Analytical Biochemistry 408 (2011) 226–234

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Analytical Biochemistry

journal homepage: www.elsevier .com/locate /yabio

Real-time bioluminescent assay for inhibitors of RNA and DNA polymerasesand other ATP-dependent enzymes

Kalvin J. Gregory ⇑, Ye Sun, Nelson G. Chen, Valeri GolovlevSci-Tec, Knoxville, TN 37932, USA

a r t i c l e i n f o a b s t r a c t

Article history:Received 11 June 2010Received in revised form 4 August 2010Accepted 12 August 2010Available online 18 August 2010

Keywords:Polymerase assayBioluminescenceReal-timeFragment-based drug discovery (FBDD)Viral polymerase inhibitors

0003-2697/$ - see front matter � 2010 Elsevier Inc. Adoi:10.1016/j.ab.2010.08.016

⇑ Corresponding author. Fax: +1 865 671 2167.E-mail address: kjgreg@aurogene.com (K.J. Gregor

1 Abbreviations used: NI, nucleoside inhibitor; NNI, noenzyme-linked immunosorbent assay; ECL, electrochempyrophosphate; HTS, high-throughput screening; FBDcovery; NMR, nuclear magnetic resonance; SPR, surfdithiothreitol; RLU, relative light units; CV, coefficient

Viral polymerases are important targets for drug development. However, current methods used to iden-tify and characterize inhibitors of polymerases are time-consuming, use radiolabeled reagents, and arecost-inefficient. Here we present a bioluminescent assay for the identification and characterization ofinhibitors of polymerases, as well as other ATP-dependent enzymes, that monitors the decrease of ATPor dATP in real time, allowing detection of enzyme inhibition based on differences in ATP/dATP consump-tion. The assay works with a variety of RNA and DNA polymerases, using both RNA and DNA templates.The assay measures changes in substrate concentration in real time and provides a faster alternative forkinetic studies of inhibition. Michaelis–Menten plots were obtained from a single reaction, yielding Km

values that compared well with literature values. The assay could identify the mechanism of inhibitionand determine inhibition constants (Ki) for a weak competitive inhibitor of Klenow fragment and twostrong noncompetitive inhibitors of HIV-1 reverse transcriptase with one series of inhibitor concentra-tions, reducing the total number of experiments that would normally be needed. The assay is also sensi-tive enough to detect a weak inhibitor with Ki > 100 lM, making it a viable technique for fragment-baseddrug discovery.

� 2010 Elsevier Inc. All rights reserved.

Viral polymerases are targets for antiviral drugs because of theimportant role they play in integrating the viral genome into thehost organism genome and in the production of new viral particles[1]. Both nucleoside and non-nucleoside inhibitors (NIs and NNIs,respectively)1 have been effective against HIV-1 [2], and thedevelopment of both classes of inhibitors remains an active field ofresearch [3]. In addition, an increasing number of both academicand industrial investigators are focusing on the discovery of NI andNNI compounds effective against other viruses such as HCV andHBV [4–9], herpes [10], and measles [11].

Virus adaptability, such as the ability of HIV-1 to rapidly devel-op resistance to drugs, is well known and is a constant challenge indeveloping new treatments [12–14]. Resistance to drugs active to-ward viral polymerases, as well as other drug targets, has been ob-served repeatedly in viruses such as HCV [15,16], HBV [17], HSV-1[18,19], and measles [20,21]. Clearly, there is an imperative to de-velop new DNA and RNA polymerase inhibitors to replace thosethat get selected out, to improve existing drug design methodolo-

ll rights reserved.

y).n-nucleoside inhibitor; ELISA,iluminescence; PPi, inorganicD, fragment-based drug dis-

ace plasmon resonance; DTT,of variation.

gies, and to develop more efficient and robust methods of detectingand characterizing potential polymerase inhibitors.

The technology used to screen for potential polymerase inhibi-tors has changed very little over the past two decades. The pre-dominant method of assaying polymerases is a primer extensionassay, which measures the incorporation of radiolabeled NTPs ordNTPs. Such assays require reagents in large quantities, are time-consuming, rely on radioactive labeling, and are not very adaptableto high-throughput applications. The cost of reagents for such as-says is high, and characterization of inhibitors is labor-intensive[5].

Some efforts to address the drawbacks in the primer extensionassay have been made. An enzyme-linked immunosorbent assay(ELISA)-based colorimetric assay for HIV-1, HIV-2, SIV-1, AMV,and M-MulV reverse transcriptases was developed by Roche [22].However, ELISA formats require immobilization of the target prod-ucts on a plate, incubation times of 30 min or longer for each anti-body used, and extensive washes to achieve a good signal/background ratio and acceptable reproducibility.

Merck Research Laboratories reported a reverse transcriptasepolymerase assay based on electrochemiluminescence (ECL) tech-nology [23]. Because ECL is an end-point assay, the assay is time-consuming and labor-intensive. A large number of replicates arerequired to identify the mechanism of inhibition. Finally, the pro-tocol includes seven major processing steps and a 3-h assay time,

Fig.1. Real-time bioluminescent polymerase assay principles. (A) Assay schematicin which polymerase and primer/template compete with luciferase for ATP or dATP.A higher polymerase activity results in a more rapid decline of luminescence overtime. (B) Sample real-time data (three replicates) showing luminescence measuredwithout T7 polymerase (curve 1) and with polymerase added (curve 2). (C)Michaelis–Menten plot generated from the polymerase reactions shown in panel Busing the data analysis algorithm. Both the reaction velocity and the substrateconcentrations are given in arbitrary units (a.u.). The assay can generate a completeMichaelis–Menten plot from a single microplate well (single reaction).

Real-time bioluminescent assay / K.J. Gregory et al. / Anal. Biochem. 408 (2011) 226–234 227

making implementation of the assay in a high-throughput formatdifficult.

Lopes and coworkers [24] reported a polymerase assay usingfluorescein-labeled primer/template in conjunction with an auto-mated DNA sequencer. Once again, this was an endpoint-type as-say that also required a separation step on a sequencing gel andrequired costly automated sequencing instrumentation to beadapted to high-throughput applications. The authors also men-tioned that the primer/template required several hours of coolingfor proper annealing.

Lahser and Malcom reported a three-enzyme assay systembased on luminescence detection [25]. The conversion of the poly-merase side product inorganic pyrophosphate (PPi) to ATP by ATPsulfurylase generated a luminescence signal with luciferin/lucifer-ase proportional to the amount of product generated by the poly-merase reaction. However, the requirement for three enzymesadds to the cost, the assay requires a-thio-ATP to avoid back-ground luminescence from the luciferin/luciferase, and the assaydoes not take full advantage of the real-time format (as demon-strated elsewhere in this article).

The development of new enzyme inhibitors is susceptible toproblems inherent in current high-throughput screening (HTS)strategies. Conventional HTS strategies suffer from a low hit rateas well as a failure to optimize many hits into a usable drug[26,27]. In addition, the optimization process tends to reducedrug-likeness as defined by Lipinski’s ‘‘rule of five,” thereby dimin-ishing the developability of the optimized drug [28,29]. Fragment-based drug discovery (FBDD) addresses the issues of hit rate andoptimizing leads to compounds with drug-like properties [30].FBDD identifies low-molecular-weight molecules (‘‘templates” or‘‘scaffolds”), which are weakly binding (Ki > 10 lM) compoundswith high ligand efficiency, and has a higher hit rate than conven-tional HTS methods [31]. However, the molecules being screenedbind very weakly, requiring very sensitive detection methods suchas nuclear magnetic resonance (NMR), high-throughput X-raycrystallography, and surface plasmon resonance (SPR). In addition,both the drug targets and the compounds being screened must beat high concentrations. Finally, these methods measure bindingevents only, and they do not measure biological activity. For thisreason, binding sites that result in modulation of activity mustbe known beforehand.

In this article, we describe a bioluminescent assay for detectingand characterizing inhibitors of polymerases. The assay works withthe major classes of polymerases and is based on a two-enzymesystem that includes a target enzyme (polymerase) and a detectionenzyme (luciferase). The luciferase produces luminescence propor-tional to the concentration of the substrate for the target enzyme.In the presence of the polymerase and a suitable template (Fig. 1A),the luminescence decreases in proportion to the polymerase activ-ity, producing a luminescence reaction progress curve (Fig. 1B) andallowing the easy identification of inhibitors. The consumption ofdATP/ATP can be monitored in real time, and these data can beused to generate a Michaelis–Menten curve from a single reaction(Fig. 1C). By running several parallel reactions with varying con-centrations of a known inhibitor, simple analytical and numericaldata analysis tools can be used to analyze the resulting reactionprogress curves and Michaelis–Menten plots to determine kineticparameters such as Km and Ki (inhibition constant). In addition, asingle set of these measurements can be used to determinewhether the inhibitor is noncompetitive, competitive, or uncom-petitive. The assay can detect weakly binding inhibitors(Ki = 105 ± 30 lM in our example) and, therefore, provides a goodalternative for FBDD for ATP- or dATP-dependent enzymes. The as-say can be carried out with any single tube or microplate lumino-meter, and it reduces reagent consumption because it can be run ina total volume of as little as 5 to 10 ll per replicate. Finally, this

assay reduces the time needed to screen a significant number ofcompounds. Data sufficient to determine Ki and the mechanismof inhibition of many inhibitors can be acquired from a singleexperiment in 1.0 to 1.5 h.

Materials and methods

Enzymes and reagents

Enzymes were obtained from the following manufacturers:DNA polymerase I (Klenow fragment), M-MulV reverse transcrip-tase, and phi6 RNA polymerase from New England Biolabs(Ipswich, MA, USA); HIV-1 reverse transcriptase from WorthingtonBiochemical (Lakewood, NJ, USA); and T7 RNA polymerase fromFermentas Life Sciences (Burlington, ON, Canada). Other reagentswere obtained from the following manufacturers: plasmid pTZ19Rand ExtremePure dNTPs/NTPs from Fermentas Life Sciences; poly-(dT) and poly-(rU) from Midland Certified Reagent (Midland, TX,USA); oligo-(dA18) from Operon (Huntsville, AL, USA); coenzymeA and the ATP Bioluminescence Assay Kit from Sigma–Aldrich(St. Louis, MO, USA); and efavirenz from Toronto ResearchChemicals (North York, ON, Canada). The inhibitor MT-4S

228 Real-time bioluminescent assay / K.J. Gregory et al. / Anal. Biochem. 408 (2011) 226–234

(3-aryl-2-methyl-2,3-dihydrobenzo[d]isothiazole-1,1-dioxide, U.S.patent 6562,850) was a gift from D. C. Baker (Department ofChemistry, University of Tennessee).

Instrumentation

All luminescence measurements were performed on an Orion IIMicroplate Luminometer (Berthold Detection Systems, Pforzheim,Germany) at 25 �C. All reactions were carried out in a Corning384-well low-volume microplate, and luminescence was measuredcontinuously using 1-s read times for 30 min to 1.5 h. Data wererecorded with Simplicity software (version 4.0) included with theluminometer.

Polymerase assays

Reaction buffers for each polymerase were supplied by themanufacturer as 10� concentrates. The final buffer compositionsfor each polymerase were as follows: for T7 RNA polymerase,40 mM Tris–HCl (pH 7.9), 6 mM MgCl2, 10 mM dithiothreitol(DTT), and 2 mM spermidine; for Klenow fragment, 50 mM Tris–HCl (pH 8.0), 5 mM MgCl2, and 1 mM DTT; for Phi6 RNA polymer-ase, 50 mM Tris–HCl (pH 8.9) and 50 mM ammonium acetate; forHIV-1 reverse transcriptase and M-MulV reverse transcriptase,50 mM Tris–HCl (pH 8.3), 75 mM KCl, 3 mM MgCl2, and 10 mMDTT.

The luciferase/luciferin (ATP Detection Kit, cat. no. FLAA, Sigma–Aldrich) was prepared according to the manufacturer’s instructionsand stored in aliquots at �20 �C until needed; from this point on-ward, it is referenced here as luc/luc. Each polymerase assay wascarried out in a total volume of 10 ll, and RNase-free H2O was usedfor dilutions where needed. Template negative controls were pre-pared for all polymerases by assembling the reactions describedbelow but leaving out the polymerase template. All assays wererun in triplicate, and unless stated otherwise, all reagent concen-trations given are for the final 10 ll of total volume.

For the T7 DNA-dependent RNA polymerase assay, polymerase(0.25 lg/ll) was combined with 1 ll of 10� reaction buffer, coen-zyme A (1 mM final concentration), and 0.5 lg of plasmid pTZ19Rand then was diluted to 8 ll. After incubation at 37 �C for 10 min,the reaction was initiated with 0.5 ll of luc/luc (diluted 1:25) and1.5 ll of NTPs (0.75 mM each of CTP, GTP, and TTP and 0.375 mMATP final concentration). Luminescence was recorded as describedabove.

The phi6 RNA-dependent RNA polymerase was assayed using0.5 lg of poly-(rU) template and 0.05 lg of oligo-(rA18) primer in1� buffer with MnCl2 added (1.5 mM final concentration). The pri-mer/template was annealed for 5 min at 90 �C and then on ice for3 min. After equilibration to room temperature, 0.22 lg/ll phi6polymerase was added to the total 10-ll reaction. After incubatingat 25 �C for 3 min, the reaction was triggered by adding ATP (finalconcentration 15 lM) and 1 ll of luc/luc. Luminescence was re-corded as described above.

Klenow fragment was assayed using a poly-(dT)/oligo-(dA18)template/primer. A volume of 1 ll of 10� reaction buffer was com-bined with 1.5 lg of poly-(dT) and 40 ng of oligo-(dA18) and thenwas diluted to a final volume of 7 ll. The primer was annealed at90 �C for 5 min and then on ice for 3 min. After equilibration toroom temperature, Klenow fragment was added (0.0125 lg/ll)with 1 ll of luc/luc. The reaction was then triggered with dATP (fi-nal concentration 8 lM). Luminescence was recorded as describedabove.

HIV-1 reverse transcriptase reactions were performed by com-bining the 10� reaction buffer with 1 lg of poly-(rU) and 1 lg ofoligo-(dA18), diluting to 7 ll, and annealing the primer/templateat 80 �C for 5 min and then on ice for 3 min. HIV-1 reverse

transcriptase was then added (0.027 lg/ll final) along with 1 llof luc/luc. M-MulV reverse transcriptase was assayed in a similarfashion but with 40 ng of oligo-(dA18) and 0.06 lg/ll enzyme.The polymerase reactions were triggered with 5 lM dATP (finalconcentration). Luminescence was recorded as described above.

Kinetic studies of polymerase inhibitors

Inhibition of Klenow fragment by dGTP was assessed by spikingdGTP (100–500 lM final concentrations) in the Klenow fragmentassay described above after the primer annealing step and additionof the polymerase. A further incubation at 20 �C for 10 min wascarried out before the addition of luc/luc and dATP.

The inhibition of HIV-1 reverse transcriptase by the noncompet-itive inhibitors efavirenz and MT-4S was assessed in a similar way.After the primer annealing step and the addition of the polymerase,the inhibitors were added in various concentrations (2–12.5 nM forefavirenz and 0.13–11 nM for MT-4S). The reactions were thenincubated for 10 min at 20 �C before the addition of luc/luc anddATP. Luminescence was recorded as described above. Preliminaryto each study, template-negative controls were run both with andwithout inhibitors in the concentration ranges used so as to verifythat no inhibition of luciferase was taking place.

Data analysis

The concentration of substrate [S] (where S = dATP or ATP) atany time is directly proportional to the luminescence intensityI(t), and o[S]/ot is proportional to oI(t)/ot. The reaction velocityfor the polymerase reaction is then given by

V ¼ � o½S�ot¼ � oIðtÞ

ot¼ VmaxIðtÞ

IðtÞ þ Kmþ bIðtÞ; ð1Þ

where b is a term that accounts for consumption of substrate byluciferase, as well as inhibition of luciferase due to PPi, o-luciferin,or thermal degradation, causing an apparent increase in the reac-tion velocity at high [S] that is not due to the polymerase reaction.The change in luminescence intensity oI(t)/ot was calculated by ourpreviously reported algorithm for fitting temporal kinetic data by atwo-exponential curve [32] and was used to generate velocity ver-sus substrate plots (Michaelis–Menten plots) from raw lumines-cence data that could be fit to Eq. (1). This equation was modifiedfor competitive, noncompetitive, and uncompetitive inhibition,and parameters of interest (Km and Vmax) were calculated by nonlin-ear least squares fits.

Results

Real-time bioluminescent detection of RNA and DNA polymerases

To demonstrate that the real-time bioluminescent assay wouldwork with different classes of polymerases, we selected one DNA-dependent DNA polymerase (Klenow fragment), one DNA-depen-dent RNA polymerase (T7 RNA polymerase), one RNA-dependentDNA polymerase (HIV-1 reverse transcriptase), and one RNA-dependent RNA polymerase (phi6 RNA polymerase). We focusedon commercially available polymerases that can serve as goodmodel enzymes to study, optimize, and validate this new assayas a platform for screening clinically relevant enzymes and enzymeinhibitors. Fig. 2 shows representative reaction progress curves forT7 RNA polymerase, Klenow fragment, HIV-1 reverse transcriptase,and phi6 RNA polymerase (panels A–D, respectively). Each curve isplotted as the logarithm of the luminescence signal (I) in relativelight units (RLU) versus time. As expected, the luminescencedecreased over time due to consumption of ATP/dATP by the

Fig.2. Real-time bioluminescent detection of different classes of polymerases, (A) DNA-dependent RNA polymerase (T7 RNA polymerase). (B) DNA-dependent DNApolymerase (Klenow fragment). (C) RNA-dependent DNA polymerase (HIV-1 reverse transcriptase). (D) RNA-dependent RNA polymerase (phi6 RNA polymerase). The blue(lower) curve denotes the sample containing polymerase and template, and the red (upper) curve indicates the template-negative control. (For interpretation of thereferences to color in this figure legend, the reader is referred to the web version of this article.)

Real-time bioluminescent assay / K.J. Gregory et al. / Anal. Biochem. 408 (2011) 226–234 229

polymerase. The template negative control for each assay, in con-trast, remained very steady throughout the duration of the mea-surement, resulting in a typical dynamic range of 2.5 to 3.0 logsbetween the template-negative control and the template-positivereaction. Experiments with other polymerases, including M-MulVreverse transcriptase and Bst DNA polymerase (data not shown),produced similar results but do suggest that variations in this dy-namic range are dependent on the particular polymerase beingstudied.

For the T7 reaction, we chose the plasmid pTZ19R, which con-tains a T7 initiation sequence. Because this template was nothomopolymeric in nature, it was necessary to use an NTP mix,not just ATP. The assay worked in a broader dynamic range (i.e.,no steady-state luminescence background after the reaction hadbeen exhausted) when the ratio of the concentration of ATP was1:2 relative to the other NTPs. In the case of the other polymerases,we chose a more universal template, either poly-(rU) or poly-(dT),with a primer consisting of either oligo-(dA)18 or oligo-(rA)18,depending on the requirements of the polymerase. Because the as-say depends on the monitoring of dATP or ATP incorporation exclu-sively, the use of poly-(dT) or poly-(rU) yields a more efficient andless expensive assay because the other dNTPs or NTPs are not re-quired. Both dATP and ATP worked well in this assay, so that it isapplicable to both DNA and RNA polymerases. ATP is a much bettersubstrate for luciferase than dATP, with dATP producing onlyapproximately 5% of the Vmax produced by ATP [33], and in practicewe found it necessary to dilute the luc/luc, whereas dilution wasnot necessary in the dATP-based assays.

The luciferase reaction can be inhibited by o-luciferin, whichcan compete with luciferin for its binding site. In both the ATP-and dATP-based assays, the luminescence signal of the template-negative control did decrease over time, although the differencebetween the total decrease of it and that of the positive controlswas always approximately 2.5 to 3.0 logs. For the ATP-based T7 as-say, the decrease in the template-negative control luminescence

could be stabilized with 1 mM coenzyme A, which forms a complexwith o-luciferin, thereby freeing up the luciferase active site [34].However, the addition of coenzyme A to the dATP-based assaymade no noticeable difference. According to Steghens and cowork-ers [35], luciferase appears to have a separate binding site for dATP,and this could possibly explain the lower efficiency of coenzyme Ato enhance the stability of the template-negative control signal.Under the conditions listed here, the assay could measure ATP con-centrations as low as 30 pM and dATP concentrations down to200 pM.

Estimation of the Michaelis constant Km

The data analysis algorithm was used to determine the steady-state kinetic constant Km, yielding values that compared well withvalues reported in the literature. The Km value, or any other con-stant such as kcat or kcat/Km, varies depending on what templateis being used or which dNTP/NTP the constant is being reportedfor. Templates may be homo- or heteropolymeric, and the Km fora given dNTP/NTP may vary significantly depending on the tem-plate [36]. In this assay, the template was always oligo-(dA18 orrA18)/poly-(dT or rU), with the exception of the double-strandedheteropolymeric T7 template, and the calculated Km values werealways based on dATP or ATP incorporation. The substrate/velocitydata generated by the data analysis algorithm were fitted to Eq. (1)by nonlinear least squares, and the resulting Km values in arbitraryunits were calibrated against the known starting concentrations ofdATP/ATP. Table 1 shows values calculated for Klenow fragment,HIV-1 reverse transcriptase, T7 RNA polymerase, and M-MulVreverse transcriptase versus reported literature values (those ob-tained from Klenow fragment [37,38], HIV-1 reverse transcriptase[36,39,40], T7 RNA polymerase [41], and M-MulV [42]). The litera-ture values reported were for dTTP incorporation into poly-(dA)templates except for the T7 value, which measured ATP incorpora-tion specifically, and so this can be regarded as only an

Table 1Comparison of calculated Km values with literature values.

Polymerase Km (lM) Literature value(s) (lM)a

Klenow fragment 1.2 ± 0.3 1–2HIV-1 reverse transcriptase 2.0 ± 0.3 2.1–6.3T7 RNA polymerase 58 ± 5 35b

M-MulV reverse transcriptase 13 ± 2 18

a See text for literature references.b The reported value was said to vary by a factor of 2.

230 Real-time bioluminescent assay / K.J. Gregory et al. / Anal. Biochem. 408 (2011) 226–234

approximate comparison, although the resulting Km values are inreasonable agreement with the reported values.

Real-time bioluminescent kinetic studies of a weak competitiveinhibitor

In FBDD, the small-molecular-weight scaffolds that arescreened are typically very weak binders [43]. To demonstrate thatthe real-time bioluminescent assay had the sensitivity required tobe used in FBDD, we selected dGTP as a weak competitive inhibitorfor Klenow fragment. For a poly-(dT) template, dGTP functions as acompetitive inhibitor. Fig. 3A shows the reaction progress curvesfor competitive inhibition of Klenow fragment by dGTP. The datawere normalized to a starting value of 1.0 by dividing the lumines-cence intensity at a given time (I) by the initial luminescenceintensity (Io), and then plotted on a semi-logarithmic scale. Asexpected, dGTP weakly inhibited Klenow fragment in a dose-dependent manner, with the time required for full consumptionof dATP increasing with an increasing concentration of inhibitor.

Fig.3. Competitive inhibition of Klenow fragment by the weak inhibitor dGTP. (A)Luminescence versus time plots of Klenow fragment polymerization reactions withan oligo-(dA18)/poly-(dT) primer/template and 0 to 500 lM dGTP (‘‘Neg” denotesthe template-negative control). Luminescence data were plotted as luminescence attime t, I divided by initial luminescence, Io, and scaled semi-logarithmically. Dataare representative of three separate experiments. (B) Michaelis–Menten plots foreach reaction generated with the data analysis algorithm, where both the reactionvelocity V and the dATP concentration are in arbitrary units (a.u.).

Reaction velocity versus substrate data for each reaction wasgenerated using our data analysis algorithm, and the resultingMichaelis–Menten plots are shown in Fig. 3B. Here both the reac-tion velocity and the substrate concentrations, which were calcu-lated from the raw luminescence data, are in arbitrary units. Theplots shown here very clearly indicate competitive inhibition.The steepness of each curve is decreasing (due to an increasingKm) with an increasing concentration of dGTP, whereas at highersubstrate concentration the curves are clearly converging to thesame Vmax. Each curve was generated from one reaction, whereasa conventional kinetic study would require a separate reactionfor each data point.

In competitive inhibition, the apparent Km, K�m, increases withinhibitor concentration by a factor of (1 + [I]/Ki), where [I] is theinhibitor concentration and Ki is the competitive inhibition con-stant. Km and K�m values were determined by fitting the data inFig. 3B to Eq. (1) (modified for competitive inhibition) by nonlinearleast squares. The ratio K�m=Km was plotted versus [dGTP] (lM),and Ki was determined by linear least squares (Fig. 4). From thisplot, a Ki of 105 ± 30 lM was obtained.

Comparison of two strong competitive inhibitors

To demonstrate that the real-time bioluminescent assay couldidentify noncompetitive inhibitors and directly compare twoinhibitors, kinetic studies were carried out with the noncompeti-tive HIV-1 reverse transcriptase inhibitors efavirenz and MT-4S(see Materials and methods for information on MT-4S). The dataanalysis algorithm was used to generate Michaelis–Menten plotsfrom the raw luminescence data for efavirenz (Fig. 5A) and MT-4S (Fig. 5B). Noncompetitive inhibition is suggested for both inhib-itors, with the Vmax values noticeably decreasing with an increasein inhibitor in both cases.

In noncompetitive inhibition, the reciprocal of the apparentmaximum velocity 1=V�max increases by a factor of (1 + [I]/Ki),where both [I] and Ki are defined as before. Values of V�max weredetermined by nonlinear least squares for each concentration ofinhibitor by fitting the data in Fig. 5A and B to Eq. (1) modifiedfor noncompetitive inhibition. For both inhibitors, the ratioð1=V�maxÞ=ð1=VmaxÞ, where Vmax is the maximum velocity with noinhibitor, was plotted versus the inhibitor concentrations for efavi-renz (Fig. 6A) and MT-4S (Fig. 6B). The Ki values (inverse of the

Fig.4. Calculation of an inhibition constant Ki for the weak competitive inhibitordGTP against Klenow fragment. Shown is a plot of the ratio K�m=Km versus [dGTP](lM), where Km is the Michaelis–Menten constant with no inhibitor added and K�mis the apparent Michaelis–Menten constant in the presence of a competitiveinhibitor. The inhibition constant Ki is the inverse of the slope and was determinedto be 105 ± 30 lM. All error bars denote standard deviations.

Fig.5. Noncompetitive inhibition of HIV-1 reverse transcriptase by efavirenz andMT-4S. (A) Michaelis–Menten plots for efavirenz. (B) Michaelis–Menten plots forMT-4S. Polymerase reactions were run with an oligo-(dA18)/poly-(rU) primer/template. Data are representative of three separate experiments. Michaelis–Mentenplots were generated using the data analysis algorithm, where both the reactionvelocity V and the dATP concentrations are in arbitrary units (a.u.).

Fig.6. Calculation of inhibition constants Ki for the strong noncompetitive inhib-itors efavirenz and MT-4S against HIV-1 reverse transcriptase. (A) Plot of the ratioð1=V�maxÞ=ð1=VmaxÞ versus [efavirenz] (nM), where 1/Vmax is the inverse of themaximum reaction velocity with no inhibitor added and 1=V�max is the same ratioin the presence of a noncompetitive inhibitor. The inhibition constant Ki is theinverse of the slope. Here Ki was determined to be 3.0 ± 0.3 nM. (B) The same plotfor MT-4S. Here Ki was determined to be 2.40 ± 0.06 nM. All error bars denotestandard deviations.

Real-time bioluminescent assay / K.J. Gregory et al. / Anal. Biochem. 408 (2011) 226–234 231

slope 1/Ki) were determined by linear least squares to be 3.0 ± 0.3and 2.40 ± 0.06 nM for efavirenz and MT-4S, respectively.

Graphical verification of mechanism of inhibition

The mechanisms of inhibition for dGTP, efavirenz, and MT-4Swere verified by a simple graphical analysis of the raw lumines-cence data. In the region of the data where ATP or dATP has largelybeen consumed, an asymptotic trend line can be drawn; on a semi-logarithmic plot of luminescence versus time, this is representedby a straight line. Fig. 7A shows theoretical plots of trend linesfor competitive, noncompetitive, and uncompetitive inhibition. Incompetitive inhibition (panel 1), the trend lines intersect at a com-mon point corresponding to the unchanging Vmax. The vertical axisintercepts, however, decrease as Km increases with inhibitor con-centration. In noncompetitive inhibition (panel 2), the lines con-verge to the same Km on the vertical axis but do not intersectbecause Vmax decreases. In uncompetitive inhibition (panel 3), bothVmax and Km vary, yielding the parallel lines seen in the figure. Notethat this is entirely analogous to determining an inhibition mech-anism by comparing Lineweaver–Burk plots. Fig. 7 illustrates thismethod for competitive inhibition of Klenow fragment by dGTPand noncompetitive inhibition of HIV-1 reverse transcriptase byefavirenz (panels B and C, respectively). The luminescence datafor the noncompetitive inhibitor MT-4S yielded similar results tothose of efavirenz (data not shown).

Discussion

A novel polymerase assay based on real-time monitoring ofsubstrate consumption with bioluminescence detection has been

demonstrated as an alternative assay for use in conventionalscreening of polymerase inhibitors, a new approach to general ki-netic studies of polymerases, and a potential method for use inFBDD for polymerases. The assay cuts down on reagent costs con-siderably, using low volumes (5–10 ll) as well as correspondinglysmaller amounts of polymerase, dNTPs/NTPs, and the like. None ofthe reagents used was radioactively labeled, and the assay does notrequire specialized equipment, using a microplate or single tubeluminometer.

The assay worked with both RNA and DNA polymerases usingeither RNA or DNA templates. The template could be either homo-or heteropolymeric. Monitoring the consumption of dATP or ATP inreal time yields data from which kinetic parameters of interest,such as Km, can be extracted. Normally, in the determination ofsteady-state kinetic parameters, each point on a Michaelis–Mentenplot represents a separate reaction and one whole plot representsseveral separate reactions. In contrast, in the assay presented here,one Michaelis–Menten plot represents only one separate experi-ment. Clearly, this implies a significant reduction in cost as wellas time.

The performance of the assay is dependent on which enzyme isused, the enzyme concentration, and the potency of the inhibitor.The assay was able to detect polymerase concentrations in thenanomolar range. However, to give the highest possible changein signal between a positive (template added) and negative (notemplate) control, enzyme concentrations in the range of a fewhundred picomolars were used, so that inhibition could readilybe observed, the inhibitors could be characterized, and the assaycould be completed in a reasonable amount of time. The potency

Fig.7. Graphical verification of inhibition mechanisms. (A) Asymptotic extrapolations of the low substrate concentration regions of raw luminescence data for competitiveinhibition (1), noncompetitive inhibition (2), and uncompetitive inhibition (3). (B) Competitive inhibition of Klenow fragment by dGTP. On a semi-logarithmic plot, theextrapolated lines intersect at (I/Io) = 1, and they intercept the vertical axis at successively lower values of (I/Io), characteristic of competitive inhibition (unchanging Vmax andincreasing Km). (C) Noncompetitive inhibition of HIV-1 reverse transcriptase by efavirenz. The lines do not intersect, and they converge at the same (I/Io) values (Km isunchanged and Vmax decreases), characteristic of noncompetitive inhibition.

232 Real-time bioluminescent assay / K.J. Gregory et al. / Anal. Biochem. 408 (2011) 226–234

of a given inhibitor will determine its detection limit. For example,for inhibitors with Ki values in the nanomolar range (efavirenz andMT-4S), inhibitor concentrations in the range of 0.1 to 1 nM couldbe detected. Weak inhibitors with large Ki values (in this casedGTP) will have much higher detection limits. Theoretically, aninhibitor with a picomolar Ki could probably be detected in thepicomolar or sub-picomolar range.

In the results presented here, the more potent inhibitors yieldedbetter reproducibility between assays. The Ki values were averagevalues calculated from three independent experiments along withthe propagated error. The reproducibility of results, as measuredby the coefficient of variation (CV), seemed to improve with lower(more potent) Ki values. The most potent inhibitor, MT-4S, had a Ki

of 2.40 ± 0.06 nM (CV = 2.5%), whereas efavirenz yielded a Ki of3.0 ± 0.3 nM (CV = 10%). Although dGTP was measured against adifferent enzyme, it is useful to compare the CV of a very poorinhibitor in the micromolar range with more potent inhibitors inthe nanomolar range. In this case, dGTP yielded a Ki of105 ± 30 lM (CV = 29%). However, this should not be surprising.As discussed above, the less potent inhibitors will be more difficultto detect by any assay. Therefore, it is not surprising to observebetter statistical performance between assays when working withstrong inhibitors. The assay was able to produce a Ki value for

efavirenz that was within reasonable agreement with the literaturevalue of 2.93 ± 0.17 nM obtained through a typical radiolabelednucleotide incorporation assay [44].

The data obtained for competitive inhibition of Klenow frag-ment with dGTP demonstrates that this assay is a very good candi-date for use in FBDD for polymerases as well as ATP-dependentenzymes in general. Leads detected with FBDD will generally beweak binders and require very sensitive methods of detection.The currently used methods, as discussed in the introductory para-graphs, are both costly and cumbersome. The assay presented hereis far cheaper and does not require any specialized instrumentationbeyond a simple microplate or single tube luminometer. Despitethis, it was able to detect a competitive inhibitor with a high Ki

value (105 ± 30 lM). Some recent data have indicated Ki valuesin excess of 1000 lM. In addition, no information about the Klenowbinding sites that result in modulation of activity was requiredbecause this assay measures the change in activity.

Determination of the mechanism of inhibition with this assaywas straightforward, with one set of reactions with varyinginhibitor concentrations being sufficient to produce a series ofMichaelis–Menten plots for each inhibitor. By observing the effectof the inhibitor on the Michaelis–Menten plots and applying thesimple graphic analysis method described in this article, the

Real-time bioluminescent assay / K.J. Gregory et al. / Anal. Biochem. 408 (2011) 226–234 233

different modes of inhibition can readily be determined. In addi-tion, through simple least squares fitting of the Michaelis–Mentendata to equations of competitive, noncompetitive, and uncompeti-tive inhibition, the relative potencies of a group of inhibitors can becompared. Because the data analysis algorithm used here greatlyreduces the number of experiments required to get meaningfuldata, inhibition studies for several molecules can easily be runsimultaneously on a 384-well plate, and this assay can conceivablybe scaled down to a 1536-well plate.

Obviously, this assay could be adapted to an HTS format. Wepreviously reported a method for reduction of sample evaporation[45], so that an open-well format compatible with automatedinjectors could be used with small volumes. Therefore, scalingdown this reaction and using it for HTS should be readily achiev-able. One pitfall in using this two-enzyme system in an HTS formatis the possibility that a given inhibitor may interfere with lucifer-ase itself, resulting in a false hit. The most likely interference wouldbe NIs that can compete well with ATP or dATP for binding to lucif-erase. In such cases, validation of hits would be necessary. Theadvantage here is that the real-time assay measures differences be-tween samples and whatever controls are used; therefore, hits canbe validated by factoring out the lower luciferase efficiencythrough normalization of the raw luminescence data [I(t)/I(t = 0)].For the case in which substantial inhibition of luciferase is ob-served, there are alternative phenotypes of luciferase that havedemonstrated inhibition profiles that differ greatly from thePhotinus pyralis luciferase [46].

Acknowledgments

This work was supported in part by National Institutes ofHealth (NIH) Small Business Innovation Research (SBIR) GrantsR43AI082745 and R43CA141858.

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