# Design of stimuli-responsive polymer capsules by interfacial complexation

Lucas Sixdenier

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## To cite this version:

Lucas Sixdenier. Design of stimuli-responsive polymer capsules by interfacial complexation. Polymers. Sorbonne Université, 2022. English. NNT: 2022SORUS126. tel-03735777v2

HAL Id: tel-03735777

https://theses.hal.science/tel-03735777v2

Submitted on 21 Jul 2022

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Sorbonne Université

École doctorale 388

Chimie Physique et Chimie Analytique de Paris Centre

École Normale Supérieure – Département de Chimie

Laboratoire PASTEUR – Pôle de Chimie Physique et Biologique de la Matière Vivante

# Design of stimuli-responsive polymer capsules by interfacial complexation

Par Lucas SIXDENIER

Thèse de doctorat en Chimie Physique

Dirigée par Emmanuelle MARIE

Présentée et soutenue publiquement le 28 janvier 2022 devant un jury composé de :

M. LECOMMANDOUX Sébastien

Mme MARIE Emmanuelle

Mme RAVAINE Valérie

Mme RIEGER Jutta

M. TRIBET Christophe

M. TSAPIS Nicolas

M. ZHAO Yue

Professeur

Directrice de recherche

Professeure

Directrice de recherche

Directeur de recherche

Directeur de recherche

Professeur

Examinateur

Directrice de thèse

Rapportrice

Examinatrice

Membre invité

Rapporteur

Membre invité

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Remerciements

Voici venu le temps des rires et des chants, mais aussi et surtout celui des remerciements. Et si une thèse ce n'est pas exactement tous les jours le printemps ou le pays joyeux des enfants heureux (cette métaphore filée a beaucoup trop duré), je dois avouer qu'elle fut pour moi une période particulièrement réjouissante. À travers les quelques mots de remerciements qui suivent, j'espère rendre un juste hommage à mon entourage, personnel comme professionnel, pour tout ce qu'il m'a apporté durant ces trois belles années.

Je voudrais d'abord remercier les rapporteurs(trices) et examinateurs(trices) de mon jury, Sébastien Lecommandoux, Valérie Ravaine, Jutta Rieger et Nicolas Tsapis, ainsi que notre collaborateur Yue Zhao, d'avoir accepté d'évaluer cette thèse. Vos remarques, questions et autres suggestions m'ont été précieuses pour peaufiner mon travail et imaginer son développement futur. Nos discussions lors de la soutenance ont été particulièrement plaisantes et constructives, je vous remercie d'avoir été un jury aussi impliqué et bienveillant !

Un grand, un immense, que dis-je, un gigantesque merci Emmanuelle d'avoir accepté de diriger cette thèse, ainsi que le stage de Master qui l'a précédée. Merci pour la confiance que tu m'as accordée si rapidement et que tu as maintenue durant ces quatre années passées à tes côtés. Nous avons construit ce projet ensemble, en partant de pas grand-chose, si ce n'est rien, et je suis assez content (ce qui pour moi est déjà pas si mal) de ce qu'on a pu produire ! Ces années n'ont fait que renforcer mon goût pour la recherche, non seulement grâce à l'environnement scientifique que tu m'as offert mais aussi, et surtout, grâce à la complicité qu'on a pu nouer tous les deux et qui m'a convaincu que le monde de la recherche était avant tout fait de relations humaines. Je me rends compte aujourd'hui de la chance que j'ai eu d'avoir une directrice de thèse comme toi. Je pense qu'on ne manquera pas de garder contact à l'avenir, compte sur moi pour cela en tout cas !

Un immense merci également à Christophe. Bien plus qu'un chef d'équipe, tu as été un véritable mentor sur le plan scientifique. Merci pour tous tes conseils, avis et idées sans lesquels cette thèse n'aurait définitivement pas eu la même consistance. J'ai énormément appris à tes côtés et j'espère faire honneur à la rigueur que tu m'as apportée dans la suite de mon parcours.

Je souhaitais également remercier chaleureusement les collaborateurs avec qui j'ai eu la chance de travailler durant cette thèse : Yue Zhao et Amélie Augé pour la partie UCST, Guillaume Baffou pour la partie thermoplasmonique, Wladimir Urbach et Chloë Thimonier pour les parties tension de surface et microfluidique, et Atef Asnacios pour les manips préliminaires de micro-rhéologie. J'espère qu'on aura l'occasion de travailler à nouveau ensemble à l'avenir !
Merci à tous les permanents du pôle CPBMV pour toutes les discussions (plus ou moins professionnelles) qu'on a pu partager, et pour tous vos conseils divers et variés. L'avantage d'évoluer dans un groupe aussi éclectique est qu'on finit toujours par avoir une solution à nos problèmes ! Merci à Ludovic Jullien pour sa constante bienveillance et pour avoir partagé son expérience du monde de la recherche qui me sera pour sûr éclairante. Un merci tout particulier à Thomas Le Saux pour ses connaissances scientifiques et pratiques dans tous les domaines m'ont toujours à la fois fasciné et bien dépanné (et aussi pour nos discussions moins sérieuses sur les vieilleries en tout genre, films, musiques et sketchs, dont personne d'autre n'avait la référence). Merci à Zoher Gueroui pour ses conseils en microscopie et pour avoir partagé ton expérience new-yorkaise qui n'a fait que me convaincre un peu plus d'aller y passer quelque temps moi aussi. Merci à Isabelle Aujard, pour sa présence, sa gentillesse, et sa patience surtout. Je ne vais même pas parler de manips, de commandes ou de grand ménage printanier, ce n'est rien à côté de toutes les discussions plus ou moins sérieuses qu'on a partagé durant tout ce temps et qui ont rendu bien plus agréables ces longues heures de labo. Merci à Marie-Aude Plamont pour toutes ces fois où je suis venu l'embêter, parfois juste pour le plaisir, en attendant de sa part un retour de bâton bien mérité.

Je souhaite également remercier les membres des autres pôles du laboratoire PASTEUR et même plus largement du département de Chimie de l'ENS, pour leur aide ponctuelle qui me fût si précieuse. En particulier merci à Ayako Amada pour le SEM, Nicolas Delsuc pour le lyoph, Jacques Fattaccioli pour le Zeta, Sergii Rudiuk pour l'AFM. Et puis merci également à Rodolphe Vuilleumier, directeur du laboratoire puis du département, avec qui j'ai eu la chance et le plaisir de partager de nombreuses séances d'enseignement à Jussieu. Merci également au personnel administratif qui a toujours su faciliter ma vie au quotidien, merci à Anne, Delphine, Elza, Nathalie, Pauline pour toutes ces fois où je suis venu solliciter leur aide.

Merci à tous les doctorant.es, postdocs, stagiaires que j'ai côtoyé.es au labo et en dehors durant ces quatre années. La liste est longue et j'en oublierai forcément quelques-un.es, veuillez me pardonner. Merci à Aliénor, Alison, Annamaria, Antoine, Baptiste, Bea, Chems, Chloë, Damien, Emilie, Enzo, Fayrouz, Julien, Kätlin, Laura, Léonard, Marc, Pauline, Raja, Tiphaine, Yoan, Yuriy. A special thanks to Alison, Annamaria, Bea and Kätlin, the four fantastic postdocs with whom I spent a lot of time in the lab, in the office or outside. I will miss you so much (and already do actually)! Merci Fayrouz d'avoir accepté de faire ton stage avec moi et d'avoir gardé ta constante bonne humeur malgré des manips particulièrement délicates (et qui ont finalement bien servi à orienter ma thèse !). Je garde le meilleur pour la fin, le « noyau dur » comme on l'appelle : Audrey, Fanny, Hela, Louise et Mary. À ce stade on ne parle plus de collègues, mais de véritables amies. Cette thèse n'aurait évidemment pas été la même sans vous, elle n'aurait pas eu ce petit arrière-goût persistant de rire et de joie. Merci pour tous ces goûters, ces soirées endiablées (devenues de plus en plus pépères avec l'âge), ces sorties, ces vacances. Merci de m'avoir supporté tel que je suis, sans trop souvent lever les yeux au ciel, enfin je crois... J'espère que tous ces moments ne feront que se multiplier à l'avenir, car je dois vous avouer que je ne saurais plus m'en passer.
Une dédicace toute particulière à toi OD, ma fidèle acolyte depuis le tout premier jour de cette aventure normalienne. Je ne sais pas vraiment comment tu as fait pour me supporter tout ce temps sans déposer une seule main courante (je t'achèterai une madeleine pour te remercier). J'espère qu'on s'est suffisamment côtoyé pour avoir un peu déteint l'un sur l'autre, moi étant devenu plus chou et toi plus méchante.

Merci également à tous mes ami.es de lycée, de prépa et de PC, qui m'ont accompagné, de près ou de loin, durant ces trois ans. Pourquoi se demander à quoi rime la vie alors que la réponse est si simple : elle rime avec passer du temps avec vous. La liste est trop longue pour être énumérée mais sachez qu'en écrivant ces mots, ce sont tous les souvenirs partagés ensemble qui me reviennent à l'esprit et me rendent plus qu'heureux.

Je vais tout de même m'oser à faire quelques dédicaces ciblées. Merci Simon et Théo pour toutes ces vacheries balancées depuis toutes ces années mais qui au fond, on le sait, cachent une amitié toute dégoulinante d'affection. Merci à mes trois colocataires, que dis-je, à mes trois fratés, avec qui je partage mon quotidien depuis maintenant un bon bout de temps. Merci Romain de m'avoir fait rencontrer la personne la plus drôle qui existe sur cette planète : toi. Et puis surtout pour le plus beau cadeau que vous auriez pu me faire, Juliette et toi, à savoir faire partie de la famille. Merci Gwen, pour tous ces fous rires, ces jeux de mots (plus ou moins validés), ces soirées blind test (plus ou moins arrosées), et puis toutes ces vacances (plus ou moins glorieuses), passées, présentes et surtout à venir ! Merci Vivi pour toutes ces années de vie commune qui ont sûrement affecté de manière irréversible notre santé mentale mais qui en tout cas n'ont fait qu'emplir mon cœur de joie. C'est simple, j'ai passé la moitié de ma vie adulte en coloc avec toi, et j'ai l'impression qu'une autre vie n'est tout simplement pas envisageable. Mé se ki é cool vivi c kon va vivre tou lé 2 le reve amérikain a la nouvel yorke dé zéta zuni damérike, sa va etre super de mangé dé donut ensemble. Et enfin promis Clément je passerai te voir à Toulouse. Dans trois ou quatre ans peut-être. Un conseil, attends assis.

Enfin, la vie ne serait rien sans un peu d'amour. Merci Axelle, tu as rendu cette période de thèse bien plus que supportable en rendant ma vie bien plus que belle. Rien ne vaut les mots sublimes de Julien Gracq pour le dire :
> « Il se demandait s'il l'avait aimée. C'était moins et mieux : il n'y avait eu de place que pour elle. »

Et enfin, merci à ma famille, mes grands-parents, mes parents, mon frère, qui n'ont pas forcément toujours suivi ce que je faisais (je les comprends) mais qui ont toujours suivi avec attention ce que je devenais. Merci de m'avoir laissé tracer ma route, à ma guise, en me maintenant les pieds sur terre et la tête sur les épaules. Merci d'avoir fait la personne que je suis devenue. Je vous dois tout. J'espère vous avoir rendu aussi fiers que je vous suis reconnaissant.

Allez, sans transition, matière molle.
# Table of Content

<table><tr><td>1</td><td>Chapter 1 : General introduction</td><td>17</td></tr><tr><td>2</td><td>Chapter 2 : State of the art and positioning of the present work</td><td>25</td></tr><tr><td>2.1</td><td>The specific challenge of protein encapsulation and release</td><td>27</td></tr><tr><td>2.2</td><td>Micro- and nanocontainers developed for the encapsulation of hydrophilic payloads</td><td>29</td></tr><tr><td>2.3</td><td>Remote control of payload release from microcontainers</td><td>48</td></tr><tr><td>2.4</td><td>Objective of the PhD work</td><td>60</td></tr><tr><td>3</td><td>Chapter 3 : Formation of stimuli-responsive polymer shells by interfacial complexation: poly(N-isopropylacrylamide) shells as a first example</td><td>63</td></tr><tr><td>3.1</td><td>Introduction</td><td>65</td></tr><tr><td>3.2</td><td>Evidence for interfacial complexation of PLL comb-like derivatives in W/FO emulsions</td><td>67</td></tr><tr><td>3.3</td><td>Microscopy imaging of emulsions</td><td>75</td></tr><tr><td>3.4</td><td>Characterization of PLL-g-PNIPAM shells formed upon interfacial complexation</td><td>80</td></tr><tr><td>3.5</td><td>Evaluation of the thermoresponsive behavior of PLL-g-PNIPAM shells</td><td>90</td></tr><tr><td>3.6</td><td>Encapsulation of a model protein in the aqueous core</td><td>107</td></tr><tr><td>3.7</td><td>Conclusion</td><td>109</td></tr><tr><td>4</td><td>Chapter 4 : Formation of polymer-nanoparticles mixed shells by interfacial complexation</td><td>113</td></tr><tr><td>4.1</td><td>Introduction</td><td>115</td></tr><tr><td>4.2</td><td>Interactions between PLL derivatives and nanoparticles</td><td>117</td></tr><tr><td>4.3</td><td>Design of “composite” polymer shells by thermo-enhanced interfacial complexation</td><td>123</td></tr><tr><td>4.4</td><td>Thermoplasmonic effects in AuNPs-loaded droplets</td><td>128</td></tr><tr><td>4.5</td><td>Conclusion</td><td>156</td></tr><tr><td>5</td><td>Chapter 5 : UCST-type polymer capsules formed by interfacial complexation</td><td>159</td></tr><tr><td>5.1</td><td>Introduction</td><td>161</td></tr><tr><td>5.2</td><td>Synthesis of PLL-g-PAAMAN polymer and formation of PLL-g-PAAMAN shells by interfacial complexation</td><td>162</td></tr></table>
<table>
  <tr>
    <td>5.3</td>
    <td>Thermoresponsive behavior of PLL-g-PAAMAN shells</td>
    <td>168</td>
  </tr>
  <tr>
    <td>5.4</td>
    <td>Towards the formation of UCST composite capsules</td>
    <td>173</td>
  </tr>
  <tr>
    <td>5.5</td>
    <td>Encapsulation of a model protein</td>
    <td>176</td>
  </tr>
  <tr>
    <td>5.6</td>
    <td>Conclusion</td>
    <td>178</td>
  </tr>
  <tr>
    <td>6</td>
    <td>Chapter 6: Conclusion and outlooks</td>
    <td>180</td>
  </tr>
  <tr>
    <td>6.1</td>
    <td>Conclusion</td>
    <td>182</td>
  </tr>
  <tr>
    <td>6.2</td>
    <td>Outlooks</td>
    <td>185</td>
  </tr>
  <tr>
    <td>7</td>
    <td>Appendix A: Materials, instrumentation, and experimental protocols</td>
    <td>195</td>
  </tr>
  <tr>
    <td>8</td>
    <td>Appendix B: Influence of the formulation of water-in-oil emulsions on the size and stability of emulsion droplets</td>
    <td>211</td>
  </tr>
  <tr>
    <td>9</td>
    <td>References</td>
    <td>219</td>
  </tr>
</table>
# List of abbreviations and symbols

## Abbreviations

**ANS**: 8-anilinonaphthalene-1-sulfonic acid

**AuNP(s)**: gold nanoparticle(s)

**CarboxyNP(s)**: carboxylate-modified polystyrene nanoparticle(s)

**CMC**: Critical micellar concentration

**Coum**: Coumarin

**CTA**: Chain transfer agent

**DLS**: Dynamic light scattering

**DP<sub>n</sub>**: Degree of polymerization

**eGFP**: enhanced Green Fluorescent Protein

**FC-40, FC-70**: Fluorinert™ fluorinated oil

**FO**: fluorinated oil

**FRAP**: Fluorescence recovery after photobleaching

**GSPM**: Grating-shadow phase microscopy

**GUV(s)**: Giant unilamellar vesicle(s)

**LbL**: Layer-by-layer

**LCST**: Lower critical solution temperature

**NeutravNP(s)**: NeutrAvidin-modified polystyrene nanoparticle(s)

**NHS**: N-hydroxysuccinimide

**NP(s)**: Nanoparticle(s)

**OPD**: Optical path difference

**PAAMAN**: Poly(acrylamide-co-acrylonitrile)

**PEG**: Poly(ethylene glycol)

**PISA**: Polymerization-induced self-assembly

**PLL**: Poly(L-lysine)

**PNIPAM**: Poly(N-isopropylacrylamide)

**PVP**: Poly(vinylpyrrolidone)
**QPM**: Quantitative phase microscopy

**RAFT**: Reversible addition-fragmentation chain-transfer

**Rho**: Rhodamine

**SEM**: Scanning electron microscopy

**SUV(s)**: Small unilamellar vesicle(s)

**UCST**: Upper critical solution temperature

**W/FO**: water-in-fluorinated oil (emulsion)

**W/O**: water-in-oil (emulsion)

**W/W**: water-in-water (emulsion)

## Symbols

**C**: concentration

**d**: droplet diameter

**e**: droplet flattening thickness

**I**: fluorescence intensity or laser irradiance

**$M_f$**: polymer mobile fraction

**n**: refractive index

**Q**: heat power

**P**: light power

**R**: droplet radius

**T**: temperature

**V**: droplet volume

**$\delta$**: optical path difference

**$\Delta T$**: Temperature rise

**$\kappa$**: thermal conductivity

**$\lambda$**: light wavelength

**$\sigma_{abs}$**: absorption cross-section
# Chapter 1

## General introduction
In everyday life, packing objects in containers, moving them, and unpacking them wherever and whenever we want is a common thing. Transposing this process at the micro- or nanoscale has been a crucial, albeit challenging, issue in many application fields, the most important one being probably the vectorization of drugs or bioactive substances for pharmacological and medical purposes. Mastering the steps of encapsulation and release of molecules or nanomaterials is much more difficult to achieve at these small scales since a direct manipulation of such tiny objects is out of the reach of human hands. As a consequence, alternative levers have been sought to remotely control the packing and unpacking of molecular payloads in micro- and nanocarriers.* For the last 20 years, the scientific research relative to the encapsulation of materials at microscales has skyrocketed due to the versatility of the approach and the diversification of the potential applications (see **Figure 1.1**).

<figure><img src="image_7.png" /><figcaption>**Figure 1.1.** Annual number of publications (articles, reviews, and book chapters) referring to “encapsulation” in the fields of Chemistry, Materials and Life Science from 1960 to current 2021 (Source: Scopus, Nov. 2021).</figcaption></figure>

The recent large-scale development of mRNA vaccines to fight against COVID-19 has been an illustrative example of recent achievements made in the field of microencapsulation. The effective encapsulation of negatively-charged mRNA in lipid-based nanoparticles has been enabled by a precise optimization of the composition of the nanocontainers.¹ This example illustrates the general issue of multiscale optimization of the formulation to enhance the efficiency of both steps of encapsulation and delivery. It is all a matter of (i) chemical design at the molecular scale, (ii) colloidal stability at the mesoscopic scale and

<footnote>* In this thesis, the discussion will be restricted to the physico-chemistry relative to the design and characterization of colloidal systems (with typical sizes between 10 nm and 100 µm). As a consequence, the terms relative to “encapsulation” (including “capsules”, “containers”, “compartments”, “carriers”) may be used without specifying the prefixes “micro” or “nano”.</footnote>
(iii) spatiotemporal control of the delivery at the scale of the target, while overall preserving the activity of interest of the encapsulated payload.

mRNA vaccines are only one example among the tremendous variety of encapsulation systems that have been explored so far for potential medical and biotechnological applications such as bioimaging (e.g. via the encapsulation of fluorescent reporters,²⁻⁴ or contrast agents for magnetic resonance imaging⁵⁻⁸), in-situ release or production of therapeutic agents⁹⁻¹¹ (with a particular attention focused on cancer therapy,¹²⁻¹⁴ see **Figure 1.2**), tissue engineering and repairing,¹⁵⁻¹⁷ or multimodal approaches such as theranostics that combines in-situ diagnostics and therapy.¹⁸⁻²¹ Besides the biomedical field, encapsulation processes have also been extensively developed for various applications in agriculture (e.g. for soil fertilization or for safer pesticide release),²²,²³ in food science (e.g. for food preservation or additive delivery),²⁴,²⁵ in cosmetics (e.g. for fragrance encapsulation or for cosmeceutical applications),²⁶,²⁷ or in material science (e.g. for self-healing properties²⁸,²⁹ or for thermal energy storage³⁰).

<figure><img src="image_8.png" /><figcaption>**Figure 1.2.** Example of in-situ production of therapeutic agents from engineered hybrid vesicles for cancer therapy. Polymeric vesicles entrapping a glucose-sensitive enzyme and iron nanoparticles are injected in the blood circulation. At the contact with tumor pH, the permeability of the vesicles dramatically increases and subsequent reactions between diffusing glucose and oxygen and the vesicles' content yield hydroxyl radicals that are able to kill tumor cells.¹⁴</figcaption></figure>

Beyond delivery applications, the design of micro- and nanocompartments has been extensively investigated in emergent topics of fundamental research. For instance, nanoreactors have been designed for the study of confinement effects on chemical reaction rates,³¹,³² for the study of reactivity at the single-molecule level,³³,³⁴ for the production of nanomaterials,³⁵,³⁶ for energy production, storage and conversion,³⁷ or for the development of catalysis for “greener” chemistry.³⁸
At the interface between chemistry and biology, the topic of “artificial cells” (or “protocells”) has also recently drawn increasing attention. Because of the intrinsic complexity of living matter, the precise role of biological processes can be difficult to decipher in vivo. To simplify the approach and better understand the general principles underlying life, chemists have proposed to design artificial microcompartments that could in a sense mimic cellular structures and functions but with a limited number of components and a better control on the physico-chemistry of the system. As a first step towards the imitation of cellular structures, multi-compartmentalized and multi-components artificial structures have been successfully designed since the end of the 2000s (see **Figure 1.3**).³⁹⁻⁴¹ Elegant works have evidenced the implementation of life-inspired functions in such protocells, including self-replication,⁴² differentiation,⁴³ photosynthesis,⁴⁴ phagocytosis,⁴⁵ or intracellular signalization.⁴⁶ The issue of cell-cell communications and interactions has also been explored within protocell communities,⁴⁷⁻⁴⁹ and the formation of prototissues by controlled aggregation of artificial cells has also been recently demonstrated.⁵⁰ As living cells are highly out-of-equilibrium machineries, recent efforts have been made to design active artificial cells that integrate out-of-equilibrium chemical reactions to mimic for instance biological self-sustained motion,⁵¹ or chemical exchanges with the extracellular environment.⁵²

<figure><img src="image_9.png" /><figcaption>**Figure 1.3.** Synthetic multicompartmentalized nanocontainers have been developed to mimic the structures and functions of living cells with a lower level of complexity and a better control on the physico-chemistry of the system.³⁹</figcaption></figure>
In this context of bio-delivery and bio-mimicking, biomacromolecules such as proteins, peptides, or DNA
can play a crucial role, but their encapsulation in increasingly complex and functional containers can be
limited by their intrinsic fragility. The objective of the present work was to develop a versatile approach
for the design of functional (more precisely stimuli-responsive) polymer capsules, in conditions which are
compatible with the presence of potentially fragile biomacromolecules.

For that purpose, we used the phenomenon of interfacial complexation in water-in-oil emulsions to
achieve the spontaneous formation of capsules composed of an aqueous core and a thermoresponsive
polymer shell. Thermally-induced changes of the physico-chemical properties of the capsules were
essentially assessed with *in situ* fluorescence imaging. We also evaluated the compatibility of the interfacial
complexation process with the presence of a model protein for potential applications such as
biomacromolecule vectorization. In addition, we demonstrated that the thermal properties of the polymers
used in the formulation enable the straightforward incorporation of nanoparticles in the polymer shells,
extending the functionalities that can be imparted to the capsules. This was in particular applied to gold
nanoparticles that exhibit optical properties that make them efficient light-to-heat converters at the
nanoscale, paving the way for photothermal stimulation of the capsules.

This manuscript has been organized in the following way: Chapter 2 presents a selective state of the art on the field of encapsulation at small scales, with a special focus on methods compatible with the encapsulation of fragile biomacromolecules such as proteins, before introducing the interfacial complexation approach that has been specifically developed in this work. Chapter 3 illustrates the formation of polymer shells by interfacial complexation with a first thermoresponsive copolymer that includes poly(N-isopropylacrylamide) moieties exhibiting a lower critical solution temperature (LCST). That chapter also details the imaging protocols that have been used to characterize *in situ* the physico-chemical properties of the so-formed polymer shells. Chapter 4 focuses on the incorporation of nanoparticles in the polymer capsules, and details the specific and original thermometry protocol that has been developed to characterize photothermal effects in presence of gold nanoparticles. Chapter 5 presents the results obtained with a second thermoresponsive polymer that includes poly(acrylamide-co-acrylonitrile) moieties exhibiting an upper critical solution temperature (UCST), and that seem to be more promising for the encapsulation and on-demand release of payloads upon thermal and light stimulation. Chapter 6 is the conclusion of this thesis and gives some outlooks and preliminary experiments that pave the way for future developments of this topic.
# Chapter 2

State of the art and positioning
of the present work
## 2.1 The specific challenge of protein encapsulation and release

The highly specific bioactivity of proteins and peptides, optimized by billions of years of evolution, makes them ideal candidates for advanced therapeutic applications. The rise of protein-based medical treatments has been associated to the development of two techniques, the DNA recombinant technology and the solid phase peptide synthesis, which have enabled the production and purification of tailor-made synthetic proteins and peptides at large scale. As a non-exhaustive list of proteins of interest for biomedical or biotechnological applications, we can mention protein drugs for the treatment of chronic disease (such as insulin for diabetes),⁵³ growth factors for tissue repairing or engineering,⁵⁴ antibodies for bio-diagnostics or immunotherapy,⁵⁵ or neuropeptides for the treatment of neurological disorder or the investigation of neuronal signaling pathways.⁵⁶

The specific activity of a protein is correlated to its well-defined structural organization which is dictated by (i) the chemical composition of the amino-acid sequence (primary structure), (ii) the local conformations of the chain backbone (secondary structure) and (iii) the tridimensional folding of the entire structure resulting from intrachain interactions and reorganization of the solvation sphere (tertiary structure). Any small alteration of this optimized structure may induce a complete loss of the protein bioactivity and may be prejudicial for therapeutic goals, making these biomacromolecules highly sensitive to their environment. As an illustration, the first formulations that have been developed for protein delivery were made of aqueous solutions of “raw” proteins (i.e. unprotected or unmodified). Upon injection *in vivo*, these proteins were readily exposed to (i) the recognition and elimination by the immune system, and/or (ii) multiple pathways of chemical degradation (hydrolysis or β-elimination along the backbone, deamidation of asparagine or glutamine residues, oxidation of methionine residues, cleavage of cysteine disulfide bonds).⁵⁷ Chemical degradation induces denaturation of the protein, i.e. misfolding or unfolding of the tridimensional structure, resulting in the loss of its biological activity and the formation of potentially harmful side-products.⁵⁸ In addition, due to their high molecular weight and high hydrophilicity, non-modified proteins are less able (compared to conventional small molecular drugs) to cross the amphiphile cellular membrane, resulting in lower levels of cellular internalization. Altogether, liquid formulations of raw proteins showed poor therapeutic efficiency, and significant side-effects for the patients.

To circumvent these pitfalls, the microencapsulation of proteins appeared as a relevant strategy to protect them from environmental degradation, to maximize their stability, to decrease their immunogenicity, to prolong their lifetime in biological conditions, to increase their cellular uptake and eventually to optimize their therapeutic effect. However, the issue of protein denaturation can also be faced during the formulation of the encapsulation system, since it may involve physical of chemical treatments
that may be harmful for the proteins, such as high temperatures, mechanical stress, chemical reactions, or
presence of incompatible solvents. Thus, achieving an efficient encapsulation of such fragile
biomacromolecules while maintaining their stability and bioactivity is still a challenge in the field of drug
delivery, and more generally in biotechnologies.

Different encapsulation systems have been explored for the stabilization and vectorization of proteins. Only organic systems will be discussed in this thesis, and some of them are represented in **Figure 2.1**. The encapsulation systems can be classified according to their structure (matrix-like vs reservoir-like) and their nature (hydrophobic vs hydrophilic). In matrix-like systems, such as colloidal hydrogels, complex coacervates, polymer microspheres, or polymer micelles, the protein is embedded in a solid-like environment usually made of entangled polymer chains. In reservoir-like systems, like emulsion droplets, vesicles such as liposomes and polymersomes, or polymer capsules, the protein is entrapped in a liquid core surrounded by a protective layer.

Depending on the nature of the polymer matrix or of the liquid reservoir, the systems may be more
appropriate for the encapsulation of either hydrophobic or hydrophilic payloads. Polymer microspheres,
polymer micelles, oil droplets and oil-core polymer capsules are usually suitable for the encapsulation of
hydrophobic compounds. Liposomes, polymersomes, colloidal hydrogels and aqueous-core polymer
capsules are usually appropriate for the encapsulation of hydrophilic species. As most of therapeutic
proteins are hydrophilic, the discussion will only focus on hydrophilic encapsulation systems throughout
this chapter.

<figure><img src="image_10.png" /><figcaption>**Figure 2.1.** Various nano- and microcontainers that have been proposed for the encapsulation and delivery of proteins.⁵⁹</figcaption></figure>
Three crucial issues must be considered for the design of protein encapsulation systems: the efficiency of protein loading during the encapsulation process, the preservation of the stability/activity of the protein within the container, and the control of the release kinetics for delivery applications.

Therapeutic applications do not always require high amounts of delivered proteins to be effective. However, the effects of dilution upon release *in vivo* may decrease the local concentration of delivered protein and thus reduce the efficiency of its biological action. As a consequence, high encapsulation efficiencies (typically > 50 %) of proteins are usually targeted upon formulation. In any case, the waste of product related to poor encapsulation may be regrettable (especially for proteins that are difficult and/or expensive to produce) and motivates the optimization of encapsulation efficiencies. The second major issue is the preservation of the protein integrity upon encapsulation. As previously mentioned, proteins are highly sensitive to their environment and can be impaired in presence of mechanical stress or chemical reactions. The encapsulation of proteins may thus require mild conditions of formulation.

In the next section, we will describe the most common encapsulation systems that have been developed in the literature for the vectorization of hydrophilic payloads (as most of therapeutic proteins are hydrophilic), and concomitantly discuss their relevance regarding protein encapsulation. Then, we will address the issue of controlled delivery in relation to these encapsulation systems. At the light of this state of the art, we will eventually settle upon a system that seems promising for the encapsulation of proteins in mild conditions and for their controlled release upon stimulation.

## 2.2 Micro- and nanocontainers developed for the encapsulation of hydrophilic payloads

### 2.2.1 Vesicles: liposomes and polymersomes

Vesicles are core-shell systems composed of an aqueous core protected by a molecular bilayer (also termed "membrane") formed by the self-assembly of amphiphilic molecules. Two main types of vesicles have been extensively studied for encapsulation applications: liposomes in which the membrane is made of a lipid bilayer, and polymersomes in which the membrane is made of a bilayer of amphiphilic copolymers, most of the time block copolymers (see **Figure 2.2**). The self-organization of these amphiphilic molecules in a
well-defined bilayer is entropically driven by the water desolvation of their hydrophobic moieties (the hydrophobic tail of lipids and the hydrophobic block of amphiphilic copolymers). In the resulting membrane, the hydrophobic moieties are buried in the middle of the bilayer and the hydrophilic moieties are exposed to water at the inner and outer surfaces of the bilayer.

A structural parameter, called packing parameter, enables to rationalize the packing of amphiphiles and predict the morphology of the resulting self-assembled structures. For lipids, this packing parameter is defined as $p = \frac{v}{a_0 l_c}$ (where $v$ and $l_c$ are respectively the volume and the length of the hydrophobic tail, and $a_0$ is the surface area of the polar head).⁶⁰ By comparing the aspect shape of the amphiphile to the case of an ideal cylinder (for which $v = a_0 l_c$), the value of $p$ hence contains the information on the curvature of the lipidic assemblies. Lipidic vesicles are low-curvature objects that are typically obtained for a value of $p$ comprised between 1/2 and 1, i.e. for relatively hydrophobic lipids. For amphiphilic block copolymers, the notion of packing parameter used for lipids has less sense since a surface area cannot be precisely defined for the hydrophilic block. The weight ratio between the two blocks is a more relevant parameter to predict the morphology of the polymeric assemblies. Accordingly, polymersomes are typically formed for a mass fraction of the hydrophobic block higher than 60 %.⁶¹

The main advantage of the vesicular structure is not only the possibility to encapsulate hydrophilic species in the aqueous core (or lumen) of the vesicle, but also to entrap hydrophobic species inside the bilayer membrane. The typical diameter of both types of vesicles can vary between 10-20 nm (for Small Unilamellar Vesicles, or SUVs) and 100-200 µm (for Giant Unilamellar Vesicles, or GUVs), depending on the method of preparation (see **Figure 2.2**).⁶²

<figure><img src="image_11.png" /><figcaption>**Figure 2.2.** Comparison of the structures of liposomes (left) and polymersomes (right). SUV/LUV/GUV: small/large/giant unilamellar vesicles.⁶²</figcaption></figure>
In terms of structural and physico-chemical properties, the main differences between liposomes and polymersomes are essentially due to the intrinsic differences in size and chemical composition of their respective molecular building blocks (lipids vs polymer chains). As natural lipids (mostly phospholipids) have a typical molecular weight of ~ 100-1000 g/mol, the membrane thickness of liposomes is limited to 3-5 nm. As the molecular weight of amphiphilic copolymers can be tuned between ~ 1000 and 10 000 g/mol during their synthesis, polymersomes can exhibit a membrane thickness ranging from 5 to 50 nm.

Since lipids are small and highly mobile molecules, the membrane of liposomes is characterized
by a high lateral fluidity and consequently a high permeability (see **Figure 2.3**). This may be an advantage
to mimic the biophysical properties of cell membranes or to interact with such membranes, but this may be
a drawback regarding the encapsulation of molecules for on-demand delivery as they could be subject to
significant and uncontrolled leakage. Lipid molecules have the major advantage to be biocompatible, but
they are easily subject to chemical degradation (via hydrolysis of ester bonds and oxidation of unsaturated
tails especially), making liposomes relatively fragile and unstable in biological environments over long
time periods.⁶²

On the other side, the membranes of polymersomes are characterized by low lateral fluidity and consequently low permeability and better mechanical properties (compared to liposomes), essentially due to the high degree of entanglement of the polymeric hydrophobic moieties within the bilayer (Figure 2.3). These properties make polymersomes robust candidates for encapsulation applications. In addition, the synthetic control over the composition and functionality of the polymer blocks gives more modularity and versatility to tune the physico-chemical properties of the polymersome membrane (compared to natural lipids which show lower potential for functionalization). As an example, reactive groups can be introduced along the polymer chains to cross-link the polymersome membrane, either for improving the encapsulation and retention efficiency (including for protein encapsulation),⁶³,⁶⁴ or for modulating the permeability.⁶⁵ As a limitation, the polydispersity inherent to the polymer synthesis can affect the structural properties of the polymersomes.⁶⁶
<figure><img src="image_12.png" /><figcaption>**Figure 2.3.** Comparison of the dynamics and chemical stability of the membranes in liposomes and polymersomes.⁶²</figcaption></figure>

The formation of liposomes and polymersomes can be achieved via similar methods, the most used being probably film hydration and solvent displacement methods.

Film hydration consists in dissolving the amphiphile in a suitable organic solvent and to let the solution dry on a substrate, leading to the deposition of an amphiphile multilayer film on the substrate. Then, the addition of water (that may contain hydrophilic species to encapsulate) on the film induces its progressive swelling and budding resulting in the detachment of spherical vesicles (see **Figure 2.4**). Film hydration can be assisted by different external fields: sonication, intercalation of a gel interlayer between the substrate and the amphiphilic film (the swelling of this gel interlayer facilitates the detachment of vesicles upon rehydration), or application of an electric field on the substrate that supports the film (the electric field induces fluctuation in the amphiphile film and favors the formation of vesicles, especially GUVs).
Film hydration is easy to implement but the encapsulation yield can be limited for hydrophilic
molecules since they have to pervade and intercalate between the densely packed layers of the amphiphile
film to be readily entrapped in the lumen of the vesicles during their detachment. This limitation is
particularly true for proteins (and other biomacromolecules) which are high molecular weight and highly
charged molecules, and thus poorly disposed to be inserted in the dried film.⁶⁷⁻⁶⁹ Some techniques have
been proposed to increase the encapsulation yield of such macromolecules in particular conditions, such as
in presence of polymeric crowding agents.⁷⁰ Besides, the use of physical treatments such as sonication or
electric field can be harmful for fragile biomacromolecules to encapsulate.

<figure><img src="image_13.png" /><figcaption>**Figure 2.4.** Formation of liposomes by the film hydration method.⁷¹</figcaption></figure>

Solvent displacement methods rely on the formation of emulsion droplets containing the amphiphiles, followed by the removal of organic solvents temporary used during the process. The simplest solvent displacement method consists in dissolving the amphiphiles in a volatile compatible solvent (e.g. ether) and to rapidly inject this solution in a water bath under vigorous stirring, resulting in the formation of organic nano- or microdroplets that contain the amphiphiles. Then, the progressive “displacement” of the organic solvent (i.e. its removal, by evaporation or dialysis for instance) induces the self-assembly of the amphiphiles in spherical membranes, eventually resulting in vesicles (usually small, multilamellar and
polydisperse) suspended in an aqueous solution. This method is easy to implement but not suitable for the encapsulation of hydrophilic species since the amphiphiles are initially solubilized in an organic solvent.

As an alternative, the emulsion phase transfer method can be used. An initial water-in-oil emulsion is formed with the amphiphile decorating the water droplets (that may contain hydrophilic species). Then this emulsion is slowly injected in a water-organic biphasic solution separated by an interfacial amphiphile monolayer. As the emulsion water droplets sink towards the bottom aqueous phase, they cross the water/organic interface and collect a thin organic layer stabilized by amphiphiles on its two sides. The removal or dewetting of the organic layer eventually leads to the formation of the vesicular bilayer membrane.⁷²

The recent generalization and improvement of microfluidic techniques have enabled considerable progress in the control of emulsion phase transfer for the production of vesicles, in terms of (i) optimization of size and dispersity of the objects,⁷³ (ii) formation of multicompartmentalized structures,⁷⁴ (iii) tunability of the membrane and lumen compositions (see **Figure 2.5**),⁷⁵ and (iv) encapsulation efficiency.⁷⁶

<figure><img src="image_14.png" /><figcaption>**Figure 2.5.** Microfluidics-generated sequential bottom-up assembly of transmembrane and cytoskeletal proteins by high throughput pico-injection into droplet-stabilized GUVs with complex membrane (scale bar = 50 mm).⁷⁵</figcaption></figure>

These approaches based on water-in-oil emulsions have usually better encapsulation yields than film hydration since the hydrophilic payloads are initially confined in water droplets that are “converted” into vesicle lumen upon formation of the membrane.
However, these approaches can be inappropriate for the encapsulation of sensitive and fragile proteins due to the presence of organic solvents that can alter their stability.⁷⁷ Indeed, in native aqueous environment, the tertiary structure of proteins is mostly maintained by intrachain hydrophobic interactions. Upon contact with an organic solvent, proteins can be subject to unfolding due to the affinity of their hydrophobic domains with the organic solvent. This denaturation process usually induces the aggregation of proteins at water/organic interfaces and the loss of their bioactivity.⁷⁸⁻⁸⁰ This interface-induced degradation of proteins is likely to occur in any encapsulation approach that involves water-in-oil or oil-in-water emulsions. It is the case for the preparation of vesicles by phase transfer methods but also for the preparation of other encapsulation systems as it will be mentioned later.

To limit the phenomenon of interface-induced degradation, some formulation adjustments have been proposed such as the use of highly concentrated protein solutions to promote the formation of a “sacrificial” protein layer at harmful interfaces that may shield the rest of the aqueous solution (so-called “self-protecting effect”), or the use of surfactant- or polymer-based protective coatings of proteins that may reduce their direct contact with organic solvents, eventually enhancing their stability.⁸¹,⁸² However, these solutions are usually not generalizable and may be ineffective, or even prejudicial, when switching from one protein of interest to another. As a general statement, the effects of additives such as surfactants, co-solvents, sugar, polymers, salts (added for the necessity of the formulation or as additional ingredients) can have either positive or negative effects on the stability of proteins (depending on its nature) and are difficult to rationalize.⁵⁷ However, some mild conditions have already been found to effectively encapsulate specific proteins in polymersomes, even in presence of chemical reactions used to cross-link (stabilize) the vesicular structure.⁶⁴

As an alternative to emulsion-based approaches, the phenomenon of polymerization-induced self-assembly (PISA) has led to breakthroughs in the design of functional polymersomes and other types of self-assembled nanostructures in water.⁸³,⁸⁴ In the PISA process, the sequential growth of amphiphilic polymer chains triggers their self-assembly in *situ* in well-defined nanostructures (spherical micelles, worm-like micelles, vesicles). The rise of this technique has been largely enabled by the optimization of reversible-deactivation radical polymerization (also termed “controlled” or “living” radical polymerization).⁸⁵,⁸⁶ Briefly, after the synthesis of a “living” hydrophilic block in water, the polymerization is carried on by the addition of monomers whose corresponding homopolymer is hydrophobic and insoluble in water. Upon extension of the hydrophobic block, an amphiphilic nature is progressively imprinted to the growing chains, resulting in their self-assembly in nanostructures whose morphology can be controlled by variations of the nature of the monomers, the degrees of polymerization, or by the use of additives (see **Figure 2.6**).⁸⁷
Regarding the production of vesicles, PISA has been for long restricted to the formation of nano-
polymersomes (typically 10-100 nm), but the formation of 10-µm GUVs has been recently achieved with
this approach.⁸⁸ Recent works have also demonstrated the potential of PISA-based polymersomes for
protein encapsulation, either in the lumen⁸⁹ or in the membrane of the vesicles.⁹⁰

<figure><img src="image_15.png" /><figcaption>**Figure 2.6.** Illustration of the polymerization-induced self-assembly (PISA) process. The reversible addition-fragmentation chain-transfer (RAFT) polymerization of a hydrophobic block of poly(2-hydroxypropyl methacrylate) from a hydrophilic macromolecular chain transfer agent (macro-CTA, either poly(glycerol monomethacrylate), poly[2-(methacryloyloxy)ethyl phosphorylcholine], or poly(ethylene glycol)) produces amphiphilic chains that self-assemble in nanostructures. Spheres, worms, or vesicles can be produced by judicious variation of the packing parameter (P) which is determined by the relative mass fractions of the hydrophilic and hydrophobic blocks (see second paragraph of this section).⁸⁷</figcaption></figure>

2.2.2 Colloidal hydrogels

Colloidal hydrogels are micro- or nanospheres (termed microgels or nanogels respectively) obtained by physical or chemical cross-linking of hydrophilic polymer chains, resulting in a colloidal network that can swell in water without falling apart. Their internal cross-linked structure provides mechanical properties which facilitate their manipulation and make them robust cargos for delivery applications.

Colloidal hydrogels are commonly prepared according to one of the three following conventional approaches: the polymerization of a colloidal network from monomer and cross-linker building blocks, the chemical cross-linking of pre-formed reactive polymer chains, and the physical cross-linking of pre-formed interactive polymer chains (see **Figure 2.7**).⁹¹,⁹²
<figure><img src="image_16.png" /><figcaption>B. Chemical/Physical Crosslinking of Polymer in Emulsion</figcaption></figure>

<figure><img src="image_17.png" /><figcaption>C. Self-Assembly through Physical interaction</figcaption></figure>

<figure><img src="image_18.png" /><figcaption>D. Self-Assembly through Electrostatic Interaction</figcaption></figure>

<figure><img src="image_19.png" /><figcaption>**Figure 2.7.** Various approaches for the formation of micro- and nanogels.⁹³</figcaption></figure>

The formation of colloidal hydrogels by polymerization of monomer and cross-linker units (usually hydrophilic acrylate derivatives) can take place in water directly (homogenous conditions), or in emulsion (heterogenous conditions).⁹⁴

In homogeneous conditions, the formation of colloidal hydrogels has been markedly enhanced by the development of reversible-deactivation radical polymerization which provides a better control on the propagation reactivity and favors the intrachain cross-linking over interchain cross-linking (this latter resulting in macrogelation and not in the formation of individualized colloidal gels).⁹⁴,⁹⁵ An other widespread method for producing colloidal gels in homogenous conditions is the precipitation polymerization, relying on the difference of solubility between the polymer precursors (water-soluble monomers and cross-linkers) and the final network (insoluble in water after reaching a critical size).⁹⁶

The most common method for preparing colloidal hydrogels is however the polymerization in emulsion, and especially in water-in-oil inverse emulsion. An aqueous solution containing the polymer precursors (monomers, cross-linker, and initiator) is emulsified in an immiscible organic solvent containing a surfactant. The polymerization occurs within surfactant-stabilized water droplets that play the role of a template, yielding spherical hydrogels. After polymerization, the organic phase is removed (by evaporation or centrifugation) and the hydrogels are redispersed in water to eventually recover a colloidal suspension. This versatile approach enables a good control on the chemical composition and on the cross-linking density of the final gels.
Colloidal hydrogels can also be achieved by chemical or physical cross-linking of preformed polymer chains. Chemical cross-linking relies on the presence of reactive chemical functions along the polymer backbone, that can react either together or with additional cross-linking species to form covalent networks.⁹⁷,⁹⁸ This approach turned out to be favored when the reactive polymer chains were either confined in emulsion droplets, pre-self-assembled in nanostructures (such as micelles), or adsorbed on solid core templates.

As an alternative to chemical reactions, colloidal hydrogels can also be formed upon physical cross-linking of polymer chains, i.e. via non-covalent interactions (hydrophobic interactions, electrostatic interactions, hydrogen bonds, metal-ligand complexation, host-guest interactions).⁹⁹⁻¹⁰² They however may suffer from poorer stability compared to covalently cross-linked analogs. Despite this limitation, one particular type of physical cross-linking has recently aroused increasing attention, and is called complex coacervation. The phenomenon of complex coacervation designates the liquid-liquid phase separation that can spontaneously occur between aqueous solutions of oppositely charged macromolecules. This process results in the formation of nanocondensates maintained by electrostatic interactions, which can be seen as physically cross-linked hydrogels. Being highly versatile, tunable and suitable for the efficient entrapment and stabilization of biomacromolecules, including proteins (see **Figure 2.8**), complex coacervation has become a promising platform for the engineering of biocompatible and easily cell-internalizable nanocarriers.⁵⁹,¹⁰³,¹⁰⁴

<figure><img src="image_20.png" /><figcaption>**Figure 2.8.** Encapsulation of a model protein (BSA) in complex coacervate microdroplets formed by liquid-liquid phase separation of an aqueous solution of oppositely charged polypeptides.¹⁰³</figcaption></figure>

To broaden the applicability of colloidal hydrogels, a progressive diversification in the choice of the monomer or polymer precursors has been proposed in the literature. The diversification of the monomer units has enabled the design of amphiphilic hydrogels to favor the incorporation of more hydrophobic drugs (in matrices which are mostly hydrophilic).¹⁰⁵,¹⁰⁶ The development of physically cross-linked hydrogels has more and more involved the use of biopolymers (such poly(amino acids) or polysaccharides) in the particle
scaffolds to favor the accommodation of biomacromolecules (including proteins), and to improve the biocompatibility of the carriers.<sup>107-111</sup>

The incorporation of proteins in colloidal gels can be achieved concomitantly with the formation of the polymer matrix by simply mixing the protein with the polymer precursors.<sup>112</sup> In that case, the encapsulation efficiency of the protein is markedly governed by the polymer properties (chain molecular weight or polymer concentration in solution).<sup>113</sup> This approach may expose the proteins to organic solvents and/or radical species that can be prejudicial for their stability. As an alternative, it has been proposed to dissociate the steps of gel formation and protein encapsulation, by loading the proteins of interest in pre-formed colloidal gels, either by impregnation (i.e. by diffusion through the polymer matrix, see **Figure 2.9**),<sup>114</sup> or by adsorption at their surface.<sup>115</sup> The efficiency of the impregnation process (in terms of yield and uniformity) depends on the chemical nature and the internal structure of the gels, especially on the crosslink density which dictates the size of the voids in the polymer network.<sup>115,116</sup> The process of adsorption usually suffers from limited encapsulation yields and does not protect the protein of interest from the environment.

In colloidal gels loaded with proteins, cross-talks between the protein and the polymer matrix may have a crucial influence on both the protein stability and the gel properties. Protein-polymer interactions, such as electrostatic attraction or hydrogen bonding, can be beneficial to increase the loading and retention of proteins,<sup>116-118</sup> to improve their thermal stability by hindering thermally-induced conformational changes, or to enhance their bioactivity.<sup>119</sup> In this line, proteins can even be used as cross-linking agents of the polymer network, and then released in suitable chemical conditions without significant structural alteration as demonstrated recently with redox-sensitive microgels.<sup>120</sup> *A contrario*, the potential reactivity of the polymer network may also lead to protein deterioration. For instance, in polyester matrices – which are interesting for their biocompatibility and biodegradability – the hydrolysis of polymer ester groups into carboxylic acids induces a progressive acidification of the microenvironment (to pH as low as 3),<sup>121</sup> that may eventually trigger the destabilization of encapsulated proteins. Note that if the polymer properties may influence the structure of the protein, reciprocally the presence of proteins may change the structural properties of the polymer matrix. In colloidal gels, a deswelling of the matrix is usually observed upon the loading of oppositely charged proteins, essentially due to protein-polymer electrostatic attraction that favor water expulsion from the matrix.<sup>115,116</sup>
<figure><img src="image_21.png" /><figcaption>**Figure 2.9.** Schematic presentation of a post-fabrication encapsulation strategy of protein drugs in microgels. Microgels were obtained from cross-linking of the pre-designed macromers. After microgel preparation, drug loading was performed by soaking aqueous solution of proteins with microgels at 4 °C (below volume phase transition temperature). The microgels collapsed after drying and thus entrapped proteins in their network. When released in buffer at human body temperature, striking early release of protein was decreased because of the shrinkage of microgels at 37 °C with respect to at 4 °C. The drug release proceeded with hydrolysis of microgels and underlying diffusion. All of proteins were eventually released due to complete biodegradation of the polymeric network.<sup>114</sup></figcaption></figure>

### 2.2.3 Polymer microcapsules

Polymer microcapsules are hollow microspheres composed of an internal reservoir surrounded by a polymer envelope. They can be seen as “core-shell” systems with a core made of an organic or aqueous solution, and a shell made of polymer chains that can be chemically cross-linked or maintained by non-covalent interactions. In contrast to the precise structuration of the bilayer membrane in polymersomes, capsule shells are usually made of a more chaotic arrangement of polymer chains in intertwined mesh, network, or multiple layers. The shell thickness of polymer capsules is highly dependent on the preparation method but can range from a few tens of nm to several microns, and is usually much thicker than polymersome membrane which is restricted to 5-50 nm (as roughly given by twice the length of the polymeric building block, see Section 2.2.1). Polymer capsules thus associate (i) the reservoir structure of vesicles with (ii) shell physical properties that are close to hydrogels’ ones.
In absence of intrinsic amphiphilicity, polymer chains have no reason to spontaneously self-organize into hollow structures with a liquid core. The formation of polymer capsules thus requires a template to drive the assembly or the growth of polymer chains into shells with well-defined size and morphology. The main techniques that have been developed for that purpose use solid nanoparticles or emulsion droplets as spherical template for the design of liquid-core hollow capsules (see **Figure 2.10**).

<figure><img src="image_22.png" /><figcaption>**Figure 2.10.** Various template-based strategies for the design of liquid-core hollow polymer capsules.<sup>122</sup></figcaption></figure>

Solid cores can be used as sacrificial templates for the successive deposition of oppositely charged polymers, resulting in a multi-layered polymer shell self-maintained by electrostatic interactions after removal of the solid core. This technique, usually referred to as layer-by-layer (LbL) assembly, enables a good control on the composition and thickness of the shells but at the expense of numerous preparation steps.<sup>123</sup> Note that other types of interactions have also been used to drive the LbL assembly, such as hydrogen-bonding (see **Figure 2.11**),<sup>124</sup> or host-guest supramolecular interactions,<sup>125</sup> but the electrostatic interaction remains the most studied one.

Despite numerous examples of bio-applications of LbL capsules (including for protein delivery),<sup>126,127</sup> this encapsulation technique is not necessarily appropriate for any payload since (i) the encapsulation efficiency is intrinsically limited by the presence of the solid template all along the capsule formation and (ii) the removal of the solid template usually involves drastic chemical treatments (including strong acidic or alkaline conditions) that may be harmful for fragile biomacromolecules such as proteins.<sup>128</sup> To circumvent these issues, some examples of LbL assembly on liquid core templates have been proposed.<sup>129,130</sup>
<figure><img src="image_23.png" /><figcaption>**Figure 2.11.** Schematics for the multilayer assembly of hydrogen-bonded (PVPON/PMAA) capsules. (a) Silica cores (a) were exposed alternately to PVPON and PMAA polymer solutions in methanol with two rinsing steps in methanol between the layers. (b) After a desired number of (PVPON/PMAA) bilayers (denoted as n) were assembled, silica cores were dissolved in hydrofluoric acid and dialyzed at pH 3, leaving behind hollow hydrogen-bonded (PVPON/PMAA)n capsules. The dashed rectangle highlights the functional groups interacting through hydrogen bonding. (c) SEM image and (d) infrared spectrum of (PVPON/PMAA)5 capsules assembled in methanol. Atomic force microscopy topography images of (e) 5 bilayer and (f) 10 bilayer (PVPON/PMAA) capsules dried on silica wafer surfaces from methanol.¹²⁴</figcaption></figure>

Most of the techniques of capsule formulation rely on emulsion droplets as the template for the formation or the self-assembly of polymer shells. It is the case for internal phase separation, which consists in progressively triggering the precipitation of polymer chains confined in emulsion droplets by changing its affinity with the solvent in the droplet core. Initially the polymer is dissolved in a mixture of good and poor solvent, which is then emulsified in an immiscible continuous phase. The good solvent is progressively removed from the droplets by evaporation, leading to the phase separation of the polymer from the poor solvent remaining in the droplets. This process eventually results in polymer segregation at the droplet surface, leaving behind liquid-core rigid capsules dispersed in the continuous phase. This approach has been used to produce aqueous-core poly(methyl methacrylate) capsules in oil (with acetone/water as good/poor polymer solvent).¹³¹

Internal phase separation can also be triggered in droplets containing only a good solvent for the polymer, the poor solvent coming from the continuous phase. This process was for instance observed in an aqueous dispersion of chloroform droplets containing polyester chains and a suitable surfactant.¹³² The presence of the surfactant in the chloroform droplets triggers the self-emulsification of water (coming from the continuous phase) within the droplets. Water being a poor solvent for the polymer chains, the growth of water droplets formed in the chloroform droplet, concomitant with the evaporation of the chloroform, progressively leads to the formation of polyester shells (see **Figure 2.12**).
<figure><img src="image_24.png" /><figcaption>**Figure 2.12.** Illustration of microcapsules formation by internal phase separation induced. (a) Chloroform droplets containing polylactide chains and bis(2-ethylhexyl) sulfosuccinate surfactant were formed upon emulsification in an aqueous phase. Rapidly, water diffused in the chloroform droplets and formed aqueous inclusions upon internal self-emulsification. (b) Count and size of the internal water droplets increase with the increase of time; (c) Internal tiny droplets coalesce into bigger ones; (d) Double emulsion droplets were formed via coalescence of the internal water droplets; (e) Polylactide microcapsules were obtained after complete evaporation of chloroform.¹³²</figcaption></figure>

In these approaches of internal phase separation, the structural features of the polymer microcapsules (capsule size, shell thickness and porosity) can be tuned by varying the size of the templating emulsion droplets, the mass of the polymer initially dissolved in the dispersed phase and the evaporation rate of the good solvent. However, these approaches require the use of organic solvents in the dispersed phase, which is barely compatible with the presence of proteins.

Another approach for emulsion-templated capsule formation consists in directly growing or cross-linking polymer chains *in situ*, i.e. at the surface of emulsion droplets. This general strategy is usually referred to as interfacial polymerization.†

Polyaddition reactions offer a broad choice of monomer or polymer building blocks for the design of capsules by interfacial polymerization. Polymer capsules can be achieved by coupling reactive species that are initially soluble in their own phase but that can meet and react at the water/oil interface after emulsification. Common click or metathesis reactions between difunctional complementary (macro)monomers have been used in that sense, such as azide-alkyne cycloaddition with or without Cu catalyst,¹³³,¹³⁴ thiol-ene coupling,¹³⁵ or olefin cross-metathesis.¹³⁶

<footnote>† In this thesis, interfacial polymerization will only refer to reactions that produce *covalent bonds* between monomers or polymer chains at the interface between two immiscible media. Some authors may also include non-covalent cross-linking (e.g. via hydrophobic or host-guest supramolecular interactions) or metal-ligand coordination among the list of interfacial polymerization reactions.</footnote>
Radical or ionic interfacial polymerization has also been proposed to build polymer shells *in situ*.
In this approach, the main challenge is to confine the chain propagation at the interface while limiting its
extension to the bulk phases of the emulsion. For that goal, the polymerization must be mediated by
reactants that are surface active, i.e. that have a high affinity with the water/oil interface. Effective
interfacial radical polyaddition have been demonstrated using amphiphilic or surface-active monomers,¹³⁷
radical initiators,¹³⁸ initial radical species,¹³⁹ or RAFT agents¹⁴⁰ (see **Figure 2.13**). The interfacial
sequestration of polymer can also be achieved if the growing chains have no affinity neither for water nor
for oil, resulting in the formation of a polymer shell at the water/oil interface.¹⁴¹ As an alternative to radical
polymerization, anionic polymerization has also been successfully implemented at water/oil interfaces to
produce aqueous-core capsules (especially derived from poly(alkylcyanoacrylates)),¹⁴² that may be suitable
for protein encapsulation.¹⁴³

<figure><img src="image_25.png" /></figure>

<figure><img src="image_26.png" /><figcaption>**Figure 2.13.** Interfacial radical polyaddition based on the confinement of reactive species at the water/oil interface of emulsion droplets. The reaction was confined thanks to the surface-active properties of a) monomers,¹³⁷ b) radical initiators,¹³⁸ c) initial radical species,¹³⁹ or d) RAFT agent.¹⁴⁰</figcaption></figure>
Polycondensation reactions can also be used to form polymer capsules at the surface of emulsion droplets. These reactions usually involve addition-elimination reactions between water-soluble diamines or diols and oil-soluble diacyl chloride or diisocyanates, yielding polyester, polyamide, polyurea (and other derivatives) capsule shells.¹⁴⁴ This approach can be extended to the formation of cross-linked polymer shells by starting from (at least) tri-functional reactive species, such as lignin.¹⁴⁵ As for click polyaddition, these polycondensation reactions involve complementary but immiscible reactive species, naturally restricting the polymerization at the water/oil interface. Note that this interfacial condensation strategy has also been implemented in sol-gel processes to readily form aqueous-core silica hollow nanocapsules.¹⁴⁶ Interfacial polymerization techniques usually lead to the formation of robust polymer shells despite their low thickness (typically below 100 nm because the diffusion of reactive species towards the interface is quickly hindered by the presence of the growing shell). However, since these strategies of capsule formation are based on *in situ* chemical reactions, they may be inappropriate for the encapsulation of fragile payloads, especially for proteins that may be degraded in presence of radicals, thiol groups, carboxylic acid derivatives or organic solvents that are commonly encountered in interfacial polymerization processes.

An alternative method to the interfacial polymerization may be to assemble pre-formed polymer chains at the surface of emulsion droplets to readily produce polymer shells without resorting to *in situ* chemical reactions. The formation of polymer capsules may be thus induced by non-covalent (weak) interactions between complementary interactive chains that are present on the two sides of the water/oil interface. This approach, termed interfacial self-assembly, includes multiple variants such as interfacial complexation, coordination, supramolecular polymerization, depending on the interactions which are involved in the shell structuration. The most studied ones are probably electrostatic interactions that are implicated in interfacial complexation: when two oppositely charged species are solubilized in their corresponding phase of an emulsion, they spontaneously accumulate at the surface of the emulsion droplets, driven by coulombic attraction across the liquid-liquid interface. The formation of aqueous-core polymer capsules by interfacial complexation has already been reported in water-in-oil (W/O) (see **Figure 2.14**),¹⁴⁷,¹⁴⁸ and in water-in-water (W/W) emulsions.¹⁴⁹⁻¹⁵²
<figure><img src="image_27.png" /><figcaption>**Figure 2.14.** Capsule fabrication by interfacial complexation of polyelectrolytes in W/O emulsion in a microfluidic device. (a) Aqueous phase (turquoise) containing a water soluble polyelectrolyte and an oil phase (yellow) containing oil soluble polyelectrolyte are brought into contact to fabricate aqueous-core microcapsules with a polymeric shell. (b) Electrostatic interaction of the polycation (red) and polyanion (blue) across the droplet interface generates a robust shell with thickness in the range of 1-2 µm. (c) Optical image of monodisperse stable capsules (diameter ~ 200 µm) in the exit flow capillary.<sup>148</sup></figcaption></figure>

W/W emulsions (also termed “all-aqueous emulsions” or “aqueous two-phase systems”) can be generated by spontaneous demixing of solutions of hydrophilic polymers that become incompatible above a critical concentration (typically occurring with poly(ethylene glycol) and dextran aqueous solutions).<sup>153</sup> The introduction of oppositely charged polymers in the two immiscible aqueous phases enables the production of aqueous-core polymer capsules by interfacial complexation in absence of organic solvents, which is major advantage for the design of biocompatible nanocarriers, and which enabled the encapsulation of proteins and cells with success.<sup>154,155</sup> However, controlling the partitioning of water-soluble species between the inner and outer aqueous phases, and thus the encapsulation efficiency, remains a crucial challenge in all-aqueous systems.<sup>155</sup>

Beyond simple emulsions, polymer capsules have also been achieved by interfacial complexation in multiple emulsions such as W/O/W,<sup>156,157</sup> or W/W/W.<sup>154,158</sup> For the first case, the formation of a polymer shell by complexation at the inner O/W interface can trigger the spontaneous dewetting of the oil intermediary layer, resulting in the production aqueous-core capsules dispersed in water (see **Figure 2.15**).<sup>156,157</sup>
<figure><img src="image_28.png" /><figcaption>**Figure 2.15.** A) Schematic illustration for the formation of microcapsules by Nanoscale Interfacial Complexation in Emulsion (NICE) from a water-in-oil-in-water (W/O/W) double emulsion. B) Schematic illustration showing complete dewetting of a NICE microcapsule from an oil droplet. C) Optical microscopy image of W/O/W double emulsions generated using a glass capillary microfluidic device. D) Optical microscopy image of NICE microcapsules completely separated from oil droplets (pink). The oil droplets contain 0.1 wt % Nile red (a fluorescent dye).¹⁵⁶</figcaption></figure>

The phenomenon of interfacial complexation has been extensively described through the formation of electrostatic interactions but other interactions have also been used to produce rigid microcapsules such as interfacial metal-ligand coordination,¹⁵⁹ or interfacial hydrogen bonding¹⁶⁰. The formation of multiple interpolymer hydrogen bonds have also been used to drive the generation of hollow polymer capsules from compact nanoparticles by spontaneous maturation at an appropriate pH.¹⁶¹ This self-structuration phenomenon has the advantage to readily produce polymer capsules without the use of an external template.
## 2.3 Remote control of payload release from microcontainers

As mentioned previously, the release of encapsulated content is one of the main motivations for the design of encapsulation systems. Any encapsulation system is subject to payload release via a “natural” mechanism of passive diffusion out of the container scaffold. The kinetics of this mechanism is intrinsically related to the size of the payload and to the internal structure of the container. For instance, considering polymer scaffolds of the same nature, the delivery of proteins is trivially faster from hollow containers than from dense ones, because the thickness of the polymer matrix to cross is lower in the first systems than in the second.¹⁶² In colloidal gels, the release rate is essentially governed by the swelling capacities of the polymer network, and can be lowered for example by increasing the cross-link density or by introducing more hydrophobic polymer moieties in the network.¹⁶³ For any polymer-based encapsulation systems, the protein-polymer interactions play also an important role on the kinetics and efficiency of the release. Specific attractive interactions as well as non-specific adsorption of proteins in the polymer matrix may reduce the release fraction and lower the delivery kinetics.¹⁶⁴

Delivery systems based on passive diffusion suffer from a poor spatio-temporal control of the release since the kinetics is “pre-recorded” in the structure of the encapsulation system and cannot be modulated *in situ*. The possibility to trigger and modulate the release *on-demand* has been a hot topic in the field of molecular delivery for the last 20 years. Major progress has been made thanks to the design of stimuli-responsive nanocontainers, i.e. structures that can be modified or destabilized upon specific chemical or physical stimulation. The on-demand release can proceed via two types of response of the encapsulation system upon stimulation: either *via* a change in porosity/permeability of the scaffold (leading to the enhancement of the diffusion kinetics) or via a partial or total disruption/disassembly of the scaffold.

### 2.3.1 Internal and external stimuli

Two categories of stimuli have guided the design of responsive microcontainers: internal stimuli which are intrinsically present in the delivery area, leading to environmentally-controlled release, and external stimuli which are triggered by an operator out of the delivery area, leading to remotely-controlled release.¹⁰

Common internal stimuli for *in vivo* applications are the temperature of the living organism (usually higher than the ambient temperature) and the chemical environment (acidic pH, redox conditions, presence of reactive biomolecules such as sugars or enzymes). External stimuli include light irradiation, magnetic field, and ultrasounds. The advantage of these stimuli is the possibility of a remote activation (i.e. without
physical intrusion in the biological sample), but on condition that the propagation of the stimulus in the biological sample is efficient enough. This may require high magnetic fields for magnetic-induced delivery or specific irradiation wavelengths for light-induced delivery for instance.

The challenge for chemists and material scientists has been to design microcontainers that can be sensitive to these stimuli, and to be able to tune the rate and the efficiency of the release upon such sti. In the following, the discussion will focus on temperature and light as they turned out to be promising levers for on-demand delivery.

### 2.3.2 Temperature as an internal trigger

The internal temperature of the biological target can be used as a trigger for the destabilization of encapsulation systems. Temperature-sensitive containers rely on changes in physical properties of their scaffold upon heating to trigger the release of their payloads. For that, thermoresponsive polymers are ideal tools since they are characterized by a drastic change of their physico-chemical properties in solution, concomitant with a transition of the chain conformation, at a critical temperature.¹¹ This critical temperature can be a Lower Critical Solution Temperature (LCST), associated to a transition of the polymer chains from hydrophilic soluble coils below the LCST to hydrophobic collapsed globules above the LCST. This transition is due to an entropically-driven desolvation of the polymer chains upon heating resulting in their association via hydrophobic interactions (see **Figure 2.16.a**).¹⁶⁵ At the macroscale, this transition manifests through the formation of polymer aggregates which precipitate in the solvent upon heating. Polymers exhibiting this type of behavior are called “LCST polymers” by misuse of language.

Conversely, the critical temperature can be an Upper Critical Solution Temperature (UCST) associated to the reverse transition: polymer chains are insoluble below the UCST and soluble above the UCST, and the precipitation of the polymer occurs upon cooling. Polymers exhibiting this type of behavior are called “UCST polymers”. Below the UCST, the insolubility of the polymer is due to multiple inter-chains weak interactions (mostly electrostatic interactions and hydrogen bonds) that maintain the polymer chains in an aggregative state. Upon heating, this network of interactions is progressively broken, resulting in the solubilization of the polymer chains.¹⁶⁶

The critical temperature of transition depends on the concentration of the thermoresponsive polymer in solution. The curve giving the evolution of the critical temperature as a function of the polymer concentration is called the binodal curve. This curve separates the experimental domain where the polymer is soluble from the domain where it phase-separates from the solvent. To be rigorous, the notions of LCST and UCST correspond to the extrema of the binodal curve (the minimum for the LCST and the maximum
for the UCST) and is thus valid for one polymer concentration (see Figure 2.16.b). The generalization of this notion of critical temperature at any polymer concentration is called the “cloud point” since it separates the temperature domain where the polymer is soluble in the solvent (transparent solution) from the temperature domain where the polymer precipitates in the solvent (turbid solution) (see Figure 2.16.c).

<figure><img src="image_29.png" /><figcaption>**Figure 2.16.** Illustration of a LCST behavior in the case of poly(N-isopropylacrylamide) (PNIPAM). a) Coil-to-globule transition of PNIPAM, corresponding to a switch from a hydrophilic state with solvated chains below the LCST to a hydrophobic state dominated by inter-chains interactions above the LCST. b) Binodal diagrams for UCST and LCST polymers depending on polymer fraction. c) Typical turbidity heating curve of a PNIPAM solution. The solution is transparent (100% transmittance) below the cloud point, and turbid (0% transmittance) above the cloud point.<sup>165</sup></figcaption></figure>

In aqueous environment, LCST polymers have been studied since decades. The most popular one has undoubtedly been the poly(N-isopropylacrylamide) (PNIPAM),<sup>167</sup> but increasing attention has been recently paid to other families of LCST polymers such as oligo(ethylene glycol)s,<sup>168</sup> poly(oxazoline)s<sup>169</sup> or poly(N-vinylcaprolactam)s.<sup>170</sup> Their implementation in delivery systems essentially relies on the deswelling or rigidification of the compartments at 37 °C (above the cloud point), resulting from the hydrophilic-to-hydrophobic transition of the polymer. Typically, a “squishing” release mechanism was observed during deswelling of PNIPAM microgels upon heating.<sup>171</sup> This thermal behavior usually leads to passive, slow and incomplete release.

For reservoir-like systems, a heat-induced on-demand delivery from LCST polymer membranes/shells seems counterintuitive since the increase of temperature makes the chains more hydrophobic, i.e. less permeable to hydrophilic payloads. Accordingly, polymersomes or polymer capsules based on LCST polymers usually exhibit a delivery upon cooling rather than upon heating.<sup>172,173</sup> However,
some aqueous-core systems have been successfully designed for a release at high temperature, such as polymersomes destabilized by heat-induced collapse of LCST polymer chains,¹⁷⁴,¹⁷⁵ or polymer capsules which include PNIPAM colloidal hydrogels as thermoresponsive gates in their shells (see **Figure 2.17**).¹⁷⁶

<figure><img src="image_30.png" /><figcaption>**Figure 2.17.** Thermoresponsive polymer capsules made of a PNIPAM hydrogel core and a porous ethylcellulose (EC) shell embedded with PNIPAM microgels. The collapse of the PNIPAM microgels above a critical transition (called Volume Phase Transition Temperature, VPTT here) acts as gate opening in the polymer shell and triggers the release of the content which is encapsulated in the capsule core.¹⁷⁶</figcaption></figure>

In contrast, UCST polymers are obviously more suitable for heat-triggered permeabilization of encapsulation systems, because they form insoluble materials at low temperature (below UCST), which can then be dissolved upon heating. Yet, only a handful of UCST-based delivery systems has been proposed.

Although UCST is most likely observed in non-aqueous and hydro-organic solvents, polymers with UCST behavior in pure water are not rare.¹⁷⁷,¹⁷⁸ Several ones exhibit a thermal transition in temperature windows relevant for most practical application (typically 10-50 °C) and/or in physiological conditions.¹⁷⁹,¹⁸⁰ Their main drawback to date has been their sensitivity to environmental and structural features: the transition temperature depends on the polymer structure (composition, chain length, end-groups), and on minor chemical degradation or changes in the environment (e.g. effect of pH, presence of salts).¹⁸¹ For instance, acidic or salt impurities may shift the working temperature of UCST polymers by more than 10 °C.¹⁷⁷ In this line, the UCST of poly(N-acryloyl glycinamide), one of the earliest non-ionic copolymers with chemically adjustable properties, can be dramatically shifted upon monomer degradation (e.g. via hydrolysis) or in presence of poorly controlled chain ends.¹⁸² In addition, the UCST transition shows a strong hysteresis upon heating/cooling cycles. Polyzwitterionic chains are more robust alternatives, and the permanent hydration of ionic comonomers makes them particularly convenient for the design of hydrophilic materials, such as temperature-responsive colloidal hydrogels,¹⁸³,¹⁸⁴ but the high density of ionic groups may be prejudicial for the stability of encapsulated proteins. One class of non-ionic copolymers, the poly(acrylamide-co-acrylonitrile) (PAAMAN), combines the desirable properties of (i) becoming hydrophobic at low temperature, (ii) having a tunable UCST via simple balance of monomer
composition, and (iii) exhibiting a robust UCST transition in water (minimal hysteresis and minimal shift upon temperature cycling).<sup>185</sup> However, the UCST of PAAMAN remains sensitive to the presence of additives such as salts or drug loading.<sup>186,187</sup>

To date, most of the microcontainers exhibiting a UCST behavior have been prepared under the form of colloidal gels,<sup>178,184,188,189</sup> and only a few examples of aqueous-core systems, essentially polymersomes, have been reported.<sup>179,190,191</sup> To the best of our knowledge, aqueous-core polymer capsules with UCST behavior has never been reported.

### 2.3.3 Light as an external trigger

Using light as an external trigger is interesting since it allows a precise spatial and temporal control of the stimulation. Indeed, the irradiation is instantaneous (contrary to temperature triggers that may suffer from heating inertia) and the spatial resolution is essentially dictated by the size of the light source spot (that can reach the microscale for focalized laser beam). For biological applications, the high tunability offered by light excitation (in terms of wavelength and power) can be limited by two factors: the strong absorption of biological tissues in the visible range and the potential phototoxicity of energetic irradiations such as UV light. These issues usually restrain the use of light-induced delivery to the study of transparent or poorly absorbent model organism (such as zebrafish larva) or to the design of near-infrared-sensitive systems (as the near infrared region is a transparent window for biological tissues).<sup>192</sup>

Intrinsically light-controllable release systems are essentially based on two families of light-sensitive molecular building blocks: photo-isomerizable groups and photo-cleavable groups.<sup>193</sup>

Photo-isomerizable groups can undergo a reversible conformational change upon light irradiation, inducing a drastic switch of shape and polarity. The most common photo-isomerizable groups are derivatives of azobenzene which are characterized by a cis-trans isomerization, or of spiropyran, dithienylethene, or donor-acceptor Stenhouse adducts (DASA), which are characterized by an open-closed isomerization. The incorporation of photo-isomerizable groups in the structure of polymeric building blocks may enable to form light-responsive structures that can undergo drastic changes in structural organization and/or polarity/hydrophilicity upon light irradiation, providing a simple way to control the release of pre-encapsulated content.

The permeability of polymer capsules or colloidal gels can be photo-regulated by using azobenzene groups as supramolecular or physical cross-links that can be disrupted when azobenzene are in their cis
form (see **Figure 2.18**).¹⁹⁴,¹⁹⁵ The modulation of permeability has also been demonstrated in polymersomes bearing spiropyran or DASA groups (the permeability being higher when these groups are in their hydrophilic open form).¹⁹⁶,¹⁹⁷ The release of pre-encapsulated content can also be triggered by more drastic phenomena such as the partial or total destabilization of self-assembled polymeric compartments as it was demonstrated for polymersomes containing azobenzene groups (dramatic change in the membrane curvature upon isomerization).¹⁹⁸,¹⁹⁹

<figure><img src="image_31.png" /><figcaption>**Figure 2.18.** Light-responsive microcapsules made by layer-by-layer supramolecular assembly of poly(acrylic acid-g-adamantane-g-azobenzene) (PAA-g-AD-g-Azo) and poly(aspartic acid-g-β-cyclodextrin) (PASP-g-β-CD). Upon UV irradiation, the trans-to-cis isomerization of azobenzene moieties triggers the dissociation of the host-guest cross-links formed with β-CD, leading to an increase of the permeability of the capsule shell and the release of the pre-encapsulated content (here fluorescent poly(ethylene glycol), PEG).¹⁹⁵</figcaption></figure>

Photo-cleavable groups can undergo irreversible splitting in (at least) two residues upon light irradiation. The most common photo-cleavable groups are *o*-nitrobenzyl esters, polyaromatic hydrocarbon methyl esters (pyrene, perylene, ...), or *p*-methoxyphenacyl esters.¹⁹³ Introducing photo-cleavable groups in encapsulation/release polymeric systems allows the partial or total degradation of the structure of the container, resulting in the release of the pre-encapsulated content.

Polymer chains containing *o*-nitrobenzyl ester units along their backbone have thus been used to form polymer micelles,²⁰⁰ polymersomes,²⁰¹ or polymer microspheres,²⁰² that can be rapidly dismantled upon light irradiation. Photo-cleavable cross-links based on *o*-nitrobenzyl ester units have also been used in colloidal gels,¹¹² or polymer capsules,²⁰³ enabling the light-induced release of pre-encapsulated payloads including proteins. Photo-cleavable groups can also be used as “capping” agents that can be removed on-
demand to change the properties of polymer chains, such as the hydrophilicity, or to trigger chain reactions. For instance, Sun et al. showed that the photo-cleavage of *o*-nitrobenzyl ester groups in polymersome membranes could trigger the hydrophobic-to-hydrophilic transition of the bilayer concomitantly to *in situ* cross-linking that prevents any denaturation of the scaffold. This photo-activation eventually resulted in an increase of the membrane permeability and a subsequent release of the pre-encapsulated payload.²⁰⁴

Photo-cleavable groups have also been used as triggers for the depolymerization of “self-immolative” polymers. These polymers are designed to be crumbled by a succession of intramolecular reactions, provided an initial fragmentation trigger. Photo-cleavable units have been introduced for this purpose, as side-groups or end-groups on the polymer backbone, to promote the self-immolative degradation of polymer microspheres,²⁰⁵ or polymersomes,²⁰⁶ upon light irradiation (see **Figure 2.19**).

<figure><img src="image_32.png" /><figcaption>**Figure 2.19.** Schematic illustration of self-immolative polymersomes self-assembled from poly(benzyl carbamate)-b-poly(N,N-dimethylacrylamide) (PBC-b-PDMA) amphiphilic block copolymers, containing hydrophobic poly(benzyl carbamate) blocks which are subjected to self-immolative depolymerization into small molecules upon cleavage of “capping” moieties (here perylen-3-yl, 2-nitrobenzyl or disulfide for stimulation with visible light, UV light, or reductive milieu, respectively).²⁰⁶</figcaption></figure>

Photo-cleavable groups have also been used in a diverted way to induce the burst of liquid-core polymer compartments. Instead of being incorporated in the polymer scaffold, photo-cleavable molecules can be directly loaded or dispersed in the lumen of vesicles or capsules upon encapsulation. The photo-degradation of these molecules may produce a high concentration of side-products in the core of the compartment, resulting in a drastic increase of the internal osmotic pressure. At some point, the stress exerted by this internal pressure on the polymer envelope may lead to its burst. This approach has been successfully
implemented in polymersomes,²⁰⁷ and polymer capsules.²⁰⁸ In the latter case, the photo-cleavable molecules were designed to release CO₂ upon fragmentation. The pressure-induced rupture of the polymer shell wall was thus attributed to the rapid expansion of gas bubbles in the core of the capsules.

### 2.3.4 Combining photo- and thermo-sensitivity with plasmonic nanoparticles

The design of polymers that are intrinsically light-responsive can turn out to be complicated as it may require complex synthesis, and complete isolation from parasitic light to avoid sample degradation. In comparison, thermoresponsive polymers are easier to manipulate and can even be commercially available. Being able to stimulate a thermoresponsive scaffold with light as a primary trigger may thus enable to combine the high spatio-temporal control offered by light with the versatility offered by thermoresponsive polymers. For that purpose, plasmonic nanoparticles are interesting formulation additives as they can convert light excitation into heat generation as described in the following.

At the nanoscale, noble metal particles (essentially silver and gold) exhibit a very specific optical resonance phenomenon called localized surface plasmon resonance. Upon light excitation at an appropriate wavelength, collective electron oscillations are produced at the surface of the nanoparticles (see **Figure 2.20.A**). The damping of these oscillations generates heat (by Joule effect) in the vicinity of the nanoparticles with a typical time scale of 10 ns,²⁰⁹ i.e. quasi instantaneously. Noble metal nanoparticles can thus be seen as efficient light-to-heat converters at the nano- and microscales, and have paved the way to the development of a new field called “thermoplasmonics”.²¹⁰

Gold nanoparticles (AuNPs) are particularly interesting plasmonic structures since their plasmon resonance occurs in the visible wavelength range (400-800 nm). More interestingly, the spectral location of this resonance can be tuned by changing the size and morphology of the particles (see Figure 2.20.B).²¹¹ For instance, the excitation wavelength for gold nanospheres is typically around 530 nm (in the green), whereas it is red-shifted towards near infrared for gold nanorods (the shifting amplitude being roughly proportional to the aspect ratio of the nanoparticles).²¹² Gold nanorods are thus good candidates for biological applications of thermoplasmonics since the near infrared region is a “transparent” window for biological tissues (compared to the rest of the visible range).¹⁹²
<figure><img src="image_33.png" /><figcaption>**Figure 2.20.** A) Schematic representation of the thermoplasmonic effect in a gold nanoparticle.<sup>209</sup> B) Typical absorption spectra of gold nanoparticles with different morphologies.<sup>213</sup></figcaption></figure>

For the last 15 years, many efforts have been made to design photo-responsive encapsulation systems based on thermoplasmonic excitation. A first strategy has been focused on the photothermal destabilization of microcompartments doped with AuNPs. Upon thermoplasmonic excitation, sufficiently high temperatures can be reached to trigger the rupture (or at least the poration) of the compartment membrane/shell as it has been demonstrated in liposomes,<sup>214</sup> polymersomes,<sup>215</sup> or multi-layer polymer capsules.<sup>216,217</sup> Another approach has been based on the change of permeability of the container induced by gold nano-heaters. For instance, the temperature rise induced by irradiated AuNPs has been used to trigger glassy-to-rubbery transition in polyester microcapsules,<sup>218</sup> therefore leading to a higher diffusion of the payload across the capsule shell.

AuNPs have also been coupled with intrinsically thermoresponsive polymers in hybrid compartments. The first systems were based on LCST polymers, essentially PNIPAM. Amstad et al. have designed a polymersome containing PNIPAM-based copolymers and AuNPs embedded in the membrane.<sup>175</sup> Upon light irradiation, the heat generated by AuNPs triggers the hydrophilic-to-hydrophobic thermal transition of PNIPAM blocks, resulting in the destabilization and eventually the rupture of the vesicle within a few minutes (see **Figure 2.21**). Hybrid PNIPAM-AuNPs colloidal gels have also been designed, either by incorporation of AuNPs in preformed gels,<sup>219</sup> or by formation of the polymer network around AuNPs used as a template.<sup>220</sup> In both cases, light irradiation was able to trigger the thermal collapse of the PNIPAM matrix and the subsequent release of molecules which were entrapped in the gel network (via a “sponge squishing” mechanism). PNIPAM-AuNPs hybrid nanogels were also used as photo-responsive gates in amorphous shells to modulate the permeability of polymer capsules upon light irradiation.<sup>221</sup>
<figure><img src="image_34.png" /><figcaption>**Figure 2.21.** Photo-responsive polymersomes based on thermoplasmonic effects of gold nanoparticles (AuNPs). The membrane of the polymersomes is composed of a bilayer of PEG-b-PLA and PNIPAM-b-PLGA diblock copolymers, embedded with dodecylthiol-stabilized AuNPs. Upon light irradiation, AuNPs generate heat that trigger the thermal transition of PNIPAM blocks from hydrophilic to hydrophobic, leading to the destabilization of the polymersome membrane and the release of the pre-encapsulated content.<sup>175</sup></figcaption></figure>

In contrast, only a few examples of light-responsive nano-carriers combining UCST polymers and AuNPs have been proposed.<sup>222,223</sup> As an alternative to AuNPs, organic dyes that are also able to generate heat by energy dissipation upon light irradiation have been used as photothermal agents in most of the UCST-based photo-sensitive systems that have been reported in the literature (see **Figure 2.22**).<sup>224-226</sup> However, these dyes usually exhibit lower light-to-heat conversion efficiency and thermo-stability than metal plasmonic NPs.

<figure><img src="image_35.png" /><figcaption>**Figure 2.22.** Schematic illustration on the preparation of UCST polymer-based drug delivery system, in which ICG and DOX (the photothermal agent and anticancer drug, respectively) are simultaneously preloaded within a micelle of poly(NAGA-co-BA) (the UCST polymer) followed by coating with red blood cell (RBC) membrane via a vesicle–micelle fusion process. The resulting nanoparticle, RUID, exhibits significantly reduced drug self-leakage but responsive drug release upon irradiation with a near-infrared laser.<sup>224</sup></figcaption></figure>
### 2.3.5 The issue of release in the second or sub-second range

For some specific applications, the release of a high dose of bioactive molecules can be required to circumvent dilution effects that may occur upon delivery *in vivo*. It may be the case for the local activation of cell receptors or the local stimulation of neuron networks. For such applications, being able to deliver high amounts of molecules in a few ms can be advantageous to actuate biological targets with a high spatial resolution. Unfortunately, only a few systems have been designed to undergo on-demand *burst* release of their contents. This is essentially due to the fact that sudden structural changes in soft nano-containers are usually hindered by long relaxation times and/or multiple local reorganizations of the system.

Note that the term of “burst release” should be used cautiously since, in the jargon of drug delivery, the “burst release effect” corresponds to a sudden and early release of a fraction of the payload immediately after the introduction of the carriers *in vivo*, and by extension in any biological conditions.²²⁷ This “burst release effect” is by definition not controlled: it occurs accidentally at relatively short time scales and, most of the time, far from the biological targets. In addition, it leads to a loss of the payload and a shortening of the controlled release period. That is why many efforts have been made to reduce this undesired effect, mostly by tuning the structure and permeability of the encapsulating scaffold. In the following the notion of burst release will only refer to the *on-demand* sudden release of pre-encapsulated content, i.e. voluntarily triggered upon precise stimulation.

Most of the effective burst-release systems have been based on the increase of the internal pressure of hollow containers until sudden rupture of their rigid membrane/shell. This mechanical effect can be mediated by the formation of gas bubbles in liquid cores, either by ultrasound-induced evaporation of a volatile solvent (such as perfluoroalkanes),²²⁸ or by light-induced production of CO₂ from photo-degradable molecules.²⁰⁸ Another approach consists in increasing the internal osmotic pressure by generating high concentrations of solutes in the core, as previously mentioned in section 2.3.3. For instance photo-degradable molecules have been loaded in the core of liposomes,²²⁹ or polymersomes,²⁰⁷ to induce the rupture of the membrane upon light irradiation in less than 1 s (see **Figure 2.23**). As an alternative to photo-stimulation, Zhang et al. showed that asymmetric polymer capsules can undergo rapid shell fracturing when placed in hypotonic conditions (lower osmotic pressure outside the capsule than inside), as a consequence of water infiltration through the thinnest part of the shell wall.²³⁰
<figure><img src="image_36.png" /><figcaption>**Figure 2.23.** a) Light-induced rupture of polymersome membrane by sudden increase of internal osmotic pressure triggered by photo-degradation of encapsulation light-sensitive molecules. b) Photocleavage of coumarin derivative under irradiation and corresponding absorption spectrum in aqueous solution before (solid line) and after (dashed line) 30 min of irradiation at 365 nm. c) Confocal observation of a coumarin-loaded GUV (green channel, emission range of coumarin, 485 nm). The vesicle undergoes a fast (few ms) rupture upon 50-mW irradiation at 405 nm (scale bar = 10 mm).²⁰⁷</figcaption></figure>

On-demand burst release can also be induced by an intrinsic perturbation of the polymer scaffold of the container upon external stimulus. Mabrouk and al. showed that polymersomes containing azobenzene-based liquid crystal moieties can experience a burst curling of the membrane in less than 1 s upon light irradiation.¹⁹⁹ This process is induced by a dramatic change in the membrane curvature induced by the photo-isomerization of the azobenzene moieties. Hybrid polymersomes combining AuNPs and thermoresponsive polymers such as PNIPAM can also undergo rapid disassembly upon plasmonic excitation of AuNPs.¹⁷⁵

The previous examples highlight the fact that light is an optimal trigger for controlled burst release since the stimulation is instantaneous. The efficiency of the bursting is then only dictated by the responsiveness of the chemical components of the nano-container. In this context, vesicles turn out to be the most efficient systems since a small mechanical perturbation of the membrane can lead to a dramatic disruption of the entire structure at short time scales, typically below 1 s. Alternative systems relying on temperature or pH triggering have also been proposed for rapid release but were less efficient in terms of temporal response than light-sensitive systems.²³¹,²³²
To the best of our knowledge, this issue of controlled burst release has not been extensively explored for protein delivery. However, the sudden release of high amounts of proteins may be interesting in many biological applications. For instance, the delivery of neuropeptides at specific neuronal interfaces with a high spatio-temporal control may be helpful to decipher specific neuronal pathways. This topic of controlled burst release of proteins combines two challenges that have been mentioned throughout this chapter: the encapsulation of proteins without altering their stability, and their on-demand release in the sub-second range.

## 2.4 Objective of the PhD work

The objective of this PhD work was to develop a chemical platform to design stimuli-responsive and functional polymer capsules suitable for the versatile encapsulation and on-demand (including burst) release of proteins or other fragile biomacromolecules. **Table 2.1** sums up the main advantages and drawbacks of the different hydrophilic encapsulation systems presented throughout this chapter (polymersomes, microgels, capsules) regarding the encapsulation of fragile biomacromolecules. Regarding this information and the whole state of the art presented in this chapter, we could then imagine a promising formulation approach.

<table><caption>**Table 2.1.** Main advantages and drawbacks of the different hydrophilic encapsulation systems presented in this chapter, regarding the encapsulation and release of fragile biomacromolecules.</caption><thead><tr><th></th><th>Polymersomes</th><th>Microgels</th><th>Capsules</th></tr></thead><tbody><tr><th>Advantages</th><td><ul><li>Core-shell structure</li><li>Robustness</li></ul></td><td><ul><li>Chemical and physical tunability/functionalities</li><li>Mechanical properties</li></ul></td><td><ul><li>Core-shell structure</li><li>Chemical and physical tunability/functionalities</li><li>Mild preparation methods accessible</li></ul></td></tr><tr><th>Limitations</th><td><ul><li>Use of organic solvents/chemical reactions</li><li>Limited encapsulation yield</li></ul></td><td><ul><li>Use of organic solvents/chemical reactions</li><li>Polymer/protein interactions</li><li>Slow release kinetics</li></ul></td><td><ul><li>Colloidal stability</li></ul></td></tr></tbody></table>
To limit potentially undesired interactions between the proteins and the container scaffold, reservoir-like structures with an aqueous core should be favored. Along this line, we decided to focus on aqueous-core polymer capsules. The use of chemical reactions should be limited during the formation of the capsule shell to avoid potential degradation of the payloads. For that purpose, the self-assembly of preformed polymer chains is an interesting approach and we decided to explore the strategy of interfacial complexation. The diversity of the polymer chains that can be used in interfacial complexation is also an advantage regarding the design of functional, especially stimuli-responsive, capsules.

The formulation of aqueous-core polymer capsules by interfacial complexation requires emulsion droplets as a template. As previously mentioned, the presence of organic solvents in water-in-oil emulsions can induce interfacial aggregation of the proteins that can impair their biological activity. The use of water-in-water emulsions is interesting but may suffer from limitations in encapsulation yield due to partitioning issues between the two aqueous phases. As an alternative, the use of fluorinated oils (FO) as the continuous phase of water-in-oil emulsions is a promising approach for several reasons: first, as aqueous and hydrocarbon species show an extremely poor solubility in fluorinated oils,²³³ the issue of phase partitioning as well as molecular cross-talks between droplets are limited in water-in-FO emulsions,²³⁴,²³⁵ suggesting that high yields of encapsulation may be achieved in the aqueous core of the capsules. Second, fluorinated oils are more biocompatible than their hydrocarbon analogs.²³⁶,²³⁷ Fluorinated solvents have even been considered as promising candidates for the design of artificial blood substitutes, thanks to their high capacity to solubilize oxygen.²³⁸ Issues of protein aggregation at water/FO interfaces have been addressed and partially solved with the utilization of suitable fluorinated surfactants.²³⁹⁻²⁴¹ Finally, fluorinated oils are compatible with the microfluidic technologies, especially with poly(dimethyl siloxane) (PDMS) devices as they do not induce PDMS swelling contrary to common hydrocarbon oils. The phenomenon of interfacial complexation in water-in-FO has not been extensively studied for the design of polymer capsules and we proposed to explore this strategy during this PhD, whose main results are presented and discussed in the following chapters.
# Chapter 3

## Formation of stimuli-responsive polymer shells by interfacial complexation: poly(N-isopropylacrylamide) shells as a first example

**Note:** This chapter contains results that have been published in the following peer-reviewed article:

L. Sixdenier, C. Tribet, E. Marie. Emulsion-Templated Poly(N-Isopropylacrylamide) Shells Formed by Thermo-Enhanced Interfacial Complexation. *Adv. Funct. Mater.* **2021**, *2105490*. https://doi.org/10.1002/adfm.202105490

To clarify the research approach and to make the results more comprehensive for a PhD manuscript, the content of this article has been reorganized and enriched with complementary, non-published results.
## 3.1 Introduction

According to the specifications set at the end of Chapter 2, we will describe in this Chapter the experimental approach that we developed to design stimuli-responsive polymer microcapsules by interfacial complexation in water-in-fluorinated oil (FO) emulsions.

Interfacial complexation in water-in-FO emulsions (W/FO) has not been extensively explored but has been well characterized in a representative system by DeJournette et al.²⁴² They evidenced the formation of a complex layer by polymer-surfactant interfacial complexation at the edge of water droplets between a water-soluble diamine (Jeffamine) and a FO-soluble macrosurfactant (a carboxylate-terminated perfluoro-poly(ethylene glycol), perfluoro-PEG, called Krytox) (see **Figure 3.1**). The tight interactions between Jeffamine and Krytox were evidenced at the molecular level (using infrared spectroscopy, mass spectrometry and NMR), and *in operando* at the water/oil interface in inverse emulsion (using fluorescence microscopy). These interfacial interactions are likely due to a combination of electrostatic and hydrogen bonding between amino groups of the polyamine and carboxylic groups of Krytox, as previously described by O'Neal and al. on analogous systems.²⁴³,²⁴⁴

<figure><img src="image_37.png" /><figcaption>**Figure 3.1.** Interfacial complexation in water-in-fluorinated oil emulsion between a cationic diamine (Jeffamine ED 900) present in water droplets and an anionic surfactant (Krytox 157 FSH) present in the continuous fluorinated oil phase.²⁴²</figcaption></figure>

More recently, Abell and co-workers used a similar polymer-surfactant interfacial complexation between a cationic polymer and Krytox to favor the formation of supramolecular microcapsules. They showed that interfacial complexation enables to increase the concentration of the polycation at the water/FO interface, and subsequently to favor supramolecular cross-linking of adjacent chains by cucurbituril units to form thin hydrogel shells (see **Figure 3.2**).²⁴⁵,²⁴⁶
<figure><img src="image_38.png" /><figcaption>**Figure 3.2.** A) Microfluidic-based formation of supramolecular hydrogel microcapsules by host-guest cross-linking between anthracene-functionalized hydroxyethyl cellulose (Ant-HEC) and cucurbit[8]uril (CB[8]) units. B) The cross-linking is enabled by an increase of the Ant-HEC concentration at the interface of a W/FO emulsions via interfacial complexation between cationic Ant-HEC in water and anionic Krytox in FO (namely FC-40).²⁴⁶</figcaption></figure>

Inspired by these examples, we decided to implement polymer-surfactant interfacial complexation in W/FO emulsions between cationic poly(L-lysine) (PLL) – which is nothing else than a polyamine – and anionic Krytox. PLL has the advantage to be biocompatible and to be easily functionalizable by reactions involving its side amino groups, giving access to a wide library of PLL-based functional copolymers.²⁴⁷ In a first approach, we prepared a LCST PLL-based copolymer by grafting LCST PNIPAM strands onto PLL. We demonstrated that the resulting PLL-g-PNIPAM comb-like copolymer leads to the formation of thermos-responsive shells at the surface of water droplet in W/FO emulsions.
## 3.2 Evidence for interfacial complexation of PLL comb-like derivatives in W/FO emulsions

### 3.2.1 Measurement of water/FO interfacial tension by the rising drop method

As a complement to the experimental evidence of polyamine-Krytox interactions reported by DeJournette and O'Neal, we proposed to specifically reveal the PLL/Krytox complexation by interfacial tension measurements. To be close to the experimental conditions of an inverse emulsion, a water droplet containing PLL was grown in a bath of fluorinated oil FC-40 containing Krytox and the water/oil interfacial tension was deduced from the shape of the droplet (see Appendix A for experimental details). Since the aqueous phase is less dense than the fluorinated oil phase (density 1.0 vs 1.9), the experiments were performed using the rising droplet configuration (see Figure 3.3.A).

The complexation between cationic amino groups of PLL and anionic carboxylate groups of Krytox is expected to occur at the surface of the droplet as the Krytox molecules may deprotonate at the contact with water. Note that in bulk fluorinated oil, Krytox molecules are likely under the form of dimers connected via hydrogen bonds between carboxylic acid groups, as it has been previously reported for other perfluoro-carboxylic acids in fluorinated solvents.²⁴⁸ This is likely due to the commonly low solubility of H⁺ in fluorinated solvents and to the low basicity of fluorinated amines such as the constituents of the FC-40 oil.²⁴⁹ Having that in mind and in order to limit the complexity of the schematics, Krytox molecules will always be represented under its deprotonated monomer form – even in bulk oil – in the schematics of this manuscript, as in Figure 3.3.A.
<figure><img src="image_39.png" /><figcaption>**Figure 3.3.** Water/fluorinated oil (W/FO) interfacial tension measurements using the rising drop method. A) Schematics of the rising drop configuration in the case of interfacial complexation between PLL present in the water droplet and Krytox surfactant molecules present in the continuous oil phase made of FC-40 fluorinated oil. B) Evolution of the W/FO interfacial tension as a function of the Krytox concentration in FO for different PLL concentrations in water (error bars = standard errors on 6 droplets per condition). The dashed lines are guides for the eyes. C) Representative images of rising droplets for the conditions indicated with a letter in the graph.</figcaption></figure>

Figure 3.3.B shows the evolution of the water/oil interfacial tension as a function of the Krytox concentration in oil for a given PLL concentration in water. The dashed lines are arbitrary guides for the eyes and are not fitting curves taken from any theoretical model.

For droplets containing pure water, the water/oil interfacial tension was constant around 37 mN/m for Krytox concentrations below 0.04 wt% in the oil phase, and showed a slight decrease to ~30 mN/m for Krytox concentrations higher than 0.05 wt%. Considering these raw data, a critical micellar concentration (CMC) of 0.05 wt% for Krytox in FC-40 may be deduced, i.e. 200 µM considering the molar
mass of 2.5 kDa for Krytox. However, considering the experimental limitations for the measurement of the interfacial tension (see **Appendix A**), this value of CMC should be taken with caution.

In her PhD manuscript, Birte Riechers reported the same experimental issues concerning the reliability of water/FO interfacial tension measurements using the pendant drop method.²⁵⁰ As an alternative, she used the Wilhelmy plate method to measure more precisely the CMC of Krytox and eventually found a value of 4 µM,²⁵¹ i.e. 0.001 wt%. This value is 50 times lower than the approximate value we obtained with the rising drop method. Note that Riecher used a different fluorinated oil (Novec™ 7500 instead of Fluorinert™ FC-40), what could partially explain the discrepancy in the CMC ranges.

As a complement, Wagner et al. measured the CMC of Krytox derivatives for which the polar head was functionalized with different linear polyglycerol moieties²⁴¹. The pendant drop method gave CMC values between 0.01 and 0.05 wt%, what is more consistent with our result. Nevertheless, the experimental conditions in Wagner's work were different from ours as (i) the measurements were performed using the pendant (and not rising) drop method for which the phases were inverted (an oil droplet was suspended in a water bath), (ii) the fluorinated oil was different (Novec 7500 instead of FC-40), and (iii) the chemical modification of Krytox polar head can drastically change the adsorption properties of the surfactant.

Considering all the variations in the experimental approaches, the comparison between all the CMC values must be taken cautiously and we will conclude this discussion saying that the Krytox CMC is probably below 0.05 wt% in FC-40 and analogous fluorinated oil (such as FC-70).

When PLL was added in the aqueous phase, the evolution of the interfacial tension with Krytox concentration was significantly different from pure water. The interfacial tension fell quickly from 50 mN/m in pure oil to 10 mN/m in oil containing 0.04 wt% Krytox, and that for all the PLL concentrations tested (in the 0.1-0.5 wt% range). The first remarkable observation is that the interfacial tension in pure oil was higher for PLL solutions compared to pure water (53 vs 37 mN/m), likely because PLL participates to the hydrogen-bond network within the aqueous phase, thus contributing to increase the energetic cost of creating water/oil interface.

As soon as Krytox was present in the oil phase, the interfacial tension drastically decreased, suggesting that a strong PLL/Krytox interaction developed across the water/oil interface. From a Krytox concentration of 0.02 wt% and above, the stabilization of the droplets (before detachment from the needle tip) was particularly arduous due to the low interfacial tension (see droplet (c) in Figure 3.3.B). At concentration higher than 0.04 wt% the formation of well-defined droplets was even impossible as the oil phase was forming a quasi-continuous flow out of the needle upon injection. This qualitative observation confirms the significant decrease of interfacial tension when more than 0.05 wt% of Krytox was added in the oil phase. The dashed lines used in the graph of Figure 3.3.A do not correspond to any theoretical fit
but arbitrarily indicates the drastic diminution of the interfacial tension, probably plateauing around 5 mN/m at higher Krytox concentrations. This observation qualitatively supports the idea of a tight interfacial complexation between PLL and Krytox in emulsion-like configuration, as previously described in the introduction of this chapter.

By taking advantage of the PLL/Krytox complexation across water/FO interfaces, we proposed to form polymer shells at the surface of water droplets by modifying the PLL backbone with polymer chains of interest (assuming that the modification of PLL would not drastically change its complexation capability). In the following, the term of “oil” will only refer to fluorinated oil and the adjective “fluorinated” will be omitted for the sake of simplicity.

### 3.2.2 Synthesis of PLL comb-like derivatives: PLL-g-PNIPAM and PLL-g-PEG

Given the propensity of PLL to adsorb at the water/oil interface when combined with Krytox, we decided to implement interfacial complexation with PLL derivatives in order to create functional polymer layers at the surface of water droplets dispersed in the oil phase. We were particularly interested in developing thermoresponsive polymer shells in this inverse emulsion configuration as they can be seen as precursors of aqueous-core polymer capsules for controlled delivery upon temperature changes. In a first approach, the strategy was tested and optimized with the LCST-type poly(N-isopropylacrylamide) (PNIPAM, already mentioned in section 2.3.2 of Chapter 2). The idea was to graft PNIPAM chains onto PLL and to accumulate the so-formed comb-like copolymer at the surface of water droplets by interfacial complexation of the PLL backbone with Krytox. The expected result was the formation of droplet-templated PNIPAM shells that can change of physicochemical properties upon heating.

We took advantage of the presence of side amine groups on PLL to easily design comb-like copolymers by the “grafting to” strategy. As amine groups can be involved in nucleophilic addition on activated ester groups, we used PNIPAM chains terminated with a highly reactive N-hydroxysuccinimide (NHS) ester to promote the formation of amide bonds between PLL and PNIPAM. The resulting comb-like copolymer comprising a PLL backbone and PNIPAM macrographs was denoted PLL-g-PNIPAM. Its architecture is represented in **Figure 3.4**. The same strategy was adopted to graft poly(ethylene glycol) (PEG) chains on PLL, resulting in a PLL-g-PEG comb-like copolymer. As PEG chains are hydrophilic and non-
thermoresponsive, PLL-g-PEG was used as a “control” during the characterization of the emulsion-based polymer shells.

<figure><img src="image_40.png" /><figcaption>**Figure 3.4.** Chemical structure of the poly(L-lysine)-g-poly(N-isopropylacrylamide) (PLL-g-PNIPAM) and poly(L-lysine)-g-poly(ethylene glycol) (PLL-g-PEG) comb-like copolymers used in this chapter. The grafting approach was based on the formation of amide bonds between side amines of PLL and N-hydroxysuccinimide-activated esters (NHS) end-groups of the polymer strands to graft.</figcaption></figure>

Note that the number of repeat units have not been specified in the polymer structures represented in Figure 3.4, especially because a repeat unit is not easy to define in a randomly grafted comb-like copolymer. The exact composition of the comb-like derivatives is detailed in the following: the backbone was made of a polydisperse PLL with a molar mass of 15-30 kDa (72-144 lysine units/chain), the mean mass being 22 kDa ($DP_{n,PLL}$ = 105 lysine units/chain). Both PNIPAM and PEG macrografts had a molar mass of 2 kDa (72 NIPAM units/chain for PNIPAM, and 144 EG units/chain for PEG). The grafting ratio was typically of 0.22 for the two comb-like copolymers, meaning that around 1/5 of the lysine units were functionalized with a macrograft (see **Appendix A** for experimental details, and **Table 3.1**).

<table><caption>**Table 3.1.** Typical characteristics of a representative PLL-g-PNIPAM (or PLL-g-PEG) copolymer.</caption><thead><tr><td></td><td>PLL DP<sub>n</sub></td><td>Grafting ratio τ</td><td>Copolymer molar mass</td></tr></thead><tbody><tr><td>PLL-g-PNIPAM</td><td>~ 105</td><td>~ 0.22</td><td>~ 68 kg/mol</td></tr></tbody></table>
During the synthesis of PLL comb-like derivatives, a fluorophore possessing an NHS-ester reactive group could be added to label the PLL backbone. This dye-functionalization was used to localize the polymer in emulsion and to characterize its properties by fluorescence imaging, especially with confocal microscopy. Two different fluorophores were used (see **Figure 3.5**): a carboxy-X-rhodamine (simplified to “rhodamine”) and a diethylamino-coumarin (simplified to “coumarin”). The first one absorbs in the yellow range (575 nm) and emits in the orange range (600 nm), the second absorbs in the dark blue range (445 nm) and emits in the light blue range (482 nm).

<figure><img src="image_41.png" /><figcaption>**Figure 3.5.** Chemical structure of the rhodamine and coumarin fluorophores covalently attached to PLL derivatives for confocal imaging. Rhodamine-NHS = 5(6)-carboxy-X-rhodamine N-succinimidyl ester, coumarin-NHS = 7-(diethylamino)coumarin-3-carboxylic acid N-succinimidyl ester.</figcaption></figure>

For some control experiments, free PNIPAM chains (non-grafted on PLL) were used in the aqueous phase of the emulsions. In order to localize these chains by fluorescence microscopy, they were also functionalized with a fluorophore. This labeling was performed by strain-promoted azide–alkyne cycloaddition (a “click chemistry” technique) between azide-terminated PNIPAM chains and dibenzocyclooctyne-cyanine 3 (see **Appendix A** for experimental details). The Cy3 fluorophore absorbs in the green-yellow range (553 nm) and emits in the yellow range (569 nm).

Before considering the possibility to form thermoresponsive polymer capsules made of PLL-g-PNIPAM, the first step was to verify if interfacial complexation between the PLL comb-like derivatives and Krytox was effective in water-in-oil emulsions.
### 3.2.3 Stabilization of water-in-oil emulsions by interfacial complexation of PLL derivatives

The effectivity of interfacial complexation between PLL comb-like derivatives and Krytox was directly assessed *in operando* in water-in-oil emulsions. All these experiments have been carried out with PLL-g-PNIPAM as a representative type of PLL comb-like derivatives. We qualitatively compared the stability of emulsions with and without PLL-g-PNIPAM in the aqueous phase to evidence the formation of polymer/surfactant complexes across the water/oil interface.

In this experiment, as well as throughout all this manuscript, the water-in-oil emulsions were obtained as follows: 5 µL of aqueous phase were manually emulsified in 95 µL of oil phase in a 0.5 mL Eppendorf tube. If not otherwise noticed, the aqueous phase was prepared in a 7.7 mM phosphate buffer at pH = 7.3 (see Appendix A) and the oil phase was composed of FC-70 fluorinated oil containing Krytox (FC-70 is an analog of FC-40).

To assess the role of interfacial complexation in the stability of the emulsions, two emulsions were compared: E1 containing 1 wt% of coumarin-labeled PLL-g-PNIPAM in the aqueous phase and E2 containing no PLL-g-PNIPAM but 50 µM of free coumarin in the aqueous phase, both with 0.03 wt% of Krytox in the oil phase. The coumarin staining of both aqueous phases was used to better distinguish the two phases under UV light.

**Figure 3.6** shows the aspect of both emulsions under UV light, a few seconds after emulsification. Directly after emulsification both samples became turbid, indicating the formation of an emulsion. Since the aqueous phase is less dense than the oil phase, the water droplets rapidly creamed on top of the oil phase. For E1 the homogenous turbidity was recovered upon gentle stirring of the sample. This easy redispersion of the water droplets is an indication of the stability of the emulsion. For E2, additionally to the creaming process, signs of demixing were observed a few seconds after emulsification. An aqueous supernatant rapidly formed on top of the oil phase, and could not be readily redispersed upon gentle stirring, suggesting that the water droplets were strongly subject to coalescence. The difference in stability between E1 and E2 was highlighted under UV light thanks to the coumarin-staining of the aqueous phase (see Figure 3.6.B and C).
<figure><img src="image_42.png" /><figcaption>**Figure 3.6.** Stability of water-in-oil emulsions in the presence or absence of 1 wt% of PLL-g-PNIPAM in the aqueous phase (emulsion E1 and E2 respectively). For both emulsions, the oil phase contained 0.03 wt% of Krytox. A) Schematic representation of the PLL-g-PNIPAM copolymer and the Krytox surfactant. B) Emulsion E1: in the presence PLL-g-PNIPAM in water, a stable emulsion was formed as indicated by its uniform turbidity under UV light (the fluorescence is due to the coumarin-labeling of the polymer). C) Emulsion E2: In the absence of PLL-g-PNIPAM, the water droplets formed upon emulsification rapidly creamed and coalesced, leading to the destabilization of the emulsion as highlighted under UV light (the fluorescence is due to free coumarin added in water for staining).</figcaption></figure>

The high stability observed in E1 compared to E2 can be attributed to the formation of tight interactions between PLL and Krytox across the water/oil interface. As previously described in section 3.2.1, this interfacial complexation is responsible for a sharp decrease of the water/oil interfacial tension and thus facilitates the dispersion and stabilization of the water droplets in the fluorinated oil.

In addition to this qualitative evaluation, the issue of emulsion stabilization by interfacial complexation was quantitatively addressed with dynamic light scattering experiments. More precisely, we assessed the influence of the PLL and Krytox concentrations on the size of the emulsion droplets produced upon interfacial complexation. The results are presented in **Appendix B**, and confirm that a minimal concentration of Krytox and PLL are required to obtain stable droplets.
## 3.3 Microscopy imaging of emulsions

### 3.3.1 Stability of emulsions injected in capillary for microscopy experiments

Microscopy imaging of the droplets was performed on emulsions injected in sealed rectangle glass capillaries. This device has been preferred to classical slide mounting to avoid any mechanical breakage of the droplets during sample preparation, or any hydrodynamic flows that could destabilize the emulsions during the observation. Since PLL derivatives are positively charged in relevant experimental conditions (neutral pH), they may spontaneously and strongly adsorb onto anionic surfaces including bare glass.²⁴⁷,²⁵² To prevent PLL derivatives present in the water droplets from adsorbing on the capillary sides, a surface treatment of the capillary was performed prior to the injection of the sample emulsion. Taking advantage of the adsorption properties of PLL, a solution of PLL-g-PEG was injected and dried in the capillary to form a “cushioning” polymer monolayer on the internal side of the capillary (see **Figure 3.7**). The presence of hydrophilic PEG strands pointing out of the surface would thus prevent any breakage of the water droplets due to the adsorption of PLL derivatives used in the emulsion.

An emulsion containing 2 wt% of PLL-g-PNIPAM in the aqueous phase and 0.05 wt% of Krytox in the FC-70 oil phase was prepared as described in section 3.2.3 (see also **Appendix A**), and injected either in a bare glass capillary or in a glass capillary previously coated with PLL-g-PEG. As the aqueous phase is less dense than the oil phase, the water droplets were in contact with the upper internal side of the capillary. Figure 3.7 shows a bright field micrograph and a side-view schematic representation of the emulsion in each capillary.
<figure><img src="image_43.png" /><figcaption>**Figure 3.7.** Effect of the capillary coating on the stability of an injected emulsion containing 2 wt% of PLL-g-PNIPAM in the aqueous phase and 0.05 wt% of Krytox in the oil phase. Bright field micrographs (scale bars = 30 µm) and a side-view schematic representation of the samples are shown for the bare capillary (A) and for the capillary previously treated with PLL-g-PEG (B). The PLL-g-PEG coating is represented by the monolayer of comb-like molecules in the schematics of the treated capillary. For more readability, PLL-g-PNIPAM and Krytox molecules have not been represented.</figcaption></figure>

For the bare capillary, most of the droplets have popped and spread at the contact with the capillary side. The affinity of PLL-g-PNIPAM present in the droplets for the bare glass surface probably favored the disruption of the water/oil interface at the contact with the capillary, leading to the destabilization of the emulsion (see Figure 3.7.A). In contrast, in PLL-g-PEG-coated capillary, intact spherical droplets lying onto the upper internal side of the capillary could be observed (see Figure 3.7.B). The PLL-g-PEG monolayer previously adsorbed on the capillary internal sides likely acts as a “cushion” that keeps the droplets out of direct contact with the glass substrate. As this coating method efficiently preserved the stability of the emulsions in capillaries, it was implemented for all the microscopy samples that will be presented throughout this manuscript.

To evaluate the stability of the emulsions once injected in a capillary, micrographs of a representative sample were taken every hour for 5h. The micrographs were taken in phase contrast mode to highlight the edges of the droplets (see **Figure 3.8**). No bursting or coalescence events were noticed in the sample during the 5h of incubation at ambient temperature, suggesting that the emulsion remained intact at least at the time scale of a microscopy experiment.
<figure><img src="image_44.png" /><figcaption>**Figure 3.8.** Phase contrast micrographs over time of a water-in-oil emulsion with 2 wt% of PLL-g-PNIPAM in the aqueous phase and 0.05 wt% of Krytox in the oil phase (scale bar = 30 µm). The emulsion has been injected in a PLL-g-PEG-coated glass capillary.</figcaption></figure>

### 3.3.2 Confocal microscopy on emulsions and evidence for the formation of polymer shells by interfacial complexation

Confocal laser scanning microscopy has the great advantage to provide high spatial resolution (in the micron range) within the sample thickness. It would allow to probe the equatorial plane of the water droplets and thus to image the physico-chemical processes taking place *inside* the droplets, and especially at the inner surface. First, confocal imaging was used to evaluate the effectivity of surfactant-polymer interfacial complexation by comparing the distribution of fluorophore-labeled PLL-g-PNIPAM and “free” PNIPAM chains (i.e. non-grafted on PLL) in emulsion droplets.

The emulsion samples were prepared in capillary as described in the previous section (see also **Appendix A**). Briefly, 5 µL of aqueous phase containing either 3 wt% of rhodamine-labeled PLL-g-PNIPAM or 2.5 wt% Cy3-labeled free PNIPAM were emulsified manually in 95 µL of oil phase containing 0.05 wt% Krytox, and injected in a glass capillary pre-treated with PLL-g-PEG. The resulting emulsion samples were observed in confocal microscopy (see **Figure 3.9** and **Appendix A** for details).
<figure><img src="image_45.png" /><figcaption>**Figure 3.9.** Formulation of water-in-oil emulsions with free PNIPAM chains or PLL-g-PNIPAM copolymer in the aqueous phase and Krytox surfactant in the oil phase. A) Schematics of the emulsion and the expected distribution of polymer species inside water droplets. B) Confocal micrographs of emulsions containing either 2.5 wt% of Cy3-labeled free PNIPAM chains (left) or 3 wt% of rhodamine-labeled PLL-g-PNIPAM (right) in the aqueous phase (scale = 20 µm), and 0.05 wt% of Krytox in the oil phase (FC-70). C) Fluorescence radial profiles of the droplets encircled in white in the micrographs.</figcaption></figure>

In the emulsion containing 3 wt% of rhodamine-labeled PLL-g-PNIPAM and 0.05 wt% Krytox, the fluorescence of rhodamine was essentially localized at the edge of the water droplets, i.e. at the water/oil interface, as shown in Figure 3.9.B. In contrast, the fluorescence of Cy3-labeled PNIPAM free chains (in the absence of PLL) was homogenously distributed in the core of the water droplets (see Figure 3.9.B). The difference of distribution between free PNIPAM and PLL-g-PNIPAM is highlighted in the radial fluorescence profiles (see Figure 3.9.C), with a sharp peak localized at the droplet edge for PLL-g-PNIPAM
and a uniform level of fluorescence within the droplet core for free PNIPAM. The higher density of PLL-g-PNIPAM at the interfaces (as compared to both the droplet cores and the continuous oil phase) is indicative of the formation of an interfacial polymer layer by complexation between cationic PLL and anionic Krytox across the interface. In contrast, non-grafted PNIPAM chains were neutral and could not be involved in interfacial electrostatic complexation. As consequence, no accumulation of free PNIPAM was observed at the inner surface of the water droplets. Thus, confocal imaging gives visual evidence of PLL/Krytox interfacial complexation in emulsion and suggests that this process can be used to accumulate PNIPAM chains at the surface of water droplets.

For our confocal experiments, we worked at relatively low magnification (x 20) to image our emulsions with a large field of view and characterize a high number of droplets at the same time. In these conditions, the imaging of small objects (typically lower than 10 µm) was limited by the optical resolution. In the case of droplets containing rhodamine-labeled PLL-g-PNIPAM, the fluorescent corona could not be distinguished at the periphery of droplets smaller than 10 µm in diameter. Instead, the fluorescence signal appeared uniform in the whole droplets because of a lack of resolution. As a consequence, the analysis of all the physico-chemical processes relying on fluorescence signal was restricted to droplets larger than 10 µm in diameter. **Figure 3.10** shows the distribution of droplet size based on the measurement of 130 droplets from the emulsion containing 3 wt% of rhodamine-labeled PLL-g-PNIPAM showed in Figure 3.9, and used as a representative sample of manually-prepared emulsion. Almost 2/3 of the droplets have a diameter smaller than 10 µm. Consequently, image analysis was performed on the other 1/3 of the sample.

<figure><img src="image_46.png" /><figcaption>**Figure 3.10.** Droplet size distribution for 130 droplets of the emulsion showed in **Figure 3.9**, containing 3 wt% of PLL-g-PNIPAM in the aqueous phase and 0.05 wt% of Krytox in the oil phase. The solid line corresponds to a lognormal fit.</figcaption></figure>
## 3.4 Characterization of PLL-g-PNIPAM shells formed upon interfacial complexation

### 3.4.1 Optimization of the formulation and observation of well-formed polymer capsules

For each batch of PLL derivative, the Krytox/PLL-g-PNIPAM concentration ratio was adjusted to achieve an optimal deposition of PLL-g-PNIPAM at the droplet surface by interfacial complexation, evidenced by (i) the stability of the emulsion and (ii) the formation of a fluorescent layer of PLL-g-PNIPAM at the droplet periphery. This optimization is illustrated in **Figure 3.11** in the case of a coumarin-labeled PLL-g-PNIPAM.

<figure><img src="image_47.png" /><figcaption>**Figure 3.11.** Partial formulation diagram of water-in-oil emulsions injected in a glass capillary and imaged by confocal microscopy (scale bar = 50 µm). PLL-g-PNIPAM was labeled with coumarin (in red) for fluorescence imaging.</figcaption></figure>

At low Krytox concentration (typically ≤ 0.01 wt%) the coalescence of droplets onto the capillary side revealed that the surfactant concentration was insufficient to stabilize the water droplets. At high Krytox concentration (typically ≥ 0.05 wt%), the fluorescence of coumarin-labeled PLL-g-PNIPAM was essentially observed in the continuous phase, indicating a massive extraction of polymer chains in the oil phase. As pure fluorinated oils do not solubilize the hydrophilic hydrocarbon-based PLL derivatives, this
observation suggests that the dispersion of these hydrophilic compounds in the oil was mediated by excess Krytox, likely under the form of inverse micelles.<sup>253,254</sup> At intermediate Krytox concentration (typically 0.03 wt%), fluorescence micrographs showed a bright corona of PLL-g-PNIPAM localized at the droplet edges and negligible fluorescence in the oil. This observation is consistent with a process of interfacial complexation where the PLL-g-PNIPAM copolymer is attracted towards the interface upon the formation of coulombic complexes with Krytox as schematized in Figure 3.9. At high amount of polymer in the aqueous phase (typically ≥ 0.05 wt%), the interface was saturated in PLL-g-PNIPAM, and excess polymer chains remained soluble within the droplet core, leading to a low fluorescence contrast between the edge and the core of the droplets. Because of the excessive amount of internal polymer content, this condition was not suitable for the formation of hollow polymer capsules.

In addition, scanning electron microscopy (SEM) imaging was performed to visualize the shell at a higher resolution and to determine the influence of the formulation on the capsule morphology. SEM imaging was performed on 5 µL of emulsion dried on a glass coverslip for three days at ambient air. Prior to imaging, the dried capsules were metalized by deposition of a 30-nm thin layer of Ag (see **Appendix A** for experimental detail). **Figure 3.12** shows SEM images of microcapsules prepared from an emulsion containing 1 wt% of either PLL-g-PNIPAM or PLL-g-PEG in the aqueous phase.

<figure><img src="image_48.png" /><figcaption>**Figure 3.12.** SEM imaging of dried emulsions showing the effective formation of polymer capsules by interfacial complexation. A) SEM micrographs of dried emulsions with 1 %wt of PLL-g-PNIPAM or PLL-g-PEG in the aqueous phase and 0.05 wt% of Krytox in the oil phase (scale bars = 20 µm). B) Thickness profiles of the shell folds indicated by colored arrows in the micrographs, giving an estimation of twice the apparent thickness of the capsule shells (as the folds are presumably formed by the collapse of two shell layers).</figcaption></figure>
SEM images on dried emulsions reveal the presence of hollow microcapsules, suggesting that drying-resistant polymer layers were initially formed at the periphery of the droplets. This result confirms that interfacial complexation is an effective technique for the formation of polymer capsules using an emulsion as a template. Their flattened aspect – presumably due to membrane collapsing during the drying step – allows for an estimate of the shell wall thickness, which is typically of 0.6 - 0.8 µm (the apparent thickness of a collapsed membrane being assumed to correspond to two stacked membrane layers, see Figure 3.12.B).

**Figure 3.13** details the influence of the polymer and Krytox concentrations on the capsule morphologies observed by SEM in the case of PLL-g-PNIPAM.

<figure><img src="image_49.png" /></figure>

<figure><img src="image_50.png" /></figure>

**Figure 3.13.** Influence of the PLL-g-PNIPAM and Krytox concentrations on the structure of PLL-g-PNIPAM capsules formed by interfacial complexation. A) SEM micrographs of dried emulsions containing 1 wt% of PLL-g-PNIPAM in the aqueous phase and an increasing concentration of Krytox in the oil phase. B) SEM micrographs of dried emulsions containing 0.05 wt% of Krytox in the oil phase and an increasing concentration of PLL-g-PNIPAM in the aqueous phase.
Figure 3.13.A shows that the concentration of Krytox has to be balanced to readily stabilize well-defined polymer capsules by interfacial complexation (at a fixed polymer concentration). At 0.05 wt% Krytox concentration, microcapsules can be observed with a typical 1 µm-thick shell wall. Their flattened aspect presumably comes from the drying step. At lower Krytox concentration (typically 0.01 wt%), no capsules were formed as not enough surfactant was present to trigger interfacial complexation. At higher Krytox concentration (typically 0.10 wt%), a messy phase composed of intertwined capsular structures, small aggregates and filaments was observed. The complexity of this phase and the low number of well-defined individual capsules suggests that the polymer was not distributed at the surface of the droplets in the initial emulsion but may have been extracted into the continuous oil phase by the excess of Krytox, which agrees with the high fluorescence level observed in the oil phase in confocal micrographs of Figure 3.11.

Figure 3.13.B shows the evolution of the capsule morphology with the concentration of PLL-g-PNIPAM in the aqueous phase, at a fixed concentration of Krytox in the oil phase. At 0.2 wt% in polymer, no rigid capsules were formed, and some circular spots observed in SEM suggest that the droplets were not stable to drying. At 1 and 2 wt%, microcapsules were obtained with a typical shell wall thickness of ~ 1 µm. At 5 wt%, closed microspheres were obtained instead of hollow capsules, suggesting that the high amount of polymer chains inside the droplets formed a bulky polymer mesh that probably hindered the polymer accumulation at the interface.

Cryogenic electron microscopy techniques would have undoubtedly enabled a more precise characterization of the polymer membranes as it would have given access to a much higher spatial resolution (down to the nm). A collaboration with microscopists from the Charles Sadron Institute (Strasbourg) who are specialists of these techniques has been launched to analyze our polymer capsules (prepared by ultrasonication to decrease the capsule diameter below 500 nm). Unfortunately, the first attempts of vitrification of the emulsion were unsuccessful and no suitable conditions were found to properly observe the capsules in cryo-SEM. Obviously, the fluorinated oil phase seems less adapted to this technique than the more common hydrocarbon oils.
### 3.4.2 Quantification of the polymer surface excess from fluorescence measurements

The previous results obtained by confocal imaging provided qualitative arguments in favor of the formation of a polymer shell by interfacial complexation but did not give any quantitative results concerning the fraction of polymer involved in the interfacial layer or the residual fraction of polymer that remained soluble in the core of the droplets. To access this type of information, we implemented a robust and reliable quantification method based on the measurement of the intrinsic fluorescence of dye-labeled PLL derivatives.

Taking advantage of the Beer-Lambert law relating fluorescence intensity to polymer concentration in solution, we determined the concentration in soluble polymer chains that were not involved in interfacial complexation by measuring the core fluorescence of the droplets. Once this residual polymer concentration in core was obtained, the fraction of polymer recruited in the interfacial layer was calculated as being equal to the loss of the soluble polymer amount. Eventually, this fraction of interfacial polymer was converted into surface excess by normalizing by the droplet interfacial area.

Raw values of fluorescence at the droplet surface could however be optically biased by curvature effects since the water/oil interface can deviate the light beam of the microscope. To account for this optical bias, the core fluorescence of dye-labeled PLL-g-PNIPAM was normalized following a suitable calibration process which is described below.

As Cy3-labeled PNIPAM (PNIPAM-Cy3) was homogeneously distributed within droplet cores (see Figure 3.9 in section 3.3.2), it was used as a reference for the calibration of the fluorescence in the droplet cores. Based on the fluorescence of PNIPAM-Cy3 in droplets, we determined a corrective factor accounting for droplet size effects on the fluorescence signal. First, the fluorescence intensity $I_{bulk}^{Cy3}$ of aqueous solutions of PNIPAM-Cy3 at various concentrations $C$ was measured by confocal imaging, yielding a Beer-Lambert linear relation (see Figure 3.14.A). In the same conditions of irradiation, PNIPAM-Cy3 was loaded – at the same concentrations than for the bulk experiments – in water-in-oil emulsion droplets (see Figure 3.14.B). The fluorescence intensity $I_{drop}^{Cy3}$ was measured in the core of the droplets for different droplet diameters $d$ (see Figure 3.14.C). The normalization of the droplet core intensity by the bulk intensity at a given PNIPAM-Cy3 concentration resulted in a corrective factor

$$I_{norm}^{Cy3}(d) = \frac{I_{drop}^{Cy3}(C,d)}{I_{bulk}^{Cy3}(C)} \text{ which only depends on the droplet diameter } d \text{ (see Figure 3.14.D).}$$
<figure><img src="image_51.png" /><figcaption>**Figure 3.14.** Calibration process accounting for optical bias on the measurement of fluorescence level in the droplet cores by confocal imaging. A) Beer-Lambert calibration curve of PNIPAM-Cy3 fluorescence measured in bulk solution by confocal fluorescence imaging. B) Confocal micrograph of an emulsion containing 2.5 wt% of PNIPAM-Cy3 in the aqueous phase and 0.05 wt% of Krytox in the oil phase at 18 °C (scale bar = 50 µm). C) Fluorescence level of PNIPAM-Cy3 measured in the core of water droplets as the function of the droplet diameter and for different concentrations of PNIPAM-Cy3. D) Normalized fluorescence level in droplets (corresponding to the ratio between core fluorescence and bulk fluorescence at the same concentration) as a function of the droplet diameter. To simplify the distribution, each droplet diameter has been rounded down to the nearest integer (error bars = standard deviation for each integer diameter).</figcaption></figure>

The same normalization approach of the droplet core intensity was followed for the fluorescence of dye-labeled PLL-g-PNIPAM. In the following, the method is detailed for rhodamine-labeled PLL-g-PNIPAM (PLLrho-g-PNIPAM), but the same approach was applied for coumarin-labeled PLL-g-PNIPAM.

First, the fluorescence intensity $I_{bulk}^{rho}$ of PLLrho-g-PNIPAM solutions at different concentrations $C$ was measured in bulk. The same polymer solutions were emulsified in oil containing Krytox. The fluorescence intensity $I_{drop}^{rho}$ of PLLrho-g-PNIPAM was then measured in the droplet cores and normalized by $I_{bulk}^{rho}$ corresponding to the same polymer concentration. The ratio between the actual PLLrho-g-PNIPAM concentration in droplet cores $C_{core}$ and the analytical concentration $C$ introduced in the aqueous phase before emulsification was eventually determined by the following formula:
$$ \frac{C_{\text{core}}}{C} = \frac{I_{\text{drop}}^{\text{rho}}(C, d)}{I_{\text{bulk}}^{\text{rho}}(C)} \times \frac{I_{\text{bulk}}^{\text{Cy3}}(C)}{I_{\text{drop}}^{\text{Cy3}}(C, d)} = \frac{I_{\text{norm}}^{\text{rho}}(d)}{I_{\text{norm}}^{\text{Cy3}}(d)} $$

The factor $I_{\text{norm}}^{\text{rho}}(d)$ evaluates the inhomogeneity of the PLLrho-g-PNIPAM distribution in water droplets due to the significant adsorption of the polymer at the water/oil interface and the factor $I_{\text{norm}}^{\text{Cy3}}(d)$ accounts for the correction of optical bias due to the droplet size. Eventually, the $\frac{C_{\text{core}}}{C}$ ratio obtained via this calibration method is no longer dependent on the droplet size, and physically corresponds to the residual fraction of polymer chains that have not been involved in the interfacial complexation with Krytox and have remained soluble in the droplet core. Accordingly, it will be referred to as “residual core concentration” in the following.

Eventually, the PLLrho-g-PNIPAM surface excess was defined as the excess number of polymer chains $N_{\text{excess}}$ adsorbed at the interface per surface unit $S$ in conditions of interfacial complexation (compared to a hypothetic uniform distribution of the chains in the droplet volume $V$). It is given by the following formula:

$$ \frac{N_{\text{excess}}}{S} = \frac{(C - C_{\text{core}})V}{S} = \left(1 - \frac{C_{\text{core}}}{C}\right) \frac{d}{6} C = \left(1 - \frac{I_{\text{norm}}^{\text{rho}}(d)}{I_{\text{norm}}^{\text{Cy3}}(d)}\right) \frac{d}{6} C $$

In the following, both the residual core concentration and the surface excess will be analyzed to conclude on the influence of different factors on the efficiency of the interfacial complexation process, such as the ionic strength and pH (see section 3.4.3), or the temperature (see section 3.5.3).

### 3.4.3 Influence of the ionic strength and the pH on the interfacial complexation

As interfacial complexation involves coulombic interactions between carboxylate-terminated Krytox and ammonium groups of PLL, the pH as well as the ionic strength of the aqueous phase should have an important impact on the interfacial complexation equilibrium, and may enable to optimize the properties of the interfacial assemblies. Beyond qualitative comparisons, the quantitative approach presented in the previous section was applied to rhodamine-labeled PLL-g-PNIPAM to conclude on the efficiency of its complexation for the different formulations of the aqueous phase.
To assess the effect of the ionic strength on the interfacial complexation of PLL-g-PNIPAM, we compared polymer-loaded droplets containing either a 7.7 mM phosphate buffer (ionic strength = 17 mM, pH = 7.3), or this buffer supplemented with 100 mM or 500 mM NaCl (see **Figure 3.15**). Irrespective of the ionic strength condition, a bright corona was visible at the edge of the droplets with a low residual fluorescence in the core (see Figure 3.15.A) and a non-zero surface excess was measured, suggesting that the interfacial complexation was effective even at high NaCl concentration.

<figure><img src="image_52.png" /><figcaption>**Figure 3.15.** Influence of the ionic strength on the interfacial complexation between 1 wt% of coumarin-labeled PLL-g-PNIPAM and 0.05 wt% Krytox at 18 °C. A) Confocal micrographs of emulsions prepared with different ionic strengths in the aqueous phase (scale bars = 20 µm). PB = phosphate buffer. B) Polymer surface excess as a function of the residual core concentration (normalized by the analytical concentration of 1 wt%). Dashed lines indicate the mean level of surface excess for each condition. Box plots give the min/Q1/median/Q3/max values of the distribution of residual concentration in core for each condition. C) Polymer surface excess as a function of the droplet diameter. The light orange dashed line reported on each plot corresponds to the trend for PB without NaCl (top layer) which was used as a reference.</figcaption></figure>

However, uneven distributions of fluorescence around the droplet edges were noticed in NaCl-containing emulsions, and particularly at 500 mM NaCl where “crescent moon-like” shapes of fluorescence became obvious specifically in small droplets (< 20 µm). This observation suggests that the presence of salts at high concentration in the aqueous phase led to a patchy segregation of the polymer chains that significantly
differs from the uniform coverage of the interface at low ionic strength. The corresponding polymer surface
excesses are shown as a function of the residual core concentration and droplet diameter in Figure 3.15.B
and C, respectively. The presence of NaCl slightly diminished the surface excess, and markedly broadened
the distribution of residual core concentrations. Some droplets containing 100 mM or 500 mM NaCl had
up to 80 % of residual polymer chains in their core, suggesting the weakening of interfacial binding with
increasing salt. Recent articles on polycation:polyanion (PC:PA) complexes consistently reported that ionic
strength play a similar role as temperature in the phase diagram of complex coacervation. With increasing
salt concentration solid-like PC:PA coacervates first turn to more fluid phases (gels or liquid-like) prior to
become soluble.²⁵⁵⁻²⁵⁷ As a tentative interpretation of "crescent-like" distribution of the polymer at
interfaces, we propose that a transition to fluid phase enabled fusion and gathering of the complexes at one
pole of the droplets. To prepare capsules, robust membranes are thus likely obtained at low ionic strength
as this offers more homogeneous interfaces and ideally a solid-like interfacial complex phase.

To assess the effect of the pH on the interfacial complexation of PLL-g-PNIPAM, we substituted the
phosphate buffer by 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 (see **Appendix A**). 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, and emulsified in the oil phase (see **Figure 3.16**).

Significant accumulation of polymer at the interface was observed in both acidic and alkaline
conditions as shown by the fluorescent corona formed at the edge of the droplets (see Figure 3.16.A). The
lower surface excess as well as higher residual fluorescence observed in droplet cores at pH = 9.4 as
compared to pH = 4.3 clearly indicate that the PLL/Krytox complexation is less efficient in alkaline than in
acidic conditions (see Figure 3.16.B). This contrast in surface excess and the poor interfacial capture of
PLL-g-PNIPAM at high pH is consistent with the decrease of the degree of ionization of PLL at higher pH:
in alkaline conditions, the pH was close to the pKa of the side amino groups of PLL (~ 10.5) and therefore
a significant fraction of amine groups was deprotonated, what is expected to weaken coulombic association
with Krytox. In contrast, all the side amino groups were protonated at pH = 4.3, and thus available to
associate with Krytox across the water/oil interface.
<figure><img src="image_53.png" /><figcaption>**Figure 3.16.** Influence of the pH on interfacial complexation between 1 wt% coumarin-labeled PLL-g-PNIPAM and 0.03 wt% Krytox at 18 °C in Britton-Robinson buffer. A) Confocal micrographs of emulsions at pH = 4.3 and 9.4 (scale bar = 30 µm). B) Correlation between polymer surface excess and residual core concentration (normalized by the analytical concentration of 1 wt%) at pH = 4.3 (black circles) and pH = 9.4 (red circles). Colored domains are guide for the eye to better visualize the scatter distributions. C) Surface excess of coumarin-labeled PLL-g-PNIPAM as a function of the droplet diameter at pH = 4.3 (black circles) and pH = 9.4 (red circles), calculated from fluorescence measurements. Dashed lines are guides for the eye. The blue dashed line is the surface excess measured in BR buffer prepared at pH = 7.3 (raw data are not shown for more readability).</figcaption></figure>
## 3.5 Evaluation of the thermoresponsive behavior of PLL-g-PNIPAM shells

### 3.5.1 Thermoresponsive behavior of PLL-g-PNIPAM in solution

First, the thermoresponsive behavior of PLL-g-PNIPAM was characterized in solution by turbidimetry and compared to free PNIPAM (see **Figure 3.17**). The transmittance profiles of either PLL-g-PNIPAM or free PNIPAM solutions decreased between 33 °C and 40 °C, indicating that PNIPAM strands – grafted or not – undergo a coil-to-globule transition (or collapse transition) in the same temperature window. A linear extrapolation of the curve around the inflexion point indicates the same cloud point of 34 °C for both PNIPAM forms (coinciding with the beginning of the decrease in transmittance).

<figure><img src="image_54.png" /><figcaption>**Figure 3.17.** Turbidity heating curves of free PNIPAM (dashed line) and PLL-g-PNIPAM (solid line) solutions at 1 wt% in phosphate buffer.</figcaption></figure>
### 3.5.2 Direct visualization of the thermal transition of PLL-g-PNIPAM shells

The thermal transition of PLL-g-PNIPAM in emulsion was probed *in operando* using 8-anilinonaphthalene-1-sulfonic acid (ANS) as an extrinsic fluorescent reporter. The fluorescence of ANS is strongly polarity-dependent and can be used as a marker of the hydrophilic-to-hydrophobic transition of PNIPAM. In polar environments, the excited state of ANS undergoes a twisted intramolecular charge transfer (TICT) to a second and more polar excited state that rapidly relaxes to the ground state via non-radiative decay.²⁵⁸ As a consequence, the quantum yield of ANS is very low in polar solvents such as water. Conversely, the fluorescence level of ANS is enhanced when the molecule is confined in more apolar or hydrophobic environments as the radiative emission from the first excited state is favored compared to the TICT transition. These polarity-dependent fluorescence properties of ANS have been used for many characterizations of macromolecular systems, including the staining of hydrophobic protein domains,²⁵⁸,²⁵⁹ the study of surfactant-polymer interactions,²⁶⁰ or the evidence of thermal transitions in thermoresponsive polymers such as PNIPAM.²⁶¹⁻²⁶³

In our system, the ANS fluorescence was used as a marker of (i) the PLL-g-PNIPAM localization in droplets since polymer-enriched regions are more hydrophobic (less polar) than the aqueous background, and (ii) the thermoresponsive properties of PNIPAM since its collapse transition to a more hydrophobic state would be evidenced by an increase in ANS fluorescence intensity.²⁶³

Before using ANS to characterize the thermal properties of PLL-g-PNIPAM at the water/oil interface, a first experiment was performed in emulsion with free PNIPAM chains to control the fluorescent behavior of ANS in presence of thermoresponsive chains. For this experiment, two emulsions were independently prepared and then mixed together: one containing only ANS in phosphate buffer (denoted E1), the other containing ANS and free PNIPAM chains in phosphate buffer (denoted E2). Both emulsions were prepared in an oil phase containing 0.05 wt% of Krytox. The mixed emulsion was observed in confocal microscopy at 20 °C (below the cloud point) and at 45 °C (above the cloud point) to monitor the evolution of ANS fluorescence in the two types of droplets (E1 and E2).

**Figure 3.18** shows confocal micrographs of the mixed emulsion at 20 °C and 45 °C, with the E1 droplets (absence of PNIPAM) encircled in green and the E2 droplets (presence of PNIPAM) encircled in red. At 20 °C, a low fluorescence signal due to ANS could be detected in the core of both droplet types, with almost no contrast between E1 and E2. At 45 °C, the ANS fluorescence remained low in E1 droplets but was drastically enhanced in E2 droplets, revealing the hydrophilic-to-hydrophobic transition of the
PNIPAM chains present in these droplets. The difference in ANS behavior between E1 and E2 environments is highlighted in the fluorescence radial profiles drawn for a representative droplet of each nature in Figure 3.18.B.

<figure><img src="image_55.png" /><figcaption>**Figure 3.18.** ANS fluorescence contrast upon hydrophilic-to-hydrophobic transition of PNIPAM in emulsion. A) Confocal micrographs at 20 °C and 45 °C of a mixture of two emulsions: E1 containing 500 µM ANS in phosphate buffer (droplets encircled in green) and E2 containing 100 µM ANS and 2 wt% PNIPAM in phosphate buffer (droplets encircled in red), both with 0.05 wt% Krytox in the oil phase (scale bars = 50 µm). B) Fluorescence radial profiles of a representative droplet of E1 and E2 (indicated with colored arrows in the top micrographs) at 18 °C and 45 °C. C) Evolution of the fluorescence ratio between E1 and E2 droplets of comparable size with temperature (error bars = standard error on 8 couples of droplets).</figcaption></figure>

To confirm the identification of the thermal transition of PNIPAM by the fluorescence enhancement of ANS, the potential effect of the aqueous background was suppressed by calculating the E2/E1 fluorescence ratio for droplets of similar size (same diameter ±5%). This ratio was plotted as a function of the temperature upon heating in Figure 3.18.C. The ratio increases with the temperature until reaching a maximum at 35 °C, corresponding to the cloud point of PNIPAM. Beyond this temperature, the ratio decreases but remains significantly higher than at 20 °C. This fall seems contradictory with the fact that the
hydrophobicity of PNIPAM increases with temperature so that the ANS fluorescence signal should be even higher at 45 °C compared to 35 °C. This discrepancy is probably neither due to the intrinsic photophysical properties of ANS nor to an anomaly in the PNIPAM thermal behavior, but is rather due to a combination of three other factors. The first one is the photobleaching underwent by ANS molecules throughout the experiments that may explain a progressive decrease of the emission intensity in highly fluorescent droplets upon heating. The second factor is the destabilization of the mixed emulsion which was evidenced by multiple coalescence events (droplets encircled in both green and red at 45 °C in Figure 3.18.A), and which may have led to the dilution of ANS molecules. This loss of stability can be explained by the absence of interfacial complexation in this sample (no cationic species were loaded in the aqueous phase to interact with Krytox). The third factor, which is probably the predominant one, was the progressive sedimentation of PNIPAM aggregates formed upon collapse transition at high temperature (evidenced by the granular aspect of the E2 droplets at 45 °C). Due to the accumulation of polymer at the bottom of the droplets, the apparent fluorescence intensity in the equatorial plane was lower than expected in the case of a uniform distribution of the polymer in the droplets.

Despite these experimental limitations, this experiment confirmed that the hydrophilic-to-hydrophobic transition of PNIPAM chains can be evidenced in aqueous droplets by an increase of the fluorescence intensity of ANS added in the aqueous phase.

To probe the thermal transition of PNIPAM strands grafted on PLL and in condition of interfacial complexation with Krytox, ANS was added in the aqueous solution of PLL-g-PNIPAM and its fluorescence was monitored upon heating. **Figure 3.19**.A shows confocal micrographs of an emulsion containing 2 wt% of PLL-g-PNIPAM and 500 µM of ANS in the aqueous phase at 20 °C and 45 °C, respectively below and above the cloud point.
<figure><img src="image_56.png" /><figcaption>**Figure 3.19.** Evidence for the thermal collapse transition of PLL-g-PNIPAM in conditions of interfacial complexation. A) Top: Confocal micrographs at 20 °C and 45 °C of an emulsion containing 2 wt% of PLL-g-PNIPAM and 500 µM of ANS in the aqueous phase and 0.05 wt% of Krytox in the oil phase (scale bars = 50 µm). Bottom: Zoom on the droplet framed in white in the top micrographs, at intermediary temperatures (scale bars = 10 µm). B) Fluorescence radial profiles of the droplet shown in B at increasing temperature. C) Plot of the droplet edge/core intensity ratio – extracted from the fluorescence radial profiles – as a function of the temperature (error bars = standard error on 8 droplets).</figcaption></figure>

At 20 °C the fluorescence signal of ANS in polymer-loaded droplets was non uniform: the intensity was higher at the droplet edge, as highlighted by the sharp peak of fluorescence (~ 3-fold higher than the core level) observed in the radial profile (see Figure 3.19.B). This observation, suggesting that the droplet edges are more hydrophobic than the droplet cores, is consistent with an initial deposition of PLL-g-PNIPAM at the water/oil interface as shown in Figure 3.9. As temperature was increased from 20 °C to 45 °C, we observed an increase in fluorescence contrast between the edge and the core of the droplets (see Figure 3.19.A), due to a concomitant decrease of intensity in the core and increase of intensity of the peak at the edge (see Figure 3.19.B). The ratio between edge and core fluorescence intensities plotted in Figure 3.19.C shows a continuous increase with temperature, with a significant change of slope at 34 °C corresponding to the cloud point. The significant values of the edge/core ratio at high temperature (T > 34 °C) confirm the occurrence of the thermal collapse transition of PNIPAM at the water/oil interface at 34 °C. Moreover, the fluorescence level of ANS fell down to almost zero in droplet cores at T > 34 °C, suggesting that PLL-g-PNIPAM chains that were initially soluble in the droplet cores have enriched the interface upon heating.
The reversibility of this process was assessed by submitting an emulsion to temperature cycles while recording the edge/core ratio of ANS fluorescence (see **Figure 3.20**).

<figure><img src="image_57.png" /><figcaption>**Figure 3.20.** Assessment of reversibility of ANS fluorescence in presence of PLL-g-PNIPAM. A) Confocal micrographs of an emulsion with 2 wt% of PLL-g-PNIPAM and 500 µM of ANS in the aqueous phase and 0.05 wt% of Krytox in the oil phase (scale bars = 50 µm). The emulsion was alternatively thermalized at 18 °C (steps 1 and 3, blue frames) and 45 °C (steps 2 and 4, red frames). Inserts: Zoom on the droplet indicated with a white arrow in the micrograph (scale bars = 10 µm). B) Fluorescence radial profiles of the droplet shown in inserts in A (same color code). C) Edge/core fluorescence ratio calculated from the curves in B (same color code, error bars = standard error on 8 droplets).</figcaption></figure>

Initially set at 18 °C, the emulsion was heated up to 45 °C, then cooled back to 18 °C and heated again to 45 °C. In samples brought back to 18 °C, the fluorescence level in the droplet cores slightly increased compared to 45 °C but did not come back to the initial value measured before heating. After the second heating step the fluorescence profile was identical to the one after the first heating step. These observations point out that after a first collapse transition in the interfacial layer, PLL-g-PNIPAM that had accumulated at the droplet edge remained stuck in the interfacial layer and did not significantly redissolve when the sample was brought back below the transition temperature (at least on experimental time scale of ~ 1 hour).

As a control experiment, the same imaging process was performed with a non-thermoresponsive poly(L-lysine)-g-poly(ethylene glycol) (PLL-g-PEG) copolymer with the same 0.22 grafting ratio than PLL-g-
PNIPAM. A solution of 2 wt% of PLL-g-PEG and 500 µM of ANS was emulsified in oil phase, and the emulsion was observed in confocal microscopy at 18 °C and 45 °C (see **Figure 3.21**).

<figure><img src="image_58.png" /><figcaption>**Figure 3.21.** Comparison of PLL-g-PNIPAM and PLL-g-PEG regarding ANS fluorescence. A) Confocal micrographs of an emulsion containing 2 wt% of either PLL-g-PNIPAM or PLL-g-PEG and 500 µM of ANS in the aqueous phase, and 0.05 wt% of Krytox in the oil phase, at 20 °C and 45 °C (scale bars = 50 µm for PLL-g-PNIPAM and 30 µm for PLL-g-PEG). B) Edge/core intensity ratio calculated from the radial fluorescence profile of 8 droplets (error bars = standard error) plotted as a function of the temperature for PLL-g-PNIPAM (black) and PLL-g-PEG (red).</figcaption></figure>

After formulation at 18 °C, the droplet edge/core ratio of ANS fluorescence was greater than 1, indicating that PLL-g-PEG has been adsorbed at the water/oil interface by complexation with Krytox, but significantly lower than for PLL-g-PNIPAM, probably because both PLL and PEG moieties are highly hydrophilic. When observed with SEM, the capsules made of PLL-g-PEG at 18 °C looked very similar both in size and effective membrane thickness (see Figure 3.12 in section 3.4.1 above). Moreover, no significant change of the edge/core ratio in droplets containing PLL-g-PEG was observed between 18 °C and 45 °C, indicating that temperature had no effect on the hydrophobicity of the interfacial polymer layer. These results confirm that the thermoresponsiveness of PLL-g-PNIPAM highlighted by ANS fluorescence at the droplet edges was intrinsically related to the collapse transition of PNIPAM strands.

The thermoresponsiveness of the PNIPAM strands being preserved once adsorbed at the water/oil interface, we presume that these results illustrate a generic approach for imparting various chemical properties to interfacial polymer layers via the use of the PLL-based comb-like architecture. Other functional polymer strands (such as light- or redox-responsive ones) could thus be grafted on PLL to design new functional polymer shells by interfacial complexation.
### 3.5.3 Effect of droplet radius and temperature on characteristic features of the PLL-g-PNIPAM shells

To characterize the effect of the temperature on the interfacial polymer layer, we implemented the quantitative fluorescence analysis presented in section 3.4.2 to measure the polymer surface excess in the droplets at low and high temperature. As the fluorescence of carboxy-X-rhodamine is neither subject to bleaching,²⁶⁴ nor sensitive to temperature (see **Figure 3.22**), it was used to evaluate the concentration of rhodamine-labeled PLL-g-PNIPAM (PLLrho-g-PNIPAM) in emulsion over temperature changes.

<figure><img src="image_59.png" /><figcaption>**Figure 3.22.** Fluorescence intensity of carboxy-X-rhodamine solutions measured with a confocal microscope in the same irradiation conditions at 18 °C and 45 °C.</figcaption></figure>

**Figure 3.23.**A shows confocal micrographs of a water-in-oil emulsion with water droplets containing 3 wt% of PLLrho-g-PNIPAM at 18 °C and 45 °C. As previously evidenced, the high rhodamine fluorescence signal at the droplet edge indicates that PLLrho-g-PNIPAM had accumulated at the water/oil interface by conventional interfacial complexation upon the emulsification process. At 45 °C (above the cloud point), the interface was still highly fluorescent, but the fluorescence signal had almost disappeared in the droplet core. This sharp decrease in core intensity upon heating is highlighted in the radial profiles drawn in Figure 3.23.A. This observation supports the idea of a thermo-induced transfer of core-soluble PLLrho-g-PNIPAM chains towards the interface, probably associated to the progressive immobilization of diffusive chains onto the excess interfacial PLLrho-g-PNIPAM layer via hydrophobic interactions. This hypothesized process is depicted in Figure 3.23.B.
<figure><img src="image_60.png" /></figure>

<figure><img src="image_61.png" /></figure>

<figure><img src="image_62.png" /></figure>

<figure><img src="image_63.png" /><figcaption>**Figure 3.23.** Effect of temperature on the amount of interfacial PLLrho-g-PNIPAM. A) Top: Confocal micrographs of a representative water droplet containing 3 wt% of PLLrho-g-PNIPAM dispersed in oil containing 0.05 wt% of Krytox at 18 °C and 45 °C (scale bar = 10 µm). Bottom: Fluorescence radial profiles of the top droplets. B) Schematic illustration of the accumulation of core-soluble PLLrho-g-PNIPAM chains at the droplet surface upon collapse transition in emulsion (occurring at 34 °C). C) Evolution of the surface excess of PLLrho-g-PNIPAM as a function of the droplet diameter at 18 °C and 45 °C, calculated from fluorescence measurements. The maximum surface excess (black dashed line) corresponds to the ideal case of interfacial adsorption of all the PLL-g-PNIPAM chains initially present in the droplet.</figcaption></figure>

From the fluorescence radial profiles of the droplets (as shown in Figure 3.23.C), we could define an “apparent” thickness of the interfacial polymer layer as the width at half height of the fluorescence peak at the droplet edge. A mean value of ~ 1.5 µm was measured independently of the polymer concentrations (in the 2-5 wt% range) and at both low and high temperatures (see **Figure 3.24**). Taking into consideration the lack of resolution of fluorescence microscopy, the width of fluorescence peak measured on the confocal micrographs is rather consistent with the thickness observed by SEM (see Figure 3.12 in section 3.4.1).
<figure><img src="image_64.png" /><figcaption>**Figure 3.24.** Apparent shell thickness measured from fluorescence imaging. A) Fluorescent shell thickness measured on an emulsion containing 2 wt% of PLLrho-g-PNIPAM in the aqueous phase and 0.05 wt% of Krytox in the oil phase at 18 °C and 45 °C. B) Mean shell thickness measured on emulsions with PLLrho-PNIPAM concentration in the range 2-5 wt% (error bars = standard deviation).</figcaption></figure>

The evolution of the polymer surface excess with the droplet diameter at 18 °C and 45 °C is shown in Figure 3.23.D. Experimental data were compared to the maximum reachable surface excess, corresponding to the adsorption at the interface of all the polymer chains contained in the droplets. This maximum excess increases linearly with the droplet diameter as it is proportional to the droplet volume/surface ratio. At 18 °C, the experimental surface excess is significantly below the maximum curve and increases with droplet diameter until reaching a plateau for big droplets (> 30 µm). These observations suggest that a significant fraction of the polymer was not involved in interfacial complexation and that a saturation equilibrium between core-soluble and edge-adsorbed chains was presumably achieved at high volume/surface ratios.

Interestingly, when switching temperature from 18 °C to 45 °C the surface excess almost doubled at a given droplet diameter. For small droplets (< 30 µm), the surface excess coincides with the maximum excess curve, indicating that all the polymer chains contained in the droplet have been segregated at the water/oil interface. This observation is in agreement with an enrichment of the polymer interfacial layer above the cloud point, confirming the hypothesis of a core draining and an accumulation of polymer chains at the interface via hydrophobic interactions. For big droplets (> 30 µm), the experimental surface excess deviates from the maximum excess curve, suggesting that a saturation level has been reached in the interfacial complexation process, and that a fraction of soluble polymer chains could not be recruited in the interfacial complex layer.

The evolution of surface excess as a function of the residual concentration in droplet cores is shown in **Figure 3.25**. From 18 °C to 45 °C, the fraction of core-soluble polymer chains decreased from 0.4-0.6 to less than 0.1 concomitantly to an increase in the mean surface excess, confirming that (i) a high
proportion of polymer chains were not involved in interfacial complexation in conditions of conventional interfacial complexation (below the cloud point) and (ii) a massive accumulation of polymer chains at the interface occurred beyond the collapse transition (above the cloud point).

<figure><img src="image_65.png" /><figcaption>**Figure 3.25.** Evolution of PLLrho-g-PNIPAM surface excess with droplet diameter (A) and residual core concentration (B) at 18 °C and 45 °C. The emulsion was made of 3 wt% of PLLrho-g-PNIPAM in the water phase and 0.05 wt% of Krytox in the oil phase.</figcaption></figure>

The microscopy experiments shown above did not give access to the molecular organization of the polymer shell. The interfacial complexation between PLL-g-PNIPAM and Krytox probably led to the formation of a complex coacervate phase growing across the liquid/liquid interface upon both interactions and diffusion of the molecular species. The formation of a hierarchically ordered membrane²⁶⁵ with a thickness of several hundreds of nm is much more likely than the formation of molecular monolayers on both side of the interface (monolayer of Krytox on the external side and monolayer of polymer on the internal side of the droplet). To reject this latter hypothesis (monolayer distribution), the mass of polymer located in the shell – experimentally deduced from the previous fluorescence measurements – was compared to the mass of polymer required to exactly compensate the charge of a monolayer of Krytox decorating the droplet surface. This mass can be derived from the following equation:

$$m_{\text{polymer}} = \frac{\# \text{ Krytox per droplet (monolayer)}}{\# \text{ charged lysine per copolymer chain}} \times m_{\text{polymer chain}} = \frac{1}{(1 - \tau)\text{DP}_{n,\text{PLL}}} \times \frac{\pi d^2}{a_{\text{Kr}}} \times \frac{M_{\text{PLL-g-PNIPAM}}}{N_A}$$

$(1 - \tau) \text{DP}_{n,\text{PLL}} \sim 82$ is the average number of positive charges (i.e. of free lysine units) per copolymer chain (see Table 3.1 in section 3.2.2), $M_{\text{PLL-g-PNIPAM}} \sim 68$ kg/mol is the average molar mass of a copolymer chain, $N_A = 6.02 \times 10^{23} \text{ mol}^{-1}$ is the Avogadro number, and $a_{\text{Kr}}$ is the surface area of the Krytox polar head, which has not been experimentally measured but was reasonably fixed to $4 \AA^2$. Using the appropriate numerical values, the equation above becomes: $m_{\text{polymer}} [\text{ng}] = 1.09 \times 10^{-4} \times d^2 [\mu\text{m}^2]$.
**Figure 3.26** shows the polymer mass accumulated in the droplet shells for the dataset presented in Figure 3.23 and Figure 3.25, simply obtained by multiplying the polymer surface excess by the droplet surface area. These experimental data are compared to the model of charge compensation of a Krytox monolayer as derived in the previous paragraph. The experimental values of polymer mass in the shells are way higher than in the model of charge compensation, both at 18 °C and at 45 °C, confirming that the droplet shells are not made of well-defined molecular monolayers but are rather made of a more complex coacervate phase.

<figure><img src="image_66.png" /><figcaption>**Figure 3.26.** Evolution of the polymer mass in the shell with the droplet diameter at 18 °C and 45 °C. The black dashed line corresponds to the case of a charge compensation of a monolayer of Krytox decorating the droplet surface</figcaption></figure>

Figure 3.26 also gives a revisited illustration of the process of thermo-induced enrichment of the polymer membrane since the polymer mass in the shells doubles as the temperature increases from 18 °C to 45 °C. As the apparent shell thickness was more or less the same at 45 °C than at 18 °C (see Figure 3.24), the enrichment of the polymer membrane at high temperature was probably concomitant with a compaction of the complex coacervate phase (which has become hydrophobic above the cloud point).

In addition, the process of thermo-induced accumulation of polymer at the interface seems to be mainly irreversible since the surface excess remained at the same high level when a sample was cooled back at 18 °C after a first heating step at 45 °C (see **Figure 3.27**). This suggests that the enrichment of the polymer shell is not subject to significant redispersion as PNIPAM chains are brought back below the cloud point.
<figure><img src="image_67.png" /><figcaption>**Figure 3.27.** Assessment of the reversibility of PLL-g-PNIPAM adsorption at the water/oil interface upon collapse transition. A) Confocal micrographs of an emulsion containing 3 wt% of PLLrho-g-PNIPAM in the aqueous phase and 0.05 wt% of Krytox in the oil phase, heated up at 45 °C and cooled back to 18 °C (scale bars = 20 µm). Thermalization was ensured for 5 min before measurement. B) PLLrho-g-PNIPAM surface excess calculated from the measurement of rhodamine fluorescence in the droplet cores at 45 °C and 18 °C.</figcaption></figure>

As a complement, **Figure 3.28** shows the effect of heating on polymer surface excess for emulsions containing 2 wt% or 5 wt% of PLLrho-g-PNIPAM in the aqueous phase.

The results of the emulsion with 2 wt% of polymer are similar to the one with 3 wt% presented above: a massive thermo-induced clustering of soluble polymer chains was observed at the droplet periphery upon heating, resulting in an increase of the polymer surface excess at higher temperature (see Figure 3.28.A).

For the emulsion with 5 wt% of polymer, the results are slightly different. First, the fluorescence contrast between the core and the edge of the droplets was hardly detectible at 18 °C, suggesting that the droplets were overloaded with PLLrho-g-PNIPAM to the point where the complexation-induced deposition at the interface was concealed (the interface was oversaturated). As a consequence, the polymer surface excess deduced from the core fluorescence was well below the maximum reachable surface excess (see Figure 3.28.B). An increase of the surface excess was observed at 45 °C but was likely due to a polymer sedimentation at the bottom of the droplets rather than to a progressive and uniform accumulation at the surface of the droplets. Indeed, the confocal micrographs at 45 °C revealed the formation of a large amount of polymer aggregates that probably sedimented before reaching the interface by diffusion, causing a decrease of the fluorescence in the equatorial plane of the droplets. This effect was misinterpreted as an increase of the surface excess in the fluorescence quantification process.
<figure><img src="image_68.png" /><figcaption>B) 5 wt% PLLrho-g-PNIPAM</figcaption></figure>

<figure><img src="image_69.png" /></figure>

**Figure 3.28.** Influence of the polymer concentration on the surface excess. Confocal micrographs and evolution of the polymer surface excess as a function of the droplet diameter at 18 °C and 45 °C for emulsions containing either 2 wt% (A) or 5 wt% (B) of PLLrho-g-PNIPAM in the aqueous phase (scale bars = 20 µm).

Considering only the surface excess below the LCST, the quantitative results are consistent with the qualitative observations made in section 3.4.1 on the influence on the polymer concentration on the morphology of the polymer capsules. At polymer concentrations < 3 wt%, high surface excesses (compared to the reachable maximum) are consistent with the formation of hollow capsules. At polymer concentration > 3 wt%, poor surface excesses (due to saturation in polymer) is consistent with the formation of filled microparticles (see SEM images in Figure 3.13).

As a control of the PNIPAM behavior, the same quantification of the polymer surface excess was performed on a non-thermoresponsive PLL-g-PEG labeled with carboxy-X-rhodamine (PLLrho-g-PEG) and the results are shown in **Figure 3.29**.
<figure><img src="image_70.png" /><figcaption>**Figure 3.29.** Effect of temperature on a non-thermoresponsive PLLrho-g-PEG layer. A) Confocal micrographs of an emulsion containing 2 wt% of PLLrho-g-PEG in the aqueous phase and 0.05 wt% of Krytox in the oil phase, at 18 °C and 45 °C (scale bars = 20 µm). B) PLLrho-g-PEG surface excess calculated from the measurement of rho fluorescence in the droplet cores as a function of the droplet diameter at 18 °C and 45 °C (error bars = propagation of uncertainty). C) PLLrho-g-PEG surface excess as a function of the concentration fraction in the droplet cores at 18 °C and 45 °C.</figcaption></figure>

The curves of surface excess superimpose at 18 °C and 45 °C and plateau at a saturation level which is twice lower than for PLL-g-PNIPAM at 18 °C (and 4 times lower at 45 °C) indicating that (i) PLL-g-PNIPAM has a higher affinity with the interface than PLL-g-PEG presumably because PNIPAM strands are shorter than PEG ones (18 vs 44 repeat units/chain respectively) and more hydrophilic, and (ii) there is no effect of the temperature on the distribution of PLL-g-PEG in droplets, confirming that the process observed for PLL-g-PNIPAM is intrinsic to the thermoresponsive nature of PNIPAM strands.
### 3.5.4 Evidence for thermo-induced enrichment of the polymer shell with neutral PNIPAM chains

The results presented in the two last sections suggested that PLL-g-PNIPAM chains can be accumulated at the interface beyond the initial deposition by interfacial complexation. This phenomenon, that we called “thermo-enhanced interfacial complexation” was likely due to the progressive accretion of soluble PLL-g-PNIPAM chains after having switched to a hydrophobic state upon heating. To evaluate this hypothesis of thermo-induced accumulation of matter at the interface, we studied the behavior of neutral PNIPAM chains (non-grafted on PLL) in droplets which also contain PLL-g-PNIPAM. Below the LCST, PLL-g-PNIPAM was expected to form a polymer shell by interfacial complexation, but free PNIPAM chains had no reason to be involved in this interfacial phenomenon since they were uncharged.

An emulsion was prepared with 1 wt% of coumarin-labeled PLL-g-PNIPAM and 0.5 wt% of Cy3-labeled PNIPAM in the aqueous phase (and 0.03 wt% of Krytox in the oil phase). Figure 3.30.A shows the distribution of these two polymer species in a representative droplet at 18 °C and at 45 °C.

At 18 °C, PLL-g-PNIPAM (in red) was located at the droplet periphery and free PNIPAM (in blue) was uniformly distributed in the droplet core, what is consistent with the fact that interfacial complexation occurs for cationic PLL-g-PNIPAM and not for neutral PNIPAM. At 45 °C, the fluorescence of free PNIPAM has almost disappeared from the droplet core and has accumulated at the droplet periphery as indicated by the formation of a bright corona. This result is highlighted in the fluorescence profiles shown in Figure 3.30.B and suggests that the thermo-induced accumulation of hydrophobic chains at the water/oil interface also occurred for free PNIPAM chains. Upon the collapse transition of the PNIPAM strands, the interfacial layer of PLL-g-PNIPAM became hydrophobic and “sticky”. Free PNIPAM chains also switched to their hydrophobic and aggregative state. Upon diffusion from the droplet core to the droplet edge, free PNIPAM chains remained trapped in the interfacial layer by hydrophobic interactions, leading to a progressive accumulation of polymer at the interface.
<figure><img src="image_71.png" /><figcaption>**Figure 3.30.** Thermo-induced accumulation of free PNIPAM chains into the PLL-g-PNIPAM shell. A) Confocal micrographs of a representative droplet containing 1 wt% coumarin-labeled PLL-g-PNIPAM (red) and 0.5 wt% Cy3-labeled PLL-g-PNIPAM (blue) at 18 °C, 45 °C and 18 °C successively (scale bar = 10 µm). B) Fluorescence radial profiles indicating the distribution of free PNIPAM chains (Cy3 channel) at the different temperatures.</figcaption></figure>

As a control, an emulsion containing only Cy3-labeled neutral PNIPAM chains (an no PLL-g-PNIPAM) in the aqueous phase was observed in confocal microscopy at 18 °C and 45 °C (see **Figure 3.31**). At 18 °C the uniform distribution of polymer chains in the droplet cores was observed as expected. At 45 °C, no evidence of polymer accumulation at the edge of the droplets was noticed, but the formation of hydrophobic polymer aggregates was clearly visible in the core of the droplets. The progressive sedimentation of these micrometric aggregates at the bottom of the droplets could explain the decrease of the core fluorescence detected upon heating, as highlighted in the fluorescence radial profiles in Figure 3.31.B.

<figure><img src="image_72.png" /><figcaption>**Figure 3.31.** Thermal behavior of free PNIPAM chains in droplets. A) Confocal micrographs of an emulsion containing 2 wt% of Cy3-labeled PNIPAM in the aqueous phase at 18 °C and 45 °C (scale bars = 30 µm). B) Fluorescence radial profiles of the droplet indicated with a white arrow in the micrographs.</figcaption></figure>
These results confirm that the hydrophilic-to-hydrophobic transition of the PLL-g-PNIPAM shell – initially formed by interfacial complexation – enables to drive the accumulation of polymer chains towards the interface upon gentle heating. This phenomenon can be viewed as a thermo-induced enrichment of the shell as it can involve polymer chains that were initially dispersed in the droplet cores. The example of thermo-accumulation of uncharged PNIPAM chains (non-grafted on PLL) suggests that the massive polymer clustering at the interface is likely due to hydrophobic interactions and not to electrostatic complexation with Krytox. This phenomenon turned out to be reversible as the free PNIPAM chains were redispersed in the core of the droplets after cooling back the emulsion below the cloud point (see Figure 3.30).

## 3.6 Encapsulation of a model protein in the aqueous core

The possibility to encapsulate proteins in PLL-g-PNIPAM core-shell droplets was tested using eGFP (enhanced Green Fluorescent Protein) as a fluorescent model protein (27 kDa, pI = 5.6).²⁶⁶ The eGFP was kindly provided by a colleague (R. Chouket), who produced and purified it according to a protocol described in the literature.²⁶⁷

An aqueous solution containing 2 wt% of coumarin-labeled PLL-g-PNIPAM and 10 µM of eGFP was prepared in phosphate buffer (pH = 7.3) and emulsified in oil containing 0.03 wt% of Krytox. The polymer and protein distributions within droplets were determined by confocal imaging at 18 °C and 45 °C (see **Figure 3.32**).

<figure><img src="image_73.png" /><figcaption>**Figure 3.32.** Encapsulation of eGFP in PLL-g-PNIPAM capsule precursors. A) Confocal micrographs of an emulsion containing 1 wt% of PLLcoum-g-PNIPAM and 10 µM of eGFP in the aqueous phase and 0.03 wt% of Krytox in the oil phase, at 18 °C and 45 °C (scale bars = 50 µm). B) Fluorescence profiles of the representative droplet indicated by a white arrow in the micrographs.</figcaption></figure>
The bright coumarin fluorescence signal at the droplet edge (in magenta) shows that PLL-g-PNIPAM was concentrated at the water/oil interface as expected, demonstrating that the presence of the protein in the aqueous phase did not affect the formation of the polymer shell by complexation with Krytox. The uniform distribution of eGFP within droplets (in green) showed that the protein had been efficiently encapsulated in the aqueous droplet cores. The overlay of confocal images clearly shows the “core-shell” distribution of the species, with the polymer at the droplet edge and the protein entrapped in the droplet core. The high fluorescence signal of the protein as well as the absence of visible protein aggregates suggest that the integrity of the protein has been preserved upon encapsulated and even after heating at 45 °C.

As a first approximation, the preservation of the protein integrity may be quantified by comparing the fluorescence level in the droplets with the fluorescence level of the initial protein solution. For that, the normalized core concentration was calculated with the fluorescence calibration method described in section 3.4.2. Briefly, the fluorescence level of eGFP in the droplets was normalized by the fluorescence level of the bulk aqueous solution and corrected by the geometrical factor measured on Cy-3 labeled PNIPAM-Cy3 (accounting for size and curvature biases). Around 70-75 % of the eGFP initial fluorescence was detected in droplets bigger than 25 µm (see **Figure 3.33**), indicating that the protein had not been dramatically denatured during the encapsulation in water droplets. In droplets smaller than 25 µm, the higher loss of fluorescence intensity in water droplets may be reasonably attributed to optical distortion of the fluorescence signal (not compensated by the correction process) and to bleaching of eGFP, rather than to a significant alteration of eGFP.

<figure><img src="image_74.png" /><figcaption>**Figure 3.33.** Normalized fluorescence level of eGFP in an emulsion containing 1 wt% of coumarin-labeled PLL-g-PNIPAM and 10 µM of eGFP in the aqueous phase and 0.03 wt% of Krytox in the oil phase.</figcaption></figure>
## 3.7 Conclusion

In this chapter, we evidenced the ability of PLL derivatives to form concentrated polymer layers at the surface of water droplets by interfacial complexation with a surfactant (Krytox) across the water/oil interface of inverse emulsion in fluorinated oils. We showed that tight PLL/Krytox coulombic interactions led to a strong decrease of the water/oil interfacial tension, enabling the formation of stable inverse emulsions.

We synthesized PLL-g-PNIPAM and PLL-g-PEG comb-like copolymers via the grafting of PNIPAM and PEG strands onto PLL. We showed that the interfacial complexation of these polymers with Krytox led to the formation of hollow polymer microcapsules – provided the optimization of the formulation – with a polymer shell thickness of ~1 µm and an aqueous core of controlled pH and ionic strength. In mild conditions of formulation, a model protein (the fluorescent eGFP) could be readily encapsulated in the core of the droplets without interfering with the formation of the polymer shell, demonstrating that our approach is promising for the storage of potentially fragile biomolecules. Other proteins should be tested to prove the robustness and versatility of the formulation approach developed here.

In the case of PLL-g-PNIPAM copolymer, we showed that the LCST thermoresponsive behavior of PNIPAM strands – switching from hydrophilic coils to hydrophobic globules at a cloud point of 34 °C – was preserved in the interfacial complexation state, suggesting that this thermoresponsiveness was successfully imparted to the polymer shells. In addition, the LCST transition of PNIPAM strands was used to thermo-induce an additional accumulation of polymer chains on the pre-formed interfacial layer of PLL-g-PNIPAM. This phenomenon has been termed “thermo-enhanced interfacial complexation” (see **Figure 3.34**). Above the temperature of collapse transition, we observed a massive clustering of PLL-g-PNIPAM at the edge of the droplets, likely due to hydrophobic interactions between PNIPAM strands. This process contributed to enrich the interfacial polymer layer as demonstrated by the increase in polymer surface excess measured from fluorescence images. Interestingly, the persistence of high surface excesses when droplets were cooled back to room temperature suggests that the thermo-induced accumulation of polymer chains in the shell wall was irreversible.

This thermo-induced polymer accumulation was also successfully implemented on neutral PNIPAM chains that could not be involved in primary interfacial complexation (as non-grafted onto PLL cationic chains). Upon heating, a massive accretion of free PNIPAM chains was observed onto the germinal interfacial PLL-g-PNIPAM layer. This process turned out to be reversible as the free PNIPAM chains turned out to redisperse in the droplet core after cooling the sample back to ambient temperature. These
results suggest that materials can be easily captured and released from interfacial layers of PLL derivatives,
thus opening new perspectives in the design of more complex and functional polymer shells. These
perspectives will be explored in Chapter 2 with the design of polymer shells that can incorporate
nanoparticles.

<figure><img src="image_75.png" /><figcaption>**Figure 3.34.** Description and applications of the phenomenon of thermo-enhanced interfacial complexation described in this chapter.</figcaption></figure>
Chapter 4

Formation of polymer-
nanoparticles mixed shells by
interfacial complexation
## 4.1 Introduction

In Chapter 3, we showed that thermoresponsive polymer shells, made of PLL-g-PNIPAM, can be readily formed by interfacial complexation in water-in-oil emulsions and in mild conditions. We showed that the collapse transition of PNIPAM chains localized in the polymer shell can induce the interfacial accumulation of PNIPAM chains that are soluble in the core of the emulsion droplets, and named this phenomenon “thermo-enhanced interfacial complexation”. In this Chapter, we will extend this phenomenon to the interfacial segregation of nanoparticles in order to design “composite” or “hybrid” polymer-particles shells.

The incorporation of nanoparticles (NPs) in the shell of polymer capsules or vesicles has been extensively explored in the literature. A first advantage provided by the presence of NPs is the improvement of the mechanical stability of the shells, due to the intrinsic mechanical properties of the NPs and/or their role of physical cross-links between polymer chains.²⁶⁸ For delivery applications, the introduction of NPs can be advantageous to modulate the permeability of the carrier,²⁶⁹ and to limit the undesired “burst release effect” in biological conditions.²⁷⁰ The introduction of NPs also enables to impart new functionalities to the polymer compartments, especially additional stimuli-responsiveness. For instance, gold nanoparticles can be used as light-to-heat converters at the nanoscale, enabling to trigger thermal transitions upon light irradiation (see section 2.3.4 in Chapter 2). Magnetic nanoparticles (such as superparamagnetic iron oxide nanoparticles, SPIONs) have also been introduced in polymer capsules to trigger on-demand release upon application of an alternating magnetic field.²⁷¹

The incorporation of NPs in polymer shells can be achieved via different strategies:

* Metal NPs can be grown within the polymer shell by reduction of soluble precursors that have impregnated the polymer scaffold. This strategy has been used to load polymer shells with silver or gold NPs for instance.²⁷²,²⁷³ The reduction process can also be specifically directed at the outer surface of the capsules, which can be interesting for catalysis applications.²⁷⁴,²⁷⁵ This strategy of *in situ* reduction has also been implemented in the membrane of polymersomes,²⁶⁹ or in the core of microgels.²⁷⁶

* Pre-formed NPs can be adsorbed at the surface of the compartments, as it has been demonstrated on polymer capsules,²⁷⁷ and more importantly on microgels.²⁷⁸,²⁷⁹

* NPs can be embedded in the polymer shell during its formation. An important illustration of this approach is the formation of layer-by-layer capsules by alternating deposition of polymer chains and NPs.²⁸⁰