pattern is imprinted on the grating such that adjacent windows alternatively induce a phase delay of 0 or $\pi$ to the transmitted light (see Figure 4.15).

<figure><img src="image_93.png" /><figcaption>**Figure 4.15.** Grafting-Shadow Phase Microscopy imaging system composed of a bidimensional $0 - \pi$ chessboard grating placed just in front of a CCD camera. The grating is typically made of 300 x 300 $\begin{vmatrix} 0 & \pi \\ \pi & 0 \end{vmatrix}$ unit cells.</figcaption></figure>

The image formed on the camera sensor upon illumination of the grating is called interferogram. It is made of fringes resulting from the interference between the multiple diffracted beams coming out of the grating. Thanks to the $0 - \pi$ chessboard pattern and the specific dimensions of the grating, only symmetrical first diffraction orders along two orthogonal directions participate significantly to the interference phenomenon
(Figure 4.16.A). In these conditions, the interferogram generated by a planar irradiation is the exact shadow of the grating (i.e. a periodic bidimensional arrangement of bright squares separated by dark lines), as if light passed in straight line through the windows without undergoing diffraction effects (see Figure 4.16.B).²⁹⁰

<figure><img src="image_94.png" /><figcaption>**Figure 4.16.** Wavefront sensing with grating-shadow phase microscopy. A) Formation of the chessboard grating's shadow by interference of 4 first diffraction orders along two orthogonal directions upon transmission of a planar incident light. B) Distortion of the shadow pattern induced by a distortion of the incident light wavefront: tilting or curving the wavefront may induce the translation or spreading/shrinking of the shadow, respectively.²⁹⁰</figcaption></figure>

Interestingly, a local distortion of the incident wavefront results in a local deformation of the shadow pattern (see Figure 4.16.B). As a consequence, the interferogram generated by the illumination of a complex sample is not a regular but a distorted grating shadow that “geometrically” contains the phase information of the sample. An OPD image of the sample can be retrieved from the distorted interferogram thanks to an algorithm based on image processing in the Fourier space (Figure 4.17).²⁹⁵
<figure><img src="image_95.png" /><figcaption>**Figure 4.17.** Main steps involved in the GSPM algorithm that enables to retrieve a phase (or OPD) map from the distortion of the interferogram fringes. The 2D Fourier transform (FT) of the interferogram reveals orthogonal first order diffraction spots ($\tilde{H}_x$ and $\tilde{H}_y$) that can be demodulated (isolated) in the Fourier space. The inverse Fourier transform (FT$^{-1}$) of these spots yields two complex images, whose arguments ($\mathrm{Arg}(H_x)$ and $\mathrm{Arg}(H_y)$) correspond to wavefront gradient maps along the two orthogonal directions. A 2D integration of these gradient maps eventually yields the phase image of the sample.²⁹⁵</figcaption></figure>

GSPM has many advantages compared to other common QPM techniques. Thanks to the high pixel density on the grating, GSPM benefits from a high spatial resolution, which is eventually limited by the optical resolution of the camera. It also benefits from a high sensitivity, the typical standard deviation of an OPD noise image being ~ 0.1 nm.²⁹⁰ With these performances, GSPM turned out to be particularly appropriate for the imaging and characterization of nanoparticles (with sizes down to 60 nm in diameter).²⁹⁶

As previously mentioned, the light coming out of the grating behaves as if it has not been subjected to diffraction. A reliable image of the interferogram can thus be formed by placing the sensor of the camera only 1 mm behind the grating. The interference phenomenon occurring on short distances, GSPM is not sensitive to environmental perturbations and is thus more stable and robust than other interference-based QPM techniques. In addition, GSPM does not require temporal coherence for the illumination, and is even more effective with a polychromatic source.²⁹⁰ Various GSPM devices have been commercialized by the company PHASICS, and can be easily implemented on any optical set-ups, including on the side port of commercial microscopes.

#### 4.4.2.4 Use of GSPM for imaging of thermoplasmonic systems

Based on wavefront sensing of refractive index variations as depicted in Figure 4.14, GSPM has been successfully implemented for the characterization of thermoplasmonic systems and the measurement of temperature gradients at microscales. Baffou et al. developed a suitable algorithm to retrieve, from the OPD image obtained by GSPM, the distribution of heating power and temperature gradients generated by gold nanostructures upon plasmonic excitation (see **Figure 4.18**).²⁹³ Note that two contributions may be observed
in raw OPD images of a plasmonic nanostructure: the OPD due to the nanostructure itself (highly refractive compared to its environment) and the OPD due to the local variations of refractive index induced by heating. This latter contribution can be isolated by simply subtracting the OPD image of the nanostructure from the OPD image during photo-excitation.

<figure><img src="image_96.png" /><figcaption>**Figure 4.18.** GSPM mapping of light-induced heat generation in a gold nanostructure excited at 808 nm and illuminated with a white source. a) SEM image of the targeted gold nanostructure. b) Map of OPD generated by the local heating of the nanostructure upon plasmonic excitation (the OPD due to the nanostructure itself was subtracted). c) Map of heating power density dissipated by the nanostructure, retrieved from image processing of the OPD map with a suitable algorithm. d) Temperature distribution around the nanostructure, also deduced from image processing of the OPD map.²⁹³</figcaption></figure>

The image processing developed by Baffou et al. to retrieve temperature maps is restricted to the characterization of 0D, 1D or 2D heat sources, i.e. individual nanoparticles (nanospheres or nanowires) or arrays of such nanoparticles. This approach is no longer suitable for 3D heat source distributions (i.e. sources much larger than the illumination wavelength along the three spatial dimensions). Indeed, since the OPD maps correspond to a 2D projection of the sample phase, all the contributions gathered along the direction of light propagation are overlapped in the final image. Complex deconvolution processes may be required to retrieve such 3D source distributions, and have never been developed so far. For now, direct 3D temperature mapping from raw images of OPD has been restricted to the study of 2D heat sources (by reconstruction from stacked OPD images).²⁹⁷

For more complex heat source distributions, simulations are required to correlate OPD images and temperature distributions. Our system (AuNPs-loaded emulsion droplets) is a typical example of complex 3D heat source distribution for which the temperature field cannot be analytically retrieved from OPD maps provided by GSPM. COMSOL simulations of temperature distributions on an approximate model may be
helpful to correlate the OPD images of the sample to actual temperatures reached *in situ*. The following
sections will present and discuss this approach.

4.4.2.5 Experimental set-up and measurement of OPD profiles

The GSPM experiments were performed in Guillaume Baffou's lab at the Fresnel Institute in Marseille.
The experimental set-up is shown in **Figure 4.19**. It was used to image the OPD induced by AuNPs-loaded
emulsion droplets upon thermoplasmonic excitation. In the following, the sample will consist in a water-
in-oil emulsions containing 1 wt% of PLL-g-PNIPAM and 0.025 wt% of PVP-coated AuNPs in the aqueous
phase and 0.05 wt% of Krytox in the oil phase. The emulsion was injected in a capillary with an internal
height of 100 µm.

The sample was illuminated from the top with a 625-nm (red) planar illumination. The OPD image of the sample was recorded with the GSPM wavefront sensing device presented in Figure 4.15. The plasmonic excitation of AuNPs was performed by irradiated the sample from the bottom with a 532-nm (green) laser. The diameter of the laser spot in the focal plane of the objective was set to 50 µm (a uniform irradiation was thus ensured for droplets smaller than 50 µm). The laser power delivered to the sample was typically between 0 and ~ 40 mW, which correspond to an irradiance (i.e. a power per unit area) between 0 and ~ 20 µW/µm².

<figure><img src="image_97.png" /><figcaption>**Figure 4.19.** Experimental grating-shadow phase microscopy set up. The sample is a water-in-oil emulsion injected in a glass capillary. The water droplets (blue disks) are loaded with 1 wt% of PLL-g-PNIPAM (not showed) and 0.025 wt% AuNPs (red dots). The sample is illuminated by the top with a 625-nm diode and excited by the bottom with a 532-nm laser (in green). The OPD image of the sample is recorded with the GSPM device previously described.</figcaption></figure>
In the following, the protocol for imaging and post-processing will be detailed for a representative droplet with a diameter of 30 µm. **Figure 4.20** shows the interferogram of the droplet provided by GSPM. A zoom on the edge of the droplet highlights the distortion of the grating shadow. From this interferogram, intensity and OPD images of the droplet are generated by the GSPM software. In the OPD image, the droplet appears as a bright white disk standing out of the dark background generated by the continuous oil phase. This sharp contrast suggests that the presence of the water droplet imprints a large OPD signal, generated by the mismatch of refractive index between water and oil. A crucial point to have mind for the quantitative interpretation of such image is the fact that an OPD value measures a “difference”. Accordingly, OPD values can be retrieved from a sample image by measuring differences of “grey level” between two regions on the same image.

From the raw OPD image showed in Figure 4.20, we could deduce the OPD generated by the water droplet using the oil background as a “zero” baseline. The droplet profile indicates a maximum OPD of $\delta = 900$ nm reached at the center (projection of the longest droplet portion crossed by the light). This positive value confirms that the refractive index of water is higher than the oil’s one. This latter can even be deduced from the formula presented in Figure 4.13: $n_O = n_W - \delta/d = 1.30$ with $d = 30$ µm the droplet diameter and $n_W = 1.33$ the refractive index of the water droplet assimilated to pure water (the presence of the polymer and the AuNPs are neglected). This experimental value of $n_O$ fits exactly with the value indicated by the commercial datasheet of the oil.

<figure><img src="image_98.png" /><figcaption>**Figure 4.20.** Interferogram, intensity image, and OPD image of a 30-µm droplet recorded by GSPM (scale bar = 15 µm). The zoom on the droplet edge in the interferogram reveals the distorted grating shadow (scale bar = 2 µm). The OPD profile of the droplet highlight the high contrast between the water droplet and the continuous oil phase ($\delta = 900$ nm at the center of the droplet, in $X = 0$).</figcaption></figure>
The previous images were obtained in absence of plasmonic excitation. When the green laser was turned on, heat was dissipated by the AuNPs dispersed in the water droplet, inducing an additional contribution to the OPD signal. To isolate this thermoplasmonic contribution, a reference image of the droplet was taken before laser irradiation and was subtracted from the image recorded upon laser irradiation. Concretely, the gradients of OPD were uniformly set to 0 just before irradiation in order to mask the water/oil mismatch of refractive index (see **Figure 4.21.A**).

Upon laser irradiation, an OPD pattern appeared and was characterized by a dark and diffuse corona at the periphery of the droplet, surrounded by a bright oil background (see Figure 4.21.A). This distribution of grey values suggests that the OPD is negative in the droplet (the reference being taken in the oil phase), which is consistent with a local decrease of the refractive index of the medium upon heating (the $\frac{dn}{dT}$ coefficient of liquids is negative). As the manipulation and graphical representation of negative values can be counterintuitive and confusing, the data treatment of GSPM images was systematically done with *absolute* values of OPD.

To retrieve quantitative values, the OPD reference (the “zero”) was taken in a region which has not been perturbed by temperature gradients generated by thermoplasmonic effects. The ideal choice would be to take this reference infinitely far from the droplet but, since the imaging field of view was limited, we were compelled to arbitrarily set the OPD reference as far as possible from the droplet, but we were aware that it did not correspond to a real “zero”.

Figure 4.21.B shows the OPD radial profile of the droplet upon laser irradiation, plotted in *absolute* values with the OPD origin set *arbitrarily* far from the droplet center. This radial profile shows that a high OPD is generated in the droplet (with a maximum value reached at the periphery) and then diffuses in the oil phase with a characteristic spatial decay.

<figure><img src="image_99.png" /><figcaption>**Figure 4.21.** OPD images of the droplet before and during laser irradiation ($\lambda$ = 532 nm, 12.7 µW/µm²) and the corresponding radial profiles (plotted in absolute value and arbitrarily set to 0 far from the droplet). The OPD was uniformly set to 0 before irradiation to specifically isolate the contribution of the thermoplasmonic effect on the wavefront distortion upon irradiation (scale bars = 20 µm).</figcaption></figure>
### 4.4.2.6 Mathematical modelling of the decay portion of the OPD radial profile

In theory, the OPD decay observed in the oil phase should asymptotically reach 0 infinitely far from the droplet but it was arbitrarily set to 0 at a finite distance due to the spatial limitation of the field of view. A theoretical fit of this decay should enable to properly rescale the OPD curve and to retrieve the actual values of OPD. A simple model is proposed to describe the system and predict the evolution of the OPD in the oil phase. An axisymmetric side-view representation of the droplet is shown in **Figure 4.22**. The z axis corresponds to the axis of light propagation (downwards for the illumination and upwards for the excitation laser) and its origin is set on the upper capillary side. Since the aqueous phase is less dense than the oil phase, the water droplet is in contact with the upper capillary side. According to the side view micrographs shown in section 3.3.1 (Chapter 3, Figure 3.7), the top of the droplet is slightly flattened by the contact with the glass slide, from a distance denoted *e* and which is typically ~ 1 µm. Accordingly, the center of the droplet is shifted in *z* = −*R* + *e* where *R* is the droplet radius. The x axis goes along the upper capillary side and its origin is set vertically above the center of the droplet. Since the system show a rotational invariance around the z axis, the analysis of the system can be restricted to the (*xz*) plane to deduce radial properties.

<figure><img src="image_100.png" /><figcaption>**Figure 4.22.** Axisymmetric side-view representation of the system. The water droplet of radius *R* (containing the AuNPs) is in contact with the upper capillary side (the droplet is flattened on a distance *e*). The center of the water droplet is set at the position *x* = 0, *z* = −*R* + *e*. The heat generated by the droplet (assimilated to a uniform spherical heat source delivering a power *Q*) creates a local increase of temperature *δT*(*x*, *z*) at each distance *r*(*x*, *z*) from the droplet center. This local temperature rise induces a local variation of refractive index in the oil phase *δn*₀(*x*, *z*) that can be integrated along the *z* axis to retrieve the OPD radial profile.</figcaption></figure>

Upon plasmonic excitation of AuNPs, a certain amount of heat, denoted *Q*, is produced in the droplet and is assumed to be uniformly distributed such that the droplet can be seen as an isotropic spherical heat source
(we neglect the droplet flattening in the model). Following the heat equation, the progressive diffusion of heat through the medium generates temperature gradients. Assuming that the steady state is rapidly reached, a simple expression of the temperature field in the oil phase can be derived:

$$T(r > R) = \frac{Q}{4\pi\kappa_o r} + T_\infty$$

In this expression, r designates the radial distance from the droplet center. By restricting this equation to the (xz) plane, the distance r is expressed in cartesian coordinates as follows: $r = \sqrt{x^2 + (z + R - e)^2}$ (see schematic in Figure 4.22). $\kappa_o = 0.07 \text{ W} \cdot \text{m}^{-1} \cdot \text{K}^{-1}$ designates the thermal conductivity of the oil phase (commercial datasheet) and $T_\infty$ corresponds to the temperature infinitely far from the heat source (for $x \to \infty$) and is equal to room temperature, i.e. $T_\infty = 273 \text{ K}$. This equation can be rewritten in terms of temperature rise $\Delta T = T(r) - T_\infty$ and in cartesian coordinates for $x > R$:

$$\Delta T(x > R, z) = \frac{Q}{4\pi\kappa_o \sqrt{x^2 + (z + R - e)^2}}$$

From a physical point of view, an infinitesimal variation of temperature $\delta T$ in the oil phase locally induces an infinitesimal variation of refractive index of the oil $\delta n_o$, which is given by $\delta n_o = \frac{dn_o}{dT} \delta T$ with $\frac{dn_o}{dT} = -3.16 \times 10^{-4} \text{ K}^{-1}$ (determined by GSPM, see section 7.9.3 in Appendix A).

Local variations of refractive index are at the origin of the OPD signature observed upon laser irradiation. The OPD radial profile in the oil phase (along the x direction) can be calculated by integration of the infinitesimal variations of refractive index of the oil along the z direction (i.e. along the direction of light transmission): $\text{OPD}(x > R) = \int_{-H}^{0} \delta n_o(x, z) dz = \int_{-H}^{0} \frac{dn_o}{dT} \delta T(x, z) dz$

Assuming that $\frac{dn_o}{dT}$ is a constant and that $\delta T(x, z)$ can be assimilated to $\Delta T(x, z)$, we obtain:

$$\text{OPD}(x > R) = \frac{dn_o}{dT} \frac{Q}{4\pi\kappa_o} \int_{-H}^{0} \frac{dz}{\sqrt{x^2 + (z + R - e)^2}}$$

The calculation of the integral eventually yields an analytical expression for the OPD in the oil phase for $x > R$:

$$\text{OPD}(x > R) = \frac{dn_o}{dT} \frac{Q}{4\pi\kappa_o} \ln \left[ \frac{R - e + \sqrt{x^2 + (R - e)^2}}{R - e - H + \sqrt{x^2 + (R - e - H)^2}} \right] = A \times f(x)$$
where $A = \frac{dn_O Q}{dT 4\pi\kappa_O}$ is a constant depending on the heat source power and on physico-chemical parameters of the oil phase, and $f(x) = \ln\left[\frac{R-e+\sqrt{x^2+(R-e)^2}}{R-e-H+\sqrt{x^2+(R-e-H)^2}}\right]$ is a decreasing function of $x$ depending on the structural parameters of the droplet (radius $R$, flattening distance $e$) and on the capillary interior thickness $H$.

Since $\frac{dn_O}{dT} < 0$, the constant $A$ as well as the final OPD are negative, which is consistent with the experimental observations. But as previously mentioned, the OPD data are treated in *absolute* values for the sake of convenience. With that in mind, the theoretical OPD is a decreasing function that tends to 0 for $x \to \infty$. This asymptotical behavior was not observed in the experimental OPD profile (since the curve was artificially set to 0 at a finite distance from the droplet center), but can be retrieved by applying a suitable offset to the experimental curve. The offset value can be introduced in the form of an additive constant $B$ in the analytical expression of the OPD used for the fit, now reading: $OPD(x) = A \times f(x) - B$.

In the fitting procedure, the constants $A$ and $B$ were let free to optimization but the definition of the function $f$ was fed with the experimental values of $R$, $e$ and $H$. The value of $B$ given by the fit was used to shift up the experimental curve and thus retrieve the asymptotical behavior of the OPD profile. The fitting procedure is demonstrated on a 30-µm droplet ($R = 15$ µm) for different laser irradiances (see **Figure 4.23**). The values of $e$ and $H$ were set to 1 µm and 100 µm respectively. The fit was only applied to the tail of the decay portion as it gave less reliable results in the region near the droplet. This hiatus is likely due to an over-simplification of the model to derive the analytical expression of the radial OPD. In particular, we used an expression for the temperature field which is only valid for an isotropic source heat in a uniform medium. We did not consider the presence of the capillary sides that (i) break the isotropy of heat diffusion and (ii) induces a local modification of the thermal conductivity (along the oil/glass interface, the effective conductivity is the mean of the conductivity of the two media). These effects are predominant near the droplet but can be neglected far from it.
<figure><img src="image_101.png" /><figcaption>**Figure 4.23.** Rescaling of the experimental OPD radial profiles by an appropriate fit of the decay portion. Left: Experimental OPD radial profiles of the 30-µm droplet under study for different laser irradiances. All the curves were artificially set to 0 at a distance arbitrarily far from the droplet center. The tail of the decay portion of the profiles was fitted with the analytical function derived from the mathematical model: $A \times f(x) - B$ where A and B were unfixed parameters and $f(x) = \ln\left[\frac{R-e+\sqrt{x^2+(R-e)^2}}{R-e-H+\sqrt{x^2+(R-e-H)^2}}\right]$ with R = 15 µm, e = 1 µm, H = 100 µm. Right: The curves were shifted up by addition of the offset parameter B deduced from the fit. Doing that, the physically expected asymptotical behavior of the OPD profiles was retrieved, i.e. $\text{OPD} \to 0$ when $x \to \infty$.</figcaption></figure>

After applying the suitable offset, the experimental OPD profiles exhibit the expected asymptotical behavior (see Figure 4.23). The thermal signature of the droplet was quantified by the measurement of the OPD value at the center of the droplet, termed “core OPD”. This parameter was plotted as a function of the irradiance for the 30-µm droplet under study in **Figure 4.24**.

<figure><img src="image_102.png" /><figcaption>**Figure 4.24.** Evolution of the droplet core OPD value extracted from the rescaled experimental profiles as a function of the laser irradiance.</figcaption></figure>
At this stage, GSPM only gives quantitative OPD values in the water droplet upon thermoplasmonic excitation. A quantitative value of temperature cannot be directly deduced from the raw OPD measurement. Reciprocally, an OPD radial profile could be easily calculated from a given temperature distribution, for instance generated by simulation tools. This “retro-active” approach was used to correlate the core OPD values obtained experimentally to the temperature locally reached in the droplet.

### 4.4.2.7 Simulation of OPD profiles with COMSOL

COMSOL simulations were performed on the same 2D axisymmetric geometrical configuration than presented in the previous section to predict the temperature distribution in the droplet and the surrounding oil. To enable the direct comparison with the experimental shown in the previous section, the simulation will be demonstrated on a droplet with a diameter of 30 µm. **Figure 4.26.**A shows the simulation domain, with the different media and the corresponding thermal diffusion coefficients. The water droplet (radius *R*, volume *V*) is defined as a uniform heat source<sup>‡</sup> which delivers a heating power *Q* (set as an input parameter in the simulation).

To estimate the relevant range of heat power to test in the simulation, the heat power experimentally generated in the droplets was derived from simple considerations regarding the formulation of the droplets and the conditions of irradiation. We assumed that the total heat power *Q* which was dissipated in a droplet was the sum of the heat power dissipated by each individual AuNP loaded in the droplet. Hence, in a droplet of volume *V* containing a concentration of AuNPs *C*<sub>AuNPs</sub> (0.05 wt% = 0.45 NPs/µm<sup>3</sup> according to the datasheet) and uniformly irradiated by an irradiance *I*, *Q* was estimated by the following formula: *Q* = *C*<sub>AuNPs</sub>*Vσ*<sub>abs</sub>*I* where *σ*<sub>abs</sub> is the absorption cross-section of a single AuNP, which is equal to 6 × 10<sup>-3</sup> nm<sup>2</sup> for a AuNP with a diameter of 50 nm.<sup>210</sup> **Figure 4.25** shows that the heat power dissipated in droplets of different size during the irradiation experiments is of the order of magnitude of a fraction of mW. This is typically the values of heating power which were tested in the simulation (0.1 – 0.8 mW).

<footnote>‡ The relevancy of the choice of a uniform heat source distribution will be discussed at the end of this section (see **Figure 4.28**).</footnote>
<figure><img src="image_103.png" /><figcaption>**Figure 4.25.** Heat power dissipated in AuNPs-containing droplets of different size upon uniform irradiation at 532 nm with varying irradiance.</figcaption></figure>

In the simulation, the temperature field is obtained by solving the stationary heat equation in all the domain ($-\kappa_W \Delta T = Q/V$ in the droplet and $\Delta T = 0$ out of the droplet). For that, COMSOL uses the finite element method on a triangular integration mesh which is shown in Figure 4.26.B. The mesh size was refined at the droplet periphery to precisely assess the temperature distribution around the water/oil interface.

Figure 4.26.C shows the temperature map resulting from the numerical simulation for $Q = 0.3 \text{ mW}$. The highest temperature is reached in the bottom half-portion of the droplet (here ~ 12 °C for $Q = 0.3 \text{ mW}$). Due to the presence of the capillary side on top of the droplet, a “pseudo-radial” temperature gradient is generated in the oil phase surrounding the side and bottom part of droplet. The spatial extension of this gradient is proportional to the heating power (here typically ~ 70 µm from the droplet center for $Q = 0.3 \text{ mW}$).
<figure><img src="image_104.png" /><figcaption>**Figure 4.26.** COMSOL simulation of the temperature field generated by a 30-µm droplet. A) Axisymmetric projection of the simulation domain (restricted to the direct vicinity of the droplet). The thermal diffusion coefficients $κ$ and the expression of the heat equation are indicated for each medium. The simulation domain was contained in a sphere which was 500 µm in radius (way larger than the droplet radius), used to fix the boundary condition $T_∞ = 273 K$. B) Integration mesh used to solve the heat equation. C) Temperature map obtained for a heating power $Q = 0.3 \text{ mW}$ uniformly distributed in the water droplet.</figcaption></figure>

The temperature map provided by simulations can be post-processed to retrieve the local temperature rise $\Delta T$ and the corresponding variation of refractive index $\Delta n$ in the droplet and the oil phase. The $\Delta n$ map was obtained by multiplying each pixel of the $\Delta T$ map with the appropriate $\frac{dn}{dT}$ coefficient, namely $-0.90 \times 10^{-4} \text{ K}^{-1}$ in the water droplet and $-3.16 \times 10^{-4} \text{ K}^{-1}$ in the oil phase. **Figure 4.27.** A and B show the $\Delta T$ and $\Delta n$ maps of a 30-µm droplet for $Q = 0.3 \text{ mW}$ (the domain has been restricted to the capillary interior volume on a length of 100 µm). The $\Delta n$ map reveals that the largest variations of refractive index (in absolute values) are obtained in a thin oil layer surrounding the water droplet, and not in the droplet core
where the highest temperatures are yet reached. This corona-like pattern is essentially due the highest thermal sensitivity of the oil phase compared to the water phase (the $\frac{dn}{dT}$ coefficient of oil is 3.5 times higher than the one of water).

<figure><img src="image_105.png" /><figcaption>A)</figcaption></figure>

<figure><img src="image_106.png" /><figcaption>B)</figcaption></figure>

<figure><img src="image_107.png" /></figure>

$$OPD(x) = \sum_{px/z} |\Delta n(x, z)|$$

<figure><img src="image_108.png" /><figcaption>C)</figcaption></figure>

**Figure 4.27.** Image processing of the temperature map to retrieve numerical OPD profiles (demonstrated on a 30-µm droplet). A) Maps representing the variations of temperature and refractive index in the domain, deduced from the raw temperature map provided by the simulation (for a heating power of 30 mW). B) OPD radial profile calculated by numerical integration of the map of variations of refractive index along the z direction, i.e. by summing the intensity of the pixels (px) along the columns of the map. C) Correlation between the OPD (measured on profile C) and the maximum temperature variation (measured on map A) in the core of the droplet. This process has been reproduced for different values of heating power used in the simulation.

A numerical radial OPD profile can be deduced from the $\Delta n$ image by summing the intensity of the pixels along the columns (which is the numerical equivalence of an integral along the z direction). This operation is made with absolute values of $\Delta n$ to get positive OPD profiles (to be consistent with the analysis of the experimental data). Figure 4.27.C shows the OPD radial profile calculated from the $\Delta n$ map showed in
Figure 4.27.B (for $Q = 0.3 \text{ mW}$). This profile generated numerically shows the same qualitative features than the profiles obtained experimentally (see Figure 4.23), suggesting that our system can be effectively simulated. In addition, the value of the core OPD measured on the numerical profile (at the center of the droplet) can be directly correlated to the maximum temperature rise in the droplet core $\Delta T_{max}$ (measured on the $\Delta T$ map). A linear relationship between these two parameters can be eventually obtained by running the simulation and numerical processing on the same droplet geometry but for different heating powers. Figure 4.27.D shows this correlation curve for the model 30-µm droplet under study. For any droplet of radius $R$, a linear relationship reading $\Delta T_{max} = \alpha(R) \times \text{OPD}_{core}$ (with $\alpha(R)$ a coefficient depending on the droplet radius) can be deduced from simulation. This relationship will be applied to the experimental OPD profiles to retrieve the temperature rise reached in the droplet.

The simulations presented above have been performed considering the droplet as a uniformly distributed heat source. This would correspond to a uniform distribution of the AuNPs within the volume of the droplet and not to a distribution in the polymer shell at the periphery of the droplet. This uniform distribution of the AuNPs should be relevant at low temperature (below the cloud point of PLL-g-PNIPAM) as demonstrated with fluoNPs in Section 4.3.1. After exceeding the cloud point of PLL-g-PNIPAM upon thermoplasmonic excitation (at sufficiently high laser intensity), the AuNPs – as the fluoNPs – are likely to accumulate in the polymer shell despite the relatively low interaction between AuNPs and PLL-g-PNIPAM (see Section 4.2.1). Accordingly, the distribution of the heat source is likely to be modified during the irradiation experiments, resulting in a modification of the correlation between the core OPD (which is measured experimentally) and the maximum temperature rise (which is predicted numerically).

To address this issue, different configurations of the heat source distribution were simulated in a model droplet of 30 µm in diameter: the heat source ($Q = 0.3 \text{ mW}$) was either uniformly distributed in the droplet core (as in the previous section), or confined in a 1 µm-thick shell at the periphery of the droplet (as expected after co-segregation of the AuNPs in the polymer interfacial layer above the cloud point). The influence of the contact between the droplet and the upper capillary side was also tested at the same time than the heat source distribution: the droplet was either in flattening contact (on a thickness of 1 µm as used in the previous section), or in point contact with the capillary. These four configurations are shown in **Figure 4.28.A**. The corresponding maximum temperature rise in the droplet core and radial OPD profile, obtained by simulation, are shown in Figure 4.28.B and Figure 4.28.C, respectively.

The two different heat source distributions (core vs shell) give approximately the same temperature rise and the same core OPD (see Config 1 vs 2, or 3 vs 4 in Figure 4.28.B and C), suggesting that these parameters are essentially dictated by the *value* of the heating power (here set at $0.3 \text{ mW}$) and not by the *distribution* of the heat source. Interestingly, the geometrical nature of the contact between the droplet and
the upper capillary side has a stronger influence on the temperature rise and core OPD: both are almost 10 % higher for a point contact than for a flattening contact (see Config 1 vs 3, or 2 vs 4). The lower temperature rise in the case of a flattening contact is due to the fact that glass dissipates heat more efficiently than oil ($κ_G = 1 W.m⁻¹.K⁻¹$ vs $κ_O = 0.07 W.m⁻¹.K⁻¹$). The higher core OPD in the case of a point contact is due to the fact that a thin oil layer is partially intercalated between the droplet and the capillary, and is on the pathway of the light which crosses the droplet, hence contributing to the core OPD. This thin oil layer does not exist in the case of a flattening contact. Experimentally, the flattening contact is more relevant than a point contact, as previously mentioned in Section 4.4.2.6.

<figure><img src="image_109.png" /><figcaption>**Figure 4.28.** Influence of the distribution of the heat source on the maximum temperature rise and OPD in the droplet core obtained by simulation (performed on a 30-µm diameter droplet and a heat power of $Q = 0.3 mW$). A) Schematic representation of the different configurations tested (blue = water droplet, yellow = oil phase, red = heat source location). The droplet is either in flattening contact (Config 1 and 2) or in point contact (Config 3 and 4) with the upper capillary side. The heat source is either uniformly distributed in the droplet (Config 1 and 3) or confined in a 1 µm-thick shell at the periphery of the droplet (Config 2 and 4). B) Simulated maximum temperature rise generated in the droplet core for the different configurations. C) Simulated core OPD profiles for the different configurations.</figcaption></figure>

Eventually, the temperature rise and core OPD obtained with the Configuration 1 (already used in the previous section) can be effectively used to describe our experimental system whatever the actual distribution of the AuNPs in the droplets (uniformly distributed in the core or segregated in the polymer shell).
### 4.4.2.8 Conversion of experimental OPD in temperature rise

Thanks to the $OPD_{core}-\Delta T_{max}$ correlation obtained numerically in the previous section, we can convert the core OPD measured in the experimental profiles into a maximum temperature rise reached in the droplet core, as demonstrated in **Figure 4.29** for the droplet under study. The values of temperature rise (right axis) have been deduced from the core OPD values (left axis) via the linear relationship introduced in Figure 4.27.C.

<figure><img src="image_110.png" /><figcaption>**Figure 4.29.** Determination of the temperature rise reached in the droplet core from the correlation between the experimental OPD value (left axis) and the simulated temperature map (right axis, see Figure 4.27.C) in a 30-µm droplet for different laser irradiances.</figcaption></figure>

This approach coupling experimental data and simulation enables to retrieve quantitative values of temperature rise in AuNPs-loaded droplets subjected to thermoplasmonic effects. For the 30-µm droplet under study, the temperature in the core of the droplet can reach 45 °C (rise of 20 °C) for the higher irradiance that was tested, i.e. a temperature well above the LCST of PLL-g-PNIPAM (34 °C).

This thermometry approach was used for droplets of different diameters: 20, 23, 27 and 30 µm (the latter being the droplet under study in the previous figures). For each droplet size, the evolution of the core OPD as a function of the laser irradiance was measured experimentally. Then the $OPD-\Delta T$ correlation was determined with simulations and was used to retrieve the temperature rise in the droplet core as function of the laser irradiance (see **Figure 4.30**).
<figure><img src="image_111.png" /><figcaption>**Figure 4.30.** Evolution of the maximum temperature rise reached in the droplet core as a function of the laser irradiance for different droplet diameters.</figcaption></figure>

A general trend appears in Figure 4.30: the largest the droplet, the highest the temperature rise in the droplet core. It may be trivially explained by the fact that smaller droplets contain a lower amount of AuNPs, resulting in a lower production of heat upon irradiation. As an illustrative example, the temperature increases by less than 8 °C in a 20-µm droplet versus more than 20 °C in a 30-µm droplet for the same irradiance of 20 µW/µm². Moreover, the quasi-superimposition of the curves for the 23 and 27 µm droplets suggests that these two droplets contain a similar number of AuNPs, betraying concentration heterogeneities between droplets within the same emulsion. To achieve a better control on the droplet size and AuNPs loading, an interesting way to explore would be the production of the emulsion by microfluidics. Thanks to this approach, a proper calibration of the temperature rise as a function of the droplet size and the AuNPs concentration may be easier to implement.

These results demonstrate that GSPM can be successfully used as a micro-thermometry technique to access the temperature generated *in situ* by thermoplasmonic excitation of individual droplets. However, this approach can face some experimental difficulties that need to be mentioned.

First, the technique is highly sensitive to any small movement of the droplet upon irradiation. Since a reference OPD image is taken *at a given droplet position* before irradiation, any translational movement of the droplet during the measurements imprints a strong OPD signature on the image. This signal is associated to the high water-oil phase contrast that may mask the contribution of thermoplasmonic effects and alter the associated OPD radial profiles (especially at the periphery of the droplet).

Second, low concentrations of PLL-g-PNIPAM are required to avoid the formation of micron-size polymer aggregates in the core of the droplets at high temperature (i.e. at high irradiance). Indeed, these
aggregates may also contribute to the wavefront distortion and may perturb the quantitative analysis of the heating-induced OPD. This effect was typically observed at 2 wt% and 5 wt% in PLL-g-PNIPAM. Accordingly, we used a concentration of 1 wt% for which the formation of visible aggregates was not observed.

Third, the droplets can undergo stability issues over prolonged periods of irradiation, that can eventually lead to droplet burst or polymer leakage in the oil phase. These negative effects were particularly observed at high irradiance (> 15 µW/µm²) and/or high AuNPs concentration. That is why the irradiance range was limited (< 25 µW/µm²) and the AuNPs concentration was restricted to 0.05 wt%.

### 4.4.3 Application: light-induced accumulation of nanoparticles in PLL-g-PNIPAM shells

In the previous sections, we demonstrated that the collapse transition of PLL-g-PNIPAM in emulsion droplets can be induced by the photoexcitation of AuNPs and that the temperature reached in the core of the droplets can be measured thanks to grating-shadow phase microscopy. We proposed to apply this light responsiveness to trigger the thermo-induced accumulation of NPs in the polymer shell (described in section 4.3.1) with a better spatio-temporal control. The idea was to mix fluorescent NeutravNPs and plasmonic AuNPs with PLL-g-PNIPAM in the emulsion droplets. As demonstrated in section 4.3.1, an accumulation of NeutravNPs can be observed at the droplet periphery upon heating, mediated by the collapse transition of PLL-g-PNIPAM. In absence of AuNPs, the heating process was performed at the scale of the whole sample within a few minutes. The addition of AuNPs in the aqueous phase may enable to trigger the thermo-induced interfacial accumulation upon laser excitation at the level of an individual droplet and quasi-instantaneously.

As a proof of concept, an aqueous solution containing 1 wt% of PLL-g-PNIPAM, 0.01 wt% of NeutravNPs and 0.05 wt% of AuNPs was prepared and emulsified in an oil phase containing 0.05 wt% of Krytox. As in section 4.3.1, the distribution of NeutravNPs in the droplets was assessed with confocal imaging. The plasmonic excitation could not be directly implemented within the confocal microscope since it was not equipped with sufficiently powerful laser source in the green range. The plasmonic excitation was performed on the commercial microscope used for bright field imaging (see Chapter 3), equipped with a 532-nm laser diode that can deliver a nominal power up of 25 mW at the level of the sample (see Appendix A). The diameter of the laser spot was set to 20 µm in focal plane.
**Figure 4.31.**A shows confocal micrographs of the emulsion at 25 °C before and after laser irradiation of an individual droplet (indicated by a white arrow) for 30 s with an irradiance of 31.8 µW/µm². The fluorescence signal detected in the micrographs was only due to NeutravNPs. Before irradiation, they were uniformly distributed in all the droplets of the emulsion, as expected at room temperature according to the results shown in section 4.3.1.

<figure><img src="image_112.png" /></figure>

**Figure 4.31.** Light-induced accumulation of NeutravNPs in the PLL-g-PNIPAM shell via thermoplasmonic excitation of AuNPs. A) Confocal micrographs of an emulsion containing 1 wt% PLL-g-PNIPAM, 0.01 wt% of NeutravNPs and 0.05 wt% of AuNPs in the aqueous phase and 0.05 wt% of Krytox in the oil phase. The emulsion was imaged at 25 °C before and after 532-nm light irradiation of an individual droplet (indicated with a white arrow) for 30 s at 31.8 µW/µm², and then uniformly heated at 45 °C (scale bars = 30 µm). B) Fluorescence radial profiles of the droplet indicated with a white arrow. C) Core fluorescence level of the droplet indicated with a white arrow expressed as a fraction of the initial value at 25 °C (error bars = standard deviation of the mean).

After laser irradiation, a bright corona was observed at the periphery of the targeted droplet, in addition to a decrease of the fluorescence level in the core (Figure 4.31.A, middle image). This pattern, highlighted in the fluorescence radial profile in Figure 4.31.B, is characteristic of the thermal-induced accumulation of NeutravNPs in the PLL-g-PNIPAM shell which has been described in section 4.3.1. In comparison, a uniform volume distribution of the NeutravNPs was still observed in the other droplets. This result
demonstrates that the interfacial capture of NPs in the PLL-g-PNIPAM shell can be triggered with light in
individual droplets. After the irradiation experiment, the whole emulsion was thermalized at 45 °C, i.e.
above the LCST of PLL-g-PNIPAM. At this temperature, the thermo-induced accumulation of NeutravNPs
at the interface was observed in all the droplets, as expected according to the results of section 4.3.1. In the
droplet previously targeted with the laser, this step of macroscopic heating induced a further decrease of
the core fluorescence level which almost fell to 0 (see the radial profile in Figure 4.31.B). This result
suggests that the migration of NeutravNPs towards the interface was only partially achieved upon light
irradiation and was less efficient than a macroscopic heating of the system. This result is quantitatively
confirmed by the comparison of the fluorescence level in the droplet core after irradiation at 25 °C and after
heating at 45 °C. Figure 4.31.C shows that 60 % of the fluorescence has disappeared after irradiation,
compared to 95 % after heating.

This result suggests that the temperature has not reached 45 °C in the droplet upon irradiation of
the AuNPs, at least non uniformly or not for long enough. However, considering the size of the droplet
(34 µm diameter) and the irradiance used during the experiment (31.8 µW/µm²), the irradiation should
generate at least 20 °C in the core of the droplet according to the GSPM results (since 20 °C were reached
with 20 µW/µm² in a 30-µm droplet, see Figure 4.30). The lower efficiency observed in the confocal
experiments was probably due to the non-uniform irradiation of the droplet, as the diameter of the laser spot
(20 µm) was smaller than the droplet diameter (34 µm). Accordingly, the temperature reached in the sample
was not uniform, and likely lower at the periphery of the droplet, which could explain the partial migration
of the NeutravNPs to the interface. This issue of non-uniform irradiation was not experienced in the GSPM
experiments since the diameter of the laser spot in the focal plane was of 50 µm, i.e. larger than the droplet
size.

The same experiment was performed with higher irradiances on droplet of similar sizes and on
smaller droplets, but the decrease of the fluorescence level in the core was always around 60 %. We thus
demonstrated that the segregation of NPs in the polymer shell can be induced by light in presence of AuNPs,
but some additional experiments should be made to precisely assess the efficiency of this process. In
particular, the optical set-up should be optimized to enable the plasmonic excitation and the confocal
observation on the same device. This may enable to observe the evolution of the NPs distribution
concomitantly to the laser excitation, especially to determine the influence of the irradiation time on the
process.
## 4.5 Conclusion

In this Chapter we demonstrated that our system based on interfacial complexation of PLL derivatives enables the straightforward formation of “composite” shells associating polymer and NPs. We evidenced the propensity of PLL to adsorb at the surface of NPs of different nature such as gold NPs, or carboxylate- or protein-modified polystyrene NPs. In particular, we showed that PLL-g-PNIPAM can coat the NPs and induce their aggregation upon hydrophilic-to-hydrophobic transition of the PNIPAM strands. We transposed this phenomenon in emulsions, in conditions of interfacial complexation of PLL-g-PNIPAM. We demonstrated that NPs, that were initially dispersed in the core of the droplets at ambient temperature, spontaneously accumulated at the surface of the droplets upon heating above the cloud point of PLL-g-PNIPAM. We attributed this process to a thermo-induced aggregation of the NPs and their progressive capture in the polymer shell, driven by hydrophobic interactions between PNIPAM chains localized both at the surface of the NPs and at the water/oil interface. Interestingly, the NPs remained segregated in the polymer shell when the emulsion was cooled back to ambient temperature. We thus evidenced a straightforward way to incorporate NPs in polymer shells without resorting to chemical reactions or capsule post-processing.

In the context of “composite” capsules, gold NPs (AuNPs) are particularly attractive due to their capability to convert light into heat at the microscale via thermoplasmonic excitation. Accordingly, we demonstrated that, in presence of AuNPs, the thermal transition of PLL-g-PNIPAM can be achieved upon light irradiation. We applied this technique to successfully trigger the thermo-induced accumulation of NPs in the polymer shell upon light irradiation for a few seconds in individual droplets, showing the potential of the technique to achieve a better spatio-temporal control of the capsule stimulation.

We also developed a protocol based on quantitative phase imaging to measure the temperature that can be reached in AuNPs-loaded droplets upon thermoplasmonic excitation. This protocol relies on the measurement of optical path difference induced by local temperature gradients generated upon AuNPs irradiation. Thanks to a calibration obtained by simulation, the optical path difference can be converted in a local measurement of the temperature rise. This technique revealed that the temperature can be increased by more than 20 °C in a few ms in sufficiently large droplets (typically 30 µm), which is promising for on-demand delivery upon photothermal stimulation.
Chapter 5

UCST-type polymer capsules
formed by interfacial
complexation
## 5.1 Introduction

In Chapter 3, we evidenced the potential of PLL comb-like derivatives to form polymer shells by interfacial complexation. We made an exhaustive study on a LCST derivative, namely PLL-g-PNIPAM, to characterize physico-chemical changes of the shell upon temperature variations. In Chapter 4, we demonstrated that the thermal response of PLL-g-PNIPAM enables the capture of nanoparticles (NPs) in the polymer shell when it turns hydrophobic, i.e. at high temperature.

In this chapter, we propose to extend our approach with the study of a UCST comb-like derivative of PLL, namely PLL-g-PAAMAN, obtained by grafting UCST poly(acrylamide-co-acrylonitrile) (PAAMAN) chains (presented in Chapter 2, section 2.3.2) on PLL. We evidenced the formation of PLL-g-PAAMAN shells by interfacial complexation, switching from a hydrophobic state at ambient temperature to a hydrophilic state upon heating. This work constitutes, as far as we know, the first demonstration of the formation of aqueous-core polymer microcapsules exhibiting a UCST behavior. This system is particularly promising for delivery applications since it implies the solubilization of the capsule shell upon heating, and consequently the release of the encapsulated content.

We demonstrated that NPs can be incorporated in the PLL-g-PAAMAN shell during the preparation of the emulsion, resulting in “composite” capsules at ambient temperature. Upon heating, the NPs were redispersed in the droplet core, resulting from the solubilization and partial redispersion of the PLL-g-PAAMAN shell. In addition, we demonstrated that the phase transition of the polymer shell can be induced upon light irradiation in presence of AuNPs, paving the way to photothermal delivery from UCST polymer capsules.
## 5.2 Synthesis of PLL-g-PAAMAN polymer and formation of PLL-g-PAAMAN shells by interfacial complexation

### 5.2.1 PAAMAN polymers

The PAAMAN polymers (poly(acrylamide-co-acrylonitrile)) used in this work were synthesized in the group of Prof. Yue Zhao (Sherbrooke University, Canada). Only two polymers, named P1 and P2, will be presented as potential candidates for the formation of UCST capsules (see structures in **Figure 5.1**). P1 and P2 were synthesized by RAFT from two different RAFT agents, with final acrylamide/acrylonitrile ratios of 0.72/0.28 and 0.66/0.34 respectively (see **Appendix A** for experimental details). The molecular masses were 12 kDa and 15 kDa respectively.

The cloud point of each polymer was evaluated by turbidity measurements upon heating on solutions at 1 wt% in water (see **Figure 5.1**). The cloud point was ~39 °C for P1 and ~45 °C for P2 (temperature corresponding to 50% of transmittance). At this stage, P1 seems more appropriate for delivery applications since its cloud point is lower and easier to reach in biological contexts.

<figure><img src="image_113.png" /><figcaption>**Figure 5.1.** Structure of the two PAAMAN polymers considered as candidates for the formation of polymer capsules and the corresponding turbidimetry heating curves (at 1 wt% in pure water).</figcaption></figure>
The physical behavior of the two PAAMAN polymers in solution was further analyzed by bright field confocal microscopy (see **Figure 5.2**). Solutions at 10 wt% in pure water were heated at 48 °C (above their cloud point) and then cooled back to 18 °C to trigger the collapse transition of the polymer.

<figure><img src="image_114.png" /><figcaption>**Figure 5.2.** Confocal micrographs of solutions of the two PAAMAN polymers at 10 wt% in pure water upon cooling from 48 °C to 18 °C (scale bars = 50 µm).</figcaption></figure>

Upon cooling, each polymer undergoes a liquid-liquid phase separation characteristic of a UCST-type behavior, but the dynamics of their polymer-enriched phase was drastically different. P1 formed liquid droplets that rapidly coalesced and relaxed, whereas P2 formed gel-like irregular aggregates that partially and slowly merged. This difference of behavior can be due to several factors: the difference in chain composition (monomer ratio, end-groups, chain length), the potential presence of residual traces of solvent synthesis, or the degree of hydration of the initial polymer powder.

The gel-like behavior of P2 seems more appropriate for the formation of robust/rigid polymer shells at ambient temperature and was selected for the rest of the experiments (despite its higher cloud point). Accordingly, in the following the term of PAAMAN will exclusively refer to the polymer P2.
### 5.2.2 Synthesis of PLL-g-PAAMAN copolymer

As for non-ionic PNIPAM in Chapter 3, non-ionic PAAMAN chains were grafted onto PLL to drive its interfacial complexation at the surface of emulsions droplets. However, the PAAMAN chains were terminated by carboxylic acid groups at both ends and a direct coupling on PLL (via amide bond formation) would thus promote the formation of cross-links, and not the formation of comb-like copolymers. As an alternative, the central trithiocarbonate unit of PAAMAN chains was cleaved by aminolysis to yield two shorter chains terminated by a thiol group at one end.²⁹⁸ Then the grafting of thiol-terminated PAAMAN strands onto PLL was mediated by an intermediary short PEG linker (3 EG units) terminated with an NHS-ester group at one end (for the anchoring on PLL) and a maleimide group at the other (for thiol-ene click reaction). The synthesis of PLL-g-PAAMAN was achieved in one-pot according to a two-step procedure described in **Appendix A** and schematized in **Figure 5.3**.

<figure><img src="image_115.png" /><figcaption>**Figure 5.3.** Synthesis procedure for the grafting of PAAMAN strands (x=0.66, y=0.34, M=15 kDa) on a PLL backbone using an intermediary poly(ethylene glycol) difunctional linker. MEA = monoethanolamine, TCEP = tris(2-carboxyethyl)phosphine.</figcaption></figure>

In this synthesis, a slight excess of PEG linkers was used compared to aminolyzed PAAMAN moieties to achieve the complete incorporation of the PAAMAN chains introduced in the reactive bath (typically 7 mol% of the side amino groups of PLL were functionalized with the linker, for 2-4 mol% of thiol-terminated PAAMAN strands produced *in situ*). Fluorescent derivatives of PLL-g-PAAMAN were obtained by addition of a reactive fluorophore that can be grafted onto PLL (rhodamine-NHS or coumarin-NHS) during the first step of the synthesis.
The composition of the PLL-g-PAAMAN copolymer van be tuned by adjusting the PLL/PAAMAN weight ratio during the synthesis. In the following, the results will refer to two copolymers synthesized with different PLL/PAAMAN weight ratio :

* PLL<sub>0.53</sub>-g-PAAMAN<sub>0.47</sub> with a PLL/PAAMAN weight ratio of 53/47 wt%, obtained by grafting 6 PAAMAN strands on a 52 kDa PLL.

* PLL<sub>0.42</sub>-g-PAAMAN<sub>0.58</sub> with a lower PLL/PAAMAN ratio of 42/58 wt%, obtained by grafting 4 PAAMAN strands grafted on a 22 kDa PLL.

The thermoresponsiveness of PLL-g-PAAMAN in solution will be illustrated in the case of PLL<sub>0.53</sub>-g-PAAMAN<sub>0.47</sub>. The cloud point of this copolymer (at 1 wt% in pure water) was measured at ~ 49 °C, i.e. 4 °C above the cloud point of non-grafted PAAMAN (dashed lines in **Figure 5.4**), indicating that the grafting on PLL slightly shifted the UCST of PAAMAN chains towards higher temperature. This behavior can be attributed to the participation of PLL in the inter-PAAMAN hydrogen-bond network, hence favoring the aggregative state of the polymer on a broader temperature range.

<figure><img src="image_116.png" /><figcaption>**Figure 5.4.** Adjustment of the cloud point of PLL-g-PAAMAN solutions with thiocyanate salt. A) Turbidimetry heating curves of PAAMAN and PLL<sub>0.53</sub>-g-PAAMAN<sub>0.47</sub> solutions at 1 wt% in either pure water or 20 mM KSCN solution. B) Cloud points extracted from the turbidimetry curves at 50 % transmittance (the bars indicate the initial and final temperatures of the UCST transition).</figcaption></figure>

For practical reasons related to microscopy observations, we needed to decrease the cloud point of PLL-g-PAAMAN solutions, typically below 45 °C. For that, we supplemented the polymer solutions with 20 mM of the chaotropic salt KSCN.<sup>186</sup> In these conditions, a “salting out” effect was observed for non-grafted PAAMAN for which the cloud point was shifted from 45 °C to 40 °C (see **Figure 5.4**). Interestingly, this effect was more important for PLL-g-PAAMAN with a decrease of the cloud point from 49 °C to
37 °C and a significant broadening of the temperature range of the UCST transition (now ranging from 30 °C to 48 °C). Thanks to this cloud point adjustment, the thermoresponsive behavior of PLL-g-PAAMAN could be observed in a temperature window which was (i) more accessible in the confocal microscopy set-up and (ii) more appropriate for biological applications.

### 5.2.3 Formation of PLL-g-PAAMAN capsules

The formation of PLL-g-PAAMAN shells was achieved by interfacial complexation in water-in-oil emulsions as presented in Chapter 1, except that the emulsification was not performed at room temperature but well above the cloud point of PLL-g-PAAMAN (typically at 60 °C). Briefly, an aqueous phase containing PLL-g-PAAMAN (typically in the wt% range) and 20 mM KSCN was emulsified at 60 °C in a continuous oil phase made of FC-70 oil containing Krytox surfactant (typically at 0.05-0.1 wt%) and then cooled down to 18 °C (well below the cloud point) for microscopy imaging.

The distribution of the polymer in the emulsion droplets at 18 °C was assessed by confocal microscopy using rhodamine-labeled PLL-g-PAAMAN. **Figure 5.5.A** and **B** shows the results for $PLL_{0.53-g-PAAMAN_{0.47}}$ and $PLL_{0.42-g-PAAMAN_{0.58}}$, respectively.

For $PLL_{0.53-g-PAAMAN_{0.47}}$, the bright fluorescent corona observed at the periphery of the droplets was indicative of the formation of a polymer shell (see **Figure 5.5.A**). For $PLL_{0.42-g-PAAMAN_{0.58}}$, micrometric globular aggregates punctuating both the core and the periphery of the droplets were observed (see **Figure 5.5.B**). These results suggest that the ability of PLL-g-PAAMAN to adsorb at the water/oil interface is highly dependent on the composition of the polymer, and seems to require a relatively high PLL/PAAMAN weight fraction (or at least a sufficiently long PLL chain length). The predominance of PAAMAN in the polymer (in terms of weight fraction) may favor phase separation under the form of globules below the cloud point (as do non-grafted PAAMAN chains), hence promoting clustering in solution instead of complexation at the water/oil interface.
A) PLL/PAAMAN weight ratio = 53/47 wt%

<figure><img src="image_117.png" /></figure>

B) PLL/PAAMAN weight ratio = 42/58 wt%

<figure><img src="image_118.png" /></figure>

**Figure 5.5.** Influence of the composition of PLL-g-PAAMAN on the ability to form polymer shells by interfacial complexation. Water-in-oil emulsions were prepared at 60 °C with 1 wt% of rhodamine-labeled PLL-g-PAAMAN and 20 mM of KSCN in the aqueous phase and 0.05 %wt of Krytox in the oil phase. A) Polymer shells were effectively formed at the periphery of the water droplets in the case of PLL<sub>0.53</sub>-g-PAAMAN<sub>0.47</sub>, after cooling the emulsion down to 18 °C. B) In the case of PLL<sub>0.42</sub>-g-PAAMAN<sub>0.58</sub>, the flat deposition of the polymer at the interface was supplanted by a globular association of the polymer within the droplet core. Scale bar in the confocal micrographs = 30 µm.

The impact of chain length and grafting ratio on the behavior of PLL-g-PAAMAN in emulsion would
deserve further studies to rationalize the synthesis of the copolymer and optimize the formation of uniform
capsule shells. In this study however, PLL<sub>0.53</sub>-g-PAAMAN<sub>0.47</sub> has been used for the proof of concept. In the
following, this polymer will be simply referred to as PLL-g-PAAMAN for the sake of readability.
## 5.3 Thermoresponsive behavior of PLL-g-PAAMAN shells

### 5.3.1 UCST behavior of PLL-g-PAAMAN shells assessed with ANS fluorescence

As for PLL-g-PNIPAM in Chapter 3, the fluorescence of ANS was used as an extrinsic polarity/hydrophobicity probe. **Figure 5.6** shows confocal micrographs of a representative droplet containing PLL-g-PAAMAN (non-fluorescent) and ANS below and above the cloud point. High fluorescence intensity at the surface of the droplets at 18 °C validates the hydrophobic nature of the polymer shell below the cloud point. In contrast, a low and homogeneous fluorescence was observed in the droplet at 48 °C, suggesting a highly hydrophilic environment both in the core and near the interface. When the temperature was brought back below the cloud point, the high fluorescence level was recovered at the surface of the droplet. These results confirm the existence of a reversible UCST-type hydrophobic-to-hydrophilic transition of the PLL-g-PAAMAN upon heating.

<figure><img src="image_119.png" /><figcaption>**Figure 5.6.** Visualization of the UCST-type transition of PLL-g-PAAMAN shell using ANS as extrinsic fluorescent probe. A) Confocal micrographs of a representative droplet containing 1 wt% of PLL-g-PAAMAN and 100 µM of ANS in 20 mM KSCN, dispersed in oil containing 0.05 wt% of Krytox. The emulsion was thermalized at 18 °C, 48 °C and 18 °C successively. B) Fluorescence radial profiles of the droplet.</figcaption></figure>
### 5.3.2 Measurement of polymer surface excess with intrinsic fluorescence measurements

The thermoresponsiveness of the shell was quantitatively assessed *in situ* by measuring the surface excess while increasing the temperature above the cloud point (from 18 °C to 48 °C). Upon heating, a decrease of the fluorescence level was observed in the interfacial layer concomitantly with an increase of the fluorescence level in the core of the droplets, as illustrated by the fluorescence radial profiles obtained by confocal imaging (see **Figure 5.7.A** and B). This qualitative evolution was attributed to the hydrophobic-to-hydrophilic transition of the PAAMAN strands that may induce a partial solubilization of the interfacial polymer layer and redispersion in the core, as depicted in Figure 5.7.A.

<figure><img src="image_120.png" /><figcaption>**Figure 5.7.** Quantification of the polymer surface excess by intrinsic fluorescence measurements. A) Confocal micrographs of a representative water droplet containing 1 wt% of rhodamine-labeled PLL-g-PAAMAN (in 20 mM KSCN) dispersed in FC-70 oil containing 0.05 wt% of Krytox at 18 °C and 48 °C (scale bars = 20 µm). B) Fluorescence radial profiles of the droplets. C) Evolution of the surface excess of PLL-g-PAAMAN as a function of the droplet diameter at 18 °C and 48 °C, calculated from fluorescence measurements. D) Evolution of the surface excess of PLL-g-PAAMAN as a function of the residual polymer concentration in the droplet core at 18 °C and 48 °C (colored domains are guides for the eye). Box plots give the min/Q1/median/Q3/max values of the distribution of the residual core concentration.</figcaption></figure>
The quantitative measurement of the polymer surface excess previously introduced in Chapter 3 (section 3.4.2) was implemented here in the case of PLL-g-PAAMAN. The evolution of the surface excess as a function of the droplet diameter or the residual core concentration at 18 °C and 48 °C are shown in Figure 5.7.C and D, respectively. These data confirm that the rise of temperature induces (i) a decrease of the surface excess for any droplet size (of 45 % in average) and (ii) an increase of the polymer concentration in the core of the droplets (of 130 % in average). These results are consistent with a partial solubilization of the hydrophilic PLL-g-PAAMAN shell upon heating.

### 5.3.3 Evaluation of the fluidity of the polymer shell by fluorescence recovery after photobleaching

Changes in fluidity/dynamics of the polymer shell with temperature was assessed by fluorescence recovery after photobleaching (FRAP). FRAP experiments were performed on coumarin-labeled PLL-g-PAAMAN shells (prepared in the same concentration and temperature conditions as described above) at 18 °C and 48 °C (see Figure 5.8.A). The fraction of mobile chains within the polymer shell, denoted $M_f$, was deduced from the following equation:

$$M_f = \frac{I_\infty - I_0}{I_i - I_0}$$

where $I_i$ was the fluorescence intensity before bleaching (fixed at 1 by normalization), $I_0$ the intensity just after bleaching and $I_\infty$ the intensity after 2 min (assuming that a steady state was approximately reached at this time).

Representative curves are shown in Figure 5.8.B. At 18 °C, almost no fluorescence recovery was observed in the photobleached area, indicating that mobile chain fraction was below 10 %. In these conditions, the diffusion of polymer chains within the shell below the cloud point was likely frozen. In contrast at 48 °C, a partial recovery was observed after 2 min, suggesting a diffusion of the chains in the polymer shell (an estimate of the mobile fraction being here slightly above 50%). These results are consistent with a thermo-induced transition of the polymer shell from a solid-like (or frozen) state to a fluid-like state upon heating.
<figure><img src="image_121.png" /><figcaption>**Figure 5.8.** Fluorescence recovery after photobleaching (FRAP) experiments on thermoresponsive PLL-g-PAAMAN shells. A) Confocal micrographs of representative droplets containing 1 wt% of coumarin-labeled PLL-g-PAAMAN (in 20 mM KSCN) during FRAP experiments at 18 °C and 48 °C (scale bars = 5 µm). B) Left: FRAP curves from the top experiments. Right: polymer mobile fraction $M_f = \frac{I_\infty-I_0}{I_i-I_0}$ calculated from the fluorescence intensity levels measured along the FRAP curves at 18 °C and 48 °C (error bars = standard deviation on 5 and 8 droplets respectively).</figcaption></figure>

As a comparison, FRAP experiments were conducted in the same conditions on a non-thermoresponsive coumarin-labeled PLL-g-PEG (see **Figure 5.9**). Compared to PAAMAN strands, PEG strands do not develop inter-chain hydrogen bonds that may affect the dynamics of the chains within the shell.

At 18 °C the mobile fraction of PLL-g-PEG reaches ~45 %, which is far above the mobile fraction obtained for PLL-g-PAAMAN (< 10 %). This difference in dynamics at low temperature confirms the frozen nature of the PLL-g-PAAMAN shell, likely associated to a robust network of interchain hydrogen bonds. At 48 °C, the mobile fraction of PLL-g-PEG increased up to ~70 %. This fluidification was probably due to the temperature-dependent nature of the diffusion process (the diffusion coefficient increases with temperature). This evolution was also observed in the case of PLL-g-PAAMAN but at a lower level (up to ~50 % of mobile chains), suggesting that the PLL-g-PAAMAN shell becomes more fluid above the cloud
point, but remains less fluid than a “model” polymer (PLL-g-PEG) exhibiting no specific polymer
interactions.

<figure><img src="image_122.png" /><figcaption>**Figure 5.9.** FRAP experiments on non-thermoresponsive PLL-g-PEG shells. A) Confocal micrographs of representative droplets containing 1 wt% of coumarin-labeled PLL-g-PEG (in 20 mM KSCN) during FRAP experiments at 18 °C and 48 °C (scale bars = 5 µm). B) *Left:* FRAP curves from the top experiments. *Right:* polymer mobile fraction $M_f = \frac{I_\infty - I_0}{I_i - I_0}$ calculated from the fluorescence intensity levels measured in the FRAP curves at 18 °C and 48 °C (error bars = standard deviation on 6 droplets).</figcaption></figure>
## 5.4 Towards the formation of UCST composite capsules

### 5.4.1 Incorporation of NeutravNPs in PLL-g-PAAMAN shells

We assessed the capacity of PLL-g-PAAMAN to drive the co-precipitation of nanoparticles in the polymer layer, and accordingly the straightforward formation of polymer-particles mixed shells. We used NeutravNPs as a proof of concept, as we did for PLL-g-PNIPAM shells in Chapter 4.

NeutravNPs were added in the aqueous phase containing PLL-g-PAAMAN prior to emulsification at 60 °C. Cooling down the emulsion to ambient temperature led to the segregation of the NeutravNPs in the PLL-g-PAAMAN shell, ascribed to a hydrophobic co-aggregation of polymers chains and NPs at the interface (see **Figure 5.10.A**). Heating the system up to 48 °C enabled to recover soluble NeutravNPs uniformly distributed in the droplet cores (see Figure 5.10.B). Interestingly, when the sample was cooled back below the cloud point, NeutravNPs were mostly recaptured in the shell. This reversible process associated to the reversible transition of PLL-g-PAAMAN is schematized in Figure 5.10.A, and highlighted in the radial profiles of a representative droplet shown in Figure 5.10.B. These results show that NPs can be spontaneously embedded in the polymer shell during its formation. This approach is promising for the straightforward design of UCST-type composite capsules.
<figure><img src="image_123.png" /><figcaption>**Figure 5.10.** Reversibility of the capture of NeutrAvidin-coated fluorescent nanoparticles (NeutravNPs) upon UCST transition of PLL-g-PAAMAN. A) Top: Confocal micrographs of droplets containing 1 %wt of PLL-g-PAAMAN and 0.025 wt% of NeutravNPs at 18 °C, 48 °C and 18 °C successively (scale bars = 30 µm). Bottom: Schematics representing the state of PLL-g-PAAMAN and the distribution of the NeutravNPs in the droplets as observed in the top micrographs (Krytox molecules are not represented for the sake of readability and the scale was not respected). B) Fluorescence radial profiles of the droplets indicated with a white arrow in the top micrographs (the droplet at step 3 is different from the one at steps 1 and 2).</figcaption></figure>

### 5.4.2 Encapsulation of AuNPs and photothermal stimulation

The possibility to incorporate NPs in PLL-g-PAAMAN adlayers was extended to the formation of polymer shells doped with AuNPs to impart light-responsiveness to the capsules. As AuNPs were not fluorescent, their presence in the polymer shells at low temperature was assessed by comparing SEM pictures of droplets subjected to (i) homogenization at 60 °C with or without AuNPs present in the aqueous phase, (ii) cooling down to room temperature, and (iii) drying at ambient air (see **Figure 5.11**). In absence of AuNPs, the surface of PLL-g-PAAMAN capsules was smooth, except the presence of cracks and folds that probably appeared during the drying step. In presence of AuNPs, white dots and aggregates were clearly visible at the surface of the PLL-g-PAAMAN capsules, and are ascribed to the presence of AuNPs although it cannot
be ruled out that their distribution was markedly affected by the drying process. However, this observation
suggests that AuNPs can be (at least partially) embedded in the PLL-g-PAAMAN shell upon formation
below the cloud point.

<figure><img src="image_124.png" /><figcaption>**Figure 5.11.** SEM micrographs of dried PLL-g-PAAMAN capsules prepared from aqueous phases with or without PVP-coated AuNPs (scale bars = 10 µm). The original emulsions contained 1 wt% of PLL-g-PAAMAN with/without 0.05 wt% of AuNPs and 20 mM KSCN in the aqueous phase and 0.1 wt% of Krytox in the oil phase.</figcaption></figure>

We assessed the possibility to trigger the dissolution of the PLL-g-PAAMAN shell via light irradiation of encapsulated AuNPs. An emulsion was prepared with 2 wt% of PLL-g-PAAMAN and 0.05 wt% of AuNPs in the aqueous phase (20 mM KSCN), and 0.1 wt% of Krytox in the oil phase. The emulsion was observed at 25 °C under phase contrast microscopy (see **Figure 5.12**). At this temperature, no polymer globules were visible in the droplet cores, and the droplet edges were slightly irregular, suggesting the presence of PLL-g-PAAMAN shells. When a 532-nm laser beam was focused on a given droplet, we should expect an increase of temperature via thermoplasmonic effect of AuNPs, that may eventually trigger the solubilization transition of the PLL-g-PAAMAN shell. This light-induced process was not clearly visible in phase contrast microscopy, except through a slight shrinkage of the droplet. When the laser was turned off however, polymer granules instantaneously appeared in the droplet core. Then the granules progressively migrated from the core to the periphery of the droplet, and eventually merged at the water/oil interface to retrieve the initial shell configuration. This suggests that (i) the temperature was raised above the cloud point upon light irradiation, (ii) polymer chains detached from the interface upon heating and redispersed in the core, and (iii) upon cooling they precipitated under the form of aggregates which eventually migrated towards the interface. The entire process is depicted below the micrographs in Figure 5.12.
<figure><img src="image_125.png" /><figcaption>**Figure 5.12.** Light-triggered dissolution of PLL-g-PAAMAN shell induced by thermoplasmonic effects of co-encapsulated AuNPs. The phase contrast micrographs represent a water droplet containing 2 wt% of PLL-g-PAAMAN and 0.05 wt% of AuNPs (in 20 mM KSCN) dispersed in oil phase containing 0.1 wt% of Krytox, and thermalized at 25 °C (scale bar = 10 µm). The droplet was irradiated with a 532-nm laser at 15 mW for a few seconds. After turning the laser off, polymer granules spontaneously formed in the droplet core and progressively migrated towards the interface in a few minutes.</figcaption></figure>

## 5.5 Encapsulation of a model protein

The compatibility of the PLL-g-PAAMAN shells with the presence of a protein encapsulated in the core of the droplets was tested on the model eGFP, as it was done in Chapter 3 (see section 3.6) for PLL-g-PNIPAM shells.

**Figure 5.13** shows confocal micrographs of an emulsion containing 1 wt% of rhodamine-labeled PLL-g-PAAMAN and 50 µM of eGFP in the aqueous phase, at 18 °C and 48 °C. At 18 °C, PLL-g-PAAMAN was not localized at the periphery of the droplet under the form of a flat shell but formed globules in the droplet core. These globules also contained a fraction of eGFP as revealed in the protein fluorescence channel. This observation suggests that, unfortunately, protein-polymer interactions drove the co-precipitation of eGFP with PLL-g-PAAMAN in the droplet core, and prevented the deposition of the polymer at the interface via interfacial complexation. At 48 °C, the polymer-protein-enriched globules were
dissolved upon hydrophobic-to-hydrophilic transition of the polymer. Consequently, eGFP was uniformly distributed within in the droplet core and PLL-g-PAAMAN formed irregular spots at the interface.

<figure><img src="image_126.png" /><figcaption>**Figure 5.13.** Encapsulation of eGFP in PLL-g-PAAMAN capsule precursors. Confocal micrographs of an emulsion containing 1 wt% of rhodamine-labeled PLL-g-PAAMAN (red channel) and 50 µM of eGFP (green channel) in the aqueous phase (20 mM KSCN) and 0.05 wt% of Krytox in the oil phase, at 18 °C and 48 °C (scale bars = 50 µm). The focal plane was set below the equatorial plane of the droplets to show the formation and sedimentation of PLL-g-PAAMAN globules at 18 °C.</figcaption></figure>

These results suggest that the presence of eGFP strongly interferes with the formation of a PLL-g-PAAAMN shell by interfacial complexation, contrary to the case of PLL-g-PNIPAM. With a lower protein concentration (10 µM), the adsorption of PLL-g-PAAMAN at the interface seemed to be partially preserved but under the form of granules containing the protein rather than under the form of an even shell surrounding core-dispersed protein (see **Figure 5.14**). Conversely, with a higher protein concentration (150 µM), most of the protein was dispersed in the droplet core, but the polymer essentially formed protein-enriched granules and did not adsorb at the interface (see Figure 5.14), as in the case of 50 µM protein. Further optimization should be done to guarantee the formation of a polymer shell and to limit non-specific polymer-protein interactions in presence of a protein of interest for delivery applications.
<figure><img src="image_127.png" /><figcaption>**Figure 5.14.** Encapsulation of eGFP in PLL-g-PAAMAN capsule precursors for two different protein concentrations. Confocal micrographs of emulsions containing 1 wt% of rhodamine-labeled PLL-g-PAAMAN (red channel) and either 10 or 150 µM of eGFP (green channel) at 18 °C.</figcaption></figure>

## 5.6 Conclusion

In this Chapter, we reported the formation by interfacial complexation in water-in-oil emulsion of aqueous-core PLL-g-PAAMAN microcapsules exhibiting a UCST behavior (see **Figure 5.15**). After emulsification above the cloud point of the polymer (typically 37 °C) and cooling down to room temperature, a hydrophobic, gel-like polymer shell was readily formed by complexation of PLL-g-PAAMAN chains with an oil-soluble surfactant at the surface of water droplets. We showed that this process enabled the straightforward incorporation of nanoparticles (previously dispersed in the aqueous phase) in the polymer shell upon temperature switch. The hydrophobic-to-hydrophilic transition of PAAMAN chains upon heating triggered a partial redispersion of the polymer shell and a drastic change in its fluidity, switching from a gel-like state to a liquid-like state. These thermoresponsive properties combined with the ability to readily encapsulate nanoparticles show the high potential of our approach for advanced delivery applications, such as photothermal delivery with gold nanoparticles, or magnetic hyperthermia (local heating induced by alternating magnetic fields) with paramagnetic nanoparticles.

However, the encapsulation of eGFP as a model protein has not been achieved yet since the presence of the protein interfered with the formation of PLL-g-PAAMAN shell. Polymer-protein
interactions seemed to favor the co-precipitation of eGFP with PLL-g-PAAMAN into globular aggregates
in the droplet core, at the expense of interfacial complexation of PLL-g-PAAMAN at the droplet surface.

<figure><img src="image_128.png" /><figcaption>**Figure 5.15.** Formation and functionalities of UCST-type polymer shells made of PLL-g-PAAMAN chains by interfacial complexation in water-in-oil emulsion.</figcaption></figure>
# Chapter 6

## Conclusion and outlooks
## 6.1 Conclusion

Many bio-applications require the encapsulation and the controlled release of hydrophilic payloads, such as localized drug delivery, tissue regeneration or exploration of cell signaling. For such goals, ideal encapsulation systems would be able to (i) preserve the integrity and bioactivity of the encapsulation content during all the steps of formulation, (ii) trigger the on-demand release of the payload upon application of an external stimulus, (iii) be versatile, i.e. appropriate for the encapsulation of a large range of hydrophilic payloads (including fragile biomacromolecules such as proteins or DNA). In this context, hollow capsules composed of an aqueous core surrounded by a polymer shell are promising encapsulation systems. An ideal polymer capsule could be described as the combination of a polymer shell that plays the dual role of protecting envelop during encapsulation/storage and stimuli-responsive gate for on-demand release of the payload, and a liquid core that enables to maintain mild aqueous conditions within the capsule. The objective of the present work was to elaborate a versatile approach for the design of stimuli-responsive aqueous-core polymer capsules that meet these specifications.

For that, aqueous-core capsules were formed by structuration of a polymer shell around water droplets via interfacial complexation in water-in-fluorinated oil emulsions. The complexation process involved two oppositely charged species that were initially solubilized in their own phase: a cationic comb-like copolymer in the water phase and an anionic surfactant called Krytox in the oil phase. Upon emulsification of the water phase into the oil phase, the anionic surfactant and the cationic polymer interacted at the surface of the water droplets through coulombic attraction. In appropriate conditions of formulation, these interactions resulted in the formation of a surfactant-polymer mixed shell with a thickness of typically 0.5-1 µm. Interfacial complexation has the advantage to be a spontaneous self-assembly process that does not require any *in situ* chemical reactions (potentially prejudicial for the stability or integrity of co-encapsulated payloads).

We demonstrated via interfacial tension measurements that the cationic poly(L-lysine) (PLL) is an interesting building block for the design of functional shells as it can be effectively adsorbed at the water/oil interface via complexation with Krytox. We thus used this strategy to generate thermoresponsive polymer shells by interfacial complexation of comb-like copolymers composed of a PLL backbone and thermoresponsive comb strands, namely either the LCST-type poly(N-isopropylacrylamide) (PNIPAM) which is hydrophilic and soluble in water at ambient temperature and becomes hydrophobic and insoluble upon heating, or the UCST-type poly(acrylamide-co-acrylonitrile) (PAAMAN) which exhibits the inverse behavior.
A detailed study has been performed on the PLL-g-PNIPAM comb-like copolymer to evidence the formation of thermoresponsive polymer shells by interfacial complexation. PLL-g-PNIPAM exhibits a LCST behavior similar to the initial PNIPAM thermoresponsive strands, i.e. characterized by the same cloud point of 34 °C. We showed that PLL-g-PNIPAM shells could be readily formed at the surface of water droplets by interfacial complexation, provided a balanced concentration ratio between polymer and surfactant in the emulsion. Furthermore, we confirmed by fluorescence measurements that hydrophilic-to-hydrophobic transition of the polymer shell occurs in emulsion when heated above 34 °C. Interestingly, this transition was able to drive the accumulation of materials within the polymer shell via hydrophobic interactions. This was illustrated by an increase of the surface excess of PLL-g-PNIPAM upon heating, i.e. an enrichment of the polymer shell. More interestingly, we observed the co-precipitation at the interface of non-grafted PNIPAM chains that were initially soluble in the droplet core at ambient temperature. We also demonstrated that nanoparticles (NPs) initially dispersed in the droplet core at ambient temperature can be irreversibly sequestered in the polymer shell upon heating. This process was ascribed to the propensity of PLL to adsorb at the surface of the NPs, leading to the capture of the NPs in the hydrophobic polymer shell at high temperature. Through this process, we evidenced a straightforward mean to access polymer-particle "composite" shells with no other treatment than a gentle heating of the templating emulsion.

In a parallel study, we demonstrated that PLL-g-PAAMAN can also form polymer shells by interfacial complexation, provided an appropriate chemical structure of the copolymer. We evidenced a UCST behavior of PLL-g-PAAMAN shells, characterized by a reversible transition from a hydrophobic gel-like state at ambient temperature to a hydrophilic liquid-like state at high temperature (typically 45 °C). This thermal transition triggers a partial redispersion of the shell upon heating. In addition, we demonstrated that NPs can be embedded in the PLL-g-PAAMAN shells during its formation, and can then be redispersed in the droplet core upon heating. These capsules are promising for the design of composite capsules which are robust at low temperature and partially soluble at high temperature, i.e. appropriate for a controlled release upon heating.

The thermoresponsiveness of PLL-g-PNIPAM and PLL-g-PAAMAN was initially induced by a macroscopic heating of the sample. To achieve a better spatio-temporal control, we proposed to use light as an external trigger. Since the PLL derivatives were not intrinsically light-sensitive but only thermosensitive, we supplemented the aqueous solutions with gold nanoparticles (AuNPs) which played the role of light-to-heat converters via thermoplasmonic effects. We qualitatively showed that, in presence of AuNPs in the droplet core, the thermal transition of PLL-g-PNIPAM of PLL-g-PAAMAN could be triggered in individual droplets upon green light irradiation (at 532 nm). In the case of PLL-g-PNIPAM, light irradiation was able to both trigger the aggregation of PLL-g-PNIPAM chains and induce the segregation of NPs at the interface similarly to what was observed upon macroscopic heating of the sample.
In the case of PLL-g-PAAMAN, polymer granules appeared in the core of irradiated droplets just after stopping the irradiation, revealing that a fraction of the polymer shell had been solubilized and detached from the interface upon light irradiation. In the experimental conditions tested in this work, the light excitation at the droplet scale was slightly less efficient than a macroscopic heating to induce physical changes in the polymer shells. However, light triggering benefits from a high degree of tunability since the irradiation power, the irradiation time, and the amount of AuNPs could be adjusted to optimize the response of the system.

In addition to the qualitative results obtained upon light irradiation, we developed a protocol to measure *in situ* the temperature that can be reached in AuNPs-loaded droplets upon light excitation. This protocol was based on quantitative phase imaging of droplets upon irradiation with a technique called grating-shadow phase microscopy. This technique enabled to map the optical path difference induced by local heating within an irradiated droplet and in its vicinity. The pattern of optical path difference was correlated to the increase of temperature reached in the droplet core thanks to simulations. This approach showed that the highest temperature rises were observed in the largest droplets. Typically, a temperature rise of 20 °C could be reached in 30-µm droplets with an irradiance of 20 µW/µm² (easily accessible with commercial laser sources).

The possibility to encapsulate fragile biomacromolecules such as proteins was an initial motivation of this work. The formation of PLL-based polymer capsules by interfacial complexation did not require strong physical or chemical treatments that could damage the encapsulated payloads (no sonication, no chemical reactions), and enabled to keep a mild aqueous environment in the droplets which are capsule precursors. We used eGFP as a model protein to assess the compatibility of our formulation protocol with the presence of proteins. eGFP was successfully encapsulated in PLL-g-PNIPAM capsule precursors as evidenced by the presence of the protein in the droplet core and the effective formation of a polymer shell, with no signs of polymer-protein interactions. In the case of PLL-g-PAAMAN, the presence of the protein hindered the formation of a polymer shell by interfacial complexation. Instead, globules containing both the polymer and the protein were formed in the droplet core, suggesting that strong polymer-protein interactions interfered with the deposition of the polymer at the surface of the droplets. These results suggest that the nature of the polymer used to form the capsule shell is crucial regarding the encapsulation of a protein of interest.
## 6.2 Outlooks

### 6.2.1 Possible improvements

PLL-g-PNIPAM was used to demonstrate the feasibility of our approach, namely the formation of functional polymer shells by interfacial complexation of PLL derivatives. In terms of applications however, the LCST behavior of PLL-g-PNIPAM shells is not ideal for on-demand release since the shell becomes hydrophobic upon heating. For applications, the efforts should focus on UCST-type shells, such as PLL-g-PAAMAN, that are gel-like at ambient temperature and undergo solubilization upon heating.

Regarding the formation of UCST-type polymer capsules, a first way of improvement would consist in the optimization of the composition of the PLL derivatives. As demonstrated in the case of PLL-g-PAAMAN, the size of the PLL backbone, as well as the grafting ratio of functional strands, have a drastic effect on the propensity of PLL to adsorb at curved water/oil interfaces upon interfacial complexation. A rationalization of the influence of these structural parameters on the amount of polymer adsorbed at the interface should enable to maximize the uniformity and the thickness of the polymer shells. The intrinsic thermal properties of UCST-type polymers are also key parameters that dictate the behavior of the shell. As demonstrated in the beginning of Chapter 5, the cloud point of PAAMAN and the dynamics of the polymer-enriched phase upon phase separation can be tuned by changing the polymer composition. The synthesis protocol of PAAMAN chains should be optimized to get (i) a cloud point closer to practical temperatures (typically 35 °C) without need of additional salts, and (ii) a gel-like dynamics in the polymer-enriched phase to improve the mechanical properties of the resulting shell.

Regarding the experiments of light triggering, two ways of improvement can be mentioned:

* In this work, the plasmonic excitation and the confocal imaging of emulsion droplets loaded with AuNPs were performed in separated set-ups. The laser sources of the confocal microscope were not sufficiently powerful to trigger significant thermoplasmonic effects, and the apparatus was not appropriate for the addition of an external laser source. Therefore, the plasmonic excitation was performed on a commercial microscope which did not work in confocal mode, but which could be supplemented with an external laser source. This set up suffered from two limitations: 1) the irradiation laser beam was passing through the internal optical system of the microscope, resulting in losses of power and optical deformations of the laser spot due to multiple reflections, 2) the impact of thermoplasmonic effects on the properties of the polymer shells were not easy to visualize since it was
restricted to bright field imaging (the fluorescence imaging was forbidden by the presence of the
irradiation laser). A home-made optical set-up combining confocal detection and orthogonal laser
irradiation should enable to properly control the shape and size of the laser spot in the focal plane, and
to simultaneously generate thermoplasmonic effects and follow the distribution of polymer and NPs
species within the droplets.

o Regarding the thermometry protocol developed with grating-shadow phase microscopy, further studies are required to precisely quantify the influence of the droplet size and the AuNPs concentration on the temperature reached upon irradiation. For that, monodisperse emulsions with a uniform distribution of AuNPs in the droplet core are required to assess the repeatability and the reliability of the measurements. The production of droplets by microfluidics is an interesting outlook for that purpose.

6.2.2 Transfer of the polymer capsules in aqueous environment

At this stage of the work, the polymer shells formed by interfacial complexation are only “capsule precursors” since they are still surrounded by an oil phase. To envisage biological applications, the polymer capsules should be transferred into an aqueous environment without falling apart. This extraction step is challenging since the mechanical stability of the polymer shell is difficult to evaluate in emulsion and even more difficult to extrapolate in all-aqueous environment. Two approaches have been tested to recover the capsules in water : the first one consisted in removing the oil phase from the original emulsion and redispersing the residual aqueous content in water, the second was based on the “cushion method” developed by He et al.²⁹⁹ In the following, we will briefly describe how we tried to adapt these strategies to our system and the experimental difficulties that we faced.

The first approach of capsule extraction that has been tested relied on the removal of the continuous
oil phase by simple evaporation. The FC-70 oil phase used so far was not appropriate for that purpose since
its boiling point was of 215 °C. This oil was thus substituted with another fluorinated oil, called Novec™
7100 (Sigma-Aldrich), characterized by a lower boiling point of 61 °C. The extraction was first tested on
PLL-g-PNIPAM shells. The emulsion was exposed to ambient air to let the oil phase evaporate. Then, a
large amount of water was added to the aqueous pellet to recover the capsules in an aqueous environment.
This ideal protocol is schematized in **Figure 6.1.A**. To assess the efficiency of the capsule recovery by
microscopy imaging, we supplemented the inner aqueous phase with 5 % of Cy3-labeled PNIPAM (playing
the role of fluorescent payload). The experimental results are schematized in Figure 6.1.B. The original
emulsion was characterized by stable micrometric water droplets. The presence of PNIPAM in the droplets
was evidenced by a high fluorescence level in the droplet core and the formation of polymer aggregates at 45 °C (see Figure 6.1.C).

<figure><img src="image_129.png" /><figcaption>**Figure 6.1.** Attempt of capsule extraction based on the evaporation of the continuous oil phase and the redispersion of the capsule precursors in water. A) Representation of the ideal protocol. Polymer capsule shells are formed in water-in-oil emulsion as described in the previous Chapters, but with Novec 7100 as the fluorinated oil (characterized by a low boiling point of 61 °C). The evaporation of the oil phase and the addition of a large amount of water should enable to redisperse the capsules precursors in an aqueous environment. B) Representation of the actual result. The evaporation of the oil phase led to the destabilization and coalescence of the water droplets. The presence of residual traces of oil resulted in the formation of an undesired direct emulsion upon redispersion of the pellet in water. C) Micrographs of the initial emulsion containing 1 wt% of PLL-g-PNIPAM and 5 wt% of Cy3-labeled PNIPAM (used as a fluorescent payload) in the aqueous phase and 0.05 wt% of Krytox in the Novec 7100 oil phase. D) Micrographs of the aqueous medium after redispersion of the pellet in water. The scale bar in C corresponds to 10 µm and is valid for all the micrographs.</figcaption></figure>

Upon oil evaporation, the water droplets were subjected to destabilization and coalescence, resulting in the disruption of the polymer shells. This process was confirmed by the absence of polymer capsules in the final aqueous phase. Instead, the “outer” aqueous phase was composed of dispersed polymer residues
(highlighted in fluorescence images) and small droplets that are probably oil droplets resulting from the emulsification of residual traces of oil contained in the evaporation pellet (Figure 6.1.D).

This preliminary result suggested that the PLL-g-PNIPAM shells have a poor mechanical stability. We tried to strengthen the polymer shells by introducing polymerizable groups along the PLL backbone of the copolymer. As an example, we synthesized a copolymer by co-grafting PNIPAM strands and acrylate-terminated PEG strands onto PLL. After formation of a polymer shell at the surface of water droplets, the copolymer chains were cross-linked via photo-polymerization of the acrylate groups. Upon application of the extraction protocol as described in Figure 6.1.A, the capsules shells were however subjected to the same destabilization issues than PLL-g-PNIPAM shells and were not successfully recovered in the aqueous phase.

As an alternative to the evaporation protocol that probably induces mechanical stress that disrupts the polymer shells, we tried to adapt the “cushion method” developed by He et al.²⁹⁹ This method consists in mixing the original emulsion containing the capsule precursors with a metastable emulsion that triggers the macroscopic phase separation of the mixture. During this demixing process, the fusion of the “metastable” aqueous droplets with the capsule precursors should lead to the recovery of the capsules in the water phase. The addition of a “cushion agent” (typically citrate) in the aqueous phase of the metastable emulsion enables to stabilize the polymer capsules during their transfer from oil into water. This method was originally implemented for the recovery of mechanically robust polymer capsules prepared by interfacial polymerization in water-in-toluene emulsions.²⁹⁹ Since toluene is less dense than water, the polymer capsules were easily collected in the bottom phase after the transfer process. In addition, due to the mechanical stability of their polymerized shell, the capsules could be subjected to washing and centrifugation steps without major degradation.

We tried to adapt this “cushion method” approach on our system. Briefly, PLL-g-PNIPAM shells were formed by interfacial complexation in an original water-in-oil emulsion (with FC-70 as the oil). Then, a metastable water-in-oil emulsion was prepared with (i) citrate in the aqueous phase, used as “cushion agent” and (ii) perfluorooctanol in the oil phase, used as a destabilizing agent. The metastable emulsion was injected in the initial emulsion to trigger the macroscopic phase separation of the water and oil phases. The polymer capsules were expected to be recovered at the bottom of the aqueous phase. Because the aqueous phase was lying onto the oil phase, the collection of the capsules – if they were intact – was particularly difficult. At the end of the process, microscopy imaging of an aqueous fraction picked up at the water/oil interface only showed residual traces of oil and no rigid capsules. This protocol may require further optimization to be able to properly recover polymer capsules in the water phase.
### 6.2.3 Production of capsules in microfluidics

As an alternative to the transfer of polymer capsules from the oil phase to an aqueous environment, we envisage to produce polymer shells in a microfluidic device in order to facilitate the recovery of the capsules. A promising approach is depicted in **Figure 6.2** and consists in the microfluidic generation of a water-in-oil-in-water (W/O/W) double emulsion in which the capsule shell is formed at the inner W/O interface and is recovered in the continuous water phase by evaporation of the thin intermediary oil layer.

<figure><img src="image_130.png" /><figcaption>**Figure 6.2.** Microfluidic approach for the production of polymer capsules by interfacial complexation in W/O/W double emulsion. A) Representation of the microfluidic device used to produce W/O/W double emulsion. A primary aqueous phase containing the polymer of interest (here PLL-g-PNIPAM) flows through the main channel of the device. In a first flow-focusing junction, an oil phase (with low boiling point) containing Krytox is injected orthogonally to pinch the aqueous flow, resulting in the formation of a W/O emulsion in which the interfacial complexation takes place. In a second flow-focusing junction, a second aqueous phase containing a semi-fluorinated surfactant called $F_8$-TAC$_{13}$ is injected to pinch the oil flow, resulting in the formation of a W/O/W double emulsion with a polymer shell formed at the inner W/O interface. B) Ideal illustration of the composition of the W/O/W emulsion coming out from the microfluidic device. Heating this emulsion would enable to (i) collapse and accumulate PLL-g-PNIPAM at the inner water/oil interface and (ii) evaporate the intermediary oil phase, eventually yielding a polymer capsule dispersed in an aqueous environment.</figcaption></figure>
Figure 6.2.A shows a promising microfluidic device and the composition of the phases that could be used to generate W/O/W emulsions. An aqueous solution containing the polymer of interest (here PLL-g-PNIPAM) is injected in the device and pinched at a first flow-focusing junction by an orthogonal flow of oil phase containing Krytox, resulting in the formation of a W/O emulsion where interfacial complexation can take place. In a second flow-focusing junction, an aqueous solution containing a surfactant that can stabilize oil-in-water emulsions (here the semi-fluorinated surfactant called $F_8$-TAC$_{13}$ is given as an example) is used to pinch the initial W/O emulsion, resulting in the formation of a W/O/W emulsion in which the inner aqueous droplet is surrounded by an intermediary oil layer.

If the oil used in this process has a low boiling point, the intermediary oil layer can be removed by evaporation upon heating, resulting in the dispersion of the polymer shells in an aqueous environment. This process is depicted in Figure 6.2.B in the ideal case of PLL-g-PNIPAM shells. For this system, the heating step is also expecting to trigger the collapse and accumulation of PLL-g-PNIPAM in the shell, which could enhance the mechanical properties of the capsule upon oil removal.

For the moment, this approach has not been tested yet but is envisaged through a collaboration with physicist colleagues (W. Urbach at the Physics Department of ENS and N. Taulier at the LIB lab of Sorbonne University), who already master the formation of W/O/W double emulsions involving fluorinated oils, essentially for the design of ultrasound-responsive carriers.

As the microfluidics generation of well-defined W/O/W emulsions in presence of polymeric additives would require important optimization, we started our collaborative work by focusing on a simpler, albeit promising, delivery system based on the intrinsic responsiveness of W/O/W emulsions. In absence of any interfacial polymer layer, W/O/W emulsions can still be seen as a core-shell system composed of an aqueous core surrounded by a protective oil shell. In this configuration, the evaporation of the oil shell would be the analogous of a polymer shell disruption, and would trigger the release of payloads pre-encapsulated in the aqueous core.

As a preliminary proof of concept, we demonstrated that the evaporation of the oil layer in W/O/W emulsions can be triggered by thermoplasmonic excitation of AuNPs encapsulated in the aqueous core. This experiment was performed on a "multi-core" double emulsion generated by microfluidics by Chloë Thimonier, a PhD student in the team of W. Urbach (ENS). The multi-core double emulsion, consisting in oil microdroplets containing multiple smaller water droplets in their core (see **Figure 6.3**), was produced in two steps: 1) an aqueous phase containing 0.05 wt% of AuNPs was emulsified in perfluorohexane (a fluorinated oil with a boiling point of 57 °C) containing 5 wt% of Krytox, yielding a nanometric primary W/O emulsion. The volume ratio of this W/O emulsion was 40/60 and the size of the water droplets was typically 500 nm. 2) This primary emulsion was injected in a microfluidic flow-focusing junction to be
pinched by an aqueous phase containing 0.1 wt% of F₈TAC₁₃, resulting in the multi-core W/O/W double emulsion showed in Figure 6.3. The “multi-core” configuration was preferred to a “simple-core” double emulsions in order to maximize the contact between the volatile oil and the AuNPs-loaded water phase, and consequently the probability of oil evaporation upon light irradiation.

The multi-core double emulsion was dispersed in a polymer gel solution (poloxamer 407 at 30 wt% in water) to immobilize the oil droplets, and was thermalized at 30 °C. When a 532-nm laser beam (15 mW) was focused on an individual oil droplet, a gas bubble appeared in less than 500 ms in the droplet core (see Figure 6.3), suggesting that the rise of temperature induced by thermoplasmonic excitation of encapsulated AuNPs has triggered the local evaporation of the oil phase. After a few seconds of laser irradiation, the droplet was totally disrupted. This illustrative experiment is a first step towards the formation of light-responsive delivery systems based on the combination of AuNPs and liquid shells made of volatile fluorinated oils.

<figure><img src="image_131.png" /><figcaption>**Figure 6.3.** Light-triggered destabilization of a multi-core W/O/W double emulsion, consisting in AuNPs-loaded nanometric water droplets dispersed in micrometric oil droplets exhibiting a low boiling point (57 °C). The focalization of a 532-nm laser beam (green disk) on a multi-core droplet induces a temperature rise that locally triggers the evaporation of the oil phase. A gas bubble (grey disk) rapidly grows within the droplet, resulting in its disruption in a few seconds. Scale bar in the micrographs = 15 µm.</figcaption></figure>
Microfluidics approaches appear thus as a promising extension to the work presented in this thesis for the design of various stimuli-responsive micro-carriers from water-in-fluorinated oil emulsions. The possibility to address physico-chemical changes in these systems with temperature, light or even ultrasounds, will be further explored via a collaboration between the two teams.
### 7.1 Materials

#### 7.1.1 Polymers and organic compounds

**Polymers:** Poly(L-lysine) hydrobromide (PLL) was purchased from Sigma-Aldrich (Mw = 15-30 kDa) and Alamanda Polymers (Mw = 6 kDa, 52 kDa). (N-hydroxysuccinimidyl ester)-terminated poly(N-isopropylacrylamide) (PNIPAM-NHS, Mw = 2 kDa), azide-terminated poly(N-isopropylacrylamide) (PNIPAM-N₃, Mw = 5 kDa), O-[N-(3-Maleimidopropionyl)aminoethyl]-O'-[3-(N-succinimidyloxy)-3-oxopropyl]triethylene glycol (Mal-PEG4-NHS) were purchased from Sigma-Aldrich. (N-hydroxysuccinimidyl ester)-terminated poly(ethylene glycol) (PEG-NHS, Mw = 2 kDa) was purchased from Rapp Polymere.

**Fluorophores:** 5(6)-carboxy-X-rhodamine N-hydroxysuccinimidyl ester (rho-NHS), 7-(diethylamino)coumarin-3-carboxylic acid N-succinimidyl ester (coum-NHS), dibenzocyclooctyne-Cy3 (DBCO-Cy3) and 8-anilino-1-naphthalenesulfonic acid (ANS) were purchased from Sigma-Aldrich.

**Protein:** Enhanced Green Fluorescent Protein (eGFP) has been produced and purified by a colleague (R. Chouket) according to a protocol described in the literature.²⁶⁷

**Other compounds:** Tris(2-carboxyethyl)phosphine (TCEP), monoethanolamine (MEA) and the different buffer ingredients (NaH₂PO₄, Na₂HPO₄, CHES, KSCN) were purchased from Sigma-Aldrich.

#### 7.1.2 Fluorinated species

Three types of Fluorinert™ fluorinated oil were purchased from Sigma-Aldrich: FC-40 (mixture of perfluoro-N-methyldibutylamine and perfluoro-tributylamine, density = 1.885), FC-70 (perfluoro-
tripentylamine, density = 1.940), and **Novec 7100** (methoxy-nonafluorobutane, density = 1.5). FC-40 was only used in surface tension experiments. FC-70 was used for the preparation of all the water-in-oil emulsions described in Chapter 3, 4 and 5. Novec 7100 was only used for preliminary experiments on capsule extraction presented in Chapter 6.

**Krytox** 157 FSL surfactant (Mw = 2.5 kDa) was purchased from Samaro.

### 7.1.3 Nanoparticles

Two types of fluorescent polystyrene nanoparticles were purchased from ThermoFisher Scientific (FluoSpheres™ range): NeutrAvidin™-coated nanoparticles (**NeutravNPs**) and carboxylate-modified nanoparticles (**CarboxyNPs**) purchased at 1 wt% in water. For both types, 200-nm NPs were used for zeta potential and DLS measurements, and 50-nm NPs were used for emulsion formulations and microscopy experiments. Both types of nanoparticles were displaying yellow-green fluorescence (505/515 nm).

Two types of gold nanoparticles (**AuNPs**) were purchased from NanoComposix: BioPure™ bare AuNPs (40-nm diameter, 1 mg/mL in water) and Econix™ AuNPs coated with 40 kDa poly(vinyl pyrrolidone) (50-nm diameter, 5 mg/mL in water).

Prior to use, NPs solutions were sonicated with a Sonics Vibra-Cell VCX 750 equipped with a 3 mm microtip for 2 min at 225 W.

### 7.1.4 Aqueous buffers

If not otherwise specified, aqueous solutions were systematically prepared in a home-made 7.7 mM phosphate buffer, prepared by dissolving 360 mg of NaH₂PO₄ (3 mmol) and 670 mg of Na₂HPO₄ (4.7 mmol) in 1 L of deionized water ([phosphate] = 7.7 mM, pH = 7.3, ionic strength = 17 mM, Debye length = 2.3 nm).

For the specific experiments about the pH-dependance of interfacial complexation (Chapter 3, section 3.4.3), we used a 30 mM Britton-Robinson (BR) buffer – made of 10 mM NaH₂PO₄/10 mM NaOAc/10 mM CHES – allowing for a buffering effect on a large range of pH. This buffer was prepared by dissolving 480 mg of NaH₂PO₄ (4 mmol), 328 mg of NaOAc (4 mmol) and 828 mg of CHES (4 mmol) in 0.4 L of deionized water (pH = 5.7, ionic strength = 30 mM, Debye length = 1.7 nm). The pH was adjusted at 4.3 (resp. 9.4) by addition of 150 µL of 1 M H₃PO₄ (resp. 500 µL of 1 M NaOH) in 30 mL of initial BR buffer. Aqueous phases containing the polymer were prepared in BR buffer at either acidic or alkaline pH, namely at pH = 4.3 or pH = 9.4.
## 7.2 Synthesis of polymers

### 7.2.1 Synthesis of PLL-g-PNIPAM and PLL-g-PEG

PLL-g-PNIPAM and PLL-g-PEG polymers were obtained by grafting NHS-terminated strands (PNIPAM or PEG) onto PLL. The number of NHS-terminated strands was adjusted as a molar fraction of the total number of lysine units ($H\cdot Br$ lysine, M = 209 g/mol) to reach the targeted theoretical grafting ratio $\tau$ (number of functionalized lysine units/total number of lysine units). Fluorescence properties were provided by the labeling of the PLL backbone with NHS-terminated fluorophores (rhodamine or coumarin). The synthesis protocol will be detailed for the preparation of PLL-g-PNIPAM. PLL-g-PEG was prepared by adjusting this protocol with reactive PEG chains instead of reactive PNIPAM chains).

PLL hydrobromide ($M_w$ = 15-30 kDa, 15 mg) and NHS-terminated PNIPAM ($M_w$ = 2 kDa, 48 mg to get $\tau$ = 0.33) were dissolved in sodium tetraborate alkaline buffer (5 mL, pH = 8.5) under stirring at 4 °C to facilitate the dissolution of PNIPAM. Then the reaction was completed under stirring for 4h at room temperature. The resulting solution was dialyzed against deionized water (regularly renewed) for 3 days in a Thermo Scientific Slide-A-Lyzer cassette (3-12 mL, $M_w$ cutoff = 3.5 kDa) and finally freeze-dried for 2 days in a Labconco Freezone Plus 2.5 apparatus, yielding a white fluffy powder (144 mg, yield = 83%). For the synthesis of fluorescent derivatives, the appropriate reactive dye (rho-NHS or coum-NHS) was added in the reaction batch at the same time than the reactive PNIPAM chains. The dye was prepared in solution at 1 g/L in DMSO and the appropriate volume was added in the reaction batch to target ~1 graft/PLL chain.

A representative ¹H NMR spectrum of PLL-g-PNIPAM is showed in **Figure 7.1**. Peak identification (chemical shift in ppm, proton identification, integration): 1.16, PNIPAM [CH₂-CH-CO-NH-CH-(CH₃)₂], 55.2H; 1.4-2.2, lysine [β,γ,δ-CH₂] + PNIPAM [CH₂-CH-CO-NH-CH-(CH₃)₂], 35.7H; 2.45-2.52, lysine-CO-CH₂-CH₂-S-PNIPAM + PNIPAM [CH₂-CH-CO-NH-CH-(CH₃)₂, 1.3H; 2.70-2.75, lysine-CO-CH₂-CH₂-S-PNIPAM + PNIPAM [CH₂-CH-CO-NH-CH-(CH₃)₂], 2.1H; 2.98, lysine [non-grafted α-CH₂], 1.7H; 3.18, lysine [grafted α-CH₂], 0.4H; 3.92, PNIPAM [CH₂-CH-CO-NH-CH-(CH₃)₂], 9.2H; 4.32, lysine backbone [CONH-CHR-CONH], 1H (*reference for integration*).
<figure><img src="image_132.png" /><figcaption>**Figure 7.1.** ¹H NMR spectrum of PLL-g-PNIPAM at 400 MHz in D₂O.</figcaption></figure>

The experimental grafting ratio $\tau_{exp}$ of the PLL backbone by PNIPAM was determined by the signal integration value of the protons characterizing the grafted lysine units (protons e') with the proton of the PLL backbone (proton a) as a reference (2 protons e' for 1 proton a): $\tau_{exp} = 0.40/2 = 0.20$. This result means that 1/5 of the lysine units have been grafted with a PNIPAM strands. For a PLL chain with a mean molar mass of 22 kDa (105 lysine units), it means that 21 PNIPAM chains have been grafted in average. In total, the PLL-g-PNIPAM copolymer has a molar mass of around 64 kDa, the PLL backbone representing 34 wt% and the PNIPAM strands representing 66 wt%.

### 7.2.2 Fluorescent labeling of “free” PNIPAM

Controlled experiments presented in sections 3.3.2 and 3.5.4 have required the use of fluorescent “free” PNIPAM chains (i.e. non-grafted on PLL). A typical protocol for such fluorescence labeling is reported below:

Azide-terminated PNIPAM ($M_w$ = 5 kDa, 100 mg) was dissolved in deionized water (5 mL). DBCO-Cy3 (23 µL at 1 g/L in PBS buffer) was added and the solution was stirred for 5h at room temperature. The resulting solution was dialyzed against deionized water (regularly renewed) for 3 days (Slide-A-Lyzer cassette, $M_w$ cutoff = 3.5 kDa) and finally freeze-dried for 2 days, yielding a pink fluffy powder.
### 7.2.3 Synthesis of PLL-g-PAAMAN

#### Synthesis of PAAMAN chains

The UCST poly(acrylamide-co-acrylonitrile) (PAAMAN) used in Chapter 5 were synthesized in the group of Prof. Yue Zhao at Sherbrooke University (Canada). A typical synthesis protocol for the PAAMAN named P2 in Chapter 5 is reported below:³⁰⁰

N-acrylamide was recrystallized in chloroform, acrylonitrile was purified in basic alumina column. The RAFT chain transfer agent, S,S'-bis(α,α'-dimethyl-α''-acetic acid)trithiocarbonate (BTC), was synthesized according to Lai et al.³⁰¹ To prepare the random copolymer, a mix of acrylamide (2.8 g, 1eq), acrylonitrile (760 mg, 0.36eq), BTC (74.4 mg, 0.007eq), AIBN (8.7 mg, 0.001eq), and DMSO (12.3 mL) was degassed by freeze-vacuum-thaw (3 cycles), then heated to 70 °C for 2 hours. The final polymer was precipitated twice from DMSO into methanol, then dialyzed against deionized water for 3 days and recovered by lyophilization. The AAm:AN feed ratio was of 73:27 mol/mol but yielded a 66:34 mol/mol ratio in the copolymer.

The ¹H NMR spectrum of PAAMAN P2 in d₆-DMSO is represented in **Figure 7.2**.

<figure><img src="image_133.png" /><figcaption>**Figure 7.2.** ¹H NMR spectrum of PAAMAN P2 with x=0.66 and y=0.34 at 400 MHz in d₆-DMSO.</figcaption></figure>
Aminolysis of PAAMAN chains

For the synthesis of PLL-g-PAMAAN, we performed the *in situ* aminolysis reaction of the trithiocarbonate unit present in the middle of the PAAMAN chains. The efficiency of this reaction was previously controlled with UV-Vis spectrophotometry. The spectrum of PAAMAN chains at 0.1 wt% in DMSO was measured before and after addition of an excess of monoethanolamine (MEA, 30eq) and tris(2-carboxyethyl)phosphine (TCEP, 10eq) (Figure 7.3.A). The absorbance peak characteristic of the trithiocarbonate unit at 280 nm disappeared after addition of MEA and TCEP, suggesting that it was readily cleaved by aminolysis to yield two thiol-terminated PAAMAN strands.

This reaction was then conducted in water retrieve the aminolyzed polymer after purification and
lyophilization (according to the precipitation-washing protocol presented above for PLL-g-PAAMAN).
The cloud point of the aminolyzed polymer was measured in water by turbidimetry and was rigorously
equal to the cloud point of the original polymer (Figure 7.3.B), suggesting that the UCST-type behavior of
PAAMAN chains was not affected by the aminolysis reaction.

<figure><img src="image_134.png" /><figcaption>**Figure 7.3.** A) Absorbance spectrum of the PAAMAN polymer P2 used in Chapter 3, before and after aminolysis with monoethanolamine (MEA) and tris(2-carboxyethyl)phosphine (TCEP) in DMSO. B) Turbidimetry heating curves of the polymer before and after aminolysis.</figcaption></figure>
Synthesis of poly(L-lysine)-g-poly(acrylamide-co-acrylonitrile) (PLL-g-PAAMAN)

A typical synthesis of PLL<sub>0.53</sub>-g-PAAMAN<sub>0.47</sub> (PLL/PAAMAN weight ratio of 53/47 %) is reported below.
A similar protocol was used for the synthesis of PLL<sub>0.42</sub>-g-PAAMAN<sub>0.58</sub> with a 22 kDa PLL instead of
52 kDa, and with an appropriate polymer ratio.

52 kDa PLL (20 mg, 250eq lysine) and Mal-PEG4-NHS (2.0 mg, 10eq) were dissolved in 3 mL D₂O in a 10 mL round-bottom flask. The solution was stirred for 4 h at room temperature. 300 µL of solution were taken and mixed with 300 µL of D₂O for NMR analysis of the intermediary PLL-g-(PEG4-Mal). After heating the solution at 60 °C, PAAMAN (17.2 mg, 3eq), MEA (4 µL, 175eq) and TCEP (2.9 mg, 30eq) were introduced, and the solution was stirred at 60 °C for 4 h. After cooling back to room temperature, the solution was purified by 3 centrifugation/redispersion cycles. The centrifugation was performed at 4 °C for 10 min at 15 000 rcf. After removing the supernatant, the pellet was redispersed in DI water by sonication using a Sonics Vibra-Cell VCX 750 equipped with a 3-mm microtip. The final solution was lyophilized overnight using a Labconco Freezone Plus 2.5 apparatus, yielding a white powder.

For the synthesis of fluorescent derivatives, the appropriate reactive dye (rho-NHS or coum-NHS) was added in the reaction batch at the same time than the PEG linker. The dye was prepared in solution at 1 g/L in DMSO and the appropriate volume was added in the reaction batch to target ~2 grafts/PLL chain.

**Figure 7.4** shows a typical ¹H NMR spectrum of the maleimide-grafted PLL intermediary in D₂O (chemical shift in ppm, proton identification, integration): 1.5-1.7 (lysine [β,γ,δ-CH₂], 6.22H), 2.5 (linker [CH₂-CONH], 0.23H), 2.8 (released NHS units, 0.33H), 3.0 (lysine [non-grafted α-CH₂], 1.94H), 3.2 (lysine [grafted α-CH₂], 0.15H), 3.3 (linker [CONH-CH₂], 0.16H), 3.6-3.8 (PEG backbone, 1.5H), 4.3 (lysine backbone, 1H), 6.9 (maleimide double-bond, 0.14H).
<figure><img src="image_135.png" /><figcaption>**Figure 7.4.** ¹H NMR spectrum of PLL-g-(PEG4-Mal) at 400 MHz in D₂O.</figcaption></figure>

The experimental grafting ratio $\tau_{exp}$ of PLL by Mal-PEG4-NHS linkers was determined by comparing ¹H NMR integration values of protons characterizing grafted and non-grafted lysine units (e' and e respectively), with protons of PLL backbone (a) as reference protons: $\tau_{exp} = 0.15/2 \sim 0.07$.
A typical ¹H NMR spectrum of PLL<sub>0.53</sub>-g-PAAMAN<sub>0.47</sub> in d<sub>6</sub>-DMSO is represented in **Figure 7.5**.

<figure><img src="image_136.png" /><figcaption>**Figure 7.5.** ¹H NMR spectrum of PLL<sub>0.53</sub>-g-PAAMAN<sub>0.47</sub> at 400 MHz in d<sub>6</sub>-DMSO.</figcaption></figure>

The UCST behavior of PLL-g-PAAMAN (0.53/0.47 wt%) was compared to the one of non-grafted PAAMAN chains in presence of PLL (see **Figure 7.6**). Compared to pure PAAMAN and PAAMAN in presence of PLL (but non grafted), the cloud point of PLL-g-PAAMAN is shifted from 4-5 °C towards higher temperatures.

<figure><img src="image_137.png" /><figcaption>**Figure 7.6.** Turbidity heating curve of solutions of PAAMAN (1 wt%), PAAMAN + PLL (1 wt% + 1 wt%) and PLL-g-PAAMAN (1 wt%) in water.</figcaption></figure>
## 7.3 Preparation of emulsions

Water-in-oil emulsions were obtained by manually emulsifying 5 µL of aqueous phase (containing polymers, NPs, and other water-soluble compounds) in 95 µL of oil phase (containing Krytox) in a 0.5 mL Eppendorf tube.

For microscopy experiments, the emulsions were observed in hollow rectangle glass capillaries (CMScientific) with the following dimensions: inner height = 100 µm, width = 1 mm, length ~ 1 cm (adjusted manually). Prior to emulsion injection, the internal sides of the capillary were coated with PLL-g-PEG to prevent the breakage of water droplets at the contact with bare glass side (see Chapter 2). Briefly the capillary was filled with an aqueous solution of PLL-g-PEG ($M_w$ = 2 kDa, 1 g/L in 7.7 mM phosphate buffer) and incubated for 30 min before evaporation on a heating plate at 60 °C. Then, deionized water was injected and evaporated at 60 °C. The microscopy samples were prepared by injecting a few µL of emulsion in a PLL-g-PEG-coated capillary. After injection, the capillary was sealed at its extremities with Vitrex putty.

## 7.4 Coating of NPs with PLL derivatives

200-nm NeutravNPs and CarboxNPs were coated with PLL-g-PNIPAM for dynamic light scattering (DLS) and zeta potential measurements. The coating protocol was the following: a 250-µL solution containing 0.5 wt% of PLL-g-PNIPAM and 0.02 wt% of NPs in phosphate buffer (~ $10^{16}$ polymer chains for ~ $10^{10}$ NPs) was gently sonicated for 5 min prior to 1h incubation at room temperature. The excess of polymer was then removed by filtration on Amicon® Ultra 0.5 mL centrifugal filters (cut-off 100 kDa), following a sequential procedure of i) centrifugation (10 min at 14 000 rcf), ii) removal of the filtrate and iii) redispersion of the NPs (retained in the filter) by addition of 250 µL of fresh phosphate buffer. After three cycles of this filtration procedure, the pellet was retrieved from the filter and diluted in 250 µL of buffer + 500 µL of pure water to yield 1 mL of solution useable in DLS and Zeta potential measurements. Control solutions of commercial NPs at 0.02 wt% were obtained by simple dilution and sonication of the commercial solutions in 500 µL phosphate buffer + 500 µL pure water.

AuNPs (40-nm bare or 50-nm PVP-coated) were too small to be submitted to the same filtration process on Amicon® filters. Consequently, DLS and zeta potential measurements were directly performed on solutions containing both AuNPs and PLL derivatives (first taking care that residual polymer chains were not contributing to the signal of interest). The solutions typically contained 0.1 wt% of polymer and 0.05 wt% of AuNPs for incubation, and were diluted 100 times for DLS and zeta measurements.
## 7.5 Interfacial tension measurements

The interfacial tension measurements were performed at room temperature using a TRACKER™ automatic drop tensiometer (Teclis Scientific). The oil phase was made of FC-40 containing Krytox surfactant. The aqueous phase was made of 6 kDa PLL in phosphate buffer. Pure (deionized) water was also used as a reference aqueous medium. Rising aqueous droplets were formed at the tip of a J-shaped curved needle submerged in 2-mL bath of oil phase. The droplet shape was imaged with a camera and processed with the TRACKER apparatus software. A value of the interfacial tension was obtained from an automatic fit of the droplet profile using the Young-Laplace model.<sup>302</sup>

The reliability of the profile fit depends on the shape and the stability of the droplet. First, the Young-Laplace model remains valid as long as gravity and capillarity effects are balancing each other. When the droplet starts to grow out of the needle, its shape is quasi-spherical because essentially dictated by capillarity. The action of gravity starts to be visible as soon as the droplet elongates at the tip of the needle, what requires a minimum injected volume.<sup>303</sup> Thus, the interfacial tension can be accurately measured provided a droplet shape which is significantly different from the spherical shape. Then, the droplet surface needs to be at equilibrium to properly fit its profile and to extract a reliable value of interfacial tension.<sup>303</sup>

In our case, due to the high difference of density between the two phases, the water droplets tend to quickly detach from the needle and to flow up in the oil phase. The optimal shape and the interfacial equilibrium are probably not reached before the droplet is released. As a consequence, the raw values of interfacial tension measured with this approach could be skewed. Nevertheless, droplets were imaged after the maximum stabilization time before release and approximate values of the interfacial tension were extracted from the fit of the droplet profile in this condition. For the interpretation, the focus was put on the qualitative evolution of the interfacial tension rather than on the quantitative data.
## 7.6 UV-Vis spectrophotometry

Absorbance and transmittance spectra were recorded on a single cell Thermo Scientific Evolution Array UV-Vis spectrophotometer equipped with a Peltier temperature-controlled cell holder (+/- 0.1 °C). The sample solutions were injected in a 150 µL quart micro-cuvette with an optical path length of 1 cm.

### UV-Vis spectrophotometry of AuNPs

Absorbance spectra of AuNPs were recorded at 25 °C on solutions containing 0.005 wt% of AuNPs and a variable concentration of 6 kDa PLL in the 0-5 wt% range.

### Turbidity measurements of thermoresponsive polymer solutions

Thermoresponsive polymers (PNIPAM, PAAMAN and their PLL comb-like derivatives) were dissolved at 1 wt% in the appropriate buffer (pure water, phosphate buffer or KSCN buffer). The solutions were submitted to a sweep of temperature between 20 °C and 60 °C by steps of 2 °C. A spectrum was recorded at each step temperature after 3 min thermalization. The mean transmittance between 650 and 750 nm was used to plot the turbidimetry curves.

## 7.7 Zeta potential measurements

Zeta potential measurements were performed on NPs solutions prepared according to the protocol described in section 7.4, and injected in DTS 1060C Zeta cells. The measurements were performed at 25 °C using a Zetasizer Nano-ZS (Malvern Instruments). For each sample, the average zeta distribution was obtained from the average of three measurements (22 runs each), separated by 2 min stabilization.
## 7.8 Dynamic light scattering (DLS) measurements

DLS experiments were performed at 90° angle and at room temperature on a BI-200SM Goniometer System (Brookhaven Instruments Corp.) equipped with a 30 mW LED laser operating at 637 nm and a BI-CROSS CORR dual photomultiplier as the detector. The scattered light intensity and the correlation function were monitored using BIC Dynamic Light Scattering software. The mean hydrodynamic diameter was automatically extracted from the correlogram using the cumulant expansion analysis.

### Characterization of water-in-oil emulsions

DLS was used to determine the influence of polymer and Krytox concentrations on the stability of water-in-oil emulsions, following a suitable design of experiments. The results are given in Appendix B. For these experiments, DLS samples were prepared as follows: 5 %vol water-in-oil emulsions were prepared by mixing 5 µL of aqueous phase (PLL-g-PNIPAM in phosphate buffer) and 95 µL of oil phase (Krytox in FC-70 oil) in a 0.5 mL Eppendorf tube. The two phases were emulsified by tip sonication using a Sonics Vibra-Cell VCX 750 for 3 min at 300 W (the sample was placed in an ice bath to limit undesired heating). The resulting emulsions were diluted 20 times in pure FC-70 oil prior to DLS measurements. The mean hydrodynamic diameter of emulsion droplets and the associated standard deviation were measured at 25 °C.

### Characterization of nanoparticles

DLS measurements were performed on solutions of AuNPs prepared according to the protocol described in section 7.4. The hydrodynamic diameter of the samples was measured at temperature intervals of 5 °C between 20 °C and 50 °C (the acquisition was made after 5 min thermalization at each temperature step).
## 7.9 Microscopy techniques

### 7.9.1 Bright field and phase-contrast microscopy

Bright field and phase contrast imaging were performed with a LEICA DM IRE2 microscope equipped with a long-focal ×63 air objective. Images were acquired with a Retina 6000 Q-imaging camera and processed with Micro-Manager 1.4 software (ImageJ).

The plasmonic excitation of AuNPs was achieved with a 532-nm diode laser with 40 mW nominal power (DJ532-40 DPSS model from Thorlabs) that was mounted on the side port of the microscope. The laser was controlled with a TED200C Thermoelectric Temperature Controller and a LDC210C Laser Diode Controller (from Thorlabs).

### 7.9.2 Confocal Laser Scanning Microscopy

#### Confocal microscope

Confocal images were acquired on a Zeiss LSM 710 META Laser Scanning Microscope equipped with a Plan Apochromat x20 (0.8 NA) objective and a temperature-controlled heating stage (25-50 °C). To ensure emulsion thermalization, capillaries filled with the emulsion were placed on a pierced supporting copper slide that was fixed on the heating stage of the confocal microscope. Low temperatures (16-20 °C) were achieved by placing a glass petri dish full of ice on top of the supporting copper slide close to the capillary. The actual temperature of the supporting copper slide was measured with a REED ST-640B thermocouple. The 405 nm excitation of ANS or coumarin was performed using a 30 mW CW/pulsed diode laser. The 458 nm excitation of eGFP or FluoSpheres™ NeutrAvidin™-Labeled Microspheres, and the 514 nm excitation of rhodamine or Cy3 were performed using a 25 mW argon laser. Images were acquired using LSM ZEN 2009 software and processed with Fiji software (ImageJ).

#### Fluorescence recovery after photobleaching (FRAP) experiments

FRAP experiments were performed on shells made of coumarin-labeled polymers (PLL-g-PAAMAN, PLL-g-PEG) at 18 °C and 48 °C on the Zeiss LSM 710 confocal microscope presented above. The sample was imaged upon 405-nm excitation at 0.15 mW (0.5 % of the maximum intensity) with a scanning speed of 0.39 µs/px. The photobleaching was performed in a circular region of interest (ROI) upon 405-nm excitation at 30 mW (100 % of the maximum intensity) with a scanning speed of 6.30 µs/px. The acquisition rate was set at 1 image/second and the mean fluorescence level in the ROI was monitored for 10 s before bleaching and 120 s after.
### 7.9.3 Grating-shadow phase microscopy (GSPM)

#### Microscopy set up

GSPM experiments were performed in the lab of Guillaume Baffou at the Fresnel Institute (Marseille). The OPD images were recorded on a home-made optical set up including : a 700 mW LED illumination at 625 nm (M625L3, Thorlabs), a x60 objective (LUC Plan FLN, NA 0.7, Olympus), a and a SID4-sC8 wavefront sensor and imaging device (Phasics). The plasmonic excitation of the sample was achieved with a MGL-FN-532 DPSS laser operating at 532 nm with a nominal power up to 1500 mW (CNI Optoelectronics Tech). The power at the sample level was limited to 40 mW with a laser spot of 50 µm in diameter. The whole optical set-up is represented in Chapter 4 (section 4.4.2.5, Figure 4.19).

#### Measurement of the $\frac{dn_0}{dT}$ coefficient of FC-70 oil

The measurement of the $\frac{dn_0}{dT}$ coefficient of FC-70 was performed using GSPM. A 12 µm-deep crater etched in a glass slide was filled with FC-70 oil. An OPD image of the crater was taken at different temperature upon heating. **Figure 7.7.A** shows the OPD image at 90 °C. At each temperature, the refractive index of the oil $n_0$ was deduced from the measurement of the OPD ($\delta$) between the bottom of the crater and the surface of the glass slide, according to the formula: $n_0 = n_{air} + \delta/h$ with $n_{air} = 1.00$ and $h = 12$ µm. The evolution of $n_0$ with the temperature is shown in Figure 7.7.B. The slope of this curve gives $\frac{dn_0}{dT} = -3.16 \times 10^{-4} K^{-1}$. This experiment was performed by Sadman Shakib, PhD student in the team of Guillaume Baffou.

<figure><img src="image_138.png" /></figure>

<figure><img src="image_139.png" /><figcaption>**Figure 7.7.** GSPM measurement of the $\frac{dn_0}{dT}$ coefficient of FC-70 oil. A) OPD image of a 12-µm crater filled with FC-70 oil at 90 °C. The refractive index of the oil is deduced from the measurement of the OPD generated by the crater. B) Evolution of the refractive index of the oil with temperature (triplicate) and corresponding linear regression.</figcaption></figure>
## 7.10 Scanning electron microscopy (SEM) imaging

SEM imaging was performed on isolated capsules resulting from the drying of water-in-oil emulsions (5 µL deposited on a glass coverslide) for three days at ambient air. To prevent the degradation of dried capsules during SEM imaging to enhance their contrast, the samples were metalized by deposition of a 30 nm-thin layer of Ag using an Emitech K675X Sputter Coater (Ag sputtering for 60 s at 120 mA). SEM images were recorded with a Hitachi TM3030 Tabletop Microscope either at 15 kV or in EDX mode.
# Appendix B

## Influence of the formulation of water-in-oil emulsions on the size and stability of emulsion droplets

As a complement to the surface tension measurements and the qualitative observation of the emulsions made in Chapter 1 (sections 3.2.1 and 3.2.3), the influence of the Krytox and PLL-g-PNIPAM concentrations on the water/oil interface stability was investigated by measuring the droplet size for different emulsion formulations by dynamic light scattering (DLS). This quantitative data will be analyzed using the statistical approach of fractional design of experiment (DOE).

The Krytox and PLL-g-PNIPAM concentrations were studied as two factors (factors A and B respectively) in a 2² factorial DOE. Two levels – one low denoted “-1” and one high denoted “+1” – were assigned to both factors, leading to 4 different formulations for which the mean droplet size was measured by DLS. The levels of the two factors were chose to cover an experimental range of concentration that seemed relevant for the formulation of the emulsions. The detailed formulations of the DOE are summarized in Table 8.1.

<table><caption>**Table 8.1.** Composition of the 2² factorial design of experiment implemented to determine the effect of Krytox and PLL-g-PNIPAM on the droplet size of water-in-oil emulsions.</caption><thead><tr><th></th><th>Factor A<br>Krytox concentration (wt%)</th><th>Factor B<br>Polymer concentration (wt%)</th></tr></thead><tbody><tr><td>Low level (-1)</td><td>0.01</td><td>0.5</td></tr><tr><td>High level (+1)</td><td>0.05</td><td>5.0</td></tr></tbody></table>

For each emulsion formulation, the *mean* water droplet diameter was measured on 3 different samples to determine an *average* diameter and an experimental variance (accounting for the experimental residual error). Note the difference between the terms “mean” and “average” in this case: the *mean* diameter refers to the mean diameter *within a given size distribution* (resulting from the measurement of a given polydisperse sample); and the *average* diameter results from the calculation of the *arithmetic* mean of the (mean) diameters measured for the 3 samples. For a given formulation *i*, the mean diameter measured by
DLS on the sample j will be denoted $d_{i,j}$ and the average diameter calculated from the 3 samples will be denoted $\bar{d}_i$.

The experimental results of the DOE are gathered in the **Table 8.2**. The average diameters are also graphically reported below in **Figure 8.1.A**.

<table><caption>**Table 8.2.** DLS measurements of the water droplet diameter for all the samples prepared in the DOE.</caption><thead><tr><th>Formulation i</th><th>Factor A</th><th>Factor B</th><th>Crossed factor AB</th><th>Mean diameter Sample 1 d<sub>i,1</sub> (nm)</th><th>Mean diameter Sample 2 d<sub>i,2</sub> (nm)</th><th>Mean diameter Sample 3 d<sub>i,3</sub> (nm)</th><th>Average diameter d&#773;<sub>i</sub> (nm)</th><th>Experimental variance σ<sup>2</sup><sub>i</sub> (nm<sup>2</sup>)</th><th>Std Dev σ<sub>i</sub> (nm<sup>2</sup>)</th></tr></thead><tbody><tr><td>1</td><td>-1</td><td>-1</td><td>+1</td><td>657</td><td>276</td><td>428</td><td>454</td><td>36826</td><td>192</td></tr><tr><td>2</td><td>+1</td><td>-1</td><td>-1</td><td>100</td><td>96</td><td>140</td><td>112</td><td>597</td><td>24</td></tr><tr><td>3</td><td>-1</td><td>+1</td><td>-1</td><td>227</td><td>347</td><td>166</td><td>358</td><td>8494</td><td>92</td></tr><tr><td>4</td><td>+1</td><td>+1</td><td>+1</td><td>80</td><td>70</td><td>73</td><td>74</td><td>29</td><td>5</td></tr></tbody></table>

<figure><img src="image_140.png" /><figcaption>**Figure 8.1.** DLS analysis of water-in-oil emulsions. A) Evolution of the hydrodynamic diameter of water droplets as a function of the PLL-g-PNIPAM concentration in the aqueous phase and the Krytox concentration in the oil phase (error bars = standard deviation on 3 samples). B) Evolution over time of the droplet diameter and the scattered intensity for the emulsion with 2 wt% PLL-g-PNIPAM and 0.03 wt% Krytox.</figcaption></figure>
The results of the formulations of interest are located at the corners of the experimental domain showed in Figure 8.1.A. In presence of the lowest Krytox concentration (0.01 wt%), relatively big droplets and high standard deviations were observed, suggesting that the emulsions were highly polydisperse and barely repeatable, which are signs of poor stability. On the other hand, in presence of the highest Krytox concentration (0.05 wt%), significantly smallest droplet sizes were reached (around 100 nm) with low standard deviations, indicating an efficient dispersion of the water phase in the oil phase. At first sight, the effect of PLL seems not to be significative compared to the effect of Krytox, what is consistent with the results of interfacial tension reported in Figure 3.3 in Chapter 3 (the same decrease of the interfacial tension with Krytox concentration was observed independently of the PLL concentration).

To validate these observations, a statistical analysis of the effects of Krytox and PLL-g-PNIPAM concentrations was developed as reported in the following.

As 3 independent samples were tested for each formulation *i*, the experimental variance $\hat{\sigma}_i^2$ for each formulation was determined with 2 degrees of freedom (dof). In total, the DOE involves 8 dof (4 formulations with 2 dof per formulation). Assuming that the hypothesis of homoscedasticity is verified (all the $\hat{\sigma}_i^2$ estimate the same residual variance), the intraclass variance $\hat{\sigma}^2$ can be used as an optimal estimation of the residual variance. It is calculated as the arithmetic mean of the variances, weighted by their respective number of dof:

$$ \hat{\sigma}^2 = \frac{\sum_i 2\hat{\sigma}_i^2}{8} = 11487 \text{ nm}^2 $$

The average effect $\bar{E}$ of each factor was deduced from the results by calculating the arithmetic mean of the diameter measured for each formulation but weighted with the suitable $\pm 1$ factor indicated in Table 8.2.

As an example: $\bar{E}(A) = \frac{-\bar{d}_1+\bar{d}_2-\bar{d}_3+\bar{d}_4}{4} = -128.6 \text{ nm}$. The effects were calculated for A, B and the cross-factor AB (accounting for the interaction between A and B) and are reported in Table 8.3. Given that the homoscedasticity is assumed for the 12 experiments, the variance on the effects $\hat{\sigma}_E^2$ is equal to $\hat{\sigma}^2/12 = 957 \text{ nm}^2$. From that, we can deduce an estimation for the standard deviation on the effects which is $\hat{\sigma}_E = 31 \text{ nm}$.

To evaluate the significativity of the effects, we assumed that the effect of a given factor *X* is not significant, i.e. $E(X) = 0$ (null hypothesis) and we proceeded to a Student test to verify it from a statistical point of view. Under this condition, the experimental ratio $\bar{E}(X)/\hat{\sigma}_E$ follows a Student law with 8 dof ($\bar{E}(X)/\hat{\sigma}_E \sim t_8$). We calculated the experimental Student parameter $t_{exp} = \bar{E}(X)/\hat{\sigma}_E$ for each factor *X* and
compared it to the corresponding tabulated value $t_8$ (which depends itself on a confidence parameter $p$). If $|t_{exp}| > t_8(p)$ for a given $p$, the null hypothesis can be rejected with a level of confidence of $1 - p$, indicating that the factor $X$ has a significant effect on the droplet diameter (with a certain level of confidence).

The experimental Student parameters $t_{exp}$ for the factors A, B and AB are reported in Table 8.3. The tabulated values of the Student parameters with 8 dof $t_8$ for a given confidence parameter $p$ are given in Table 8.4.

<table><caption>**Table 8.3.** Average effect and experimental Student parameters calculated for all the factors of the DOE.</caption><thead><tr><th>Factor<br><b>X</b></th><th>Effect<br>Sample 1</th><th>Effect<br>Sample 2</th><th>Effect<br>Sample 3</th><th>Average effect<br><span style="text-decoration: overline;">E</span> (nm)</th><th>Experimental Student<br>parameter (with 8 dof)<br><b>t</b><sub>exp</sub> = <span style="text-decoration: overline;">E</span>/<span style="text-decoration: overline;">σ</span><sub>E</sub></th></tr></thead><tbody><tr><td>A</td><td>-176,05</td><td>-114,45</td><td>-95,25</td><td><b>-128.6</b></td><td><b>-4.1</b></td></tr><tr><td>B</td><td>-112,55</td><td>11,25</td><td>-82,35</td><td><b>-61.2</b></td><td><b>-2.0</b></td></tr><tr><td>AB</td><td>102,5</td><td>-24,25</td><td>48,7</td><td><b>+42.3</b></td><td><b>+1.4</b></td></tr></tbody></table>

<table><caption>**Table 8.4.** Tabulated values of Student parameters with 8 degrees of freedom as a function of the confidence parameter.</caption><thead><tr><th>Confidence parameter<br><b>p</b></th><th>Student parameter with 8<br>dof<br><b>t</b><sub>8</sub></th></tr></thead><tbody><tr><td><b>0.01</b></td><td><b>3.355</b></td></tr><tr><td>0.02</td><td>2.896</td></tr><tr><td>0.05</td><td>2.306</td></tr><tr><td>0.10</td><td>1.860</td></tr><tr><td>0.20</td><td>1.387</td></tr></tbody></table>

The data will be analyzed taking $p = 0.01$, so that the results of the Student test can be guaranteed with a confidence of 99 % (see Table 8.4). According to the results given in Table 8.3, the condition $|t_{exp}| > t_8(0.01) = 3.355$ is verified for the factor A ($t_{exp} = -4.1$ in green) but not for the factors B ($t_{exp} = -2.0$ in red) and AB ($t_{exp} = +1.4$ in red). Consequently, the null hypothesis can be rejected for the factor A with a confidence of 99 % but not for the factors B and AB. In other words, the factor A has a significant effect (from a statistical point of view) on the water droplet diameter, what is not the case for
the factor B or the crossed-factor AB. Coming back to the physics of the system, these results indicate that the Krytox concentration has a significant influence on the water droplet size and that this size decreases when the Krytox concentration increases ($E(A) < 0$). In contrast, the PLL-g-PNIPAM concentration does not have a major influence on the water droplet size and there is no interaction between the effects of Krytox and PLL-g-PNIPAM concentrations.

In Section 3.2.1, we reported that, in presence of PLL as complexation partner, an increase in Krytox concentration led to a sharp decrease in water/oil interfacial tension. Considering the process of droplet formation upon emulsification, we will show that the interfacial tension is a key parameter that dictates the size of the emulsified droplets. The driving force of droplet formation upon emulsification is the shearing stress produced by the sonication, which stands as $\eta\dot{\gamma}$ with $\eta$ the viscosity of the continuous oil phase and $\dot{\gamma}$ the shear rate. In opposition, the droplet formation is hindered by the energetic cost related to the creation of water/oil interfaces, which is nothing else than the interfacial stress $\Gamma/R$ with $\Gamma$ the water/oil interfacial tension and $R$ the radius of the droplets. The competition between shearing and interfacial stresses can be described by a dimensionless number, called the capillary number $Ca$, which reads:

$$Ca = \frac{\text{shearing stress}}{\text{interfacial stress}} = \frac{\eta\dot{\gamma}R}{\Gamma}$$

The regime of droplet formation is set when the shearing stress balances the interfacial stress, i.e. for $Ca \sim 1$. Given that $\eta$ and $\dot{\gamma}$ are fixed in the experimental conditions (the oil viscosity is a constant and the same sonication power was used for all the emulsions), the droplet radius $R$ would thus be proportional to the interfacial tension $\Gamma$. Then an increase in Krytox concentration leads to a decrease in interfacial tension, and consequently a decrease in the droplet size formed upon emulsification. This result makes the connection between the interfacial tension measurements reported in Chapter 3 (section 3.2.1) and the droplet size measurements reported in this Appendix.

In addition to the 4 formulations used in the DOE, a formulation with “intermediate” concentrations in additives (2 wt% PLL-g-PNIPAM and 0.03 wt% Krytox) was characterized and added to the data as an example of a “central” point in the experimental domain defined by the DOE (see Figure 8.1.A). The mean droplet diameter of this formulation was measured for consecutive days to follow the stability of the emulsion over time (see Figure 8.1.B). Starting from a value of 100 nm just after emulsification, the average droplet diameter increases to 200 nm after one day and seems to stabilize at this value afterwards. Note that a standard deviation of around 100 nm was determined, suggesting a high degree of polydispersity (what was confirmed by a relatively high polydispersity index around 0.2 in the size distribution of a given
sample) and probably a partial destabilization of the emulsion over time. This process can also be pointed
out regarding the progressive decrease of the scattering intensity of the samples over time (see Figure
8.1.B). This result can be attributed to a less efficient redispersion of the water droplets in the oil over time,
suggesting a progressive – but not drastic – destabilization of the emulsion.
References

(1) Khurana, A.; Allawadhi, P.; Khurana, I.; Allwadhi, S.; Weiskirchen, R.; Banothu, A. K.; Chhabra, D.; Joshi, K.; Bharani, K. K. Role of Nanotechnology behind the Success of mRNA Vaccines for COVID-19. *Nano Today* **2021**, 38, 101142. https://doi.org/10.1016/j.nantod.2021.101142.

(2) Kim, S. B.; Ito, Y.; Torimura, M. Bioluminescent Capsules for Live-Cell Imaging. *Bioconjug. Chem.* **2012**, *23* (11), 2221–2228. https://doi.org/10.1021/bc300323x.

(3) Liang, K.; Gunawan, S. T.; Richardson, J. J.; Such, G. K.; Cui, J.; Caruso, F. Endocytic Capsule Sensors for Probing Cellular Internalization. *Adv. Healthc. Mater.* **2014**, *3* (10), 1551–1554. https://doi.org/10.1002/adhm.201400139.

(4) Jeyaraman, J.; Shukla, A.; Sivakumar, S. Targeted Stealth Polymer Capsules Encapsulating Ln3+-Doped LaVO4 Nanoparticles for Bioimaging Applications. *ACS Biomater. Sci. Eng.* **2016**, *2* (8), 1330–1340. https://doi.org/10.1021/acsbiomaterials.6b00252.

(5) Martina, M. S.; Fortin, J. P.; Ménager, C.; Clément, O.; Barratt, G.; Grabielle-Madelmont, C.; Gazeau, F.; Cabuil, V.; Lesieur, S. Generation of Superparamagnetic Liposomes Revealed as Highly Efficient MRI Contrast Agents for in Vivo Imaging. *J. Am. Chem. Soc.* **2005**, *127* (30), 10676–10685. https://doi.org/10.1021/ja0516460.

(6) Jagielski, N.; Sharma, S.; Hombach, V.; Mailänder, V.; Rasche, V.; Landfester, K. Nanocapsules Synthesized by Miniemulsion Technique for Application as New Contrast Agent Materials. *Macromol. Chem. Phys.* **2007**, *208* (19–20), 2229–2241. https://doi.org/10.1002/macp.200700254.

(7) Pisani, E.; Tsapis, N.; Galaz, B.; Santin, M.; Berti, R.; Taulier, N.; Kurtisovski, E.; Lucidarme, O.; Ourevitch, M.; Doan, B. T.; Beloeil, J. C.; Gillet, B.; Urbach, W.; Bridal, S. L.; Fattal, E. Perfluorooctyl Bromide Polymeric Capsules as Dual Contrast Agents for Ultrasonography and Magnetic Resonance Imaging. *Adv. Funct. Mater.* **2008**, *18* (19), 2963–2971. https://doi.org/10.1002/adfm.200800454.

(8) Cheng, Z.; Al Zaki, A.; Jones, I. W.; Hall, H. K.; Aspinwall, C. A.; Tsourkas, A. Stabilized Porous Liposomes with Encapsulated Gd-Labeled Dextran as a Highly Efficient MRI Contrast Agent. *Chem. Commun.* **2014**, *50* (19), 2502–2504. https://doi.org/10.1039/c3cc48939f.

(9) De Cock, L. J.; De Koker, S.; De Geest, B. G.; Grooten, J.; Vervaet, C.; Remon, J. P.; Sukhorukov, G. B.; Antipina, M. N. Polymeric Multilayer Capsules in Drug Delivery. *Angew. Chemie - Int. Ed.* **2010**, *49* (39), 6954–6973. https://doi.org/10.1002/anie.200906266.

(10) Hu, X.; Zhang, Y.; Xie, Z.; Jing, X.; Bellotti, A.; Gu, Z. Stimuli-Responsive Polymersomes for Biomedical Applications. *Biomacromolecules* **2017**, *18* (3), 649–673.
https://doi.org/10.1021/acs.biomac.6b01704.

(11) Bordat, A.; Boissenot, T.; Nicolas, J.; Tsapis, N. Thermoresponsive Polymer Nanocarriers for Biomedical Applications. *Adv. Drug Deliv. Rev.* **2019**, *138*, 167–192.
https://doi.org/10.1016/j.amdr.2018.10.005.

(12) Sun, T.; Zhang, Y. S.; Pang, B.; Hyun, D. C.; Yang, M.; Xia, Y. Engineered Nanoparticles for Drug Delivery in Cancer Therapy. *Angew. Chemie - Int. Ed.* **2014**, *53* (46), 12320–12364.
https://doi.org/10.1002/anie.201403036.

(13) Senapati, S.; Mahanta, A. K.; Kumar, S.; Maiti, P. Controlled Drug Delivery Vehicles for Cancer Treatment and Their Performance. *Signal Transduct. Target. Ther.* **2018**, *3* (1), 1–19.
https://doi.org/10.1038/s41392-017-0004-3.

(14) Ke, W.; Li, J.; Mohammed, F.; Wang, Y.; Tou, K.; Liu, X.; Wen, P.; Kinoh, H.; Anraku, Y.; Chen, H.; Kataoka, K.; Ge, Z. Therapeutic Polymersome Nanoreactors with Tumor-Specific Activable Cascade Reactions for Cooperative Cancer Therapy. *ACS Nano* **2019**, *13* (2), 2357–2369.
https://doi.org/10.1021/acsnano.8b09082.

(15) Saunders, B. R.; Fraylich, M.; Alexander, C.; Wang, W.; Liang, H.; Cheikh Al Ghanami, R.; Hamilton, L.; Shakesheff, K. M. Biodegradable Thermoresponsive Microparticle Dispersions for Injectable Cell Delivery Prepared Using a Single-Step Process. *Adv. Mater.* **2009**, *21* (18), 1809–1813.
https://doi.org/10.1002/adma.200800382.

(16) Facca, S.; Cortez, C.; Mendoza-palomares, C.; Messadeq, N.; Dierich, A.; Johnston, A. P. R.; Mainard, D. Active Multilayered Capsules for in Vivo Bone Formation. **2009**.
https://doi.org/10.1073/pnas.0908531107.

(17) Zheng, Z.; Wang, H.; Li, J.; Shi, Q.; Cui, J.; Sun, T.; Huang, Q.; Fukuda, T. 3D Construction of Shape-Controllable Tissues through Self-Bonding of Multicellular Microcapsules. *ACS Appl. Mater. Interfaces* **2019**, *11* (26), 22950–22961. https://doi.org/10.1021/acsnano.9b05108.

(18) Wang, F.; Xiao, J.; Chen, S.; Sun, H.; Yang, B.; Jiang, J.; Zhou, X.; Du, J. Polymer Vesicles: Modular Platforms for Cancer Theranostics. *Adv. Mater.* **2018**, *30* (17).
https://doi.org/10.1002/adma.201705674.

(19) Delcassian, D.; Luzhansky, I.; Spanoudaki, V.; Bochenek, M.; McGladrigan, C.; Nguyen, A.; Norcross, S.; Zhu, Y.; Shan, C. S.; Hausser, R.; Shakesheff, K. M.; Langer, R.; Anderson, D. G. Magnetic Retrieval of Encapsulated Beta Cell Transplants from Diabetic Mice Using Dual-Function MRI Visible and Retrievable Microcapsules. *Adv. Mater.* **2020**, *32* (16).
https://doi.org/10.1002/adma.201904502.

(20) Shakeri, S.; Ashrafizadeh, M.; Zarrabi, A.; Roghanian, R.; Afshar, E. G.; Pardakhty, A.; Mohammadinejad, R.; Kumar, A.; Thakur, V. K. Multifunctional Polymeric Nanoplatforms for
Brain Diseases Diagnosis, Therapy and Theranostics. *Biomedicines* **2020**, *8* (1).
https://doi.org/10.3390/biomedicines8010013.

(21) Chiang, W. H.; Huang, W. C.; Chang, C. W.; Shen, M. Y.; Shih, Z. F.; Huang, Y. F.; Lin, S. C.; Chiu, H. C. Functionalized Polymersomes with Outlayered Polyelectrolyte Gels for Potential Tumor-Targeted Delivery of Multimodal Therapies and MR Imaging. *J. Control. Release* **2013**, *168* (3), 280–288. https://doi.org/10.1016/j.jconrel.2013.03.029.

(22) Schoebitz, M.; López, M. D.; Roldán, A. Bioencapsulation of Microbial Inoculants for Better Soil-Plant Fertilization. A Review. *Agron. Sustain. Dev.* **2013**, *33* (4), 751–765. https://doi.org/10.1007/s13593-013-0142-0.

(23) Camara, M. C.; Campos, E. V. R.; Monteiro, R. A.; Do Espirito Santo Pereira, A.; De Freitas Proença, P. L.; Fraceto, L. F. Development of Stimuli-Responsive Nano-Based Pesticides: Emerging Opportunities for Agriculture. *J. Nanobiotechnology* **2019**, *17* (1), 1–19. https://doi.org/10.1186/s12951-019-0533-8.

(24) Blanco-Padilla, A.; Soto, K. M.; Hernández Iturriaga, M.; Mendoza, S. Food Antimicrobials Nanocarriers. *Sci. World J.* **2014**, *2014*. https://doi.org/10.1155/2014/837215.

(25) Ezhilarasi, P. N.; Karthik, P.; Chhanwal, N.; Anandharamakrishnan, C. Nanoencapsulation Techniques for Food Bioactive Components: A Review. *Food Bioprocess Technol.* **2013**, *6* (3), 628–647. https://doi.org/10.1007/s11947-012-0944-0.

(26) Lohani, A.; Verma, A.; Joshi, H.; Yadav, N.; Karki, N. Nanotechnology-Based Cosmeceuticals. *2018*, *2014*.

(27) He, L.; Hu, J.; Deng, W. Preparation and Application of Flavor and Fragrance Capsules. *Polym. Chem.* **2018**, *9* (40), 4926–4946. https://doi.org/10.1039/c8py00863a.

(28) Vimalanandan, A.; Lv, L. P.; Tran, T. H.; Landfester, K.; Crespy, D.; Rohwerder, M. Redox-Responsive Self-Healing for Corrosion Protection. *Adv. Mater.* **2013**, *25* (48), 6980–6984. https://doi.org/10.1002/adma.201302989.

(29) Zhu, D. Y.; Rong, M. Z.; Zhang, M. Q. Self-Healing Polymeric Materials Based on Microencapsulated Healing Agents: From Design to Preparation. *Prog. Polym. Sci.* **2015**, *49–50*, 175–220. https://doi.org/10.1016/j.progpolymsci.2015.07.002.

(30) Salunkhe, P. B.; Shembekar, P. S. A Review on Effect of Phase Change Material Encapsulation on the Thermal Performance of a System. *Renew. Sustain. Energy Rev.* **2012**, *16* (8), 5603–5616. https://doi.org/10.1016/j.rser.2012.05.037.

(31) Shchukin, D. G.; Möhwald, H. Urea Photosynthesis inside Polyelectrolyte Capsules: Effect of Confined Media. *Langmuir* **2005**, *21* (12), 5582–5587. https://doi.org/10.1021/la050429+.

(32) Küchler, A.; Yoshimoto, M.; Luginbühl, S.; Mavelli, F.; Walde, P. Enzymatic Reactions in Confined
Environments. *Nat. Nanotechnol.* **2016**, *11* (5), 409–420. https://doi.org/10.1038/nnano.2016.54.

(33) Musyanovych, A.; Mailänder, V.; Landfester, K. Miniemulsion Droplets as Single Molecule Nanoreactors for Polymerase Chain Reaction. *Biomacromolecules* **2005**, *6* (4), 1824–1828. https://doi.org/10.1021/bm050084+.

(34) Comellas-Aragonès, M.; Engelkamp, H.; Claessen, V. I.; Sommerdijk, N. A. J. M.; Rowan, A. E.; Christianen, P. C. M.; Maan, J. C.; Verduin, B. J. M.; Cornelissen, J. J. L. M.; Nolte, R. J. M. A Virus-Based Single-Enzyme Nanoreactor. *Nat. Nanotechnol.* **2007**, *2* (10), 635–639. https://doi.org/10.1038/nnano.2007.299.

(35) Dong, R.; Liu, W.; Hao, J. Soft Vesicles in the Synthesis of Hard Materials. *Acc. Chem. Res.* **2012**, *45* (4), 504–513. https://doi.org/10.1021/ar200124g.

(36) Cacace, D. N.; Rowland, A. T.; Stapleton, J. J.; Dewey, D. C.; Keating, C. D. Aqueous Emulsion Droplets Stabilized by Lipid Vesicles as Microcompartments for Biomimetic Mineralization. *Langmuir* **2015**, *31* (41), 11329–11338. https://doi.org/10.1021/acs.langmuir.5b02754.

(37) Tian, H.; Liang, J.; Liu, J. Nanoengineering Carbon Spheres as Nanoreactors for Sustainable Energy Applications. *Adv. Mater.* **2019**, *31* (50). https://doi.org/10.1002/adma.201903886.

(38) De Martino, M. T.; Abdelmohsen, L. K. E. A.; Rutjes, F. P. J. T.; Van Hest, J. C. M. Nanoreactors for Green Catalysis. *Beilstein J. Org. Chem.* **2018**, *14* (2), 716–733. https://doi.org/10.3762/bjoc.14.61.

(39) Marguet, M.; Bonduelle, C.; Lecommandoux, S. Multicompartmentalized Polymeric Systems: Towards Biomimetic Cellular Structure and Function. *Chem. Soc. Rev.* **2013**, *42* (2), 512–529. https://doi.org/10.1039/c2cs35312a.

(40) Peters, R. J. R. W.; Marguet, M.; Marais, S.; Fraaije, M. W.; van Hest, J. C. M.; Lecommandoux, S. Cascade Reactions in Multicompartmentalized Polymersomes. *Angew. Chemie* **2014**, *126* (1), 150–154. https://doi.org/10.1002/ange.201308141.

(41) Schoonen, L.; Van Hest, J. C. M. Compartmentalization Approaches in Soft Matter Science: From Nanoreactor Development to Organelle Mimics. *Adv. Mater.* **2016**, *28* (6), 1109–1128. https://doi.org/10.1002/adma.201502389.

(42) Kurihara, K.; Tamura, M.; Shohda, K. I.; Toyota, T.; Suzuki, K.; Sugawara, T. Self-Reproduction of Supramolecular Giant Vesicles Combined with the Amplification of Encapsulated DNA. *Nat. Chem.* **2011**, *3* (10), 775–781. https://doi.org/10.1038/nchem.1127.

(43) Tian, L.; Li, M.; Patil, A. J.; Drinkwater, B. W.; Mann, S. Artificial Morphogen-Mediated Differentiation in Synthetic Protocells. *Nat. Commun.* **2019**, *10* (1), 1–13. https://doi.org/10.1038/s41467-019-11316-4.

(44) Berhanu, S.; Ueda, T.; Kuruma, Y. Artificial Photosynthetic Cell Producing Energy for Protein
Synthesis. *Nat. Commun.* **2019**, *10* (1). https://doi.org/10.1038/s41467-019-09147-4.

(45) Rodríguez-Arco, L.; Kumar, B. V. V. S. P.; Li, M.; Patil, A. J.; Mann, S. Modulation of Higher-Order Behaviour in Model Protocell Communities by Artificial Phagocytosis. *Angew. Chemie - Int. Ed.* **2019**, *58* (19), 6333–6337. https://doi.org/10.1002/anie.201901469.

(46) Hindley, J. W.; Zheleva, D. G.; Elani, Y.; Charalambous, K.; Barter, L. M. C.; Booth, P. J.; Bevan, C. L.; Law, R. V.; Ces, O. Building a Synthetic Mechanosensitive Signaling Pathway in Compartmentalized Artificial Cells. *Proc. Natl. Acad. Sci. U. S. A.* **2019**, *116* (34), 16711–16716. https://doi.org/10.1073/pnas.1903500116.

(47) Wang, X.; Tian, L.; Du, H.; Li, M.; Mu, W.; Drinkwater, B. W.; Han, X.; Mann, S. Chemical Communication in Spatially Organized Protocell Colonies and Protocell/Living Cell Micro-Arrays. *Chem. Sci.* **2019**, *10* (41), 9446–9453. https://doi.org/10.1039/c9sc04522h.

(48) Qiao, Y.; Li, M.; Qiu, D.; Mann, S. Response-Retaliation Behavior in Synthetic Protocell Communities. *Angew. Chemie* **2019**, *131* (49), 17922–17927. https://doi.org/10.1002/ange.201909313.

(49) Villringer, S.; Madl, J.; Sych, T.; Manner, C.; Imberty, A.; Römer, W. Lectin-Mediated Protocell Crosslinking to Mimic Cell-Cell Junctions and Adhesion. *Sci. Rep.* **2018**, *8* (1), 1–11. https://doi.org/10.1038/s41598-018-20230-6.

(50) Bekir, M.; Jelken, J.; Jung, S. H.; Pich, A.; Pacholski, C.; Kopyshev, A.; Santer, S. Dual Responsiveness of Microgels Induced by Single Light Stimulus. *Appl. Phys. Lett.* **2021**, *118* (9). https://doi.org/10.1063/5.0036376.

(51) Nijemeisland, M.; Abdelmohsen, L. K. E. A.; Huck, W. T. S.; Wilson, D. A.; Van Hest, J. C. M. A Compartmentalized Out-of-Equilibrium Enzymatic Reaction Network for Sustained Autonomous Movement. *ACS Cent. Sci.* **2016**, *2* (11), 843–849. https://doi.org/10.1021/acscentsci.6b00254.

(52) Xu, Z.; Hueckel, T.; Irvine, W. T. M.; Sacanna, S. Transmembrane Transport in Inorganic Colloidal Cell-Mimics. *Nature* **2021**, *597* (7875), 220–224. https://doi.org/10.1038/s41586-021-03774-y.

(53) Reis, C. P.; Ribeiro, A. J.; Houng, S.; Veiga, F.; Neufeld, R. J. Nanoparticulate Delivery System for Insulin: Design, Characterization and in Vitro/in Vivo Bioactivity. *Eur. J. Pharm. Sci.* **2007**, *30* (5), 392–397. https://doi.org/10.1016/j.ejps.2006.12.007.

(54) Tayalia, P.; Mooney, D. J. Controlled Growth Factor Delivery for Tissue Engineering. *Adv. Mater.* **2009**, *21* (32–33), 3269–3285. https://doi.org/10.1002/adma.200900241.

(55) Slastnikova, T. A.; Ulasov, A. V.; Rosenkranz, A. A.; Sobolev, A. S. Targeted Intracellular Delivery of Antibodies: The State of the Art. *Front. Pharmacol.* **2018**, *9* (OCT), 1–21. https://doi.org/10.3389/fphar.2018.01208.

(56) Frézard, F.; Silva-Barcellos, N. M.; dos Santos, R. A. S. A Novel Approach Based on