NMR-Based Activity Assays To Characterize Enzymes in the

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Chapter 3

NMR-Based Activity Assays To Characterize Enzymes in the Biochemistry Laboratory and in Undergraduate Research Brian J. Stockman* Department of Chemistry, Adelphi University, Garden City, New York 11530, United States *E-mail: [email protected]

Pragmatic applications of 1H and 19F NMR-based activity assays in the teaching and research environments are described. NMR-based activity assays are a logical extension from prerequisite organic chemistry NMR analyses. Relationships between chemical structure and chemical shift, and between peak area and quantity, are extended to distinguish between substrate and product resonances and to determine their corresponding concentrations, respectively. The equilibrium constant for a reaction can be determined by integrating peak areas. The more intensive setting of undergraduate research provides an opportunity to introduce students to the practical aspects of these methods in the context of an antitrichomonal drug discovery project. NMR-based activity assays are being used to screen essential enzymes for inhibitors, determine IC50 values, weed out aggregation-based or otherwise false-positive inhibitors, and determine structure/activity relationships.

Introduction Adelphi University purchased a 500 MHz NMR spectrometer in 2012. It is introduced to students in the first-semester organic chemistry laboratory when groups collect 13C NMR spectra while getting an overview of the instrumentation. Then in second-semester organic chemistry laboratory, each student obtains hands-on experience collecting 1H NMR spectra on three occasions: nitration of methylbenzoate, reduction of camphor with sodium © 2016 American Chemical Society Soulsby et al.; NMR Spectroscopy in the Undergraduate Curriculum: Upper-Level Courses and Across the Curriculum Volume 3 ... ACS Symposium Series; American Chemical Society: Washington, DC, 2016.


borohydride, and identification of an unknown (1). The 1H NMR spectrum of methyl 3-nitrobenzoate demonstrates the relationships between chemical shift and type of proton, splitting patterns and neighboring protons, and peak area and number of protons on a given carbon. The 1H NMR spectrum of the products from the camphor reduction demonstrates that chemical shift is conformation dependent and that peak area can also be correlated with the amounts of multiple products formed (borneol and isoborneol). Second-semester organic chemistry is a prerequisite for both biochemistry laboratory and undergraduate research where these principles are applied to biochemical systems. Biochemistry laboratory is co-requisite with first-semester biochemistry lecture. Lab meets for one five-hour block each week during the 14-week semester with a maximum of 12 students in each section. The lab is project-based (2), with students working in groups of three to select a protein, design and carry out a purification protocol, and design and carry out a protein characterization protocol. Typical sources of protein include animal tissues or products, plants, or E. coli expression. Typical characterizations include SDS gel electrophoresis; specific activity; activity or structure as a function of pH, salt concentration, temperature, or organic solvents; and determination of substrate Km values. Each group is also required to spend at least one lab period developing and using NMR methods to characterize their protein. NMR has been used by others to characterize numerous enzyme reactions in the biochemistry laboratory including the stereospecificity of dehydrogenases that use NAD+/NADH (3, 4), acetylcholinesterase (5), acylase I (6–9), fumarase (10), β-glucosidase (11), invertase (12), and neuraminidase (13). These applications are extended here as a method to characterize and assay lactate dehydrogenase and β-amylase purified by students. Students explore the function of the enzyme by monitoring the reaction in the NMR tube. Chemical shifts and splitting patterns are used to identify the resonances of substrate and product. In the case of lactate dehydrogenase, peak areas of the pyruvate and lactate resonances are integrated and used to determine the absolute concentrations at equilibrium and the equilibrium constant for the reaction. In the case of β-amylase, incomplete conversion of starch to maltose is readily observed in the superposition of broad and narrow linewidths for substrate and product resonances, respectively. For proteins that do not possess enzymatic activity, such as the cytochrome c described here, students monitor the structural integrity of the protein in water/methanol mixtures. The relationship between chemical shift and conformation is extended to incorporate the spatial effects of a folded protein. In the case of cytochrome c, the loss of resonance dispersion and the sharpening of the heme resonances are used as indicators of protein denaturation. NMR spectroscopy provides an alternative to circular dichroism that has been used by others to study protein denaturation in the biochemistry laboratory (2, 14–16). Students that carry out undergraduate research (required of chemistry and biochemistry majors) can gain industry-relevant experience applying NMR-based activity assays. These assays are used routinely in the biotechnology and pharmaceutical industries (17–19). These single-enzyme assays are not prone to coupled-assay false positives, thus making them useful as orthogonal assays, complementing traditional high throughput screening assays and benchtop triage assays. They are also often used as stand-alone assays for fragment 34

Soulsby et al.; NMR Spectroscopy in the Undergraduate Curriculum: Upper-Level Courses and Across the Curriculum Volume 3 ... ACS Symposium Series; American Chemical Society: Washington, DC, 2016.


screening since they are ideally suited to work at the higher concentrations of test compounds, without worry of readout interference, required to detect these weaker inhibitors. In the research laboratory, students use NMR-based activity assays to identify inhibitors of Trichomonas vaginalis nucleoside ribohydrolases. This parasite causes the most prevalent non-viral sexually transmitted disease (20). Increasing resistance to existing therapies (21) is driving the need for novel, mechanism-based treatments. Essential nucleoside salvage pathway enzymes represent prime targets (22). NMR-based activity assays are described for two enzymes, uridine nucleoside ribohydrolase and adenosine/guanosine preferring nucleoside ribohydrolase. The assays are being used to screen for chemical starting points, determine IC50 values, weed out aggregation-based or otherwise false-positive inhibitors, and determine structure/activity relationships. Five applications of using NMR to characterize enzymes or proteins in the teaching and research environments are detailed below. In the biochemistry laboratory, 1H NMR-based activity assays are described for monitoring the reactions catalyzed by lactate dehydrogenase and β-amylase. The use of 1H NMR to monitor the denaturation of cytochrome c is also presented. In the undergraduate research setting, 1H and 19F NMR-based activity assays are described for two enzymes, uridine nucleoside ribohydrolase and adenosine/guanosine preferring nucleoside ribohydrolase.

Biochemistry Laboratory: Lactate Dehydrogenase Lactate dehydrogenase (LDH) catalyzes the NADH-dependent reduction of pyruvate to L-lactate during lactic acid fermentation, thus regenerating NAD+ required for glycolysis to continue under anaerobic conditions. The ΔG′° for the reaction favors the reaction in the direction shown in Figure 1.

Figure 1. NADH-dependent reduction of pyruvate to L-lactate catalyzed by the enzyme lactate dehydrogenase (LDH). Separate solutions containing pyruvate and lactate were prepared first in order to assign the 1H resonances as shown in Figure 2. Students were asked to predict what the spectra will look like while the data is collecting using their knowledge of chemical shifts and coupling patterns from organic chemistry. The 1H NMR spectrum of pyruvate contains the expected singlet resonance at 2.27 ppm while that for lactate contains the expected doublet at 1.23 ppm and quartet at 4.02 ppm. The smaller singlet at 1.38 ppm in Figure 2A is thought to arise from an acetaldehyde impurity in the pyruvate sample. Resolved resonances for substrate and product indicate the feasibility of monitoring the reaction by 1H NMR. 35



Figure 2. 1H NMR spectra of (A) pyruvate and (B) lactate solutions. The CH3 resonances of pyruvate and lactate are clearly distinguishable as a singlet at 2.27 ppm and doublet at 1.38 ppm, respectively. The CH group of lactate appears as a quartet at 4.02 ppm. The smaller singlet at 1.38 ppm marked by an asterisk is likely an acetaldehyde impurity in the pyruvate sample.

Two solutions were then prepared in parallel, each containing 1 mM pyruvate and 1 mM NADH. To the first was added 50 μL of buffer, while to the second was added 50 μL of LDH solution. The samples were incubated for 30 minutes at room temperature prior to NMR data collection. Figure 3A shows the 1H NMR spectrum after incubation in the absence of LDH. The pyruvate resonance at 2.27 ppm is clearly visible, while resonances for lactate at 1.23 and 4.02 ppm are not observed. Doublets at 2.59 and 2.71 ppm are also observed in this region of the 1H NMR spectrum for the two C4 protons of the nicotinamide ring of NADH. Figure 3B shows the 1H NMR spectrum after incubation in the presence of LDH. Now the pyruvate resonance is significantly diminished, while lactate resonances are now observed and NADH signals are now absent. Integration of the methyl resonances indicates an 8.6 to 1.0 ratio of lactate to pyruvate, corresponding to concentrations of 0.89 mM and 0.11 mM, respectively. Based on the reaction stoichiometry this corresponds to a K′eq of 36



approximately 65. The acetaldehyde impurity (marked by an asterisk) is also significantly diminished in Figure 3B suggesting that some of the NADH has been consumed reducing it to ethanol, either by LDH or another enzyme in the LDH sample. Thus the calculated K′eq for the LDH reaction is an underestimate of the actual value.

Figure 3. 1H NMR spectra of pyruvate and NADH solution after 30 minutes (A) without and (B) with LDH. The pyruvate CH3 and NADH resonances are still observed after 30 minutes in the absence of LDH. However, in the presence of LDH, these resonances are markedly reduced (or absent) after 30 minutes, while new resonances corresponding to the lactate CH and CH3 groups appear.

Experimental Details for Lactate Dehydrogenase Lactate dehydrogenase (LDH) was purified from bovine heart using a combination of homogenization, centrifugation, ammonium sulfate precipitation, anion exchange chromatography, and gel filtration chromatography (23). Fractions from the gel filtration column with LDH activity were pooled and concentrated to about 5 mL final volume for the characterization experiments. Stock solutions of 10 mM lactate, 10 mM pyruvate, and 10 mM NADH were prepared in 200 mM sodium phosphate buffer at pH 7.0. NMR samples were 37 Soulsby et al.; NMR Spectroscopy in the Undergraduate Curriculum: Upper-Level Courses and Across the Curriculum Volume 3 ... ACS Symposium Series; American Chemical Society: Washington, DC, 2016.


then prepared by combining appropriate amounts of stock solutions, D2O, and buffer to give 1 mM solutions of the desired compound(s) and 10% D2O in a final volume of 600 μL. For samples containing LDH, 50 μL of the final concentrated sample were also added. Wilmad 507-PP NMR tubes were used. All NMR data sets were collected on a Bruker Avance III 500 MHz NMR spectrometer at 298 K using a 5 mm BBFO room temperature probe. All 1H NMR spectra were collected with 64 scans. The water resonance was suppressed using excitation sculpting with gradients (24).

Biochemistry Laboratory: β-Amylase β-Amylase catalyzes the partial hydrolysis of starch into maltose and can be easily purified from plant sources such as soy beans or sweet potatoes. The reaction catalyzed is shown in Figure 4.

Figure 4. Hydrolysis of starch to maltose catalyzed by the enzyme β-amylase.

38 Soulsby et al.; NMR Spectroscopy in the Undergraduate Curriculum: Upper-Level Courses and Across the Curriculum Volume 3 ... ACS Symposium Series; American Chemical Society: Washington, DC, 2016.


Separate solutions containing maltose and starch were prepared first in order to ascertain if 1H resonances from maltose would be distinguishable from those of starch. The 1H NMR spectra for starch and maltose are shown in Figure 5A and Figure 5E, respectively. It is clearly observed that maltose has a unique doublet of doublets resonance at 3.18 ppm and a triplet resonance at 3.33 ppm. These signals correspond to the C2 proton of the reducing glucose and the C4 proton of the non-reducing glucose. This comparison between starch and maltose provides another interesting teachable moment in that the resonance linewidths observed for the starch polymer are significantly broader compared to those for the individual maltose monomers. The relationship between correlation time and linewidth can make for an interesting discussion, especially for those students that are keenly interested in math or physics. Then a second starch solution was prepared containing β-amylase. The sample was placed into the NMR spectrometer immediately upon the addition of enzyme and data collection was initiated. The dead time between adding enzyme and the first NMR scan (including putting the sample into the NMR tube and magnet, locking, and shimming) was about three minutes. Thus the first 1H NMR spectrum was considered to be at 5 minutes. Subsequent spectra were then acquired every 5 to 10 minutes until no further increase in the maltose signal intensities were observed. Figure 5B-D show the spectra acquired at 5, 10, and 40 minutes. The reaction is essentially complete after 40 minutes as the spectrum remained unchanged at 70 minutes (data not shown). The spectrum in Figure 5D suggests that a mixture of starch and maltose is present. The reaction does not fully convert starch to maltose, most likely because β-amylase is not capable of fully hydrolyzing the branched amylopectin component of starch. This might also result from product inhibition or the limited solubility of the starch solution. As a control, the 1H NMR spectrum of the starch solution without added β-amylase was also collected after 40 minutes (data not shown).



Figure 5. 1H NMR spectra of (A) starch, (B-D) after addition of β-amylase at time points of 5, 10, and 40 minutes, and (E) maltose. Starch and maltose are clearly distinguishable by both the significant difference in linewidths and by the appearance of new resonances in the maltose spectrum corresponding to the C2H of the reducing glucose and the C4H of the non-reducing glucose. After addition of β-amylase to the starch solution, the narrower maltose resonances increasingly superimpose with the broad starch resonances, and the maltose C2H and C4H resonances increase in intensity over time.

Experimental Details for β-Amylase β-Amylase was purified from soy flour using a combination of acid extraction, centrifugation, ammonium sulfate precipitation, anion exchange chromatography, and gel filtration chromatography (25). Fractions from the gel filtration column with β-amylase activity were pooled and used for the characterization experiments. Stock solutions of 10 mM maltose and 1% starch were prepared in water. NMR 40 Soulsby et al.; NMR Spectroscopy in the Undergraduate Curriculum: Upper-Level Courses and Across the Curriculum Volume 3 ... ACS Symposium Series; American Chemical Society: Washington, DC, 2016.


samples were then prepared by combining 540 μL of these solutions with 60 μL of D2O in a final volume of 600 μL. For samples containing β-amylase, 100 μL of the stock solution was replaced with 100 μL of the final purified sample. Wilmad 507-PP NMR tubes were used. All NMR data sets were collected on a Bruker Avance III 500 MHz NMR spectrometer at 298 K using a 5 mm BBFO room temperature probe. All 1H NMR spectra were collected with 16 scans. The water resonance was suppressed using excitation sculpting with gradients (24).

Biochemistry Laboratory: Cytochrome c Cytochrome c is a small, soluble heme-containing protein that shuttles electrons from Complex III to Complex IV in the mitochondrial electron transport chain. Cytochrome c is also involved in apoptosis when released from the mitochondria (26). The 1H NMR spectrum of the oxidized form of the protein is shown in Figure 6C. The spectrum took just under one hour to acquire which afforded a sequestered learning environment to discuss the basics of protein NMR spectroscopy with the team of students (27). By this point in the semester the students have been thoroughly introduced to amino acids and protein structure in the co-requisite lecture. The central themes of the small-group discussion were two-fold. First, since proteins are polymers of organic compounds, the 1H NMR spectrum should look like that of a huge organic molecule. Chemical shift ranges for aliphatic, aromatic, amide, and alpha protons were discussed. Some examples of 2D protein NMR spectra from on-going research projects were discussed in the context of removing overlap and making sense of such crowded spectra. Second, the concept of ring current and other anisotropic effects were discussed in the context of a ‘folded’ protein and the relationship between resonance dispersion and ‘structure’. A range of interest was encountered in this small group discussion, but the students began to appreciate the complexities and capabilities of protein NMR spectroscopy and how it is being applied to challenging problems today. The students were not disappointed when the first spectrum finished and the guesstimated ‘thousand-or-so’ resonances are observed as shown in Figure 6C. For most students, this is their first experience with resonances with a negative chemical shift. The high-field signals at -2.55 and -2.79 ppm most likely arise from side chain protons of Met-80 that coordinates the heme iron. NMR spectra were then collected in the presence of 20% and 40% methanol which is known to destabilize and denature proteins (28). Progressive loss of structure is observed going from 20% methanol (Figure 6B) to 40% methanol (Figure 6A) by the loss of resonance dispersion and by the appearance of several sharp resonances between 6.9–8.5 ppm that likely arise from increased rotation of the heme as the protein denatures. This series of spectra provides a very good demonstration of the hydrophobic effect driving protein folding. It can also be very satisfying to collect a 1H NMR spectrum on commercial cytochrome c for comparison (data not shown).



Figure 6. 1H NMR spectra of cytochrome c in the presence of (A) 40% methanol, (B) 20% methanol, and (C) buffer. High-field resonances at -2.55 and -2.79 ppm that are indicative of folded protein structure in the absence of methanol are indicated by arrows. These resonances shift and disappear with added methanol. Sharp resonances between 6.9–8.5 ppm that appear with added methanol, likely arising from increased rotation of the heme, are marked by asterisks.

Experimental Details for Cytochrome c Cytochrome c was purified from bovine heart using a combination of homogenization, centrifugation, ammonium sulfate precipitation, cation exchange chromatography, and gel filtration chromatography (29). Fractions from the gel filtration column were pooled and concentrated to about 4 mL final volume, resulting in a final protein concentration of approximately 230 μM. NMR samples with 0%, 20%, and 40% methanol were then prepared by combining 360 μL of the cytochrome c solution and 60 μL of D2O with appropriate amounts of methanol-d4 and water to give 600 μL final volumes. The approximate final concentration of cytochrome c in these samples was 140 μM. Wilmad 528-PP NMR tubes were used. NMR data sets were collected on a Bruker Avance III 500 MHz NMR spectrometer at 298 K using a 5 mm BBFO room temperature probe. All 1H NMR spectra were collected with 1,024 scans. The water resonance was suppressed using excitation sculpting with gradients (24). 42



Undergraduate Research: Uridine Nucleoside Ribohydrolase Trichomoniasis affects an estimated 170 million people worldwide with as many as 85% of cases occurring in developing nations (20). More than 1 million new cases are reported each year in the United States, with infection prevalence estimated to be almost 4 million (30). Trichomoniasis is typically treated with the 5-nitroimidazole drug metronidazole, the first-line treatment used in the United States since the 1960s (21, 31). Resistance to metronidazole and related 5-nitroimidazole drugs is increasing, with an estimated 5% of trichomoniasis clinical cases resulting from T. vaginalis strains with some resistance (21). New targets with a defined mechanism of action are needed to develop novel antitrichomonal agents. T. vaginalis is an obligate parasite in that it is incapable of the de novo synthesis of purine (32) and pyrimidine rings (33, 34). It must scavenge nucleosides from host cells and then use salvage pathway enzymes to obtain the nucleobases. The parasite requires the activity of these enzymes to metabolize the nucleosides. The first step in this pathway is the hydrolysis of nucleosides to release the nucleobases. Purine and pyrimidine nucleoside ribohydrolases (NHs) comprise a superfamily of structurally related calcium-dependent enzymes that hydrolyze the N-glycosidic bond of β-ribonucleosides producing ribose and the free nucleobase (35). The pyrimidine-specific uridine nucleoside ribohydrolase (UNH) and the purine-specific adenosine/guanosine preferring nucleoside ribohydrolase (AGNH) have distinct substrate specificities and thus recognize distinct pharmacophores. An NMR-centric approach is being used to identify and characterize inhibitors for both enzymes in the context of undergraduate research. An 19F NMR-based activity assay was developed for UNH, utilizing 5-fluorouridine in place of uridine as shown in Figure 7. NMR-based assays avoid many complications associated with screening compounds at high concentrations by other methods (17). While 1H NMR was sufficient to monitor the hydrolysis of the natural substrate uridine, it was found that monitoring the hydrolysis of 5-fluorouridine with 19F NMR was superior for several reasons. First, the 19F NMR spectra were much simpler because they were not complicated by the presence of test compound resonances or large signals from the non-deuterated DMSO used to plate some compounds. Second, the Km values for uridine and 5-fluorouridine were determined to be 54 μM and 15 μM, respectively, indicating that 5-fluorouridine would provide a lower concentration hit threshold compared to uridine (36).

Figure 7. Hydrolysis of 5-fluorouridine catalyzed by the enzyme uridine nucleoside ribohydrolase (UNH). 43 Soulsby et al.; NMR Spectroscopy in the Undergraduate Curriculum: Upper-Level Courses and Across the Curriculum Volume 3 ... ACS Symposium Series; American Chemical Society: Washington, DC, 2016.


UNH enzyme assays were carried out in batch mode to test multiple compounds or multiple concentrations simultaneously. Reactions typically ran for 30–45 minutes and were then quenched with HCl which inactivates the enzyme and allows data to be collected sequentially. The 19F NMR spectra at 0 minutes and 30 minutes are shown at the top and bottom of Figure 8, respectively. The 19F chemical shifts for the 5-fluorouridine substrate (-165.8 ppm) and 5-fluorouracil product (-169.2 ppm) are clearly resolved allowing either or both to be used to monitor the reaction. The reaction is about 50% complete after 30 minutes. This assay was used to screen the NIH Clinical Collection for inhibitors (37). Figure 8 also shows the 19F NMR spectra collected in the presence of 15 test compounds from the NIH collection screened in five mixtures of three test compounds each. The 19F spectra for Mixtures 1–4 resemble the 30 minute control indicating lack of UNH inhibition. No product signal is observed for Mixture 5, however, indicating the presence of an inhibitor. Mixture 5 was deconvoluted by testing the three compounds individually as shown in Figure 9. Lack of product signal in the spectrum for well G3 identifies the active compound, cefatrizine. Using this approach, a total of 23 compounds were identified that significantly inhibited UNH. Interestingly, the proton-pump inhibitors omeprazole, pantoprazole, and rabeprazole were among these (37). IC50 values (concentrations resulting in 50% inhibition) were determined for selected compounds using the same assay carried out on serially diluted compounds. For these experiments, however, compounds were obtained as solids and diluted accordingly with DMSO to provide final assay concentrations ranging from 200 μM to 0.04 μM. Dose-dependent assays for pantoprazole are shown in Figure 10. Curve-fitting of substrate peak intensities resulted in an IC50 value of 14.5 μM for pantoprazole. The 19F NMR-based activity assay continues to be used to test fragments and synthesized compounds to develop structure-activity relationships.



Figure 8. Representative 19F NMR spectra monitoring the UNH reaction in the presence of compound mixtures. Spectra are shown for 0 and 30 minute control solutions plus five compound mixtures. Resonances of the 5-fluorouridine substrate (S) and 5-fluorouracil product (P) are labeled in the 30 minute control spectrum. Lack of product resonance for Mixture 5 indicates that an inhibitor is present in this mixture. The additional resonance marked with an asterisk in the Mixture 2 spectrum arises from a fluorine-containing compound in this mixture.



Figure 9. 19F NMR spectra monitoring the UNH reaction for 0 and 45 minute control solutions and the three compounds in Mixture 5 from Figure 8. Resonances of the 5-fluorouridine substrate (S) and 5-fluorouracil product (P) are labeled in the 45 minute control spectrum. Lack of product resonance for G3 identifies this compound as the inhibitor in Mixture 5.



Figure 10. 19F NMR spectra monitoring the UNH reaction for 0 and 30 minute control solutions and various concentrations of pantoprazole. Resonances of the 5-fluorouridine substrate (S) and 5-fluorouracil product (P) are labeled in the 30 minute control spectrum. The amount of product formation decreases as inhibitor concentration increases. Experimental Details for Uridine Nucleoside Ribohydrolase Uridine nucleoside ribohydrolase (UNH) was cloned and expressed in E. coli and was purified using a combination of affinity chromatography and gel filtration chromatography (38). A stock assay solution containing 50 mM phosphate and 0.3 M KCl at pH 6.5, 80 nM UNH, and 10% D2O was prepared. Stock solutions of 10 mM 5-fluorouridine in buffer and 10 mM test compounds in DMSO were prepared. Reaction samples were prepared by first adding 5-fluorouridine and test compounds to microcentrifuge tubes in appropriate amounts to give final concentrations of 50 μM 5-fluorouridine and the desired amount of test compound in a final volume of 600 μL (50 μM for initial screens and deconvolution, variable amounts for the dose-dependent assays). The reaction was then initiated with the addition of stock assay solution, mixed twice by aspirating and dispensing, 47



and left to sit on the benchtop for 30–45 minutes. Reactions were then quenched with the addition of 10 μL 1.5 M HCl and transferred to Norell 502 NMR tubes for data collection. These inexpensive NMR tubes are sufficient for the simple 1D NMR experiments carried out. The tubes can be washed and reused about 10 times before becoming either chipped or too dirty. All NMR data sets were collected on a Bruker Avance III 500 MHz NMR spectrometer at 298 K using a 5 mm BBFO room temperature probe. The 19F{1H} NMR spectra were collected with 256 scans for the initial screening experiments and 1,024 scans for the dose-dependent assays. 19F chemical shifts were referenced to external 50 μM CF3CD2OH in the identical quenched buffer solution at -76.7 ppm.

Undergraduate Research: Adenosine/Guanosine Preferring Nucleoside Ribohydrolase It is hypothesized that UNH and AGNH represent targets that will be inhibited by distinct molecules. To examine this hypothesis, an NMR-based activity assay was developed to monitor the AGNH reaction. By analogy to the UNH screening assay, an initial attempt was made to monitor the AGNH reaction using 2-fluoroadenosine as the substrate. However, this compound was a surprisingly poor substrate with the reaction taking many hours to reach the midpoint. Since other fluorinated analogs of either adenosine or guanosine are not commercially available, a 1H NMR-based activity assay was developed instead using adenosine as the substrate (the Km value for adenosine is 54 μM) as shown in Figure 11 (39).

Figure 11. Hydrolysis of adenosine catalyzed by the enzyme adenosine/guanosine preferring nucleoside ribohydrolase (AGNH). Figure 12 shows the control 1H NMR spectra at 0 and 30 minutes. The reaction is about 50% complete during this time period. Drawbacks to the 1H NMR-based assay compared to an 19F NMR-based will be the large residual signal from the protonated DMSO used to plate the NIH Clinical Collections, and the potential overlap of substrate and product resonances by test compound resonances. Figure 12 suggests that both of these issues are surmountable. The dynamic range of the NMR spectrometer allows signals from both adenosine and adenine to be detected without distortion in the presence of the large DMSO signal (spectral region not 48



shown). Further, the reaction can be monitored using the resolved adenosine resonances at 6.10 and 8.48 ppm or the resolved adenine resonance at 8.33 ppm. It is unlikely that test compounds, even in mixtures of three, will overlap with all of these resonances simultaneously.

Figure 12. 1H NMR spectra of adenosine after incubating with AGNH for 0 and 30 minutes. Resolved resonances of the adenosine substrate (S) and adenine product (P) are labeled in the 30 minute spectrum. The unlabeled resonance in the 30 minute spectrum arises from both substrate and product. Experimental Details for Adenosine/Guanosine-Preferring Nucleoside Ribohydrolase Adenosine/guanosine preferring nucleoside ribohydrolase (AGNH) was cloned and expressed in E. coli and was purified using a combination of affinity chromatography and gel filtration chromatography (40). A stock assay solution containing 50 mM phosphate and 0.3 M KCl at pH 6.5, 45 nM AGNH, and 10% D2O was prepared. A stock solution of 5 mM adenosine in buffer was prepared. Reaction samples were prepared by first adding adenosine to microcentrifuge tubes to give a final concentration of 100 μM in a final volume of 600 μL. The reaction was then initiated with the addition of stock assay solution, mixed twice by aspirating and dispensing, and left to sit on the benchtop. Reactions were then quenched with the addition of 10 μL 1.5 M HCl and transferred to Norell 502 NMR tubes for data collection. All NMR data sets were collected on a Bruker Avance III 500 MHz NMR spectrometer at 298 K using a 5 mm BBFO room temperature probe. The 1H NMR spectra were collected with 64 scans.

Conclusion The NMR-based activity assays carried out in the biochemistry laboratory are straightforward and can be easily carried out for a given enzyme in a single afternoon. NMR spectroscopy provides a more tangible enzyme reaction assay than can be obtained using spectrophotometry or fluorescence spectroscopy. In the latter methods, the reaction time course is indicated by a change in 49 Soulsby et al.; NMR Spectroscopy in the Undergraduate Curriculum: Upper-Level Courses and Across the Curriculum Volume 3 ... ACS Symposium Series; American Chemical Society: Washington, DC, 2016.


absorbance attributable to some chemical species being consumed or created. In the NMR spectra, both the consumption of substrate and creation of product are directly observed. The association of observed signals with discrete chemical entities learned in organic chemistry laboratory is translated to enzyme-catalyzed chemical transformations. This concept should be widely applicable to other enzymes typically encountered in the biochemistry laboratory. NMR-based activity assays are also used routinely in the biotechnology and pharmaceutical industries. The more intensive setting of undergraduate research provides an opportunity to introduce students to the practical aspects of this approach in the context of a drug discovery project. The assays described here for UNH and AGNH are being used to screen chemistry space, define the pharmacophore of each enzyme, and interface with medicinal chemistry efforts. Students are driving the discovery of tool compounds to test the linkage between enzyme inhibition and antitrichomonal activity, and the discovery of lead chemical series to initiate structure-based drug design.

Acknowledgments I thank the many biochemistry laboratory students whose projects have been described here. The UNH and AGNH characterizations were carried out by Sierra Beck, Simona I. Bekker, Annie Laurie Benzie, Paola J. Burburan, Colleen S. Humes, Vivian N. Matubia, Samantha N. Muellers, Sandy S. Ramcharan, Irving Rosario Jr., Tara A. Shea, and Victoria L. Violo. The T. vaginalis drug discovery project is a collaboration with David W. Parkin and Melissa A. VanAlstine-Parris. Research has been supported by Faculty Development Grants and a Frederick Bettelheim Research Award from Adelphi University to BJS, and by Horace G. McDonell Summer Research Fellowships to SB and TAS.

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