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. Author manuscript; available in PMC: 2021 Jun 11.
Published in final edited form as: Methods Enzymol. 2019 Mar 14;622:431–448. doi: 10.1016/bs.mie.2019.02.037

Bioorthogonal oncometabolite ligation

Chloe A Briney 1, Susana Najera 1, Jordan L Meier 1,*
PMCID: PMC8195447  NIHMSID: NIHMS1703243  PMID: 31155064

Abstract

Dysregulated cellular metabolism is an emerging hallmark of cancer. Improved methods to profile aberrant metabolic activity thus have substantial applications as tools for diagnosis and understanding the biology of malignant tumors. Here we describe the utilization of a bioorthogonal ligation to fluorescently detect the TCA cycle oncometabolite fumarate. This method enables the facile measurement of fumarate hydratase activity in cell and tissue samples, and can be used to detect disruptions in metabolism that underlie the genetic cancer syndrome hereditary leiomyomatosis and renal cell cancer (HLRCC). The current method has substantial utility for sensitive fumarate hydratase activity profiling, and also provides a foundation for future applications in diagnostic detection and imaging of cancer metabolism.

1. Introduction

The ability of metabolism to influence tumorigenesis is an emerging paradigm in oncology (Vander Heiden, Cantley, & Thompson, 2009). In addition to providing building blocks for cell division, in several instances metabolism has also been found to directly fuel malignant signaling through the production of “oncometabolites.” One such example occurs in the genetic cancer disposition syndrome hereditary leiomyomatosis and renal cell cancer (HLRCC) (Launonen et al., 2001; Tomlinson et al., 2002). Patients with HLRCC are born with inactivating mutations in one allele of the TCA cycle enzyme fumarate hydratase (FH) (Fig. 1A). Loss of heterozygosity leads to an absence of FH activity, hyperaccumulation of fumarate, and a predisposition to aggressive kidney cancer (Sudarshan, Pinto, Neckers, & Linehan, 2007). Inactivation of FH is associated with distinct changes in gene expression. This is believed to reflect fumarate’s ability to reversibly and irreversibly interact with proteins, which can influence transcription factor stability, chromatin demethylation, and cysteine-dependent protein–protein interactions (Kulkarni, Bak, et al., 2019; Sudarshan et al., 2009; Xiao et al., 2012; Zengeya, Kulkarni, & Meier, 2015). In addition to HLRCC, hyperaccumulation of fumarate has also been characterized in diabetes, non-hereditary kidney cancer, neuroblastoma, colorectal cancer, and tumors of the adrenal gland (Ashrafian et al., 2010; Bateman et al., 2017; Cancer Genome Atlas Research Network et al., 2016; Fieuw et al., 2012; Lehtonen et al., 2006; Nagai et al., 2007). These findings illustrate the ability of FH activity to be altered in the absence of mutation, and suggest fumarate may drive pathophysiological signaling in additional disease contexts.

Fig. 1.

Fig. 1

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(A) Mutations in the TCA cycle enzyme fumarate hydratase (FH) cause hyperaccumulation of the oncometabolite fumarate. (B) Fumarate is a metabolic dipolarophile. Reaction of fumarate with nitrileimine dipoles forms fluorescent pyrazoline cycloadducts, making this reaction useful for reporting fumarate levels in samples of interest.

Several methods for the analytical detection of fumarate have been developed. In clinical metabolomics, fumarate is most often detected by GC–MS or NMR (Collins, Patel, Putnam, Kapur, & Rakheja, 2017; Yang et al., 2013). However, these approaches require extensive derivatization and/or access to advanced instrumentation which may not always be available. FH activity has also been directly assessed using UV spectroscopy (Massey, 1955). This approach takes advantage of the reversibility of the FH reaction and monitors conversion of malate to fumarate by evaluating the latter’s absorbance at 240nm. However, this method is of limited utility in cell extracts which contain proteins, nucleic acids, and other metabolites that also absorb in this region. Furthermore, this approach does not have the potential to inform cellular imaging methods, whose development could lead to a better understanding of how fumarate’s subcellular localization correlates with its mechanistic effects.

To advance the study of aberrant FH activity in HLRCC and other cancers, we recently developed a fluorogenic approach to detect fumarate (Kulkarni, Briney, et al., 2019; Zengeya et al., 2016). Our method takes advantage of the ability of nitrileimines and electron-rich olefins to undergo [3+2] cycloaddition reactions to form pyrazoline cycloadducts. This reaction was first discovered by Huisgen and coworkers in the 1960s, before being significantly developed as a bioorthogonal protein ligation strategy by Lin’s group (Clovis, Eckell, Huisgen, & Sustmann, 1967; Lim & Lin, 2011). Inspired by these efforts, we hypothesized nitrileimines may also possess substantial reactivity with fumarate because it is an endogenous metabolic dipolarophile (Fig. 1B). Despite fumarate’s relatively high lying LUMO, we discovered nitrileimines readily react with the metabolite and generate strongly fluorescent pyrazoline products upon covalent bond formation. In our initial studies, we have applied this “bioorthogonal oncometabolite ligation” to sensitively detect FH-loss in HLRCC cell and tumor samples, as well as to image the accumulation of dipolarophile drugs and metabolites (Kulkarni, Briney, et al., 2019; Zengeya et al., 2016). More broadly, these studies established that the selectivity inherent to bioorthogonal reactions may, in some cases, be co-opted to study endogenous biological contexts (Devaraj, 2018).

To facilitate the further application of these tools, here we describe methods for the fluorogenic profiling of FH activity. First, we specify considerations important to the design, synthesis, and kinetic evaluation of fumarate-detecting dipoles. Next, we describe two approaches to specifically quantify FH activity in cell and tumor samples, using either hydrolytically-labile or photoinducible nitrileimine precursors. Finally, we discuss future opportunities for the application of these reactions to in the study of HLRCC, inborn errors of metabolism, and dipolarophile drugs.

2. Technical aspects

Fluorogenic reagents for the reaction-based detection of fumarate must balance several properties including (1) facile nitrileimine formation, (2) rapid reaction with fumarate, (3) slow decomposition in buffer, and (4) formation of highly fluorescent pyrazoline cycloadducts. Before describing protocols for fumarate detection using optimized reagents, we briefly describe some of these design considerations.

2.1. Considerations in the design of fluorogenic fumarate detection reagents

2.1.1. Synthesis, fluorescence, kinetics, and stability of nitrileimine precursors

Nitrileimine dipoles are inherently unstable in aqueous buffers and must be generated in situ from stable precursors for bioorthogonal labeling applications. The two classes of nitrileimine precursors that have been most well-characterized for these purposes are hydrazonyl chlorides and 2,5-diaryl tetrazoles (Fig. 2A) (Lee et al., 2013; Wang, Hu, Song, Lim, & Lin, 2008). Hydrazonyl chlorides spontaneously form nitrileimines in aqueous buffer upon loss of hydrochloric acid. Mechanistic studies by Liu and coworkers have found this reaction is base-catalyzed and highly sensitive to pH and chloride ion concentration (Wang, Lee, & Liu, 2014). In contrast, 2,5-diaryl tetrazoles can produce nitrileimines via photolysis of the tetrazole ring and loss of nitrogen (Wang et al., 2008). The wavelengths required for this photolytic reaction range from high energy UV light to two-photon excitation, and can be greatly influenced by the structure of the diaryl tetrazole substituents (Lim & Lin, 2011; Yu, Ohulchanskyy, An, Prasad, & Lin, 2013).

Fig. 2.

Fig. 2

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(A) Nitrileimines can be produced in situ from hydrazonyl chloride (left) and diaryl tetrazole (right) precursors. Each of these reagents have advantages that make them useful as fluorescent fumarate detection agents. (B) Molecular orbital diagram depicting energy gap between dipole HOMO and dipolarophile LUMO required for [3+2] cycloaddition. Careful selection of dipole precursors such as in (A) allows for optimization of reaction kinetics and sensitivity.

The reaction rate of nitrileimines with dipolarophiles such as fumarate is thought to be primarily dictated by the energy gap between the nitrileimine HOMO and dipolarophile LUMO (Fig. 2B) (Wang, Song, Hu, & Lin, 2009). Fumarate is less polarizable and has a relatively higher lying LUMO than the acrylate dipolarophiles commonly used in other applications, necessitating the identification of nitrileimine precursors with optimized reaction rates, stability, and fluorescence properties. Hydrazonyl chlorides such as 1 (Fig. 3) can be readily synthesized in two steps by microwave-assisted condensation of aldehydes and aryl hydrazines, followed by chlorination under Corey-Kim conditions (Zengeya et al., 2016). This facile synthetic entry allowed the production of a small panel of analogs, which we used to define structure–reactivity relationships and fluorescence properties of the resulting fumarate-derived pyrazolines (Zengeya et al., 2016). Calculation of molar extinction coefficients and quantum yields revealed that the brightness of the pyrazoline fluorophore correlated well with fumarate detection sensitivity. In contrast, kinetic studies found that nitrileimines capable of more rapid reaction with fumarate also exhibited more rapid degradation by buffer, exemplifying the challenge of balancing on- and off-target reactivity. An important aspect of this approach is that it is not intrinsically limited to fumarate and can also detect a number of other metabolic dipolarophiles. Selectivity is therefore attained in these assays by the high dynamic range of fumarate levels in HLRCC (measured to be ~1000-fold greater than adjacent tissues) as well as by the addition of enzyme substrates such as malate, whose conversion and detection provides a signal-amplified readout of FH activity (Zengeya et al., 2016).

Fig. 3.

Fig. 3

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Exemplary hydrazonyl chloride (1) and tetrazole (2) nitrileimine precursors used in the protocols described in this chapter.

Very recently, we reported the development of second-generation diaryl tetrazoles for fumarate detection (2, Fig. 3) (Kulkarni, Briney, et al., 2019). While tetrazoles have long been the nitrileimine-precursors of choice for bioorthogonal labeling applications (Lim & Lin, 2011), their preparation is more challenging than that of hydrazonyl chlorides, requiring the synthesis of custom sulfonyl hydrazone and aryl diazonium building blocks. Synthetic protocols for 2 are reported elsewhere (Kulkarni, Briney, et al., 2019). In this study we used our knowledge of hydrazonyl chloride structure–activity relationships to rapidly identify a pair of diaryl tetrazoles that displayed suitable reactivity and fluorescence properties for fumarate detection (Kulkarni, Briney, et al., 2019). Comparative analyses revealed diaryl tetrazoles provide more sensitive readout than hydrazonyl chlorides of related structure, a finding attributed to their decreased susceptibility to nucleophilic decomposition. This illustrates the complementary nature of these two classes of dipolarophile reporters: hydrazonyl chlorides can be used to rapidly identify lead scaffolds with optimized kinetic and fluorescence properties, thereby facilitating the design of tetrazole detection agents with improved stability and sensitivity. We currently apply these approaches interchangeably, using hydrazonyl chloride 1 for the routine quantification of fumarate in biochemical assays (Protocol 2.2), and employing tetrazole 2 to sensitively profile endogenous fumarate and FH activity in cell and tumor extracts (Protocol 2.3).

2.2. Fluorogenic detection of fumarate using hydrazonyl chloride-derived nitrileimines

Hydrazonyl chloride 1 is readily synthesized in two steps from the commercially available aldehyde and hydrazine. Detailed synthetic protocols for this molecule are reported elsewhere (Zengeya et al., 2016). The reaction of 1 with fumarate is more rapid than other hydrazonyl chlorides, and forms a laser-excitable pyrazoline fluorophore characterized by a large Stokes shift (λex = 390nm, λem = 530nm; Fig. 3). While lacking the sensitivity of tetrazole reagents, hydrazonyl chloride 1 holds the advantage of utilizing a slightly simpler reaction protocol, as it does not require UV-based photolysis to uncage the reactive species. This makes 1 an ideal reagent for in vitro studies, and in the past we have applied it to monitor the reversibility of cysteine S-succination, as well as to quantify recombinant and cellular FH activity (Kulkarni, Bak, et al., 2019; Zengeya et al., 2016).

2.2.1. Materials

Hydrazonyl chloride 1 (Zengeya et al., 2016)

Fumaric acid (Alfa Aesar, A10976)

Dimethyl sulfoxide (DMSO, Sigma-Aldrich, D4540)

Acetonitrile (ACN, Sigma-Aldrich, 34851)

Sodium phosphate buffer, pH 7 (Teknova, P2070)

96-well microplate (Greiner BioOne, 655801)

Parafilm (VWR, 52858–076)

Multichannel pipet

Fluorimeter (Photon Technology International QuantaMaster 400)

3 × 3mm quartz cuvette (Hellma Analytics, 101–015-40)

2.2.2. Quantitative detection of fumarate using hydrazonyl chloride 1

  1. Dissolve an aliquot of purified hydrazonyl chloride 1 in acetonitrile at 200 mM. Concentrated stock solutions of 1 can be stored at −20°C for weeks without noticeable loss of signal. However, care should be taken to protect the compound from light and water, and prolonged storage in solution should be avoided to limit degradation.

  2. Make a stock solution of fumarate at 1 M. Fumaric acid may be dissolved in either DMSO or equimolar base (2 M NaOH). If fumarate is dissolved in DMSO, care should be taken to keep the final percentage of this organic co-solvent at or below 1%.

  3. Serially dilute solutions of fumarate in a 96-well plate for construction of a standard curve. For these studies, we typically first make an intermediate stock solution of 100 mM fumarate. This is used to make 50μL samples of 10 mM fumarate in 100 mM sodium phosphate buffer, pH 7, which are loaded into a 96-well plate and subjected to twofold serial dilutions. Ten dilutions cover a range of 10 mM to 10μM fumarate, and typically suffices for determination of an unknown sample concentration.

  4. In parallel, plate 50μL of experimental sample (unknown) into the same 96-well plate. The sample buffer should be identical to that used for the calibration curve, in this case 100 mM sodium phosphate buffer, pH 7.

  5. Using a multichannel pipet, add 50μL of 300μM hydrazonyl chloride in acetonitrile to bring each well to a final volume of 100μL. Mix by pipetting up and down once without bubbles.

  6. Cover the 96-well reaction block with Parafilm or a lid to prevent evaporation and place in the dark at room temperature for 1h. This incubation time has been empirically determined to allow reactions containing between 10 and 100μM fumarate to approach completion when performed in 1:1 acetonitrile: 100 mM sodium phosphate buffer, pH 7. Reaction times may need to be re-optimized for samples of alternative chemical composition.

  7. While samples are incubating, power up the Photon Technology International QuantaMaster 400 fluorimeter and turn on the lamp. The fluorimeter should be warmed up for 10–20min to maximize lamp life and consistency of measurements.

  8. Following incubation, transfer individual samples (100μL) to a quartz cuvette for collection of fluorescence spectra. For hydrazonyl chloride 1, samples should be excited at 390nm and emission spectra collected from 410 to 650nm. The maximum emission wavelength for each sample should be 530nm. Note: depending on the concentration of fumarate in each sample, the excitation and emission slit widths may need to be adjusted in order to avoid oversaturation. Fully open slit widths are appropriate for samples up to 10 mM fumarate with hydrazonyl chloride 1, but may need to be adjusted based on individual sample composition. Evaluating the most concentrated sample from the standard curve first can help determine the appropriate slit widths.

  9. Use the reported relative fluorescence units (RFUs) at 530nm to plot a standard curve of fumarate concentration versus fluorescence. Perform a linear regression on this curve, and use the equation to determine the amount of fumarate in the experimental (unknown) samples.

2.3. Fluorogenic detection of FH activity using tetrazole-derived nitrileimines

To expand the applications of fluorescent fumarate detection we recently reported the use of diaryl tetrazole 2 as a reporter of oncometabolite accumulation in HLRCC (Kulkarni, Briney, et al., 2019). This probe generates a nitrileimine almost identical in structure and electronics to hydrazonyl chloride 1. However, unlike hydrazonyl chlorides, the tetrazole of probe 2 is extremely stable and does not form nitrileimines or undergo hydrolytic degradation reactions in the absence of high energy UV light. Additionally, the carboxylate moiety of 2 is modified with an azide group, which allows for its conjugation to tags for subcellular targeting. These enhancements greatly improved the sensitivity of 2, and allowed us to detect FH activity and for the first time differentiate HLRCC tumor specimens from healthy adjacent tissue based on endogenous dipolarophile metabolite levels (Kulkarni, Briney, et al., 2019). The stability of 2 also facilitated its application in proof-of-concept cell imaging studies, although these are limited by the short wavelengths required for tetrazole uncaging. Here we describe a protocol for the application of 2 to quantify cellular FH activity (Fig. 4). This method relies on the ability of endogenous FH to catalyze the formation of fumarate from malate. Monitoring this “reverse reaction” using 2 converts FH activity into an increase in fluorescence intensity, and allows for straightforward control of enzymatic signal amplification by increasing incubation time. FH activity can be determined from cell or tissue samples without fractionation, provided extracts are prepared in appropriate non-denaturing buffers. Calibration to a standard curve of commercially available FH is used to convert production of fumarate by cell extracts to quantitative FH activity units, facilitating comparison of multiple samples.

Fig. 4.

Fig. 4

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Experimental protocol for profiling cellular FH activity using tetrazole 2. Cells or tissue samples of interest are gently lysed under non-denaturing conditions. Crude cell extracts are treated with malate, which is converted to fumarate by the “reverse reaction” of FH. Samples are then developed by adding a solution of 2 in acetonitrile. Photoirradiation at 302nm uncages a nitrileimine, which reacts with newly formed fumarate to form a fluorescent cycloadduct. Samples are then analyzed via fluorimetry and quantified using a standard curve of FH activity.

2.3.1. Materials

Extracts are prepared from cells cultured under standard conditions. For FH activity determination it is critical to avoid protein denaturation. This necessitates lysis be performed in buffers at a pH optimal for enzyme activity. Soluble FH can be isolated with other proteins from cells and tissues by gentle sonication on ice. Alternatively, for rapid profiling of cell lines in microplates, extracts with active FH activity may be prepared by simple incubation with the non-denaturing detergent NP-40. After lysis by sonication, insoluble material is cleared from solution by centrifugation at 4°C. Microplate extracts should be assayed immediately, while cell and tissue samples may be stored at −80°C and thawed on ice prior to use.

Plated cells

Cell scraper (Biologix, 70–2180)

PBS (Thermo Fisher, 10010)

Sodium phosphate buffer pH 7 (Teknova, P2070)

Potassium phosphate buffer pH 7.6 (G Biosciences, 786–488/786–487)

Protease Inhibitor Cocktail (Cell Signaling, 5871S)

NP40 (Sigma-Aldrich, NP40S)

Ultrasonicator (Qsonica Q700)

Refrigerated benchtop centrifuge

Qubit Protein Assay Kit (Q33212)

Qubit Fluorometer (Q32866)

Tetrazole 2 (Kulkarni, Briney, et al., 2019)

Fumarate hydratase from porcine heart (Sigma Aldrich, F1757)

Malate (TCI, M0022)

Dimethyl sulfoxide (DMSO, Sigma-Aldrich, D4540)

Acetonitrile (Sigma-Aldrich, 34851)

96-well microplate (Greiner Bio-One, 655801)

Black sided clear bottomed 96-well microplate (Greiner Bio-One, 655090)

Multichannel pipet

Parafilm (VWR, 52858–076)

Fluorescence-capable plate reader (Biotek Synergy MX microplate reader)

Fluorimeter (Photon Technology International QuantaMaster 400)

UV transilluminator (UVP MultiDoc-It Imager Benchtop UV Transilluminator)

2.3.2. Preparation of extracts with active FH from cell and tissue samples by sonication

  1. Plate cell line of interest and allow to grow to confluency. As a control we typically assess two well-characterized HLRCC cell lines: (1) UOK262 cells, which lacks FH activity, and (2) UOK262WT cells, in which FH activity has been rescued (Yang et al., 2012).

  2. To ensure isolation of sufficient proteome containing FH for this method, we recommend plating HLRCC cells in six-well plates at a density of ~300,000 cells/well. This provides confluent cells ready to be harvested in ~48h.

  3. Gently wash adherent cell lines twice with ice cold PBS, being careful not to dislodge cells.

  4. Harvest cells by gentle lifting using a cell scraper. Care should be taken to scrape the bottom and sides of the dish to ensure complete sample collection.

  5. Pellet cells by centrifugation for 5min at 700×g, 4°C. Discard the supernatant.

  6. Prepare lysis buffer: we recommend using 100 mM potassium phosphate buffer pH 7.6 for FH activity determinations using tetrazole 2, and 100 mM sodium phosphate buffer pH 7 for FH activity determinations using hydrazonyl chloride 1. Add 1 × Protease Inhibitor Cocktail to prevent protein degradation during sample preparation.

  7. Add lysis buffer to cell pellet. We recommend adding ~40μL buffer for cells harvested from a six-well plate, or ~5mL buffer per 1mL packed tissue sample.

  8. Lyse cells via ultrasonication on ice, using three pulses (1s each at 5J) with 30s on ice between each pulse.

  9. Centrifuge 30min at 4°C, 30,279×g (maximum speed) in a pre-chilled benchtop centrifuge.

  10. Transfer supernatant to a fresh tube.

  11. Quantify protein concentration using the Qubit Protein Assay according to the manufacturer’s directions.

  12. Dilute samples to 1–2mg/mL using the same lysis buffer and store at −80°C.

2.3.3. Alternative procedure for preparation of cell extracts via direct lysis in microplates

  1. Plate cell lines in sterile, clear-bottomed, black-sided 96-well plates and allow to adhere overnight.

  2. Prepare detergent-containing lysis by adding 0.8% NP-40 to the 100 mM potassium phosphate buffer pH 7.6 used for FH activity determinations by tetrazole 2.

  3. Remove media from the wells.

  4. Gently wash cells once with 100μL room temperature PBS.

  5. Add 47.5μL detergent-containing lysis buffer, and mix by pipetting up and down without bubbles.

  6. Microscale FH activity analyses by 2 should be performed immediately, directly in this microplate.

2.3.4. Quantitative determination of FH activity using diaryl tetrazole 2

  1. Dissolve an aliquot of purified diaryl tetrazole 2 in DMSO at 10 mM. Concentrated stock solutions of 2 are stable and can be stored at −20°C for months without noticeable change in activity. Care should be taken to protect the compound from light. Use this concentrated stock to prepare a diluted working solution of 2 at 300μM in acetonitrile.

  2. Prepare FH activity assay buffer: 100 mM potassium phosphate buffer, pH 7.6. Be sure to prepare at least 50μL of buffer per 5μg of cell extract to be analyzed. Assay buffer can be kept at room temperature prior to use.

  3. Construct a standard curve of FH activity: For these studies, we dilute commercial FH enzyme derived from porcine heart (Sigma) of known activity to a final volume of 47.5μL in FH activity buffer. Ten twofold serial dilutions of 2.5mU FH covers a range that typically suffices for determination of FH activity in 5–20μg cell or tissue extract. Be sure to include a no-enzyme control.

  4. Calculate the volume of sonicated cell extract required to add 5–20μg to each reaction. Add this amount to separate wells of the microplate to bring the total volume to 47.5μL in FH activity buffer. For cellular samples prepared by detergent lysis in 96-well plates (Section 2.2.2), proceed directly to step #5. Samples should be done in triplicate and can be plated in the same microplate as the FH activity standard curve.

  5. Prepare a solution of 1M malate in 2M NaOH. The use of neutralized malate at pH 7 is essential to preserve FH activity, as acidic samples perturb FH activity.

  6. Incubate purified commercial FH as well as FH-containing cell extracts with malate. For initial activity determinations, we typically add 2.5μL of a 1M solution of malate to the 47.5μL in each well (50 mM final concentration) and measure conversion to fumarate via the procedure below at 0, 1, and 2h. It is only necessary to create an FH standard curve at one of the timepoints. The microplate plate should be kept covered with Parafilm or a lid at room temperature between enzyme activity determinations.

  7. At the end of each timepoint, add 50μL of tetrazole 2 (300μM working stock in acetonitrile) to standards and unknown samples using a multichannel pipet. This will bring each well to a final volume of 100μL. Mix by pipetting up and down once without bubbles. Note: hydrazonyl chloride 1 (300μM in acetonitrile) can also be used for FH activity measurements. In this case, addition of the hydrazonyl chloride should be performed as above, and step #8 of this protocol replaced with a 1h incubation of the covered microplate in the dark at room temperature.

  8. 10–20min prior to use, power up the Photon Technology International QuantaMaster 400 fluorimeter and turn on the lamp. The fluorimeter should be warmed up in order to maximize lamp life and consistency of measurements.

  9. Place the microplate into the UV transilluminator and shut the door. Irradiate the samples at 302nm for 2min. Note: If a UV transilluminator is not available, a handheld UV lamp equipped with a logwave 302nm bulb may also be used.

  10. Following incubation, transfer individual samples (100μL) to quartz cuvettes for collection of fluorescence spectra. For tetrazole 2, samples should be excited at 410nm and emission spectra collected from 430 to 630nm. The maximum emission wavelength for each sample should be 540nm. Note: depending on the concentration of fumarate produced by FH in each sample, the excitation and emission slit widths may need to be adjusted in order to avoid oversaturation. Fully open slit widths are appropriate for samples up to 25 mM fumarate or 2.5mU FH with tetrazole 2, but may need to be adjusted based on individual sample composition. Evaluating the standard or unknown sample predicted to have the highest FH activity can help determine the appropriate slit widths.

  11. Alternatively, if a fluorescently-capable plate reader is available, place the plate in the reader and excite at 410nm and collect emission at 540nm. This method is less sensitive but provides reliable results for samples with abundant FH activity, and works best in a black 96-well plate with a clear bottom.

  12. Use the reported relative fluorescence units (RFUs) at 540nm to plot a standard curve of FH activity versus fluorescence. Perform a linear regression on this curve, and use the equation to determine the units of FH enzyme activity present in the experimental (unknown) samples.

3. Discussion

3.1. Critical parameters and troubleshooting

One aspect critical for FH activity determinations by fluorogenic probes 1 and 2 is the use of high quality cell and tissue extracts. The factors we have found most commonly cause loss of FH activity in extracts are (1) use of strong detergents (e.g., SDS) during lysis, (2) subjection of samples to repeated freeze-thaw cycles, and (3) addition of unbuffered acidic malate during the activity measurement step. This can be mitigated by careful selection of buffers and preparation of samples immediately prior to analysis. In addition, analysis of recombinant commercial FH, as well as extracts prepared from isogenic wild-type and mutant FH cell lines (i.e., UOK262 and UOK262 WT) can be used as valuable controls to verify proper assay procedure and extract activity, respectively. The quality of the nitrileimine precursor is also paramount for these studies. Stock solutions and freshly dissolved samples should be checked for purity periodically using thin layer chromatography, analytical HPLC, or LC–MS. Finally, when adapting this procedure for new applications care should be taken to determine the compatibility of buffer components with these reagents as well as their influence on the fluorescence spectra of 1 and 2. We typically use 1 and 2 to develop solutions of fumarate by incubation in solutions of 1:1 acetonitrile: phosphate buffer (Kulkarni, Briney, et al., 2019; Zengeya et al., 2016). The use of an organic co-solvent such as acetonitrile is critical for fumarate detection, as it facilitates hydrazonyl chloride solubility and slows competing hydrolysis reactions. However, recent studies indicate nitrileimines can directly react with acetonitrile, suggesting DMSO may be a preferable co-solvent in some cases (Li et al., 2016). This study found that nitrileimines can also react directly with thiols and carboxylates. Therefore, control reactions of 1 and 2 with either fumarate or a more rapidly reacting molecule such as ethyl acrylate should always be performed to determine the influence of new buffer components on reporter sensitivity and background fluorescence.

3.2. Applications and future directions

Here we have presented a detailed set of protocols which employ the fluorogenic nitrileimine-olefin cycloaddition to profile the oncometabolite fumarate. Our strategy hybridizes bioorthogonal and reaction-based sensing strategies, and defines a novel disease context in which this chemoselective ligations can be harnessed to study endogenous biology (Kulkarni, Briney, et al., 2019; Zengeya et al., 2016). As currently constituted, these methods are well-suited for in vitro measurements of fumarate as well as determination of FH activity from cell, xenograft, and patient tumor samples. Importantly, fluorogenic reagents 1 and 2 are not limited to detection of fumarate, and in theory may be used to assay any enzyme that produces or consumes a dipolarophile of interest. Additional coupling opportunities can be envisioned by using these species to follow metabolites involved in the TCA cycle (cis-aconitate), immunometabolism (itaconate), and lipid oxidation (9-hydroxynonenal) (Fig. 5). We also envision several improvements that may expand the applications of this technology. For example, the relatively high energy UV light required to uncage tetrazole 2 can crosslink cellular DNA, which has so far limited its utility as an imaging agent in proof-of-concept experiments. In the future this may be addressed by the release of nitrileimines using two-photon light, which is less damaging to biological structures (Yu et al., 2013). A challenge to sensitivity is the rate of reaction, which is inherently limited by fumarate’s low lying LUMO (Zengeya et al., 2016). While this can in theory be addressed by designing nitrileimines with high energy HOMO, in practice such structural alterations can also increase the electrophilicity of the dipole, causing cognate destruction by biological nucleophiles which nullify any gains in sensitivity. Lin and coworkers recently reported an elegant strategy to circumvent this side reaction through steric hindrance of the nitrileimine dipole (An, Lewandowski, Erbay, Liu, & Lin, 2018; An & Lin, 2018). In the future it will be interesting to see whether these reagents may display improved detection of endogenous dipolarophiles in HLRCC. Other potential approaches to improve the sensitivity of this method include the exploration of alternative classes of dipoles and new reporter strategies, such as using the nitrileimine-fumarate cycloaddition to trap diffusable radiotracers and fluorophores in HLRCC cells (Kulkarni, Briney, et al., 2019). With further development, such methods have the potential to greatly influence our knowledge of how metabolism influences the development, progression, and treatment of cancer.

Fig. 5.

Fig. 5

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Examples of other metabolite dipolarophiles whose enzymatic production or consumption may be assayed by 1 and 2.

Acknowledgments

We are grateful to members of the Meier lab for providing helpful discussions and critical feedback on this chapter. This work was supported by the Intramural Research Program of the NIH, National Cancer Institute, Center for Cancer Research (ZIA-BC011488-06).

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