Naked-Eye Thiol Analyte Detection via Self-Propagating, Amplified Reaction Cycle

Abstract

We present an approach for detecting thiol analytes through a self-propagating amplification cycle that triggers the macroscopic degradation of a hydrogel scaffold. The amplification system consists of an allylic phosphonium salt that upon reaction with the thiol analyte releases a phosphine, which reduces a disulfide to form two thiols, closing the cycle and ultimately resulting in exponential amplification of the thiol input. When integrated in a disulfide cross-linked hydrogel, the amplification process leads to physical degradation of the hydrogel in response to thiol analytes. We developed a numerical model to predict the behavior of the amplification cycle in response to varying concentrations of thiol triggers and validated it with experimental data. Using this system, we were able to detect multiple thiol analytes, including a small molecule probe, glutathione, DNA, and a protein, at concentrations ranging from 132 to 0.132 μM. In addition, we discovered that the self-propagating amplification cycle could be initiated by force-generated molecular scission, enabling damage-triggered hydrogel destruction.

Figure 1

Schematics of the signal amplification system (SAS), its conditions, components, and material degradation mechanism. (a) Generic SAS, consisting of nucleophilic substitution and disulfide reduction reaction. (b) Chemical structure of allylic phosphonium salt 1, substitution product 2, free phosphine (TPPTS), disulfide 3, thiol 4, and oxidized phosphine (OTPPTS). Specifically, TPPTS is liberated from 1 upon SH-signal (compound 4), which initiates disulfide reduction. The reduction reaction produces additional thiols, which themselves continue to liberate more TPPTS. This cascade results in an amplification of the starting thiol signal. (c) BAC cross-linked DMA hydrogels used for naked-eye SH-analyte detection. Upon signal addition, the liberated TPPTS inside the hydrogel matrix reduces disulfide cross-links, which themselves liberate more TPPTS. This results into a signal-triggered self-propagating amplification and ultimately into the degradation of the hydrogel material.

Figure 2

Kinetic control over TPPTS release. (a) Nucleophilic substitution reaction of compound 1 with S-, N-, or O-terminal nucleophiles (l-glutathione, N-acetyl cysteine, l-proline, l-phenylalanine, and p-nitrophenol), forming the nucleophilic substitution product and releasing TPPTS. (b) Reaction of compound 1 with a range of 4 (SH-signal) concentrations, forming 2-(acetylamino)ethanethiol (2) and TPPTS. TPPTS concentration vs time for a range of 4 concentrations. (c) SH-signal input (%) and TPPTS output (%) diagram, showing the control of TPPTS release upon the addition of 4. Reactions were monitored by UV–vis at 260 nm and performed in duplicate. Conditions: phosphate buffer (0.1 M, pH 7.6), RT, 20 h, 2 mM of 1 and indicated amounts of 4. (d) NMR reactivity study using 0.067 mM of 1 (1.0 equiv) and 0.20 equiv of a range of different nucleophiles (indicated in the figure) in aqueous buffer (2:8 D₂O: phosphate buffer (0.1 M, pH = 7.6)) at 25 °C. Error bars represent standard deviation from duplicate runs. Solid lines represent the k-value model fit to the experimental data.

Figure 3

Nucleophilic substitution and disulfide reduction kinetics. (a) Schematic representation and full reaction pathway overview for nucleophilic substitution (I) and disulfide reduction (II) in the amplification system. (b) SH-triggered substitution kinetics of compound 1 (2.0 mM), measured by the appearance of TPPTS from UV–vis at 260 nm. (c) Disulfide reduction kinetics for 3 (12.0 mM) at varying concentrations of TPPTS, measured by the disappearance of TPPTS from UV–vis at 300 nm. All reactions were measured in 0.1 M phosphate buffer (pH = 7.6) at 25 °C. Representative samples from duplicate runs (green line). Solid black lines correspond to the model fits to each condition. Statistical evaluation between the kinetic model and experimental data can be found in Supporting Figures 13–16.

Figure 4

Kinetic experiments of the signal amplification system and model predictions for species variation. (a) Schematic representation and full reaction pathway overview for nucleophilic substitution and disulfide reduction in the amplification system. (b) UV–vis kinetic experiments using compound 4 as SH-trigger, at concentrations of 0.95 mM (10%), 1.40 mM (15%), and 2.30 mM (25%). Conditions: 9.0 mM of compound 1 with 13.5 mM of 3 in phosphate buffer (pH = 7.6) at 25 °C. (c) NMR kinetic experiments using 13.5 mM of 1 (1.0 equiv), 1.5 equiv of disulfide 3, and 0.05 equiv of 4 in aqueous buffer (2:8 D₂O: phosphate buffer (0.1 M, pH = 7.6)) at 25 °C. Error bars represent standard deviation from duplicate runs. Solid lines correspond to the model fits to each varying condition or species. (d) Schematic representation of the signal amplification system output using 0.05 equiv of 4.

Figure 5

Time lapse photographs of hydrogel degradation using the amplification system triggered by SH-analytes. (a) Control gels with (1) no additives, (2) with 1.5 equiv of 1, (3) 1.5 equiv of TPPTS, and (4) 1.5 equiv of 1 and 5% (0.05 equiv) of 4. (b) Gels with 1.5 equiv of 1 and SH-trigger addition of (1) 1.0% (0.01 equiv) of l-glutathione, (2) 0.01% (0.0001 equiv) of bovine serum albumin, and (3) 0.001% (0.00001 equiv) of thiol-functionalized DNA. Conditions: gels were submerged in 1.5 mL of phosphate buffer (0.1 M, pH = 7.6). All measurements were done in duplicate (see Supporting Figures 17 and 18).

Figure 6

Time lapse photographs of hydrogel degradation using the self-amplification system triggered by damage (cut). Control gels with (1) no additives and 1× horizontal cut, (2) 1.5 equiv of 1 and 1× horizontal cut, and (3) 1.5 equiv of 1 and 2× cut (1× horizontal and 1× vertical). Conditions: gels were submerged in 1.5 mL of phosphate buffer (0.1 M, pH = 7.6). All measurements were done in duplicate (see Supporting Figure 19).