Polymer Mechanochemistry Using Ultrasound: From Fundamental Reactivity to Controlled Drug Delivery

Citation

Luo, Meng (Stella) (2026) Polymer Mechanochemistry Using Ultrasound: From Fundamental Reactivity to Controlled Drug Delivery. Dissertation (Ph.D.), California Institute of Technology. doi:10.7907/mnbj-xq97. https://resolver.caltech.edu/CaltechTHESIS:01212026-163903186

Abstract

Polymer mechanochemistry harnesses mechanical force to drive specific chemical transformations in stress-sensitive molecules known as mechanophores. Through judicious chemical design, these force-driven reactions have enabled functional polymeric materials capable of sensing, self-healing, and catalysis. Research in this field encompasses expanding the mechanophore repertoire, elucidating the fundamental principles governing mechanically induced reactivity, and translating force-responsive systems into practical applications.

This thesis advances the field by contributing both fundamental insight and applied functionality in mechanophore activation under ultrasonication, with a particular focus on controlled drug release in biological environments. First, we develop an improved methodology for characterizing mechanophore reactivity, addressing sensitivity limitations in sonication experiments and, combined with computational modeling, revealing underlying principles that govern force-induced bond activation. Separately, we establish a synergistic platform that couples cargo-releasing mechanophores with biocompatible focused ultrasound, enabling controlled release of a fluorophore and a chemotherapeutic agent under physiological conditions. Finally, we demonstrate mechanochemically triggered drug delivery in vivo and validate this process using an inducible protein-expression system as a biological readout, achieving the first constructive modulation of cellular function enabled by covalent polymer mechanochemistry. Together, these studies deepen the fundamental understanding of mechanophore reactivity and illustrate the substantial biomedical potential of mechanochemical approaches using ultrasound activation.

Item Type:

Thesis (Dissertation (Ph.D.))

Subject Keywords:

Polymer Mechanochemistry, Mechanochemistry, Ultrasonication, focused ultrasound, in vivo drug delivery, gas vesicle

Degree Grantor:

California Institute of Technology

Division:

Chemistry and Chemical Engineering

Major Option:

Chemistry

Thesis Availability:

Public (worldwide access)

Research Advisor(s):

Robb, Maxwell J.

Thesis Committee:

Stoltz, Brian M. (chair)
Nelson, Hosea M.
Shapiro, Mikhail G.
Robb, Maxwell J.

Defense Date:

5 December 2025

Funders:

Funding Agency	Grant Number
CCE Innovation Grant	UNSPECIFIED
Beckman Young Investigator Award	UNSPECIFIED
National Institute of General Medical Sciences of the National Institutes of Health	R35GM150988

Record Number:

CaltechTHESIS:01212026-163903186

Persistent URL:

https://resolver.caltech.edu/CaltechTHESIS:01212026-163903186

DOI:

10.7907/mnbj-xq97

Related URLs:

URL	URL Type	Description
https://doi.org/10.1021/acspolymersau.2c00047	DOI	Article adapted for Ch.2
https://doi.org/10.1073/pnas.2309822120	DOI	Article adapted for Ch.3

ORCID:

Author	ORCID
Luo, Meng (Stella)	0000-0003-4003-7468

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No commercial reproduction, distribution, display or performance rights in this work are provided.

ID Code:

17830

Collection:

CaltechTHESIS

Deposited By:

Meng Luo

Deposited On:

28 Jan 2026 17:53

Last Modified:

09 Mar 2026 21:30

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Polymer Mechanochemistry Using Ultrasound: From Fundamental Reactivity to Controlled Drug Delivery Thesis by Meng (Stella) Luo In Partial Fulfillment of the Requirements for the degree of Doctor of Philosophy CALIFORNIA INSTITUTE OF TECHNOLOGY Pasadena, California 2026 (Defended Dec 5, 2025) ii ã 2026 Meng Luo ORCID: 0000-0003-4003-7468 iii ACKNOWLEDGEMENTS My journey as a graduate student at Caltech has been indescribable. Every challenge, failure, and success has shaped me into a better scientist and a more resilient person. First and foremost, I would like to express my deepest gratitude to my advisor, Professor Maxwell Robb. Over the past five years, he has taught me how to think about and solve problems as a chemist, supported me as my project expanded into unfamiliar territory, and guided me toward understanding what I hope to accomplish as a researcher. Working with Max, I learned how to identify and focus on important scientific questions when I felt lost in the possibilities, how to communicate progress with clarity and confidence, and how to cultivate the creative and meticulous approach that I will carry forward in my career. I am also profoundly grateful to Professor Mikhail Shapiro, who welcomed me into his lab as a collaborating researcher, provided opportunities for me to explore a completely new field, and inspired me with his curiosity, positivity, and passion for advancing science. My committee chair, Professor Brian Stoltz, and committee member, Professor Hosea Nelson, encouraged me to think deeply and critically about chemistry and offered invaluable guidance toward my academic and personal growth. I also want to thank my undergraduate advisor, Professor Dennis Cao. Thank you for offering me a position in the Cao Lab at Macalester College. This opportunity became the starting point and foundation of my entire academic career. Under your guidance, I was trained to become an independent chemist, and I am grateful for your continued support well beyond my time at Macalester. I am deeply appreciative of the staff in the Division of Chemistry and Chemical Engineering for their help over the years. Dave Vander Velde assisted with the characterization of many complex and enormous molecules. Scott Virgil provided indispensable help with the Robb group instruments, and I greatly enjoyed serving as his teaching assistant. Alison Ross helped me navigate several complicated situations. Annette Luymes kept the Robb group running smoothly and was a wonderful friend throughout my PhD. Joe Drew diligently kept the Church third floor functional and safe. Additionally, I am thankful to the lab management team in the Shapiro Lab, the OLAR office, the Caltech Imaging Facility, and the Caltech iv Flow Cytometry and Cell Sorting Facility for helping me navigate my new journey into biology. I feel fortunate to have met such incredible colleagues in the Robb group. Even though I joined during the pandemic, the senior members welcomed me warmly. Ross Barber was a dedicated mentor from whom I learned countless synthetic chemistry skills and how to have a resilient attitude toward research. Molly McFadden offered tremendous help when I joined her collaboration project with the Shapiro Lab and taught me everything about managing lab operations. Anna Overholts was always insightful in mechanistic discussions and taught me the most rigorous approach to kinetic experiments. I was beyond lucky to join the group with Skylar Osler. I learned invaluable attitudes and perspectives from her and felt blessed to have gone through everything over the five years together. She consistently engaged in deep discussions about my project, offered critical support and advice, and shaped me into the researcher I am today. Yan and Debbie joined just a year after me, yet both were so talented and experienced that I was able to learn from them and grow alongside them. I also feel fortunate to have had the opportunity to mentor Madie Patch, an experience that helped me grow into a more mature scientist. I was equally lucky to become close friends with the Robb group members. I will always cherish my first camping trip, our outings for boba and ice cream, and summer softball games, which I struggled with but had so much fun. I will forever remember the Chinese holidays with May, Yan, Debbie, Xiaoran, and Peng, and all the good food that made me feel at home. I especially treasure the days spent in office 311 with my dearest friends Skylar, Molly, Ross, and Yan. That room holds countless memories, as we shared research insights, talked deeply about life, and supported one another through joy and tears. I would like to mention Skylar again, who has been my greatest support in graduate school. We navigated many challenges together, and I am deeply grateful for her care, wisdom, and the joy she brought into my life. Outside of the Robb group, I was lucky to have met many wonderful people in the Shapiro Lab. Yuxing Yao was an invaluable mentor, collaborator, and a good friend. His scientific v insight, career guidance, and personal encouragement helped me grow in countless ways. I would also like to thank, in no particular order, Ann, George, Richard, Ernesto, Daniel, Brian, Hao, Jee Won, Ishaan, Josh, Trevor, Julio, Kaiwen, Baptist, Dina, and Zhiyang for teaching me biology from the ground up, providing creative solutions from different perspectives, and filling my time in the Shapiro Lab with so much laughter. I felt an unprecedented sense of belonging at Caltech because of these friends. With the original team: Yuxing, Zhiyang, Jee Won, and Yan, we shared countless game nights, always fighting together as a team. Karaoke nights, dinners, and gatherings with Yuxing, May, Jee Won, Yan, Ziyang, Hao, Kaiwen, Kun, Hengyu, and Changfan created some of the happiest moments of my time at Caltech. Living in the LA area also allowed me to enjoy many unforgettable concerts with like-minded friends such as Yuxing, Jee Won, Natalie, Hao, and many others. I am also grateful to all the friends who helped look after my cats. I am incredibly fortunate to have support that extends beyond Caltech as well. Meihua has been my closest friend for the past nine years. I was lucky to spend high school, college, and my graduate school years in the same city with her. She has listened to every experience, celebrated each milestone, and encouraged me through every stage of this journey. Our band performances, game nights, dinner and boba trips, and deep conversations have brought me immense groundedness, comfort, and joy. Above all, I am deeply grateful to my parents for their unwavering love and support. Their faith and trust in me have empowered me to become the person I aspire to be and have given me the courage to pursue my goals wholeheartedly. During the past five years, when I was unable to return home, they took many 16-hour overseas flights to visit me, take care of me, and bring the feeling of home across the world. Their presence during the most challenging moments reminded me of where I come from and what truly matters. Everything I have achieved is built upon their love. I want to thank my partner, Jincheng. He supported me every single day throughout graduate school. Because of his wholehearted care and understanding, I was able to focus on research, maintain a healthy life, and have a true home. He has encouraged me, stood by me, and vi supported me as I grew into a braver and more confident person. He has been my greatest companion, my constant supporter, and my best friend. I treasure every moment we have shared and every journey we have taken together. Lastly, I want to extend my gratitude to all of you. Words and tears cannot fully express how thankful I am. Your guidance, kindness, and support have shaped me into the person I am today. The successes and achievements in this thesis were made possible because of you. I will carry forward everything you have given me, and I thank you sincerely for being part of my life during my time at Caltech. vii ABSTRACT Polymer mechanochemistry harnesses mechanical force to drive specific chemical transformations in stress-sensitive molecules known as mechanophores. Through judicious chemical design, these force-driven reactions have enabled functional polymeric materials capable of sensing, self-healing, and catalysis. Research in this field encompasses expanding the mechanophore repertoire, elucidating the fundamental principles governing mechanically induced reactivity, and translating force-responsive systems into practical applications. This thesis advances the field by contributing both fundamental insight and applied functionality in mechanophore activation under ultrasonication, with a particular focus on controlled drug release in biological environments. First, we develop an improved methodology for characterizing mechanophore reactivity, addressing sensitivity limitations in sonication experiments and, combined with computational modeling, revealing underlying principles that govern force-induced bond activation. Separately, we establish a synergistic platform that couples cargo-releasing mechanophores with biocompatible focused ultrasound, enabling controlled release of a fluorophore and a chemotherapeutic agent under physiological conditions. Finally, we demonstrate mechanochemically triggered drug delivery in vivo and validate this process using an inducible protein-expression system as a biological readout, achieving the first constructive modulation of cellular function enabled by covalent polymer mechanochemistry. Together, these studies deepen the fundamental understanding of mechanophore reactivity and illustrate the substantial biomedical potential of mechanochemical approaches using ultrasound activation. viii PUBLISHED CONTENT AND CONTRIBUTIONS Luo, S. M.; Barber, R. W.; Overholts, A. C.; Robb, M. J. Competitive Activation Experiments Reveal Significantly Different Mechanochemical Reactivity of Furan– Maleimide and Anthracene–Maleimide Mechanophores. ACS Polym. Au. 2023, 3, 202–208. https://doi.org/10.1021/acspolymersau.2c00047. S.M.L participated in the design of the project, synthesized materials, ran experiments, analyzed data, and wrote the manuscript. Yao, Y.;‡ McFadden, M. E.;‡ Luo, S. M.; Barber, R. W.; Kang, E.; Bar-Zion, A.; Smith, C. A. B.; Jin, Z.; Legendre, M.; Ling, B.; Malounda, D.; Torres, A.; Hamza, T.; Edwards, C. E. R.; Shapiro, M. G.; Robb, M. J. Remote Control Of Mechanochemical Reactions Under Physiological Conditions Using Biocompatible Focused Ultrasound. Proc. Natl. Acad. Sci. U.S.A. 2023, 120, e2309822120. https://doi.org/10.1073/pnas.2309822120. S.M.L participated in the design of the project, synthesized materials, ran experiments, and analyzed data. 5 TABLE OF CONTENTS ACKNOWLEDGEMENTS ............................................................................................... III ABSTRACT ....................................................................................................................... VII PUBLISHED CONTENT AND CONTRIBUTIONS .................................................. VIII Chapter 1. INTRODUCTION..............................................................................................6 REACTIVITY STUDIES OF MECHANOPHORES .............................................................................8 BIOCOMPATIBLE DEMONSTRATIONS OF CARGO-RELEASING MECHANOPHORES .................. 10 REFERENCES .......................................................................................................................... 12 Chapter 2. COMPETITIVE ACTIVATION EXPERIMENTS REVEAL SIGNIFICANTLY DIFFERENT MECHANOCHEMICAL REACTIVITY OF FURAN–MALEIMIDE AND ANTHRACENE–MALEIMIDE MECHANOPHORES ............................................................................................................................................... 24 2.1 INTRODUCTION ................................................................................................................. 26 2.2 RESULTS AND DISCUSSION ............................................................................................... 28 2.3 CONCLUSION .................................................................................................................... 33 2.4 ACKNOWLEDGMENTS ....................................................................................................... 34 2.5 EXPERIMENTAL DETAILS ................................................................................................. 34 2.6 REFERENCES..................................................................................................................... 72 Chapter 3. REMOTE CONTROL OF MECHANOCHEMICAL REACTIONS UNDER PHYSIOLOGICAL CONDITIONS USING BIOCOMPATIBLE FOCUSED ULTRASOUND................................................................................................................... 79 3.1 INTRODUCTION ................................................................................................................. 81 3.2. RESULTS AND DISCUSSION .............................................................................................. 83 3.3 CONCLUSIONS .................................................................................................................. 88 3.4 MATERIALS AND METHODS ............................................................................................. 89 3.5 ACKNOWLEDGEMENTS ..................................................................................................... 90 3.6 EXPERIMENTAL DETAILS ................................................................................................. 90 3.7 REFERENCES................................................................................................................... 124 Chapter 4. IN VIVO CONTROL OF POLYMER MECHANOCHEMISTRY USING BIOCOMPATIBLE FOCUSED ULTRASOUND REPORTED BY GENE EXPRESSION ................................................................................................................... 130 6 Chapter 1 INTRODUCTION Chemical reactions fundamentally rely on the input of energy to overcome activation barrier and promote bond rearrangement. Traditionally, thermal, photochemical and electrochemical energies have been recognized as major sources of activation, enabling reactions through heat-induced molecular motion, photon absorption, or electron transfer processes.1 In contrast, mechanical forces represent one of the most fundamental forms of energy in nature, as seen in processes such as friction, tension and compression. However, the use of mechanical energy as a direct and controllable driving force to activate specific chemical bonds and trigger well-defined molecular transformations has only gained systematic attention recently.2 The field that explores mechanically induced transformations is known as mechanochemistry. Broadly defined, mechanochemistry studies how mechanical energy can be converted into chemical potential, enabling bond scission, rearrangement, or even formation through applied stress or strain.3 As a natural driving force, mechanochemistry contributes to Earth’s chemical cycles by initiating redox and decomposition reactions through shear and fracture in rocks and minerals.4,5 Early chemical studies primarily sought to harness mechanical energy for solvent-free, solid-state chemical synthesis,6 in which grinding or milling promotes reactions through friction and compression between particles, offering environmentally sustainable routes to chemical production.7 More recent advances have shifted the focus toward molecular-level control, enabling chemoselective activation of specific bonds and well-defined mechanochemical transformations.8,9 This evolution has paved the way for polymer mechanochemistry, where mechanical force is transmitted along covalent backbones to selectively activate embedded mechanosensitive motifs.10,11 Polymer mechanochemistry originated from the longstanding observation that mechanical force can rupture covalent bonds within polymer backbones, a process traditionally associated with material fatigue and degradation.12,13 Recognizing that such force-induced bond scission reflects a direct input of energy, researchers began to envision 7 ways to redirect this unselective and destructive chemistry towards precise and productive molecular responses. Encina and co-workers observed that some chemical bonds are more readily cleaved under force than others.14 This work led to the principle of bond-specific activation and served as the foundation for the development of mechanophores, which are mechanically reactive species that undergo predictable and useful chemical transformation in response to applied force.15,16 Since the first report of an azo mechanophore in 2005,17 the field of polymer mechanochemistry has expanded to encompass a wide range of mechanochemically active motifs, involving processes such as heterolytic and homolytic bond cleavage, retrocycloadditions, and electrocyclic ring opening reactions.18 These chemical transformations are judiciously designed to impart practical functions, including force-induced changes in color or luminescence,19–23 release of small molecules or functional groups,24–28 and modulation of electrical conductivity.29,30 These mechanochemically responsive behavior has enabled the development of force-sensitive materials that can be exploited for stresssensing,31–33 enhanced mechanical properties,34–36 or mechanically controlled catalysis.37,38 To enable force transduction to mechanophore units, these motifs are typically embedded within linear or crosslinked polymer networks, which allow their activation under different modes of applied stress. In solid-state materials, mechanophores are most commonly activated by tensile, compressive, or shear force during macroscopic material deformation39–42. In solution, the predominant method for inducing force along polymer chain is ultrasonication, which generates elongational forces capable of selectively activating mechanophores.43–45 Across these experimental platforms, major aims of the field include elucidating the fundamental principles governing force-induced bond activation,46,47 developing new mechanophore structures,10,40 advancing methodologies to analyze and quantify mechanophore reactivity,48–51 and exploring practical36,52 and translational53–55 application of polymer mechanochemistry. In this thesis, we focus specifically on evaluating mechanophore activation using ultrasound. We develop methodologies to analyze and compare mechanophore reactivities under ultrasonic conditions and demonstrate cargo release from mechanophorefunctionalized polymers activated by biocompatible focused ultrasound. 8 Reactivity Studies of Mechanophores When beginning to investigate mechanophore systems, computational methods provide predictions of how mechanical forces reshape their chemical reactivity.56 Among these, advanced quantitative approaches such as Force-Modified Potential Energy Surface (FMPES),57 which simulates how external force directly alters the potential energy surface and reaction barrier; External Force Explicitly Included (EFEI),58 which generates forcemodified molecular structures, and Ab initio Steered Molecular Dynamics (AISMD),59 which captures the dynamic response of molecular systems under mechanical load, have proven useful in explanatory investigations. Although these methods provide detailed mechanistic insights into mechanophore activation, their implementation often demands specialized computational proficiency and time investment. In contrast, the Constrained Geometries simulate External Force (CoGEF) method provides a simpler, qualitative alternative for the initial screening of mechanophore candidates and for visualizing molecular structures under explicitly applied strain.60 By incrementally elongating the distance between two terminal atoms of a molecule and optimizing its constrained geometry at each step, this methods predicts the mechanochemical outcome that accompany mechanical distortion, and estimates the maximum force (Fmax) associated with the chemical transformation.60 Recently, CoGEF has been validated as a predictive method through comparison with empirically recognized mechanochemical transformations such as retrocycloadditions, electrocyclic ring-opening reactions, and homolytic bond cleavages.61 Experimentally, the reactivity of mechanophores can be determined and compared through several approaches. Single-Molecule Force Spectroscopy (SMFS)62 provides the most direct measurement of mechanochemical activation forces by stretching a mechanophore-containing polymer chain under controlled conditions.63–65 However, SMFS requires specialized instrumentation and is limited by its loading-rate dependence and challenges in reproducibility. In addition, some mechanophores present synthetic difficulties when incorporated into polymers compatible with SMFS setups. Ultrasonication, in contrast, is a highly reproducible and experimentally accessible technique for characterizing mechanophore reactivity in solution.44,66,67 During ultrasonication, mechanical force is transduced along the polymer backbone through the solvodynamic shear field generated by 16 cavitation events of dissolved gas bubbles. 9 Conveniently, conventional analytical methods like UV-Vis spectroscopy, gel permeation chromatography (GPC) or highperformance liquid chromatography (HPLC), enable quantitative characterization of the reactivities of mechanophores incorporated into linear polymer backbones. However, it is challenging to validate and compare the reactivates predicted by the CoGEF method with respect to those measured in ultrasonication experiments. The maximum force predicted by CoGEF does not account for the influence of thermal energy during sonication experiments.60,61 As a result, the predicted rupture force does not accurately reflect the combined effects of strain rate, thermal contribution, and dynamic pathways during sonication experiments. Moreover, the observed activation kinetics in solution-phase ultrasonication depend strongly on the molecular weight and dispersity of the polymer backbone,51,68,69 making it difficult to directly compare rate constants across different polymer samples. These limitations highlight the need for experimental approaches that enable meaningful comparison of reactivity across different mechanophores and provide a clearer connection to computational predictions. To enable direct comparison between different mechanophores, competitive experiments, either between a mechanophore candidate and a reference unit with known reactivity, or between two mechanophore candidates, have proven highly informative in many studies70–72. For instance, the Craig group frequently benchmarked the reactivity of new mechanophores against gem-dichlorocyclopropane (gDCC) mechanophores embedded in multimechanophore polymers synthesized via ring opening metathesis polymerization (ROMP).71,73,74 However, this strategy is typically applied to comparison between nonscissile mechanophores, whose activations do not alter polymer chain length under ultrasound activation. In contrast, chain cleavage events associated with scissile mechanophores alter the local force environment of the nearby mechanophore units, thereby convoluting the resulting reactivity analyses. In chapter 2, we extend the competitive approach to compare two scissile mechanophores, furan–maleimide (FM) and anthracene-maleimide (AM), under ultrasonication.75 By covalently linking the two mechanophores into a bis-adduct architecture, both units are positioned at the center of the same polymer chain and thus 10 experience identical mechanical forces during sonication. The results revealed that the FM adduct exhibits significantly higher mechanochemical reactivity than the AM adduct, despite their CoGEF-predicted rupture forces being nearly indistinguishable. We further examined the CoGEF modeling for both adducts and identified differences in their mechanochemical coupling efficiencies, which offered a potential explanation for their different reactivities under force. Together, these findings demonstrated that competitive sonication experiments, combined with computational analysis, provide a robust and generalizable methodology for evaluating the reactivity of mechanophores. Biocompatible Demonstrations of Cargo-Releasing Mechanophores Although ultrasonication is a convenient method to activate mechanophores in the chemistry laboratory, its practical applicability is limited when compared to solid-state activation modes that can be readily translated into developing functional materials. In solution, low-frequency ultrasound (20 kHz) is highly efficient in promoting cavitations of dissolved gas bubbles, generating large solvodynamic forces that activate mechanophores.16,67 However, the ultrasound intensities required to achieve this cavitation also produce extreme localized heating and violent mechanical effects, rendering these conditions highly destructive to living tissue.76,77 As a result, traditional probe ultrasonication is fundamentally incompatible with biological environments and present significant barriers to translational applications of polymer mechanochemistry.78 In contrast to low-frequency laboratory ultrasonication, ultrasound at higher frequencies is widely used in clinical settings.76,79 Diagnostic ultrasound, typically operating in the megahertz (MHz) range, is routinely employed for noninvasive imaging,80 such as obstetric sonography, and provides high spatial and temporal resolution with excellent safety. Beyond imaging, therapeutic applications of ultrasound have been developed through the use of focused ultrasound (FUS).81 Sound waves emitted from a transducer with a concave geometry can converge acoustic energy at a defined focal point, achieving sub-millimeter precision. At the focal zone, therapeutic ultrasound can induce biological effects through thermal heating or mechanical effects, enabling noninvasive clinical applications including tissue and tumor ablation,82,83 neuromodulation,84,85 and drug delivery.86,87 Despite these advances, coupling clinically relevant ultrasound frequencies and intensities to drive 11 chemical transformations remain a significant challenge. The acoustic conditions accessible in clinical ultrasound are orders of magnitude less intense than those required to induce mechanochemical reactions via laboratory sonication, and the mechanisms of force delivery differ substantially. As a result, the translation of ultrasound-activated mechanochemistry into in vivo or therapeutic applications has remained elusive. Efforts within polymer mechanochemistry have increasingly focused on bridging this translational gap by exploring the use of higher-frequency, clinically relevant ultrasound for mechanophore activation. Early work by the Moore group demonstrated that high-intensity focused ultrasound (HIFU) could activate naphthopyran mechanophores in PDMS films, likely through an acoustic radiation force mechanism.88 However, the acoustic intensities required for activation were approximately two orders of magnitude above the clinically safe range and produced substantial local heating (> 30 °C observed above baseline at the focal point), making this approach unsuitable for biological applications. More recently, a highly labile azo mechanophore was shown to generate free radicals under focused ultrasound, enabling mechanochemical dynamic therapy (MDT) in hydrogels89 and in vivo.53 Nonetheless, this demonstration is limited to the generation of radicals and reactive oxygen species (ROS), hindering constructive therapeutic applications. In parallel, work from the Göstl and Hermann groups demonstrated FUS-triggered release of therapeutic oligonucleotides to modulate biological function.90 However, these findings were not applied to promote covalent mechanochemical transformations under fully biocompatible FUS conditions, leaving a significant need for more modular and productive ultrasound-activated chemistries. In 2019, our group introduced a modular cargo-releasing platform based on from a masked 2-furylcarbinol motif.25 These furan-maleimide derived mechanophores exhibited high thermal stability and broad cargo compatibility,91 making them promising candidates for translational uses in ultrasound-controlled drug delivery. In Chapter 3, we describe the investigation of the remote activation of these mechanophores using biocompatible focused ultrasound in physiological conditions. By leveraging Gas Vesicles (GVs), an air-filled protein structure that nucleates cavitation events to promote force transduction,92 we established biocompatible acoustic conditions that enable efficient mechanophore activation. 12 Under these conditions, we demonstrated the mechanochemical release of fluorogenic reporters and a chemotherapeutic payload for targeted cancer treatment.93 To further illustrate the biomedical potential of this platform, we extend these efforts to demonstrate small-molecule release in complex in vitro and in vivo environments. A central challenge in these settings is that quantifying small molecules in biological samples require extensive sample processing,94 and can be confounded by intracellular sequestration, metabolism or nonspecific interactions.95 To overcome these limitations, we proposed an inducible protein-expression system96 that responds selectively to mechanochemically released small molecule, effectively translating and amplifying otherwise elusive chemical signals into robust biological readouts that can be directly measured. In Chapter 4, we employ this system to validate FUS-induced mechanochemical delivery of small molecules from hydrogels in the presence of living cells and in in vivo models. This work represents the first demonstration of in situ polymer mechanochemistry controlled by FUS in living systems to enable constructive modulation of cell function, underscoring the broader potential of ultrasound-activated mechanochemical technologies. References (1) Silberberg, M. S. 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Au 2023, 3, 202–208 25 Abstract During the past two decades, our understanding of mechanochemical reactivity has advanced considerably. Nevertheless, an incomplete knowledge of structure–activity relationships and the principles that govern mechanochemical transformations limits molecular design. The experimental development of mechanophores has thus benefited from simple computational tools like CoGEF, from which quantitative metrics like rupture force can be extracted to estimate reactivity. Furan–maleimide (FM) and anthracene–maleimide (AM) Diels–Alder adducts are widely studied mechanophores that undergo retro-Diels– Alder reactions upon mechanical activation in polymers. Despite possessing significantly different thermal stability, similar rupture forces predicted by CoGEF calculations suggest that these compounds exhibit similar mechanochemical reactivity. Here, we directly probe the relative mechanochemical reactivity of FM and AM adducts through competitive activation experiments. Ultrasound-induced mechanochemical activation of bis-adduct mechanophores comprising covalently tethered FM and AM subunits reveals pronounced selectivity—as high as 13:1—for reaction of the FM adduct compared to the AM adduct. Computational models provide insight into the greater reactivity of the FM mechanophore, indicating a more efficient mechanochemical coupling for the FM adduct compared to the AM adduct. The methodology employed here to directly interrogate the relative reactivity of two different mechanophores using a tethered bis-adduct configuration may be useful for other systems where more common sonication-based approaches are limited by poor sensitivity. 26 2.1 Introduction Mechanical force is used as a stimulus to promote specific chemical reactions of force-sensitive molecules called mechanophores.1,2 In polymer mechanochemistry, polymer chains covalently attached to the mechanophore facilitate force transduction and the mechanically selective scission of labile bonds in mechanophores enables productive activation pathways to outcompete the typical degradative process of nonspecific polymer backbone scission.3 Over the past two decades, a wide variety of mechanophores have been developed that provide functional polymers with unique force-responsive behaviors such as stress and damage reporting through changes in color or luminescence,4–8 release of small molecules,9,10 conductivity switching,11 catalyst activation,12,13 and many others.14,15 However, despite the rapidly expanding repertoire of mechanophores and force-responsive polymers, intuition of mechanochemical reactivity is still limited, particularly as trends in mechanophore reactivity often diverge from those expected under more conventional reaction manifolds.16–20 Computational methods are commonly employed to provide insight into mechanophore reactivity and support an evolving but incomplete understanding of structure– mechanochemical activity relationships that include stereochemistry21–25 and regiochemistry17,25–29 of the mechanophore as well as its connectivity within the polymer backbone.30–33 In our group and in many others,3 mechanophore candidates are typically first evaluated with density functional calculations using the constrained geometries simulated external force (CoGEF) method.34,35 This simple and powerful technique enables the qualitative prediction of mechanochemical reactions while also providing an estimate of the maximum force (Fmax) associated with the chemical transformation. Importantly, the forces calculated using CoGEF provide a framework for comparing the relative reactivity of different mechanophores that agrees well with experimental trends.35 For example, a recent study investigating the selectivity of mechanophore activation compared to nonspecific polymer chain scission confirmed that a coumarin dimer mechanophore36 is much less reactive than an anthracene–maleimide (AM) mechanophore,17,37–39 consistent with the significantly different predicted Fmax values of 5.9 and 4.1 nN, respectively.40 It should be noted that relevant Fmax values from CoGEF are in a narrow range < 6.0 nN, which represents 27 an apparent upper limit for selective mechanophore reactivity. On the other hand, it is 35 much more challenging to make reliable predictions about differences in mechanochemical reactivity for mechanophores with similar predicted values of Fmax. For instance, Wang and Craig demonstrated that the endo and exo isomers of a furan–maleimide (FM) mechanophore41 exhibit significantly different reactivity upon ultrasound-induced mechanical activation,21 despite similar predicted Fmax values of 3.9 nN (endo) and 4.1 nN (exo).35 A typical approach to assess mechanophore reactivity experimentally is to measure the rate of mechanophore reaction upon ultrasound-induced mechanochemical activation of polymers.22 For scissile mechanophores, rates of chain scission in ultrasonication experiments are frequently used as a proxy for mechanophore activation to compare relative reactivity.42 However, the insensitivity of this method has proven intractable in some cases for distinguishing the reactivity of different mechanophores. In an elegant study by Stevenson and De Bo, the rate of fragmentation was shown to vary for FM adducts with proximal and distal pulling geometries, but no discernable difference was observed for endo and exo isomers of the FM mechanophore with proximal pulling geometry.25 A more sensitive methodology pioneered by the Craig group relies on the internal competition between the activation of a scissile mechanophore and the ring-opening reaction of gemdichlorocyclopropane mechanophores distributed along the polymer chain.19 As mentioned earlier, this approach was able to clearly distinguish the mechanical lability of endo and exo stereoisomers of the FM mechanophore, highlighting the enhanced sensitivity provided by internal competition experiments as opposed to comparing absolute rates of chain scission.21 Alternatively, covalently coupling two mechanophores in a tandem configuration would ensure that each is exposed to the same mechanical forces and provide an approach for directly comparing their relative reactivity. This methodology was employed by De Bo and coworkers to study the impact of having adjacent FM mechanophores on mechanochemical coupling,43 but to the best of our knowledge, it has not been used to compare the relative reactivity of distinct mechanophores. 40 In the course of other investigations, 28 we became interested in the relative mechanochemical reactivity of the AM and FM mechanophores. Despite significantly different thermal stabilities, the similar values of Fmax predicted by CoGEF for these Diels– Alder adducts would suggest similar mechanical sensitivity. Here we study the selectivity of mechanochemical activation between FM and AM Diels–Alder adducts through direct competitive activation experiments. We designed two bis-adduct Scheme 2.1. Competitive Retro-Diels–Alder Reaction Induced by Ultrasonication in a Tandem Bis-Adduct mechanophores comprising an AM Mechanophore Design. and a FM Diels–Alder motif with either endo or exo stereochemistry connected in tandem by an alkyl tether (Scheme 2.1). Due to the scissile nature of each Diels–Alder adduct, only a single retro-Diels– Alder reaction will take place per mechanochemical activation event, allowing the extent to which each AM or FM adduct has reacted upon mechanical extension to be quantified. We find that the product distribution following ultrasound-induced mechanical activation of the bis-adduct mechanophores exhibits significant selectivity—as high as 13:1—for activation of the FM adduct. Computational modeling indicates that the selectivity for FM activation originates from a more efficient mechanochemical coupling. 2.2 Results and Discussion CoGEF calculations were initially performed to evaluate the mechanochemical reactivity of bis-adducts AM–FM(endo) and AM–FM(exo) with either endo or exo stereochemistry of the FM subunit, respectively (Figure 2.).34,35 The retro-Diels–Alder reaction of the FM subunit is predicted to occur in both cases, with calculated Fmax values of 4.0 nN for AM–FM(endo) and 4.2 nN for AM–FM(exo). These Fmax values differ minimally from those of the isolated endo and exo FM adducts, which again are 3.9 and 4.1 nN, respectively.35 CoGEF calculations performed on analogous bis-adducts incorporating an ethylene tether instead of 29 the propylene tether provided identical results, suggesting that the reactivity is not significantly affected by tether length (Figure 2.S1). To further probe the mechanochemical activity of the AM subunit in the tethered bis-adduct design, CoGEF calculations were performed in which the junction bonds of the FM adduct were constrained to their equilibrium length. In this case, a Figure 2.1 DFT calculations using the CoGEF method predict the retro-Diels–Alder reaction of the furan– maleimide (FM) subunit upon mechanical elongation of the bis-adduct mechanophores for both endo and exo isomers. The predicted products shown correspond to the data points indicated with arrows. Calculations were performed at the B3LYP/6-31G* level of theory. retro-Diels–Alder reaction of the AM adduct was predicted to occur with an Fmax of 4.4 nN for both stereoisomers (Figure 2.S2). These results confirm that there are not geometrical or structural constraints in the bis-adduct configuration that significantly bias the reactivity of the AM mechanophore unit. Despite the similar Fmax values for reactions of the isolated FM and AM subunits, the calculations performed on the bis-adduct mechanophores suggest unambiguously that the FM adduct is more mechanochemically reactive than the AM adduct. Following these computational predictions, we synthesized a pair of poly(methyl acrylate) (PMA) polymers incorporating the AM–FM bis-adducts near the chain center to experimentally evaluate the selectivity of ultrasound-induced mechanochemical retro-Diels– Alder reactions. Bis-adduct initiators were synthesized and subsequently used in the controlled radical polymerization of methyl acrylate to afford PMA polymers incorporating AM–FM(endo) (Mn = 106 kDa; Đ = 1.20) and AM–FM(exo) (Mn = 103 kDa; Đ = 1.19) bisadducts near the center of the chains where mechanical force is maximized (Figures 2.S3 and 2.S4).3 The similar molecular weight and dispersity of the two polymers ensures that 30 each bis-adduct mechanophore will experience comparable forces during ultrasonication (Figure 2.S5).44 Two separate sets experiments quantify of were the ultrasonication performed to mechanochemical generation of anthracene and furan products via the respective retro-Diels– Alder reactions of the AM and FM subunits (Figure , see the SI for details). To determine AM activation, dilute solutions of each polymer were subjected to pulsed ultrasonication (2 mg/mL in MeCN, 1 s on/ 2 s off, 6–9 Figure 2.2 (a) Ultrasound-induced mechanochemical activation of polymers containing chain-centered AM– FM mechanophores produces either an anthracene or a furan terminated polymer. (b) Activation of the AM adduct is characterized using photoluminescence spectroscopy (λex = 365 nm) monitored at 413 nm, and (c) activation of the FM adduct is characterized using 1H NMR spectroscopy. Representative data are shown for the AM–FM(endo) mechanophore. °C, 20 kHz, 8.2 W/cm2) in the presence of BHT (30 mM), which is necessary to prevent anthracene.42 the degradation Formation of of the anthracene product was monitored by photoluminescence (PL) spectroscopy and quantified using a calibration curve (Figure a).42 A separate set of sonication experiments was performed under nearly identical conditions to measure the mechanochemical activation of the FM subunit in each bis-adduct mechanophore using 1H NMR spectroscopy,25,40 but in the absence of BHT to prevent spectral contamination (Figure 2.2c). Ultrasonication experiments performed on a low molecular weight control polymer (Mn = 18 kDa; Đ = 1.10) containing a chain-centered AM–FM(endo) bis-adduct mechanophore demonstrate negligible reactivity, consistent with a mechanochemical transformation (Figure 2.S6). The selectivity of the mechanically-induced retro-Diels–Alder reactions was determined by evaluating the percent activation of the FM and AM subunits in each bis- adduct 31 upon mechanophore ultrasonication.40,42 The plateau value obtained from fitting the time- dependent activation data for each adduct to an expression of first order kinetics provides an estimate of the total mechanophore activation, which is illustrated for the AM and FM adducts in Figure 2.3a and 2.3b, respectively. Both bis-adducts exhibit a much higher selectivity for activation of the FM subunit than for the AM subunit. Specifically, ultrasonication of AM–FM(endo) results in ~80% activation of the FM subunit and ~6% activation of the AM subunit. For AM– FM(exo), ultrasonication leads to ~66% retro-Diels–Alder reaction of the FM subunit and ~9% activation of the AM subunit. The differences in activation between the stereoisomers Figure 2.3 Activation of (a) AM and (b) FM subunits as a function of sonication time quantified using PL and 1H NMR spectroscopy, respectively. Data points are averages from two experiments with error bars denoting the range of the two measurements. Dashed lines are fits to a firstorder rate expression. are consistent with the greater reactivity of the endo FM adduct in comparison to the exo FM adduct,21 which results in slightly greater competition in the case of AM–FM(exo). Regardless, FM activation outcompetes AM activation by a 7:1 ratio for AM–FM(exo), and by a 13:1 ratio for AM–FM(endo). The total mechanophore activation efficiency is ≥ 75% for both bis-adducts, confirming the relatively high mechanochemical reactivity of each Diels–Alder adduct relative to nonspecific backbone cleavage, as expected.40 However, the significant selectivity for FM activation is 32 somewhat surprising given the similar Fmax values calculated for the retro-Diels–Alder reactions of the isolated adducts from CoGEF. To understand the significant selectivity for FM activation in the bis-adduct mechanophores, we again turned to CoGEF to evaluate the efficiency with which mechanical force is transduced to achieve a mechanochemical reaction. approaches retro-Diels–Alder Rigorous experimental to quantifying such mechanochemical coupling have invoked the restoring force of strained molecules45,46 and single molecule force spectroscopy techniques,30,47,48 in addition to comprehensive theoretical models.49–51 Alternatively, De Bo and coworkers proposed a simple approach to characterize mechanochemical coupling using CoGEF by monitoring the sensitivity with which the scissile bond of a mechanophore elongates (d/d0) in response to global molecular elongation, as represented by increasing displacement between the terminal atoms (D/D0).25,52 Applying Figure 2.4 (a) Change in the scissile bond length for indicated Diels–Alder adducts in response to molecular extension from CoGEF calculations. (b) Strain distribution across AM–FM(endo) and AM–FM(exo) models immediately prior to bond rupture. Values of bond strain (d/d0) for the scissile bond in each adduct are indicated. this methodology to the AM–FM bisadduct mechanophore system provides some insight into the observed selectivity for FM activation (Figure 2.4a, see section 2.5 for details). A steeper curve indicates better mechanochemical coupling, again suggesting greater reactivity. Notably, the response–displacement curves for AM–FM(endo) and AM– 33 FM(exo) closely match those of isolated FM adducts, indicating that the tethered AM subunit does not significantly impact mechanochemical coupling of the FM subunit in the bis-adduct mechanophore. The response–displacement curve for an isolated AM mechanophore exhibits weaker mechanochemical coupling than any of the FM adducts. While this particular method of analysis is unable to differentiate the reactivity of endo and exo FM isomers established previously,21 it nevertheless successfully captures the differences in coupling between the FM and AM units in the bis-adduct mechanophores. De Bo and coworkers have previously proposed that differences in mechanochemical coupling should lead to observable differences in reactivity, even for mechanophores with similar predicted values of Fmax.52 In contrast to the results for mechanophores comprising adjacent FM adducts,43 here the pronounced difference in mechanochemical coupling predicted for the FM and AM adducts does indeed manifest in significantly greater reactivity for FM compared to AM in direct competitive activation experiments. The greater reactivity of the FM adducts compared to the AM adduct is also represented by mapping the strain distribution across each bis-adduct mechanophore at the point in the CoGEF calculations immediately prior to bond rupture (Figure 2.4b). These strain maps visually illustrate the efficiency with which force applied across the molecule is coupled to the elongation of specific bonds, and serve to reinforce the conclusions stated above. 2.3 Conclusion In summary, we find significant differences in the mechanochemical reactivity of furan–maleimide (FM) and anthracene–maleimide (AM) Diels–Alder adducts despite similar rupture forces predicted by CoGEF calculations. Competitive activation experiments performed on bis-adduct mechanophores comprising covalently tethered FM and AM subunits demonstrate pronounced selectivity for reaction of the FM adduct in polymers subjected to ultrasound-induced mechanical force. Specifically, mechanochemical activation of FM outcompetes that of AM by a factor of 13:1 in the bis-adduct with endo stereochemistry of the FM subunit and 7:1 for the exo isomer. Density functional calculations provide insight into the experimentally observed selectivity, indicating a more efficient mechanochemical coupling for the FM adduct compared to the AM adduct and highlighting the greater compliance of the former mechanophore under force. The competitive activation 34 of covalently tethered mechanophores is demonstrated to provide a reliable methodology for resolving the mechanochemical reactivity of otherwise similar motifs that may be difficult to ascertain by more common methods such as comparing absolute rates of chain scission for which sensitivity is limited. 2.4 Acknowledgments Financial support from Caltech is gratefully acknowledged. A.C.O was supported by a NSF Graduate Research Fellowship (DGE-1745301) and an Institute Fellowship from Caltech. We thank the Center for Catalysis and Chemical Synthesis of the Beckman Institute at Caltech for access to equipment. 2.5 Experimental Details 2.5.1 General Experimental Details Reagents from commercial sources were used without further purification unless otherwise noted. Methyl acrylate was passed through a short plug of basic alumina to remove inhibitor immediately prior to use. Copper wire was cleaned prior to use by soaking in 1 M HCl for 15 min and then rinsed consecutively with deionized water, acetone, and dichloromethane. Dry solvents were obtained from a Pure Process Technology solvent purification system. All reactions were performed under a N2 atmosphere unless specified otherwise. Column chromatography was performed on a Biotage Isolera system using SiliCycle SiliaSep HP flash cartridges. NMR spectra were recorded using a 400 MHz Bruker Avance III HD with Prodigy Cryoprobe or a 400 MHz Bruker Avance Neo spectrometer. All 1H NMR spectra are reported in δ units, parts per million (ppm), and were measured relative to the signal for residual chloroform (7.26 ppm) in deuterated solvent using a 5 s acquisition time. All 13C NMR spectra were measured in CDCl3 and are reported in ppm relative to the signal for chloroform (77.16 ppm). High resolution mass spectra (HRMS) were obtained from an Agilent 6230 series time-of-flight mass spectrometer equipped with an Agilent G1958 Jet Stream Electrospray Ionization Source. 35 Analytical gel permeation chromatography (GPC) was performed using an Agilent 1260 series pump equipped with two Agilent PLgel MIXED-B columns (7.5 x 300 mm), an Agilent 1200 series diode array detector, a Wyatt 18-angle DAWN HELEOS light scattering detector, and an Optilab rEX differential refractive index detector. The mobile phase was THF at a flow rate of 1 mL/min. Molecular weights and molecular weight distributions were calculated by light scattering using a dn/dc value of 0.062 mL/g (25 °C) for poly(methyl acrylate). Photoluminescence spectra were recorded on a Shimadzu RF-6000 spectrofluorophotometer using a quartz microcuvette (Starna 18F-Q-10-GL14-S). Ultrasound experiments were performed inside a sound abating enclosure using a Vibra Cell 505 liquid processor equipped with a 0.5 in. diameter solid probe (part #6300217), sonochemical adapter (part #830-00014), and a Suslick reaction vessel made by the Caltech glass shop (analogous to vessel #830-00014 from Sonics and Materials). Compounds 2.S1,53 PMA-Furan,53 2.S2,54 and 2.255 were prepared according to previously reported methods. 36 2.5.2. Supplementary Figures Chart 2.S1. Structures of the initiators and small molecules used in this study. Chart 2.S2. Structure of polymers used in this study. 37 Figure 2.S1. DFT calculations using the constrained geometries simulate external force (CoGEF) method for bis-adducts AM–C2H2–FM(endo) and AM–C2H2–FM(exo) incorporating a shorter ethylene linker predict the retro-Diels–Alder reaction of the furan– maleimide subunit upon mechanical elongation with Fmax values of 4.0 and 4.2 nN, respectively. Calculations were performed at the B3LYP/6-31G* level of theory. 38 Figure 2.S2. DFT calculations using the constrained geometries simulate external force (CoGEF) method for bis-adducts in which the lengths of the furan–maleimide junction bonds (colored red) were constrained to their equilibrium length. In this case, CoGEF predicts a retro-Diels–Alder reaction of the anthracene–maleimide adduct upon mechanical elongation for models of AM–FM(endo) and AM–FM(exo), each occurring with an Fmax value of 4.4 nN. Calculations were performed at the B3LYP/6-31G* level of theory. 39 Figure 2.S3. GPC traces (RI response) normalized by peak height for pristine polymers AM– FM(endo) (Mn = 106 kDa, Đ = 1.20) and AM–FM(exo) (Mn = 103, Đ = 1.19). 40 Figure 2.S4. 1H NMR spectra (400 MHz, CDCl3) confirming the successful incorporation of mechanophore initiator 2.1(endo) into PMA polymer AM–FM(endo) (Mn = 106 kDa, Đ = 1.20). 41 Figure 2.S5. 1H NMR analysis confirming the successful incorporation of mechanophore initiator 2.1(exo) into PMA polymer AM–FM(exo) (Mn = 103 kDa, Đ = 1.19). 42 Figure 2. S6. Control sonication experiments performed on a low molecular weight polymer (Mn = 18 kDa; Đ = 1.10) containing a chain-centered AM–FM(endo) bis-adduct mechanophore. (a) PL measurements (λex = 365 nm) after 40 min ultrasonication (2 mg/mL polymer) demonstrating negligible reactivity of the AM adduct compared to AM–FM(endo) (Mn = 106 kDa; Đ = 1.20). Calculated yield of anthracene product is given for each reaction based on the calibration curve in Figure S10. The purple dashed line represents the maximum PL signal achieved for AM–FM(endo) after 220 min sonication. Note: the molar concentration of mechanophore in the experiment using the low molecular weight control polymer is ~6× higher. (b) Partial 1H NMR (400 MHz, CDCl3) spectra demonstrating negligible reactivity of the FM adduct in the low molecular weight control polymer after 40 min ultrasonication compared to AM–FM(endo) (Mn = 106 kDa; Đ = 1.20). (c) GPC traces (RI response) normalized by peak area for the low molecular weight control polymer before and after ultrasonication for 40 min demonstrating negligible chain scission. 2.5.3. DFT Calculations (CoGEF) CoGEF calculations were performed using Spartan ’18 Parallel Suite according to previously reported methods.56,57 A truncated model of the mechanophore with terminal acetoxy groups was used. Ground state energies were calculated using DFT at the B3LYP/6- 43 31G* level of theory. Starting from the equilibrium geometry of the unconstrained molecule (energy = 0 kJ/mol), the distance between the terminal methyl groups of the structure was increased by 0.05 Å increments and the energy was minimized at each step. The maximum force associated with the mechanochemical retro-[4 + 2] cycloaddition reaction was calculated from the slope of the curve immediately prior to bond cleavage. There are four possible diastereomers for each bis-adduct with either endo or exo stereochemistry of the FM subunit (Charts 2.S3 and 2.S4). CoGEF calculations performed on each set of diastereomers predict the same overall mechanochemical transformation (activation of the FM subunit) with identical values of Fmax (Figure 2.S7). Chart 2.S3. Possible truncated diastereomer structures of AM–FM(endo). Chart 2.S4. Possible truncated diastereomer structures of AM–FM(exo). 44 Figure 2.S7: DFT calculations using the constrained geometries simulate external force (CoGEF) method predict the retro-Diels–Alder reaction of the furan-maleimide adduct upon mechanical elongation of (a) all four diastereomeric models of AM–FM(endo) (Fmax = 4.0 nN for all isomers), and (b) all four diastereomeric models of AM–FM(exo) (Fmax = 4.2 nN for all isomers). Structures of the diastereomers are shown above in Charts S3 and S4. Calculations were performed at the B3LYP/6-31G* level of theory. Figure 2.S8. Schematic representation of d, d0, D, and D0 used for analysis of mechanochemical coupling. Bond lengths were extracted from CoGEF calculations. 45 Mechanochemical coupling was analyzed according to the methodology by De Bo and coworkers.58,59 As depicted in Figure 2.S8, the normalized bond length (d/d0) of the primary scissile Diels–Alder adduct bond predicted to lengthen selectively in the CoGEF calculation was plotted against the overall end-to-end displacement of the mechanophore model (D/D0) for AM–FM(endo), AM–FM(exo), FM(endo), FM(exo), and AM. Only the enthalpic stretching portion of the calculation was considered, starting where d ³ 0.001 Å at each incremental step, until the step immediately prior to bond rupture. Bond length data for FM(endo), FM(exo), and AM were determined from the literature.57 Diastereomer 1 was used as the model for both AM–FM(endo) and AM–FM(exo). For the strain maps shown in Figure 2.4 in the main text, bond lengths in the CoGEF model immediately prior to rupture were represented by percent bond elongation (100 × d/d0) relative to equilibrium and mapped for all bonds in the AM–FM(endo) and AM–FM(exo) models. Strain maps using these data were generated using PyMOL.60 The molecular model immediately prior to bond rupture derived from the CoGEF calculation was exported from Spartan ’18 Parallel Suite a .pdb file and imported into the PyMOL interface. Each bond in the molecule was selected individually and color coded based on the bond strain. 46 2.5.4. Synthetic Details Scheme 2.S1. Synthesis of Anthracene–Maleimide and Furan–Maleimide Bis-Adduct Mechanophores AM–FM(endo) and AM–FM(exo). 2,2'-(propane-1,3-diyl)bis(3a,4,7,7a-tetrahydro-1H-4,7-epoxyisoindole-1,3(2H)-dione) (2.3). Compound 2.3 was synthesized using a method adapted from Sun and coworkers.61 A flame dried 2-neck round bottom flask equipped with a stir bar and a condenser was charged with 2.2 (6.0 g, 35.97 mmol), propane-1,3-diamine (1.38 mL, 16.35 mmol), and anhydrous methanol (200 mL). The solution was heated to reflux followed by the dropwise addition of triethylamine (4.59 mL, 32.7 mmol). After refluxing for 20 h, the reaction mixture was cooled by placing it in a freezer. The solid precipitate was collected by vacuum filtration and washed with cold methanol to afford 2.3 as a white solid (2.38 g, 39% yield). Rf = 0.57 (10:90 methanol/DCM). 1 H NMR (400 MHz, CDCl3) δ: 6.50 (s, 4H), 5.26 (s, 4H), 3.47 (t, J = 7.1 Hz, 4H), 2.84 (s, 4H), 1.91 (p, J = 7.2 Hz, 2H) ppm. 47 13 1 C{ H} NMR (101 MHz, CDCl3) δ: 176.2, 136.6, 81.0, 47.5, 36.3, 25.4 ppm. HRMS (ESI, m/z): calcd for [C19H18N2NaO6]+ (M+Na)+, 393.1063; found, 393.1062. 4-(hydroxymethyl)-2-(3-(9-(hydroxymethyl)-12,14-dioxo-9,10-dihydro-9,10[3,4]epipyrroloanthracen-13-yl)propyl)-3a,4,7,7a-tetrahydro-1H-4,7-epoxyisoindole1,3(2H)-dione (2.4). A round bottom flask equipped with a stir bar and a condenser was charged with 2.3 (1.96 g, 5.3 mmol), 9-hydroxymethylanthracene (367 mg, 1.77 mmol), and toluene (15 mL). The mixture was heated to reflux and stirred for 20 h before being concentrated under reduced pressure. The resulting solid was dissolved in ethyl acetate (10 mL) followed by the addition of 2-(hydroxymethyl)furan (1.042 g, 10.62 mmol). The reaction was stirred at room temperature for 72 h before being concentrated under reduced pressure onto celite. The crude mixture was purified by column chromatography on silica (50–100% EtOAc/hexanes, then 0–10% MeOH/EtOAc) to afford 2.4 as a mixture of stereoisomers (392 mg, 39% yield). This isomeric mixture was used directly in the next step. HRMS (ESI, m/z): calcd for [C31H29N2O7]+ (M+H)+, 541.1897; found, 541.1972. (13-(3-((3aR,4R,7S,7aS)-4-(((2-bromo-2-methylpropanoyl)oxy)methyl)-1,3-dioxo1,3,3a,4,7,7a-hexahydro-2H-4,7-epoxyisoindol-2-yl)propyl)-12,14-dioxo-9,10[3,4]epipyrroloanthracen-9(10H)-yl)methyl 2-bromo-2-methylpropanoate (2.1(endo)) and (13-(3-((3aS,4R,7S,7aR)-4-(((2-bromo-2-methylpropanoyl)oxy)methyl)-1,3-dioxo1,3,3a,4,7,7a-hexahydro-2H-4,7-epoxyisoindol-2-yl)propyl)-12,14-dioxo-9,10[3,4]epipyrroloanthracen-9(10H)-yl)methyl 2-bromo-2-methylpropanoate (2.1(exo)). A flamed-dried round bottom flask equipped with a stir bar was charged with isomeric mixture 2.4 (336 mg, 0.621 mmol), DCM (10 mL), and triethylamine (0.606 mL, 4.35 mmol). The reaction was cooled in an ice bath followed by the addition of α-bromoisobutyryl bromide (0.385 mL, 3.11 mmol). The reaction was stirred at room temperature for 5 h and then diluted with ethyl acetate (50 mL). The organic layer was washed consecutively with 10% NH4Cl (50 mL), H2O (3 x 50 mL), and brine (50 mL). The organic layer was then dried over Na2SO4, filtered, and concentrated under reduced pressure. The crude mixture was purified by column 48 chromatography on silica (10–80% EtOAc/hexanes) to afford 2.1(endo) as a white solid (156 mg, 30% yield) and 2.1(exo) as a white solid (187 mg, 38% yield). 2.1(endo): Rf = 0.38 (50/50 EtOAc/hexanes). 1 H NMR (400 MHz, CDCl3) δ: 7.44 – 7.33 (m, 2H), 7.33 – 7.27 (m, 1H), 7.24 – 7.13 (m, 5H), 6.52 (ddd, J = 5.3, 3.3, 1.7 Hz, 1H), 6.38 (dd, J = 5.8, 1.2 Hz, 1H), 5.70 – 5.53 (m, 1H), 5.53 – 5.36 (m, 1H), 5.30 (dd, J = 5.5, 1.7 Hz, 1H), 4.93 (d, J = 12.6 Hz, 1H), 4.81 – 4.73 (m, 1H), 4.70 (dd, J = 12.5, 1.2 Hz, 1H), 3.61 (ddd, J = 7.7, 5.5, 1.5 Hz, 1H), 3.39 (dd, J = 7.7, 1.4 Hz, 1H), 3.34 – 3.19 (m, 2H), 3.08 (t, J = 6.3 Hz, 2H), 2.97 (dd, J = 7.3, 1.5 Hz, 2H), 1.98 (m, 12H), 0.98 – 0.79 (m, 2H) ppm. 13 C{1H} NMR (101 MHz, CDCl3) δ: 176.3, 175.2, 174.5, 174.3, 171.4, 171.3, 141.8, 141.0, 138.8, 138.1, 136.1, 134.6, 127.4, 127.2, 127.1, 126.9, 125.5, 124.3, 123.3, 122.3, 89.8, 79.7, 63.3, 63.2, 55.55, 55.53, 55.52, 48.0, 47.8, 47.7, 46.8, 45.8, 36.2, 36.0, 30.91, 30.88, 30.84, 30.81, 25.3 ppm. HRMS (ESI, m/z): calcd for [C39H42Br2N3O9]+ (M+NH4)+, 854.1282; found, 854.1275. 2.1(exo): Rf = 0.24 (50/50 EtOAc/hexanes). 1 H NMR (400 MHz, CDCl3) δ: 7.43 – 7.33 (m, 2H), 7.33 – 7.27 (m, 1H), 7.24 – 7.12 (m, 5H), 6.55 (dt, J = 5.7, 1.7 Hz, 1H), 6.47 (dd, J = 5.7, 1.7 Hz, 1H), 5.61 (d, J = 11.0 Hz, 1H), 5.53 – 5.36 (m, 1H), 5.25 (t, J = 2.0 Hz, 1H), 4.95 (dd, J = 12.6, 1.4 Hz, 1H), 4.81 – 4.70 (m, 1H), 4.56 (dd, J = 12.6, 2.4 Hz, 1H), 3.34 – 3.23 (m, 2H), 3.18 (q, J = 6.5 Hz, 2H), 3.04 (t, J = 6.5 Hz, 2H), 2.97 (dd, J = 6.5, 3.3 Hz, 1H), 2.91 (dd, J = 6.4, 3.9 Hz, 1H), 2.05 – 1.89 (m, 12H), 1.17 – 0.90 (m, 2H) ppm. 13 C{1H} NMR (101 MHz, CDCl3) δ: 176.3, 175.6, 175.2, 174.1, 171.4, 171.2, 141.8, 141.0, 138.7, 138.0, 137.5, 137.1, 127.4, 127.3, 127.0, 126.8, 125.6, 125.5, 124.3, 123.3, 89.4, 81.2, 63.3, 62.7, 55.6, 55.5, 49.94, 49.92, 48.4, 48.0, 47.7, 45.8, 36.4, 35.7, 30.89, 30.86, 30.84, 30.83, 25.0 ppm. HRMS (ESI, m/z): calcd for [C39H42Br2N3O9]+ (M+NH4)+, 854.1282; found, 854.1277. 49 Scheme 2.S2. Synthesis of Poly(Methyl Acrylate) (PMA) Containing Chain-Centered Bis-Adduct Mechanophore FM–AM(endo). Scheme 2.S3. Synthesis of Mechanophore FM–AM(exo). PMA Containing Chain-Centered Bis-Adduct Representative procedure for the synthesis of PMA polymers containing a mechanophore near the chain midpoint. PMA polymers were synthesized by controlled radical polymerization following the procedure by Nguyen et al.62 A 25 mL Schlenk flask equipped with a stir bar was charged with initiator 2.1(endo) (24.1 mg, 0.0287 mmol), DMSO (4.5 mL), methyl acrylate (4.5 mL), and freshly cut copper wire (1.5 cm length, 20 gauge). The flask was sealed, the solution was deoxygenated with three freeze-pump-thaw cycles, and then allowed to warm to rt and backfilled with nitrogen. Me6TREN (31.5 μL, 0.115 mmol) was added via microsyringe to initiate the polymerization. After stirring at rt 50 for 2 h, the flask was opened to air and the solution was diluted with dichloromethane. The polymer solution was precipitated into cold methanol (3x) and the isolated polymer was dried under vacuum to provide 2.15 g (50%) of AM–FM(endo). Mn = 106 kDa, Đ = 1.20. Table S1. Summary of Mn and Đ data for AM–FM(endo), AM–FM(exo) and PMA-Furan. Mn (kDa) Đ 106 1.20 18 1.10 AM–FM(exo) 103 1.19 PMA-Furan 57 1.07 AM–FM(endo) 2.5.5. Description of Sonication Experiments General procedure for ultrasonication experiments. An oven-dried sonication vessel was fitted with rubber septa, placed onto the sonication probe, and allowed to cool under a stream of dry nitrogen. The vessel was charged with a dilute solution of the polymer (2.0 mg/mL, 20 mL) and submerged in an ice bath. Sonications of AM–FM(endo) and AM–FM(exo) for the purpose of monitoring anthracene–maleimide activation were performed in MeCN with 30 mM BHT.63 The solution was sparged continuously with nitrogen beginning 30 min prior to sonication and for the duration of the sonication experiment. Pulsed ultrasound (2 mg/mL in MeCN, 1 s on/ 2 s off, 6–9 °C, 20 % amplitude, 20 kHz, 8.2 W/cm2) was then applied to the system. Aliquots (1.0 mL) were removed at specified time points (sonication “on” time) and filtered through a 0.45 µm PTFE syringe filter prior to analysis by GPC and fluorescence spectroscopy. Sonications were performed for 220 min in total. A representative example of sonication time-dependent GPC data (RI response) from the sonication of AM–FM(endo) and AM–FM(exo) are shown in Figure 2.S9. Tabulated data are provided in Tables 2.S2 and 2.S3. Ultrasonic intensity was calibrated using the method described by Berkowski et al.64 Analysis of sonicated polymer samples by 1H NMR spectroscopy. Sonications of AM– FM(endo) and AM–FM(exo) for the purpose of quantifying furan–maleimide activation were performed in MeCN (without BHT) according to the same procedure described above. 51 Samples were prepared by subjecting 20 mL of the sonicated polymer solution to the specified amount of sonication time. The solution was then filtered through a 0.45 µm PTFE syringe filter and concentrated under reduced pressure. The entire sample was then dissolved in CDCl3 (700 µL) and analyzed by 1H NMR spectroscopy to determine the extent of furan– maleimide activation. Analysis of sonicated polymer samples by fluorescence spectroscopy. As described above, aliquots from the sonication experiment were added to a quartz microcuvette and analyzed by fluorescence spectroscopy to determine the extent of anthracene–maleimide activation. Emission spectra for AM–FM(endo) and AM–FM(exo) were recorded at 375– 500 nm using an excitation wavelength of 365 nm. Excitation and emission slit widths were 5 nm. Figure 2.S9. Time-dependent GPC traces (RI response) normalized by peak area for the sonication of (a) AM–FM(endo) (Mn = 106 kDa, Đ = 1.20) and (b) AM–FM(exo) (Mn = 103 kDa, Đ = 1.19), illustrating decreasing molecular weight of the polymer sample upon ultrasonication. 52 Table 2.S2. Determined Mn for AM–FM(endo) (Mn = 106 kg/mol; Đ = 1.20) upon ultrasonication in MeCN. Trial 1 Trial 2 Sonication time (min) Mn Mn 0 103 100 10 79.1 78.9 20 67.4 65.8 30 58 56.6 40 56.1 54.6 60 49 47.3 80 44.4 44.3 120 39.3 41.8 160 34.2 37.3 220 31 29.6 Table 2.S3. Determined Mn for AM–FM(exo) (Mn = 103 kg/mol; Đ = 1.19) upon ultrasonication in MeCN. Trial 1 Trial 2 Sonication time (min) Mn Mn 0 102 100.3 10 82.1 79.8 20 70.2 64.5 30 64.5 63.3 40 51.2 55.4 60 51 50.4 80 43.4 46.2 120 38.7 38.2 160 28.2 29.3 220 17.3 18.3 53 2.5.6. Determination of Mechanophore Activation Efficiency Characterization of activation efficiency for the anthracene–maleimide mechanophore. Activation efficiency was calculated according to the literature method.63 Samples of small molecule 2.S2 in MeCN at various concentrations were prepared and PL spectra were acquired to construct the calibration curve shown in Figure 2.S10. The theoretical PL intensity for each sonication experiment was calculated from this calibration curve based on the total mechanophore concentration in the solution and used as the value for 100% activation. A representative example is shown below in Figure 2.S11. Data are provided below in Tables 2.S4 and 2.S5 for each sonication experiment. Correction for background fluorescence. To control for an increase in PL intensity during ultrasonication resulting from sonication of the solvent, a blank of MeCN was sonicated according to the methods described above. The PL intensity was found to increase linearly with sonication time. Fitting this time-dependent PL response to a line provided a background correction factor that was subtracted from each PL measurement (Figure 2.S12). 54 Figure 2.S10. Construction of a calibration curve for experimental determination of the concentration of anthracene-containing polymer. (a) Photoluminescence emission spectra (λex = 365 nm), and (b) PL intensity at 413 nm for solutions of compound 2.S2 in MeCN as a function of concentration. A linear regression of the data in (b) gives the calibration function. 55 b) Sonication Activation PL (a.u.) Conc. (𝜇M) 0 97.3 -- -- 10 2925.6 0.28 1.49 20 4935.6 0.48 2.50 30 6638.4 0.64 3.38 40 7516.2 0.69 3.66 60 8994.4 0.87 4.63 80 10480 1.01 5.36 120 13291.4 1.13 5.98 160 14648.5 1.14 6.06 220 15210.8 1.15 6.07 time (min) (%) Figure 2.S11. (a) Representative PL measurements (λex = 365 nm) for AM–FM(endo) (Mn = 106 kg/mol; Đ = 1.20) upon ultrasonication. (b) Photoluminescence intensities at 413 nm, determined concentration of anthracene, and determined percent activation of AM. 56 Figure 2.S12. Construction of a calibration curve for determining background PL upon sonication of MeCN solvent. (a) Photoluminescence emission spectra (λex = 365 nm), and (b) PL intensity at 413 nm for pure MeCN as a function of sonication time. A linear regression of the data in (b) gives the calibration function used for background subtraction. 57 Table 2.S4. Determined PL response (λem = 413 nm) for AM–FM(endo) (Mn = 106 kg/mol; Đ = 1.20) upon ultrasonication in MeCN. [Mechanophore]0 = 19.1 μM. Trial 1 Trial 2 Sonication time (min) PL (a.u.) PL (a.u.) 0 97.3 139.2 10 2925.6 3202.3 20 4935.6 5300.5 30 6638.4 7152.8 40 7516.2 7821.7 60 8994.4 9961 80 10480 11621.4 120 13291.4 13274.2 160 14648.5 13880.1 220 15210.8 14555.4 Table 2.S5. Determined PL response (λem = 413 nm) for AM–FM(exo) (Mn = 103 kg/mol; Đ = 1.19) upon ultrasonication in MeCN. [Mechanophore]0 = 19.8 μM. Trial 1 Trial 2 Sonication time (min) PL (a.u.) PL (a.u.) 0 80.4 137.9 10 4994.8 4793.7 20 8187.2 8060.9 30 11402.5 10810.8 40 13686.5 12726.9 60 16692.3 15786.5 80 18633.1 17741.3 120 20511 19627.8 160 21703.3 20387.2 220 22080.9 21221.1 Characterization of activation efficiency for the 58 furan–maleimide mechanophore. As mentioned above, radical inhibitor BHT was excluded in sonication experiments for the purpose of determining furan–maleimide activation to prevent spectral contamination from the strong BHT signals in 1H NMR analysis. First, to confirm that the furan moiety was stable under these conditions, a polymer containing a chain-end furan moiety, PMA-Furan (Mn = 57 kg/mol; Đ = 1.07), was synthesized and sonicated according to the procedure described above in Section 2.4.5. The sonicated polymer and a sample of pristine PMA-Furan were dissolved separately in 650 μL of CDCl3, and 50 μL of a dilute solution of 1,3,5-trimethoxybenzene in CDCl3 was added to each sample as an internal standard for 1H NMR analysis. Integration of all signals for PMA-Furan are consistent before and after sonication, indicating the stability of the chain-end furan moiety under these conditions (Figure 2.S13). An additional control experiment was performed to confirm that furan–maleimide consumed in the reaction is converted to the retro-Diels–Alder products without significant side reactions or degradation under the sonication conditions. A sample of AM–FM(exo) was prepared and sonicated for 220 min in the absence of BHT and analyzed by 1H NMR spectroscopy. 1,3,5-trimethoxybenzene was used as an internal standard, as described above. The integration of signals corresponding to protons a and b at 0 min is consistent with the total integration of signals corresponding to protons Ha, Hb, Hc, and Hd after 220 min of sonication (Figure 2.S14). For AM–FM(endo), integration of signals corresponding to the residual FM adduct (6.34 ppm, Ha + Hc, 2H) and the furan mechanochemical reaction product (6.37 ppm, Hb, 1H) were used to calculate mechanophore conversion. For AM–FM(exo), integration of signals corresponding to residual mechanophore (6.52 ppm, Ha, 1H and 6.46 ppm, Hb, 1H) and the furan mechanochemical reaction product (6.37 ppm, Hd, 1H and 6.34 ppm, Hc, 1H) were used to calculate mechanophore conversion. Assignments are based on the literature and illustrated in Scheme 2.S4.53 Representative examples are illustrated in Figures 2.S15 and 2.S16. 59 Figure 2.S13. Partial 1H NMR spectra (400 MHz, CDCl3) of PMA-Furan (Mn = 57 kg/mol; Đ = 1.07) before and after 220 minutes of sonication in MeCN in the absence of BHT confirming stability of the chain-end furan moiety. The peak at 6.1 ppm corresponds to 1,3,5trimethoxybenzene internal standard. 60 Figure 2.S14. Partial 1H NMR spectra (400 MHz, CDCl3) of AM–FM(exo) (Mn = 103 kg/mol; Đ = 1.19) before and after 220 min of sonication in MeCN in the absence of BHT to prevent spectral contamination as described above. The peak at 6.1 ppm corresponds to 1,3,5trimethoxybenzene internal standard. 61 Scheme 2.S4. Protons Selected in the 1H NMR Anaysis for the Characterization of Furan–Maleimide Activation Sonication time (min) 0 10 20 30 40 60 80 120 160 220 Hb Hd 1 0.70 0.53 0.56 0.49 0.21 0.32 0.04 0.15 -- 0 0.30 0.44 0.44 0.51 0.65 0.79 0.84 0.84 0.83 Figure 2.S15. (a) Partial 1H NMR (400 MHz, CDCl3) spectra of AM–FM(endo) (Mn = 106 kg/mol; Đ = 1.20) indicating the proton signals selected for integration analysis, and (b) 1H NMR signal integrations used to calculate the extent of furan–maleimide activation. 62 Sonication time (min) 0 10 20 30 40 60 80 120 160 220 Ha + Hb Hc + Hd 1 0.88 0.72 0.63 0.59 0.63 0.59 0.35 0.37 0.32 0 0.12 0.28 0.37 0.41 0.37 0.41 0.65 0.63 0.68 Figure 2.S16: (a) Partial 1H NMR (400 MHz, CDCl3) spectra of AM–FM(exo) (Mn = 103 kg/mol; Đ = 1.19) indicating the proton signals selected for integration analysis, and (b) 1H NMR signal integrations used to calculate the extent of furan–maleimide activation. Table 2.S6. Determined 1H NMR signal integrations for AM–FM(endo) (Mn = 106 kg/mol; Đ = 1.20) upon ultrasonication in MeCN. Trial 1 Trial 2 Sonication time (min) Hb Hd Hb Hd 0 1 0 1 0 10 0.70 0.30 0.69 0.30 20 0.53 0.44 0.63 0.39 30 0.56 0.44 0.51 0.51 40 0.49 0.51 0.43 0.58 60 0.21 0.65 0.36 0.65 80 0.32 0.79 0.31 0.69 120 0.04 0.84 0.26 0.77 160 0.15 0.84 0.23 0.78 220 -- 0.83 0.24 0.78 63 Table 2.S7. Determined H NMR signal integrations for AM–FM(exo) (Mn = 103 kg/mol; Đ = 1.19) upon ultrasonication in MeCN. Trial 1 Trial 2 Sonication time (min) Ha + Hb Hc+ Hd Ha + Hb Hc+ Hd 1 0 1 0 1 0 10 0.88 0.12 0.83 0.17 20 0.72 0.28 0.73 0.27 30 0.63 0.37 0.68 0.32 40 0.59 0.41 0.64 0.36 60 0.63 0.37 0.59 0.41 80 0.59 0.41 0.48 0.52 120 0.35 0.65 0.41 0.59 160 0.37 0.63 0.40 0.60 220 0.32 0.68 0.34 0.66 Calculation of percent activation for AM–FM(endo) and AM–FM(exo). Determined percent activation values for AM and FM adducts were averaged from two replicate trials and fitted to eq 2.1 following a previously described procedure.63 The fitdetermined plateau value (A) was used as the maximum percent activation. Representative examples are illustrated in Figures 2.S17–2.S20 and the data are recorded in Table 2.S8. 𝐼 = 𝐴(1 − 𝑒 !"# ) (2.1) 64 Figure 2.S17. Activation of the furan–maleimide subunit in AM–FM(endo) (Mn = 106 kg/mol; Đ = 1.20), determined by 1H NMR spectroscopy as a function of ultrasonication time. Fitting the data to eq 2.1 provides a plateau percent activation, A, of 80.2%. Data points are averages from two experiments with error bars denoting the range of the two measurements. 65 Figure 2.S18. Activation of the furan–maleimide subunit in AM–FM(exo) (Mn = 103 kg/mol; Đ = 1.19), determined by 1H NMR spectroscopy as a function of ultrasonication time. Fitting the data to eq 2.1 provides a plateau percent activation, A, of 65.1%. Data points are averages from two experiments with error bars denoting the range of the two measurements. 66 Figure 2.S19. Activation of the anthracene–maleimide subunit in AM–FM(endo) (Mn = 106 kg/mol; Đ = 1.20), determined by PL spectroscopy as a function of ultrasonication time. Fitting the data to eq 2.1 provides a plateau percent activation, A, of 6.2%. Data points are averages from two experiments with error bars denoting the range of the two measurements. 67 Figure 2.S20. Activation of the anthracene–maleimide subunit in AM–FM(exo) (Mn = 103 kg/mol; Đ = 1.19), determined by PL spectroscopy as a function of ultrasonication time. Fitting the data to eq 2.1 provides a plateau percent activation, A, of 9.3%. Data points are averages from two experiments with error bars denoting the range of the two measurements. Table 2.S8. Determined percent activation of the AM and FM adduct for AM–FM(endo) (Mn = 106 kDa; Đ = 1.20) upon ultrasonication in MeCN. 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Macromolecules 2005, 38, 8975–8978. https://doi.org/10.1021/ma051394n. 79 Chapter 3 REMOTE CONTROL OF MECHANOCHEMICAL REACTIONS UNDER PHYSIOLOGICAL CONDITIONS USING BIOCOMPATIBLE FOCUSED ULTRASOUND This chapter has been reprinted from Yao, Y.; McFadden, M.E.; Luo, S.M.; Barber, R.W.; Kang, E.; Bar-Zion, A.; Smith, C.A.B.; Jin, Z.; Legendre, M.; Ling, B.; Malounda, D.; Torres, A.; Hamza, T.; Edwards, C.E.R.; Shapiro, M.G.; and Robb, M.J.; Proc. Natl. Acad. Sci. U.S.A. 2023, 120, e2309822120 80 Abstract External control of chemical reactions in biological settings with spatial and temporal precision is a grand challenge for noninvasive diagnostic and therapeutic applications. While light is a conventional stimulus for remote chemical activation, its penetration is severely attenuated in tissues, which limits biological applicability. On the other hand, ultrasound is a biocompatible remote energy source that is highly penetrant and offers a wide range of functional tunability. Coupling ultrasound to the activation of specific chemical reactions under physiological conditions, however, remains a challenge. Here, we describe a synergistic platform that couples the selective mechanochemical activation of mechanophore-functionalized polymers with biocompatible focused ultrasound by leveraging pressure-sensitive gas vesicles as acousto-mechanical transducers. The power of this approach is illustrated through the mechanically triggered release of covalently bound fluorogenic and therapeutic cargo molecules from polymers containing a masked 2furylcarbinol mechanophore. Molecular release occurs selectively in the presence of gas vesicles upon exposure to focused ultrasound under physiological conditions. These results showcase the viability of this system for enabling remote control of specific mechanochemical reactions with spatiotemporal precision in biologically relevant settings and demonstrate the translational potential of polymer mechanochemistry. 81 3.1 Introduction External control of specific chemical reactions with spatial and temporal precision is a grand challenge for biomedical applications. For example, the targeted delivery of therapeutic compounds to precise locations in the body maximizes accumulation of drugs at the sites of disease and reduces undesirable side effects to healthy tissues.1–4 Likewise, spatially triggered reactions can establish patterned engineered tissues and functional living materials.5–8 Among various ways to drive chemical transformations remotely, light is a prototypical external stimulus for achieving high spatiotemporal resolution,9,10 but its limited tissue penetration depth hampers biological utility.11 Research in the nascent field of polymer mechanochemistry has focused on the development of force sensitive molecules termed mechanophores,12 including those that release covalently bound small molecules upon mechanochemical activation.13,14 Mechanical force, which can also be applied with spatial and temporal precision, is transduced to the mechanophore through covalently linked polymer chains to elicit a chemoselective response. Mechanophores have been developed that release carbon monoxide,15,16 acid,17,18 catalysts,19 and a variety of other payloads.13 Research from the groups of Göstl and Herrmann20,21 and the Robb group22–24 has focused on the design of more general and modular mechanophore platforms enabling the mechanically triggered release of functionally diverse molecules that are especially promising for biomedical applications.13 Ultrasonication is an effective technique for achieving mechanophore activation in the laboratory. High intensity low frequency sonication (typically 20 kHz) using an immersion probe is commonly employed to exert mechanical forces on polymers in solution via acoustic cavitation, a process in which the nucleation, growth, and collapse of gas bubbles produces solvodynamic shear and results in the rapid elongation of polymer chains (Figure 3.1a).12,25 However, the cavitation of dissolved gases under these conditions is highly destructive to tissues, making it incompatible with most biological applications.13,26 Recent studies have begun to explore the use of clinically relevant focused ultrasound (FUS) operating at higher frequencies for mechanophore activation in polymer networks27 and hydrogels.28 Focused ultrasound is an ideal external stimulus as it can be applied with submillimeter spatial resolution and is highly penetrant to biological tissues.29 Nevertheless, 82 while these earlier studies provide a promising direction, the high acoustic intensities employed cause unsafe heating, which represents a major obstacle for applications.27,28,30 biomedical Moreover, coupling biocompatible ultrasound with Figure 3.1. Activation of mechanochemical reactions under physiological conditions using biocompatible focused ultrasound enabled by gas vesicles as acousto-mechanical transducers. a) Conventional methods of achieving ultrasound-induced mechanochemical activation rely on the cavitation of dissolved gases in solution using hazardous sonication at low frequency and high acoustic intensity. b) Using gas vesicles, which function as acousto-mechanical transducers, mechanochemical activation of mechanophore-functionalized polymers is achieved using biocompatible focused ultrasound under physiological conditions, enabling the controlled release of molecular cargo. the remote activation of mechanochemical reactions in solution an remains unsolved challenge. New methodologies are therefore required to realize the translational potential of polymer mechanochemistry.13 Gas vesicles (GVs) are a unique class of genetically encodable, air-filled protein nanostructures that have been developed as novel contrast agents for non-invasive biomedical imaging, including high-frequency diagnostic ultrasound.31–34 GVs typically have diameters of 45– 250 nm and lengths of 100–600 nm and comprise amphiphilic protein shells that are gaspermeable but exclude liquid water from their hydrophobic interior.35 Recently, the Shapiro group discovered that GVs act as seeds for bubble formation and cavitation upon exposure to biocompatible focused ultrasound in vivo.36 Under sustained ultrasound pulses with relatively low pressure levels, larger gas bubbles liberated from the rupture of GVs undergo rapid growth followed by intense collapse in an inertial cavitation process with pronounced mechanical effects. With the ultrasound parameters used, these effects take place only in the vicinity of GVs and are benign to other tissues.36 Based on these results, we hypothesized that GVs could function as acousto-mechanical transducers to effectively couple focused ultrasound operating under physiological conditions with the mechanochemical activation of mechanophore-containing polymers (Figure 3.1b). 83 Herein, we describe an approach to achieve remote control of mechanochemical reactions in aqueous environments using biocompatible focused ultrasound. Using GVs as acousto-mechanical transducers, focused ultrasound selectively triggers the release of fluorogenic and therapeutic cargo molecules from mechanophore-functionalized polymers. This methodology represents a promising step forward for bridging the gap between the technological potential of polymer mechanochemistry and translational applications in biology, medicine, and living materials. 3.2. Results and Discussion We initially set out to identify focused ultrasound parameters for safe operation under physiological conditions with a particular emphasis on maintaining pressure low-to-moderate levels and avoiding temperature increases greater than ~6 °C to stay within a biomedically relevant regime. Focused ultrasound applied at high acoustic intensities can induce coagulative necrosis in tissues resulting disruption.37 from We thermal performed Figure 3.2. Analysis of biocompatible focused ultrasound conditions for mechanochemical activation. experiments with 330 kHz focused a) Equilibrated maximum temperature increment (max. ΔT) at the focus of ultrasound in a tissue-mimicking 1% agarose ultrasound on either an 800 µL water hydrogel phantom with varying peak negative pressure and sample in a sealed duty cycle. b) Temperature profile of an 800 μL water plastic sample at the focus of ultrasound inside a 2 mL plastic microcentrifuge tube or a tissue- microcentrifuge tube or a tissue-mimicking agarose gel (phantom) with different ultrasound conditions (1.47 MPa mimicking agarose gel inside a water PNP, 330 kHz, pulsed wave, 3000 cycles each pulse, 5% duty cycle, 3.6 W/cm2 acoustic intensity, or 1.9 MPa PNP, tank, and monitored peak negative 330 kHz, continuous wave, 10 s on / 20 s off, 4 repeats, 40 2 pressure (PNP) and sample W/cm acoustic intensity). c) Reaction scheme illustrating mechanically triggered molecular release from a masked 2temperature using a hydrophone and furylcarbinol mechanophore incorporated near the center of a polymer. 84 internal thermocouple, respectively (Figure 3.2a). An upper limit for PNP of 1.47 MPa was established with a 4.5% duty cycle (3000 cycles per pulse with 5 Hz pulse repetition frequency), which resulted in a maximum temperature increase of only 3.6 °C in the agarose phantom (7.3 °C inside the plastic tube) that equilibrated in < 2 min (Figure 3.2b). This biocompatible upper limit results in a spatial peak-temporal average acoustic intensity (Ispta) of 3.6 W/cm2 that is within the safety limits established for the use of ultrasound in therapeutic applications.38 Similar ultrasound parameters have been successfully applied for therapeutic purposes to delicate tissues like the brain in both live animal studies and human clinical trials.39,40 We note that using similar focused ultrasound parameters as those previously employed for mechanophore activation,27,28 at both 330 and 916 kHz, resulted in an unsafe temperature increase exceeding 27 °C above baseline due to the significantly higher acoustic intensity (Figure 3.2b and Section 3.6, Figure 2.S1). Having identified biocompatible focused ultrasound parameters, we next sought to determine if mechanophore activation could be achieved under physiologically relevant conditions. We employed a masked 2-furylcarbinol mechanophore developed in the Robb group for mechanically triggered molecular release (Figure 3.2c).22,23 Briefly, mechanochemical activation of the furan‒maleimide Diels‒Alder adduct reveals a latent 2furylcarbinol derivative that spontaneously decomposes to release a covalently bound cargo molecule. Release occurs efficiently in polar protic environments and the molecular design is amenable to payloads conjugated to the mechanophore via a range of common functional groups including alcohols and amines.23 The incorporation of cargo molecules through covalent linkages increases resistance to nonspecific payload release in the absence of a specific triggering event compared to non-covalent encapsulation strategies,41 while the ability to precisely modulate the reactivity of mechanophores through structural modification using the tools of organic chemistry affords excellent selectivity and control over molecular release.13 We further identified the water soluble, nontoxic polymer poly(2(methylsulfinyl)ethyl acrylate) (PMSEA)42 that can be synthesized using controlled radical polymerization, enabling straightforward incorporation of the mechanophore near the middle of the polymer chain where force is maximized during solvodynamic extension.12 We synthesized PMSEA model polymers without incorporating mechanophores and measured 85 the refractive index increment (dn/dc) in MilliQ water as 0.153 ± 0.003 mL/g using gel permeation chromatography (GPC) (Table 3.S1, Figure 3.S9). We note that alternative polymer conjugation strategies are also readily available, such as “click” coupling,43,44 to further broaden the scope of accessible materials. We first investigated a PMSEA polymer containing a chaincentered mechanophore loaded with a fluorogenic aminocoumarin payload Figure 3.3. Mechanochemically-mediated release of an aminocoumarin small molecule payload triggered using biocompatible focused ultrasound. a) Biocompatible FUS (330 kHz, 1.47 MPa PNP, 3.6 W/cm2, 3000 cycles, 5% duty cycle, 10 min) triggers the release of aminocoumarin small molecule from PMSEA-CoumNH2 (2 mg/mL in water) selectively in the presence of GVs (1.4 nM) resulting in a fluorogenic response. b,c) Release of aminocoumarin characterized by photoluminescence (PL) spectroscopy. Error bars represent standard deviation from three replicate experiments for (-GV -FUS) and (-GV +FUS), and from four replicate experiments for (+GV +FUS). Asterisks represent statistical significance by ordinary one-way AVOVA with multiple comparison (****P < 0.0001; ***P < 0.001; **P < 0.01; ns, not significant). (PMSEA-CoumNH2, Mn = 260 kg/mol; Ð = 1.47) (Figure 3.3a, see the SI for details).23 Release of the aminocoumarin small molecule results in a pronounced increase in fluorescence that is easily detected and quantified spectroscopically. Exposing a dilute solution of the polymer (2 mg/mL) in water to 330 kHz focused ultrasound under the conditions identified above in the presence of GVs (1.4 nM) results in a strong fluorogenic response indicating the successful release of aminocoumarin (Figure 3.3a,b). Approximately 15% release was observed after 10 min of focused ultrasound exposure (Figure 3.3c). In aqueous solution, the rate of cargo release after mechanochemical retro-Diels–Alder reaction was determined to be instantaneous, consistent with the highly polar protic environment (Figure 86 3.S1). Given their relative concentrations, ~800 equivalents of PMSEA-CoumNH2 were activated per GV within this timeframe. Exposure to sonication conditions did not lead to degradation of the aminocoumarin cargo (Figure 3.S2). Extended exposure to focused ultrasound resulted in additional aminocoumarin release (Section 3.6, Figure 3.S2). Importantly, a fluorogenic response was not observed in the absence of GVs under the same conditions, confirming the GVs function as essential acousto-mechanical transducers, enabling mechanophore activation and selective cargo release under biocompatible conditions. Additional control experiments were performed on an analogous polymer with the mechanophore located at the chain end, which is not subjected to mechanical force.12 Minimal aminocoumarin release was observed under otherwise identical experimental conditions, confirming the mechanochemical origin of molecular release from PMSEACoumNH2 upon exposure to focused ultrasound in the presence of GVs (Figure 3.3b,c and Section 3.6, Figure 3.S3). Next, we set out to investigate the mechanically triggered release of a small molecule therapeutic to demonstrate the capability of this system for future drug delivery applications. We selected camptothecin as a model chemotherapeutic anticancer agent.45 Following a similar procedure as above, camptothecin was conjugated to the mechanophore bis-initiator through a carbonate linkage followed by polymerization to afford PMSEA-CPT (Mn = 319 kg/mol; Ð = 1.47) (Figure 3.4a, see the Section 3.6 for details). Preliminary experiments performed on an isolated small molecule furfuryl carbonate model compound resembling the intermediate that is unmasked upon mechanophore activation confirmed that camptothecin is successfully released in aqueous media, as evidenced by 1H NMR spectroscopy and highperformance liquid chromatography (HPLC) measurements (Section 3.6, Figure 3.S4 and 3.S5). Importantly, PMSEA-CPT is stable in aqueous solution, with negligible release of camptothecin detected over a period of 2 months under ambient conditions (Section 3.6, Figure 3.S6). The triggered release of camptothecin from PMSEA-CPT was then investigated using biocompatible focused ultrasound in the presence of GVs under the physiologically relevant conditions established above (Figure 3.4a). Approximately 8% release of camptothecin was achieved after 10 min exposure to focused ultrasound in the presence of 87 Figure 3.4. Mechanochemically triggered release of camptothecin using biocompatible focused ultrasound. a) Biocompatible FUS (330 kHz, 1.47 MPa PNP, 3.6 W/cm2, 3000 cycles, 5% duty cycle, 10 min) triggers the release of the small molecule topoisomerase inhibitor camptothecin (CPT) from PMSEA-CPT (2 mg/mL in water) selectively in the presence of GVs (1.4 nM). b) Characterization of CPT release using HPLC. Release of CPT occurs selectively upon exposure to FUS only in presence of GVs. c) Cell viability assays (MTT assay on Raji cell line, incubated 2 days at 37 °C) illustrating a significant decrease in viability for PMSEA-CPT exposed to FUS +GV with a half-maximal effective concentration (EC50) of ~250 nM. The EC50 for pure CPT is 25 nM, consistent with the extent of CPT release from PMSEA-CPT. Cell viability is not diminished in control experiments across the concentration range studied. Error bars represent standard deviation from three measurements each on samples from three replicate experiments. GVs, as evidenced by quantitative HPLC measurements (Figure 3.4b). Similar to the results above for the release of aminocoumarin, no camptothecin release was detected upon exposure of PMSEA-CPT to focused ultrasound in the absence of GVs. To assess the feasibility of this synergistic platform for targeted chemotherapy, cytotoxicity was investigated in vitro on the viability of Raji cells, a diffuse large B-cell lymphoma model of non-Hodgkin lymphomas, using an MTT colorimetric assay.46 Cells were treated with samples of PMSEA-CPT before and after mechanochemical activation with focused ultrasound at various concentrations and incubated for 2 days at 37 °C (Figure 3.4c). Cells 88 incubated with PMSEA-CPT that was previously exposed to focused ultrasound in the presence of GVs exhibited a significant decrease in viability with a half-maximal effective concentration (EC50) of ~250 nM. This EC50 is approximately one order of magnitude higher than cells treated with isolated camptothecin small molecule, fully consistent with the extent of cargo release after 10 min of insonation characterized above. In contrast, no significant cytotoxicity was observed across all polymer concentrations in control experiments performed with PMSEA-CPT that was not exposed to focused ultrasound, with PMSEACPT that was exposed to ultrasound in the absence of GVs, or with a chain-end functional control polymer (PMSEA-Control) subjected to identical focused ultrasound conditions in the presence of GVs. 3.3 Conclusions In summary, we describe a synergistic platform that couples biocompatible focused ultrasound with the solution-phase activation of mechanochemical reactions under physiologically relevant conditions. Critical to this methodology is the use of pressuresensitive air-filled protein nanostructures called gas vesicles that function as acoustomechanical transducers, effectively converting focused ultrasound energy into a coordinated mechanical stimulus for the activation of mechanophore-containing polymers. The mechanically triggered release of molecular payloads aminocoumarin and camptothecin from polymers in aqueous media illustrates the power of this approach for noninvasive bioimaging and therapeutic applications of polymer mechanochemistry. More broadly, this study highlights a novel approach for remote control of specific chemical reactions under biomedically relevant conditions with the spatiotemporal precision and tissue penetration afforded by focused ultrasound. Future research will aim to translate these proof-of-principle results to in vivo settings for drug delivery and other biomedical technologies. One potential administration route involves systemic co-injection of the GVs and polymer, relying on these synergistic components to accumulate at sufficient concentration and leveraging the ability of GVs to mechanically activate polymers at ~800-fold lower stoichiometry. Strategies that colocalize the GVs and polymers through non-covalent interactions or even covalent anchoring could 89 further enhance the local concentration of the two components necessary to achieve mechanochemical transduction. Another approach is to confine the GVs and polymer within a semipermeable implant, enabling temporal control of molecular release using ultrasound. To go beyond the fundamental findings of this study, future research will investigate these approaches in specific in vivo applications. 3.4 Materials and Methods Preparation of PMSEA-CoumNH2 and PMSEA-CPT. The masked 2furylcarbinol mechanophore with an aminocoumarin payload was synthesized according to the literature.23 A similar procedure was used to synthesize the mechanophore incorporating camptothecin as the payload as described in the Section 3.6. The monomer 2-methylsulfinyl ethyl acrylate (MSEA) was prepared according to the literature.42 Water soluble linear PMSEA polymers incorporating the mechanophores were synthesized by controlled radical polymerization from the mechanophore initiator according to a procedure adapted from Nguyen et al.47 described in the Section 3.6. Polymers were purified by precipitation into diethyl ether three times and subsequently dialyzed against deionized water for at least five cycles prior to use. General protocols for FUS experiments. Gas vesicles (GVs) were produced and purified from cyanobacteria Anabaena flos-aquae as described previously.32,36 Polymer solutions and GV suspensions were prepared using MilliQ water and separately placed under vacuum at 0.5 atm overnight to remove transiently trapped air bubbles prior to treatment with FUS. The GV suspension and polymer solution were gently mixed in a 2 mL microcentrifuge tube to achieve the desired concentrations of each component without introducing observable air bubbles. The total sample volume was 800 μL. The microcentrifuge tube was stabilized and exposed to FUS on a custom stage that positions the center of the sample at the focus of the FUS transducer (Precision Acoustics, H115) operating at 330 kHz (fundamental frequency) or 916 kHz (third harmonic). After FUS exposure, samples were subsequently analyzed by fluorescence spectroscopy or high-performance liquid chromatography, or evaluated for cytotoxicity. Additional experimental details are provided in the Section 3.6. Cell viability assays. Raji cells were incubated in RPMI 1640 media with 10% fetal 90 bovine serum and 1% antibacterial Penicillin/Streptomycin (PenStrep) at 37 °C and 5% CO2. An MTT colorimetric assay was used to quantify cell viability.46 Experimental details are provided in the Section 3.6. 3.5 Acknowledgements The authors gratefully acknowledge financial support from Caltech through a CCE Innovation Grant (M.J.R. and M.G.S.), the Arnold and Mabel Beckman Foundation through a Beckman Young Investigator Award (M.J.R.), the David and Lucille Packard Foundation (M.G.S.), the Resnick Sustainability Institute (M.G.S.), the Institute for Collaborative Biotechnologies (W911NF-19-D-0001 to M.G.S.), and the National Institute of General Medical Sciences of the National Institutes of Health (R35GM150988 to M.J.R.). M.E.M. was supported by an NSF Graduate Research Fellowship (DGE-1745301) and a Barbara J. Burger Fellowship from Caltech. A.B.-Z. was supported by the Lester Deutsch Fellowship and the Marie Skłodowska-Curie Postdoctoral Fellowship. C.A.B.S. was supported by the Human Frontiers Science Program Cross-Disciplinary Fellowship. We thank Dr. Scott Virgil for technical assistance and the Center for Catalysis and Chemical Synthesis of the Beckman Institute at Caltech for access to equipment. We gratefully acknowledge the Alfred P. Sloan Foundation for a Sloan Research Fellowship (M.J.R.) and the Camille and Henry Dreyfus Foundation for Camille Dreyfus Teacher-Scholar Awards (M.J.R. and M.G.S.). We thank the Kornfield group at Caltech for use of their GPC. M.J.R. thanks Prof. Peter Dervan for helpful discussions. 3.6 Experimental Details 3.6.1 General Experimental Details and Methods Reagents from commercial sources were used without further purification unless otherwise stated. Dry dichloromethane was obtained from a Pure Process Technology solvent purification system. All reactions were performed under a N2 atmosphere unless specified otherwise. Column chromatography on silica gel was performed on a Biotage Isolera system using SiliCycle SiliaSep HP flash cartridges. NMR spectra were recorded using a 400 MHz Bruker Avance III HD with Prodigy 91 Cryoprobe. All H NMR spectra are reported in δ units, parts per million (ppm), and were 1 measured relative to the signals for residual chloroform (7.26 ppm) or toluene (2.08 ppm) in deuterated solvent. All 13C NMR spectra were measured in deuterated solvents and are reported in ppm relative to the signals for chloroform (77.16 ppm). Multiplicity and qualifier abbreviations are as follows: s = singlet, d = doublet, t = triplet, q = quartet, dd = doublet of doublets, dq = doublet of quartets, ABq = AB quartet, m = multiplet, br = broad. High resolution mass spectra (HRMS) were obtained via direct injection on an Agilent 1260 Infinity II Series HPLC coupled to a 6230 LC/TOF system in electrospray ionization (ESI+) mode. Polymer molar mass and dispersity was determined on a GPC system with an Agilent pump (G7110B), equipped with an autosampler (G7129A), a Wyatt DAWN 8 multi-angle laser light scattering detector (λ = 658.9nm), a Wyatt Optilab refractive index detector (RI) (λ = 658nm), and an Agilent PL Aquagel-OH MIXED-H column. Aqueous buffer was prepared containing 0.2 M NaNO3 with 200 ppm NaN3. Filtered aqueous buffer was used as the eluent at a flow rate of 0.3 mL/min at 25 °C. The data were analyzed using Wyatt Astra 7 with dn/dc = 0.153 mL/g to obtain average molar masses (Mw and Mn) for each PMSEA polymer reported. Photoluminescence spectra were recorded on a Shimadzu RF-6000 spectrofluorophotometer using a quartz microcuvette (Starna Cells 18F-Q-10-GL14-C, 10 x 2 mm). Excitation and emission slit widths were 5 nm and 3 nm, respectively. A Tecan Spark microplate reader was also used to acquire photoluminescence spectra in some experiments with 96-well microplate (Costar). Excitation and emission slit widths were both 5 nm. High-Performance Liquid Chromatography (HPLC) measurements were performed with an Agilent Eclipse Plus C18 column (959961-902) or a C8 column (993967-906) using an in-line UV-vis detector and acetonitrile/water as the eluent. Sonication experiments with an immersion probe were performed using a 500 Watt Vibra Cell 505 liquid processor (20 kHz) equipped with a 0.5-inch diameter solid probe (part #630-0217), sonochemical adapter (part #830-00014), and a Suslick reaction vessel made by the Caltech glass shop (analogous to vessel #830-00014 from Sonics and Materials). LCMS measurements were performed with an Agilent 6140 Series Quadrupole 92 LCMS Spectrometer System equipped with an Agilent Eclipse Plus C18 column using an in-line UV-vis detector and acetonitrile/water as the eluent. Compounds 3.DA-1, 3.DA-2, 3.1-CoumNH2, and 3.2-CoumNH2 were synthesized as reported previously.23 Internal standard 4-methyl-2-oxo-2H-chromen-7-yl acetate (IS) was prepared according to the literature.48 Monomer 2-methylsulfinyl ethyl acrylate (MSEA) was prepared according to the literature.42 All polymer solutions were dialyzed against deionized water for at least 5 cycles to remove any residual small molecule in the reaction mixture to prevent any aminocoumarin (CoumNH2) or camptothecin (CPT) from affecting functional assays. Gas vesicles (GVs) were produced and purified from cyanobacteria Anabaena flosaquae as described previously.32,36 GvpC was removed from the GVs by urea treatment followed by dialysis process. As a quality assurance step to confirm successful GvpC removal, pressure-dependent optical density (OD) measurements were performed for each sample at 0–7 bar to match previously reported measurements using an echoVis Vis-NIR light source coupled with an STS-VIS spectrometer (Ocean Optics) and a 176.700-QS sample chamber (Hellma Analytics). GV concentrations were measured as OD at 500 nm using a spectrophotometer (NanoDrop ND-1000, Thermo Scientific). OD500 = 1 corresponds to 114 pM of GV particles.32,36 Both polymer solution and GV suspension were placed in a vacuum environment at 0.5 atm overnight to remove transiently trapped air bubbles before FUS treatment. GV suspension and polymer solution were gently mixed to desired concentration without introducing observable air bubbles in a 2 mL microcentrifuge tube. The total volume of sample was 800 µL. The microcentrifuge tube was stabilized on a homemade holder which positions the center of the sample to be at the focus of a focused ultrasound transducer (Precision Acoustics, H115) operating at 330 kHz (fundamental frequency) or 916 kHz (third harmonic). All cell lines were ordered from American Type Culture Collection. 93 3.6.2 Supplementary Figures Chart 3.S1. Structures of all compounds used in this study. 94 330 kHz, 1.9 MPa, 40 W/cm2 40 Temperature increment (°C) 916 kHz, 1.9 MPa, 40 W/cm2 30 20 10 0 0 50 100 150 Time (s) Figure 3.S3 Temperature increase above baseline of an 800 µL water sample inside a plastic microcentrifuge tube under 330 kHz or 916 kHz FUS. The temperature probe was inserted into the tube with the probe tip positioned at the tube center. Continuous-wave FUS was applied at 1.9 MPa peak negative pressure (PNP), 10 s on / 20 s off, 4 repetitions, corresponding to a spatial peak-temporal average acoustic intensity (Ispta) of 40 W/cm2. Note that our Ispta value differs from the literature6 because we considered the time average acoustic intensity with 33% duty cycle. Figure 3.S4. Release of aminocoumarin (CoumNH2) from PMSEA-CoumNH2 (2 mg/mL, Mn = 260 kg/mol, Ð = 1.47) increases as a function of temporal exposure to biocompatible FUS (330 kHz, 3.6 W/cm2, 1.47 MPa PNP, 4.5% duty cycle) in the presence of GVs (1.4 nM). Release of CoumNH2 does not occur in the absence of GVs even after 30 min of insonation. 95 Figure 3.S5. No significant increase in PL intensity is observed from chain-end functionalized polymer Control-CoumNH2 (2 mg/mL, Mn = 244 kg/mol, Ð = 1.51) upon exposure to biocompatible FUS (330 kHz, 3.6 W/cm2, 1.47 MPa PNP, 4.5% duty cycle, 10 min) in the presence or absence of GVs (1.4 nM). 96 Figure 3.S6. Release of camptothecin (CPT) is observed following the thermally-induced retro-Diels–Alder reaction of 3.1-CPT and subsequent dilution in polar protic solvent. Comparison of partial 1H NMR spectra of (a) pristine 3.1-CPT, (b) 3.1-CPT after heating for 24 h at 100 °C, and (c) maleimide M1 indicates significant retro-Diels–Alder reaction occurs thermally. Subsequent analysis by LCMS with in-line UV-vis absorption 97 spectroscopy of (d) the NMR solution of 3.1-CPT after heating for 24 h at 100 °C diluted into 3:1 acetonitrile/methanol, and (e) pure CPT standard supports that CPT is released from thermally-activated 3.1-CPT in polar protic solvent. Good agreement with the CPT standard is observed in retention time, mass-to-charge ratio, and absorption spectrum. Figure 3.S7. Analysis of thermally activated 3.1-CPT (24 h at 100 °C in toluene-d8) immediately after dilution in either (a) aprotic acetonitrile, or (b) 3:1 acetonitrile/methanol using LCMS with in-line UV-vis absorption spectroscopy. The amount of CPT relative to remaining unreacted 3.1-CPT is significantly greater after dilution in protic solvent consistent with the expected reactivity of the furfuryl carbonate. 98 Figure 3.S8. No release of camptothecin (CPT) from PMSEA-CPT (4 mg/mL, Mn = 319 kg/mol, Ð = 1.47) is observed in aqueous solution, even after 2 months in ambient conditions. The same sample volume was used for each injection. 3.6.3 Synthetic Details ((3aR,4R,7S,7aS)-2-(2-((2-bromo-2-methylpropanoyl)oxy)ethyl)-7-(1-(((((S)-4ethyl-3,14-dioxo-3,4,12,14-tetrahydro-1H-pyrano[3',4':6,7]indolizino[1,2-b]quinolin-4yl)oxy)carbonyl)oxy)ethyl)-1,3-dioxo-6-phenoxy-1,2,3,3a,7,7a-hexahydro-4H-4,7epoxyisoindol-4-yl)methyl 2-bromo-2-methylpropanoatebromo-2-methylpropanoate (3.1-CPT). A flame-dried 25 mL round bottom flask was charged with 4dimethylaminopyridine (DMAP) (544 mg, 4.43 mmol) and camptothecin (302 mg, 0.867 mmol). The flask was evacuated and refilled with N2 three times, after which 5 mL of 99 dichloromethane was added. Phosgene (15 wt% in toluene, 700 µL, 0.981 mmol) was added to the off-yellow suspension and the reaction was allowed to stir at room temperature. After 1 h the reaction had become homogenous and dark red, and 3.DA-123 was added as a solution in 5 mL dichloromethane. The reaction continued to stir for 20 h, after which it was quenched with sat. NH4Cl (8 mL). The products were extracted into dichloromethane (3 x 5 mL) and the combined organic layers were washed with brine (5 mL), dried with sodium sulfate, and filtered. The crude mixture was purified via silica gel chromatography (0–10% methanol/dichloromethane), followed by separation of the diastereomers by reverse-phase HPLC (70% acetonitrile/water). The title product (single diastereomer) was isolated as a light-yellow solid (84 mg, 27%). TLC (5% methanol/dichloromethane): Rƒ = 0.55 1 H NMR (400 MHz, CDCl3) δ: 8.38 (s, 1 H), 8.07 (d, J = 8.5 Hz, 1H), 7.91 (d, J = 8.1 Hz, 1H), 7.84–7.76 (m, 1H), 7.68–7.59 (m, 1H), 7.36 (s, 1H), 7.25–7.18 (m, 2H), 7.13– 7.05 (m, 1H), 6.90–6.81 (m, 2H), 5.69 (d, J = 17.3 Hz, 1H), 5.49 (q, J = 6.5 Hz, 1H), 5.39 (d, J = 17.2 Hz, 1H), 5.26 (s, 2H), 4.98 (s, 1H), 4.66 (Abq, ΔνAB = 104 Hz, JAB = 12.5 Hz, 2H), 3.91–3.79 (m, 2H), 3.76–3.72 (m, 2H), 3.25–3.14 (m, 1H), 3.05–2.94 (m, 1H), 2.33 (dq, J = 14.9, 7.4 Hz, 1H), 2.18 (dq, J = 14.7 Hz, 7.5 Hz, 1H), 1.94 (s, 3H), 1.91 (s, 3H), 1.70 (s, 3H), 1.68 (s, 3H), 1.53 (d, J = 6.6 Hz, 3 H), 1.01 (t, J = 7.4 Hz, 3H). 13 C{1H} NMR (101 MHz, CDCl3) δ: 173.5, 172.6, 171.2, 171.1, 167.3, 162.2, 157.3, 154.4, 152.9, 152.4, 148.6, 146.5, 145.5, 131.4, 131.0, 130.0, 129.3, 128.7, 128.4, 128.23, 128.19, 125.9, 120.6, 119.4, 101.2, 95.8, 90.2, 88.3, 78.2, 71.8, 67.3, 63.6, 62.2, 55.6, 55.5, 51.4, 50.1, 47.8, 36.9, 32.0, 30.75, 30.71, 30.5, 30.4, 15.7, 7.7. HRMS (ESI, m/z): calcd for [C48H46Br2N3O14+] (M+H)+ 1046.1341, found 1046.1355. 100 ((3aR,4R,7S,7aS)-2-(2-((2-bromo-2-methylpropanoyl)oxy)ethyl)-7-(1-(((((S)-4ethyl-3,14-dioxo-3,4,12,14-tetrahydro-1H-pyrano[3',4':6,7]indolizino[1,2-b]quinolin-4yl)oxy)carbonyl)oxy)ethyl)-1,3-dioxo-6-phenoxy-1,2,3,3a,7,7a-hexahydro-4H-4,7epoxyisoindol-4-yl)methyl pivalate (3.2-CPT). A flame-dried 25 mL round bottom flask was charged with DMAP (336 mg, 2.75 mmol) and camptothecin (192 mg, 0.552 mmol). The flask was evacuated and refilled with N2 three times, after which 5 mL of dichloromethane was added. Phosgene (15 wt % in toluene, 430 µL, 0.602 mmol) was added to the off-yellow suspension and the reaction was allowed to stir at room temperature. After 1 h the reaction had become homogenous and dark red, and 3.DA-223 was added as a solution in 5 mL of dichloromethane. The reaction continued to stir for 18 h, after which it was quenched with sat. NH4Cl (8 mL). The products were extracted into dichloromethane (3 x 5 mL) and the combined organic layers were washed with brine (5 mL), dried with Na2SO4, and filtered. The crude mixture was purified via silica gel chromatography (0–10% methanol/ dichloromethane), followed by separation of the diastereomers by reverse-phase HPLC (70% acetonitrile/water). The title product (single diastereomer) was isolated as a light yellow solid (50 mg, 28%). TLC (5% methanol/dichloromethane): Rƒ = 0.57 1 H NMR (400 MHz, CDCl3) δ: 8.39 (s, 1H), 8.07 (d, J = 8.5 Hz, 1H), 7.92 (dd, J = 8.3, 1.3 Hz, 1H), 7.81 (ddd, J = 8.4, 6.9, 1.2 Hz, 1H), 7.69–7.62 (m, 1H), 7.35 (s, 1H), 7.25– 7.19 (m, 2H), 7.13–7.07 (m, 1H), 6.85 (dd, J = 7.6, 1.6 Hz, 2H), 5.70 (d, J = 17.2 Hz, 1H), 5.50 (q, J = 6.5 Hz, 1H), 5.41 (d, J = 17.3 Hz, 1H), 5.28 (s, 2H), 4.93 (s, 1H), 4.56 (Abq, ΔνAB = 120 Hz, JAB = 12.7 Hz, 2H), 3.90–3.79 (m, 2H), 3.73–3.63 (m, 2H), 3.26–3.13 (m, 101 1H), 3.04–2.92 (m, 1H), 2.35 (dq, J = 14.9, 7.4 Hz, 1H), 2.19 (dq, J = 14.8, 7.5 Hz), 1.71 (s, 3H), 1.69 (s, 3H), 1.54 (d, J = 6.6 Hz, 3H), 1.21 (s, 9H), 1.01 (t, J = 7.5 Hz, 3H). 13 C{1H} NMR (101 MHz, CDCl3) δ: 177.8, 173.5, 172.6, 171.2, 167.3, 162.1, 157.3, 154.5, 153.0, 152.5, 148.8, 146.6, 145.4, 131.4, 131.0, 130.1, 129.4, 128.7, 128.4, 128.24, 128.22, 125.9, 120.8, 119.4, 101.3, 95.7, 90.2, 88.6, 78.2, 71.8, 67.4, 62.3, 62.2, 55.5, 51.4, 50.1, 47.9, 39.0, 36.9, 32.1, 30.53, 30.46, 27.2, 15.7, 7.8. HRMS (ESI, m/z): calcd for [C49H49BrN3O14+] (M+H)+ 982.2392, found 982.2408. Representative procedure for the synthesis of poly(2-(methylsulfinyl)ethyl acrylate) (PMSEA) polymers. PMSEA polymers were synthesized by controlled radical polymerization following an adaptation of the procedure by Nguyen et al.47 A flame-dried Schlenk flask was charged with freshly cut 20 G copper wire (2 cm), initiator 3.1-CoumNH2 (3.5 mg, 0.0040 mmol), dimethyl sulfoxide (DMSO) (5 mL), and 2-methylsulfinyl ethyl acrylate (2.46 g, 15.2 mmol). The flask was sealed and the solution was degassed via three freeze-pump-thaw cycles, then backfilled with nitrogen and warmed to room temperature. Me6TREN (5.0 µL, 0.019 mmol) was added via microsyringe and the reaction was stirred at room temperature. Upon completion of the polymerization after 6 h, the flask was opened to atmosphere and diluted with a minimal amount of methanol. The polymer was precipitated 3x into room temperature diethyl ether, dried under vacuum, dialyzed in water (10 mL in 4 L, 5×) and finally lyophilized to afford PMSEA-CoumNH2 as a foamy white solid (822 mg, 33%). Mn = 260 kg/mol, Ð = 1.47. GPC traces for all polymers used in this study are provided below in Figure 3.S10. 102 Determination of dn/dc of PMSEA polymers. Six different PMSEA polymers were synthesized via the procedure described above using the initiator 3.3. Three 2 mg/mL solutions of each polymer were prepared volumetrically and characterized by GPC (Table 3.S1). Data were analyzed using Wyatt Astra 7 to obtain dn/dc values using the 100% mass recovery method. Average dn/dc was calculated to be 0.153 ± 0.003 mL/g. GPC traces for all PMSEA-3 polymers are provided in Figure 3.S9. Table 3.S1. GPC measurements of all PMSEA-3 used for dn/dc determination. Monomer Polymeriz dn/dc Mn (kDa) Mp (kDa) Đ eq. ation time (mL/g) 46.3 55.5 1.25 0.1597 PMSEA450 1.5h 44.3 55.5 1.29 0.1529 3a 45 55.5 1.28 0.1525 147 207.5 1.42 0.1547 PMSEA1500 3.5h 146 207.3 1.42 0.1521 3b 142 207.4 1.47 0.1494 217 335.4 1.66 0.1530 PMSEA2500 4h 218 333.5 1.61 0.1508 3c 217 337.3 1.65 0.1540 285 380.7 1.62 0.1564 PMSEA3600 5h 269 389.5 1.69 0.1521 3d 268 393.3 1.71 0.1523 234 372.1 1.70 0.1517 PMSEA5000 5.5h 230 351.9 1.66 0.1583 3e 226 356.8 1.72 0.1564 376.1 545.3 1.85 0.149 PMSEA8000 1.67h 357.9 534 1.85 0.1473 3f 351.4 416.5 1.89 0.1546 103 Figure 3.S9 GPC traces (refractive index response) for polymers PMSEA-3a to PMSEA3f used to determine dn/dc of PMSEA. Each polymer was measured in triplicate. 104 Figure 3.S10. GPC traces (refractive index response) for each polymer studied. 3.6.4 General Procedure for Ultrasonication Experiments with an Immersion Probe An oven-dried sonication vessel was fitted with rubber septa, placed onto the sonication probe, and allowed to cool under a stream of dry nitrogen. The vessel was charged with a solution of the polymer in either MilliQ water or 3:1 acetonitrile/water (2 mg/mL, 20 mL total volume) and submerged in an ice bath. The solution was sparged with nitrogen for 1 h prior to sonication, and a nitrogen headspace was maintained throughout the experiment. Pulsed ultrasound (1 s on/1 s off, 20% amplitude, 20 kHz, 9.04 W/cm2) was then applied to the system. “Sonication time” is defined as the duration of exposure to sonication, which is 50% of experimental or “clock” time under 1 s on /1 s off pulsation. The solution temperature during sonication was measured to be ~10 °C by an internal temperature probe. Aliquots taken from the sonicated solution were filtered through a 0.45 µm PTFE syringe filter prior 105 to analysis. Ultrasonic intensity was calibrated using the method described by Berkowski et al.49 3.6.5 General Procedure for Experiment with Focused Ultrasound (FUS) FUS setup. A 2-mL microcentrifuge tube was placed on a 3D-printed sample stage, and the center position of the tube was determined using an L22-14vX 128-element Verasonics imaging probe. The tube was removed and exchanged with a needle hydrophone (Onda, HNR-0500) situated in the exact position as the previously registered center of the tube. A 330 kHz focused ultrasound transducer (Precision Acoustics, H115) was mounted on a computer-controlled 3D translatable stage (Velmex) orthogonally to the Verasonics probe. A previously reported MATLAB program was used to automatically scan and align the transducer’s focus at the center of the microcentrifuge tube, according to the feedback from the needle hydrophone, which reports the acoustic pressure.36 To calibrate pressure, the needle hydrophone was placed inside and at the center of a plastic microcentrifuge tube to account for the attenuation of ultrasound wave through the plastic tube. Spatial peak-temporal average acoustic intensity (𝐼$%#& ) is calculated with eq. 3.S1: 𝐼$%#& = 𝑝' ∙ 𝐷𝐶 2𝜌𝑐 (3. S1) where 𝑝 is the peak negative pressure (PNP) of ultrasound, 𝜌 is the density of aqueous media, 𝑐 is the speed of sound in aqueous media, 𝐷𝐶 is the duty cycle applied in ultrasound treatment. Temperature calibration. Temperature change inside the microcentrifuge tube was monitored by a temperature probe (Fluke T3000FC Temperature Module), which is inserted inside the tube. The volume of sample inside the microcentrifuge tube was maintained at 800 µL in all experiments. Once 330 or 916 kHz FUS was turned on, the temperature inside the tube was recorded until it reached equilibrium. The measured temperature remained relatively constant when the temperature probe was moved within the tube while the ultrasound remained on. Different ultrasound conditions, including pressure, duty cycle, cycle number in each emitting pulse were systematically investigated to find the upper limit of ultrasound conditions for maintaining biocompatible temperatures (Figure 3.2a). 106 Temperature increment in different sample container (plastic microcentrifuge tubes vs tissue-mimicking 1% agarose phantom) was also investigated with the same ultrasound conditions (Figure 3.2b). As demonstrated with the biocompatible FUS conditions, the temperature increase is higher for the sample in the microcentrifuge tube because the plastic walls absorb more ultrasound than agarose gel or water, resulting in increased heat dissipation at the interface. 3.6.6 Characterization of Aminocoumarin Release Using Photoluminescence Spectroscopy Aliquots from either the sonication (immersion probe) or FUS experiments were filtered through 0.45 µm syringe filters and added to either a quartz microcuvette or a 96well microplate for photoluminescence measurements. The samples were then analyzed by fluorescence spectroscopy on either a Shimadzu fluorimeter or an Tecan Spark microplate reader at room temperature. Emission spectra were recorded using an excitation wavelength of 365 nm. The concentration of released aminocoumarin (CoumNH2) was determined from calibration curves constructed on both the Shimadzu fluorimeter (Figure 3.S11 and Figure 3.S12) and the Tecan Spark microplate reader (Figure 3.S13). Aminocoumarin Release with Immersion Probe Sonication. As expected, efficiency of CoumNH2 release from PMSEA-CoumNH2 (2 mg/mL in water) increases as a function of exposure time to sonication (Figure 3.S16). Maximum release of CoumNH2 was observed after 120 min sonication in water (44%). In contrast to the significant release triggered from PMSEA-CoumNH2 containing a chain-centered mechanophore, sonication of chain-end functional polymer Control-CoumNH2 (2 mg/mL in water) does not result in significant release of CoumNH2 (Figure 3.S17), supporting that release occurs via a mechanochemical process.12 Aminocoumarin Release with Biocompatible FUS. While typical biocompatible FUS experiments were performed for only 10 min, CoumNH2 release efficiency was observed to increase with additional insonation of PMSEA-CoumNH2 (2 mg/mL in water, 1.4 nM GVs) (Figure 3.S4). In contrast, exposure of chain-end functional control polymer Control-CoumNH2 (2 mg/mL in water) to biocompatible FUS resulted in insignificant release of CoumNH2 (Figure 3.S5). Likewise, no significant release of CoumNH2 was 107 observed from either PMSEA-CoumNH2 or Control-CoumNH2 over 17 h upon heating to 50 °C, further supporting that release under FUS is mechanochemically triggered (Figure 3.S18). Insignificant release of CoumNH2 was also observed from PMSEACoumNH2 when peak negative pressure (PNP) was < 1.3 MPa, despite collapse of GVs occurring under these conditions (Figure 3.S19). The extent of CoumNH2 release was also dependent on the concentration of GVs, whereby a higher concentration of GVs resulted in greater percent release of CoumNH2, which can be attributed to an increased number of cavitation events and consequently a greater extent of mechanochemical activation (Figure 3.S20). Furthermore, insonation of PMSEA-CoumNH2 in the presence of pre-collapsed GVs (1.4 nM), wherein gas content inside the nanostructures and surrounding fluid is substantially depleted, resulted in insignificant CoumNH2 release, demonstrating that the gas content within intact GVs is crucial for efficient mechanochemical activation (Figure 3.S21). Finally, the extent of CoumNH2 release varied insignificantly with polymer concentration between 2 and 8 mg/mL (Figure 3.S22). Figure 3.S11. Construction of a calibration curve for experimental determination of the concentration of aminocoumarin (CoumNH2) from fluorescence measurements on a Shimadzu fluorimeter. Emission spectra (λex = 365 nm) and intensity at 440 nm for solutions of CoumNH2 in water (with ≤ 0.1% DMSO for solubility) as a function of concentration. A linear regression of the data in the inset gives the calibration function y = 5240x (R2 = 0.9996). 108 Figure 3.S12. Construction of a calibration curve for experimental determination of the concentration of aminocoumarin (CoumNH2) using a Shimadzu fluorimeter. Emission intensity at 424 nm (λex = 365 nm) for solutions of CoumNH2 in 3:1 acetonitrile/methanol as a function of concentration. A linear regression of the data gives the calibration function y = 4220x (R2 = 0.998). Figure 3.S13. Construction of a calibration curve for experimental determination of the concentration of aminocoumarin (CoumNH2) using a Tecan Spark microplate fluorescence reader. Emission intensity at 440 nm (λex = 365 nm) for solutions of CoumNH2 in water as a function of concentration. A linear regression of the data gives the calibration function y = 12000x (R2 = 0.999). 109 Figure 3.S14. PL spectra (λex = 365 nm) collected on a Shimadzu fluorimeter illustrate instantaneous aminocoumarin release during sonication of PMSEA-CoumNH2 using an immersion probe (2 mg/mL in water, 20 kHz, 9.04 W/cm2). Figure 3.S15. PL spectra (λex = 365 nm) collected on a Shimadzu fluorimeter illustrate aminocoumarin stability during sonication of PMSEA-3c using an immersion probe (2 mg/mL in water, 20 kHz, 9.04 W/cm2). 110 Figure 3.S16. Time-dependent PL spectra (λex = 365 nm) collected on a Shimadzu fluorimeter illustrate aminocoumarin release during sonication of PMSEA-CoumNH2 using an immersion probe (2 mg/mL in water, 20 kHz, 9.04 W/cm2). 111 Figure 3.S17. Release of aminocoumarin from PMSEA-CoumNH2 and ControlCoumNH2 as a function of sonication time quantified by PL measurements on a Shimadzu fluorimeter. Polymers were sonicated with an immersion probe (2 mg/mL in water, 20 kHz, 9.04 W/cm2). 112 PMSEA-CoumNH2 CoumNH2 Release (% yield) 20 Control-CoumNH2 15 10 5 0 0 5 10 Incubation time (h) 15 20 Figure 3.S18. Insignificant release of CoumNH2 from PMSEA-CoumNH2 is observed at elevated temperatures. Polymer solutions (2 mg/mL in water) were incubated at 50 °C. Timedependent CoumNH2 release was measured by PL measurements on a Tecan Spark microplate reader. PMSEA-CoumNH2 +GV +FUS 20 CoumNH2 Release (% yield) PMSEA-CoumNH2 −GV +FUS 15 10 5 0 1.1 1.2 1.3 1.4 1.5 Peak negative pressure (MPa) Figure 3.S19. Pressure-dependent release of aminocoumarin (CoumNH2) from PMSEACoumNH2 (2 mg/mL in water) after FUS treatment. FUS was maintained at 3000 cycles each pulse, 4.5% duty cycle (equivalent to 5 Hz pulse repetition frequency), 10 min total insonation at various pressures. Release yield was measured by PL intensity at 440 nm (λex = 365 nm) on a Tecan Spark microplate reader. For the samples containing GVs, the GV concentration is OD500 = 12, corresponding to ~1.4 nM. All measurements are based on an average of 3 replicates. Error bars represent standard deviation from three fluorescence measurements each on samples from three separate experiments (N=9). 113 CoumNH2 Release (% yield) 20 15 10 5 0 0 1 2 3 4 GV concentration (nM) Figure 3.S20. Effect of GV concentration on release efficiency of CoumNH2 from PMSEACoumNH2 (2 mg/mL in water). FUS was maintained at 1.47 MPa peak negative pressure, 3000 cycles each pulse, 4.5% duty cycle (equivalent to 5 Hz pulse repetition frequency), 10 min total insonation. Release yield was measured by PL measurements on a Tecan Spark microplate reader. The GV concentrations were OD500 = 6, 12, and 25 corresponding to ~0.7, 1.4 and 2.9 nM. All measurements are based on an average of 3 replicates. Error bars represent standard deviation from one fluorescence measurement each on samples from three separate experiments (N=3). 114 PMSEA-CoumNH2 ✱✱✱✱ 20 CoumNH2 Release (% yield) ✱✱✱✱ ✱✱ 15 10 5 +F U S S −G V +F U +c ol la ps ed +G V G V +F U S 0 Figure 3.S21. Effect of intactness of GVs on release efficiency of CoumNH2 from PMSEACoumNH2 (2 mg/mL in water). FUS was maintained at 1.47 MPa peak negative pressure, 3000 cycles each pulse, 4.5% duty cycle (equivalent to 5 Hz pulse repetition frequency), 10 min total insonation. Release yield was measured by PL measurements on a Tecan Spark microplate reader. Experiments were performed with intact GVs, with pre-collapsed and degassed GVs, and without GVs. The GV concentration was OD500 = 12, corresponding to ~1.4 nM. All measurements are based on an average of 3 replicates. Error bars represent standard deviation from three fluorescence measurements each on samples from three separate experiments (N=9). Asterisks represent statistical significance by ordinary one-way ANOVA with multiple comparison (****P < 0.0001; ***P < 0.001; **P < 0.01; NS, not significant). 115 Figure 3.S22. Effect of polymer concentration on the release of CoumNH2 from PMSEACoumNH2 (2 or 8 mg/mL in water) after FUS treatment. FUS was maintained at 1.47 MPa peak negative pressure, 3000 cycles each pulse, 4.5% duty cycle (equivalent to 5 Hz pulse repetition frequency), 10 min total insonation. Release yield was measured by PL measurements on a Tecan Spark microplate reader. The GV concentration was OD500 = 12, corresponding to ~1.4 nM. All measurements are based on an average of 3 replicates. Error bars represent standard deviation from three fluorescence measurements each on samples from three separate experiments (N=9). Asterisks represent statistical significance by ordinary one-way ANOVA with multiple comparison (****P < 0.0001; ***P < 0.001; **P < 0.01; NS, not significant). 3.6.7 Characterization of Camptothecin Release Using Liquid Chromatography Confirmation of CPT release via 1H NMR and LCMS. Prior to mechanochemical activation studies, we first confirmed that camptothecin was released in polar protic solvent upon unmasking of the expected unstable furfuryl carbonate. To enable characterization, the furfuryl carbonate species was generated thermally from small molecule initiator 3.1-CPT. Masked furfuryl carbonate 3.1-CPT was dissolved in toluene-d8 and analyzed by 1H NMR spectroscopy before and after heating for 24 h at 100 °C (Figure 3.S6a and Figure 3.S6b). Comparison to spectrum of the expected maleimide fragment confirmed significant thermal 116 retro-Diels–Alder reaction occurred under these conditions (Figure 3.S6c). A drop of this solution was then diluted into 3:1 acetonitrile/methanol and analyzed by LCMS (gradient of 0–50% acetonitrile/water over 4 minutes) (Figure 3.S6d). Comparison of the newlygenerated peak to a standard sample of pure camptothecin (CPT, Figure 3.S6e) reveals identical retention times, absorption spectra, and mass-to-charge ratios, supporting that CPT is released from the unmasked furfuryl carbonate upon exposure to polar protic solvent. Calculation of HPLC Relative Response Factors (RRF). Three standard solutions with known concentrations of 4-methyl-2-oxo-2H-chromen-7-yl acetate internal standard (IS) and CPT were prepared and analyzed by HPLC equipped with a UV detector. The RRF is calculated from the HPLC results (C8 column, isocratic 30% acetonitrile/water) of the standard solution using eq. 3.S2: Analyte peak area DAnalyte concentrationG Analyte Response Factor RRF = = 𝐈𝐒 peak area IS Response Factor D𝐈𝐒 concentrationG RRF at 254 nm was determined to be 6.71 ± 0.23 for CPT:IS (Figure 3.S23). (3. S2) 117 Figure 3.S23. The relative response factor (RRF) of analyte (CPT) versus internal standard (IS) was determined to be 6.71 ± 0.23 from the average of three separate volumetric solutions. Determination of the concentration of released CPT from polymers after FUS exposure. A solution of IS of known concentration was added to analyte solutions in a volumetric ratio of 1:3 (IS:analyte solution). The solution was then kept at room temperature and analyzed by HPLC (50 µL injection, C8 column, isocratic 30% acetonitrile/water) at various time intervals. The total concentration of released CPT was calculated using the relationship in eq. 3.S3: Released 𝐂𝐏𝐓 (µM) = Peak area of 𝐂𝐏𝐓 1 4 ∗ ∗ Concentration of 𝐈𝐒 (µM) ∗ (3. S3) Peak area of 𝐈𝐒 RRF 3 No release of CPT is observed from aqueous solutions of PMSEA-CPT stored under ambient conditions for 2 months (Figure 3.S8). Applying FUS (330 kHz, 3.6 W/cm2, 1.47 118 MPa PNP, 4.5% duty cycle, 10 min) to PMSEA-CPT (4 mg/mL in water) triggers release of CPT (7.732 ± 0.006%, average of three trials) in the presence of GVs (1.4 nM). No release is observed with FUS in the absence of GVs (main-text Figure 3.3, Figure 3.S24). Release of CPT is also not observed upon insonation of chain-end functional polymer Control-CPT in the presence of GVs (1.4 nM), supporting that mechanical force is responsible for activation (Figure 3.S25). Finally, free CPT was confirmed to be stable upon exposure to FUS (10 min) in either the presence or absence of GVs (1.4 nM) (Figure 3.S26). Figure 3.S24. Exposure of PMSEA-CPT (4 mg/mL in water) to biocompatible FUS (330 kHz, 3.6 W/cm2, 1.47 MPa PNP, 4.5% duty cycle, 10 min) in the presence of GVs (1.4 nM) triggers the release of CPT as measured by HPLC (C8 column, isocratic 30% acetonitrile/water). Experiments were run in triplicate, and samples were diluted 3:1 with a stock solution of IS (72.8 µM). Concentrations were determined from integration of the peak corresponding to CPT relative to that of the internal standard and scaled by 4/3 to account for dilution. No IS was used in the experiments without GVs. 119 Figure 3.S25. HPLC measurements demonstrating that exposure of chain-end functional polymer Control-CPT (4 mg/mL in water) to biocompatible FUS (330 kHz, 3.6 W/cm2, 1.47 MPa PNP, 4.5% duty cycle, 10 min) in the presence or absence of GVs (1.4 nM) does not result in the release of CPT. 120 Figure 3.S26. HPLC measurements demonstrating stability of CPT upon exposure to biocompatible FUS (330 kHz, 3.6 W/cm2, 1.47 MPa PNP, 4.5% duty cycle, 10 min) in the presence or absence of GVs (1.4 nM). Experiments were performed in duplicate on a sample of CPT in water (~3µM). After insonation, samples were diluted 3:1 with a stock solution of IS (40 µM) and concentrations were determined from integration of the peak corresponding to CPT relative to that of the internal standard and scaled by 4/3 to account for dilution. 3.6.8 Cytotoxicity Experiments MTT-based cell viability assay. Raji cells were used to evaluate the effectiveness of CPT release for potential chemotherapy. Raji cells were incubated in RPMI 1640 media, with 10% fetal bovine serum and 1% antibacterial Penicillin/Streptomycin (PenStrep) at 37 °C and 5% CO2. The instructions provided by the manufacturer (Abcam) were followed to perform the MTT assay. Approximately 10,000 Raji cells suspended in 150 µL of media were seeded into each well of a 96-well plate. Subsequently, 50 µL of polymer sample (or pure CPT) at varying concentrations was added to corresponding wells. For calibration purposes, wells without cells and wells with only cells and media were also prepared, representing 0% and 100% cell viability, respectively. After 48 hours of incubation, cells were spun down inside the plate and media were carefully aspirated. Then 50 µL of serum free media (Opti-MEM, ThermoFisher) and 50 µL of MTT solution were added to each well. 121 Following a 3 h incubation, 150 µL MTT solution was added to each well. The plate was thoroughly mixed and shaken on an orbital shaker at 37 °C for 15 min. The absorbance at 590 nm was measured by a microplate reader (TECAN Spark) to quantify the cell viability in each well. Cell viability is calculated using eq. 3.S4: 𝐶𝑒𝑙𝑙 𝑣𝑖𝑎𝑏𝑖𝑙𝑖𝑡𝑦 = 𝐴 − 𝐴()*+& -./0 𝐴1)// -./0 − 𝐴()*+& -./0 (3. 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