Near-infrared Absorption and Emission Probes with Optimal Connection Bridges for Live Monitoring of NAD(P)H Dynamics in Living Systems

Abstract

Two NAD(P)H-biosensing probes consisting of 1,3,3-trimethyl-3H-indolium and 3-quinolinium acceptors, linked by thiophene, A, and 3,4-ethylenedioxythiophene, B, bridges are detailed. We synthesized probes C and D, replacing the thiophene connection in probe A with phenyl and 2,1,3-benzothiadiazole units, respectively. Probe E was prepared by substituting probe A's 3-quinolinium unit with a 1-methylquinoxalin-1-ium unit. Solutions are non-fluorescent but in the presence of NADH, exhibit near-infrared fluorescence at 742.1 nm and 727.2 nm for probes A and B, respectively, and generate absorbance signals at 690.6 nm and 685.9 nm. In contrast, probes C and D displayed pronounced interference from NADH fluorescence at 450 nm, whereas probe E exhibited minimal fluorescence alterations in response to NAD(P)H. Pre-treatment of A549 cells with glucose in the presence of probe A led to a significant increase in fluorescence intensity. Additionally, subjecting probe A to lactate and pyruvate molecules resulted in opposite changes in NAD(P)H levels, with lactate causing a substantial increase in fluorescence intensity, conversely, pyruvate resulted in a sharp decrease. Treatment of A549 cells with varying concentrations of the drugs cisplatin, gemcitabine, and camptothecin (5, 10, and 20 μM) led to a concentration-dependent increase in intracellular fluorescence intensity, signifying a rise in NAD(P)H levels. Finally, fruit fly larvae were treated with different concentrations of NADH and cisplatin illustrating applicability to live organisms. The results demonstrated a direct correlation between fluorescence intensity and the concentration of NADH and cisplatin, respectively, further confirming the efficacy of probe A in sensing changes in NAD(P)H levels within a whole organism.

Graphical Abstract

1. Introduction

The redox pair nicotinamide adenine dinucleotide, comprising the reduced NADH and oxidized NAD⁺ forms, is an essential coenzyme involved in numerous reduction-oxidation transformations within eukaryotic cells.[1, 2] Similarly, NADPH, representing the reduced version of the coenzyme nicotinamide adenine dinucleotide phosphate, plays a vital role in various anabolic pathways and antioxidant defense mechanisms.[3] These coenzymes are critical for cellular redox balance and participate in various biological processes.[4-7] Given the significant role of NADH and NADPH in biological systems,[4-7] accurately determining their concentrations is essential to unravel physiological processes and disease pathways, aiding in disease diagnosis.[8] Several analytical approaches have been established for quantifying NADH and NADPH levels, with fluorescence methods coupled with fluorescence confocal microscopes offering unique advantages, such as real-time visualization in live cells, sensitivity, selectivity, and spatiotemporal resolution, Table S1.[8-18] While intrinsic NADH and NADPH emissions have been employed for quantification and visualization, optical interference from other biological molecules can be challenging due to weak emissions at specific wavelengths and short absorption peaks.[19-21] To overcome these limitations, fluorescent probes with near-infrared emissions from 650 nm to 850 nm have been developed that reduce optical interference and enhances tissue penetration.[10, 15, 17, 22] Recently, a cyanine-based probe was documented to sense NAD(P)H with remarkable sensitivity (Scheme 1).[23] However, this probe has limited detection capabilities. Specifically, it demonstrates a fluorescence activation response at 561 nm when NADH concentrations are below 10 μM. Unfortunately, at NADH levels above 10 μM, the fluorescence is quenched, resulting in a decreased emission intensity.[23-25] While this probe shows initial promise for sensing NADH, its narrow effective detection range and fluorescence quenching behavior restrict its usefulness.[23-25] Continued development of NAD(P)H-responsive probes is needed to achieve robust turn-on fluorescence with negligible background and no saturation effects across the necessary detection range. More effective probes would enable researchers to study NADH signaling under diverse metabolic conditions.

Scheme 1.

NAD(P)H sensing mechanism of the probes.

Overcoming the limitations of current cyanine-based NADH probes remains an important goal for advancing redox and metabolic bioimaging. In this study, we report on two near-infrared NAD(P)H-sensing fluorescent probes, A and B, built upon cyanine dyes bearing 3-quinolinium units (Scheme 1). Additionally, we synthesized probes C and D, wherein phenyl and 2,1,3-benzothiadiazole units replace the thiophene connection in probe A. We also prepared probe E by substituting the 3-quinolinium unit with a 1-methylquinoxalin-1-ium unit from probe A. Probes A and B display negligible fluorescence without the presence of NAD(P)H, and upon their gradual addition, new fluorescence maxima at 742.1 nm and 727.2 nm were observed, accompanied by new absorption maxima at 690.6 nm and 685.9 nm. Conversely, probes C to E displayed no significant fluorescence response to NAD(P)H, indicating that the choice of connection bridges plays a critical role in the probes' ability to detect NAD(P)H dynamics. Probe A offers a valuable tool for studying NAD(P)H dynamics in live cells and whole organisms and provides reduced background interference. Using probe A, we have unveiled how major nutrients, metabolites, and anticancer drugs impact NAD(P)H levels in cells, and how an anticancer drug impacts NAD(P)H levels in fruit fly larvae.

2. Experimental

2.1. Instrumentation.

The investigation involved utilizing a 500 MHz Bruker NMR Spectrometer, for acquiring ¹H, ¹⁹F and ¹³C NMR spectra. An absorption spectrum of each probe was collected using a PerkinElmer Lambda 35 UV/vis spectrometer, while a Jobin Yvon Fluoromax-4 spectrofluorometer was employed to record the probe's emission spectra. A confocal fluorescence cellular imaging experiment was conducted using an Olympus IX 81 confocal fluorescence microscope (Olympus America Inc.). Acquisition and processing of the cellular image were performed with an Olympus FV10-ASW 3.1 viewer, ImageJ, and Photoshop 9.0.

2.2. Synthesis of fluorescent probes

2.2.1. Synthesis of probe A

Compound 7 (100.0 mg, 0.18 mmol) was dissolved in 10 mL of dichloromethane, and methyl trifluoromethanesulfonate (33.14 mg, 0.20 mmol) was added in a 50 mL round-bottom flask. The reaction mixture was stirred for 6 hours at room temperature under a nitrogen atmosphere and monitored by TLC. Afterward, the solvent was removed under reduced pressure, and the residue was treated with 20 mL of cold water, followed by extraction with dichloromethane (10 x 3 times). The combined organic layers were dried over sodium sulfate, and the solvent was evaporated under reduced pressure to obtain the crude product. Purification was performed using column chromatography with a dichloromethane and methanol eluent (9:1), resulting in a 45% yield of probe A. ¹H NMR (500 MHz, DMSO-d₆), δ (ppm): 1.82 (6H, s), 4.15 (3H, s), 4.73 (3H, s), 7.43 (1H, d, J = 15 Hz), 7.64 (2H, m), 7.89 (1H, m), 7.93 (1H, m), 8.11 (1H, m), 8.18 (1H, d, J = 5.0 Hz), 8.30 (1H, m), 8.35 (1H, d, J = 5 Hz), 8.52 (2H, m), 8.71 (1H, d, J = 15 Hz), 9.59 (1H, s), 10.09 (1H, s). ¹³C NMR (125 MHz, DMSO-d₆), δ (ppm): 25.73, 34.87, 46.14, 52.66, 113.15, 115.73, 119.79, 122.49, 123.44, 127.33, 129.59, 129.64, 130.01, 130.05, 131.23, 136.27, 137.96, 141.93, 142.38, 142.66, 144.15, 144.56, 144.81, 148.98, 181.38.

2.3. Cell culture, cell, and fruit fly imaging

Cell culture, cell and fruit fly imaging were performed in accordance with our previously published protocols [25, 27, 37]. Comprehensive details can be accessed in the supporting information section.

2.4. Theoretical Calculations

Models of probes were computed using Gaussian 16[26] and density functional theory (DFT), with the APFD functional and basis sets at the 6-311+g(d,p) for optimization of the geometry in a Polarizable Continuum Model (PCM) of water. Upon confirming the lack of imaginary frequencies, the CAM-B3LYP/6-311+g(d) combination was used in a TD-DFT calculation to calculate the absorption energies. Results were interpreted using GaussView 6[27] for all data and figures. Coordinates are listed in supporting information.

3. Results and Discussion

3.1. Probe synthesis

We previously developed thiophene-based cyanine dyes incorporating electron-withdrawing formyl and 3-quinolinium acceptors, specifically designed for NAD(P)H sensing applications.[25] While exhibiting a unique turn-on deep-red fluorescence response to NAD(P)H, these probes had relatively slow reaction times.[25] We hypothesized that replacing the formyl group with a stronger 1,3,3-trimethyl-3H-indolium acceptor would shift the emissions into the near-infrared spectrum and significantly improve response kinetics to NAD(P)H. To validate our hypothesis, we synthesized two cyanine dyes (designated as probes A and B) via condensation reactions of 5-(quinolin-3-yl)thiophene-2-carbaldehyde (4) and 7-(quinolin-3-yl)-2,3-dihydrothieno[3,4-b][1,4]dioxine-5-carbaldehyde (10) with 1,2,3,3-tetramethyl-3H-indol-1-ium trifluoromethanesulfonate (6), yielding compounds 7 and 11 (Scheme 2), and through further methylation of compounds 7 and 11. Key precursors 7and 11 were obtained through a palladium-catalyzed Suzuki reaction of 3-quinolineboronic acid (1) with 5-bromothiophene-2-carbaldehyde (3) and 7-bromo-2,3-dihydrothieno[3,4-b][1,4]dioxine-5-carbaldehyde (9), respectively (Scheme 2).[25] Probes C and D were synthesized by replacing compound 3 with compounds 12 and 13, respectively, in the probe A protocol. Probe E was prepared following the probe A method but starting from 2-quinoxalinyl boronic acid (2) instead of (quinolin-3-yl)boronic acid (1). By systematically modifying the acceptor strength and connection structures, we aimed to establish a robust structure-activity relationship to guide further optimization of NAD(P)H-responsive near-infrared fluorescent probes. The intermediates and probes underwent comprehensive characterization using both NMR and mass spectrometry techniques, as illustrated in Figures S1-S21. The ¹⁹F NMR spectrum of probe A unequivocally substantiates the existence of two trifluoromethane sulfonate anions within probe A's structure (Figure S5).

Scheme 2.

Synthetic method to make near-infrared cyanine dyes (probes A-E) for NAD(P)H sensing.

3.2. Optical sensing of NADH

In the absence of NADH, probes A and B reveal primary absorption peaks at 460.4 nm and 445 nm, respectively. Upon gradual addition of NADH in pH 7.4 phosphate buffer with 5% DMSO, new near-infrared absorption bands emerge at 690.6 nm for probe A and 685.9 nm for probe B, intensifying with increasing NADH concentration. This arises from NADH-mediated reduction of the 3-quinolinium acceptor to an electron-bestowing 1-methyl-1,4-dihydroquinoline donor, generating distinct donor-π-acceptor cyanine dyes (Scheme 1 and Figures 1 and S22).

Figure 1.

Absorption (on the left) and fluorescence (on the right) profiles of 10 μM probe A (in pH 7.4 phosphate buffer supplemented with 5% DMSO) with excitation and emission slit widths set to 5 nm. The spectra were measured under both absence and presence conditions of various NADH concentrations following incubation for 75 minutes and 680 nm was used as an excitation wavelength for fluorescence profile of probe A.

In the absence of NADH, probes A and B show minimal fluorescence when excited at their absorbance maxima because of the presence of two potent electron-accepting groups - the 1,3,3-trimethyl-3H-indolium and 3-quinolinium acceptors (Figures 1 and S22). However, upon gradual addition of NADH in pH 7.4 phosphate buffer with 5% DMSO, both probes exhibit a dramatic fluorescence turn-on in the near-infrared region, with emission peaks at 742.1 nm for probe A and 727.2 nm for probe B (Figures 1 and S22). This significant enhancement of near-infrared fluorescence indicates that NADH induces the reduction of the 3-quinolinium acceptor to an electron-supplying 1-methyl-1,4-dihydroquinoline donor, generating the anticipated donor-π-acceptor cyanine dyes. Reaction products, i.e., probes AH and BH were verified by high-resolution mass spectrometer (Figures S19 and S21). Importantly, probes A and B demonstrate rapid response times to NADH. Probe A shows complete activation within 75 minutes, while probe B is fully activated within 120 minutes (Figure S23). This is a considerable improvement compared to our previous cyanine dyes containing weak formyl acceptors, which required 180 minutes to elicit a fluorescence response.[25] The faster response kinetics validate our design strategy of utilizing the strong 1,3,3-trimethyl-3H-indolium acceptor to tune the optical properties and reactivity. The longer response time of probe B is attributed to the electron-abundant 3,4-ethylenedioxythiophene uniting element between the two acceptor moieties, which reduces its reactivity compared to the thiophene bridge in probe A.

Probe C, which incorporates a phenyle connection bridge, exhibits absorbance at 360 nm and weak fluorescence at 450 nm. Upon addition of NADH, both absorption at 600 nm and fluorescence at 450 nm increase, potentially due to NADH fluorescence (Figure S24). Similarly, probe D, with a 2,1,3-benzothiadiazole connection bridge, demonstrates a comparable response to NADH, also displaying interference from NADH fluorescence at 450 nm (Figure S25). In contrast, probe E, featuring a 1-methylquinoxalin-1-ium sensing unit, does not exhibit any fluorescence response to NADH (Figure S26). These findings underscore the significance of optimizing both probe connection bridges and sensing elements to achieve sensitive NAD(P)H detection.

3.3. Theoretical calculations

The optimized geometries of the probes were obtained as described previously[28] using the program Gaussian 16[26] with the APFD functional[29] and with atoms defined by the 6-311G(d,p) basis set and in water as defined by the polarizable continuum model (SCRF) option. Electronic transitions, i.e., TD-DFT were then calculated for each probe using the CAM-B3LYP[30] functional, in water as this method was previously determined[31] to result in good agreement with experimental results in complexes containing delocalized excited states. The optimized geometries indicated that the indole and central thiophene moieties were coplanar in all probes, Figures S38(A), S42 (AH), S45 (B), and S48 (BH). However, in probes A and B, the planes of the quinoline moieties were twisted in relation to the plane of the central thiophene at angles of 29.7 degrees and 20.9 degrees, respectively. It appears that the AH and BH probes are planar because the adjacent methylene group has its hydrogen atoms symmetrically displaced with respect to the adjacent sulfur atom in the central thiophene.

The primary excitation mode for each probe appears to primarily consist of a transition between HOMO (the highest occupied molecular orbital) and LUMO (the lowest unoccupied molecular orbital), which was used to model the current density charts seen in Figure 2. The data collected for the first excited state of each probe allows for speculation into the accuracy of these methods in predicting the spectral properties of these probes. The differences between the theoretical and experimental wavelengths are as follows: 3.39 nm, A; 124.69 nm, AH; 18.78 nm, B; 138.22 nm, BH. This suggests that these computational methods are very precise in predicting the spectral properties of the non-hydrogenated probes A and B but begin to drift away from high agreement with respect to the hydrogenated probes AH and BH.

Figure 2:

Mapped current density charts for probes A, AH, B, and BH showing the flow of electron density as the molecular orbitals transition from the HOMO to the LUMO during the first excited state. H* and L* refer to the HOMO and LUMO orbitals.

As is apparent in Figure 2, there is a shift in the nature of the transition from the central thiophene moieties in probes A and B to the indole section compared to probes AH and BH where the transition occurs from the more electron rich quinoline moieties to the central thiophene and indole sections and is more delocalized due to the planarity of the structure.

As is evident in the contour and relief (shows variations of electron density above and below a plane) maps, Table S12 for the probes constructed on the plane of the 3-quinolinium moieties, probes A and B attain extended conjugation after hydride addition, judging by the increased electron density in the plane. Interestingly, the images for probes C and D indicate that they could attain increased planarity upon hydride addition, so it is puzzling that these probes displayed reduced reactivity upon addition of NADH. Probe E was calculated to be already planar so nothing would be gained in this case by hydride addition. Examination of the values of the q(A) charges [32] revealed the following values for the carbon atom where the hydride could bind for probes A-D and for the N atom in probe E: A, 0.0169; B, 0.0102; C, 0.0132; D, 0.0235; E, −1.1233. By this measure, probe E with a negative q(A) would clearly repel a hydride but reactions should be possible with probes C and D. Perhaps resistance to rehybridization to achieve planarity is the issue here.

3.4. The selectivity and photostability of the probes

We evaluated the sensing specificity of probes A and B by comparing their fluorescence response to NADH and NAD(P)H against a wide range of potentially interfering analytes. As shown in Figures S28-S29, probes A and B exhibited minimal fluorescence changes when exposed to any of these potential interferents at biologically relevant concentrations. In contrast, they demonstrated a remarkable increase in fluorescence signal exclusively with the presence of NADH and NADPH. This excellent specificity confirms that probes A and B can reliably detect NAD(P)H levels even in complex biological samples containing diverse analytes.

Probe A exhibits a distinctive turn-on fluorescence response to SO₂, emitting prominently at a shorter wavelength of 502 nm, with no fluorescence response to SO₂ observed in the near-infrared region under excitation at 680 nm (refer to Figure S30 and Figure S31). The unique fluorescence response results from a well-documented addition reaction of SO₂ to the double bond within probe A, disrupting its π-conjugation and causing fluorescence emission at 502 nm, as depicted in Scheme S2. This dual-response capability of probe A allows for the separate detection of NADH and SO₂ in distinct fluorescence channels within the near-infrared and visible regions, respectively. While this paper primarily focuses on the sensitive detection of NAD(P)H in live cells, the exploration of SO₂ detection serves as a secondary aspect, falling outside the scope of the current research.

Probe A exhibits its optimal sensitivity to NADH within the pH range of 7.0 to 8.2. Any deviation from this range, such as a shift to pH 5.0 or an increase to pH 9.0, induces a noticeable fluorescence quenching effect (Figure S32). The consistent fluorescence intensity of probe A in the presence of NADH within the pH range of 7.0 to 8.2 facilitates precise determination of NAD(P)H levels in mitochondria under various chemical treatments. This accuracy is achieved without interference from pH effects, given that mitochondria inherently maintain a pH of 8.0.

To thoroughly characterize the photostability of probes A and B, we performed rigorous time-course fluorescence measurements under sustained illumination. Specifically, probe A was continuously irradiated at 680 nm excitation while submerged within a pH 7.4 PBS buffer solution containing 35 μM NADH. Similarly, probe B was constantly exposed to 633 nm excitation in a buffer containing 50 μM NADH. Fluorescence intensity was monitored over 120 minutes under these harsh illumination conditions (Figures S33-S34). Remarkably, both probes displayed outstanding photostability with less than a 5% decrease in fluorescence even after 2 hours of continuous irradiation. This minor reduction in signal indicates that probes A and B resist photobleaching remarkably well. Compared to traditional fluorophores that rapidly photobleach, the exceptional photostability of these probes enables reliable quantitative bioimaging over timescales from minutes to hours without signal deterioration. This characteristic makes probes A and B extremely well-suited for practical, reproducible biological imaging applications, both in vitro and in vivo.

3.5. Probe cytotoxicity

Prior to studies in living systems, it is imperative to rigorously assess biocompatibility and cytotoxicity. Therefore, we assessed the cytotoxicity of probes A and B using the MTT assay in the cell line A549 (Figure S35).[24, 25, 33-39] Cells were incubated with varying concentrations of probe A or probe B, ranging from 0 to 50 μM. Both probes showed minimal toxicity, with cells maintaining high viability even at the highest dose of 50 μM. This excellent cytocompatibility confirms the low cytotoxicity of probes A and B, validating their safety for bioimaging in vivo. Additionally, probe A's rapid fluorescence response to NADH further heightens its potential for investigating biological dynamics.

3.6. Fluorescence imaging applications

We conducted fluorescence imaging experiments in A549 cells using a step-wise protocol to evaluate the intracellular response kinetics of probe A. A549 cells were pretreated as described in supporting information. Fluorescence microscopy images were acquired at regular intervals during exposure to probe A (Figure S36). Intriguingly, we noted that the fluorescence intensity within the cells reached a plateau within the first 30 minutes of incubation with probe A. No further increase in fluorescence was detected with longer incubation times up to 60 minutes. This indicates a significantly faster activation of probe A inside living cells compared to the response time in solution. While requiring 75 minutes for full activation in cuvette measurements, probe A showed complete fluorescence turn-on within 30 minutes inside cells. This exciting discovery highlights the rapid intracellular response kinetics of probe A, validating its potential for time-sensitive bioimaging applications. By enabling real-time visualization of NADH fluctuations on short timescales, probe A could provide unprecedented insights into cell metabolism and redox biology dynamics. The efficient intracellular behavior further strengthens the position of probe A as an optimal probe for elucidating biological processes through rapid, selective NADH imaging in living systems.

To confirm that probe A specifically responds to intracellular NADH levels, control, and NADH-dose experiments were conducted in A549 cells. As part of the control group, cells underwent a 30-minute incubation with 10 μM of probe A in a glucose-absent DMEM medium, without prior NADH pre-treatment. Fluorescence imaging revealed minimal fluorescence under these basal conditions (Figure 3), indicating low intrinsic NADH levels. This demonstrates the lack of probe A activation in the absence of elevated NADH. In contrast, cells pre-treated with varying concentrations of NADH (5, 15, 30 μM) for 30 minutes in glucose-absent DMEM exhibited a dose-dependent increase in fluorescence when subsequently incubated with probe A (Figure S37). Higher NADH pre-treatment led to greater fluorescence intensity, correlating precisely with the extracellular NADH levels. This concentration-dependent response confirms that probe A specifically detects changes in intracellular NADH, with higher fluorescence corresponding to elevated NADH content.

Figure 3.

Emission images of A549 cells under diverse experimental conditions: (1) Having first incubated the cells with graded glucose amounts (0, 5, 10, 20 mM) in glucose-devoid DMEM for 30 minutes, this was accompanied by a further 30 minute incubation period with 10 μM of probe A in DMEM medium without glucose, and (2) Control image of A549 cells incubated with 10 μM of probe A in glucose-absent DMEM medium for 30 minutes. The emission images were acquired from 700 nm to 800 nm under excitation at 633 nm.

Our key objective was to investigate how metabolic perturbations in glucose availability impact intracellular NADH levels, as glucose is converted to pyruvate while generating cytosolic NADH.[40, 41] We hypothesized that increasing extracellular glucose would activate glycolysis and enhance NADH generation. We carried out a glucose dose experiment in A549 cells a described in supporting information. As predicted, we detected a successive intensification in fluorescence that correlated with increased glucose pre-incubation, reflecting elevated NADH levels (Figure 3). Cells exhibited minimal baseline fluorescence without glucose, indicating low intrinsic NADH. Fluorescence steadily rose in a dose-dependent manner with 5 mM, 10 mM, and 20 mM glucose pre-treatment. This suggests that increased extracellular glucose availability drove glycolytic flux and NADH production. Overall, these findings provide valuable insights into the dynamic relationship between nutrient availability and intracellular redox states.

We hypothesized that the two positive charges on probe A would confer specific mitochondrial targeting via electrostatic interactions involving the mitochondrial membrane potential characterized by negative charge. To test this, we performed co-localization experiments in A549 cells using the mitochondrial-specific dye MitoView 405. Cells were pretreated with 25 mM glucose in glucose-absent DMEM for 30 minutes to stimulate mitochondrial NADH production. They were then co-incubated with 10 μM probe A and 10 μM MitoView 405 for 30 minutes. Quantitative analysis using the Pearson correlation coefficient revealed a value of 0.961, indicating strong colocalization between probe A and MitoView 405 (Figure S38). This high degree of correlation validates our hypothesis that probe A selectively stains mitochondria due to its cationic nature, which drives specific accumulation in the negatively charged mitochondria. By enabling real-time imaging of NADH fluctuations in mitochondria, probe A can elucidate mechanisms involved in mitochondrial dysfunction.

We sought to elucidate how exogenous pyruvate and lactate impact glycolytic flux and intracellular NADH levels. Cancer cells preferentially convert pyruvate to lactate during aerobic glycolysis, which consumes NADH.[42-44] We hypothesized that pyruvate and lactate would have opposite effects on cytosolic NADH that could be dynamically monitored using probe A. A549 cells were pretreated with varying concentrations of pyruvate or lactate before incubation with probe A. Strikingly, 5 mM pyruvate pre-treatment caused a sharp decrease in probe A fluorescence compared to control cells (Figure 4). This implies rapid NADH consumption by lactate dehydrogenase during pyruvate-to-lactate conversion.[42-44] Conversely, 10 mM lactate pre-treatment significantly increased probe A fluorescence, indicating elevated NADH levels. This suggests that exogenous lactate augments intracellular NADH production.[42-44] Remarkably, combined pre-treatment with 10 mM lactate and 5 mM pyruvate elicited higher probe A fluorescence than control cells. This synergistic effect reveals the complex interplay between lactate and pyruvate in modulating glycolytic NADH. Our work highlights the tremendous value of visualizing NADH dynamics using probe A for elucidating complex metabolic processes in cancer and beyond. Understanding these mechanisms is essential, as dysregulated NADH signaling underlies various diseases.

Figure 4.

Emission images of A549 cells exposed to diverse pre-treatment conditions, encompassing lactate, pyruvate, or a blend of both, in glucose-absent DMEM medium, over a 30-minute period. Afterward, the cells were co-incubated with 10 μM of probe A in glucose-absent DMEM medium for an extra 30 minutes. The emission images were obtained employing excitation at 633 nm and detecting emission from 700 nm to 800 nm.

Cisplatin is a widely used and effective chemotherapeutic drug primarily used to treat various types of cancers by inducing DNA damage and oxidative stress in cancer cells.[45, 46] We investigated how cisplatin impacts NAD(P)H metabolism in A549 lung cancer cells and experimental details are in supporting information. We observed a dose-dependent increase in fluorescence with higher cisplatin concentrations (Figure 5), indicating elevated NAD(P)H levels. In response to cisplatin treatment, A549 cells initiated an adaptive response to counteract the drug's cytotoxic effects and promote cell survival. A prominent adaptive response observed in cancer cells treated with cisplatin is the upregulation of cellular metabolism, including glycolysis and the tricarboxylic acid (TCA) cycle.[47-49] Cisplatin has been shown to enhance glucose uptake and augment glycolytic flux in cancer cells. The increased glycolysis leads to elevated pyruvate production, which is subsequently converted to lactate through the action of lactate dehydrogenase, effectively regenerating NAD⁺ from NADH.[47] This process results in a decrease in the intracellular NADH/NAD⁺ ratio. Interestingly, despite the increase in glycolytic activity and lactate production, the overall cellular NAD(P)H levels tend to rise when cells are treated with cisplatin. This phenomenon can be attributed to the activation of various cellular stress responses and redox signaling pathways triggered by cisplatin-induced DNA damage. Notably, these pathways involve the activation of NAD(P)H-dependent enzymes, including enzymes like poly(ADP-ribose) polymerases (PARPs), which utilize NAD⁺ during DNA repair processes.[47-49] Additionally, cisplatin-induced cellular stress can activate NAD⁺ biosynthesis pathways, such as the recycling route and the newly generated synthesis pathway. These pathways entail multiple enzymatic steps that convert nicotinamide (NAM) and nicotinamide riboside (NR) into NAD⁺. The increased synthesis of NAD⁺ helps replenish the cellular NAD(P)H pool. As the concentration of cisplatin increases, the extent of DNA damage and cellular stress escalates, leading to a more robust activation of these metabolic and redox pathways.[48, 49] Consequently, the increase in NAD(P)H levels becomes more pronounced with higher cisplatin concentrations. In summary, cisplatin induces multifaceted metabolic adaptations, including heightened glycolysis, DNA repair, and NAD⁺ synthesis, which culminate in elevated NAD(P)H. This aids cancer cell survival and may promote drug resistance. Probe A provided unique real-time insights into cisplatin-induced alterations in NAD(P)H metabolism in live cells.

Figure 5.

Emission images of A549 cells exposed to different cisplatin concentrations (0, 5, 10, and 20 μM) during a 2-hour pre-treatment in glucose-absent DMEM medium. Afterward, the cells were treated with 10 μM of probe A for 30 minutes in the same glucose-absent DMEM medium. The emission images were taken within the spectral range of 700 nm to 800 nm, utilizing excitation at 633 nm.

Gemcitabine is a chemotherapeutic agent that interferes with DNA synthesis and repair in cancer cells.[50, 51] We investigated how gemcitabine impacts NAD(P)H metabolism in A549 cells using probe A. Cells underwent pre-treatment with different gemcitabine concentrations as detailed in supporting information. Much like cisplatin, we noted a dosage-related increase in fluorescence associated with increased gemcitabine concentrations (Figure 6), pointing to heightened NAD(P)H levels. This rise in NAD(P)H may be attributed to metabolic adaptations triggered by gemcitabine's disruption of DNA replication and damage.[52, 53] Gemcitabine induces cellular stress responses involving redox signaling and activation of NAD(P)H-dependent enzymes to repair DNA damage.[52, 53] Further, gemcitabine enhances glycolytic flux, reminiscent of the Warburg effect,[54] increasing lactate production and NADH consumption. However, overall NAD(P)H levels still increase, likely due to overriding activation of other NAD(P)H-dependent pathways involved in the cell stress response.[52]

Figure 6.

Emission images of A549 cells subjected to diverse gemcitabine concentrations (0, 5, 10, and 20 μM) during a 2-hour pre-treatment period in glucose-absent DMEM medium, succeeded by concurrent incubation with 10 μM of probe A in glucose-absent DMEM medium for an extra 30 minutes. Fluorescence signals were elicited employing an excitation wavelength of 633 nm, and emissions were documented within the 700 nm to 780 nm range.

Camptothecin is a natural alkaloid chemotherapeutic agent that works by inhibiting topoisomerase I, an enzyme critical for DNA replication and repair.[55-57] We investigated how camptothecin affects NAD(P)H metabolism in A549 lung cancer cells using probe A as detailed in supporting information. Similar to cisplatin and gemcitabine, we observed a dose-dependent fluorescence increase with higher camptothecin concentrations (Figure 7), reflecting elevated NAD(P)H levels. This rise in NAD(P)H can be attributed to metabolic adaptations in response to camptothecin-induced DNA damage. By inhibiting topoisomerase I activity, camptothecin causes DNA strand breaks and replication fork stalling. This triggers cell stress pathways and activation of NAD(P)H-dependent enzymes like PARP to repair DNA damage, consuming NAD⁺.[55-57] Further, camptothecin stimulates NAD⁺ biosynthesis through salvage and de novo synthesis from NAM and NR,[58] replenishing NAD⁺ pools depleted by DNA repair mechanisms. As camptothecin dose increases, DNA damage and cell stress escalate, leading to enhanced activation of glycolytic and NAD⁺ biosynthetic pathways. This culminates in dose-dependent accumulation of NAD(P)H levels. In summary, probe A enabled real-time tracking of the intricate metabolic responses to camptothecin involving DNA repair, redox signaling, and NAD⁺ metabolism that underlie the observed rise in NAD(P)H.

Figure 7.

Emission images of A549 cells initially subjected to 5, 10, and 20 μM of camptothecin in a glucose-absent DMEM medium for 2 hours, and then incubated with 10 μM of probe A in a DMEM medium without glucose for 30 minutes. The fluorescence images were captured between 700 nm to 800 nm upon excitation at 633 nm.

We utilized freshly hatched, starved fruit fly larvae to evaluate probe A's ability to detect NAD(P)H dynamics in vivo. Larvae are ideal due to their stabilized NAD(P)H levels resulting from metabolic adaptations to nutrient deprivation at hatching.[25, 33, 35, 37, 38, 59, 60] Lacking food, the larval metabolism conserves NAD(P)H, a vital coenzyme involved in cellular reactions.[25, 33, 35, 37, 38, 59, 60] This steady state provides a controlled baseline for studying NAD(P)H fluctuations in response to treatments.[25, 33, 35, 37, 38, 59, 60] In control larvae immersed in PBS with probe A, weak fluorescence confirmed the detection of intrinsic NAD(P)H. Treatment with varying concentrations of NADH elicited a dose-dependent fluorescence increase, reflecting rising NAD(P)H levels (Figure 8). Treatment with cisplatin also increased the fluorescence of probe A versus the control, implying elevated NAD(P)H levels resulting from cisplatin-induced stress (Figure 8). The NAD(P)H rise likely stems from metabolic adaptations and cell response pathways activated by cisplatin’s toxicity.

Figure 8.

Fluorescence imaging was performed on recently hatched, starved fruit fly larvae subjected to NADH and cisplatin treatments. In the NADH treatment scenario, larvae were exposed to varying NADH concentrations ranging from 0 (control) to 30 μM, utilizing a pH 7.4 PBS buffer, for a duration of 1 hour. Conversely, for the cisplatin treatment, larvae were exposed to cisplatin concentrations ranging from 0 (control) to 20 μM, within a pH 7.4 PBS buffer, for a 2-hour period. Following these treatments, the larvae were subjected to three washes with PBS buffer and then transferred into a PBS buffer containing 10 μM of probe A for an additional 2 hours. Fluorescence signals captured in the images were obtained within a spectral range of 700 to 800 nm, employing an excitation wavelength of 633 nm.

4. Conclusion

In summary, we have successfully constructed two novel fluorescent probes, A and B, with absorption and near-infrared emission, for selective NAD(P)H detection in cells and organisms. Leveraging cyanine dyes' photophysical properties, these probes provide near-infrared excitation/emission to minimize background interference and photodamage. Probe A enabled unique real-time insights into NAD(P)H signaling dynamics in response to major nutrients, metabolites, and anticancer drugs. In A549 cells, glucose increased NAD(P)H levels, while lactate and pyruvate showed distinct effects, highlighting probe A's ability to elucidate NAD(P)H fluctuations induced by metabolites. Furthermore, probe A revealed concentration-dependent NAD(P)H elevations in response to cisplatin, gemcitabine, and camptothecin, unraveling metabolic adaptations to drug-induced stress. Crucially, probe A also effectively monitored in vivo NAD(P)H changes in fruit fly larvae treated with NADH and cisplatin. Overall, probe A allowed unprecedented visualization of NAD(P)H dynamics in living systems, providing fundamental new insights into redox biology, cell metabolism, and drug response pathways. Our rationally designed near-infrared fluorescent probes fill a major technology gap for studying NAD(P)H signaling in situ. This exploration into the impact of connection bridges on probe sensitivity not only deepens our comprehension of these probes' behavior but also provides invaluable insights for shaping the design of future NAD(P)H-responsive probes. It underscores the critical importance of tailoring probe architecture to achieve optimal performance in biosensing applications. The development and successful application of these biosensing tools represent a critical advance that could propel innovative diagnostics and therapeutics targeting aberrant NAD(P)H metabolism underlying diseases like cancer. Going forward, further optimization and application of these probes will illuminate previously invisible NAD(P)H processes, accelerating biomedical research.

Highlights.

• NAD(P)H-biosensing probes for near-infrared detection

• Importance of the probe connection bridges

• Reduced background fluorescence, autofluorescence, and cellular damage

• Detect changes in NAD(P)H levels in A549 cells

• Responds to glucose, lactate, pyruvate, and various cancer drugs

• Probe applied to freshly hatched fruit fly larvae

Biographies

Sushil K. Dwivedi is a research scientist at Michigan Technological University, specializing in fluorescent probes based on different fluorophores.

Dilka Liyana Arachchige is a Ph.D. student at Michigan Technological University, specializing in optical measurements and biomedical applications of fluorescent probes.

May Waters is a undergraduate student, currently conducting optical measurements and imaging of fluorescent probes.

Sophia Jaeger is a undergraduate student, currently conducting optical measurements and imaging of fluorescent probes.

Mohamed Mahmoud is a Ph.D. student, currently focusing on synthesis of cyanine- and hemicyanne-based fluorescent probes for biomedical application.

Adenike Mary Olowolagba is a Ph.D. student, currently focusing on synthesis of coumarin-based fluorescent probes for biomedical applications.

Daniel R. Tucker is a undergraduate student, currently focusing on computation chemistry of fluorescent probes.

Micaela Geborkoff is a undergraduate student, currently focusing on use of fruit fly larvae for biomedical application.

Thomas Werner is a full professor at Michigan Technological University, specializing in field of fruit fly larvae for medical application

Rudy L Luck is an associate professor at Michigan Technological University, specializing in computation chemistry of fluorescent probes/

Bhaskar Godugu is a director of mass spec facility at University of Pittsburgh.

Haiying Liu is a full professor at Michigan Technological University. He focuses his research on creating innovative fluorescent probes utilizing various fluorophores including BODIPY, Coumarin, Rhodamine, Rhodol, Fluorescein, hemicyanine and cyanine dyes. The goal is to develop these novel probes for use in biomedical applications.

Data Availability

The data from this study will be provided to interested parties upon request.