Versatile Pre-Polymer Platform for Controlled Tailoring of Quantum Dot Surface Properties

Abstract

Quantum dots (QDs) hold immense promise for bioimaging, yet technical challenges in surface engineering limit their wider scientific use. We introduce poly(pentafluorophenylacrylate) (PPFPA) as a user-friendly pre-polymer platform for creating precisely controlled multidentate polymeric ligands for QD surface engineering, accessible to researchers without extensive synthetic expertise. PPFPA combines the benefits of both bottom-up and pre-polymer approaches, offering minimal susceptibility to hydrolysis and side reactions for controlled chemical composition, along with simple synthetic procedures using commercially available reagents. Live cell imaging experiments highlighted a significant reduction in nonspecific binding when employing PPFPA, owing to its minimal hydrolysis, in contrast to ligands synthesized using conventional prepolymers prone to uncontrolled hydrolysis. This observation underscores the distinct advantage of our pre-polymer system. Leveraging PPFPA, we synthesized biomolecule-conjugated QDs and performed QD-based immunofluorescence to detect a cytosolic protein. To effectively label cytosolic targets in such a dense and complex environment, probes must exhibit minimal nonspecific binding and be compact. As a result, QD-immunofluorescence has focused primarily on cell surface targets. By creating compact QD-F(ab’)2, we sensitively detected alpha-tubulin with ~50-fold higher signal-to-noise ratio compared to organic dye-based labeling. PPFPA represents a versatile and accessible platform for tailoring QD surfaces, offering a pathway to realize the full potential of colloidal QDs in various scientific applications.

Graphical Abstract

Introduction

Quantum dots (QDs) possess immense potential to expand the scope of bioimaging due to their superior optical properties, including bright signals, excellent photostability, and color purity with narrow emission spectra.^1–7 For example, QDs enable single molecule tracking experiments in biological samples,^{3, 8–9} which is challenging for dye-based imaging due to dim labeling signals from single molecules and prompt photobleaching under intense illumination. Additionally, short-wave infrared (SWIR)-emitting QDs offer unique opportunities for whole-organism imaging, single cell tracking, and high resolution image-guided surgery due to the increased tissue penetration of SWIR light combined with the brightness and photostability of QDs.^10–12 However, QDs have not been widely used in bioimaging, primarily limited to research groups with expertise in QD surface engineering. A major challenge is the steep technical barriers to obtaining high-quality QD probes. The stable performance of QD probes in biological systems is sensitively dependent on the precise composition and arrangement of surface functional groups.^13–16 Commercial QDs are unreliable in delivering consistent quality and performance,^{15, 17} highlighting the need for a method to reliably produce high-quality QD surfaces with intended properties. While various types of ligands have been developed to yield excellent biocompatibility and stability in complex in vivo environments,^{5, 11, 18–19} their synthesis mostly involves specialized equipment, extensive labor, and synthesis techniques. Further, QDs of various compositions are becoming available to cover various wavelength ranges and achieve reduced biotoxicity requiring versatile ligand chemistries that can incorporate various types of metal coordinating groups.^19–22

Among different classes of surface engineering methods, spanning micelle-forming ligands^{21, 23} to coordinating ligands^{21, 24–25}, multidentate polymeric ligands have become the gold standard for bioimaging with their unmatchable advantages.^{21, 26–28} The multivalency of surface-anchoring moieties ensures stable passivation of the QD surface, even in the absence of excess ligands.^{26, 29} This property is crucial for bioimaging as non-coordinating ligands must be removed to avoid their toxicity to biological systems. Stable coordination of ligands is crucial for achieving high quantum yields as well as maintaining the stability of QDs.^{13, 26–27} Furthermore, the multivalency expands the available QD surface-anchoring groups, accommodating neutral binding groups, such as imidazole and carbene, to provide quick ligand exchange with good stability.^{21, 30}

Despite the multilayered benefits of multidentate polymeric ligands, their implementation is still limited to a small group of experts. Existing methods involve a trade-off between synthetic accessibility and chemical controllability. (Figure 1A). A fundamental approach to producing multidentate ligands involves bottom-up polymerization of a specified combination of functional acryloyl or methacryloyl monomers (Figure 1B).^{5, 26, 31} This method often employs Reversible Addition-Fragmentation Chain Transfer (RAFT) polymerization, ensuring quantitative incorporation of a series of monomers as designed in a reaction (high controllability, circle i in Figure 1A). However, this approach requires laborious and low-yielding synthesis throughout ligand preparation (low accessibility, circle i in Figure 1A). The labor starts from the preparation of pure acryloyl or methacryloyl monomers with polyethylene glycol (PEG) units. PEG units are typically included in the monomers to grant biocompatibility of QD. However, PEG–containing acryloyl or methacrylol compounds are very expensive with limited options for functional groups from commercial sources. Therefore, it is typical to synthesize these monomers in-house for routine and specific use in the lab. The purification process for in-house synthesized acryloyl or methacryloyl monomers containing long PEG chains, and various required functional groups like amines, carboxylic acids, azides, etc., can be notably challenging, requiring multiple rounds of column purification with low yields. Furthermore, the bottom-up synthesis approach necessitates radical polymerization for each unique design with a different composition of monomers. This requirement poses a big roadblock for researchers without extensive synthetic expertise because radical polymerization requires specialized equipment and synthetic techniques such as a chemical fume hood equipped with a high vacuum pump and noble gas lines, freeze-pump-thaw techniques, and more. Additionally, the bottom-up approach often requires post-polymerization modification and/or additional protection-deprotection steps due to the incompatibility of certain functional groups with RAFT polymerization. For instance, monomers containing norbornene, one of the popular copper-free click reaction moieties, can inhibit the controlled addition of monomers, which is a key step required for successful RAFT polymerization. Moreover, any potential radical scavengers, such as sulfur-, amine-, and phosphorus-containing groups, can interfere with controlled RAFT polymerization, necessitating post-polymerization modification steps to incorporate such chemical handles or protection/deprotection steps to avoid undesired reactions during polymerization. More importantly, the critical assumption of the bottom-up strategy, random incorporation of the provided monomers into a polymer, does not always work; depending on a monomer structure, the reactivity of the monomers for radical polymerization varies, inhibiting the complete randomness in polymerization.³² Due to these technical difficulties, the bottom-up synthesis approach, despite its commendable control over the composition of multidentate ligands through quantitative reactions, poses a significant obstacle to researchers lacking synthetic skills.

Figure 1.

Current methods for QD surface engineering. A. Diagram illustrating two pivotal aspects in QD surface engineering: accessibility and controllability. Each circle represents a different strategy, its position reflecting the level of accessibility and controllability. The optimal strategy, marked as iii, showcases high accessibility and controllability. B. Bottom-up synthesis strategy using RAFT polymerization (i in A). C. Pre-polymer strategy using PNAS and PMA (ii in A). This method involves aminolysis with amine-containing compounds to form QD ligands. D. Undesired side reactions in PNAS and PMA. The schematic is partially reproduced with permission from ref. 39. Copyright 2017 American Chemical Society. Side reactions include hydrolysis, ring opening, and substitution imidation, which hinder the incorporation of intended functional groups and result in undesired charges on the polymer backbone.

To lower these synthesis barriers, an alternative strategy has been developed utilizing pre-polymer platforms employing activated esters. Commonly used pre-polymers include poly(N-hydroxysuccinimide) (PNAS)^{21, 28, 33} and poly(maleic anhydride) (PMA).^{30, 34} In this method, universal pre-polymers serve as the foundational backbone. Through a straightforward aminolysis process, the activated esters within the pre-polymers are replaced with amine compounds in desired ratios, yielding the final coating polymers for QDs. This approach is technically much more accessible than the bottom-up method. Firstly, it eliminates the need for synthesizing acryloyl monomers, thereby bypassing the associated synthetic challenges. Secondly, the wide availability of various amine monomers commercially streamlines the synthetic process. While pre-polymers require polymerization akin to the bottom-up synthesis, their commercially available monomers, such as N-hydroxysuccinimide acrylates or maleic anhydride acrylates, simplify the process. Notably, the single-step polymerization of PPFPA accommodates diverse designs for QD-coating polymers. In contrast to the bottom-up method, which demands air-free RAFT polymerization for each specific QD-coating polymer design, the pre-polymer method eliminates the need for individualized polymerization steps. Instead, a one-pot reaction with the pre-polymer and amino compounds enables quantitative functionalization, significantly streamlining synthesis processes (Figure 1C). ^{21, 28, 30, 33–34} To illustrate, while 10 different QD-coating polymers would require 10 separate batches of polymerization in the bottom-up strategy, a single bulk synthesis of PPFPA can produce numerous QD-coating polymers with diverse designs and quantities, as long as PPFPA is available. This bulk preparation of pre-polymer offers an additional advantage of PPFPA methods over the bottom-up approach; bulk synthesis is often more manageable in terms of controlling the reaction compared to small-scale synthesis in the bottom-up approach, where the ratio of initiator, RAFT agent, and monomer significantly impacts the quality of the polymer. These advantages are highly appealing to researchers, leading to the widespread adoption of the pre-polymer strategy for QD surface engineering (circle ii in Figure 1A). ^35–37

However, the available pre-polymer platforms, such as PNAS and PMA, have a significant weakness in compositional control due to the vulnerability of N-hydroxysuccinimide (NHS) and anhydride groups to hydrolysis and side reactions during aminolysis³⁸ (Figure 1D). Uncontrolled hydrolysis of these groups unintentionally introduces carboxylic acids into the pre-polymers, hampering the reliability of the approach. Also, both PNAS and PMA display uncontrolled side reactions. The NHS groups in PNAS can undergo ring-opening or imidation by adjacent substituted amines (Figure 1D), defying a chemical composition from the design.^39–40 These side reactions can occur in a non-reproducible manner changing the charge and chemical composition of the surface, thereby altering the biological behavior of the QDs from batch to batch. Moreover, PNAS and PMA have limited solubility in organic solvents other than DMSO and DMF, which are very difficult to obtain in a complete anhydrous condition state.^{38, 41} This limited solvent choice further increases the chance of hydrolysis for PNAS and PMA. Additionally, it is worth noting that any ring-opening reactions in PMA inevitably generate carboxyl groups along the polymer backbone, limiting the design of chemical properties for downstream experiments. In summary, although the existing pre-polymer method offers greater accessibility to the scientific community by minimizing the synthetic challenges, it falls short of meeting the critical need for fine control over the chemical composition of QD-coating polymers, which is essential for reliably producing high-quality QD-based probes.

Here, we present a new pre-polymer platform, poly(pentafluorophenylacrylate) (PPFPA), addressing the challenges of QD ligand synthesis. It offers technical simplicity and precise control for incorporating diverse functional groups onto the QD surface. With its high stability against hydrolysis and compatibility with various organic solvents, PPFPA enables modular, accessible, and controllable ligand synthesis.^{39, 42–43} Its components, including pre-polymer backbones and amine-modified functional groups, can be either purchased or easily synthesized using known procedures.²⁶ Moreover, PPFPA facilitates the quantitative incorporation of various functional groups. While the structure of amino compounds can affect the nucleophilicity of amine for aminolysis, previous reports show quantitative conversion of PFP units with the provided ratio of various types of amines.^{38, 44–47} We have created a series of multidentate QD-coating ligands from PPFPA that incorporate various moieties for metal coordination–imidazole, carbene, and carboxylic acid–, water solubilization–PEG–, and conjugation–carboxylic acid, amine, and azide. Unlike the existing platforms, which exhibit uncontrolled hydrolysis, the PPFPA-based ligand synthesis yielded QDs with precisely the desired properties. Characterizations using zeta potential, gel electrophoresis, and NMR measurements show that the PPFPA platform achieves quantitative incorporation of monomers with negligible undesired side reactions. QDs with PPFPA-derived ligands were conjugated with DNA and proteins for potential imaging applications, with QD-antibody conjugates sensitively detecting alpha-tubulin in human osteosarcoma U2OS cells.

Results and Discussion

Strategy for low technical hurdles and high controllability.

The pre-polymer strategy offers a more accessible and cost-effective alternative to bottom-up synthesis. The bottom-up approach involves polymerization of high-purity acryloyl monomers, which often need to be synthesized in-house. Different polymer compositions require a different set of monomers to be synthesized and polymerized. The acryloyl monomers used for QD ligands typically consist of an acryloyl group, a PEG segment, and additional functional groups that serve as polymerization handles, impart hydrophilicity, and provide essential chemical properties, respectively. However, obtaining high-quality monomers with >95% purity is laborious due to the polymeric and hydrophilic nature of the PEG chain; these attributes extend the elution time of the compounds through the column and cause overlap with side products whose structures are largely similar to the desired products. In our experience, yields can be as low as 10% depending on the molecular structure. Consequently, the options available for these compounds from commercial sources are very limited, with only a handful of choices for PEG length and functional moieties. Furthermore, acryloyl-PEG compounds come at a significant cost, ranging from $380 to $1350 per 500 mg (source: Broadpharm). The bottom-up method demands separate RAFT polymerization for each distinct QD-coating polymer design, even with slight variations in monomer ratios, necessitating synthetic expertise in both small molecules and polymers. To afford polymers with uniform lengths and narrow molecular weight distributions, synthetic expertise is required to handle vacuum and inert gas lines and to optimize polymerization conditions depending on monomer types. The pre-polymer strategy, on the other hand, allows the pre-polymer to be purchased or produced in bulk. A one-step incubation with a selected ratio of amino compounds produces the final QD-coating polymers via aminolysis, providing a much simpler and more manageable route than RAFT polymerization. A wide range of amino-PEG compounds are commercially available, offering numerous options in PEG length (1-36 units) and functional groups such as carboxylic acid, butyl ester, Boc-protected amine, alcohol, azide, alkyne, and more, at competitive prices, making this approach even more accessible.

As discussed, current pre-polymer platforms are prone to hydrolysis and unintended side reactions. These reactions alter the surface charge and chemical composition of QDs, impacting their binding and transport behavior in vitro and in vivo.^{14, 48} Moreover, the number of functional groups on the surface affects conjugation efficiency. As a result, ligands prepared by the pre-polymer approach have yielded less reliable performance than those produced bottom up. Our newly developed PPFPA pre-polymer system stands out due to its robustness, exhibiting minimal susceptibility to hydrolysis and undesired side reactions. The hydrophobic nature of the pentafluorophenyl (PFP) group is key to its low susceptibility to hydrolysis and broad solubility range.⁴¹ This system gains an advantage from the commercial availability of both pentafluorophenyl acrylate and methacrylate at competitive prices, facilitating widespread access to the polymerization process. Once PPFPA is produced in bulk, the only step left is a simple, one-step incubation for aminolysis, a commonly used procedure in commercial conjugation kits. Moreover, the replacement of PFP units in PPFPA can be easily monitored using ¹⁹F NMR, as fluorine groups exhibit distinct chemical shifts when present within the PPFPA backbone compared to when present as pentafluorophenol, after being replaced via aminolysis.^{39, 41} Compared to the simple ¹⁹F spectra, ¹H NMR spectra of the same polymers have multiple broad peaks, making it difficult to cleanly distinguish and integrate the monomer versus polymer peaks. These advantages have allowed PPFPA to be exploited in various contexts, such as oligopeptides,⁴² single-chain organonanoparticles,^{44, 46} fabrication of silicon surfaces with biomolecules^49–50, and so on. To leverage these exceptional properties, we evaluated the suitability of PPFPA as a dependable pre-polymer for the consistent and precise preparation of QD-coating polymers.

PPFPA as a robust pre-polymer platform allowing precise control of the QD surface.

To evaluate the performance of our PPFPA pre-polymer compared to conventional pre-polymer platforms, we conducted experiments using two sets of QDs. Each set was coated with identically designed ligands, but one set used PPFPA and the other used PNAS as the pre-polymer. We selected PNAS over PMA for this comparison based on two primary reasons. Firstly, aminolysis of PMA inevitably results in the formation of carboxylic acids, which impedes the synthesis of polymers resembling those derived from PPFPA-based or bottom-up synthesis methods and leads to altered surface charges. Secondly, aminolysis of PPFPA and PNAS follows the same preparation procedure. Specifically, during the aminolysis of PPFPA and PNAS, byproducts of pentafluorophenol and N-hydroxysuccinimide, respectively, are generated during the replacement of the activated ester. These byproducts are easily eliminated through simple polymer precipitation. Conversely, aminolysis of PMA does not produce byproducts because the process involves ring opening, with resulting carboxyl groups remaining on the backbone (Figure 1C).

Both PPFPA and PNAS were prepared by RAFT polymerization.^{33, 44} Note that the degree of polymerization was slightly different for PPFPA (DP = 100) and PNAS (DP = 120) when following published protocols. We then produced neutral poly(imidazole) ligands (PILs)²⁶ by aminolysis of either PPFPA or PNAS with 40% imidazole groups as QD-anchoring moieties, and 60% PEG11-OMe as neutral and hydrophilic moieties (Figure 2A). Aminolysis was achieved using a one-step incubation in DMF at 50°C with amino compounds required for the final structure. It is important to highlight our rationale for selecting DMF for aminolysis reactions. Firstly, we aimed to maintain uniform reaction conditions across both PNAS-based and PPFPA-based approaches to effectively compare their resistance to hydrolysis. The composition of each unit in the final polymer was controlled by the stoichiometric addition of each amino monomer, assuming quantitative conversion of the pre-polymer. To minimize differences between the two synthetic routes, we standardized experimental procedures for aminolysis, including pre-polymer storage, reagent selection, and reaction conditions for both PPFPA and PNAS. Because PNAS is highly prone to hydrolysis, we took extra precautions to maintain a dry environment. This involved storing PNAS at 4°C with drierite, drying the reagents under high vacuum, and conducting the reaction under dry N₂ gas and in an anhydrous solvent from the Solvent Purification Systems.

Figure 2.

Comparison of PPFPA- and PNAS-based methods. A. Synthetic routes to create a neutral polymer from PNAS or PPFPA. B. Nonspecific binding of QDs generated using PNAS or PPFPA in live U2OS cells. Arrows indicate the line for the intensity plot in C. The detected fluorescence from QDs in raw images is displayed in white to ensure clear contrast against the background. Both images have the same contrast range, with the minimum set to the maximum signal intensity of PPFPA-based QDs. Higher contrast images are available in Figure S9. C. Intensity plot across the arrows in image B. QDs created from PPFPA showed significantly lower nonspecific background signals compared to those of QDs generated from PNAS. D. Surface charge properties demonstrated by agarose gel electrophoresis and zeta potential. E. ¹H NMR data of PNAS-based polymer P1. F. 1H NMR data of PPFPA-based polymer P2.

We analyzed the resulting polymers using ¹H NMR to assess their chemical composition (Figure 2E–F, S5). Quantifying protons within PEG chains poses challenges due to their broad peak appearance and potential overlap with water impurity peaks. Also, dynamic configurations resulting from single bond rotation and the amorphous structure of PEG influence electron density. To address this complexity, we focused terminal methoxy protons (H_OMe), characterized by a distinct chemical shift and single peak, and the protons at the polymer terminus (H_term). As summarized in Table 1, the theoretical proton counts present in the methoxy groups in P1 and P2 are 216 (3 H per OMe group x 72 units) and 180 (3 H per OMe group x 60 units), respectively. When the integrated peaks were normalized to the terminal methyl group in a chain transfer agent (3H, H_term), P2 had 184 H_OMe, which is very close to the calculated values, suggesting quantitative incorporation. We also verified the complete replacement of all PFP units in the PPFPA pre-polymer using ¹⁹F NMR (Figure S2). In contrast, P1 contained only 172 H_OMe compared to the theoretical value of 216, showing ~80% efficiency in incorporating the corresponding amino compounds. These data suggest an unintended loss of ~20% of NHS groups, likely due to hydrolysis, despite efforts to keep the reagents involved dry. Consistently, a significantly lower number of PEG protons (H_PEG) was detected for P1 compared to P2. It is noteworthy that the theoretical number of H_PEG calculated based on the average molecular weight (M_n) of PEG is less accurate. Thus, the H_OMe to H_term ratio was used to calculate the conversion efficiency. The degree of hydrolysis is hard to control and would vary from experiment to experiment, leading to unreliable QD surface engineering. Undesired hydrolysis and unreliable incorporation efficiencies of PNAS have been reported in various studies in polymer chemistry.^{38, 45, 51}

Table 1.

Summary of ¹H NMR analysis of P1 and P2

Prepolymers	Calculated/Detected	HPEG	HOMe	Hterm	Conversion efficiency
P1 (From PNAS)	Calculated	~3600	216	3	~80%
	Detected	~2148	172	3
P2 (From PPFPA)	Calculated	~3000	180	3	~100%
	Detected	~2529	184	3

Gel electrophoresis and zeta potential measurements revealed distinct surface charge properties of QDs coated with ligands synthesized using different pre-polymers. QDs coated with ligands from the PNAS pre-polymer (P1-QDs) exhibited a negative charge (ζ: −10.3 mV), suggesting hydrolysis of NHS moieties to carboxylates (Fig. 2D and S7). In contrast, those with the ligands from our PPFPA pre-polymer (P2-QDs) were completely neutral (ζ: 0.1 mV). We further investigated how the presence of unintended carboxyl groups on the QD surface affects their nonspecific binding to cells and proteins. To this end, we conducted live-cell imaging assays using U2OS cells (Figure 2B–C and S9). Cells were treated with 500 nM QDs at 4°C in McCoy cell culture medium, washed, and then imaged to assess nonspecific adsorption of QDs (Figure 2B–C). The imaging conditions were consistent for both QD samples, with identical acquisition times and intensity settings. Two notable differences are observed. Firstly, background signals were significantly higher for P1-QDs than for P2-QDs (23000 vs 16000). Additionally, P1-QDs displayed nonspecific binding peaks with a signal-to-noise ratio (S/N) more than an order of magnitude higher than those of P2-QDs (~2964 vs. ~37). These disparities seem to arise from the use of distinct polymers in QD surface engineering. Analysis of data from NMR spectra, zeta-potential, and gel electrophoresis suggests that P1-QD carries carboxylate groups from undesired hydrolysis, while P2-QD lacks them as intended. Previous studies indicate that a net neutral charge minimizes nonspecific binding compared to charged QDs.²¹ For this comparison, P1-QDs and P2-QDs are synthesized from the same batch of QDs and tested on cells from identical batches to minimize potential variations affecting their activities. We also performed protein nonspecific binding tests by incubating QDs in fetal bovine serum (FBS) and performing gel filtration chromatography (GFC) (Figure S8). For both QD samples, the size distribution of QDs remained constant after FBS incubation indicating minimal protein adsorption even for the negatively charged P1-QDs. The nonspecific binding studies highlight the superiority of PPFPA as a reliable and easy pre-polymer platform to create QD–coating polymers, compared to the widely used PNAS pre-polymer.

Versatility and controllability of PPFPA.

To evaluate the versatility of PPFPA, we created a series of QD-coating polymers P2–P7 by aminolysis (Figure 3A). P2–P7 were designed to include various chemical moieties relevant to various applications in the nanobiotechnology field, along with different QD-coordinating moieties such as imidazole and carbene. The inclusion of different metal-binding moieties enables the polymers to be compatible with different types of QDs. Specifically, imidazole groups stably coordinate with cadmium-based QD surfaces, while carbene moieties are effective ligands for zinc-based surfaces.⁵² Furthermore, the anionic polymers P4 and P5 consist of four different monomers, including PEG-amine and PEG-carboxylic acid. This design underscores our aim to investigate increased complexity in polymer composition, featuring four distinct segments. While the amine segment was not utilized in our study, it holds potential as a functional group for post-conjugation through amide coupling. The synthesis of these polymers was straightforward, requiring only a simple one-pot incubation at room temperature with a pre-defined mix of amino compounds. The only exception was P7, which requires additional steps for carbene derivatization (SI, Figure S5). The amino compounds used were either purchased or synthesized following the reported procedures.²⁶ Regarding a potential side reaction with the secondary amine in the imidazole ring, we posited that its occurrence would be minimal based on several considerations. Firstly, aminolysis reactions typically prefer primary amines due to their heightened nucleophilicity, and our reaction setup involves an excess of primary amines compared to the secondary amine in imidazole (140 primary amines vs. 40 secondary amine in imidazole). Additionally, the nucleophilicity of the secondary amine in imidazole is further impeded by its aromaticity and steric effects. Moreover, we did not detect an additional peak in the range of imidazole protons indicative of this side reaction, affirming our rationale. Also, our decision to use DMF as the reaction solvent was influenced by the solubility of histamine rather than the presence of PPFPA. Should another QD-anchoring moiety be adopted with solubility in midrange polar solvent, post-modification of PPFPA could feasibly occur in those solvents. Our subsequent analyses and biological evaluations focused on the imidazole-based polymers P2–P6 to eliminate any variables stemming from the coordination chemistry.

Figure 3.

Fine-tuning of QD surface properties with various chemical properties and functional moieties. A. Series of polymers created using the PPFPA platform via aminolysis with designed ratios of different amine compounds. B. Agarose gel electrophoresis of QDs coated with P2-P6, demonstrating different migration patterns based on the intended QD charges. Zeta-potential measurements further confirmed the designed charge properties of each QD type. C-D. 1H NMR analysis for the ratio of imidazole protons (H_a and H_b) to protons in carboxylic acid moieties (H_c – H_e), confirming the incorporation of monomers with the intended ratios in P4 and P5.

We first investigated whether the composition of the QD-coating polymers matched the input amine ratios, especially for polymers made with a complex mixture of amine compounds. For this test, we used ¹H NMR to characterize the pendant composition of P4 and P5 produced by incubating PPFPA with four different amine compounds: imidazole, methoxy-PEG, carboxylate-PEG, and amine-PEG. We focused on analyzing the imidazole protons (H_a and H_b in Figure 3A&C–D) and the protons in the methylene groups adjacent to the carboxylate (H_c + H_e and H_d in Figure 3A&C–D) since their chemical shifts are distinct from the other protons. Polymers typically exhibit slight deviations in peak integration values, but we found that the integration ratios of these protons closely matched their theoretical values. The expected ratios of these protons in P4 and P5 are H_a:H_b:H_c+e:H_d = 1:1:2:1 (40H:40H:80H:40H) and H_a:H_b:H_c+e:H_d = 1:1:4:2 (40H:40H:160H:80H), respectively. The observed values were 1:1.1:2.1:1.1 and 1:1:3.6:1.7, confirming that the chemical moieties were incorporated in a controlled manner as designed.

We then prepared QDs coated with P2-P6 and characterized their physical properties. All QD samples had a hydrodynamic size of 11-14 nm as measured by GFC, confirming their compact size and lack of aggregation (Figure S6). Zeta potential measurements confirmed the expected charge of the QDs based on the ligand design. The QDs coated with neutral ligands showed minimal zeta potential values (P2-QDs: 0.1 mV and P6-QDs: −1.4 mV), while those coated with positive (P3-QD: 5.6 mV) and negative ligands (P4-QDs: −11.9 mV and P5-QDs: −19.3 mV) demonstrated corresponding charges. Although it is known that the zeta potential is not linearly correlated with charge density,⁵³ it accurately reflects the relative intensity of the charges. For example, P5 contains a higher net negative charge than P4, and indeed P5-QDs have a more negative zeta potential than P4-QDs. Gel electrophoresis of these QDs revealed differential migration patterns, aligning with their intended designs. As shown in Figure 3D, QDs coated with neutral ligands-P2 and P6- remained in the well, while QDs coated with negatively charged ligands-P4 and P5- migrated toward the cathode. Moreover, P5-QDs, which have a higher net negative charge than P4-QDs, exhibited faster migration. Intriguingly, P3-QDs containing 10% amino-PEG groups, which have a positive zeta potential (ζ: 5.6 mV), showed no migration toward the cathode. This result can be attributed to the basic nature of the gel running buffer (pH: 8.3) and the low content of NH₂ moiety (5%). Compared to the buffer used for zeta potential measurements (pH: 7.4), the more basic buffer used for gel electrophoresis would result in less protonation of amines.

QD-biomolecule conjugates for various applications at the nano-bio interface.

QD-based bioimaging requires conjugation of QDs with biomolecules such as DNA or proteins, which serve as targeting handles. Leveraging the facile, controlled, and reproducible synthesis offered by the PPFPA platform, we created QD–DNA and QD–protein conjugates and evaluated their performance in vitro. Conjugation was achieved via strain-promoted click chemistry between dibenzocyclooctyne (DBCO)-modified biomolecules and azide-displaying P6-QDs (Figure 4). The successful conjugation between QD and DNA was confirmed by the notable migration of the resulting QD–DNA conjugates observed in gel electrophoresis and a significant shift in zeta potential (𝜻 = –1.4 ± 2.4 → −32.5 ± 2.8, Figure 4A–B). Upon DNA conjugation, the hydrodynamic size was also increased from 15.5 nm to 37.9 nm, based on DLS measurements (Figure S10). The number of DNA strands per QD was measured by hybridizing QD–DNA with complementary DNA oligos labeled with a Hex dye. The absorption spectrum of these conjugates ((1) in Figure 4C) displayed the absorption features of both QD and Hex dye, and the deconvoluted spectrum ((1)–(2) in Figure 4C) clearly showed the absorption spectra of Hex. The absorption spectra confirm that ~7 DNA strands are coupled to each QD. The degree of DNA conjugation can be adjusted by modulating the percentage of a conjugating moiety within the polymer or by changing the stoichiometry in the conjugation reaction between DBCO–DNA and azide–QDs. We also demonstrated protein conjugation using streptavidin (Figure 4D). Streptavidin (SAV) was chosen for a representative protein conjugation because SAV can be utilized in diverse applications including pull-down assays and biotin-mediated conjugation to other biological molecules.⁵⁴ To confirm successful conjugation, QD–SAV conjugates were tested by incubation with biotin–labeled DNA oligo, followed by gel electrophoresis. As shown in Figure 4E, QD–SAV conjugates incubated with biotin-DNA showed significant migration toward the cathode (lane 4).

Figure 4.

Conjugation of QDs with biomolecules. A. Schematic of QD-DNA conjugation and its characterization. B. Agarose gel electrophoresis and zeta potential analysis demonstrating successful QD-DNA conjugation. C. Absorption spectra of (1) QD–DNA with Hex-labeled complementary DNA, (2) QD alone, and the recovered Hex dye spectra by subtracting (2) from (1). D. Schematic of QD-streptavidin conjugation and its characterization. E. Successful conjugation of QD–streptavidin evidenced by migration toward the cathode in the presence of (lane 4). Simple mixing of QDs and biotinylated DNA without streptavidin conjugation did not migrate on a gel (lane 1).

We employed this protein conjugation method to generate QD–antibody conjugates and assessed the capability of these probes for cytosolic immunofluorescence targeting alpha-tubulin. Although QD-based immunofluorescence has been established, challenges remain in improving detection efficiency, accuracy, and stability. These aspects depend critically on reliable surface engineering to regulate the chemical properties of functional groups and charges.⁵⁵ Notably, QD-antibody conjugates obtained from commercial sources often exhibit inconsistent labeling efficiency or aggregation.⁵⁶ This inconsistency may be attributed, in part, to the lack of a reliable method to create QD-based probes in a robust and reproducible manner. More importantly, most existing QD-based labeling predominantly targets protein markers on the cell surface, which are comparatively more accessible than cytosolic proteins.⁵⁵ The densely packed and complex nature of the cytosol environment, coupled with the diverse array of biomolecules within it, poses challenges for effective QD delivery and potential off-target interactions. We have leveraged the exceptional stability and biocompatibility of the PIL design, along with precise control over the chemical composition on the QD surface using the PPFPA strategy, to produce high quality QD probes with minimal non-specific binding. To achieve effective probe delivery in the dense cytosolic environment, we prepared compact antigen-targeting probes by conjugating fragmented F(ab’)2 antibodies with P6-QDs (Figure 5A). The resulting QD–F(ab’)2 had a hydrodynamic diameter of ~20 nm as determined by DLS measurement (Figure S12).

Figure 5.

QD–based immunofluorescence. A. Schematic of QD–F(ab’)2 conjugation. B-E. Fluorescence images of dye–based (B&D) and QD–based (C&E) labeling of alpha-tubulin. Dye and QD images were acquired at 500 ms, and 250 ms, respectively. Image contrast scales vary between images to best visualize the signals in each image with fine filamentous structures of tubulin. D-E. Enlarged views of B and C. The arrows indicate the line for the intensity plot in F. F. Intensity analysis plot for alpha-tubulin labeling, focusing on peripheral filaments to minimize signal interference from neighboring actin filaments.

These QD-F(ab’)2 probes sensitively labeled the cytosolic target alpha-tubulin in U2OS cells detecting its fine filamentous structure (Figure 5 and S13). The signal intensity plot in Figure 5F shows that the fluorescence signal from QD-based immunofluorescence is significantly brighter and more distinct compared to dye-based labeling. Despite slightly higher background signals due to the intense fluorescence of QDs, the signal-to-noise ratio over single tubulin was 46-fold higher with QD–F(ab’)2 than with Alexa568–F(ab’)2 even though the acquisition time for the QD images was half that of the dye, a measure taken to prevent intensity saturation. This result confirms the effective diffusion of QD-F(ag’)2 due to its compact size and minimal nonspecific interactions with other cellular components.

Conclusion

The introduction of the PPFPA pre-polymer platform offers exceptional controllability while minimizing technical complexity. Using this platform for QD-coating polymer synthesis eliminates the laborious processes including monomer synthesis, column purification, and ligand-by-ligand polymerization, required by the bottom-up approach. Instead, straightforward one-pot aminolysis using commercially available amino compounds simplifies QD surface engineering. The PPFPA addresses the limitations of current pre-polymer methods, such as uncontrolled hydrolysis and side reactions, and provides precise control over the chemical composition of the ligands. In essence, PPFPA acts as the pre-polymer platform, ultimately yielding products equivalent to PILs synthesized via bottom-up methods, albeit with considerably streamlined procedures and minimal side reactions. PPFPA pre-polymer leverages the advantages of currently available methods by merging precise control over chemical composition from the bottom-up approach with the simplicity and ease of synthesis inherent in the pre-polymer method. Also, it is crucial to understand that it is PILs that passivate the QD metal surface, not PPFPA. Through comprehensive characterization using ¹H NMR, zeta-potential, gel electrophoresis, and live cell imaging, we confirmed that QDs produced via the PPFPA approach exhibit the intended physicochemical properties. In contrast, QDs produced by the commonly used PNAS-based method suffer from unintended negative charges and increased nonspecific binding due to uncontrollable hydrolysis.

Using the PPFPA-based chemistry, we produced QD-DNA and QD-protein conjugates and performed immunofluorescence. The PPFPA-based synthesis consistently yielded compact QDs with desired charges and surface functional groups, enabling efficient conjugation to both DNA and proteins. Our QD-F(ab’)2 probes effectively detected the cytosolic target, alpha-tubulin, with minimal background and ~50 times higher signal-to-noise ratio compared to traditional organic dye-based immunofluorescence. PPFPA significantly lowers the synthetic barrier for creating high-quality QD probes due to its simple synthetic procedure, commercial availability of reagents, and capability to generate stable, biocompatible, and compact QDs with precise control over chemical composition. The PPFPA backbone, synthesized in bulk using commercial PFPA monomers, offers a one-step reaction to generate ligands tailored for specific research. Unlike existing pre-polymer systems, PPFPA is soluble in common organic solvents, expanding its use in hydrophobic ligand synthesis. The advantages offered by PPFPA extend beyond bioimaging and have broad implications as a tool for controlled surface engineering in diverse nanoparticle applications. Its utility ranges from facilitating drug delivery systems and enhancing biosensing applications to serving in device applications that demand precise surface engineering.

Materials

All reagents were purchased from Acros Organics, Fisher Scientific, AK Scientific, TCI America, Chem-Impex Inc. or Sigma-Aldrich, and used without further purification unless otherwise noted. DNA was purchased from Integrated DNA Technologies. For bench synthesis, DMF and other solvents were taken from Solvent Purification System or stored over activated 4 Å molecular sieves. Water was obtained from a Milli-Q water purification system. Analytical thin-layer chromatography (TLC) was performed on 0.2 mm silica 60 coated on glass slides with F254 indicator. Flash column chromatography was performed on 40-63 μm silica gel (SiO₂). NMR spectra were recorded using a Varian U500, CB500,or VXR500 spectrometer in the NMR Laboratory, School of Chemical Science, University of Illinois. The data was processed in MestReNova and the spectrum were taken as PDF export directly from the application. Dynamic light scattering analysis was performed on a Malvern Zetasizer system. GFC was performed on Superdex^tm 200 Increase 10/300 GL column in a Shimadzu HPLC. Agarose gel electrophoresis was performed using Biorad horizontal gel electrophoresis apparatus.

Materials for QD synthesis:

n-tetradecylphosphonic acid (TDPA, 97%), oleylamine (OAm, 70%), 1-octadecene (ODE, 90%), trioctylphosphine oxide (TOPO, 99%), trioctylphosphine (TOP, 97%), and oleic acid (OA, 90%) were purchased from Sigma Aldrich. Selenium shot, octadecylphosphonic acid (ODPA, 97%), and cadmium oxide (CdO, 99.999%) were purchased from Alfa Aesar. Tributylphosphine (TBP) was purchased from Strem Chemicals. Bis(trimethylsilyl)sulfide ((TMS)2S) was purchased from Acros Organics. All reagents were used as received unless noted. 1.5 M trioctylphosphine selenide (TOP-Se) was prepared by dissolving 0.15 mmol of selenium shot in 100 mL of TOP under inert atmosphere and stirring vigorously overnight, forming a 1.5 M TOPSe solution. All air sensitive materials were handled in a glove box under dry nitrogen atmosphere with oxygen levels < 0.1 ppm.

Methods

QD (CdSe/CdS core/shell) synthesis.

CdSe cores with 570 nm first absorption peak were synthesized using a previously reported method.31 To summarize, 0.44 mmol (60 mg) of CdO, 0.88 mmol (0.2475g) of TDPA, 9.56 mmol (3.70 g) of TOPO were placed in 25 mL round bottom flask. The solution was degassed for 1 hr at 120oC and heated to 320oC under nitrogen until the CdO dissolved and formed a clear homogenous solution. 1 mL of TOP was injected, and the solution was reheated to 340oC under nitrogen. 1.5 mL of 1.5 M TOP-Se solution was rapidly injected. The cores were then grown further at 280oC to produce cores with the 570 nm for the first absorption feature.

CdS and ZnS shells were deposited on the CdSe core (570 nm absorption peak) using (TMS)₂S. 50 nmol of CdSe cores isolated by repeated precipitations from hexane with acetone. The CdSe QD cores were redispersed in a minimal amount of hexane and loaded in a solvent mixture of 3 mL of OAm and 3 mL of ODE. The reaction solution was degassed under vacuum at 100°C for 1 hr. The amount of each Cd or S precursor was calculated for growing 6 ML of CdS shell on the CdSe QD cores. The Cd precursor (0.39 mmol Cd-oleate, 0.78 mmol of OAm in 5.7 mL of TOP) and the S precursor (0.39 mmol of (TMS)₂S in 6 mL of TOP) were slowly injected (3 mL/hr) to the CdSe core solution simultaneously. The reaction temperature was maintained for 10 min, then increased by 5°C every 5 minutes up to 130°C. The reaction temperature was kept at 130°C for another 80 min. After the precursor injection, the reaction vessel was cooled down to room temperature.

Synthesis of PPFPA.

RAFT polymerization of PFPA was performed as previously reported.⁴⁵ Detailed procedure is described in supplementary information.

Synthesis of PNAS.

RAFT polymerization of PFPA was performed as previously reported.^33,57 Detailed procedure is described in supplementary information.

Aminolysis of PPFPA.

Generally, aminolysis of pre-polymer were performed in the same way as following. Depending on the designed ratio of each monomer in the final polymer, appropriate mixture of amine–compounds were added to PPPFA solution. After the reaction at 50 °C overnight, the crude ¹⁹F NMR was taken to ensure the complete replacement of PFP units. Then the reaction mixture was purified by precipitation with MeOH three to five times, followed by centrifuge filtration with H₂O/MeOH =50%/50% solution. The resulting polymers were stored in MeOH or MeOH/H₂O mixture at 4°C until later use. Then small aliquot of polymer wad dried and lyophilized as needed for the use of QD ligand exchange. As a note, we used mono-Boc protected PEG diamine was used to avoid crosslinking for P3-P5. For these reactions, the reaction solutions were proceeded to deprotection reaction by directly adding 1M HCl in MeOH/H₂O to the reaction solution before the purification. Thereafter, the reaction was neutralized with a drop of 10N NaOH. The resulting polymer was purified by precipitation with excess ethyl ether, followed by further purification using centrifuge filtration to remove pentafluoropenol and any leftover amine monomers. The resulting gel-like polymer was dissolved in H₂O/MeOH =50%/50% solution, and purified further by centrifuge filtration. Detailed procedure is described in supplementary information.

QD Ligand exchange.

Ligand exchange of native QDs with QD-coating polymers (PIL) was performed following the known procedure.^5,26 Briefly, QDs (~1 nmol) were precipitated using hexanes (30 μL), CHCl₃ (30 μL) and acetone or EtOH (150μL). The supernatant containing organic ligands was discarded, and the precipitant was re-dissolved 30-50 μL of CHCl₃. This step was repeated twice to remove the native organic ligands from QD solution. In order to ensure that the imidazole is in its unprotonated form, the PIL was prepared in CHCl3 after basification with NaOH in water/MeOH or with triethylamine in CHCl₃.Then, the QD stock solution was mixed with a solution of PIL (~6 mg) in CHCl₃(30 μL), and stirred for 20 min at RT. Then, gradual amount of MeOH wad added starting from 20 μL of MeOH till the total volume of MeOH reaches ~ 500 μL. QD sample stays clear without any noticeable aggregation or precipitation during MeOH addition. QD samples were then precipitated by the addition of EtOH (40 μL), and excess hexanes (~4mL). The sample was centrifuged at 4,000 g for 2 minutes. The clear supernatant was discarded, and the pellet was dried quickly in vacuo, followed by the addition of PBS (~300 μL, pH 7.4). Prior to any use for experiments, excess ligand was removed by three cycles of centrifuge filtration with MW 50,000 Da (Millipore) with excess amount of PBS.

Agarose gel electrophoresis of QD.

Approximately 1 pmol of QD or QD–conjugated biomolecules (either DNA or protein) were mixed with loading buffer (16% sucrose in ddH2O) and loaded to 1% agarose gel. The gel was ran for 20-30 min at 100V in 1X TAE buffer (40 mM Tris-acetate, 1 mM EDTA, pH 8.3), and visualized using Biorad Gel Doc.

SDS-PAGE of streptavidin and QD-SAv conjugates.

Approximately 1μg of SAv, SAv-DBCO or SAv-QD were prepared with Laminelle solution and 2 μL of 1M DTT. These samples were denatured by heating them in a boiling water for 10 min and loaded onto 4-20% precast polyacrylamide gel (Biorad, CAT# 4561094) along with a protein ladder (Biorad, Precision Plus Protein Dual Color Standards (CAT #1610374). The gel was allowed to run at 60 V for ~30 min, then changed to 120V for ~1.5 hours. Proteins were stained with Coomassie Blue for visualization.

ζ-Potential Measurement.

ζ-Potential was measured on Malvern Zetasizer Nano ZS instrument. Zeta-potentials were measured in 0.1X PBS in a Dip Cell. Values are reported as the average of triplicate runs consisting of 100 scans each.

Dynamic Light Scattering (DLS).

Light-scattering analysis was performed using Malvern Zetasizer Nano ZS instrument. All QD samples were prepared by centrifuge filtration (MWCO = 50 kDa) with 1X PBS to remove any polymer ligands in the storage buffer. The purified QD samples were subjected to measure hydrodynamic size. Hydrodynamic sizes were obtained from a volume-weighted size distribution analysis and reported as the mean of at least triplicate measurements.

Conjugation of QD and DNA.

QD was prepared in 100 mM NaHCO₃ by exchanging the storage solution (1X PBS) to 100 mM NaHCO₃ buffer (pH 8.5). QD concentration was determined in 100 mM NaHCO₃ buffer by measuring UV absorbance at 350 nm, and the amount of QD in the buffer was calculated in moles. 20 eq. of DBCO-functionalized DNA (30 nt) (IDT, /5DBCON/TT GAG TGG ATA CAC TAC CAC CAT TTC CTA T) was then added to QD solution and the mixture in 100 mM NaHCO₃ buffer (pH 8.5) was allowed for reaction at RT for 5 days with gentle shaking. After the reaction, leftover DNA was removed by centrifuge filtration (MWCO = 50 kDa), and the resulting solution was used as QD–DNA conjugate.

Conjugation of QD and streptavidin.

Streptavidin (SAV) (AnaSpec, ~55 kDa) was first washed from a storage buffer containing sodium azide by centrifuge filtration with 1X PBS, and was prepared to 0.5 mg/mL in 1X PBS. SAV in 1X PBS was allowed to react with dibenzocyclooctyne (DBCO) by mixing SAv with DBCO-sulfo-NHS RAFT (Click Chemistry Tools, 2.5 mM in anhydrous DMSO) at a molar ratio of 1:20 for 30 min at RT. Leftover DBCO-sulfo-NHS was removed and the SAV-DBCO conjugate was purified by centrifugal filtration using a filter with MWCO of 30 kDa. SAV-DBCO conjugate was then mixed with QD-N₃ with 1:1 ratio, and the mixture was allowed to react at room temperature overnight. The reaction solution was quenched by adding 50 eq. of 2-azidoacetic acid on ice for 15 min. The resulting solution was washed with 1X PBS using centrifuge filtration (MWCO 100 kDa) for three times, and concentration of QD-SAV was determined by UV absorbance at 350 nm. To characterize the QD–SAV, biotin–DNA (/5Biosg/TT GAG TGG ATA CAC TAC CAC CAT TTC CTA T) was incubated with QD–SAV for 2h at 4 °C. The leftover DNA was removed by centrifuge filtration with MWCO of 50 kDa. The resulting QD-SAV-Biotin-DNA were subjected to electrophoresis.

Conjugation of QD and antibody.

F(ab’)2-Goat anti-Mouse IgG (H+L) Cross-Adsorbed Antibody (Thermofisher, Cat#A24526) was first conjugated with dibenzocyclooctyne (DBCO) by reacting the antibody with DBCO-sulfo-NHS RAFT (Click Chemistry Tools, 2.5 mM in anhydrous DMSO) at a molar ratio of 1:40 for 2 hours on ice with gentle pipetting of the reaction solution every 30 min. Leftover DBCO-sulfo-NHS was removed and the antibody-DBCO conjugate was purified by centrifugal filtration using a filter with MWCO of 10 kDa. DBCO-functionalized antibody was then mixed with QD-N₃ (QD-P6) with 1:1 ratio, and the mixture was allowed to react at room temperature overnight. The reaction solution was quenched by adding 50 eq. of 2-azidoacetic acid on ice for 15 min. The resulting solution was washed with 1X PBS using centrifuge filtration (MWCO 100 kDa) for three times, and concentration of QD-antibody conjugate was determined by UV absorbance at 350 nm. QD-Ab conjugate was prepared along with immunofluorescence procedure so that QD-Ab can be prepared on the day of its use for secondary antibody incubation.

Nonspecific Binding of QDs to Serum.

QD (15μL, ~5 μM) was mixed with 100% FBS or 1× PBS (55 μL) to final concentrations of FBS of ~80% (v/v). The mixture was incubated for 4h at 37 °C. Then, the QD size distribution was analyzed using gel filtration chromatography (GFC). The mixture was injected into a Superdex 200 Increase 5/150 GL (GE Healthcare) on a Shimadzu HPLC with an in-line degasser, autosampler, diode array detector, and fluorescence detector. 1X PBS (pH 7.4) was used as the mobile phase with a flow rate of 0.75 mL/min. Fluorescence detected at 609 nm with excitation at 250 nm.

Nonspecific Binding of QDs to U2OS cells.

Cells were seeded in μ-Slide 8 well (ibidi, Cat # 80826) following the manufacturer’s recommendation. Before the treatment of QD, cells were rinsed with 1X PBS three times. QDs at 500 nM were treated to cells at 4 °C for 10 min in McCoy media without FBS. QDs were removed and the cells were rinsed again with McCoy media without FBS , then imaged in the same cell culture media using widefield microscopy (Zeiss Axiovert 200M).

Immunofluorescence.

The cells were rinsed with 1X PBS three times. Then, cells were fixed and permeabilized with chilled methanol for 10 min at −20 °C. After rinsing the cells with 1X PBS three times, the cells were then blocked with 5% BSA + 0.05% of Superblock (Thermo Scientific Cat #37515) for 10 min. Then, the cells were incubated with primary antibody (monoclonal Anti-alpha-Tubulin antibody produced in mouse, Millipore Sigma, T9026) in 5% BSA + 0.05% of Superblock (1:500) for 1 hour at room temperature. Cells were washed three times with 5% BSA + 0.05% of Superblock (Thermo Scientific Cat #37515) in 1X PBS for 10 min of rocking at each washing. Secondary antibody (either Alexa568-conjugate (Goat anti-Mouse IgG Alexa Fluor568, Invitrogen Cat#A-11031) or QD-antibody conjugate) was incubated for 1 hour at room temperature (1:1000 Alexa568-conjugate or 20 nM of QD-antibody conjugate) in the same blocking buffer. Then, the cells were washed with the blocking buffer three times for 10 min per each wash. Finally, cells were stained with DAPI for 5 min, and rinsed with 1X PBS three times before being mounted on a coverslip. When mounting a coverslip on a glass plate, ProLong Diamond Antifade Mountant (ThermoFisher, CAT #P36965) was placed for all samples for imaging using widefield microscopy (Zeiss Axiovert 200M).