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Abstract

You often hear that humans are composed predominantly of water; how this works is much like a sponge. We soak that water through and around our cells, and it carries the nutrients and chemical signaling agents needed for those cells to survive and communicate. Oxygen (O₂ gas) dissolves in the water that makes us up and gets shuttled throughout the body by our blood cells. The oxygen molecule plays a major role in energy production, at the cost of playing a supporting role in side reactions that damage cells by attacking biomolecules, including individual lipids in the phospholipid bilayers that separate contents within a cell and provide barriers between cells. We call this oxidative stress because of oxygen’s role in harming these essential dividing layers of fat. Oxidative stress occurs naturally during a living creature’s lifetime and is not unique to humans. Over time, this stress can accumulate and lead to cell mutations and inevitable death (Villapando-Rodriguez and Gibson 2021; Chen et al. 2025).

Phospholipids are single molecules composed of two portions: Roughly one-third are the “head group,” or polar region, and the other two-thirds are the “tail group,” or nonpolar region. They are the fundamental building blocks of a cell’s exterior membrane as well as the divisions between its organelles, little functional portions of a single cell that carry out essential processes. The integrity of these membranes is crucial for a cell’s stability and functionality (Meer et al. 2008; Singer and Nicholson 1972). When membrane structure is compromised, pH gradients essential for cellular function can be disrupted, vital nutrients can leach into the extracellular space, and cells can become cancerous from damage to the DNA held within the nucleus (Su et al. 2019; Nakamura and Takada 2021). Damage to the phospholipid membranes of cells and their organelles is a major factor contributing to the effects of natural aging as well as severe diseases such as cancer and cardiovascular disorders—even neurodegenerative diseases like Alzheimer’s and dementia (Su et al. 2019; Nakamura and Takada 2021; Chang et al. 2004). Therefore, tools that enable the study of phospholipid bilayers are essential to fully understand how these membranes degrade and when they have reached the critical failure point. Understanding cumulative degradation of these cellular membranes may allow for preemptive correction or identification of complications that could lead to an organism’s death.

Researchers at the White-Mathieu lab at the �鶹app seek to understand the role of oxidative stress on the cellular environment by characterizing stages of cellular demise. We use nanoscopic tools called activity-based sensors, molecules that react with specific targets, resulting in a fluorescent response in the sensor, to not only detect the presence of harmful material within the cellular environment but also to report on the morphological changes to membranes cell-wide. One key aspect of this research was to develop a synthetic phospholipid for delivery and incorporation of these sensors into the cellular environment, and that step is the focus of this article. This project was supported by the Ronald E. McNair Post-Baccalaureate Achievement Program in 2024 with subsequent funding for 2025 generously provided by a Summer Undergraduate Research Fellowship from the Hamel Center for Undergraduate Research.

Free Radicals and Oxidative Stress

Peroxides, the simplest form of which can be found in our medicine cabinets, can be the starting point of oxidative stress. In the cellular environment, hydrogen peroxide (H₂O₂) serves as a signaling molecule between cells. Peroxides typically express to cells that inflammation is necessary in a damaged area. As they accumulate in an organism, however, peroxides produce an abundance of “free radicals,” which are molecules bearing unpaired, reactive electrons. Unpaired electrons are like a plague on a unified system of individual cells; they propagate and ultimately cause destruction in cellular membranes (Su et al. 2019).

Free radical propagation is stopped only when the radical is terminated by antioxidants within the environment. Until this termination point, radicals have many opportunities to cause cumulative damage while creating more radicals in the process. Detailed microscopic imaging of free radical accumulation and the exact effect they have on cellular membranes is crucial for evaluation of a system’s health. Detecting radicals within the cells of an organism is made possible by the employment of activity-based sensors, an advancement in nanotechnology made in the early twenty-first century (Chang et al. 2004; Bruemmer et al. 2020). An activity-based sensor operates on the principle of fluorescence, the same phenomenon that allows a glow stick to glow. Fluorescent response activation is like the cracking of a glow stick, and what cracks our molecular tools is the presence of radical precursors. Imagine each molecule as a tiny unlit lantern that floats around the water within cells. As that lantern encounters the compounds it selects for, it lights up. Fluorescent biosensors serve as powerful tools for real-time cellular imaging, enabling the detection of analytes such as peroxides, which contribute to oxidative stress and conditions like Alzheimer’s disease and cancer. 

Detection of hazardous substrates within a cell is just one piece of the puzzle; the other piece is anchoring these sensors to the phospholipid bilayers so that illumination of their structure is possible with a microscope. Despite advances in biosensor chemistry, a critical gap remains in the efficient integration of these sensors into live-cell environments. This project aims to develop an optimized method for incorporating a lipid-anchored fluorescent biosensor into cellular membranes to monitor oxidative damage.

Our approach seeks to anchor these sensors directly to cellular membranes to get an idea where harmful toxins are produced and determine a timeline with distinct stages indicating the health of a cell’s phospholipid membranes. To achieve this, we attach an activity-based sensor to a synthetic phospholipid outside of the cell using a robust technique known as “click chemistry.” We call it click chemistry because it is as simple as clicking a seat belt together. All that is required are two functional groups called an azide and an alkyne. An azide consists of three nitrogen atoms, and an alkyne consists of two carbon atoms triple bonded together that can react with the azide. When these two groups encounter each other, a strong triazole ring is created that “clicks” our targets together (Agard et al. 2004; Gutierrez et al. 2023). The resulting lipid and sensor conjugate can then be shuttled into live cell environments to monitor oxidative damage.

Synthesizing an Azido-Phospholipid

To develop a synthetic phospholipid, we must connect individual molecules in a process similar to what cells do to make phospholipids from scratch. The process begins with a fat molecule that has an average biologically relevant size, consisting of a tail length between sixteen and twenty carbon atoms long, which translates to approximately two nanometers long, or, roughly, four ten-millionths of an inch. Once we have the basic lipid structure, it must be made into a phospholipid derivative capable of enabling further connectivity. Last, we add an azide-bearing amine, which provides a place for the sensor to anchor. 

This process is what we call in chemistry a convergent sequence, which is a synthesis strategy for designing molecules by independently preparing multiple, smaller fragments and then joining them together in the final product. This allows us to have checkpoints along the way and leads to higher average yields. Also, a convergent sequence can be run in tandem, allowing for efficiency in production. 

To begin, we chose a fat derived from palm oil, 1,2-Dipalmitoyl-rac-glycerol, because of its optimal tail lengths of sixteen carbons. We added the compound to a flame-dried flask to ensure a water-free reaction environment; this is important, because under the experimental conditions, water can hydrolyze reagents required in the synthesis, thereby decomposing the material before it can react. Keeping water out of the reaction flask was essential, so throughout the synthesis the reaction was sealed and kept under a nitrogen line. To this we injected benzene and a strong, noncompetitive base, triethylamine, with a syringe, then stirred for fifteen minutes. The base pulls a proton off the alcohol on our starting material, leaving a reactive site in its wake. Next, we added ethylene chlorophosphate, a phosphate molecule containing a two-carbon chain, in a dropwise fashion to ensure ample dispersion. The reaction was then stirred at 0˚C for forty-eight hours. The long time span allows for complete reaction while the cold temperature stabilizes the solution as a bond is formed between the phosphate and the lipid. We then cleaned up the reaction by separating the product from the solvent, impurities, and unreacted starting materials in what we call the crude mixture. Once isolated, our phospholipid product came out as a white solid with an 88% yield, primed and ready for azido-amine addition (Gutierrez et al. 2023).

Next, our focus shifted to the azido-amine that would go on to compose the second half of this synthetic polar head group alongside the phosphate. The head group of the lipid sticks out toward the water side of the membrane and anchors to the biosensor because of the azide functionality. We began with the amine, 2-Bromo-N,N-dimethylethanamine hydrobromide. The goal of this reaction was to swap the bromine with an azide. This amine was added to a flask, but this time, dryness was not a problem because water was used as this reaction’s solvent. To this solution, I carefully added sodium azide with the end of a glass tube to avoid using a metal spatula for transfer. Sodium azide is a dangerous salt. If metal tools are used to measure it out, it could explode like a car’s air bag. That is because if an electron is supplied to this compound, it will spontaneously decompose into nitrogen gas. Chemistry is all about stability and working with higher-energy reactants to reach lower, more stable products; to work with the natural world and achieve our goals, we must play by Mother Nature’s rules (Jao et al. 2015).

After the sodium azide salt was added to the flask containing the bromo-amine, a small volume of water followed to dissolve it. The reagents were stirred in a hot oil bath at 85˚C for forty-eight hours. In this reaction, the azide functional group takes the place of the bromine to serve as an anchor point for the sensor once combined with the lipid. I then removed the product from heat, cooled it to room temperature, dropped it into an ice bath, and carefully combined it with a strong base to neutralize any acid present. After forty-eight hours we decanted the product as the top layer of this two-layer liquid mixture for a 45% yield (Jao et al. 2015).

With both precursors prepared, we could then couple the azide to the functionalized lipid. Our goal was to make a carbon nitrogen bond with the two-carbon chain of the phosphate and the nitrogen of the azido-amine. This final stage couples the two previously described materials, thus culminating the convergent sequence. First, an excess of the azido-amine was added to a small flask. To this we added a spatula’s tip of sodium sulfate to dry the reagent. A septum was affixed to the neck of the flask to keep air out, and dry acetonitrile was added as a solvent to dissolve the lipid and prevent water from entering the reaction environment. The flask was then sealed and affixed with a nitrogen inlet needle. The tip of the nitrogen needle was plunged into the liquid and left to bubble gas through the solution for three hours (Gutierrez et al. 2023).

A small, glass, pressure-resistant tube was then flame dried, and this served as the final reaction vessel. To this pressure vessel we added our phospholipid followed by more dry acetonitrile. The dried azide was then transferred by syringe to the pressure tube from the small flask. We sealed the pressure vessel and then submerged it in a 65˚C oil bath for 120 hours, after which the reaction was taken off heat and cooled. The final product precipitated into a solid as it cooled to room temperature in the pressure vessel. This was then transferred to a vial and separated from residual solvent by affixing a vacuum to the material. The vacuum caused the solvent to evaporate, leaving behind our desired azido-phospholipid product, ready for anchor with an activity-based sensor, in a 40% total yield. 

Findings Drawn from Azido-Phospholipid Experiment

Initial yields for the azido-amine reaction were very low. This led to a small-scale attempt at the final reaction, which also gave a low yield of 2%. We examined the final azido-lipid product under ¹H NMR spectroscopy, a fundamental analytical technique used to determine the structure of organic molecules. In this technique a powerful magnet is used to measure the electronic environment around a molecule’s nuclei. It can be thought of as an MRI machine for our chemicals—what would give an image of a knee joint instead gives us data that corresponds to portions of a desired compound. Information is recorded in the form of “chemical shifts,” or proton peaks, which can be compared with previously established values. If the values match or are similar to those obtained in past experiments, we can conclude that our final product was made. 

Initial NMR spectroscopy results for the final azido-lipid showed chemical shift data corresponding to both the starting material and final product, implying the reaction did not reach completion. Subsequent replication of this process was altered to modify the isolation of the azido-amine compound as described herein. This increased precursor yield from 15% to 45%, enabling synthesis of the final product in a yield that also increased from 2% to 40%. 

Each subsequent NMR experiment revealed promising results when the chemical shifts were examined and compared with the literature. NMR scans also showed the presence of small messy peaks as impurities in the final azido-phospholipid product, but this is expected because final purification is still under development. Experimental spectroscopic evidence supports the conclusion that the final product was indeed synthesized.

Conclusion and Further Research

The synthetic azido-lipid I developed for this research project serves as the basis for future biosensor anchoring and subsequent integration into a cell’s phospholipid bilayers. We will attempt to shuttle the entire sensor-phospholipid conjugate into the cellular environment where it may self-integrate within native phospholipid bilayers, thus incorporating a fluorescent activity-based sensor into cellular membranes for further bilayer monitoring. 

Final Click Coupling of an Activity-Based Sensor and Synthetic Lipid

The future direction for this project is to couple this azido-phospholipid to a variety of activity-based sensors. These sensors will be selective for various free-radical-generating compounds such as iron, hydrogen peroxide, and singlet oxygen. After creating a sensor-bound lipid, the complex will be dissolved and introduced to cells. The cells will then be treated with the analytes each sensor is selective for and left to incubate. Afterward, the cells will be examined by fluorescence microscopy. The images will be processed to quantify fluorescence signal and note any changes in membrane structure at sequential time intervals. This is one of the major ongoing projects in the White-Mathieu lab.

Professionally, this project has served as my first solo research project outside of industry. The experience developed my laboratory skills as a synthetic organic chemist as well as my confidence within the laboratory. During this time, I had the opportunity to learn advanced techniques and become a laboratory asset in my own right. It is invaluable to have this new set of synthetic skills for my career as a chemist and researcher. I aim to one day apply this knowledge to my own independent research laboratory.

Undoubtedly, this work could not have been undertaken without the chemistry department here at Parsons Hall. Thank you so very much to our instrumentation center, Pat Stone, John Wilderman, Philip Place, Nancy Cherim; Cindi Rohwer, the greatest building manager and absolute living legend (you’re my rock, Cindi); our stockroom specialist, Svetlana Shuba, for always having just what we need; my lab mates, Aakriti Garg, Tom DiPhilippo (aka my great mentor who honed me into the chemist I am today; I cannot thank you enough, TD), Paige Ring, Saghar Jarollahi, Yagmur Altunsoy, Arpan Ghosh, Prasanna Ganesh, Erin McCarthy, Taylor Stock, Kylie Armor, Maddie Pageau, Muriel Lubelczyk, Mason Russell, Aliyah Krestalica, and last but certainly not least, my good friend Sam Moreau, who has been there for me in both victory and defeat. I would also dearly love to thank our mentor Brittany White-Mathieu for teaching us not only science but how to be great scientists. Doc White-Mathieu, your guidance has grown me not only as a student, researcher, and scholar, but also as a person. You really have changed my life. You taught me how to manage my time, energy, and well-being, and to not need to be everything to everybody. I promise I will sleep properly at some point, ha ha! I will never forget you and no thanks will do it justice; instead, I will do great work in grad school and beyond. I would also like to thank my friends Nayanthara Krishnan, Ian Ferraro, and now Dr. Qian Liu for being so kind to me when I first arrived at UNH. Nayan, Qian, and Ian, you’ve always encouraged me to be my very best and always asked about my journey. I just want you to know that the kindness you guys have shown me over the years really made the difference for me—I can’t thank you enough. To my good friend Dr. Hale, thank you for giving me my first shot at research and always being there for me; you were the first one who saw something in me and really took me under your wing (pun intended). And just one more thing, Doc Berda, if you ever see this, “I’m on it, Boss.” Generous funding has been given for this project from the Ronald E. McNair Post-Baccalaureate Achievement Program in 2024 with subsequent funding for 2025 generously provided by the Hamel Center’s Summer Undergraduate Research Fellowship (SURF) program. 

References

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Author and Mentor Bios

Nick Mixon will receive a bachelor of science degree in chemistry from the �鶹app in May 2026. He is from Amesbury, Massachusetts, and during his time at UNH he became a McNair Scholar, Judge William Treat Fellow, TRIO Scholar, and undergraduate researcher. He plans to pursue his graduate education in chemistry, specializing in biologically inspired “smart polymers” for catalytic applications.

Brittany White-Mathieu is an associate professor in the Department of Chemistry at the �鶹app. She uses organic chemistry for the development of fluorescent molecules that address major challenges in biology. Focusing on the essential roles of membranes in cells, these novel fluorescent scaffolds are designed to investigate cellular signaling pathways and enable super-resolution imaging of cellular structures. Inspired by the unique properties of macrocyclic strictures, researchers in her lab also aim to develop a new class of next-generation fluorophores with superior optical properties.

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