A continuous fluorescence assay for polymerase activity

Little is known about the kinetic limitations of the polymerase chain reaction (PCR). Advancements in chemistry and instrumentation have increased its speed and specificity. Further improvements will be facilitated by a more complete understanding of the rates of the individual reactions that comprise PCR. A continuous fluorescent assay is developed to study DNA polymerase extension. Nucleotide incorporation is monitored with noncovalent DNA dyes using a defined hairpin template. The extension rate is measured in nucleotides incorporated per second per molecule of polymerase and has greater relevance to PCR than traditional activity methods. This assay was developed and validated on a stopped-ow instrument and subsequently adapted for real-time PCR instruments to extend its utility to any laboratory setting. The influences of a variety of buer components were determined and optimal conditions for fast polymerase extension are recommended. The incorporation rates of each nucleotide were determined and extension was found to depend on template sequence. When DMSO was included in the reaction to reduce inhibition from secondary structure, extension rates of random sequences were closely approximated by their base composition. Extension rates as a function of temperature were determined and were applied to a kinetic model. This model accounts for extension during temperature transitions and more accurately portrays fast PCR with rapid thermal cycling. A complete model of PCR based on differential equations derived from mass action equations is provided. This can be used to incorporate experimentally derived parameters obtained for the other reactions of PCR. Knowledge of the temperature and chemistry dependence of reaction rates will enable improved thermal cycling and solution conditions for more rapid and effcient PCR.

A CONTINUOUS FLUORESCENCE ASSAY FOR POLYMERASE ACTIVITY by Jesse L. Montgomery A dissertation submitted to the faculty of The University of Utah in partial ful llment of the requirements for the degree of Doctor of Philosophy Department of Bioengineering The University of Utah December 2013 Copyright c Jesse L. Montgomery 2013 All Rights Reserved The Univers i ty of Utah Graduate School STATEMENT OF DISSERTATION APPROVAL The dissertation of Jesse L. Montgomery has been approved by the following supervisory committee members: Carl T. Wittwer , Chair 10/28/2013 Date Approved Vladimir Hlady , Member 10/28/2013 Date Approved David W. Grainger , Member 10/28/2013 Date Approved Robert A. Palais , Member 10/28/2013 Date Approved James N. Herron , Member 10/28/2013 Date Approved and by Patrick A. Tresco , Chair/Dean of the Department/College/School of Bioengineering and by David B. Kieda, Dean of The Graduate School. ABSTRACT Little is known about the kinetic limitations of the polymerase chain reaction (PCR). Advancements in chemistry and instrumentation have increased its speed and speci city. Further improvements will be facilitated by a more complete understanding of the rates of the individual reactions that comprise PCR. A continuous uorescent assay is developed to study DNA polymerase extension. Nucleotide incorporation is monitored with noncovalent DNA dyes using a de ned hairpin template. The extension rate is measured in nucleotides incorporated per second per molecule of polymerase and has greater relevance to PCR than traditional activity methods. This assay was developed and validated on a stopped- ow instrument and subsequently adapted for real-time PCR instruments to extend its utility to any laboratory setting. The in uences of a variety of bu er components were determined and optimal conditions for fast polymerase extension are recommended. The incorporation rates of each nucleotide were determined and extension was found to depend on template sequence. When DMSO was included in the reaction to reduce inhibition from secondary structure, extension rates of random sequences were closely approximated by their base composition. Extension rates as a function of temperature were determined and were applied to a kinetic model. This model accounts for extension during temperature transitions and more accurately portrays fast PCR with rapid thermal cycling. A complete model of PCR based on di erential equations derived from mass action equations is provided. This can be used to incorporate experimentally derived parameters obtained for the other reactions of PCR. Knowledge of the temperature and chemistry dependence of reaction rates will enable improved thermal cycling and solution conditions for more rapid and e cient PCR. CONTENTS ABSTRACT : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : iii LIST OF FIGURES : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : vii LIST OF TABLES: : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : ix ACKNOWLEDGMENTS : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : x CHAPTERS 1. INTRODUCTION : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : 1 1.1 Advancements in the Speed of PCR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 1.2 The Need for Speed . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 1.3 A More Accurate Model of PCR Kinetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 1.4 The Standard Polymerase Activity Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 1.5 Alternative Polymerase Activity Assays . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 1.5.1 Quench-Flow . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 1.5.2 Stopped-Flow . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 1.5.3 Microtiter Plate Format . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 1.5.4 Benchtop Fluorometer and Spectrophotometer . . . . . . . . . . . . . . . . . . . 5 1.5.5 Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 1.5.6 Quartz Crystal Microbalance . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 1.5.7 Atomic Force Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 1.6 A Continuous Fluorescence Assay Using Noncovalent Dyes . . . . . . . . . . . . . . 7 1.7 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 2. STOPPED-FLOW DNA POLYMERASE ASSAY BY CONTINUOUS MONITORING OF DNTP INCORPORATION BY FLUORESCENCE : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : 16 2.1 Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17 2.2 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17 2.3 Materials and Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17 2.3.1 Oligonucleotides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17 2.3.2 DNA Polymerases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18 2.3.3 Polymerase Extension Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18 2.3.4 Bu er Component Study . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18 2.4 Results . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18 2.4.1 Polymerase Quanti cation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 18 2.4.2 Assay Validation and Calibration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19 2.4.3 Substrate Exhaustion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19 2.4.4 Calibration with Standards . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19 2.4.5 Polymerase Comparison . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 20 2.4.6 Bu er Components . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21 2.5 Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22 2.6 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23 3. THE INFLUENCE OF PCR REAGENTS ON DNA POLYMERASE EXTENSION RATES MEASURED ON REAL-TIME PCR INSTRUMENTS : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : 24 3.1 Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25 3.2 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25 3.3 Materials and Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25 3.3.1 DNA Polymerases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25 3.3.2 Polymerase Extension Template . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26 3.3.3 Polymerase Extension Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26 3.3.4 PCR Dyes and Additives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26 3.3.5 Hot Start Activation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26 3.3.6 Assay Calibration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26 3.3.7 Polymerase Activity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26 3.3.8 Extension Rates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27 3.4 Results . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27 3.5 Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29 3.6 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 31 4. THE INFLUENCE OF NUCLEOTIDE SEQUENCE AND TEMPERATURE ON THE ACTIVITY OF THERMOSTABLE POLYMERASES : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : 32 4.1 Abstract . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32 4.2 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33 4.3 Materials and Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33 4.3.1 DNA Polymerases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33 4.3.2 Oligonucleotides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33 4.3.3 Polymerase Extension Assay . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 34 4.3.4 Assay Calibration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35 4.4 Results . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35 4.5 Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 38 4.6 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 41 5. COMPARISON OF KINETIC AND EQUILIBRIUM MODELS FOR POLYMERASE EXTENSION : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : 51 5.1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51 5.2 Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51 5.2.1 Number of Nucleotides Extended with a Kinetic Model . . . . . . . . . . . . . 52 5.2.2 Number of Nucleotides Extended with an Equilibrium Model . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 52 5.3 Results . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 53 5.4 Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 53 5.5 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 55 v 6. CONCLUSION : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : 59 6.1 Future Work . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60 6.1.1 Rates for a Kinetic Model of PCR . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60 6.1.2 Expanded Bu er Component Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . 61 6.1.3 Expanded Polymerase Studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 62 6.1.4 Reverse Transcriptase Polymerases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 62 6.1.5 Structure-Function Relationships for Nucleotide Incorporation . . . . . . . 62 6.2 References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 63 APPENDICES A. OLIGONUCLEOTIDE TEMPLATE SEQUENCES : : : : : : : : : : : : : : : : : 64 B. A PROPOSED KINETIC MODEL OF PCR : : : : : : : : : : : : : : : : : : : : : : : 68 C. LINEARITY OF FLUORESCENCE WITH LENGTH OF DOUBLE-STRANDED DNA : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : : 72 vi LIST OF FIGURES 1.1 Comparison of equilibrium and kinetic models of PCR . . . . . . . . . . . . . . . . . . . 15 2.1 Quanti cation of polymerases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 19 2.2 Linearity of extension rates with polymerase concentration . . . . . . . . . . . . . . . 19 2.3 Quantitative analysis of polymerase extension curves . . . . . . . . . . . . . . . . . . . . 20 2.4 Extension rates of polymerases in the common and vendor bu ers . . . . . . . . . . 20 2.5 Measured extension rates (nt/s) versus calculated speci c activity (U/mg) . . . 20 2.6 E ects of bu er components on KlenTaq extension rates . . . . . . . . . . . . . . . . . 21 3.1 Validation of a polymerase extension rate assay that mimics PCR with in- creased uorescence from dye incorporation performed on a real-time PCR instrument . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27 3.2 E ect of monovalent cations on extension rates . . . . . . . . . . . . . . . . . . . . . . . . 27 3.3 E ect of Tm depressors on extension rates . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28 3.4 E ect of DNA dyes on extension rates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28 3.5 Optimal conditions for fast polymerase extension . . . . . . . . . . . . . . . . . . . . . . . 29 3.6 Extension rates after activation with hot start polymerases and nucleotides . . 29 4.1 Polymerase template designs for sequence dependence studies . . . . . . . . . . . . . 44 4.2 Agarose gels of templates with varying sequences after extension at 75 C . . . . 45 4.3 Extension rates as a function of temperature for hairpin templates with varying GC contents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 46 4.4 Incorporation rates of individual nucleotides as a function of temperature . . . . 47 4.5 Comparison of measured extension rates and rates predicted from base sequence 48 4.6 Comparison of predicted extension rates with measured rates in the presence of increasing DMSO . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49 4.7 Extension rates as a function of temperature for linear templates with varying primer melting temperatures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 50 5.1 Implementing a kinetic model for polymerase extension using extension rates . 56 5.2 Implementing a kinetic model for polymerase extension using saturation rates 57 5.3 Comparison of polymerase extension calculated with kinetic and equilibrium models . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 58 C.1 Oligonucleotides used to assess linearity of uorescence with length of double- stranded DNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 73 C.2 Fluorescence of hairpin oligonucleotides increases linearly with the length of double-stranded DNA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 74 viii LIST OF TABLES 2.1 Polymerase concentrations of stock solutions purchased from the manufactur- ers and their recommended concentrations in PCR . . . . . . . . . . . . . . . . . . . . . . 19 4.1 Extension inhibition by secondary structure and oligonucleotide probes . . . . . 43 A.1 Hairpin templates with varying GC contents . . . . . . . . . . . . . . . . . . . . . . . . . . 65 A.2 Hairpin template with secondary structure . . . . . . . . . . . . . . . . . . . . . . . . . . . . 66 A.3 Hairpin template with oligonucleotide probe . . . . . . . . . . . . . . . . . . . . . . . . . . . 66 A.4 Hairpin templates with single-base repeats . . . . . . . . . . . . . . . . . . . . . . . . . . . . 66 A.5 Linear templates with primers having varying melting temperatures . . . . . . . . 67 ACKNOWLEDGMENTS This work was made possible by the kind support of many individuals. I extend my gratitude to my fellow lab mates who have provided a positive atmosphere for me to conduct my studies. They each have o ered many words of encouragement and a fresh and optimistic perspective on my challenges. I consider them my dear friends. A special thanks is due to Luming Zhou who has taught me many principles of mental discipline and perseverance required of great scientists. Words are inadequate to describe my great debt to my advisor and mentor. He has been an outstanding example of patience and generosity, one that I strive to emulate. I am grateful for the assistance of my family who have eagerly relieved me of many burdens to allow me to continue my work. Most of all, I thank my wife who has sustained me through every di culty and whose faith in my abilities has given me strength beyond my own. CHAPTER 1 INTRODUCTION 1.1 Advancements in the Speed of PCR The introduction of the polymerase chain reaction (PCR) revolutionized molecular biology. First developed in the mid-1980s [1], this technique allowed analysis of DNA in a fraction of the time and with orders of magnitude greater sensitivity than other methods available at the time such as cloning and Southern blotting [2]. PCR continues to bene t from technological advancements, increasing its speed and sensitivity. Initially, thermal cycling was accomplished by manually transferring reaction tubes between water baths set at di erent temperatures [3]. Because a heat-labile polymerase was used (commonly the Klenow fragment of DNA polymerase from Escherichia coli ), additional polymerase was required after each cycle. PCR typically required more than 4.5 h for 35 cycles. Time requirements for PCR were greatly reduced with the use of a thermostable DNA polymerase from Thermus aquaticus [4]. Because the polymerase did not need to be replenished, PCR could be automated with block thermal cycling machines. This allowed PCR to be accomplished in less than half the time, with 35 cycles typically completed in 1.5 to 2 h. Further advancements were accomplished with instrumentation. Rather than thermal cycling microcentrifuge tubes with metal blocks, reactions have been placed in thin glass capillaries and heated and cooled with forced air [5], [6]. This enabled PCR in less than 15 min. Not only did rapid thermal cycling reduce time requirements, it also improved PCR by amplifying DNA with greater yield and speci city [7]. Additional instrumentation con gurations have been developed. These include micro u- idic and continuous ow designs [8]{[14], shuttle PCR [15]{[17], magnetic ow systems with ferro uid [18], micro-electro-mechanical systems (MEMS) [19], [20], and droplet and emulsion PCR [21]{[23] including ampli cation on beads for second generation sequencing [24], [25]. Thermal cycling has been accomplished with thin- lm resistors [26], [27], infrared lamps [28]{[31], electrically conductive bu ers [32], and liquid gallium [33]. These designs address a number of factors including throughput, cost, and miniaturization. However, the 2 common objective of most instrumentation modi cations is to reduce time requirements. Advancements in PCR speed have been accomplished with several systems, demonstrating DNA ampli cation in under 5 min [13], [16], [18], [27], [29], [33]. PCR will continue to improve through advancements in both chemistry and instrumentation. 1.2 The Need for Speed Fast PCR is not just a luxury, but an essential asset in medical diagnostics. This is especially true for infectious disease. Hospital acquired infections have become a great burden on the healthcare system. These occur in about 1 in 20 hospitalizations [34] with one- third of those resulting in readmission [35]. The annual cost of these infections in the United States is estimated at between 5 and 10 billion dollars [36]. This has led many hospitals to implement screening procedures for a variety of problematic infections. Screening is only feasible with rapid and cost-e ective molecular assays. High sensitivity, low cost, and speed of PCR-based methods make them especially suited for this purpose. These have been used for screening of vancomycin-resistant Enterococcus [37], [38], Clostridium di cile [39], methicillin-resistant Staphylococcus aureus [40]{[43], Klebsiella pneumoniae [44], [45], and Mycobacterium tuberculosis [46], [47]. Rapid diagnostics are also essential for surveillance and management of in uenza out- breaks. Clinics and laboratories are already heavily burdened with seasonal in uenza. Im- proved diagnostic tools are needed to cope with potential endemic and pandemic outbreaks. PCR-based methods are viable solutions as they require little expertise to perform and are quickly adaptable to rapidly evolving pathogens. Since the pandemic outbreak of 2009 in uenza A H1N1, PCR-based methods have become the primary techniques for tracking and diagnosing in uenza [48]. Increasing the speed of PCR will bring screening solutions closer to the point of care, improving management of infectious disease and decreasing healthcare costs. This will be facilitated by a thorough understanding of PCR kinetics. 1.3 A More Accurate Model of PCR Kinetics Understanding the kinetics of PCR is essential to realizing its full potential. PCR consists of three main reactions|denaturation of template DNA, annealing of DNA primers to the template, and extension of primed template by DNA polymerase. Ampli cation of DNA is accomplished by thermal cycling between the optimal temperatures for these reactions. Traditionally, PCR is viewed from an equilibrium paradigm (Figure 1.1). The three reactions are thought to occur independently at de ned temperatures. Only hold times at set temperatures are considered while transitions between temperatures are neglected. 3 However, PCR is more accurately considered from a kinetic paradigm, especially in the case of rapid thermocycling. More time is spent in transition between temperatures than at any individual temperature. The three reactions of PCR overlap with rates that vary as a function of temperature. A more complete understanding of PCR can be obtained by measuring the kinetics of each reaction in isolation. The rates of reactions can be determined in a variety of bu er con- ditions over a range of temperatures relevant to PCR. This would allow identi cation of rate limiting processes and optimal conditions. Improved bu er chemistries and instrumentation with optimal thermal cycling parameters can be developed to achieve PCR with maximum speed and speci city. The work presented here isolates DNA polymerase extension and measures the rates of this reaction under a variety of conditions. This is accomplished with the development of a new assay for polymerase activity. This assay represents a signi cant improvement over the standard method used to measure polymerase activity. 1.4 The Standard Polymerase Activity Assay The most common assay for polymerase activity was rst used in the discovery of DNA polymerase [49]. Enzyme fractions were combined with radiolabelled dNTPs and calf thymus DNA as the extension template. Polymerization of the radiolabelled dNTPs formed an acid-insoluble product. After incubating the reaction for 30 min, the reaction was quenched. The DNA product was precipitated with acid, rinsed, and assayed for radioactivity. Radioactivity measurements were correlated to polymerase activity. This method is in frequent use today and has remained almost unchanged. It has been used to characterize the activity of a wide variety of polymerases [50]{[54], and is currently the only assay used to measure the activity of polymerases used for PCR. Early in the development of this assay, activated DNA was established as the extension template [55]. Activated DNA is prepared by enzymatic digestion or mechanical shearing of genomic DNA, usually from salmon sperm or calf thymus. The result is a random and heterogenous mixture of single-stranded and double-stranded DNA. The template for extension cannot be standardized. This is quite di erent from PCR in which polymerase extends a template of de ned length and sequence. Some studies have altered the radioactive assay by using a de ned template with primers [56]{[58]. One study compared activity measured with activated DNA and a de ned template for two polymerases [56]. Both polymerases exhibited changes in activity with temperature optimums di ering by as much as 10 C between the two templates. Activity measurements using a de ned template will 4 more accurately re ect the kinetics of PCR. The standard radioactive assay provides end-point measurements. It is incapable of providing initial rates of nucleotide incorporation that exist during PCR. In addition, the units of activity are not intuitive. Activity is described as the amount of radiolabelled nucleotides converted into acid-precipitable material in a given length of time. With such an obscure de nition, it is di cult to know how a polymerase will perform in ampli cation of a de ned template during PCR. 1.5 Alternative Polymerase Activity Assays While the standard radioactive assay is commonly used to characterize the activity of polymerases, it is not commonly used in detailed kinetic studies. This is likely due to the drawbacks of a heterogenous template and end-point measurements. Several additional assays have been developed. 1.5.1 Quench-Flow Quench- ow is capable of monitoring fast reactions. Reactants are rapidly mixed and quenched with ethylenediaminetetraacetic acid (EDTA) at speci ed time points. Either radiolabeled dNTPs or primers are used. Products are collected on lter paper or resolved on polyacrylamide gels and the radioactivity assessed. Radioactivity as a function of time is plotted and t to kinetic models to obtain nucleotide incorporation rates. This technique has been used to measure extension rates of DNA polymerases [59], [60], RNA polymerases [61]{[63], and reverse transcriptase enzymes [64], [65]. Detailed kinetic mechanisms have been elucidated with this method. However, it is laborious and time-consuming and unlikely to be adopted as a general activity assay. 1.5.2 Stopped-Flow Continuous monitoring of product formation makes stopped- ow methods a more con- venient alternative to quench- ow. There is no need for radiolabeled reagents or analysis of products on gels. Several methods have been used to monitor polymerase extension. The uorescent base analog 2-aminopurine has been placed within the extension region of the template [61], [66]{[68]. The uorescence of the base is quenched upon incorporation of the complementary nucleotide and can be monitored during extension. Rates can only be measured for bases complementary to 2-aminopurine. Incorporation of nucleotides is accompanied by the release of pyrophosphate. One study took advantage of this to follow polymerase extension indirectly [63]. An enzyme-coupled 5 reaction converted pyrophosphate to a product that could be measured by absorbance. Another study used uorescence polarization [69]. The extension template was labeled with uorescein. As the template was elongated, the end of the strand became more restricted and anisotropy increased. These assays are capable of measuring fast reactions, but require modi ed template or additional reagents not used in PCR. 1.5.3 Microtiter Plate Format Activity assays adapted for microtiter plates greatly increase throughput. The stopped- ow uorescence polarization assay discussed previously was initially developed on a uo- rescence microplate reader [70]. Because the speed of polymerization was faster than the time required for temperature equilibration, this assay was only capable of end-point mea- surements. Misincorporation of noncomplementary bases could be monitored in real-time. A radioactive assay was developed for a microtiter plate format [71]. This measures the incorporation of radiolabelled dNTPs into extension templates that are immobilized on well surfaces. The reaction is quenched with EDTA at di erent time points and the radioactivity measured. This is not a continuous assay and multiple experiments must be performed to obtain kinetic data. 1.5.4 Benchtop Fluorometer and Spectrophotometer The accessibility, convenience, and cost of assays are improved with the use of common laboratory instrumentation. Polymerase activity assays have been developed for uorom- eters and spectrophotometers available to most laboratories. One assay relied on the intrinsic uorescence of single-stranded binding protein [72]. Saturating amounts of the protein were added to the extension template. As the template is extended, single-stranded binding protein is displaced and the intrinsic uorescence of the protein increases. An enzyme-coupled reaction was developed for a spectrophotometer [73]. Like the stopped- ow assay discussed earlier, pyrophosphate was converted to a product that could be monitored by absorbance. Activated calf thymus DNA was used as the template. Both of these are continuous assays capable of providing initial rates but are complicated by the need for additional reagents. Fluorescent probes were used to screen the activity of T7 RNA polymerase variants [74]. Molecular beacons were designed complementary to transcripts formed by RNA polymerase from a synthetic DNA template. Molecular beacons hybridize to the transcripts as they are produced, resulting in an increase in uorescence. Activity is expressed as the number of molecular beacons recognizing a transcript per min per mg of polymerase. Absolute 6 extension rates are not obtained. Activity measurements obtained with this assay may be in uenced by hybridization kinetics of the molecular beacon. Another spectrophotometer assay was based on light scattering [75]. As the template is extended, the scattering of the reaction solution increases by 10%. Despite being a contin- uous assay, no kinetic constants were calculated. This may have been due to di culties in calibration because of high light scattering background. 1.5.5 Microscopy The activity of RNA polymerase was measured with light microscopy [76]. Polymerase molecules were attached to glass coverslips by nonspeci c adsorption and extension tem- plates were attached to gold particles. Extension was monitored by acquiring images averaged over 2 s intervals. The length of extension was correlated to the range of Brownian motion of the particles. Lengths of extension as a function of time provided extension rates. The e ect of nonspeci c adsorption on activity of the polymerase is unknown. The activity of the Klenow Fragment polymerase was measured with uorescence mi- croscopy [77]. Template was immobilized on glass coverslips. The template contained a hairpin structure downstream from the primer. The hairpin kept a uorophore and a quencher in close proximity. As the polymerase extended the template, the hairpin was opened and uorescence increased. Gradual opening of the hairpin during extension allowed real-time monitoring of nucleotide incorporation. An optical trap assay was used to measure the extension rate and mechanical force associated with extension of T7 polymerase [78]. Both ends of the template were attached to beads. One end was immobilized with a glass pipette and the other was held with an optical trap. Extension was monitored by imaging the distance between the beads as a function of time. These distances were compared to the distance of beads with double or single stranded DNA to obtain extension rates. All of these methods obtain extension rates at a single-molecule level but are technically di cult to perform. 1.5.6 Quartz Crystal Microbalance Quartz crystal microbalance uses thin quartz wafers to measure changes in mass. The quartz oscillates with an applied alternating current. The frequency of oscillation decreases as mass is accumulated. This technique was used to monitor extension of immobilized tem- plate [79]. Klenow Fragment was rst equilibrated to achieve a baseline. Then dNTPs were added and the change in frequency observed. Calibration provides correlation of changes in frequency to increases in mass. This allowed calculation of nucleotide incorporation rates 7 from the initial slopes of frequency decreases during polymerase extension. In this study, the extension of the majority of templates could not be monitored. This may be due to reduced sensitivity as the signal from addition of nucleotides was small compared to the signal from the binding of polymerase. 1.5.7 Atomic Force Microscopy Atomic force microscopy was used to measure transcription of a template by an RNA polymerase [80]. Polymerases were attached to mica surfaces by nonspeci c adsorption. Surfaces were scanned in a ow-through system following addition of dNTP. Templates of known length were observed at di erent time points as they were extended by the polymerase. This technique su ers from low time resolution and the concentration of dNTP was intentionally kept low to reduce extension rates. Adsorption of the polymerases in the presence of high concentrations of zinc may have an impact on polymerase activity. 1.6 A Continuous Fluorescence Assay Using Noncovalent Dyes All of the polymerase activity assays described here use reagents that are foreign to PCR. In contrast, the assay introduced here is a continuous uorescent assay that monitors nucleotide incorporation with noncovalent uorescent dyes used ubiquitously in real-time PCR. The template has a de ned length and sequence. This allows simple calibration of data and provides activity in units of nucleotides per second per molecule of polymerase and are intuitive within the context of PCR. 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In the equilibrium model of PCR, denaturation, annealing, and extension are treated as independent reactions. Reactions occur at de ned temperatures and rates are constant. Temperature changes are assumed instantaneous. The kinetic model of PCR accounts for continuously changing temperatures with temperature-dependent rate constants and more adequately describes rapid PCR. CHAPTER 2 STOPPED-FLOW DNA POLYMERASE ASSAY BY CONTINUOUS MONITORING OF DNTP INCORPORATION BY FLUORESCENCE Reprinted from Analytical Biochemistry, vol. 441, no. 2, J. L. Montgomery, N. Rejali, and C. T. Wittwer, \Stopped- ow DNA polymerase assay by continuous monitoring of dNTP incorporation by uorescence," pp. 133-139, 2013, with permission of Elsevier. Stopped-flow DNA polymerase assay by continuous monitoring of dNTP incorporation by fluorescence Jesse L. Montgomery, Nick Rejali, Carl T. Wittwer ⇑ Department of Pathology, University of Utah, Salt Lake City, UT 84132, USA a r t i c l e i n f o Article history: Received 17 April 2013 Received in revised form 3 July 2013 Accepted 6 July 2013 Available online 16 July 2013 Keywords: Activity assay Polymerase Stopped-flow Intercalating dye a b s t r a c t DNA polymerase activity was measured by a stopped-flow assay that monitors polymerase extension using an intercalating dye. Double-stranded DNA formation during extension of a hairpin substrate was monitored at 75 C for 2 min. Rates were determined in nucleotides per second per molecule of poly-merase (nt/s) and were linear with time and polymerase concentration from 1 to 50 nM. The concentra-tions of 15 available polymerases were quantified and their extension rates determined in 50 mMTris, pH 8.3, 0.5 mg/ml BSA, 2 mM MgCl2, and 200 lM each dNTP as well as their commercially recommended buffers. Native Taq polymerases had similar extension rates of 10-45 nt/s. Three alternative polymerases showed faster speeds, including KOD (76 nt/s), Klentaq I (101 nt/s), and KAPA2G (155 nt/s). Fusion poly-merases including Herculase II and Phusion were relatively slow (3-13 nt/s). The pH optimum for Klentaq extension was between 8.5 and 8.7 with no effect of Tris concentration. Activity was directly correlated to the MgCl2 concentration and inversely correlated to the KCl concentration. This continuous assay is rel-evant to PCR and provides accurate measurement of polymerase activity using a defined template with-out the need of radiolabeled substrates. 2013 Elsevier Inc. All rights reserved. The extension rates of DNA polymerases under PCR conditions have not been characterized.Awidevariety of polymerases are avail-able and many are designed for increased fidelity and speed. The conventional way to measure the activity of DNA polymerase is in terms of units, most commonly defined as the number of nanomoles of radiolabeled dNTPs incorporated into activated calf thymus or sal-mon spermDNA at 72 to 75 C for 30 min. This is a time-consuming endpoint assay and does not provide information about the initial extension rates of polymerases. In addition, assay conditions are not standardized and often differ from those used during PCR. Anumberof alternative assays have been introduced forDNAand RNA polymerases. These include methods based on atomic force microscopy [1], light microscopy [2], single-molecule optical trap-ping [3], quartz crystal microbalance [4], and radiometric assays [5]. Others use enzyme-coupled reactions to monitor pyrophos-phate release [6,7]. Fluorescence-based methods have monitored the displacement of single-stranded DNA-binding protein [8] or polarization of labeled extension templates [9,10]. Quench-flow [6] has been used and allows kinetic analysis of rapid reactions. However, this method requires stopping the reaction at several time points, followed by analyzing the products on gels or by chromatog-raphy methods. Stopped-flow [6,9,11] assays have been developed and enable continuous reaction monitoring, but these use covalent fluorescent labels or nucleotide analogs. Some of these methods are capable of providing extension rates in terms of individual nucle-otide incorporation [1-3,8,9,11]. However, they all require template modifications (fluorescent or radioactive) or immobilization of either template or polymerase onto a substrate. We introduce a fluorescent stopped-flow assay for monitoring polymerase extension that requires no modification of the template or polymerase. This method relies on the increase in fluorescence of double-stranded DNA dyes during nucleotide incorporation. These dyes are frequently used in real-time PCR, eliminating the need to change reaction chemistry. Measured extension rates are directly applicable to PCR.Weuse this assay to compare the speed of 15 poly-merases at equimolar concentrations. Because their activity was strongly dependent on the reaction buffer, we then measured the ef-fects of common buffer conditions, including pH and KCl, MgCl2, and Tris concentration. Materials and methods Oligonucleotides The sequence tagcgaaggatgtgaacctaatcccTGCTCCCGCGGCCG atctgcCGGC-CGCGGGAGCA was used as the extension template and a baseline fluorescence standard (capital letters denote self-complementary sequences). The oligonucleotide forms a hairpin with a 14-bp stem that has a free 30 end and a 25-base overhang 0003-2697/$ - see front matter 2013 Elsevier Inc. All rights reserved. http://dx.doi.org/10.1016/j.ab.2013.07.008 ⇑ Corresponding author. Fax: [phone]. E-mail address: [e-mail] (C.T. Wittwer). Analytical Biochemistry 441 (2013) 133-139 Contents lists available at ScienceDirect Analytical Biochemistry journal homepage: www.elsevier.com/locate/yabio 17 for extension. The fully extended template was also synthesized as a fluorescence standard: TAGCGAAGGATGTGAACCTAATCCCTGCTC CCGC-GGCCGatctgcCGGCCGCGGGAGCAGGGATTAGGTTCACATCCT TCGCTA. Oligonucleotides were ordered from Integrated DNA Technologies with the extension substrate purified by high-pres-sure liquid chromatography and the fully extended standard puri-fied by polyacrylamide gel electrophoresis. Each was quantified by absorbance at 260 nm following digestion by purified phosphodi-esterase [12] for accurate quantification. DNA polymerases Fifteen polymerases were included in this study: Amplitaq (Invitrogen), KOD (EMD Millipore), Taq (New England Biolabs), Platinum Taq (Invitrogen), GoTaq (Promega), Titanium Taq (Clon-tech), Paq5000 (Agilent), Herculase II (Agilent), Phusion (New Eng-land Biolabs), KAPA2G (Kapa Biosystems), MyTaq (Bioline), Ex Taq (Clontech), Taq (Roche), SpeedSTAR (Clontech), and Klentaq I (pur-chased from either AB Peptides or Washington University in St. Louis, MO, USA). Polymerases were quantified on sodium dodecylsulfate (SDS) polyacrylamide gels stained with Sypro orange (Invitrogen). Gel images were obtained using a Gel Doc XR+ with XcitaBlue (Bio- Rad) conversion screen accessory and analyzed with Image Lab (Bio-Rad) software. Prior to being loaded on the gels, samples were reduced in 30 mM Tris, pH 6.8, 12.5% glycerol, 1% SDS, and 360 mM b-mercaptoethanol at 96 C for 5 min. Klentaq I (purchased from Washington University in St. Louis) was used as the quantification standard. The standard was quantified by absorbance at 280 nm using an extinction coefficient of 6.91 104 M 1 cm 1 calculated from the amino acid content of the published sequence [13]. The purity of this standard was determined by fluorescence integration from polyacrylamide gels and the concentration adjusted propor-tionately. Two replicates of each quantity standard (50, 100, 200, and 300 ng) and four replicates of each polymerase were included on each gel. Major bands at expected molecular masses were con-sidered to be the polymerase of interest. The integrated fluores-cence intensity of these bands was used to calculate the concentration and purity of the polymerases. Molecular masses used in concentration calculations were measured from the gels or taken from the literature [14,15], including vendor product information. Klentaq I was measured by mass spectrometry (Mass Spectrometry and Proteomics core facility at the University of Utah) after dialyzing for 48 h at room temperature in PBS buffer (150 mM NaCl, 2.5 mM KCl, 10 mM disodium phosphate, 1.5 mM dipotassium phosphate, pH 7.4). A molecular mass of 62,596 Da was determined compared to 62,097 Da predicted from the amino acid sequence [13]. The specific activity in units per milligram of polymerase (U/mg) was calculated from the unit concentration provided by the manufacturer and the concentration of polymerase measured on the gels. Most manufacturers define 1 U of polymerase as the amount required to incorporate 10 nmol of dNTPs in 30 min. How-ever, the manufacturers of Klentaq I (ABPeptides) and Taq (NEB) define a unit as the incorporation of 60 and 15 nmol dNTP, respec-tively. The specific activities of these polymerases were scaled to allow comparison to other polymerases (i.e., the specific activity calculated for Klentaq I was multiplied by 6 and that of Taq (NEB) was multiplied by 1.5). The specific activities for Herculase II and Titanium were not calculated because the manufacturers do not provide the polymerase activities. Polymerase extension assay Polymerase extension studies were performed with a stopped-flow instrument (SFM-300, Bio-Logic SAS). Excitation was set at 495 nm with a monochromator and fluorescence collected with a photomultiplier tube and a 530 ± 15 nm discriminating filter. Ther-moelectric heaters separately maintained the temperature of the mixing lines and the reaction cuvette. Each line was held at 75 C. Reactants were added to two separate mixing lines and mixed in a 1:1 ratio at a flow rate of 9 ml/s. The estimated dead time for mixing was 6.6 ms. Extension reactions were carried out in 1 EvaGreen (Biotium) and either the buffer supplied by the manufacturer of each polymerase or a common buffer (50 mMTris, 0.5 mg/ml bovine serum albumin (BSA), and 2 mM MgCl2, pH 8.3). When MgCl2 was not included in the vendor buffer (KOD and Plat-inum Taq), a final concentration of 2 mM was used. Preliminary experiments determined maximal extension rates for Klentaq I with 200 lM each dNTP with a Km of 39 lM. Polymerase extension was initiated by mixing 400 lMeach dNTP with 10 nM polymerase and 200 nM oligonucleotide (final concentrations were 200 lM each dNTP, 5 nM polymerase, and 100 nM oligonucleotide). To pre-vent template degradation, extension experiments for polymerases exhibiting 30 to 50 exonuclease activity (Herculase II, KOD, and Phu-sion) were initiated by mixing the polymerase with dNTP and oli-gonucleotide. MyTaq includes dNTPs in the vendor buffer at a final concentration of 250 lM each. For this polymerase, extension reactions were initiated by mixing the polymerase with the oligonucleotide. Polymerase extension curves were calibrated either by allowing the reaction to go to completion or by using fluorescence stan-dards. Except where indicated, calibration was performed with fluorescence standards. Polymerase was omitted from reactions containing fluorescence standards and calibration was repeated for each experiment to account for influences of buffer conditions on absolute fluorescence. Seven to ten stopped-flow shots were re-peated for each experiment and the means and standard deviations reported. Data were acquired for 2 min every 50 ms. Buffer component study The effects of pH and the concentrations of Tris, KCl, and MgCl2 on extension rate were observed. One parameter was varied while the other three were kept constant. Final conditions were Tris con-centrations at 10, 20, 30, 40, and 50 mM, KCl at 0, 12.5, 25, 37.5, 50, and 62.5 mM, and MgCl2 at 1, 1.5, 2, 2.5, 3, 4, 5, and 6 mM, and pH was at 7, 7.5, 8, 8.3, 8.5, 8.7, and 9. Unless varied, Tris concentration was held at 50 mM, KCl at 0 mM, and MgCl2 at 2 mM and the pH at 8.0. Studies were done with Klentaq I at 5 nM, 200 lM each dNTP, 100 nM template, 1 EvaGreen, and 0.5 mg/ml BSA final concen-trations. Reaction completion was used to calibrate the data in this study except when the extension was so slow that saturation could not be observedwithin 2 min. This occurred only when KCl concen-tration was 62.5 mM, the pH was 7, and MgCl2 concentration was 1 mM. Results Polymerase quantification A typical quantification gel and standard curve are shown in Fig. 1. Antibody hot-start polymerases (Platinum, Titanium, MyTaq, ExTaq, and SpeedSTAR) showed characteristic heavy- and light-chain bands at around 50 and 25 kDa. Paq5000 showed a promi-nent band of unknown identity at 62.5 kDa. Phusion had a diffuse band centered around 150 kDa and a prominent band at 64.5 kDa. Neglecting bands known to be other components, the purity of all polymerases was calculated at greater than 90%, with the excep-tion of KOD (70%), Phusion (50%), and KAPA2G (65%). Measured concentrations for all polymerases are shown in Table 1. Most 134 Fluorescent DNA polymerase assay / J.L. Montgomery et al. / Anal. Biochem. 441 (2013) 133-139 18 vendors supply polymerases at a concentration around 1 lM. Addi-tionally, the concentration of polymerase in PCR is typically in the range of 5 to 20 nM. Exceptions are KOD (94.5 nM), Klentaq I (63 nM), and Titanium (197 nM), which are supplied and used at considerably higher concentrations. Assay validation and calibration Extension rates were derived from the initial slope of the exten-sion curves. Fig. 2 shows that the initial slope is proportional to polymerase concentration from at least 1 to 50 nM. To obtain rates in absolute units, extension curves were calibrated in one of two ways. Substrate exhaustion Polymerase extension reactions are allowed to proceed to satu-ration with complete extension of the template. The maximum and minimum data points of individual extension curves are normal-ized between 0 and the total number of nucleotides that each poly-merase molecule can extend. This is calculated as: ½Template L=½Poly ; ð1Þ where [Template] is the concentration of template, L is the length of extension in base pairs, and [Poly] is the concentration of the poly-merase. Normalized this way, the initial slope of extension curves directly yields extension rate in nucleotides per second per mole-cule of polymerase (nt/s). Calibration with standards Extension curves can be normalized using oligonucleotide stan-dards (Fig. 3). The baseline fluorescence is measured from the extension template without polymerase present. A synthetic ana-log of the fully extended template is used as a maximum fluores-cence standard. The average fluorescence of the baseline standard is taken as 0 and the average fluorescence of the maxi-mum standards is scaled to the value calculated by Eq. (1). The same offset and scaling factor are also applied to each experimen-tal curve. Both analyses were compared using Klentaq I at 75 C. Ten experiments of 8 to 10 shots each were acquired. Substrate exhaustion yielded an extension rate of 102 ± 4.2 nt/s, whereas calibration with oligonucleotide standards gave 99 ± 8.4 nt/s. The standard deviations of individual shots within an experiment were similar for both methods at 3.8 and 3.4%, respectively. Both A B Fig.1. Quantification of polymerases. The purity and size of polymerases were determined on reducing polyacrylamide gels after staining with Sypro orange. (A) Quantification of SpeedStar. Four replicates of the SpeedStar (lanes 3, 6, 10, and 13) were compared to Klentaq I standards at 50 ng (lanes 2 and 4), 100 ng (lanes 5 and 7), 200 ng (lanes 9 and 11), and 300 ng (lanes 12 and 14). Molecular mass markers (Precision Plus Protein, Bio-Rad) are shown in lanes 1, 8, and 15. SpeedStar is an antibody hot-start polymerase and bands corresponding to heavy and light chains are shown near 25 and 50 kDa. The top band near 90 kDa is the polymerase. (B) Quantification of polymerases from a standard curve. The integrated fluorescence intensity of the unknown polymerase (squares) is projected on a regression line through the Klentaq I quantification standards (circles). The R2 of the regression line is 0.999. Table 1 Polymerase concentrations of stock solutions purchased from the manufacturers and their recommended concentrations in PCR. Polymerase Source Stock concentration (lM) Recommended PCR concentration (nM) Taq (NEB) Native Taq 1.06 ± 0.01 5.3 ± 0.1 Taq (Roche) Native Taq 0.26 ± 0.01 5.2 ± 0.1 Amplitaq Native Taq 0.52 ± 0.02 2.6 ± 0.1 GoTaq Native Taq 0.98 ± 0.08 4.9 ± 0.4 MyTaq Native Taq 1.31 ± 0.04 26.1 ± 0.8 ExTaq Native Taq 1.28 ± 0.08 10.2 ± 0.7 Platinum Native Taq 0.65 ± 0.02 2.6 ± 0.1 Herculase II Pfu fusion variants 1.9 ± 0.1 19.4 ± 1.3 Phusion Pfu fusion variants 1.01 ± 0.05 10.1 ± 0.5 Klentaq I [13] (ABPeptides) Deletion variant 39.4 ± 1.5 63 ± 2.5 Titanium [13] Deletion variant 9.9 ± 0.2 197 ± 3.6 KAPA2G Engineered Taq 1.11 ± 0.09 4.4 ± 0.4 Paq5000 [16] Pfu 0.85 ± 0.02 8.5 ± 0.2 KOD [15,17] Thermococcus kodakaraensis 4.7 ± 0.2 95 ± 4.1 SpeedStar Proprietary 1.08 ± 0.06 5.4 ± 0.3 Fig.2. Linearity of extension rates with polymerase concentration. The initial slope of polymerase extension curves is linear with polymerase concentration. A linear regression yields R2 = 0.999. Experiments were performed with Klentaq I. Fluorescent DNA polymerase assay / J.L. Montgomery et al. / Anal. Biochem. 441 (2013) 133-139 135 19 analysis methods were concordant to within 3%. The advantage of using standards is that reactions need not proceed to exhaustion, greatly reducing acquisition time when the activity is low. How-ever, increased precision makes substrate exhaustion preferable when the activity is high. Polymerase comparison The extension rates of various polymerases were measured in their corresponding vendor buffers as well as in a common buffer composed of 50 mM Tris, pH 8.3, 0.5 mg/ml BSA, 2 mM MgCl2, and 200 lM each dNTP (Fig. 4). Most native Taq polymerases var-ied in extension rate within a factor of 4. MyTaq showed the fastest performance in either buffer, whereas Platinum Taq was the slow-est. Overall, rates for the native polymerases were faster in the common buffer with an average of 31.3 nt/s compared to 25.8 nt/ s for the vendor buffers. The fusion variants had the slowest exten-sion rates. These are Pyroccocus furiosis (Pfu) polymerases fused to a double-stranded DNA binding domain intended to improve fidel-ity. Comparing these rates to that of PAQ5000, an unmodified Pfu polymerase [16], the fused domains hinder extension. The deletion Fig.3. Quantitative analysis of polymerase extension curves. Extension curves are analyzed in one of two ways: (1) by measuring the fluorescence of oligonucleotide standards without polymerase (the minimum standard is the extension template and the maximum standard is a synthetic oligonucleotide identical to the sequence of the fully extended template) or (2) by substrate exhaustion, using time 0 as the fluorescence minimum. In both cases, the maxima and minima are scaled between 0 and the total number of bases extended by each polymerase calculated using Eq. (1). The initial slope is then the extension rate in nucleotides per second per molecule of polymerase (nt/s). Both approaches yield extension rates concordant within 3%. Fig.4. Extension rates of polymerases in the common (black bars) and vendor (gray bars) buffers. Extension rates were strongly influenced by buffer conditions. KOD, Klentaq I, and KAPA2G were the fastest polymerases. Fig.5. Measured extension rates (nt/s) versus calculated specific activity (U/mg). The specific activity of each polymerase was calculated from the unit concentration supplied by the manufacturer and the mass of polymerase measured from polyacrylamide gels. There is little relationship between the two measurements of specific activity. The correlation is positive in the common buffer (circles), with a Pearson's r coefficient of 0.32, and negative in the vendor buffers (squares), with a Pearson's r coefficient of 0.30. 136 Fluorescent DNA polymerase assay / J.L. Montgomery et al. / Anal. Biochem. 441 (2013) 133-139 20 variants are mutants of Taq with a deletion of the 50 exonuclease domain [13]. These showed faster extension, especially in the com-mon buffer. KAPA2G, an engineered variant of Taq, showed the fastest extension rates. KOD is a polymerase from the Thermococcus kodakaraensis KOD1 archae [15,17]. In the vendor buffer, it was the third fastest polymerase. SpeedStar is a polymerase from an organ-ism undisclosed by the manufacturer, but extension rates are sim-ilar to those observed for the native Taq polymerases. The specific activity was calculated from unit concentrations provided by the vendor and the mass of polymerase measured from gels. For most polymerases the specific activity was between 40,000 and 65,000 U/mg. Specific activities were higher for Amplit-aq (103,300 U/mg), Platinum Taq (81,800 U/mg), and Taq (NEB) (75,600 U/mg). The specific activities for Phusion (21,900 U/mg) and KOD (5900 U/mg) were lower. Fig. 5 contrasts the measured extension rates to calculated spe-cific activities. These are analogous measurements of polymerase speed. Both are nucleotide incorporation rates normalized to the amount of polymerase used in the assay. In this study, extension rate is a measurement of the initial rate of nucleotide incorporation using a defined template and is expressed per molecule of poly-merase. Specific activity is the rate of nucleotide incorporation into activated DNA and is expressed per milligram of the polymerase. Pearson correlation coefficients were calculated to assess the linear relationship between these two measurements. The relationship is weakly positive when measured in the common buffer and weakly negative when measured in the vendor buffers. Buffer components The difference between extension rates in the common versus the vendor buffers for many polymerases is striking. Nearly a 3- fold increase with the common buffer was observed for Klentaq I. The enhancement of KOD and KAPA2G in the vendor buffer ap-proached 100-fold. It is apparent that buffer components strongly influence extension rates. A B C D Fig.6. Effects of buffer components on KlenTaq extension rates. (A) Tris has little effect. (B) KCl strongly inhibits extension. (C) Optimal pH for extension is between 8.5 and 8.7, with rapid decreases outside these values. (D) Magnesium increases extension rates with saturation near 5 mM. Fluorescent DNA polymerase assay / J.L. Montgomery et al. / Anal. Biochem. 441 (2013) 133-139 137 21 The effects of four common components of PCR buffers on extension rates were studied for Klentaq I (Fig. 6). The concentra-tion of Tris has very little influence on extension rates (Fig. 6A). KCl concentration inhibits polymerase activity (Fig. 6B). Extension rates decline linearly between 0 and 37.5 mM with over a 70% de-crease. Only 21% of total activity was measured at 50 mM. Optimal pH is between 8.5 and 8.7, with rapid decreases outside of this va-lue (Fig. 6C). Extension is almost entirely inhibited at pH 7. Rates quickly increase with total MgCl2 concentration (Fig. 6D), saturat-ing at 5 mM. At 1.5 mM MgCl2, extension rates were 43% of the maximum at 5 mM. Discussion The homogeneous stopped-flow assay presented here provides a simple and precise measurement of polymerase activity. The use of double-stranded DNA dyes allows continuous monitoring of extension. These dyes are commonly used in real-time PCR and eliminate the need for template modifications including covalent labels, radioactivity, or nucleotide analogs that are used in other assays. Template and buffer conditions reflect those found in PCR. Performance of polymerases can easily be tested under a vari-ety of conditions and can aid in screening polymerases and buffer conditions for various applications. EvaGreen was used in these studies and inhibits PCR with increasing concentration [18]. The same effect has been observed for SYBR Green I [19] and Syto 9 [20]. Comparative studies have shown that the degree of inhibition varies across dyes [18,20,21]. The effect of DNA dyes on polymerase activity has not yet been studied. In these experiments, template was in 20-fold excess of the polymerase and each polymerase molecule bound and extended multiple templates just as in PCR. It has been shown that template binding is not a limiting step in polymerase extension [22] and is not expected to contribute to the rates measured here. Extension rates are measured in nt/s and have greater relevance to PCR than the standard unit definition. PCR amplifies templates of defined length and knowledge of the extension rates in nt/s provides better insight into the speeds obtainable during PCR. For example, this could guide optimization of thermal cycling protocols for faster and more efficient PCR. As shown in Fig. 5, vendor-claimed specific activities correlate poorly with measured extension rate per molecule. These are both normalized measurements of the rate of nucleotide incorporation into a template and should be directly comparable. Extension rate is expressed per molecule of polymerase and specific activity is ex-pressed per milligram of polymerase. However, these normaliza-tion approaches are similar because the molecular masses of all the polymerases in this study other than Klentaq and Titanium are within about 4%. Poor correlation between the two measure-ments of activity can be attributed to differences in buffer condi-tions and extension templates. Buffers used in traditional radiometric assays for polymerase activity vary widely and differ-ences in pH, denaturants, and MgCl2, KCl, template, and dNTP con-centration may contribute to disagreement in specific activities reported for polymerases. The average specific activity calculated for the native Taq polymerases in this study is nearly fivefold lower than in a study that measured the specific activity of Taq polymer-ase at 292,000 U/mg [23] under different conditions. Wide variance in assay conditions complicates comparison of specific activities across studies. Different templates also introduce variability. Radiometric as-says use activated DNA, which is prepared with a variety of tech-niques including enzymatic digestion and mechanical shearing. This results in a heterogeneous template that does not reflect PCR conditions. PCR amplifies defined templates and is processive rather than random. One study compared the activity of polymer-ase with activated salmon sperm DNA and a defined template using single-stranded M13 with a primer [23]. The activity differed between the templates by about 60% at 70 C with Taq polymerase. Activity measurements of DNA polymerases will have greater rel-evance to their intended use if assay conditions are similar. Manufacturers claim superior speed for 7 of the 15 polymerases that were studied. Of the native Taq polymerases, fast extension rates are claimed only for MyTaq. This polymerase was the fastest in the category of native Taq polymerases and the fifth fastest poly-merase overall (Fig. 4). Fast extension rates are claimed for both the fusion polymerases, Herculase II and Phusion, though they were among the slowest polymerases studied. Phusion claims to be 10-fold faster than unmodified Pfu polymerase; however, Phu-sion was 13.5 nt/s while 28.2 nt/s was observed for Paq5000, a na-tive Pfu polymerase. Speed claims are also made for Paq5000, though this polymerase exhibited only moderate activity. KAPA2G and KOD both have fast extension rates as indicated by the manu-facturer, but this is dependent on the buffer used. SpeedStar did not demonstrate superior speed as claimed, with a maximum extension rate (31.1 nt/s) only marginally faster than the average extension rate of all native Taq polymerases (28.3 nt/s). The second fastest extension rate was observed with Klentaq I, though this is not generally considered a fast polymerase. Although the native Taq polymerases should be molecularly similar, their extension rates vary by nearly a factor of 3 in the common buffer. These differences indicate there is some variability in the activity of the same polymerase prepared under different conditions. Also, polymerase extension rates are strongly depen-dent on buffer conditions and the vendor buffer is not always opti-mal (Fig. 4). Faster speeds were observed in the common buffer (with 95% confidence) for both of the Taq polymerase deletion vari-ants and five of the seven native Taq polymerases-Taq (NEB), Taq (Roche), Amplitaq, GoTaq, and Platinum Taq. In contrast, faster extension rates were observed in vendor buffers for KAPA2G and KOD. KCL and MgCl2 concentration and pH greatly influence exten-sion rates. The range of optimal pH is narrow (Fig. 6C). Below the optimum of pH 8.5 to 8.7, extension rates declined about 60% with each pH unit. Above the optimum, the decline was nearly twice as rapid. Another study using a radiometric assay found that optimal pH was dependent on the buffer system [24]. Optimums for Tris, glycine, and potassium phosphate buffers ranged between pH 7.0 and 8.0. For each buffer system, rapid decreases in activity were also observed outside the optimal pH. The highest activity for the Tris buffer was at pH 7.8 and was lower than the pH optimum in our study. This buffer contained components not included in the Tris buffer we used, including 2-mercaptoethanol, KCl, and fivefold higher MgCl2 concentration. This suggests that other components in addition to the buffer system may also influence the optimal pH for extension. Extension rates continued to increase with MgCl2 concentration until saturating at 5 mM (Fig. 6D). This is a higher concentration than is typically used in PCR, often because of concerns with non-specific amplification. Greater specificity is achieved with faster thermal cycling [25-27] and higher MgCl2 concentration may be most appropriate in rapid PCR. KCl strongly inhibits extension (Fig. 6B). A number of methods have been used to study the effect of KCl concentration on poly-merase activity, including sequencing [28] and measuring the rate of incorporation of radiolabeled dNTPs [23,24]. The outcome of these studies varied with optimal activity at 0 mM [28], 60mM [24], or either 10 or 55 mM KCl depending on the template [23]. Two studies used a defined template with primers as opposed to activated DNA [23,28]. These also showed KCl inhibition with 138 Fluorescent DNA polymerase assay / J.L. Montgomery et al. / Anal. Biochem. 441 (2013) 133-139 22 activity greatest in the absence of KCl or at the lowest concentra-tion studied. The manufacturers of Taq (NEB), Taq (Roche), and Amplitaq dis-close the contents of their PCR buffers. Each has 10 mM Tris, 50 mM KCl, and 1.5 mM MgCl2 at pH 8.3. Speeds were similar in the vendor buffer, with extension rates between 15.1 and 23.4 nt/s. These polymerases were faster in the common buffer, presumably because of lower KCl concentration (0 mM) and higher MgCl2 concentration (2 mM). In the common buffer, Klentaq is nearly twofold faster than Titanium but has slightly lower activity in the vendor buffers. This behavior is not adequately explained by the influence of the components studied. Both vendor buffers have identical MgCl2 concentration. The pH of the Klentaq buffer is 9.1 and that of the Titanium buffer is 8.0. Our results in Fig. 6Cindicate that the pH's of both buffers are suboptimal. In addition, the Tita-nium buffer contains KCl at 16 mM, whereas the Klentaq buffer does not. Also included in the Klentaq buffer is ammonium sulfate at 16 mM, which was not studied here. Further studies of this com-ponent as well as other PCR additives will allow more complete elucidation of optimal buffer components. Buffer conditions affect the fidelity of nucleotide incorporation. For example, the rate of base substitution error increases fivefold for Taq polymerase when increasing MgCl2 concentrationfrom 1 to 5 mM [29]. In contrast, the error decreases threefold for Pfu polymerase over the same concentration range [30]. The pH of buf-fers has also been shown to positively and negatively affect fidelity [29-31]. For applications sensitive to nucleotide misincorporation, additional methods should be used to verify adequate fidelity. Accurate measurement of polymerase activity under PCR condi-tions has strong implications in achieving rapid PCR. Advance-ments in instrumentation continue to decrease thermal cycling times, allowing amplification within a few minutes [32-34]. Real-izing the full potential of PCR will require optimization of both instrumentation and chemistry. Conditions that are sufficient for standard PCR may not be well suited to very fast PCR. As cycling times are reduced, even small differences in activity may have an impact on the success of amplification. Measurements of activity are more relevant when defined in terms of nucleotides per second per molecule of polymerase rather than units per milligram. Accu-rate quantification of polymerase activity under optimal reaction conditions will facilitate PCR with maximum speed and efficiency. Acknowledgment This work was supported by a Grant from BioFire Diagnostics. References [1] S. Kasas, N.H. Thomson, B.L. Smith, H.G. Hansma, X. Zhu, M. 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Imanaka, Characterization of DNA polymerase from Pyrococcus sp. strain KOD1 and its application to PCR, Appl. Environ. Microbiol. 63 (1997) 4504-4510. [16] K.S. Lundberg, D.D. Shoemaker, M.W. Adams, J.M. Short, J.A. Sorge, E.J. Mathur, High-fidelity amplification using a thermostable DNA polymerase isolated from Pyrococcus furiosus, Gene 108 (1991) 1-6. [17] M. Nishioka, H. Mizuguchi, S. Fujiwara, S. Komatsubara, M. Kitabayashi, H. Uemura, M. Takagi, T. Imanaka, Long and accurate PCR with a mixture of KOD DNA polymerase and its exonuclease deficient mutant enzyme, J. Biotechnol. 88 (2001) 141-149. [18] F. Mao, W.Y. Leung, X. Xin, Characterization of EvaGreen and the implication of its physicochemical properties for qPCR applications, BMC Biotechnol. 7 (2007) 76. [19] C.T. Wittwer, M.G. Herrmann, A.A. Moss, R.P. Rasmussen, Continuous fluorescence monitoring of rapid cycle DNA amplification, Biotechniques 22 (1997) 130-131. 134-138. [20] P.T. Monis, S. Giglio, C.P. Saint, Comparison of SYTO9 and SYBR Green I for real-time polymerase chain reaction and investigation of the effect of dye concentration on amplification and DNA melting curve analysis, Anal. Biochem. 340 (2005) 24-34. [21] H. Gudnason, M. Dufva, D.D. Bang, A. Wolff, Comparison of multiple DNA dyes for real-time PCR: effects of dye concentration and sequence composition on DNA amplification and melting temperature, Nucleic Acids Res. 35 (2007) e127. [22] R.D. Kuchta, V. Mizrahi, P.A. Benkovic, K.A. Johnson, S.J. Benkovic, Kinetic mechanism of DNA polymerase I (Klenow), Biochemistry 26 (1987) 8410- 8417. [23] F.C. Lawyer, S. Stoffel, R.K. Saiki, S.Y. Chang, P.A. Landre, R.D. Abramson, D.H. Gelfand, High-level expression, purification, and enzymatic characterization of full-length Thermus aquaticus DNA polymerase and a truncated form deficient in 50 to 30 exonuclease activity, PCR Methods Appl. 2 (1993) 275-287. [24] A. Chien, D.B. Edgar, J.M. Trela, Deoxyribonucleic acid polymerase from the extreme thermophile Thermus aquaticus, J. Bacteriol. 127 (1976) 1550-1557. [25] T.G. Mamedov, E. Pienaar, S.E. Whitney, J.R. TerMaat, G. Carvill, R. Goliath, A. Subramanian, H.J. Viljoen, A fundamental study of the PCR amplification of GC-rich DNA templates, Comput. Biol. Chem. 32 (2008) 452-457. [26] C.T. Wittwer, D.J. Garling, Rapid cycle DNA amplification: time and temperature optimization, Biotechniques 10 (1991) 76-83. [27] C.T. Wittwer, B.C. Marshall, G.H. Reed, J.L. Cherry, Rapid cycle allele-specific amplification: studies with the cystic fibrosis delta F508 locus, Clin. Chem. 39 (1993) 804-809. [28] M.A. Innis, K.B. Myambo, D.H. Gelfand, M.A. Brow, DNA sequencing with Thermus aquaticus DNA polymerase and direct sequencing of polymerase chain reaction-amplified DNA, Proc. Natl. Acad. Sci. U.S.A. 85 (1988) 9436-9440. [29] K.A. Eckert, T.A. Kunkel, High fidelity DNA synthesis by the Thermus aquaticus DNA polymerase, Nucleic Acids Res. 18 (1990) 3739-3744. [30] J. Cline, J.C. Braman, H.H. Hogrefe, PCR fidelity of pfu DNA polymerase and other thermostable DNA polymerases, Nucleic Acids Res. 24 (1996) 3546- 3551. [31] K.A. Eckert, T.A. Kunkel, Effect of reaction pH on the fidelity and processivity of exonuclease-deficient Klenow polymerase, J. Biol. Chem. 268 (1993) 13462- 13471. [32] M. Hashimoto, P.C. Chen, M.W. Mitchell, D.E. Nikitopoulos, S.A. Soper, M.C. Murphy, Rapid PCR in a continuous flow device, Lab Chip 4 (2004) 638-645. [33] E.K. Wheeler, C.A. Hara, J. Frank, J. Deotte, S.B. Hall, W. Benett, C. Spadaccini, N.R. Beer, Under-three minute PCR: probing the limits of fast amplification, Analyst 136 (2011) 3707-3712. [34] C.T. Wittwer, R.P. Rasmussen, K.M. Ririe, Rapid polymerase chain reaction and melting analysis, in: S.A. Bustin (Ed.), The PCR Revolution: Basic Technologies and Applications, Cambridge Univ. Press, New York, 2010, pp. 48-69. Fluorescent DNA polymerase assay / J.L. Montgomery et al. / Anal. Biochem. 441 (2013) 133-139 139 23 CHAPTER 3 THE INFLUENCE OF PCR REAGENTS ON DNA POLYMERASE EXTENSION RATES MEASURED ON REAL-TIME PCR INSTRUMENTS Reprinted from Clinical Chemistry, clinchem.2013.212829 Published-Ahead-of-Print, September 30, 2013, Jesse L. Montgomery and Carl T. Wittwer, \In uence of PCR Reagents on DNA Polymerase Extension Rates Measured on Real-Time PCR Instruments," with permission of the American Association for Clinical Chemistry. Influence of PCR Reagents on DNA Polymerase Extension Rates Measured on Real-Time PCR Instruments Jesse L. Montgomery,1 and Carl T. Wittwer1* BACKGROUND: Radioactive DNA polymerase activity methods are cumbersome and do not provide initial extension rates. A simple extension rate assay would enable study of basic assumptions about PCR and de-fine the limits of rapid PCR. METHODS: A continuous assay that monitors DNA poly-merase extension using noncovalent DNA dyes on com-mon real-time PCR instruments was developed. Exten-sion rates were measured in nucleotides per second per molecule of polymerase. To initiate the reaction, a nucle-otide analog was heat activated at 95 °C for 5 min, the temperature decreased to 75 °C, and fluorescence moni-tored until substrate exhaustion in 30-90 min. RESULTS: The assay was linear with time for over 40% of the reactions and for polymerase concentrations over a 100-fold range (1-100 pmol/L). Extension rates de-creased continuously with increasing monovalent cation concentrations (lithium, sodium, potassium, cesium, and ammonium). Melting-temperature depressors had vari-able effects. DMSO increased rates up to 33%, whereas glycerol had little effect. Betaine, formamide, and 1,2- propanediol decreased rates with increasing concentra-tions. Four common noncovalent DNA dyes inhibited polymerase extension. Heat-activated nucleotide analogs were 92% activated after 5 min, and hot start DNA poly-merases were 73%-90% activated after 20 min. CONCLUSIONS: SimpleDNAextension rate assays can be performed on real-time PCR instruments. Activity is decreased by monovalent cations,DNAdyes, and most melting temperature depressors. Rational inclusion of PCR components on the basis of their effects on poly-merase extension is likely to be useful in PCR, particu-larly rapid-cycle or fast PCR. © 2013 American Association for Clinical Chemistry Several methods have been developed to measure the ac-tivity ofDNApolymerases, but complexity, time require-ments, and specialized instrumentation have prevented their widespread use. Polymerase activity is most often characterized with radiometric assays. These assays mea-sure the incorporation of radiolabelled deoxyribonucleo-tide triphosphates (dNTPs)2 into mechanically sheared or enzymatically digested complex genomicDNA.Activity is measured in terms of units that are generally defined as the amount of enzyme required to incorporate 10nmolof dNTPin 30 min. However, assay conditions and unit def-initions are not standardized, making comparison be-tween measurements difficult. In addition, end-point methods do not provide initial rates and application to PCR kinetics is limited. Other methods have been used to measure poly-merase kinetics, including atomic force microscopy (1 ), light microscopy (2 ), single molecule optical trap-ping (3 ), quench flow (4 ), stopped flow (4-7), and quartz crystal microbalance (8 ). Each of these requires instrumentation not found in most laboratories and has relatively low throughput. Other assays have been adapted for more common instruments, including benchtop fluorometers and microplate readers (9-11). However, these require covalent fluorescent labels, enzyme-coupled reactions, or saturating amounts of single-stranded DNA binding protein. Previously we reported a continuous polymerase ac-tivity assay that uses a stopped-flow instrument (7 ). Nu-cleotide incorporation was monitored with DNA dyes typically used in real-time PCR, eliminating the need to alter reaction chemistry. In the investigation we report here, the assay was modified for useoncommonreal-time PCR instruments. The effects of monovalent cations, melting temperature (Tm) depressors, and DNA dyes on polymerase extension rates were measured. Materials and Methods DNA POLYMERASES Klentaq I (purchased from Wayne M. Barnes at Wash-ington University in St. Louis), FastStart™ (Roche), 1 Department of Pathology, University of Utah Health Sciences Center, Salt Lake City, UT. * Address correspondence to this author at: Department of Pathology, University of Utah Medical School, 50 N. Medical Drive, Salt Lake City, Utah 84132. Fax [phone]; e-mail [e-mail]. Received July 9, 2013; accepted August 27, 2013. Previously published online at DOI: 10.1373/clinchem.2013.212829 Clinical Chemistry 60:2 000-000 (2014) Molecular Diagnostics and Genetics 1 The latest version is at http://hwmaint.clinchem.org/cgi/doi/10.1373/clinchem.2013.212829 Papers in Press. Published October 15, 2013 as doi:10.1373/clinchem.2013.212829 Copyright (C) 2013 by The American Association for Clinical Chemistry 25 Platinum® (Invitrogen), Amplitaq® (Invitrogen), Taq (New England Biolabs), GoTaq® (Promega), Tita-nium ® Taq (Clontech), KAPA2G (Kapa Biosystems), MyTaq™ (Bioline), Ex Taq® (Clontech), Taq (Roche), SpeedSTAR™ (Clontech), KOD (EMD Millipore), Paq5000 (Agilent), Herculase II (Agilent), Phusion® (New England Biolabs), and Amplitaq® Gold (Invitro-gen) were quantified as described previously (7 ) on SDS gels stained with Sypro® Orange (Invitrogen). POLYMERASE EXTENSION TEMPLATE A self-complementary oligonucleotide with the se-quence tagcgaaggatgtgaacctaatcccTGCTCCCGCGGC CGatctgcCGGCCGCGGGAGCA was used as the exten-sion template (capital letters denote self-complementary sequences). This forms a hairpin with a 14-bp stem that has a free 3 end and a 25-base overhang for extension. The oligonucleotide was ordered from Integrated DNA Technologies and purified by high-pressure liquid chro-matography. Concentrations were determined by absor-bance at 260 nm following digestion with purified phos-phodiesterase (12). POLYMERASE EXTENSION ASSAY Extension reactions were performed with a LightCycler® 480 (Roche). Except where otherwise indicated, final concentrations were 50 mmol/L Tris (pH 8.3), 3 mmol/L MgCl2, 1 LCGreen Plus, 50 pmol/L Klentaq I, 100 nmol/L oligonucleotide template, and 200 mol/L of each nucleotide. CleanAmp™ dGTP (TriLink BioTechnologies) was mixed with unmodi-fied dATP, dCTP (deoxycytidine triphosphate), and dTTP (deoxythymidine triphosphate) (Bioline) to limit extension of the template before temperature equilibration. Preliminary studies showed that the use of a single heat-activated nucleotide with 3 unmodified nucleotides increased extension rates by a mean of 14% compared to using all 4 heat-activated nucleotides. Re-duced extension rates were likely caused by lower avail-able dNTPs due to incomplete conversion of the heat-activated nucleotides. The concentration of polymerase was reduced to 50 pmol/L to lengthen the reaction time and ensure initial rates were observed. This is at least 100-fold be-low typical PCR concentrations of 5-20 nmol/L (7 ). To reduce protein loss with serial dilutions, polymerases were diluted from the commercial stock solution im-mediately before extension reactions in 50 mmol/L Tris (pH 8.3), 300 g/mL BSA, and 0.03% Tween® 20. The reaction was initiated by activating the CleanAmp dGTP at 95 °C for 5 min, followed by fluorescence monitoring of nucleotide incorporation at 75 °C. This was accomplished by programming the LightCycler 480 for repeated holds at 75 °C for 1 s with a single acquisition. Reactions were allowed to continue to ex-haustion (30-90 min). Four replicates of each reaction were performed and the SDs reported. PCR DYES AND ADDITIVES Monovalent cations, Tm depressors, DNA dyes, and MgCl2 were titrated into extension reactions to deter-mine their effects on extension rates. LiCl, NaCl, KCl, CeCl, and (NH4)2SO4 were included at final monova-lent cation concentrations up to 50 mmol/L. Final con-centrations of betaine and 1,2-propanediol up to 2.5 mol/L, DMSO and glycerol up to 10%, and formamide up to 7.5% (v/v) were examined. LCGreen® Plus (BioFire Diagnostics), EvaGreen® (Biotium), and SYBR® Green I (Invitrogen) were studied from 0.1 to 5 (approximately 0.1-5 mol/L) (13 ), and Syto® 9 (Invitrogen) was examined from 0.4 to 10.0 mol/L. MgCl2 was studied at concentrations up to 6 mmol/L. In addition to Klentaq I, MgCl2 titrations were also performed for Platinum, Amplitaq, Taq (NEB), GoTaq, Titanium, KAPA2G, MyTaq, Ex Taq, Taq (Roche), and SpeedSTAR. HOT START ACTIVATION The activation times of 2 chemical hot start poly-merases (FastStart and Amplitaq Gold) and heat-activated nucleotide analogs (CleanAmp dNTPs) were assessed. Extension reactions were performed as de-scribed above, except that unmodified dNTPs were used with the hot start polymerases and all 4 heat-activated dNTPs were used with Klentaq I. Activation times between 5 s and 60 min at 95 °C were investi-gated. The concentration of polymerase was increased to 100 pmol/L with 60-min activation times to com-pensate for low extension rates. ASSAY CALIBRATION Linearity between fluorescence and dNTP incorpora-tion was assumed. The first 15 s of data were excluded to eliminate artifacts of initial temperature equilibration. Polymerization was allowed to proceed to substrate ex-haustion, apparent as a maximum plateau and taken as the fluorescence equivalent of 100% extension. Calibration of fluorescence data allows measure-ment of polymerase activity. Additionally, specific ac-tivity in terms of extension rates can be calculated if polymerase quantification is performed. POLYMERASE ACTIVITY Extension curves were normalized between zero and the total number of nucleotides that can be extended, given by: Template L V, (Eq. 1) where [Template] is the concentration of template in nanomoles per liter, L is the extension length of the 2 Clinical Chemistry 60:2 (2014) 26 substrate in bases, and V is the volume of the reaction in liters. The initial slope of the normalized extension curves yields polymerase activity in nanomoles of nu-cleotides per second. EXTENSION RATES Extension curves were normalized between zero and the total number of nucleotides that each polymerase molecule can extend, given by: Template L/ Poly , (Eq.2) where [Poly] is the concentration of the polymerase in nanomoles per liter. With time in seconds as the x axis, the initial slope is the extension rate in nucleo-tides per second per molecule of polymerase, or sim-ply seconds 1. Results Polymerase extension was linear with time for at least 40% of the reaction (Fig. 1A). The initial slope of ex-tension was proportional to the polymerase concentra-tions from 1 pmol/L to at least 100 pmol/L (Fig. 1B). Polymerases were diluted in a buffer containing deter-gent and BSA. When diluted without these compo-nents, variable decreases in activity were observed. This was presumably due to loss of polymerase by adsorp-tion onto surfaces during serial dilutions. Tween 20 was used here, but similar retention of activity was ob-tained with IGEPAL® CA-630, TritonTM X-100, and Brij® 58. The highest extension rates were observed with detergent at 0.03% and BSA at 0.3 g/L (data not shown). All monovalent cations decreased extension rates (Fig. 2). Lithium, sodium, potassium, and cesium had a similar effect, with a mean decrease of 57% at 25 mmol/L. Ammonium most strongly reduced rates, with a decrease of 79% at 25 mmol/L. Extension assays were also performed with divalent cations replacing magnesium. Calcium, manganese, cobalt, and zinc 00 1000 2000 3000 4000 5000 10 000 20 000 30 000 40 000 50 000 Nucleotides extended/polymerase Time (s) A 0.0000 20 40 60 80 100 0.005 0.010 0.015 B 0.020 Initial slope (arbitrary units) Concentration (pmol/L) Fig. 1. Validation of a polymerase extension rate assay that mimics PCR with increased fluorescence from dye incorporation performed on a real-time PCR instrument. Increase of fluorescence signal during nucleotide incorporation is linear with time for at least 40% of reaction completion (A). Initial slope of fluorescence data is linear with polymerase concentrations from 1 to at least 100 pmol/L with R2 0.999 (B). 00 10 20 30 40 50 20 40 60 80 100 120 Concentration (mmol/L) Extension rate (s−1) Fig. 2. Effect of monovalent cations on extension rates. Cations of lithium (circles), sodium (squares), potassium (diamonds), and cesium (triangles) all produce similar de-creases in rates. Cations of ammonium (inverted triangle) show the strongest inhibition. DNA Polymerase Extension and PCR Clinical Chemistry 60:2 (2014) 3 27 were tested at concentrations ranging from 0.3 to 10 mmol/L. Each of these produced artifacts in fluores-cence (i.e., quenching or enhancement) that precluded accurate analysis (data not shown). The effect of Tm depressors on extension rates is shown in Fig. 3. DMSO enhanced rates at concentra-tions up to 10% (1.4 mol/L), with an optimum between 5 and 7.5% (0.7 and 1.1 mol/L). Glycerol had very little effect on extension, with a small increase of 5% at 2.5% (0.3 mol/L) and a decrease of 6% at 10% (1.4 mol/L). Betaine and propanediol did not influence rates at 0.5 mol/L but showed linear decreases above this concen-tration. Extension rates decreased with betaine at a rate of 16% with every increase of 0.5 mol/L beyond 0.5 mol/L. Propanediol showed twice the inhibition, with a decrease of 33% per 0.5 mol/L. A small decrease in rate of 6% was observed with formamide at a 1% concen-tration (0.3 mol/L). At higher concentrations, exten-sion rates also declined linearly. The rates decreased 10% for every 1% increase of the formamide concentration. Each of the DNA dyes studied decreased polymer-ase extension rates, but to varying degrees (Fig. 4). SYBR Green I showed the greatest inhibition, followed by LCGreen Plus, EvaGreen, and Styo 9. Extension rates for the dyes at typical 1 concentrations varied over a 2-fold range with SYBR Green I at 101 s 1, LCGreen Plus at 124 s 1, EvaGreen at 184 s 1, and Syto 9 at 209 s 1. Extension rates increased with increasing concen-trations of MgCl2 up to 6 mmol/L for 9 of the 11 poly-merases studied (see Fig. 1 in the Data Supplement that accompanies the online version of this report at http:// www.clinchem.org/content/vol60/issue2). MgCl2 be-tween 4 and 5 mmol/L produced the fastest extension rates for Titanium and Klentaq I, 2 deletion variants of Taq polymerase, with decreasing rates at higher con-centrations. Data from 4 additional polymerases (KOD, Paq5000, Herculase II, and Phusion) could not be analyzed because the template was degraded by 3 to 5 exonuclease activity before acquisition. Optimal conditions for fast polymerase extension found here and in our previous study (7 ) are summarized in Fig. 5. Fig. 6 shows extension rates as a function of acti-vation time at 95 °C for heat-activated nucleotide analogs (CleanAmp dNTPs) and 2 chemical hot start polymerases (FastStart and Amplitaq Gold). The heat-activated nucleotides were maximally active after 20 min with an extension rate of 110 s 1, but activation was 92% complete after 5 min. Maximal activity of the hot start polymerases required 40 min, with extension rates of 45 s 1 for FastStart and 28 s 1 for Amplitaq Gold. After 20 min, activation was 90% complete for FastStart and 73% complete for AmpliTaq Gold. Con-sistent with prior findings (7 ), the maximal extension Concentration (mol/L) Extension rate (s−1) 0.0 0.5 1.0 1.5 2.0 2.5 0 50 100 150 200 Fig. 3. Effect of Tm depressors on extension rates. DMSO (circles), glycerol (squares), betaine (diamonds), pro-panediol (triangles), formamide (inverted triangles). DMSO increased extension rates at concentrations up to 10% (1.4 mol/L). Glycerol had little effect on rate. Betaine, propane-diol, and formamide each decreased rates with increasing concentration. Propanediol at 2.5 mol/L was not measured because of low activity. 0.1 0.3 1 3 10 0 50 100 150 200 250 300 Extension rate (s−1) Concentration (μmol/L) Fig. 4. Effect of DNA dyes on extension rates. Syto 9 (circles), EvaGreen (squares), LCGreen Plus (dia-monds), SYBR Green I (triangles). Typical 1 concentra-tions used in PCR are indicated by filled markers. Extension rates were not measured for SYBR Green I at 5 (3.4 mol/L) because of low activity and for Syto 9 at 0.4 mol/L because of low signal. Dye concentrations are plotted on a log scale. 4 Clinical Chemistry 60:2 (2014) 28 rate of native Taqs was lower than with deletion variants. Compared to the previously described stopped-flow assay (7 ), convenience and throughput are greatly improved, with a 96/384 well plate format without compromise of accuracy or precision (data not shown). The instrument expense and setup require-ments of a temperature-controlled stopped-flow appa-ratus also greatly exceed those of real-time PCR machines. Discussion Measurement of polymerase extension rates on com-mon real-time PCR instruments enables systematic study of numerous PCR reagents and conditions. Prior work has been hindered by laborious radiolabeled as-says or expensive instrumentation. Our previously re-ported stopped-flow assay conveniently used noncova-lent DNA dyes to measure polymerase extension (7 ). Conversion of the stopped-flow assay to real-time PCR instruments required (a) decreasing the concentration of polymerase to increase the reaction time to 30-90 min, (b) using a heat-activated nucleotide analog to prevent extension during sample preparation, and (c) activating the nucleotides at 95 °C for 5 min, followed by rapid cooling to the desired extension temperature (75 °C) to ensure that initial velocities are observed. It was also critical to dilute the polymerase in detergent and BSA, presumably to prevent polymerase adsorp-tion on vessel walls during dilution. Although we used a LightCycler 480, any instrument capable of exporting fluorescence data as a function of time can be used. We have developed an online tool to simplify analysis of the kinetic data (https://www.dna.utah.edu/ext/ ExtensionCalc.php). Extension rates are normalized to a single poly-merase molecule and are analogous to specific activity. However, unlike prior radiometric assays, initial veloc-ities are measured, templates are standardized, and the buffers mimic those in PCR. As a result, extension rates better reflect the kinetics seen in PCR with processive extension of a defined template for more reproducible activity measurements. Polymerase quantification is not necessary when only activity measurements are desired. The initial slope of calibrated curves yields polymerase activity in nanomoles of nucleotides per second. This is analo-gous to the unit definition of activity, except that initial velocities rather than end-point rates are measured. Noncovalent Dyes Lowest possible Monovalent Ions (Li+, Na+, K+, Ce+, NH4 +) 0 mmol/L DMSO Tm depressors 5 8% Glycerol 0 10% Tris 10 50 mmol/L pH 8.5 8.7 Mg++ 3 6 mmol/L 0 0.5 M Betaine 0 1% Formamide 0 0.5 M 1,2-Propanediol Fig. 5. Optimal conditions for fast polymerase extension. Fastest rates are obtained by eliminating monovalent ions, using high MgCl2 between 3 and 6 mmol/L, and including DMSO between 5% and 8%. Optimal pH is narrow, be-tween 8.5 and 8.7, and Tris has little effect (7 ). Betaine and 1,2-propanediol are best kept below 0.5 mol/L and forma-mide below 1%. Noncovalent dyes are best used at the lowest concentration that produces the desired fluores-cence intensity. 00 10 20 30 40 50 60 20 40 60 80 100 120 Activation time (min) Extension rate (s−1) Fig. 6. Extension rates after activation with hot start polymerases and nucleotides. FastStart (squares), Amplitaq Gold (diamonds), hot-start nucleotide analogs (circles). FastStart showed nearly dou-ble the extension rates of Amplitaq Gold with the same activation time. The hot start nucleotides required half the activation time as the hot start polymerases for maximum activity. DNA Polymerase Extension and PCR Clinical Chemistry 60:2 (2014) 5 29 All monovalent cations studied decreased exten-sion rates in a concentration-dependent manner (Fig. 2). Oddly enough, potassium chloride and ammonium sulfate are frequently found in PCR buffers. With these components, amplification appeared more specific with higher yields in some reported studies (14-16). Fig. 2 indicates that any benefit obtained from inclu-sion of monovalent cations in PCR does not result from enhanced extension rates. Tmdepressors are often added for PCR of GC-rich templates. We found that DMSO increases extension rates and glycerol has little effect at concentrations up to 10%. Other studies found that activity decreased 50% in the presence of 10% DMSO and 30% with 10% glycerol (17, 18 ). These prior studies used radioactive assays and activated salmon sperm DNA as a template. We observed a linear decrease in extension rates with formamide concentrations above 1%. A previous study showed no effect up to 10% (17 ). Another showed a 50% decrease in activity at 10%. Our study showed greater inhibition, with a 65% decrease in the presence of 7.5% formamide. The discrepancies in these studies suggest that greater uniformity in assay conditions and standardization of the template may improve the re-producibility of activity assays. Betaine and propanediol both produced linear de-creases in extension rates. When maximal polymerase extension rates are a concern, betaine, propanediol, and formamide should be used at the lowest concen-trations possible for successful amplification. The noncovalent DNA dyes studied here decrease polymerase extension rates (Fig. 4). Selection of the appropriate dye and concentration will depend on a number of factors, such as instrument optical require-ments, desired extension rates, and post-PCR process-ing. For example, the fastest extension rate was ob-served for SYBR Green I at 0.2 (0.14 mol/L). However, SYBR Green I does not detect heterodu-plexes in high-resolution melting analysis (19, 20 ). Faster extension rates can be obtained with each dye by lowering the concentration, but this is also accompa-nied by a lower signal and may be limiting, depending on the sensitivity of the instrument. The choice of a hot start method determines the speed of activation before PCR. Chemical hot starts show low extension rates despite very long activation times, though FastStart appears to require about half the activation time of Amplitaq Gold for the same ex-tension rate. Rates were faster for the heat-activated nucleotide analogs at all activation times, indicating a large difference in the specific activity of chemically modified hot start polymerases and Klentaq I. For faster PCR, heat-activated nucleotide analogs are more desirable than modified polymerases. Routine measurements of activity and extension rates are enabled by this continuous fluorescence assay. High throughput is attained with microtiter plates, allowing simultaneous comparison of several poly-merases and conditions. Components are easily optimized to identify suitable PCR reagents and storage buffers. Engineered polymerase variants can be screened for desired activity. Polymerase prepara-tion lots can be assayed for consistent activity to ensure reproducible PCR efficiency. Polymerases screened for high extension rates are needed for rapid PCR applica-tions. Because the extension rate is measured under PCR conditions, insight into the speed of extension obtainable during PCR can guide optimization of thermal cycling protocols for faster, more efficient amplification. Is PCR constructed rationally, or are we following the initial choices of PCR pioneers and reluctant to change familiar reagents and ingrained protocols? Considering the data obtained from this and our prior stopped-flow study (7 ), extension rates are improved by high Mg2 (3- 6 mmol/L) andDMSO(5%-10%) in a narrow pH range (8.5- 8.7) and decreased by K , (NH4)2SO4, dyes, and most Tm suppressors. Of course, PCR is much more than just polymerase exten-sion. Fidelity and specificity are also crucial. Neverthe-less, polymerase extension is a central factor in under-standing PCR and paramount to efforts to increase its speed. Author Contributions: All authors confirmed they have contributed to the intellectual content of this paper and have met the following 3 re-quirements: (a) significant contributions to the conception and design, acquisition of data, or analysis and interpretation of data; (b) drafting or revising the article for intellectual content; and (c) final approval of the published article. Authors' Disclosures or Potential Conflicts of Interest: Upon man-uscript submission, all authors completed the author disclosure form. Disclosures and/or potential conflicts of interest: Employment or Leadership: C.T. Wittwer, BioFire Diagnostics, and Clinical Chemistry, AACC. Consultant or Advisory Role: None declared. Stock Ownership: C.T. Wittwer, BioFire Diagnostics. Honoraria: None declared. Research Funding: C.T. Wittwer, BioFire Diagnostics. Expert Testimony: None declared. Patents: None declared. 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DNA Polymerase Extension and PCR Clinical Chemistry 60:2 (2014) 7 31 CHAPTER 4 THE INFLUENCE OF NUCLEOTIDE SEQUENCE AND TEMPERATURE ON THE ACTIVITY OF THERMOSTABLE POLYMERASES 4.1 Abstract Extension rates of a thermostable polymerase were measured from 50 to 90 C for templates with varying sequence using a uorescent activity assay adapted for real-time PCR instruments. Templates consisted of identical hairpins with a melting temperature of 92 C and extension regions with either single-base repeats or guanosine-cytosine (GC) contents ranging from 0 to 100%. Optimum extension temperature was 70 to 75 C for all templates with a near linear decrease in extension rates outside this range. Extension rates increased with GC content up to 60% and decreased at higher GC. Rates varied greatly for each nucleotide with guanosine (214 s-1 at 75 C) >cytidine (150 s-1 at 75 C) >adenosine (81 s-1 at 75 C) >thymidine (46 s-1 at 75 C). Predictions were within 30% of measured rates for 59% of calculations with greatest agreement among lower GC content. Templates with higher GC contents exhibited slower rates and were increased to within 4 to 20% of prediction with the addition of 7.5% dimethyl sulfoxide (DMSO), indicating inhibition due to secondary structure. In the presence of oligonucleotide probes, polymerases with and without 5' to 3' exonuclease activity exhibited similar decreases in extension rates of 70 and 65%. Extension rates are in uenced by template sequence, increasing with higher GC content and decreasing with secondary structure. Understanding these parameters across temperatures will enable improved temperature cycling for DNA ampli cation with higher yield and speci city. 33 4.2 Introduction Understanding the parameters that in uence DNA polymerase activity is essential for optimizing PCR conditions and preventing ampli cation failure. Sequence characteristics of templates such as GC content and secondary structure are known to reduce ampli cation e ciency. In addition, incorporation rates are base speci c for a variety of polymerases [1]{[6]. However, the sequence dependence of activity has not been studied for thermostable DNA polymerases. Kinetic analysis of nucleotide incorporation has been performed using stopped- ow [6], quench- ow [1], [4], [7], and quartz crystal microbalance [8]. These methods are capable of monitoring fast reaction times but have required uorescent base analogs, radiolabeled nucleotides or immobilization of the template to<