Methods in Enzymology Vol 340: Drug-Nucleic Acid Interactions

Preface Progress in molecular biology and studies of small molecule binding to nucleic acids have been inextricably li...

Author: Jonathan B. Chaires | Michael J. Waring

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Preface

Progress in molecular biology and studies of small molecule binding to nucleic acids have been inextricably linked. A testament to that fact is the inclusion of eight papers directly concerned with drug-DNA interactions among the recently published list of the 100 most cited articles in the Journal of Molecular Biology. Few other scientific areas are as well represented on that list. Small molecules have perhaps taught us more about DNA than DNA has taught us about small molecules. Watson, for example, notes in the Molecular Biology of the Gene that the "fact that intercalation occurs so readily indicates that it is energetically favored... [and] is additional evidence for the metastability of the double-helical structure--its ability to assume many inherently unstable configurations that normally revert quickly back to the standard B conformation." From that point of view, intercalation provided one of the very first indications of the plasticity of DNA, an area that has blossomed to reveal an incredible diversity of structural forms. Perhaps the most widespread interest in small molecules that bind to nucleic acids stems from their potential as useful pharmaceutical agents. Indeed, some of the very best anticancer drugs are well-documented DNA binders. While interest in drug-DNA interactions has at times waned, recent advances in chemical synthesis, analytical instrumentation to measure binding, and structural biology have greatly enhanced the potential for rational design of new therapeutic compounds. Accordingly, studies on the interaction of small molecules with nucleic acids have taken on new life and have helped spawn several emergent biotechnology companies dedicated to exploiting the promise of making new types of pharmaceuticals targeted at nucleic acids. The aim of this volume is to consolidate key methods for studying ligandnucleic acid interactions, both old and new, into a convenient source. Accordingly, we have solicited from experts in a variety of disciplines articles that concisely but completely describe useful methods and strategies for studying small molecule binding to nucleic acids. Techniques that are useful now range from biophysical and chemical approaches to methods rooted in molecular and cell biology. We hope that this volume will serve as a useful compendium of methods both to newcomers entering the field as well as to scientists already actively engaged in research in this area. JONATHANB. CHAIRES MICHAELJ. WARING

xiii

Contributors to Volume 3 4 0 Article numbers are in parentheses following the names of contributors. Affiliations listed are current.

CHRISTIAN BAILLY (24, 31), INSERM

CARLEEN M. CULL1NANE (23), Pharma-

U-524, and Laboratoire de Pharmacologie Antitumorale du Centre Oscar Lambret IRCL, 59045 Lille, France

cology and Developmental Therapeutics Unit, Peter MacCallum Cancer Institute, Victoria 3002, Australia

ALBERT S. BENIGHT (8), Department

SUZANNE M. CUTrS, (23), Department of

of Chemistry, University of Illinois, Chicago, Illinois 60607 and DNA Codes LLC, Chicago, Illinois 60601

Biochemistry, La Trobe University, Bundoora, Victoria 3083, Australia

LAWRENCE A. BOTTOMLEY(11), School of

Chemistry and Biochemistry, Georgia Institute of Technology, Atlanta, Georgia 30332 SOPHIA Y. B REUSEGEM(10), Laboratoryfi~r

Fluorescence Dynamics, Department of Physics, University of Illinois, Urbana, Illinois 61801 JONATHAN B. CHAIRES (1, 5, 27), De-

partment of Biochemistry, University of Mississippi Medical Center, Jackson, Mississippi 39216 YEN CHOO (30), Gendaq Limited, London

NW7 lAD, United Kingdom BABUR Z. CHOWDHRY(6), School of Chem-

ical and Life Sciences, University of Greenwich, London SE18 6PF, United Kingdom ROBERT M. CLEGG (10), Laboratory for

Fluorescence Dynamics, Department of Physics, University of Illinois, Urbana, Illinois 61801 DONALD M. CROTHERS (3, 23), Depart-

ment of Chemistry, Yale University, New Haven, Connecticut 06520-8107 MARK S. CUBBERLEY (28), Department of

Chemistry and Biochemistry, University of Texas, Austin, Texas 78712

JAMES C. DABROWIAK(21), Department of

Chemistry, Center for Science and Technology, Syracuse University, Syracuse, New York 13244 TINA M. DAVIS (2), Department of Chem-

istry, Georgia State University, Atlanta, Georgia 30303 PETER B. DERVAN (22), Department of

Chemistry, California Institute of Technology, Pasadena, California 91125 MAGDALENAERIKSSON(4), Department of

Physical Chemistry, Chalmers University of Technology, Gothenburg SE-41296, Sweden, and Department of BiDchemistry, University of Gothenburg, Gothenburg SE-40530, Sweden CHRISTOPHE ESCUDI~ (16), Laboratoire

de Biophysique, INSERM U201, CNRS UMR 8646, Museum National d'Histoire Naturelle, 75231 Paris Cedex 05, France IZABELA FOKT (27), M. D. Anderson Can-

cer Center, University of Texas, Houston, Texas 77030 KEITHR. Fox (20), Division of Biochemistry

and Molecular Biology, School of Biological Sciences, University of Southampton, Southampton S016 7PX, United Kingdom

x

CONTRIBUTORS TO VOLUME 340

THI~RI~SE GARESTIER (16), Laboratoire

BESIK I. KANKIA (7), Department of

de Biophysique, 1NSERM U201, CNRS UMR 8646, Museum National d'Histoire Naturelle, 75231 Paris Cedex 05, France

Pharmaceutical Sciences, University ~f Nebraska Medical Center, Omaha, Nebraska 68198

JERRY GOOD1SMAN (21), Department of

ASMITA KUMAR (33), Department of Bio-

Chemistry, Center for Science and Technology, Syracuse University, Syracuse, New York 13244 DAVID E. GRAVES (18), Department of

Chemistry, University of Mississippi, University, Mississippi 38677 KEITH A. GRIMALDI(17), CRC Drug-DNA

Interactions Research Group, Royal Free and University College Medical School, University College London, London WI P 8BT, United Kingdom VLAD1M1RM. GUELEV (28), Department of

Chemistry and Biochemistry, Universi~" of Texas, Austin, Texas 78712 Krebs Institute for Biomolecular Science, Department of Chemisto, University of Sheffield, Sheffield $3 7HF, United Kingdom

IHTSHAMUL HAG (6),

JOHN A. HARTLEY (17), CRC Drug-DNA

chemistry, University of Mississippi, Jackson, Mississippi 39216 DONALD W. KUPKE (7), Department of

Chemistrry, University of Virginia, Charlottesville, Virginia 22901 ANDREW N. LANE (12), Division of Molecu-

lar Structure, National Institute for Medical Research, London NW7 IAA, United Kingdom GREGORY H. LENO (33), lnfgen Incorpo-

rated, DeForest, Wisconsin 53532 PETER T. LILLEHEI (l l), School of Chem-

istry and Biochemistry, Georgia Institute of Technology, Atlanta, Georgia 30332 R. SCOTT LOKEY (28), Department of

Chemistry and Biochemistr); University of Texas, Austin, Texas 78712

Interactions Research Group, Royal Free and University College Medical School, University College London, London W1P 8BT, United Kingdom

FRANK G. LOONTIENS (10), Laboratory

PAUL B. HOPKINS (19), Department of

RYAN A. LUCE (19), Department of Chem-

for Biochemistry, WEVIO, University of Gent, Gent 9000, Belgium

Chemistry, University of Washington, Seattle, Washington 98195

istr); University of Washington, Seattle, Washington 98195

LAURENCE H. HURLEY (29), College of

CHRISTOPHE MARCHAND (32), Laboratory

Pharmacy, University of Arizona, Tucson, Arizona 85721 and Arizona Cancer Center, Tucson, Arizona 85724

of Molecular Pharmacology, Division of Basic Sciences, National Cancer Institute, National Institutes of Health, Bethesda, Maryland 20892

Gendaq Limited, London NW7 laD, United Kingdom

MARK ISALAN (30),

Chemistry and Biochemistry, University of Texas, Austin, Texas 78712

Department of Pharmaceutical Sciences, University of Nebraska Medical Center, Omaha, Nebraska 68198

TERENCE C. JENKINS (6), Yorkshire Cancer

CLAIRE J. MCGURK (17), CRC Drug-DNA

Research Laboratory of Drug Design, Cancer Research Group, University of Bradford, Bradford BD7 1DP, United Kingdom

Interactions Research Group, Royal Free and University College Medical School, University College London, London WI P 8BT, United Kingdom

BRENT L. IVERSON (28), Department of

LUIS A. MARKY (7),

CONTRIBUTORS TO VOLUME 340 PETER J. MCHUGH (17), CRC Drug-DNA

Interactions Research Group, Royal Free and University College Medical School, University College London, London W1P 8BT, United Kingdom MARK P. MCPIKE (21), Department of

Chemistry, Centerfor Science and Technolog); Syracuse University, Syracuse, New York 13244

xi

R. PHILLIPS (23), Department of Biochemistry, LaTrobe Universit3; Bundoora, Victoria 3083, Australia

DON

YVES POMMIER (32), Laboratory of Molec-

ular Pharmacolog); Division of Basic Sciences, National Cancer Institute, National Institutes of Health, Bethesda, Maryland 20892 JOSl~ PORTUGAL(25, 27), Departamento de

Chemistry and Biochemistry, Universi~ of Texas, Austin, Texas 78712

Biologia Molecular y Celular, lnstituto de Biologia Molecular de Barcelona, CSIC, Barcelona 08034, Spain

NOURI NEAMATI(32), Laboratory of Molec-

WALDEMAR PRIEBE (27), M. D. Ander-

ular Pharmacology, Division of Basic Sciences, National Cancer Institute, National Institutes of Health, Bethesda, Maryland 20892

son Cancer Center, University of Texas, Houston, Texas 77030

MEREDITH M. MURR (28), Department of

JAROSLAV NESETI~IL (8), Department of

Applied Mathematics, Faculty of Mathematics and Physics, Charles Universi~, 118 O0 Praha 1, Czech Republic PETER E. NIELSEN (15), Department of

Medical Biochemistry and Genetics, The Panum Institute, University of Copenhagen, Copenhagen DK-2200, Denmark BENGT NORD~N (4), Department of Phys-

ical Chemistry, Chalmers University of Technology, Gothenburg SE-41296, Sweden

TERESA PRZEWLOKA (27), M. D. Ander-

son Cancer Center, University of Texas, Houston, Texas 77030 PETER REGENFUSS (10), Laboratory for

Fluorescence Dynamics, Department of Physics, Universi~' of Illinois, Urbana, Illinois 61801 JINSONG REN (5), Department of Biochem-

istry, University of Mississippi Medical Center, Jackson, Mississippi 39216 RICCELLI (8), Department of Chemistry, University of Illinois, Chicago, Illinois 60607, and DNA Codes LLC, Chicago, Illinois 60601

PETER V.

Department of Chemistry, University of Illinois, Chicago, Illinois 60607, and Integrated DNA Technologies, Coralville, Iowa 52241

RICHARD D. SHEARDY(26), Department of

PETR PAN(~OSKA(8), Department of Chem-

School, Elmwood Park, New Jersey 07407

RICHARD OWCZARZY (8),

istry, University of Illinois, Chicago, Illinois 60607, and Center for Discrete Mathematics, Applied Computer Science and Applications DIMAT1A, Charles University, Prague, Czech Republic, and DNA Codes LLC, Chicago, Illinois 60601 MARY ELIZABETH PEEK (13), School of

Chemistry and Biochemistry, Georgia Institute of Technology, Atlanta, Georgia 30332

Chemistry and Biochemistry, Seton Hall Universit); South Orange, New Jersey 07079 ANGELA M. SNOW (26), Memorial High CHARLES H. SPINK (9), Department of

Chemistry, State University of New York, Cortland, New York 13045 DAEKYU SUN (29), Institute for Drug De-

velopment, San Antonio, Texas 78245 JIAN-SHENG SUN (16), Laboratoire de

Biophysique, INSERM U201, CNRS UMR 8646, Museum National d'Histoire Naturelle, 75231 Paris Cedex 05, France

xii

CONTRIBUTORS TO VOLUME 340

MICHAEL J. TILBY (17), Cancer Research

Unit, Medical School, University of Newcastle Upon Tyne, Newcastle NE2 4HH, United Kingdom JOHN W. TRAUGER (22), Department of Chemistry, California Institute of Technology, Pasadena, California 91125 JOHN O. TRENT (14, 27), James Graham Brown Cancer Center, Department of Medicine, University of Louisville, Louisville, Kentucky 40202 PETER M. VALLONE (8), Department of Chemistry, University of Illinois, Chicago, Illinois 60607 and National Institute of Standards" and Technology, Biotechnology Division, Gaithersburg, Mao, land 20899 MICHAEL J. WARING (20, 24), Department of Pharmacology, University of Cam-

bridge, Cambridge CB2 IQJ, United Kingdom SUSAN E. WELLMAN (9), Department of Pharmacology and Toxicolog), University of Mississippi Medical Center, Jackson, Mississippi 39216 LOREN DEAN WILLIAMS (13), School of

Chemistry and Biochemistry, Georgia Institute of Technology, Atlanta, Georgia 30332 W. DAVID WILSON (2), Department of

Chemistry, Georgia State University, Atlanta, Georgia 30303 HONGZH1XU (33), Department of Biochem-

istry, University of Mississippi, Jackson, Mississippi 39216 STEVEN M. ZEMAN (3), Department of

Chemistry, Yale University, New Haven, Connecticut 06520

[1]

ANALYSTSOF LIGAND-DNA BINDINGISOTHERMS

3

[1] Analysis and Interpretation of Ligand-DNA Binding Isotherms By JONATHANB. CHA1RES

Introduction To attain a reasonable understanding of any ligand-receptor interaction, it is necessary to answer the questions posed by Scatchard ~ more than 50 years ago: "How many? How tightly? Where? Why? What of it?" The first two Questions (and in part the third) can be answered by equilibrium binding studies, and are the primary focus of this chapter. The remaining questions concisely express the concerns of structural and functional studies, and may be addressed by X-ray crystallography, nuclear magnetic resonance (NMR) techniques, molecular modeling, and a variety of chemical and molecular biological methods. Macromolecular binding is a phenomenon of general interest, and the underlying general principles are the same for ligand binding to proteins or to nucleic acids. A number of excellent general treatments of macromolecular binding are available that explain the underlying physical chemistry in detail .2-6 What distinguishes the binding of small molecules to DNA from their binding to proteins is the need to account for behavior arising from the lattice properties of linear DNA molecules. Various neighbor exclusion models have evolved to cope with that complexity, and are described. An excellent discussion of the principles of nucleic acid binding interactions is provided by Bloomfield et al. 7 Determination of the binding constant K allows the binding free energy change, AG, to be calculated by the standard Gibbs equation, AG = - R T In K, where R is the gas constant and T is the temperature in degrees Kelvin. From studies of the temperature dependence of the binding constant, or (preferably) by calorimetric studies, the binding enthalpy (AH) may be obtained. The binding free energy may then be partitioned into its enthalpic and entropic components, AG = AH -- TAS, where AS is the entropy change. Knowledge of these thermodynamic parameters

I G. Scatchard, Ann. N.Y. Acad. Sci. 51,660 (1949). 2 j. T. Edsall and J. Wyman, "Biophysical Chemistry." Academic Press, New York, 1958. 3 j. Wyman and S. J. Gill, "Binding and Linkage." University Science Books, Mill Valley, California, 1990. 41. M. Klotz, "Ligand Receptor Energetics." John Wiley & Sons, New York, 1997. 5 E. diCera, "Thermodynamic Theory of Site-Specific Binding Processes in Biological Macromolecules." Cambridge University Press, Cambridge, 1995. 6 G. Weber, "Protein Interactions." Chapman & Hall, New York, 1992. 7 V. A. Bloomfield, D. M. Crothers, and J. Ignacio Tinoco, "Nucleic Acids: Structures, Properties and Functions," 1st Ed. University Science Books, Sausalito, California, 2000.

METHODS IN ENZYMOLOGY, VOL. 340

Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. 0076-6879/00 $35.00

4

BIOPHYSICALAPPROACHES

[ 11

provides a firm foundation for understanding the molecular forces that govern the binding reaction, allowing one to begin to address Scatchard's question "Why?" Details of attempts to parse binding free energies for ligand-DNA interactions in order to understand the contribution of various molecular forces are described in publications from this and other laboratories, s- 12 The aim of this chapter is to offer a concise guide for the analysis and interpretation of ligand-DNA binding isotherms. Methods for experimentally obtaining binding data are not discussed because detailed, practical descriptions of experimental protocols are available. 13 15 In this chapter, examples of binding data are taken from results obtained in the author's laboratory with the anticancer agent daunomycin (daunorubicin). Daunomycin is perhaps the best-characterized DNA intercalator, and its binding to a wide variety of DNA sequences and structures has been thoroughly investigated. ~6,17 Model-Independent Approaches Figure 1 shows the results from two types of binding experiments, each of which addresses one of Scatcbard's queries as directly as possible. The method of continuous variations Is-a1 may be used to construct a so-called Job plot (Fig. 1A). Binding stoichiometries may be determined from such plots without recourse to any assumed binding model. For the data shown in Fig. 1A for the interaction of daunomycin with calf thymus DNA, an inflection near 0.2 mol fraction ligand indicates a binding stoichiometry of one ligand per 3 or 4 base pairs. The exact stoichiometry from the inflection at 0.21 mol fraction is (1.0 - 0.21)/0.21 = 3.76 base pairs. This value represents the predominant binding mode, although an s j. B. Chaires, Anticancer Drug Des. 11,569 (1996). 9 j. B. Chaires, Biopolymers 44, 201 (1997). l01. Haq, J. E. Ladbury, B. Z. Chowdhry, T. C. Jenkins, and J. B. Chaires, J. Mol. Biol. 271,244 (1997). II j. Ren, T. C. Jenkins, and J. B. Chaires, Biochemistry 39, 8439 (2000). 12 S. Mazur, F. A. Tanious, D. Ding, A. Kumar, D. W. Boykin, I. J. Simpson, S. Neidle, and W. D. Wilson, J. Mol. Biol. 300, 321 (2000). 13 X. Qu and J. B. Chaires, Methods Enzymol. 321, 353 (2000). L4T. C. Jenkins, in "Drug-DNA Interaction Protocols" (K. R. Fox, ed.), Vol. 90, pp. 195-218. Humana Press, Totowa, New Jersey, 1997. 15 p. C. Dedon, in "Current Protocols in Nucleic Acid Chemistry" (S. L. Beaucage, D. E. Bergstrom, G. D. Glick, and R. A. Jones, eds.), Vol. 1, pp. 8.2.1-8.2.8. John Wiley & Sons, New York, 2000. 16 j. B. Chaires, in "Advances in DNA Sequence Specific Agents" (L. H. Hurley, ed.), Vol. 2, pp. 141167. JAI Press, Greenwich, Connecticut, 1996. 17 j. B. Chaires, Biophys. Chem. 35, 191 (1990). 18 E Job, Ann. Chim. (Paris) 9, 113 (1928). 19 C. Y. Huang, Methods Enzymol. 87, 509 (1982). 2o A. Waiters, Biomed. Biochim. Acta 44, 132t (1985). 21 E G. Loontiens, E Regenfuss, A. Zechel, L. Dumortier, and R. M. Clegg, Biochemistry 29, 9029 (1990).

[1]

ANALYSIS OF L I G A N D - D N A BINDING ISOTHERMS

I

I

I

200

I

/

I

1.2

,

,

,

,

5 ,

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,

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t

0.8

-200 0.6 -400

0.4

-600

0.2

-800

o.o

•

B

, I , I

0.0

0.2

0.4

0.6

0.8

-20

-18

Mole Fraction Daunomycin

-16

I

I

-14

-12

-t0

In Cf

FIG. I. Daunomycin binding to calf thymus DNA. (A) Job plot obtained from fluorescence titration studies. A F is the difference in fluorescence emission intensity between solutions of daunomycin alone and in the presence of DNA. The minimum indicates a binding stoichiometry of 3 or 4 base pairs. (B) Binding isotherm for the daunomycin--calf thymus DNA interaction. The fractional saturation was calculated assuming a 3-bp binding site. The abscissa is the natural logarithm of the free daunomycin concentration.

inflection near 0.5-0.6 mol fraction indicates an additional binding mode at higher drug concentrations. The results shown here, based on fluorescence data, agree well with data based on absorbance changes. 2° The Job plot thus answers the question "How many?" directly. In studies of ligand-DNA interactions, this method has been underutilized and its advantages largely unappreciated. In the case of multiple binding modes, the method of continuous variations is particularly valuable, and clearly reveals complexities in the binding process. Published examples for the groove-binder Hoechst 3325821 and for the bisintercalating anthracycline WP63122 illustrate the value of the method in cases of complicated, multimode binding interactions. Figure 1B shows a titration binding isotherm for the daunomycin-calf thymus DNA interaction. In this form, the fractional occupancy of binding sites is shown as a function of the natural logarithm of the free daunomycin concentration (Ct-). The fractional occupancy was calculated from the experimentally determined binding

22 F. Leng, W. Priebe, and J. B. Chaires, Biochemistry 37, 1743 (1998).

6

BIOPHYSICALAPPROACHES

[ 1]

ratio r (moles daunomycin bound per mole base pair) and the binding stoichiometry was determined from the Job plot shown in Fig. 1A. The form of the plot shown in Fig. 1B is regarded by some 3 as the most fundamental representation of binding data because the logarithm of the free ligand activity is proportional to the chemical potential of the ligand. For simple binding to identical, noninteracting sites, titration binding curves should be symmetric about a midpoint located at a ligand concentration that is the reciprocal of the association binding constant, and should cover a span of 1.8 lOgl0 units (4.14 In units) in going from 0.1 to 0.9 fractional saturation. 3'4'6 The data shown in Fig. 1B cover a span of 5.4 in units (2.4 log~0 units) and represent an essentially complete binding titration curve. The span is greater than expected for simple binding, which indicates negative cooperativity, neighbor exclusion, or heterogeneity of binding sites. Perhaps the main advantage of the data shown in Fig. 1B is that they may be analyzed in a model-independent way by using the Wyman concept of median ligand activity. 3'5 The free energy of ligation (AGx) to go from a state where no ligand is bound to a degree of saturation of k? is given by Eq. (1): P 2 AGx = RT Jo In Cfg~"

(1)

where RT has its usual meaning. The pronounced advantage of Eq. (1) is that it provides a free energy estimate to attain any degree of saturation without recourse to any specific binding model. Numerical integration of the data in Fig. 1B yields an estimate of AGx = - 7 . 8 kcal mol I for the full ligation of a daunomycin binding site. Free energies derived from binding constants obtained by curve fitting to specific models must agree with this model-independent value if the model is reasonable. Neighbor Exclusion Models Figure 2 shows data for the daunomycin-calf thymus DNA interaction in the form of a Scatchard plot, J by far the most common representation of binding data for ligand-DNA interactions. To explain the curvature in such plots, a variety of neighbor exclusion models were proposed, 23'24 and these have become the most commonly used models for the interpretation of binding isotherms. Neighbor exclusion models assume (in their simplest form) that the DNA lattice consists of an array of identical and noninteracting potential binding sites. The base pair is commonly defined as the lattice binding site for duplex DNA. Ligand binding to any one site occludes neighboring sites from binding as defined by the site size n. As the lattice approaches saturation, the probability of finding a stretch 23 D. M. Crothers, Biopolymers6, 575 (1968). 24j. D. McGheeand R H. yon Hippel,J. Mol.Biol. 86, 469 (1974).

[ 1]

ANALYSIS OF L I G A N D - D N A BINDING ISOTHERMS

8.0x10

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0.15

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r FIG. 2. Scatchard plot for the daunomycin-calf thymus DNA interaction. The solid line is the best fit of the neighbor exclusion model [Eq. (2)] to the experimental data yielding the parameters shown in Table I. The dashed line is the best fit with the exclusion parameter constrained to an integral value of 3.

of unoccupied DNA n base pairs long decreases, producing the curvature seen in Fig. 2. The curvature does not result from a decrease in the intrinsic binding affinity, but rather arises from the decreased probability of finding a free site of the appropriate size. McGhee and von Hippe124 derived a closed form equation that embodies the neighbor exclusion model [Eq. (2)]:

r

--

=

K(1

-

nr)

I

1;

1--"r

]°' J

(2)

where K is the association constant for ligand binding to an isolated lattice site, n is

8

BIOPHYSICAL APPROACHES

[ 1]

the neighbor exclusion parameter, and r is the binding ratio. Since its publication in 1974, the McGhee and von Hippel article has been cited more than 1350 times, and is certainly the most commonly used model for the interpretation of ligand binding to DNA. It should be noted that Crothers 23 originally derived a neighbor exclusion model 6 years before McGhee and von Hippel, using the statistical mechanics matrix method, but did not offer a convenient closed form equation for use in fitting experimental data. However, starting with Crother's characteristic equations, 23 it is straightforward to obtain an equation identical to Eq. (2) by simple algebraic rearrangement. The two models are therefore equivalent. Equation (2) is commonly used as a fitting function for nonlinear least-squares analysis of ligand-DNA binding isotherms. 13'25 In fact, Eq. (2) is not appropriate for such purposes, because nonlinear fitting by most methods assumes that the independent variable is error free, and that all of the experimental uncertainty resides in the dependent variable. 26 Because the binding ratio r is experimentally determined, it contains error. Worse, the dependent variable r/Cf is a derived quantity so that error in r is propagated into the dependent variable. Nonetheless, nonlinear fitting methods are inevitably used to extract K and n value from experimental data. Two excellent software packages are routinely used in this laboratory for nonlinear curve fitting, FitAll (MTR Software, Toronto, Canada) and Origin (Microcal, Northampton, MA). FitAll now contains a module for Monte Carlo analysis z7 of the error in parameter estimates. Origin contains a module for a rigorous evaluation of parameter error by determination of their upper and lower bounds at any chosen confidence interval.28 Table I shows the results of fits of the data shown in Fig. 2 to the neighbor exclusion model. The binding constant, K = 6.6 x 105 M 1, may be used to calculate AG = - 7.8 kcal mol -I. That value is in excellent agreement with that obtained for the model-independent approach described above. The exclusion parameter, n = 3.3, agrees well with the estimate of the site size obtained by the method of continuous variations. The nonintegral value of the exclusion parameter (n) poses a problem. For neighbor exclusion models, n should strictly be an integer quantity. A fractional value makes no physical sense for a DNA lattice composed of identical, noninteracting sites] Table I shows, however, that if n is constrained to an integer value (either 3 or 4), the standard deviation of the fit degrades significantly. Figure 2 shows the best fit obtained with n = 3. Systematic deviation between the fit and the data are clear, with the data having more curvature than the calculated function. This results in nonrandom residuals, indicating an inadequate fit of the model to the data. 28 The fractional values of n that are required to obtain a statistically 25 j. j. Correia and J. B. Chaires, Methods Enzymol. 240, 593 (1994). 26 M. L. Johnson, Methods Enzymol. 210, 106 (1992). 27 M. Straume and M. L. Johnson, Methods Enzymol. 210, 117 (1992). 28 M. L. Johnson and L. M. Faunt, Methods Enzymol. 210, 1 (1992).

[]]

ANALYSIS OF L I G A N D - D N A BINDING ISOTHERMS

9

TABLE I NONLINEARLEAST-SQUARESFITS OF DAUNOMYCINBINDINGDATATO NEIGHBOREXCLUSIONMODELSa

Model McGhee-von Hippel n = 3.0 n = 4.0

Friedman-Manning n = 3.0 n ----4.0

K/IOS(M I) 6.6 ± 5.9± 5.4 i 7.2+ 7.0 ± 7.2 ±

0.2 0.1 0.4 0.2 0.2 0.5

N (bp)

Standard deviation

3.3 + 0. l Fixed Fixed 3.1 + 0.1 Fixed Fixed

51,330 56,250 154,600 53,310 53,280 133,300

a McGhee-von Hippel refers to the neighbor exclusion model specified by Eq. (2). Friedman-Manning refers to the model specified by Eq. (3). K is the association constant and n is the neighbor exclusion

parameter expressed in base pairs. The standard deviation is of the best fit.

acceptable fit indicate that the neighbor exclusion model is not, in fact, an appropriate model for the data. Fundamental assumptions of the model must be violated. As is discussed below, the most likely assumption that is violated is that sites are identical, when in fact they are heterogeneous. An extension of the simple McGhee-von Hippel model incorporates an additional term to account for ligand-ligand cooperativity. In principle, integral values of n coupled with negative ligand-ligand cooperativity might adequately fit data such as are shown in Fig. 2. Unfortunately, we previously have shown that attempts to incorporate the added cooperativity parameter are statistically unwarranted and that attempts to force fits to integral values of n are futile. 25 Friedman and Manning derived a variant of the neighbor exclusion model that incorporates aspects of polyelectrolyte theory.29'3° Counterion condensation around DNA is dictated by the spacing of the charged phosphates along the double helix. 31'32 Counterion release coupled to the binding of a charged ligand to DNA provides an energetically favorable contribution to the binding free energy. For DNA intercalation reactions, a complexity arises that the FriedmanManning model addresses. Intercalation results in a separation of the stacked base pairs in duplex DNA, and increased phosphate spacing. This alters the polyelectrolyte properties of the duplex, and results in additional counterion release. As the DNA lattice is saturated with intercalator, there is an ever-changing increase in the 29 R. A. Friedman and G. S. Manning, Biopolymers 23, 2671 (1984). 3o R. A. G. Friedman, G. S. Manning, and M. A. Shahin, in "Chemistry and Physics of DNA-Ligand Interactions" (N. R. Kallenbach, ed.), pp. 37~64. Adenine Press, Schenectady, New York, 1988. 31 M. T. Record, Jr., C. E Anderson, and T. M. Lohman, Q. Rev. Biophys. 11, 103 (1978). 32 G. S. Manning, Q. Rev. Biophys. 11, 179 (1978).

BIOPHYSICALAPPROACHES

10

[ 1]

average phosphate spacing, resulting in a systematic decrease in the polyelectrolyte contribution to the binding free energy. A closed form equation was derived 29,3° to embody this model for a univalent intercalator binding to B-form DNA in excess univalent salt solutions:

r K(2+r]-(2+~)lO_[c~i~2o2,1~+~](l_nr)[1-nr ],,-1 C---~=

\2-~-rJ

1 -(n --~l)r

(3)

In Eq. (3) K and n have the same meaning as given above, and the added exponential terms describe the decrease in polyelectrolyte contribution to the binding free energy over the course of lattice saturation. The parameter ff0 is the dimensionless charge density parameter and is a function of the structure of duplex DNA. Specifically, ~'0 = qZ/EkTb, where q is the charge of an electron, e is the bulk dielectric constant of the liquid, k is the Boltzmann constant, T is the temperature in degrees kelvin, and b is the charge spacing on the DNA chain. For standard B-form duplex DNA, b = 1.7 A, and ~'o = 4.2. The results of fits of the data shown in Fig. 2 to the Friedman-Manning model are listed in Table I. In all cases examined, the fits are statistically worse than those obtained with the simpler McGhee-von Hippel model. The standard deviations of the fits shown in Table I are all larger for the Friedman-Manning model than for the McGhee-von Hippel model when both K and n are allowed to vary, even though the former contains an additional parameter. The curvature imposed on the fitting function by the added exponential terms in Eq. (3) evidently makes it more difficult to match the curvature in the data. Statistically, therefore, there is no justification for use of the Friedman-Manning model instead of the simpler McGhee-von Hippel neighbor exclusion model. However appealing the underlying theory, the reality of the experimental data provides the ultimate test of the model. In this case, the inclusion of added complexity of polyelectrolyte effects is statistically unwarranted. Use of the neighbor exclusion model has become the standard practice in studies of ligand-DNA interactions. The preceding discussion, however, poses some serious questions about its use. Fractional values for the neighbor exclusion parameters are inevitably required to accurately describe the curvature in experimental data, yet have no meaning in the context of the model because integral values were assumed in the derivation of the model. Fractional neighbor exclusion values signify that the model is not an appropriate description of the actual data. The specific assumption of neighbor exclusion models that is violated is most likely that lattice sites are homogeneous. Chemical and enzymatic footprinting methods have shown unambiguously that such is not the case, and that most ligands bind to DNA sites with a wide distribution of affinities. 33'34 For the specific case of daunomycin, for examples, footprinting studies revealed a strong preference for triplet binding sites with the sequence 5'-(A/T)GC or 5'-(A/T)CG, where the notation

[1]

ANALYSIS OF L I G A N D - - D N A BINDING ISOTHERMS

11

(A/T) means that either A or T can occupy the position. 35'36 Further, sequences containing runs of AT base pairs were revealed by footprinting not to bind daunomycin with appreciable affinity. 16 All DNA lattice sites are clearly not identical, in which case a central tenet of the neighbor exclusion model is violated, vitiating its use as an appropriate model for the analysis of binding isotherms. This conclusion was strongly supported in the single example available, where both macroscopic binding studies and footprinting studies were carried out on the same homogeneous fragment, the 165-bp tyrT DNA fragment. 37 In that study, the macroscopic binding isotherm was complex in shape, could not be fit to the neighbor exclusion model, and clearly revealed a class of high-affinity binding sites whose number was consistent with the number of high-affinity sites visualized by the companion footprinting titration study. The neighbor exclusion principle has also been questioned on other grounds. Rao and Kollman 38 carried out molecular mechanics and molecular dynamics studies from which they concluded that there was no stereochemical basis for neighbor exclusion, at least for the intercalation of 9-aminoacridine into DNA. NMR studies from the Wilson laboratory 39'4° showed conclusively that actinomycin D can bind to adjacent 5'-GpC dinucleotide sites within an oligonucleotide, a finding that contrasts with the 5- or 6-bp exclusion parameter normally associated with the drug. While these are limited and perhaps specialized cases, they do raise questions about the exact physical basis for the neighbor exclusion phenomenon. Dinucleotide Binding Model If the neighbor exclusion model is excluded, what model can be used for the analysis of ligand-DNA binding data? One possibility is to assume heterogeneity of binding sites at the outset. Crothers in fact introduced an early variant of the neighbor exclusion model that incorporated simple base pair selectivity, with an added parameter to account for the relative affinity of GC versus AT base pairs. 23 That simple "two-site" model has not seen wide application. Another, more radical possibility is to abandon the neighbor exclusion concept entirely and to ascribe 33 j. C. Dabrowiak, A. A. Stankus, and J. Goodisman, in "Nucleic Acid Targeted Drug Design C" (L. Propst and T. J. Perun, eds.). Marcel Dekker, New York, 1992. 34 M. J. Waring and C. Bailly, J. Mol. Recognir 7, 109 (1994). 35 j. B. Chaires, K. R. Fox, J. E. Henera, M. Britt, and M. J. Waring, Biochemist~ 26, 8227 (1987). 36 j. B. Chaires, J. E. Herrera, and M. J. Waring, Biochemistry 29, 6145 (1990). 37 C. Bailly, D. Suh, M. J. Waring, and J. B. Chaires, Biochemistry 37, 1033 (1998). 38 S. N. Rao and P. A. Kollman, Proc. Natl. Acad. Sci. U.S.A. 84, 5735 (1987). 39 W. D. Wilson, R. L. Jones, G. Zon, E. V. Scott, D. L. Banville, and L. G. Marzilli, J. Am. Chem. Soc. 108, 7113 (1986). 40 E. V. Scott, R. L. Jones, D. L. Banville, G. Zon, L. G. Marzilli, and W. D. Wilson, Biochemist~ 27, 915 (1988).

12

BIOPHYSICALAPPROACHES

[ 11

the curvature in Scatchard plots entirely to heterogeneity. We have explored the simplest case in this scenario, a model in which a dinucleotide binding site is assumed. There are 16 possible dinucleotide combinations, 10 of which are unique. These are (5' -+ 3'): AT, AA (---- TT), TA, AC ( = GT), CA ( = TG), GC, GG ( = CC), CG, GA ( = TC), and AG ( = CT). Because intercalators insert between adjacent base pairs and make contact with both, dinucleotide selectivity is not an unreasonable starting point. For binding to dinucleotides (MN), each of which has a unique affinity (KMN), the binding isotherm is described by rD

MN fMNKMNCf

(4)

1 q'- KMNCf

where the binding ratio ro is now expressed as moles of ligand per mole of dinucleotide. The remaining variables are the dinucleotide frequency (fMN) and the free ligand concentration (C0. Dinucleotide frequencies were experimentally determined and tabulated for a numerous natural DNA samples, 41 and may be fixed as constants for a given DNA. Because there are 10 unique dinucleotide steps, the equation has 10 terms, and 10 binding constants must be obtained by nonlinear least-squares fitting of experimental data. Although at first glance the exercise of resolving 10 parameters may seem hopeless or even ludicrous, we show that it can in fact be done with the return of reasonable results. Figure 3 shows the fit of daunomycin-calf thymus DNA binding data to the dinucleotide model. Note that the binding ratio is now expressed in terms of total dinucleotide concentration rather than the usual base pair concentration. The solid line represents the best fit to the dinucleotide binding model. Estimates of 10 binding constants are obtained, and are summarized in Fig. 4. The data in Fig. 4 are average values of KMN estimates obtained from fitting daunomycin binding data obtained with eight different natural DNA samples with known and widely varying dinucleotide frequencies. These samples ranged from Clostridium perfringens DNA (31% GC content) to Micrococcus lysodeikticus DNA (72% GC content). Unique KMN values are returned that range over two orders of magnitude. Figure 4 shows that AA, AT, and TG steps represent low-affinity sites. TA, GC, and CG have intermediate affinity. High-affinity sites are TC, AC, AG, and GG steps. Does this analysis make sense? Is it valid? Several observations suggest that the answer is "yes" to both questions. First, footprinting studies of the daunomycinDNA interaction showed that sequences protected from DNase I cleavage by the bound drug were enriched in the dinucleotides AC, AG, GG, GC, and CG relative to the tyrT DNA fragment alone that was used for footprinting. 35 These are among the very dinucleotide steps with the highest KMN values. In contrast, analysis of unprotected cleavage sites from the footprinting experiments showed that such 41 G. D. Fasman, "Handbook of Biochemistry and Molecular Biology," 3rd Ed. CRC Press, Cleveland, Ohio, 1976.

[l ]

ANALYSIS OF LIGAND-DNA BINDINGISOTHERMS

13

0,75 ]

0.50

0.00

t _.t.._

1E-9

1E-8

1E-7

1E-6

1E-5

1E-4

Cf, M FIG. 3. Binding isotherm for the daunomycin-calf thymus DNA interaction. The binding ratio on the ordinate is expressed as moles of daunomycin bound per dinucleotide. Free daunomycin concentration is shown on the abscissa. The solid line represents the best fit to the 10-site, dinucleotide binding model described in text.

sites were enriched in the dinucleotides AA, AT, and TA. 16'35 Two of these steps have the lowest measured KMN. Second, KMN values can be used to calculate a total binding free energy for the loading of the calf thymus DNA lattice that is in excellent agreement with the model-independent approach described earlier in this chapter. The total free energy (AGT) is specified by MN

MN

AGT = ~ fMNAGMN = Z fMN(-RT In KMN)

(5)

where A G M N is the free energy for binding to the dinucleotide step MN. Using the known fMN values for calf thymus DNA and the KMN values shown in Fig. 4, a value of AGT = --7.8 kcal mol I is obtained, in excellent agreement with that obtained by model-independent analysis. Finally, the binding constants shown in Fig. 4 may be used in combination with the known dinucleotide frequency of calf thymus DNA to simulate a binding isotherm, as shown in Fig. 5. If these simulated data are then fit to the neighbor exclusion model, values o f K = 6.5 x 105 M - I and n -- 3.3 bp are obtained. These values are in excellent agreement with those obtained by fits to actual experimental data (Fig. 2). The key point of this exercise is

14

BIOPHYSICALAPPROACHES 10 7

I

I

!

108

I

I

I

I

[ 1] I

I

l~,//'~~i,X~ i

10 s

104

t AA

AT

TA

t

I

t

t

T G T C AC AG GG GC C G

Dinucleotide Step FIG. 4. Affinity profile for the interaction of daunomycin with the 10 unique dinucleotide steps. The binding constants (K) are average values obtained by fitting of binding isotherms for the interaction of daunorubicin with eight different DNA samples with widely varying dinucleotide frequencies. The error bars show standard deviations from the mean values.

that dinucleotide heterogeneity can account entirely for the curvature in Scatchard plots at least as well as the neighbor exclusion model does. The model presented in this section is somewhat radical and vitiates the conventional wisdom of the neighbor exclusion model. But because a fundamental assumption of the neighbor exclusion model (the homogeneity of potential binding sites) is demonstrably incorrect, the development of other models is mandatory. Furthermore, a few studies have questioned the physical basis of neighbor exclusion. 38-4° The dinucleotide model is consistent with the results of more than a decade of footprinting studies, which show that ligands inevitably bind with a wide range of affinities along the DNA lattice. In one sense, we have come full circle. Early binding isotherms for proflavin42 were curved and were interpreted in term of site heterogeneity, namely with two classes of sites later attributed to intercalation and "outside" binding. The dinucleotide model assumes a different kind

42 A. R. Peackocke and J. N. H. Skerren, Trans. Faraday Soc. 52, 261 (1956).

[1]

ANALYSIS OF L I G A N D - D N A BINDING ISOTHERMS I

I

I

I

~

15

I

5 x 1 0 5.

4xl 0 5

3x10 5

0 2x 10 5

lx10 5

I

I

I

I

I

0,05

0.10

0.15

0.20

0.25

r bound

0.30 ,-,

FIG. 5. Simulated Scatchard plot for the daunomycin-calf thymus DNA interaction. Dinucleotide binding constants (Fig. 4) were used in combination with the known dinucleotide frequency for calf thymus DNA to simulate binding data (solid circles), which were then cast into the form o f a Scatchard plot. If these simulated data are then fit to the neighbor exclusion model [Eq. (2)], the best fit (solid line) yields K -- 6.5 × 105M -1 and n = 3.3 bp.

of heterogeneity involving a larger number of sites derived from a fundamental property of any DNA, its nearest neighbor dinucleotide frequency.

Coping with Cooperativity In some cases, ligand-DNA binding isotherms exhibit evidence of positive cooperativity. In these cases, data cast into the form of a Scatchard plot show positive slopes at low binding ratios (Figs. 6 and 7). Analysis and interpretation of such isotherms become even more complicated. McGhee and yon Hippe124 (and

16

BIOPHYSICALAPPROACHES 4

I

I

I

[ 11 I

I

!3 v

2

+

%÷÷**~+÷%Q~. •

- -

4~m J.

~ $$ m

I 0

I

I

I

I

0.1

0.2

0.3

0.4

r-bound FIG. 6. Allosteric binding of daunomycin to poly(dA).poly(dT). Binding data were obtained for the interaction of daunomycin with poly(dA).poly(dT) in buffered 0.2 M NaCI solutions (pH 7.0). The solid line was calculated for the parameters listed in Table II for the Crothers allosteric binding model. The -I- + + line shows the best fit to the McGhee-von Hippel Neighbor exclusion model with added ligand-ligand cooperativity [Eq. (6)].

Crothers earlier 23) presented an extended form of the neighbor exclusion model that included an additional term to account for ligand-ligand interactions. The equation for this model is

r C--f = R =

K(l_nr)[(2w-1)(l-nr)+r-R]n-l[l-(n+l)r+R]2 Y(~--- U d - n ~ 2(1 - nr) {[1 - (n + l)r] 2 + 4mr(1 - nr)} 1/2

(6)

where K, n, and r have the same meaning as given above, and co is the cooperativity parameter, co is defined as the equilibrium constant for moving two ligands bound at isolated sites into proximity, such that they occupy contiguous lattice sites. If ~o > 1.0, positive cooperativity results and ligands bind preferentially next to one another. If ~o < 1.0, negative cooperativity results, and ligand binding at adjacent sites is hampered. A distinctive feature of this model is that the DNA lattice remains and inert array of identical sites. All cooperativity results from ligandligand interactions of an unspecified nature. A contrasting model that can account for positive cooperativity is the Crothers allosteric model. 43 The underlying concept is radically different from the McGhee-von Hippel model. The allosteric model is analogous to the classic Monod-Wyman-Changeux model for allostery 44 derived to explain the cooperative 43 N. Dattagupta, M. Hogan, and D. M. Crothers, Biochemistry 19, 5998 (1980). 44 j. Monod, J. Wyman, and J.-P. Changeux, Z Mol. Biol. 12, 88 (1965).

[ l]

ANALYSIS OF LIGAND--DNA BINDING ISOTHERMS I

I

I

I

17

I

1.2x10 s

1.0xlO s

Z Form

8.0x104

"')

•

B Form

6.0xl 0 4

4.0x10 4

2.0xl 0 4

0.0 I

0.0

~

I

0.1

~

I

~

0.2

I

0.3

~

I

0.4

0.5

r

FIG. 7. Allosteric binding of daunomycin to left-handed Z-DNA. Binding data were obtained for the interactions of daunomycin with Z-form [poly(dG-dC)]2in buffered 3.0 M NaCI solutions (pH 7.0). The solid line was calculated for the parameters listed in Table II for the Crothers allosteric binding model. At the start of the binding isotherm, the polymer is in the left-handed Z form. Beyond the maximum near r = 0.3, daunomycin binding has allosterically converted the polymer to the right-hand form.

b i n d i n g of ligands to proteins, such as the b i n d i n g of o x y g e n to hemoglobin. Crothers built on the basic concept of allostery, but included specific details of the m a c r o m o l e c u l a r conformational transition and ligand b i n d i n g that were appropriate for a D N A lattice. The allosteric model assumes that the D N A lattice can exist in one of two conformational forms (1 and 2). The transition of the lattice between these forms proceeds by a nucleation step, followed by propagation steps: O-2S

Nucleation:

...111111...

< ~ ... 112111...

Propagation:...112111...

< ~ ... 112211...

S

The equilibrium constant for nucleation is cr2s and that for propagation is s. The allosteric model then assumes that ligand can bind to each conformational form

18

BIOPHYSICAL APPROACHES

[1]

with unique neighbor exclusion binding parameters, (K1, n l, o~1) and ( K 2 , t/2,092). Cooperativity arises when the ligand binds selectively or preferentially to one of the conformational forms. Binding then drives an allosteric conformational transition to the form with higher ligand affinity. Eight parameters are needed to compute binding isotherms. There is no closed form, analytic equation for the allosteric model. Crothers et al. wrote a Fortran program that calculates binding isotherms according to the derived statistical mechanical model, 43 a program that has seen modest circulation 45 47 and that has been modified on occasion 4s'49 for easier use. No nonlinear fitting routine has yet been developed that incorporates the allosteric model, and users must optimize binding parameters by successive approximation along with judicious constraint of those selected parameters that can be estimated by independent methods. 5° Nonetheless, use of the allosteric model is unavoidable in some cases, as examples will show. It is important to contrast the key features of the McGhee-von Hippel and allosteric models. In the McGhee-von Hippel model, positive cooperativity arises from ligand-ligand interactions while the DNA lattice remains in a single conformation. For protein binding to DNA, such ligand-ligand interactions might be visualized as protein-protein contacts formed when adjacent lattice sites are occupied. For small molecule ligands, although such interactions could also occur, it is less easy to ascribe a molecular picture to the process. In contrast, positive cooperativity arises in the Crothers allosteric model from an underlying conformational transition in the DNA lattice to a form with higher ligand binding affinity. The allosteric model was perhaps the first clear statement of the possibility of structural selectivity in ligand-DNA interactions. Two examples will illustrate positive cooperativity in ligand-DNA interactions. Figure 6 shows the binding of daunomycin to poly(dA) • poly(dT). Independent studies have shown that poly(dA), poly(dT) undergoes a premelting transition between two helical forms. 51-53 The binding data in Fig. 6 show a positive slope at low r values, and pass through a maximum near r = 0.05. Attempts to fit these data to the extended McGhee-von Hippel model yield unsatisfactory results. The best fit to that model (with K = 1.4 × 104M -j , n = 2.3 bp, ~o = 2.4) is shown by the 45 j. B. Chaires, J. Biol. Chem. 261, 8899 (1986). 46 G. Y. Walker, M. E Stone, and T. R. Krugh, Biochemistry 24, 7462 (1985). 47 G. T. Walker, M. E Stone, and T. R. Krugh, Biochemistry 24, 7471 (1985). 48 G. T. Walker, Ph.D. Thesis, University of Rochester, 1986. 49 D. Snh, Ph.D. Thesis, University of Mississippi Medical Center, 1993. 5o j. B. Chaires, J. Biol. Chem. 261, 8899 (1986). 51 j. E. Herrera and J. B. Chaires, Biochemistry 28, 1993 (1989). 52 S. S. Chan, K. J. Breslauer, M. E. Hogan, D. J. Kessler, R. H. Austin, J. Ojemann, J. M. Passner, and N. C. Wiles, Biochemistry 29, 6161 (1990). 53 S. S. Chan, K. J. Breslauer, R. H. Austin, and M. E. Hogan, Biochemistry 32, 11776 (1993).

[1]

ANALYSIS OF L I G A N D - D N A BINDING ISOTHERMS

19

TABLE II PARAMETER ESTIMATES FOR ALLOSTERICMODEL "FIT" TO DAUNOMYCINBINDING TO POLY(dA).POLY(dT) AND Z-DNA Parameter o" s

KI(M -I ) K2(M -I ) K2/KI ?11 n2

Description

Poly (dA).Poly(dT) a

Z-DNA ~'

Nucleation parameter Propagation constant Association constant for binding to form 1 Association constant for binding to form 2 Ratio of association constants Neighbor exclusion parameter (form 1) Neighbor exclusion parameter (form 2)

0.01 0.985 4,800 21,000 4.3 2.0 2.0

0.00 l 0.635 8,000 350,000 44 2.0 2.3

a Daunomycin binding to poly(dA).poly(dT) in buffered 0.2 M NaCI solutions (pH 7.0). Parameters result from analysis of the binding data shown in Fig. 7. b Daunomycin binding to Z-form [poly(dG-dC)]2 in buffered 3.0 M NaCI solutions (pH 7.0). Parameters result from analysis of the binding data shown in Fig. 6.

curve with the (+) symbols; no value of ~o can be found that can produce a curve that matches the steep positive slope at the start of the isotherms. The model, with an assumed inert DNA lattice, cannot match the experimental data. In contrast, the allosteric model can match the shape of the binding data well, as shown by the solid curve in Fig. 6. The best estimates of the parameters for the allosteric model that describe the binding data are collected in Table II. The key driving force for the allosteric conversion of poly(dA), poly(dT) is the >4-fold preference for binding to form 2 of the polynucleotide. A variety of additional physical and enzymatic tools was used to demonstrate that daunomycin binding was indeed coupled to a conformational change in poly(dA), poly(dT) 51 the essential feature of the allosteric model. Figure 7 shows daunomycin binding to poly(dG-dC) under solution conditions that initially favor the left-handed Z conformation of the polynucleotide. 45'54 The allosteric model with the parameters listed in Table II was used to obtain the solid curve matching the experimental data. In this case, the underlying conformational transition to which binding is coupled is the Z-to-B conversion. Compared with the first example of binding to poly(dA) • poly(dT), this system is much more cooperative. The reason for that is the greater preference of daunomycin for the right-hand form relative to the left-handed form (with K z / K I -~ 44; Table II), compared with its preference for the two right-handed helical forms of poly(dA) • poly(dT) (with K2/KI = 4; Table II).

54 X. Qu, J. O. Trent, I. Fokt, W. Priebe, and J. B. Chaires, Proc. Natl. Acad. Sci. U.S.A. 97, 12032

(2O0O).

20

BIOPHYSICAL APPROACHES

[ 1]

L i g a n d B i n d i n g to O l i g o n u c l e o t i d e s Advances in synthetic methods have made it relatively easy to prepare DNA or RNA oligonucleotides of precisely defined length and sequence, and it is now fashionable to use such molecules for ligand binding studies. The significant advantage of oligonucleotides is their homogeneity. There are, however, disadvantages. First, end effects may become a consideration in oligonucleotide studies. Neighbor exclusion models typically assume an "infinite lattice" specifically to avoid end effects, and therefore such models become inapplicable to oligonucleotide systems. Polyelectrolyte theory and experiment have shown that significant end effects exist for oligonucleotides less than about 24 bp in length. 55'56 Second, problems of appropriate representation of all possible sequence elements may arise in oligonucleotide systems. There are 10 unique dinucleotide combinations, as discussed above, and it is rare that a given oligonucleotide is appropriately designed to contain all possible dinucleotide steps. It is always possible that a high-affinity interaction may go undetected because the appropriate site is absent in the oligonucleotide chosen for study. Binding constants may thus be biased by the choice of sequence unless a large number of oligonucleotides is studied. For trinucleotides, the situation becomes even worse, because there are 64 possible triplets, 32 of which are unique. These concerns should not be taken as a call to abrogate oligonucleotide studies, but rather to acknowledge their appropriate role. Complete binding studies should be comprehensive and should systematically move from long natural DNA samples through synthetic polynucleotides of defined simple repeating sequences to oligonucleotides of precisely defined sequence. Oligonucleotide systems are of particular value for the study of the energetics of binding when other experiments have defined the sequence of the preferred binding site for a given ligand. Neighbor exclusion models are generally inapplicable for the analysis of ligand binding to oligonucleotides because they are based on an "infinite lattice" assumption. Binding isotherms in these cases are best analyzed by the classic and simple stoichiometric binding models that are described in any number of texts and monographs. 2-4 Figure 8 shows an example of daunomycin binding to a 24-bp duplex oligonucleotide with the sequence (5'-TGCATGCATGCATGCATGCATGCA)2. This oligonucleotide was designed and synthesized to contain a repetitive motif containing the preferred daunomycin binding site that emerged from footprinting studies [5'-(A/T)GC; where (A/T) means A or T]. Binding data were fit to the simple expression for multiple identical, noninteracting sites: r

~tK Cf -

-

-

-

1 +KCf

(7)

55M. C. Olmsted,C. E Anderson,and M. T. Record,Jr., Proc. Natl. Acad. Sci. U.S.A. 86, 7766(1989), 56M. C. Olmsted,C. E Anderson,and M. T. Record,Jr., Biopolymers 31, 1593(1991).

[ 1]

ANALYSIS OF L I G A N D - D N A BINDING ISOTHERMS

I

I

!

I

2

4

6

8

21

(D "O

"5

EI1)

cO

O to

E3

0

0

10

Ct~ee,gM FIG. 8. Binding of daunomycin to a 24-bp duplex oligonucleotide. Data (solid circles) are presented as a direct plot, with the best fit to Eq. (7) shown as the solid line.

where r is now expressed as moles of daunomycin bound per mole of oligonucleotide, n is the number of sites, and Kis the association constant. Nonlinear leastsquares fitting of the data yields K = 4.0 (4-0.1) × 106M - 1 and n = 4.3 (-t-0.3). Evidently only four molecules of daunomycin are binding to each oligonucleotide, even though there are six potential sites that contain the preferred triplet sequence. It is possible that the sites near the ends are disfavored, or that there is anticooperativity that disfavors binding to adjacent sites. In the latter case, more complicated models with added parameters would need to be invoked and implemented. An excellent example of such an analysis applied to daunomycin binding to hexanucleotide sequences was provided by Rizzo and co-workers. 57 Summary Binding studies provide information of fundamental and central importance for the complete understanding of ligand-DNA interactions. Studies of ligand binding to long natural DNA samples, to synthetic deoxypolynucleotides of simple repeating sequence, and to oligonucleotides of defined sequence are all needed to 57 V. Rizzo, C. Battistini, A. Vigevani, N. Sacchi, G. Razzano, F. Arcamone, A. Garbesi, E P. Colonna, M. Capobianco, and L. Tondelli, J. Mol. Recognit. 2, 132 (1989).

22

BIOPHYSICALAPPROACHES

[21

begin to understand the interaction in detail. Binding studies provide entry into the thermodynamics of the DNA interactions, which in turn provides great insight into the molecular forces that drive the binding process. This chapter summarizes both model-dependent and -independent approaches for the analysis and interpretation of binding isotherms, and should serve as a concise guide for handling experimental data. Acknowledgment W o r k in the a u t h o r ' s l a b o r a t o r y w a s f u n d e d b y g r a n t C A 3 5 6 3 5 f r o m the N a t i o n a l C a n c e r Institute.

[2] S u r f a c e P l a s m o n R e s o n a n c e of RNA-Small Molecule

Biosensor Analysis Interactions

By TINA M. DAVIS and W. DAVID WILSON Introduction Because some of the most devastating human diseases are caused by RNA viruses, such as human immunodeficiency virus (HIV) and hemorrhagic fever viruses such as dengue and Ebola, l RNA is an attractive therapeutic target in drug design. 2-4 The unique structural folds present in RNA, but not DNA, offer the possibility of much higher recognition specificity by small molecules. Structured sites in RNA can be targeted with the same specificity with small molecules as structured binding regions of proteins. Despite the potential advantages of targeting RNA, few drugs are known that target RNA, and rational design of drugs that target RNA is in the beginning stages.5-SA summary of some of the RNAs that have been successfully targeted in drug design and interaction studies is shown in Table 1.2,9-26 I M. B. A. Oldstone, "Viurses, Plagues, and History." Oxford University Press, Oxford, 1998. 2 W. D. Wilson and K. Li, Curr. Med. Chem. 7, 73 (2000). 3 K. Michael and Y. Tor, Chem. Eur. J. 4, 2091 (1998). 4 F. Walter, Q. Vicens, and E. Westhof, Curr. Opin. Chem. Biol. 3, 694 (1999). 5 K. Li, M. Fernandez-Saiz, C. T. Rigl, A. Kumar, K. G. Ragunathan, A. W. McConnaughie, D. W. Boykin, H. J. Schneider, and W. D. Wilson, Bioorg. Med. Chem. 5, 1157 (1997). 6 T. Hermann and W. Westhof, Curt Opinion Biotechnol. 8, 278 (1998). 7 C. Chow and F. M. Bogdan, Chem. Rev. 97, 1489 (1997). 8 M. Afshar, C. D. Prescott, and G. Varani, Curr. Opin. BiotechnoL 10, 59 (1999). 9 M. J. Rogers, Y. V. Bukhman, R. E McCutchan, and D. E. Draper, RNA 3, 815 (1997). 10 G. L. Conn, R. R. Gutell, and D. E. Draper, Biochemistry 37, 11980 (1998).

METHODS IN ENZYMOLOGY, VOL. 340

Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. 0076-6879f00 $35.(X)

22

BIOPHYSICALAPPROACHES

[21

begin to understand the interaction in detail. Binding studies provide entry into the thermodynamics of the DNA interactions, which in turn provides great insight into the molecular forces that drive the binding process. This chapter summarizes both model-dependent and -independent approaches for the analysis and interpretation of binding isotherms, and should serve as a concise guide for handling experimental data. Acknowledgment W o r k in the a u t h o r ' s l a b o r a t o r y w a s f u n d e d b y g r a n t C A 3 5 6 3 5 f r o m the N a t i o n a l C a n c e r Institute.

[2] S u r f a c e P l a s m o n R e s o n a n c e of RNA-Small Molecule

Biosensor Analysis Interactions

By TINA M. DAVIS and W. DAVID WILSON Introduction Because some of the most devastating human diseases are caused by RNA viruses, such as human immunodeficiency virus (HIV) and hemorrhagic fever viruses such as dengue and Ebola, l RNA is an attractive therapeutic target in drug design. 2-4 The unique structural folds present in RNA, but not DNA, offer the possibility of much higher recognition specificity by small molecules. Structured sites in RNA can be targeted with the same specificity with small molecules as structured binding regions of proteins. Despite the potential advantages of targeting RNA, few drugs are known that target RNA, and rational design of drugs that target RNA is in the beginning stages.5-SA summary of some of the RNAs that have been successfully targeted in drug design and interaction studies is shown in Table 1.2,9-26 I M. B. A. Oldstone, "Viurses, Plagues, and History." Oxford University Press, Oxford, 1998. 2 W. D. Wilson and K. Li, Curr. Med. Chem. 7, 73 (2000). 3 K. Michael and Y. Tor, Chem. Eur. J. 4, 2091 (1998). 4 F. Walter, Q. Vicens, and E. Westhof, Curr. Opin. Chem. Biol. 3, 694 (1999). 5 K. Li, M. Fernandez-Saiz, C. T. Rigl, A. Kumar, K. G. Ragunathan, A. W. McConnaughie, D. W. Boykin, H. J. Schneider, and W. D. Wilson, Bioorg. Med. Chem. 5, 1157 (1997). 6 T. Hermann and W. Westhof, Curt Opinion Biotechnol. 8, 278 (1998). 7 C. Chow and F. M. Bogdan, Chem. Rev. 97, 1489 (1997). 8 M. Afshar, C. D. Prescott, and G. Varani, Curr. Opin. BiotechnoL 10, 59 (1999). 9 M. J. Rogers, Y. V. Bukhman, R. E McCutchan, and D. E. Draper, RNA 3, 815 (1997). 10 G. L. Conn, R. R. Gutell, and D. E. Draper, Biochemistry 37, 11980 (1998).

METHODS IN ENZYMOLOGY, VOL. 340

Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. 0076-6879f00 $35.(X)

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23

TABLE I SMALL MOLECULESAND THEIR BIOLOGICALRNA TARGETS Sequence and secondary structure

RNA

Small molecule ligand

Ref.

uAA A rRNA OTPase

AOCAGccAUcAuU AGAuUCGGuuG UA,~,

A# AGcGUA cC U CGAuA

Thiostrepton

9, 10

Aminoglycosides

11-13

A U 5'-GCC GGU-3' rRNA A site

AC CC -CCAC UGC GC-5' -GGUG UCG # A - 3 '

AAa U~

tRNA

tRNA phe

Hoechst 33258 Bleomycin

14, 15, 16

Ribozymes

Hepatitis delta virus ribozyme Hammerhead RNA Group I intron

Aminoglycosides

17, 18, 19

HIV- 1 RRE

AAcuGCGAC GC CG GUA-5' uGACGCUCACGGuACAG-3'

Aminoglycosides Diphenylfuran polycations

20 21, 22

HIV-I TAR

GUC~GA'UCu" ' t tJ 5A GA-5 G G AGCUC --,- UCU-3'

Aminoglycosides Aminoquinozalines Acridine polycations

23 24 25

ACc G C G c c G C c c CC-5' CUCCUGUGG-GGUC_3' G U

Aminoglycosides

26

Thymidylatesynthase mRNA

RNA-ligand interactions can be investigated by the same techniques as DNAligand interactions. Nuclear magnetic resonance (NMR) 27-33 and X-ray crystallography34 can be used to obtain detailed structural information about RNA-ligand complexes, although obtaining the quantity of RNA necessary for such studies is Jl D. Fourmy, M. I. Recht, S. C. Blanchard, and J. D. Puglisi, Science 274, 1367 (1996). 12 D. Fourmy, S. Yoshizawa, and J. D. Puglisi, J. Mol. Biol. 277, 333 (1998). 13 M. I. Recht, D. Fourmy, S. C. Blanchard, K. D. Dahlquist, and J. D. Puglisi, J. Mol. BioL 262, 421 (1996). 14E. g. Bichenkova, S. E. Sadat-Ebrahimi, A. N. Wilton, N. O'Toole, D. S. Marks, and K. T. Douglas, Nucleosides Nucleotides 17, 1651 (1998). 15 S. E. S. Ebrahimi, A. N. Wilton, and K. T. Douglas, Chem. Commun. 4, 385 (1997). 16C. E. Holmes, R. J. Duff, G. A. van der Marel, J. van Boom, and S. M. Hecht, Bioorg. Med. Chem. 5, 1235 0997). 17 U. yon Ahsen, J. Davies, and R. Schroeder, Nature (London) 353, 368 (1991 ). I8 j. Rogers, A. H. Chang, U. yon Ahsen, and R. Schroeder, ,l. Mol. Biol. 259, 916 (1996).

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often much more costly than it is for DNA. In addition, making NMR resonance assignments for RNA is often much more difficult than it is for DNA because of decreased spectral dispersion in the NMR spectra of RNA. Chemical and nuclease footprinting can provide information regarding drug-binding site(s) and sequence specificity of binding. 32'33'35-38 Theoretical studies of RNA complexes have reached a high level of sophistication and can now independently provide important structural and dynamic information. 4 Melting experiments by spectroscopic methods such as UV and circular dichroism (CD) can provide information regarding the thermodynamics of drug binding.5,38 43 CD has the added ability to provide information regarding the

19 T. Hermann and E. Westhof, J. Mol. Biol. 276, 903 (1998). 2o M. L. Zapp, S. Stern, and M. R. Green, Cell 74, 969 (1993). 21 K. Li, G. Xiao, T. Rigl, A. Kumar, D. W. Boykin, and W. D. Wilson, in "Structure, Motion, Interaction and Expression of Biological Macromolecules: Proceedings of the 10th Conversation in the Discipline Biomoleculor Stereodynamics," University of Albany, New York (R. H. Sarma and M. H. Sarma, eds.), pp. 137-145. Adenine Press, Schenectady, New York, 1998. 22 L. Ratmeyer, M. L. Zapp, M. R. Green, R. Vinayak, A. Kumar, D. W. Boykin, and W. D. Wilson, Biochemistry 35, 13689 (1996). 23 S. Wang, P. W. Huber, M. Cui, A. W. Czaruik, and H.-Y. Mei, Biochemistry 37, 5549 (1998). 24 H.-Y. Mei, D. P. Mack, A. A. Galan, N. S. Halim, A. Heldsinger, J. A. Loo, D. W. Moreland, K. A. Sannes-Lowery, L. Sharmeen, and A. W. Czarnik, Bioorg. Med. Chem. 5, 1173 (1997). 25 E Hamy, V. Brondani, A. Florsheimer, W. Stark, M. J. J. Blommers, and T. Klimkait, Biochemisttiy 37, 5085 (1998). 26 A. R. Ferre-D'Amare, K. Zhou, and J. A. Doudna, Nature (London) 395, 567 (1998). 27 M. Katahira, S. Kobayashi, A. Matsugami, K. Ouhashi, S. Uesugi, R. Yamamoto, K. Taira, S. Nishikawa, and P. Kumar, Nucleic Acids Symp. Sel: 42, 269 (1999). 28 W. H. Gmeiner, Curt Med. Chem. 5, 115 (1998). 29 S. Yoshizawa, D. Founny, and J. D. Puglisi, EMBO J. 17, 6437 (1998). 30 Q. Chen, R. H. Shafer, and 1. D. Kuntz, Biochemistry 36, 11402 (1997). 31 L. Jiang, A. K. Suri, R. Fiala, and D. J. Patel, Chem. Biol. 4, 35 (1997). 32 E Hamy, V. Brondani, A. Florsheimer, W. Stark, M. J. Blommers, and T. Klimkait, Biochemistry 37, 5086 (1998). 33 M. I. Recht, D. Fourmy, S. C. Blanchard, K. D. Dahlquist, and J. D. Puglisi, J. Mol. Biol. 262, 421 (1996). 34 E Takusagawa, K. T. Takusagawa, R. G. Carlson, and R. F. Weaver, Bioorg. Med. Chem. 5, 1197 (1997). 35 N. Gelus, C. Bailly, E Hamy, T. Klimkait, W. D. Wilson, and D. W. Boykin, Biool2~. Med. Chem. 7, 1089 (1999). 36 N. Gelus, E Hamy, and C. Bailly, Bioorg. Med. Chem. 7, 1075 (1999). 37 G. Rosendahl and S. Douthwaite, Nucleic Acids Res. 22, 357 (1994). 38 L. Dassonneville, E Hamy, P. Colson, C. Houssier, and C. Bailly, Nucleic Acids Res. 25, 4487 (1997). 39 j. E. Draper and T. C. Gluick, Methods Enzymol. 259, 281 (1995). 4o D. S. Pilch, M. A. Kirolos, X. Liu, G. E. Plum, and K. J. Breslauer, Biochemistry 34, 9962 (1995). 41 U. Sehlstedt, P. Aich, J. Bergman, H. Vallberg, B. Norden, and A. Graslund, J. Mol. Biol. 278, 31 (1998).

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mode of drug binding to RNA (groove binding versus intercalation). Fluorescence experiments are useful for obtaining binding information provided the small molecule has fluorescent properties that are altered on binding to RNA. 23 Calorimetric techniques such as differential scanning calorimetry (DSC) and isothermal calorimetry (ITC) provide direct quantitative information regarding the thermodynamics of drug binding and are perhaps the most reliable methods for obtaining such information. 4°-42 Calorimetric methods, however, usually require substantially more material than spectrophotometric methods, and they have difficulty measuring large binding constants. Gel band shift has been the most widely used method for studying nucleic acid-ligand interactions. 5,35 Providing that the RNA-small molecule complex can be separated from free RNA (i.e., there is a detectable band shift from drug binding) and that the binding kinetics are relatively slow, the technique can provide accurate binding constants and stoichiometry with only minimal quantities of RNA and drug. The technique is particularly useful for screening focused combinatorial libraries that meet the above-described criteria. Many RNA-small molecule complexes are not easily separated on gels, however, or have kinetics that are too fast for resolution on gels. It is clear that additional methods are required for such complexes. Several newer techniques for studying nucleic acid-small molecule interactions have been described. Ibis Therapeutics (Carlsbad, CA) a division of Isis Pharmaceuticals, has developed a sophisticated mass spectrometry-based assay for screening mixtures of small molecules for binding to RNA. 44-46 The technique permits multiple ligands and multiple RNAs to be screened simultaneously. In addition, the technique can provide binding affinities and binding sites on RNA. This is an excellent method for analysis of multiple compounds when the appropriate equipment is available. Luedtke and Tor 47 have developed a novel solid-phase assay for investigating RNA-small molecule interactions. The assay was used to identify small molecules that bind to the Rev-responsive element (RRE) fragment of the HIV virus and was based on the competition between potential RNA binders and a fluorescent Rev peptide. A biotinylated RRE fragment was immobilized on insoluble agarose beads that were covalently modified with streptavidin. A ternary complex was formed 42 Z. Xu, D. S. Pilch, A. R. Srinivasan, W. K. Olson, N. E. Geacintov, and K. J. Breslauer, Bioorg. Med. Chem. 5, 1137 (1997). 43 G. Luck, C. Zimmer, and B. C. Baguley,Biochim. Biophys. Acta 782, 41 (1984).

44 S. A. Hofstadler, K. A. Sannes-Lowery,S. T. Crooke, D. J. Ecker, H. Sasmor, S. Manalili, and R. H. Griffey, Anal. Chem. 71, 3436 (1999). 45 R. H. Griffey, S. A. Hofstadler, K. A. Sannes-Lowery,D. J. Ecker, and S. T. Crooke, Proc. Natl. Acad. Sci. U.S.A. 96, 10129 (1999). 46 K. Hamasaki and R. R. Rando, Anal. Biochem. 261, 183 (1998). 47 N. W. Luedtke and Y. Tor, Angew. Chem. 39, 1788 (2000).

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on addition of fluorescein-labeled Rev peptide. Ligand binding was indirectly detected by monitoring the quantity of fluorescent peptide either released into solution or remaining on the solid support as ligand bound to the RNA. The assay offers a simple, rapid, and quantitative method that facilitates the discovery and characterization of RNA binders directed at a specific complex. Surface plasmon resonance (SPR) biosensors are another relatively new technique for characterizing nucleic acid-small molecule interactions. Although SPR biosensors have been commercially available for more than a decade, their application to nucleic acid-small molecule systems is just developing. Only a handful of SPR articles describing macromolecule-small molecule interactions have been published to date, and only a few of these focus on nucleic acid-small molecule systems.48 51 As with the methods described above, SPR offers a rapid and powerful tool for screening small molecule libraries. 49'52 An advantage of SPR is that the method simultaneously can provide kinetic and equilibrium characterization of the interactions of active compounds with macromolecules. 53 Such information gives important insight for understanding whether inhibitory activity is governed by the kinetic or equilibrium (or both) properties of the interaction. For example, Hendrix e t al. found that whereas two similar aminoglycoside antibiotics have significantly different inhibitory potency, their binding constants are rather similar9 The dissociation rates for the two molecules, however, are substantially different, accounting for the more than 100-fold difference in inhibitory activity. When one of the interacting species can be captured on a biosensor surface, the advantages of SPR over conventional interaction analyses can be numerous. SPR monitors molecular interactions in real time 53 and is a significant improvement over optical methods for systems involving strong binding and/or low absorbance or fluorescence. In addition, material requirements are minimal when compared with spectrophotometric techniques, generally requiring only picomole to nanomole quantities, while simultaneously providing accurate and reproducible data. Although excellent articles on nucleic acid-small molecules have begun to appear, to our knowledge, a comprehensive guide for investigating nucleic acid-small molecule systems through SPR has not been published to date. For this reason and because of the commercial availability of high-sensitivity SPR instruments that are amenable for studying these systems, we have focused this review on how to study RNA-small molecule interactions with SPR biosensors. 48 C.-H. Wong,M. Hendrix, W. S. Priestley, and W. A. Greenberg, Chem. Biol. 5, 397 (1998). 49C.-H. Wong,M. Hendrix, D. D. Manning, C. Rosenbohm,and W. A. Greenberg,J. Am. Chem. Soc. 120, 8319 (1998). 50M. Hendrix, E. S. Priestley, G. E Joyce, and C.-H. Wong,J. Am. Chem. Soc. 119, 3641 (1997). 51 L. Wang, C. Bailly, A. Kumar, D. Ding, M. Bajic, D. W. Boykin, and W. D. Wilson, Proc. Natl. Acad. Sci. U.S.A. 97, 12 (2000). 52 P.-O. Markgren, M. Hamalained, and U. H. Danielson,Anal. Biochem. 265, 340 (1998). 53 Biacore, "BIACoreBIAapplicationsHandbook."Biacore, Uppsala, Sweden, June 1994.

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RNA INTERACTIONS

Description of Surface Plasmon Resonance

27 Biosensors

SPR biosensors employ surface plasmon resonance to qualitatively and/or quantitatively describe molecular interactions. Although several companies now offer SPR biosensors, 54 currently, instruments from BIAcore (Uppsala, Sweden) are the most widely used. The BIAcore 2000 biosensor is used in our laboratory, and the experimental guidelines and technical details presented in this review are specific to this biosensor and the BIAcore 3000 biosensor. However, the basic principles of using SPR to study molecular interactions are universal. In biosensor experiments, the interaction between molecules in solution (the analyte in BIAcore literature) and molecules immobilized on a sensor chip surface (the ligand in BIAcore literature) is monitored in real time. ~3 Labeling is not required and there are many immobilization chemistries available depending on experimental designY Biosensor experiments involve immobilizing one of the interacting species on a sensor chip surface either covalently or through affinity capture (such as biotinstreptavidin or antibody-antigen) to produce a biospecific surface. 55 Binding of analyte to this biospecific surface is monitored through SPR in a thin gold film that is the base of all sensor chips. The change in SPR response is directly related to changes in the refractive index at the biospecific surface. The refractive index at the surface is directly related to the concentration of molecules at the surface. Binding of analyte to the immobilized ligand, for example, causes a change in the refractive index at the surface, which in turn leads to a change in the reflected light angle at which SPR is observed (the SPR angle). Binding data are presented in the form of a sensorgram, which is a plot of the SPR angle, converted to resonance units (RU), versus time 53 (Fig. 1). For the most commonly used sensor chips, a change of 1000 RU is equivalent to binding of about 1 ng of protein and 0.8 ng of nucleic acid surface concentration per millimeter squared. 53,56,57 The greater response per nanogram of nucleic acid is a result of their generally higher refractive index increment with respect to proteins. Small molecules can have quite different refractive index increments (RIIs) than proteins and nucleic acids and the importance of this difference is discussed below. Experimental

Design and Protocols

Experimental Design~Considerations

There are several important factors to consider when designing an SPR experiment. At the forefront of experimental design are the questions of which flow cell surface to use, which species to immobilize, what chemistry to use to immobilize it, 54R. L. Rich and D. G. Myszka,Curl: Opin. Biotechnol. 11, 54 (2000). 55 Biacore, "BIACoreBIAtechnologyHandbook." Biacore, Uppsala, Sweden, June 1994. 56M. Buckle, R. M. Williams, M. Negroni, and H. Buc, Proc. Natl. Acad. Sci. U.S.A. 93, 889 (1996). 57R. J. Fisher, M. Fivash, J. Casasfinet, J. W. Erickson, A. Kondoh, S. V. Bladen, C. Fisher, D. K. Watson, and T. Papas, Protein Sci. 3, 257 (1994).

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3 n-

steadyassociation state

dissociation

.1 ~

-50

0

50

100 Time

150

200

250

300

350

(second)

FIG. 1. Example sensorgramcollected on a BIAcore 2000 biosensor.The association, steady state, and dissociation regions are indicated. The data result from binding of a small molecule (5 nM) to immobilizednucleic acid. and how much of it to immobilize. In an ideal experiment, both interacting species would be immobilized in reverse experiments to ensure that any data collected is not affected by the immobilization and other experimental factors. If only one can be immobilized then it is advantageous to immobilize the lower molecular weight species because the response signal on analyte binding is dependent on the molecular weight (the greater the molecular weight, the greater the signal). 53 In RNA (or D N A ) - s m a l l molecule systems, it is often impractical or impossible to immobilize the small molecule, especially when the purpose of the experiment is to screen a small molecule library. Subtle changes in a small molecule can drastically affect binding, making perturbation-free immobilization and data interpretation difficult. In contrast, immobilizing RNA through a 5' or 3' tether is unlikely to affect specific binding at a site in the interior of the molecule. Fortunately, we have found

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excellent signal-to-noise ratios (S/N) with small molecule binding, and where it has been possible to determine the equilibrium constant through both solution and SPR methods for small molecule-nucleic acid complexes, the constants are in excellent agreement. 58 Measuring binding of the low molecular weight compound does, however, require considerable attention to baseline stability. Methods that we have found to give the necessary S/N ratios are presented throughout. For immobilizing nucleic acids, the biotin-streptavidin affinity complex has been the immobilization method used in the vast majority of BIAcore SPR experiments,48-51.59-63 although other immobilization methods are appearing. 64 In the biotin-streptavidin immobilization method, biotinylated RNA (see Preparation of RNA, below) is immobilized on a sensor chip that has been derivatized with streptavidin (see Preparation of Sensor Chip, below). The rather large affinity constant for the biotin-streptavidin complex creates a quite stable binding surface over the time course of most experiment. However, in our experience, streptavidinnucleic acid sensor chips gradually lose binding activity through repeat use. We have been able to immobilize additional nucleic acid onto the sensor chip to achieve the original binding level but the activity of the surface continues to decline. Alternative immobilization strategies, including covalent attachment on activated carboxymethylated dextran through both a primary amine and a thiol group tethered to the 5' end of the DNA, offer the potential for longer chip stability. However, in our hands, neither strategy yielded enough immobilized nucleic acid for small molecule studies. It appears that the negative charge on the carboxymethyl dextran inhibits sufficient immobilization of the negative nucleic acid even at extremes of salt and pH in reasonable time periods. We were, however, able to quantitatively immobilize a 5'-thiolated nucleic acid directly onto a gold surface, using the method outlined in Heine and Tarlov,65 but the sensor chip lost significant activity rapidly, probably due to noncovalent adsorption of the nucleic acid on the surface. The interaction between small molecule compounds and the gold sensor chip surface is a point that needs investigation as research on this 58 S. Mazur, F. A. Tanious, D. Ding, A. Kumar, D. W. Boykin, I. J. Simpson, S. Neidle, and W. D. Wilson, J. Mol. Biol. 300, 321 (2000). 59 p. j. Bates, H. S. Dosanjh, S. Kumar, T. C. Jenkins, C. A. Laughton, and S. Neidle, Nucleic Acids Res. 23, 3627 (1995). 6o G. Bischoff, R. Bischoff, E. Birch-Hirschfeld, U. Gromann, S. Lindau, W.-V. Meister, S. Bambirra, C. Bohley, and S. Hoffman, J. Biomol. Struct. Dynam. 16, 187 (1998). 61 B. Persson, K. Stenhag, E Nilsson, A. Larsson, M. Uhlen, and E-A. Nygren, Anal. Biochem. 246, 34 (1997). 62 C. Rutigliano, N. Bianchi, M. Tomassetti, L. Pippo, C. Mischiati, G. Feriotto, and R. Gambari, Int. J. Oncol. 12, 337 (1998). 63 T. M. Nair, D. G. Myszka, and D. R. Davis, Nucleic Acids Res. 28, 1935 (2000). 64 C. I. Webster, M. A. Cooper, L. C. Packman, D. H. Williams, and J. C. Gray, Nucleic Acids Res. 28, 1618 (2000~[. 65 T. M. Heme and M. J. Tarlov, J. Am. Chem. Soc. 119, 8916 (1997).

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type of immobilization proceeds. We are currently working on a new method for covalently immobilizing nucleic acid onto a modified sensor chip that will allow for regeneration of the original surface after the chip has lost activity or when it is desirable to immobilize a different nucleic acid sequence. In the meantime, however, immobilization through the biotin-streptavidin affinity complex is the recommended strategy to efficiently and reproducibly immobilize nucleic acids for SPR experiments. The next question that must be addressed in sensor chip surface preparation is how much nucleic acid to immobilize. For kinetic experiments it is usually best to immobilize the smallest amount of nucleic acid (or any ligand) possible, while maintaining the desired signal-to-noise ratio, in order to minimize mass transport effects. Mass transport of analyte to the surface will alter kinetic data when the rate of mass transport is slower than or on the same time scale as the interaction kinetics. 53 Because a high concentration of surface binding sites depletes the analyte at the surface, the more nucleic acid immobilized the greater the contribution from mass transport. Crouch et al. suggest an immobilization level less than 100 RU for kinetic analysis of RNA-protein interactions. 66 When the analyte is a small molecule, however, it becomes necessary to increase the ligand surface density because the instrument response from small molecule binding will be much less than the response from protein binding. We typically immobilize about 250 RU of hairpin nucleic acid ('v50 bases in length) for small molecule assays. In our experience, the increase in surface density does not affect kinetic data because small molecules diffuse more rapidly than macromolecules and are not as limited by mass transport as macromolecules. Our small molecule nucleic acid experiments to date have not been limited by mass transport. It is important, however, to account for mass transport when it is present, to correctly analyze kinetic data. See Refs. 67-69 for a detailed description of processing kinetic data influenced by mass transport and additional suggestions for minimizing its effects. Because mass transport does not affect steady state data, it is advantageous for signal-to-noise optimization to immobilize a higher level of nucleic acid for equilibrium analyses, especially when working with small molecules. Experimental Protocols Preparation o f Sensor Chip. For immobilizing biotin-RNA (or any biotinnucleic acid) (see Preparation of RNA, below) on a sensor chip, the sensor chip

66R. J. Crouch, M. Wakasa, and M. Haruki, in "RNA-ProteinInteractionProtocols"(S. Haynes, ed.), pp. 143-160. Totowa, New Jersey, 1999. 67B. Goldstein, D. Coombs,X. He, A. R. Pineda, and C. Wofsy,J. Mol. Recognit. 12, 293 (1999). 68D. G. Myszka, Curt: Opin. Bioteehnol. 8, 50 (1997). 69T. A. Morton and D. G. Myszka,Methods Enzymol. 295, 268 (1998).

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must be modified to a streptavidin surface. BIAcore offers high-quality premade streptavidin sensor chips (SA sensor chip) that are ready for immediate use. However, it is possible and in some cases worthwhile to prepare streptavidin sensor chips (Refs. 50 and 53, and unpublished results, 2000) using standard (CM5) dextran surfaces or pioneer chips (chips that are under development but are available from BIAcore) with features such as a low-density carboxyl surface. The low-density carboxyl surfaces uses dextran but has less negative charge and so may be advantageous when investigating the interactions between RNA and molecules with high positive charge. We use the procedure outlined below for immobilizing streptavidin on CM5 sensor chips. The procedure is straightforward, and in our experience, generates streptavidin chips of similar quality to the premade chips available from BIAcore but with lower cost and additional flexibility. REQUIRED MATERIALS AND SOLUTIONS.The following is adapted from Ref. 55. CM5 or Pioneer sensor chip HEPES-buffered saline (HBS) buffer: 10 mM HEPES (pH 7.0), 150 mM NaC1, 3 mM EDTA, 0.005% (v/v) polysorbate 20 (running buffer) N-Hydroxysuccinimide (NHS): 100 mM in water N-Ethyl-N'-(dimethylaminopropyl)carbodiimide (EDC): 400 mM in water Acetate buffer (pH --~ 5): 10 mM (immobilization buffer) Streptavidin, 200-400 #g/ml [Pierce (Rockford, IL); Molecular Probes (Engine, OR); or Sigma (St. Louis, MO)] in immobilization buffer Ethanolamine hydrochloride: 1 M in water (pH 8.5) (deactivation solution) Sodium dodecyl sulfate (SDS): 0.05% (w/v) IMPORTANTNOTES 1. The sensor chip and solutions should equilibrate to room temperature for at least 30 min prior to use. 2. Because of the small diameter of the tubes in the microfluidics, all buffers should be filtered (0.22-#m pore size filter) and degassed daily prior to use: 3. The "amine coupling kit" available from BIAcore contains the NHS, EDC, and ethanolamine hydrochloride. The kit reagents do not need to be filtered or degassed. These reagents are also available from other suppliers but must be of sufficient quality so as not to clog the microfluidics, and they must be filtered and degassed. 4. For efficient immobilization of streptavidin (or any protein), it is necessary to use an immobilization buffer with a pH below the pl of the protein. If the macromolecule has a negative charge during immobilization, the amount of

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material that can be linked to the surface will be limited. Different sources and batches of streptavidin can vary in pl (pl 5-6), resulting in the necessity to optimize the immobilization procedure for each new batch (see Preconcentration of Streptavidin, below). BIAcore reports the use of 200 #g/ml in 10 mM acetate, pH 4.5, for streptavidin from Sigma, and 400 #g/ml in 10 mM acetate, pH 5.0, for streptavidin from Molecular Probes. We currently use streptavidin at 200 #g/ml in 10 mM acetate, pH 4.5, for streptavidin from Pierce. 5. BIAcore Software requires specification of volumes rather than times when performing injections. Therefore, with a flow rate of 5 #l/min, a 2-min injection must be specified as 10 #1. PRECONCENTRAT1ON OF STREPTAVID1N. Efficient immobilization of proteins through amine coupling on a carboxymethylated dextran surface involves electrostatic attraction between the negative charges on the surface matrix (carboxymethyl dextran) and positive charges on protein when it is below its pl (termed preconcentration). Ideally, the solution should have low ionic strength (50 mM or less) to maximize preconcentration. 55 It is important to note that if the pH of the buffer is too low (pH < 3-3.5), the dextran matrix becomes protonated and preconcentration is less efficient. 55 In addition, amine coupling requires uncharged amino groups and is therefore favored by higher pH. Clearly, the optimum pH is a compromise between efficient preconcentration and efficient coupling. To optimize the immobilization of streptavidin through amine coupling, injections of streptavidin in buffers differing in pH should be done as outlined below. Three buffers at pH 4.5, 5.0, and 5.5 are generally sufficient. We currently use 10 mM acetate buffer although any buffer lacking primary amine groups should suffice (note that a Tris buffer is not suitable for amine coupling). PRECONCENTRATIONPROTOCOL. The following is adapted from Ref. 55.

1. Dock a CM5 or Pioneer sensor chip into the instrument. 2. Prime the instrument three times with HBS running buffer. 3. Start a sensorgram at 5 #l/min. 4. Perform a 2-min "QUICK INJECT" of each streptavidin-pH combination from'high to low pH. Begin with a streptavidin concentration of 200 #g/ml. 5. Perform a l-min "QUICK INJECT" with "Extra Clean" consisting of 0.05% (w/v) SDS to remove any residual streptavidin from the fluidics. 6. Perform a 1-min "QUICK INJECT" with "Extra Clean" consisting of HBS buffer to remove any residual SDS from the fluidics. 7. Using the crosshair, note the preconcentration level in RU. BIAcore suggests that a streptavidin immobilization level of ~5000 RU is suitable for most applications. 55 We currently immobilize 2500-3500 RU of streptavidin. We have found this immobilization level suitable for our applications.

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Typically, 60-80% of the protein preconcentration level is immobilized. Therefore, aim for an electrostatic preconcentration level 150% of the desired immobilization yield. 55 For example, to immobilize 3000 RU of streptavidin, use conditions that favor an electrostatic preconcentration level of ~4500 RU. Using the data from the preconcentration experiment above, determine which pH is best for immobilizing the desired amount of streptavidin, keeping in mind that a lower pH may also reduce coupling efficiency. If none of the solutions produces the desired response, it may be necessary to increase the concentration of streptavidin [see Important Notes (4) above]. IMMOBILIZATIONOF STREPTAVIDIN. After choosing the appropriate pH to immobilize the desired amount of streptavidin, follow the procedure outlined below for immobilizing streptavidin. 1. Run a sensorgram at 5 #l/min. 2. With NHS in one vial and EDC in other, use the DILUTE command to make a 1 : 1 mixture of NHS-EDC (note that the NHS-EDC solution must be prepared fresh immediately prior to use). 3. Inject NHS-EDC for 7 min (35 #1) to activate the carboxymethyl surface to reactive esters. Higher or lower activation may be desired for preparation of nonstandard surfaces. 4. Inject streptavidin in the appropriate buffer for 7 min (35 #1) to immobilize streptavidin. 5. Inject ethanolamine hydrochloride for 7 min (35 /~1) to deactivate any remaining reactive esters. If immobilization fails after achieving appropriate levels of preconcentration then it may be necessary to increase the time of activation (the volume of NHS-EDS injected) or the streptavidin concentration if the pH is relatively low. Alternatively, EDC is quite labile and if the EDC solution is not freshly made or has not been kept properly frozen it could contain extensive inactive material. We keep EDC as a dry powder in a desiceant bottle at 4 ° and make fresh solutions as needed rather than aliquot all the EDC as suggested in the BIAcore amine coupling kit. We have found that streptavidin immobilization is straightforward as long as fresh solutions are used. Preparation of RNA: Derivatization to Incorporate Biotin. The biotin-streptavidin capture method requires that the RNA be modified with biotin. The ideal level of biotinylation for BIAcore nucleic acid applications is one biotin per nucleic acid. Small RNAs can be synthetically prepared with biotin on either the 5' or 3' terminus and are commercially available. Larger RNAs must be prepared by in vitro transcription. An excellent source for the synthesis of RNA by in vitro transcription is Wyatt et al. 7° The use of guanosine 5'-monophosphorothioate (GMPS) to prime a transcription reaction is a convenient way to site-specifically

34

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incorporate a single 5' terminal thiol group into RNA. 50'70-72 The thiol can then be modified to biotin with biotin-maleimide or biotin-iodoacetamide conjugates (Molecular Probes). A concise procedure for synthesizing GMPS from guanosine can be found in Burgin and Pace. 72 Alternatively, larger RNAs can be transcribed so as to incorporate 4-thiouridine internally, 73 and then modified with biotin as described above. Drawbacks to this method are that (1) additional experiments must be performed to ensure the internal biotin moiety does not disrupt native structure or function, and (2) it is difficult to achieve a low level of biotinylation unless only a couple of uridines are present in the RNA. RNA can also be sitespecifically modified to contain a thiol group on the 5' terminus through a kinase reaction. 7t Briefly, RNA is prepared by in vitro transcription, using GMP or GTP as the priming nucleotide instead of GMPS, or obtained commercially unmodified. The Y-phosphate(s) are then removed with phosphatase followed by transfer of the thiophosphate of ATPvS to the RNA by T4 polynucleotide kinase. Other methods to incorporate biotin into RNA include oxidation of the 3' terminal cis-diol to dialdehyde by periodate followed by reaction with biotin-hydrazine conjugates (Molecular Probes). Alternatively, the Y-phosphate of RNA (or DNA) can be converted to a 5'-phosphorimidazolide with imidazole and a water-soluble carbodiimide, and subsequently reacted with biotin-amine derivatives (Molecular Probes). Detailed protocols as well as limitations and necessary considerations for all of the above-described methods can be found in Qin and Pyle 71 (that article focuses on modification of RNA with fluorophores, but biotin derivatives can be used instead). It should be noted that regardless of the method used to modify RNA, excess biotin must be efficiently removed after modification to avoid competition with biotin-RNA for streptavidin binding sites and to enable reproducible levels of immobilization. Gel filtration on a desalting column and dialysis are simple methods to separate excess biotin from biotin-RNA. 55 Immobilization o f RNA. Once derivatized with biotin, the RNA is ready to be immobilized on a streptavidin-coated sensor chip. Generally, immobilization methods that rely on rapid kinetics and high-affinity binding of the ligand to the surface, such as the streptavidin-biotin interaction, do not require preconcentration experiments to assist in immobilization. 55 To date, we have worked with relatively short oligonucleotide hairpins (< 50 bases). For this size nucleic acid, we routinely use HBS as the immobilization and running buffer. Larger RNAs may require either higher salt to minimize electrostatic repulsion or a pH lower than the pl of streptavidin to favor electrostatic interactions with the negatively charged RNA, We routinely use a concentration of ~25 nM oligonucleotide (1 : 1 biotin : RNA 7oj. R. Wyatt, M. Chastain, and J. D. Puglisi, Biotechniques 11, 764 (1991). 71 p. Z. Qin and A. M. Pyle, Methods 18, 60 (1999). 72A. B. Burgin and N. R. Pace, EMBO,I. 9, 4111 (1990). 73j. E Milligan and O. C. Uhlenbeck, Methods Enzymol, 180, 51 (1989).

[21

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ratio) in HBS buffer when immobilizing nucleic acids less than 50 bases in length. It may be necessary to increase the concentration when using larger nucleic acids. A concentration that is too high, however, will make control over the amount of ligand immobilized difficult. In addition, too high a concentration may cause the outer dextran layer to bind RNA rapidly, potentially blocking inner layers and limiting the amount of RNA that can be immobilized. As discussed in the introduction, we typically immobilize about 250 RU of nucleic acid for small molecule studies. REQUIRED MATERIALSAND SOLUTIONS Streptavidin-coated sensor chip (SA chip or prepared as outlined above). HBS buffer: 10 mM HEPES (pH 7.0), 150 mM NaC1, 3 mM EDTA, 0.005% (v/v) polysorbate 20 (running buffer) Activation buffer: 1 M NaC1, 50 mM NaOH Biotin-modified RNA: ~25 nM in appropriate running buffer Biotin (optional) IMPORTANTNOTES 1. RNA is extremely sensitive to nucleases and base hydrolysis. Many researchers strictly use diethyl pyrocarbonate (DEPC)-treated water when making solutions for RNA research. We have found water from commercial systems such as NANO (Barnstead, Inc., Dubuque, Iowa) or Milli-Q (Pharmacia, Peapack, NJ) of sufficient quality for RNA work. 2. Because of the sensitivity of RNA, it is critical to maintain a "nuclease- free" working environment. It is always necessary to run the BIAcore Desorb method just prior to working with RNA, especially if there are multiple users on the instrument. In addition, any external instrument components such as the needle and the tubing placed in the running buffer should be routinely treated with RNase Zap (Ambion, Austin, TX) or another chemical to inactivate/remove any ribonucleases. 3. Researchers at the National Institutes of Health (NIH, Bethesda, MD) routinely cleanse the fluidics of the BIAcore instrument by injecting 20 #1 of RNase Zap solution through each flow cell at 20 #l/min followed by ten 20-#1 injections of DEPC-water (Ref. 66 and Inna Gorshkova, personal communication, 2000). It may be advisable to run this procedure when there are multiple instrument users, if a ribonuclease has ever been used in an experiment, or if problems with RNA degradation during an experiment are encountered. IMMOBILIZATIONPROTOCOL. The following is adapted from Ref. 55. 1. Dock the streptavidin-coated sensor chip into the instrument. 2. Prime the instrument three times with HBS buffer (note that priming does not need to be done if the streptavidin chip was just made as outlined above,

36

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has not been removed from the instrument and HBS has been the running buffer). 3. Start a sensorgram with a 20-/zl/min flow rate. 4. Inject activation buffer for 1 min (20 ~1) three times to remove any unbound streptavidin from the sensor chip. 5. Allow buffer to flow for at least 5 min before immobilizing the RNA. Note that NaOH will facilitate hydrolysis of the RNA, so it is critical that buffer flows long enough to remove any trace of NaOH. 6. Select the desired flow cell on which to immobilize the RNA and start a new sensorgram for only that flow cell. Take care not to immobilize RNA on the flow cell chosen as the control flow cell. Generally flow cell 1 ("fcl") is used as a control and RNA is not immobilized on it. It is often desirable to immobilize a different RNA (perhaps wild-type and mutants) on different flow cells. In this situation it is necessary to immobilize ligands one at a time. Alternatively, it may be desirable, depending on specific needs, to immobilize different amounts of the same ligand on each flow cell. 7. Using M A N U A L INJECT with a flow rate of 2 #l/rain, load the loop with ~ 1 0 0 #1 of RNA and inject over the desired flow cell(s). Using the crosshair, track the number of resonance units immobilized and stop the injection after the desired level is reached. As a rule of thumb, do not inject more than 80-90% of the loaded volume, to minimize sample dispersion effects at the end of the injected sample. If necessary, reload the loop and inject more RNA. It may be necessary to alter immobilization conditions such as ionic strength, pH, and ligand concentration to obtain the desired immobilization level. 8. At the end of the injection and after the baseline has stabilized, use the crosshair to determine the resonance units of RNA immobilized and record this amount. The amount of RNA immobilized is required to determine the theoretical number of small molecule binding sites for the flow cell. 9. Repeat steps 7 and 8 for each flow celI-RNA combination according to experimental design. i0. Some researchers block the remaining biotin binding sites and the biotin binding sites on the streptavidin in the control flow cell with free biotin. This will depend on any nonspecific interactions between components in the flow solution with streptavidin on the sensor chip surface. STORAGEOF RNA SENSOR CHIP. Researchers at the NIH store oligonucleotide sensor chips in Tris-buffered saline (TBS) buffer to maintain binding activity (Inna Gorshkova, personal communication, 2000). To do so, remove the sensor chip from the white cartridge, being careful not to touch the flow cells. Immerse and store the chip in RNase-free TBS buffer. To subsequently use the sensor chip, remove it from the TBS. Shake the sensor chip a couple of times to remove the excess buffer, and then leave the chip to air dry in a sterile environment until it is completely

[2]

RNA INTERACTIONS

37

dry. Do not touch or wipe the flow cells in any way. Insert the sensor chip into the white cassette and dock. If storing sensor chips dry, it may be advisable to run a solution of RNase inhibitors over the sensor chip prior to undocking, Data Collection and Analysis We routinely obtain high-quality binding data on a BIAcore 2000 instrument, even with only 10% site saturation of small molecules (MW 300-600) with only a few hundred resonance units of immobilized nucleic acid (Fig. 1). For binding constants of 107-108 M -l, as observed with many RNA complexes (our unpublished results, 2000), small molecule concentrations from 1 nM to 1 # M in the flow solution allow accurate determination of binding constants. Methods to extract binding constants from experimental data have been described in detail. TM In addition to instrument performance, data collection and data processing influence the quality of the data and the information that can be extracted from it. Myszka published a comprehensive review outlining the steps necessary to obtain high-quality biosensor data. 75 The article discusses many of the potential pitfalls of biosensors and provides suggestions for avoiding them. The discussion below focuses on application of those principles to nucleic acid-small molecule complexes. Data Collection

Some of the issues affecting the quality and accuracy of the data obtained through SPR were addressed above. For example, the advantages of working with a homogeneous surface with as low a surface density as possible have been discussed. In addition, one flow cell should be left blank (no RNA immobilized) to provide a control flow cell. Some researchers immobilize biotin or an unstructured RNA on the control flow cell to match the control and RNA flow cells as closely as possible. However, with most compounds in our laboratory, we have not found significant adsorption on the dextran or streptavidin surface. In addition, adding a different RNA to the control surface could introduce unknown sources of nonspecific binding in the blank subtraction. Additional issues such as the mode and length of sample injection, sample concentration range and order of injection, regeneration and buffer injections, and the integrity of the surface are addressed when setting up an experiment. Surface interactions can be a serious problem and must be evaluated for each compound series to be investigated. There are several injection modes available in the BIAcore 2000 and 3000 instruments for delivering sample to the sensor chip. The injection modes differ in the amount of volume that can be injected. More importantly, they differ in

74j. j. Correiaand J. B. Chaires,Methods Enzymol. 240, 593 (1994). 75D. G. Myszka,J. Mol. Recognit. 12, 279 (1999).

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[2]

the amount of sample dispersion that occurs during transport and injection. It is generally best to use the injection mode "KINJECT" for sample injections. This mode offers the maximum number of air segments surrounding the flowing sample, minimizing the amount of sample dispersion and dilution with buffer. Myszka points out that high flow rates are advantageous since faster switching between running buffer and sample helps to deliver a consistent sample plug. 75 In addition, high flow rates help to minimize rebinding of analyte to ligand during the dissociation phase when the association rate is fast. Higher flow rates are not necessary in steady state experiments, and may not be feasible if the kinetics are slow and sample is limited because faster flow rates require more sample. Sample concentrations should vary over a wide range (at least 100-fold). In general, the middle of the sample concentration range should be near 1/Ka if the Ka can be approximated. If the Ka is unknown, a broad concentration range should be used in a preliminary experiment to obtain an estimate of the K a. Ideally, the order of sample injection should be randomized. Many of the small molecules we have worked with, however, adsorb nonspecifically to the tubing of the microfluidics and are slowly released over the course of the experiment. In this situation, we have found that injecting samples from low to high concentration is necessary for eliminating artifacts in the data from adsorption carryover. It is also important to replicate each injection at least once. At the end of the injection, a regeneration step should be performed to remove any analyte still bound to the surface. Many of our compounds dissociate rapidly enough to regenerate the surface with buffer flow alone and do not require a separate regeneration step. When working with RNA, each solution and injection increases the potential for degradation so the least number of steps the better, as long as the quality of the data is not compromised. When a regeneration step is required, we typically use 1 M NaCI. We have used 1 M NaCI under slightly acidic conditions but found that salt alone was sufficient to dissociate most cationic compounds from RNA. To subsequently remove any remaining NaCI, we perform two 1-min injections of running buffer prior to the end of the cycle followed by a 5-min wait with running buffer flowing. After the next cycle has begun, we have the instrument wait another 5 min with running buffer flowing to ensure baseline has stabilized before the next sample injection. When working with small molecules that often give changes of less than 2 RU at steady state, it is essential that the baseline not drift significantly during the injection. We typically achieve a drift less than 0.5 RU/min by putting in the 5-rain wait (instrument specification is 1 RU/min), and such attention to the baseline is essential in small molecule binding experiments. For our experiments, we identify a well-behaved and well-characterized analyte as a standard to periodically evaluate the integrity of the surface when working with both RNA and DNA. In some of our experiments the RRE hairpin was the immobilized RNA. Because the Rev-RRE complex is a well-behaved 1 : 1 complex under appropriate conditions and has previously been studied by SPR, 5° we used

[2]

RNA INTERACTIONS

39

the Rev peptide to periodically check the amount and integrity of the RRE immobilized. If a well-described analyte is not available, any well-behaved molecule that binds to the immobilized RNA can be injected after RNA immobilization and calibrated for use as a standard for that surface. The same concentration of the standard is analyzed periodically and the resonance unit response compared with that of the initial injection. By using a calibrator molecule, the amount of RNA on the chip can be routinely checked and any loss can be accounted for during data processing. We have found a calibrator essential when working with the same RNA sensor chip for extended periods of time, especially if the chip has been stored for any length of time. If there are questions about the chip, a more thorough evaluation of the surface, including a complete concentration gradient experiment with a standard, should be performed to compare kinetic rates and immobilization levels with those obtained after the RNA was first immobilized. Below is a method that we routinely use to collect small molecule data on nucleic acid surfaces. This method is set for a flow rate of 10 #l/min (FLOW 10) over flow cells 1,2, and 3 (FLOWPATH 1,2,3). The samples are injected as written (STEP) from low to high concentration and then the injection sequence is repeated (TIMES 2). Note that before any drug is injected, 10- and 5-min buffer injections are done to enable double referencing (see Data Analysis, below). 75 In addition, the volume of drug injected is set as a variable so that the least amount of volume required to reach a steady state is used for each concentration. Much less time is required for the association reactions at high concentration of analyte.

Typical Method MAIN RACK 1 thermo_c RACK 2 thermo_a FLOWCELL 1,2,3,4 LOOP DB340 STEP APROG drug %sample2%position2%volume2%conc2 ENDLOOP APPEND Continue END DEFINE APROG drug PARAM %sample2%position2%volume2 %conc2 KEYWORD Concentration %Conc2 CAPTION %conc2%sample2 over RRE (gradient surface) F L O W 10 FLOWPATI-I 1,2,3

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WAIT 5:00 KINJECT %position2 %volume2 300 QUICKINJECT r2f4 20 ! 1M NaCI EXTRACLEAN QUICKINJECT r2f3 10 !Buffer EXTRACLEAN QUICKINJECT r2f3 10 !Buffer EXTRACLEAN WAIT 5:00 END DEFINE LOOP DB340 LPARAM %sample2%position2%volume2 %conc2 TIMES 2 Buffer Buffer DB340 DB340 DB 340 DB340 DB340 DB340 DB340 DB340 DB340 DB340 DB340 DB340 DB340 DB340 DB340

r2f3 r2f3 r2al r2a2 r2a3 r2a4 r2a5 r2a6 r2a7 r2a8 r2a9 r2al0 r2bl r2b2 r2b3 r2b4 r2b5

100 50 100 100 100 100 50 50 50 50 50 50 50 50 50 50 50

0.000u 0.000u 0.025u 0.075u 0.125u 0.250u 0.375u 0.500u 0.625u 0.750u 1.000u 1.750u 2.500u 3.250u 5.000u 6.750u 10.00u

END

Data Processing After the data have been collected, there are several processing steps that must be carried out before any quantitative information can be extracted. We use BIAevaluation 3.0 software for data processing. Zeroing on the y axis (RU) and then the x axis (time) are the first steps in data processing. Because the flow cell surfaces are not identical to each other, the refractive index of each surface is different,

[2]

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41

causing the flow cells to register at different positions on the y axis. Zeroing the data on the y axis is necessary to allow the responses of each flow cell to be compared. Generally the average of a stable time region of the sensorgram, prior to sample injection, should be selected and set to zero. Because the flow cells are aligned in series, sample is not injected across the flow cells simultaneously. Zeroing on the x axis aligns the beginnings of the injections with respect to each other. Myszka has shown that artifacts, such as spikes at the beginning and end of injections and baseline fluctuations, are usually present in all flow cells. 75 The two data processing steps outlined next serve to minimize these artifacts and also to correct for the bulk shift that results when injection buffer and running buffer are not identical. In the first step, the control flow cell sensorgram is subtracted from the reaction flow cell sensorgrams. This removes the bulk shift contribution to the change in resonance units. The last step in data processing is required to remove systematic deviations that are frequently seen in the sensorgrams. Referred to as "double referencing," this process removes the systematic drifts and shifts in baseline that are frequently observed even in control cell-subtracted sensorgrams. In the data collection method shown above, two buffer injections are performed, one for each volume amount used for sample injection. In double referencing, plots are made for each flow cell separately, overlaying the control cell-corrected sensorgrams from buffer and all sample injections. The buffer sensorgram is then subtracted from the sample sensorgrams. If multiple buffer injections were done, averaging these sensorgrams before subtracting from the sample sensorgram can improve the data further. At this point, the data should be of optimum quality and are ready to be analyzed.

Data Analysis In most systems, both equilibrium and kinetic rate constants can be extracted from SPR data. The equilibrium constant can be obtained from fitting steady state data directly, or from kinetic data. The association equilibrium constant (K) is the ratio of the association (ka) and dissociation rate (kj) constants. Comparing the K value obtained both ways is a good way to evaluate the models used to fit the data. Most cationic molecules that bind RNA also interact nonspecifically with the RNA, but frequently both K, ka, and ka for the strong specific site can be determined accurately because the equilibrium constants for the two types of binding differ markedly. Having both kinetic and equilibrium information can yield important insight into understanding whether any biological activity of the small molecule is governed by the kinetic or equilibrium (or both) constants of binding to RNA. Knowledge of the stoichiometry of the system is essential for obtaining correct kinetic and binding constants as well as for obtaining a complete description of the system being studied. While at first glance determining stoichiometry seems straightforward, this is often not the case when working with nucleic acid-cation

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TABLE II REFRACTIVE INDEX INCREMENTS AND REFRACTIVE INDEX INCREMENT RATIOS WITH B S A AND d A d T

RI incrementa Compound BSA Neomycin Netropsin Quinacrine Berenil Pentamidine DB404 dGdC dAdT

RI increment ratio vs BSAh

RI increment ratio vs dGdCh

590.5nm 633nm 679.5nm 590.5nm 633nm 679.5nm 590.5nm 633nm 0.18 0.15 0.22 0.24 0.35 0.22 0.28 0.22 0.21

0.17 0.15 0.21 0.25 0.34 0.22 0.28 0.22 0.21

0.17 0.15 0.21 0.24 0.33 0.22 0.27 0.22 0.21

-0.9 1.2 1.4 2.0 1.3 1.6 1.2 1.2

-0.9 1.2 1.4 2.0 1.3 1.6 1.3 1.2

-0.9 1.2 1.4 1.9 1.3 1.6 1.3 1.2

0.8 0.7 1.0 1.2 1.7 1.1 1.3 1.0 --

0.8 0.7 1.0 1.2 1.6 1.1 1.3 1.1 --

679.5 nm 0.8 0.7 1.0 1.2 1.6 I. 1 1.3 1.1 --

RI increment (RII) = A(An)/~xC; determined at 590.5,633, and 679.5 nm. RI increment ratio = RllcompoundI/Rllcompound2 = (~n/~C)compound l/(6n/~C)compound 2. interactions because nonspecific b i n d i n g will always be present. The refractive index increments (RIIs) of small molecules can vary considerably more than those of proteins because small molecules can contain a m u c h larger variety of potential constituent units. While it is recognized that the a m o u n t of analyte b o u n d and the molecular weight of the analyte contribute to the observed b i n d i n g response, the RII is often an overlooked contributor to the b i n d i n g response. W h e n studying m a c r o m o l e c u l a r interactions, such as p r o t e i n - p r o t e i n and p r o t e i n - n u c l e i c acid systems (the m a i n focus of biosensor studies until recently), the RIIs of the interacting species are in fact generally irrelevant because the RIIs of most proteins and nucleic acids are similar. We have found, however, that the RIIs of small molecules can vary by a factor of two, and can be smaller or significantly larger than the RIIs of proteins and nucleic acids (Table II). 76 Highly conjugated molecules have significantly larger RIIs than those of unconjugated molecules. Because the RIIs of small molecules can be different from those of proteins and nucleic acids, it is essential that such a difference be accounted for during data interpretation to correctly determine stoichiometry, and subsequently kinetic and equilibrium constants. The m a x i m u m BIAcore instrument response for a 1 : 1 b i n d i n g interaction can be predicted with the following equation: (RUpred)max = RUE \ M ~ L J

L~ - ]

where (RUpred)max is the predicted m a x i m u m instrument response in resonance units for b i n d i n g at a single site, RUE is the experimental a m o u n t of R N A 76 T. M. Davis and W. D. Wilson, Anal. Biochem. 284, 348 (2000).

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immobilized on the chip in resonance units, MWA is the molecular weight of the analyte, MWL is the molecular weight of the RNA, (On/OC)Ais the RII of the analyte, and (On/OC)Lis the RII of the RNA. The maximum observed instrument response [(RUobs)max] will approach this predicted value as the concentration of analyte bound at the surface increases and saturation of the single binding site occurs. For systems with analyte-to-ligand stoichiometry greater than 1, RU(pred)rnax must be multiplied by the number of binding sites per macromolecule to obtain the maximum response: (RUpred)sat = (RUpred)maxP

(2)

where (RUpred)sat is the predicted instrument response on saturation of p binding sites in resonance units, (RUpred)maxis from Eq. (1), and p is the number of binding sites per macromolecule. The maximum instrument response on saturation of all binding sites, (RUobs)sat, is obtained from fitting SPR results, but a value for (RUpred)max is essential in order to correctly determine stoichiometry from (RUobs)sat. Small molecules generally have more than a single binding site in nucleic acid complexes. Because nonspecific binding can occur with cationic molecules and RNA, the RII ratio is critical for accurate determination of stoichiometry. Consider, for example, typical parameters for the cationic small molecule DB404 binding to DNA: RUL of 250, MWA of 467, MWL of 6465, and (On/OC)A/(On/OC)Lratio (RII ratio) of 1.3 (Table II). Without correcting for the RII difference, these parameters would predict an RUma× per binding site of 18 [Eq. (1)]. However, accounting for an RII ratio of 1.3, the actual predicted RUmax per binding site is 24. For a dimer binding mode, which we have encountered with similar compounds, 51 an experimental (RUobs)sat near 50 could incorrectly be characterized as a trimer binding mode [predicted at 54, Eq. (2)] if the RII ratio is not considered. Likewise, for a trimer binding mode, (RUobs)sat would register around 72, which could be mistaken for four binding sites per RNA [predicted at 72, Eq. (2)] if not corrected for the difference in RIIs. Clearly, the RII ratio is a critical factor to consider when interpreting SPR data involving small molecules. It should be noted that because stoichiometry is an integral number [p in Eq. (2)], it should only be necessary to have an approximate value of the RII to determine p. Available data may serve as a guide for estimating the RIIs of other small molecules if such data cannot be experimentally determined. 76

Equilibrium Analysis Although our laboratory uses the SPR biosensor to obtain kinetic constants and screen small combinatorial libraries, in many applications the binding constants of small molecule-nucleic acid systems are the primary requirement. Many of our compounds bind specifically to multiple sites on the nucleic acid, especially our

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small m o l e c u l e - R N A systems, and also display various degrees of nonspecific binding. To analyze equilibrium data with BIAevaluation software, the data should be treated in the following manner. After processing the data as outlined above, overlay the sensorgrams of all the analyte concentrations for the flow cell(s) with immobilized RNA (Fig. 2A). Using the software, calculate an average of the blankcorrected data in the steady state region of each sensorgram and then plot the average as a function of analyte concentration (Fig. 2B). The result is a recognizable binding isotherm. The simplest model system for determination of a binding constant from such results is one with a stoichiometric ratio of one binding site per

50

....

, ....

, ....

, ....

, ....

, ....

, ....

, ....

A

4o

3O

mr 20

10

0

-10

.... -50

' .... 0

~ .... 50 Time

' .... 100

' .... 150

' . . . . . . . . 200 250

' .... 300

350

(second)

FIG. 2. (A) Sensorgramsat different concentrationsof the small molecule DB244 binding to DNA. The data have been processed as described in text. (B) The average response at steady state for the curves shown in (A), plotted as a function of concentration.The result is a typical binding isotherm.

[2]

RNA INTERACTIONS 30

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45 .

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25

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1x10" 6

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3xlO" 6

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FIG.2. (Continued) RNA that can be fit with Eq. (3), using either the BIAevalution software or any suitable program, such as KaleidaGraph, for this purpose: KA[AI ] RUss = RUmax (1 ~K~A[AI)J

(3)

where RUss is the RU response at steady state, KA is the dissociation constant, RUmax is the fitted value for the maximum binding capacity of the surface in resonance units, and [A] is the concentration of analyte in the flow solution. Note that the free analyte concentration in equilibrium with bound analyte {[A] in Eq. (3)} is constant in the flow solution. The dissociation constant, Ko, is 1]KA. Note that in the ideal case, RUmax equals (RUpred)maxand (RUss)max. In some cases

46

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[2]

it may not be possible to go to a high enough analyte concentration to reach the (RUss)max plateau. Also, if the surface has significantly degraded, RUmax will be less than Rg(pred)max. As mentioned above, most small cation-nucleic acid complexes have stoichiometry greater than 1. Many of our systems involve specific binding at one or two sites followed by additional nonspecific binding at higher concentration. For these systems, we typically visualize the data in the form of both direct and Scatchard plots while fitting by nonlinear least squares, using Eq. (3) or a more complex model. After obtaining the average of the data in the steady state region as described above, we plot the data as either R U J [ A ] as a function of RU~s or r/[A] as a function of r, where r is RWss/RW(pred)max.Plotting the data in Scatchard form can visually reveal considerable information about the binding constants, stoichiometry of specific and nonspecific binding, and cooperativity.51 Figure 3A provides an excellent example of complex binding information obtained on a BIAcore 2000 and presented in a Scatchard plot. In this example, the differences in binding constants, stoichiometry, and cooperativity for binding of two low molecular weight aromatic cations, DB293 and DB270, to two different DNA hairpins, oligo 1 and oligo 2-1, illustrate the power of SPR biosensors. The cooperative binding of two molecules of DB293 to oligo 2-1 is clear. This type of information is difficult to obtain by other methods. Equilibrium constants were obtained by fitting the Scatchard plots with either a single-site model [Eq. (4) with K2 = 0] or with the two-site model in Eq. (4): r =

KI[A] + 2K1K2[A] 2 1 + KI[A] + 2K1K2[A] 2

(4)

where r represents the moles of bound compound per mole of DNA hairpin and is calculated as described above, Kl and Kz are macroscopic association binding constants, and [A] is the concentration of analyte in the flow solution. Figure 3B provides another example of SPR data presented in the form of a Scatchard plot. The data are for binding of a small aromatic tetracation to an RNA hairpin. The data indicate a stoichiometry of two specific binding sites per RNA hairpin, but additional nonspecific binding occurs as the concentration increases. The binding constant for nonspecific binding is substantially lower that the binding constants for the two specific binding events. Higher salt concentrations help to reduce the amount of nonspecific binding.

Kinetic Analysis Because binding data is collected in real time with SPR biosensors, the technique is particularly well suited for kinetic analyses. Currently, association constants in the range of 103-106 M-1 sec-I and dissociation constants in the range of 10-5-10 -1 sec -1 can be accurately obtained on BIAcore instruments. These

[2]

RNA INTERACTIONS

47

ranges can be extended, however, with proper data handling, favorable S/N, and other conditions. One of the most important parameters to consider during kinetic analysis of SPR data is mass transport, which is transport of the analyte to the solid phase covalently bound with ligand. Although it is best to optimize data collection parameters so as to remove any contribution to the kinetics from mass transport, accurate kinetic constants can still be obtained when mass transport contributions

2 10 7

A

1.5 10 7

oo

~-.~"1 10 7

5 10 6

O

O

o

0

0.5

1

1.5

2

r FIG. 3. Examples of SPR data visualized by Scatchard plots. (A) Scatchard plot for binding of the small molecules DB293 and DB270 to two different DNA olignucleotides along with best fit binding curves are shown. (A, A) DB293 and DB270 binding to oligo 1; (O, O) DB293 and DB270 binding to oligo 2-1. [These data are reprinted with permission from L. Wang, C. Bailly, A. Kumar, D. Ding, M. Bajic, D. W. Boykin, and W. D. Wilson, Proc. Natl. Acad. Sci. U.S.A. 97, 12 (2000).] (B) Scatchard plot for binding of the small molecule DB340 to RRE with the best fit for the two strong binding sites (our unpublished data, 2000).

48

BIOPHYSICAL APPROACHES

[21

8 x l 06

I

'

'

'

B 7 x l 06

6 x l 06

5 x l 06

0 4 x l 06 •

B

•

•

•

3 x l 06

2 x l 06

lx10 6

i° ' i

0

0.5

1

1.5

2

.

.

.

.

.

.

.

I

*

.

2.5

3

r

FIG,3. (Continued) cannot be entirely removed. 67-69 With small molecules, however, mass transport effects often do not exist. Indeed, we have never experienced mass transport limitations with any of the small molecules we have investigated to date. It is essential, however, to test for mass transport effects, by methods such as varying the flow rate, before proceeding with data collection and kinetic analysis. The best way to process kinetic data obtained with SPR biosensors is through global analysis, especially for complex systems. 77 Global parameters are constrained to the same value for each curve in the data set, in contrast to local 77T. A. Morton, D. G. Myszka, and I. M. Chaiken,Anal. Biochem. 227, 176 (1995).

[2]

RNA INTERACTIONS

49

parameters, which can have different values for each curve. Global analysis offers a more stringent test of reaction models and a statistically more reliable calculation of the parameters. 77 Currently, global analysis offers the most reliable fitting method for obtaining optimum fitted kinetic constants from SPR data. BIAevalution software versions 3.0 and above have the option to fit data globally. CLAMP is an excellent program for fitting SPR biosensor data that combines numerical integration and nonlinear global curve fitting routines. 69'78 The software is user-friendly and can be downloaded at http:/]www.hci.utah.edu]groups/interaction]clamp.htm. As with equilibrium analysis, the simplest system for fitting kinetic data is one with a stoichiometric ratio of one binding site per RNA. The classic 1 : 1 rate equation [Eq. (5)] is converted to SPR terms in Eq. (6). d[AB] dt dRU

dt

- ka[A][B] - kd[AB]

(5)

-- kaC(RUmax - RU) - kdRU

(6)

where [AB] is the concentration of complex and is directly related to the instrument response, resonance units in Eq. (6); [A] is the concentration of free analyte, which is the concentration of analyte in the flow, and is equal to the constant C in Eq. (6); [B] is the concentration of immobilized ligand that is not bound by analyte, and is equal to the maximum amount of B immobilized on the surface, RUmax, minus the amount of B bound with analyte, RU; ka is the association constant; and kd is the dissociation constant. For a 1 : 1 system with no mass transport, Eq. (6) should be used during fitting analyses once the data has been processed as described earlier. As mentioned above, most of our RNA-small molecule systems have stoichiometry greater than 1 and require more complex models for fitting. Complex models, however, usually have too many parameters for unique fits. Because SPR biosensors permit low analyte concentrations, it is often possible to isolate and fit only the strongest binding site. SPR biosensors can also be used to identify and rank small molecules that bind to an immobilized ligand, using data in the association and dissociation regions of a sensorgram, without actually determining the kinetic constants, s2 Once identified, a detailed kinetic analysis of the compounds that bind within the desired rate region can be determined in subsequent experiments. BIAsimulation software (BIAcore) is an excellent tool for generating simulated sensorgrams to evaluate the effect kinetic and experimental parameters have on the shape of a binding curve. For example, the program can be used to determine what amount of ligand to immobilize given a set of kinetic constants so as to minimize mass transport effects. The reader is urged to explore the applications of BIAsimulation during experimental design and data analysis. 78 D. G. Myszka and T. A. Morton, Trends Biochem. Sci. 23, 149 (1998).

50

BIOPHYSICALAPPROACHES

[2]

Thermodynamic Analysis The BIAcore 2000 biosensor has the ability to collect data at temperatures from 4 to 40 °. Dissociation and kinetic constants obtained at different temperatures can be further analyzed to obtain thermodynamic information. In a review, 79 Myszka and Morton present examples of calculating the Gibbs free energy change of binding, the enthalpy and entropy of binding through van't Hoff analysis, and the enthalpic and entropic contributions to the free energy of activation through Eyring analysis for a model protein-protein system from data collected on a BIAcore 2000 biosensor. We refer the reader to this review as a guide to performing such analyses. Chaires 8° has presented experimental considerations required to obtain accurate thermodynamic parameters through van't Hoff analyses. A powerful approach to obtaining complete thermodynamic profiles for cation-nucleic acid interactions is to determine the equilibrium constant (Gibbs energy) as a function of temperature by SPR methods, and to directly determine the reaction enthalpy as a function of temperature by calorimetery. 58 Conclusion Surface plasmon resonance biosensors are firmly established for monitoring macromolecular interactions and are becoming a popular technique for characterizing small molecule complexes. Only more recently, however, has the technology advanced to the point that small molecule systems can be studied without the need for a high molecular weight tag. SPR biosensors are amenable to the study of kinetics and equilibrium properties of many systems as well as screening of combinatorial libraries, and the potential applications of biosensors are rapidly expanding. SPR biosensors offer an excellent method for investigating small molecule-macromolecule systems because many small molecules are either not fluorescent or do not yield notable gel band shifts on binding to macromolecules. In addition, such small quantities are required that precious or sparse samples can be investigated with minimal material requirements. This is in direct contrast to UV or CD spectroscopies, where a 10- to 100-fold increase in sample quantity may be necessary to obtain a satisfactory signal-to-noise ratio. This review is intended to serve as a guide for beginning users of SPR biosensors. Other excellent reviews and articles have been noted and the reader is encouraged to explore them also. It is worth reemphasizing that the refractive index increments of small molecules must be considered for accurate analysis of SPR data for systems with stoichiometry greater than 1. The application of biosensors 79D. G. Myszka,MethodsEnzymol. 323, 325 (2000). 80j. B. Chaires,Biophys. Chem. 64, 15 (1997).

[3]

MEASUREMENT OF BINDING AND UNWINDING

51

to small molecule systems is so new that advances, pitfalls, and other important considerations in this area are likely to develop rapidly.

Acknowledgments Work from our laboratory described in this review has been supported by the NIH. We also acknowledge Farial Tanious for helpful suggestions and data for some figures and Lei Wang for data for some figures. We appreciate preprints from Dr. David Myszka.

[3] Simultaneous Measurement of Binding Constants

and Unwinding Angles by Gel Electrophoresis By STEVEN M. ZEMAN and DONALD M. CROTHERS

The perturbation of DNA topology due to ligand binding is a phenomenon of profound biological importance. Protein-induced DNA bending is an important, in many cases indispensable, factor in the regulation of gene expression, and several drugs utilized in the treatment of cancer derive their therapeutic efficacy from an ability to intercalate DNA, unwinding the double helix in the process. The methodology we describe pertains to the latter class of compounds, namely small organic molecules. It offers the advantage over related methods that D N A topological changes and drug binding constants are measured simultaneously and, perhaps most importantly, under identical reaction conditions. The original intercalation model 1describes a lengthening of the D N A helix and an associated unwinding of the phosphodiester backbone DNA on drug binding in order to accommodate a molecule of intercalator. As this unwinding varies with intercalator, the unwinding angle (¢) is a valuable biophysical parameter when evaluating the characteristics of a compound interacting with DNA. To date, most experimental approaches for measuring ¢ have exploited the special conformational characteristics of closed circular D N A (ccDNA). As isolated from prokaryotes and viruses, ccDNA is negatively supercoiled, that is to say, its writhe (Wr) and linking difference ( A L k ) are negative. *'2'3 The unwinding angle ~b is the change in I L. S. Lerman, J. Mol. Biol. 3, 18-30. * Lk = Tw + Wr, where Tw is the twist about the helical axis and Wr is the writhe or superhelicityof the helical axis through three-dimensionalspace. Whereas Tw and Wr and each geometric properties that vary with environmentalconditions such as temperature and ionic strength, the linking number Lk is a topologicalproperty that is invariantin the absenceof single-strandednicks. ALk is definedas the difference in linking number or superhelicalturns between an idealized fully relaxed, nonligated closed circular DNA for which Wr = 0, and a topoisomer j o f nonzero Wr. Whereas ALk can

METHODS IN ENZYMOLOGY, VOL. 340

Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. 0076-6879/00 $35.00

[3]

MEASUREMENT OF BINDING AND UNWINDING

51

to small molecule systems is so new that advances, pitfalls, and other important considerations in this area are likely to develop rapidly.

Acknowledgments Work from our laboratory described in this review has been supported by the NIH. We also acknowledge Farial Tanious for helpful suggestions and data for some figures and Lei Wang for data for some figures. We appreciate preprints from Dr. David Myszka.

[3] Simultaneous Measurement of Binding Constants

and Unwinding Angles by Gel Electrophoresis By STEVEN M. ZEMAN and DONALD M. CROTHERS

The perturbation of DNA topology due to ligand binding is a phenomenon of profound biological importance. Protein-induced DNA bending is an important, in many cases indispensable, factor in the regulation of gene expression, and several drugs utilized in the treatment of cancer derive their therapeutic efficacy from an ability to intercalate DNA, unwinding the double helix in the process. The methodology we describe pertains to the latter class of compounds, namely small organic molecules. It offers the advantage over related methods that D N A topological changes and drug binding constants are measured simultaneously and, perhaps most importantly, under identical reaction conditions. The original intercalation model 1describes a lengthening of the D N A helix and an associated unwinding of the phosphodiester backbone DNA on drug binding in order to accommodate a molecule of intercalator. As this unwinding varies with intercalator, the unwinding angle (¢) is a valuable biophysical parameter when evaluating the characteristics of a compound interacting with DNA. To date, most experimental approaches for measuring ¢ have exploited the special conformational characteristics of closed circular D N A (ccDNA). As isolated from prokaryotes and viruses, ccDNA is negatively supercoiled, that is to say, its writhe (Wr) and linking difference ( A L k ) are negative. *'2'3 The unwinding angle ~b is the change in I L. S. Lerman, J. Mol. Biol. 3, 18-30. * Lk = Tw + Wr, where Tw is the twist about the helical axis and Wr is the writhe or superhelicityof the helical axis through three-dimensionalspace. Whereas Tw and Wr and each geometric properties that vary with environmentalconditions such as temperature and ionic strength, the linking number Lk is a topologicalproperty that is invariantin the absenceof single-strandednicks. ALk is definedas the difference in linking number or superhelicalturns between an idealized fully relaxed, nonligated closed circular DNA for which Wr = 0, and a topoisomer j o f nonzero Wr. Whereas ALk can

METHODS IN ENZYMOLOGY, VOL. 340

Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. 0076-6879/00 $35.00

52

BIOPHYSICALAPPROACHES

[3]

twist plus writhe in a linear molecule on ligand binding; it is usually assumed that there is no net change in writhe, in which case q~ is equal to the number of degrees by which the DNA is unwound about its helical axis per ligand molecule bound. The topological properties of DNA can be altered by the enzyme topoisomerase I (Topo I), 4'5 whose discovery opened new experimental avenues for the determination of Lk and ~b.Topo I facilitates thermodynamic equilibration between supercoiled and relaxed forms of ccDNA by creating a transient single-strand nick at which superhelical strain can be relieved, as shown schematically in Fig. IA. Methods for the determination of ~b that employ Topo I and intercalating ligand are based on the processes shown in Fig. 1B-D. On intercalation by a small, usually aromatic molecule (Fig. 1, step a), naturally supercoiled ccDNA is unwound (ATw < 0), so that the writhe becomes less negative at constant Lk (Fig. 1B). For a certain critical amount of bound intercalator (Fig. 1C), all of the negative supercoiling is taken up by negative twist produced by unwinding at drug binding sites, writhe disappears, and the ccDNA exists as a relaxed circle in which the helical axis lies fiat in a plane. Further titration with intercalator past this topological equivalence point (Fig. 1D) continues to unwind the helix, introducing supercoils in an opposite sense, so that Wr is positive. When ccDNA that has been intercalated to some extent is acted on by Topo I (Fig. 1, step b), a fully relaxed species is generated whose ATw reflects the presence of bound intercalator. In this fully relaxed species, Wr = 0. When the intercalator is removed, usually by organic extraction, the resulting change in Tw is offset by compensatory Wr such that Lk remains unchanged. (Lk, the algebraic sum of Tw and Wr, is the number of times the two DNA strands wind about each other, and is an invariant property intrinsic to any nonnicked topoisomer of ccDNA.) The change in Wr on ligand removal manifests itself conformationally as a reappearance of superhelical turns, the magnitude of which is determined by the original amount of intercalator bound at the time of enzymatic religation. Specifically, the linking number deficit - A L k in the final state is equal to the twist change - A T w produced by intercalator binding in step a (Fig. 1).

assume a fractional value because of the idealized reference state, Lk is integral by definition. Excellent treatments of ccDNA topology exist in the literature. 2"3 2 A. D. Bates and A. Maxwell, in "DNA Topology" (D. Rickwood and D. Male, eds.), pp. 17-61. Oxford University Press, Oxford, 1993. 3 N. R. Cozzarelli, T. C. Boles, and J. H. White, in "DNA Topology and Its Biological Effects" (N. R. Cozzarelli and W. C. Wang, eds.), pp. 139-184. Cold Spring Harbor Laboratory Press, Plainview, New York, 1990. 4 j. C. Wang and L. E Liu, (1979) in "Molecular Genetics, Part liP' (J. H. Taylor, ed.), pp. 65-88. Academic Press, New York, 1979. 5 j. C. Wang, in "Nucleic Acid Research" (K. Mizobuchi, I. Watanabe, and J. D. Watson, eds.), pp. 549-566. Academic Press, New York, 1983.

[3]

53

MEASUREMENT OF BINDING AND UNWINDING

A

B

Wr=-5

Wr=.-5

ATw=0

ATw=0 i ~ _ ~

I

C Wr=-~

D

Ia

ATw=0 i

_

:

Wr=+2

l

c ~Lk=0

~LI(=-2

&l.k=-5

,~J..k=-7

Wr=0 ATw = 0

Wr=-2 ATw = 0

Wr=-5 ATw = 0

Wr=-7 ATw = 0

FIG. I. Schematic topo 1 relaxation reactions of ccDNA at no, low ([I]L), medium ([I]M), and high ([I]H) ligand binding, a, Drug binding; b, topo I relaxation; c, ligand removal by organic extraction. Ligand molecules are represented as nodules along the helical axis of the ccDNA shown, (A) Reaction without ligand, so that the ccDNA product is fully relaxed with ALkc = 0. (B) Reaction containing an amount of ligand that, when bound to the DNA, is sufficient to titrate out two superhelical turns (sht) so that the ccDNA prior to enzymatic relaxation contains - 3 sht. After steps b and c these two titrated sht are manifested in a final ALkc of - 2 . (C) Reaction containing just enough bound ligand to fully unwind all five sht, so that product and starting ccDNAs are indistinguishable. (D) Binding of ligand in high concentration to +2 sht beyond the equivalence point of (C). Here, the ccDNA product bears this additional superhelicity in its ALkc of - 7 . (In reality, some of the superhelical stress is taken up in Tw, so the number of Wr turns is less than the linking number deficit.)

54

BIOPHYSICALAPPROACHES

[3]

In reality, the resulting ceDNA is not a single species, but rather exists as a family of topoisomers (substantially identical DNA molecules differing relative to one another only in their topology), which differ by integral increments of the linking number. ALkj is defined for topoisomer j as the difference in superhelical turns (Tw + Wr) between an idealized, fully relaxed species for which Wr = 0, and a species j of writhe Wrj. Thermal fluctuations at the time of religation give rise to a Boltzmann distribution of topoisomers around an average ALk. 6-9 This central ALk (ALkc) is taken as the average most abundant topoisomer in the Boltzmann population, and is used in the following method as a comparative value between populations arising from Topo I reaction on ccDNA with varying intercalator concentrations. The simplest method to quantitate such topoisomeric reaction products is resolution by agarose gel electrophoresis. Separation results because the speed of topoisomer migration through an agarose gel is directly proportional to the absolute value of ALk. U n w i n d i n g a s M e a s u r e of E x t e n t of L i g a n d B i n d i n g The present method to calculate ~b and the binding constant Ka relies on resolving topoisomeric reaction products in two electrophoretic dimensions 10.~1 and analyzing the degrees of unwinding in a way formally analogous to the classic treatment of optical binding data.12 The incorporation of a second gel dimension, run perpendicular to the first, resolves topoisomers according to the handedness of their superhelicities, thus enabling resolution of negative and positive ccDNA topoisomers, which otherwise comigrate in just one electrophoretic dimension. Mathematical analysis of reaction products employs the Scatchard binding isotherm adjusted for nearest neighbor exclusion by bound ligand) 3~14expressed in closed formlS: I__/__"= Ka( 1 _ nr)F CF

1 - nr

],,-1

k 1 -- (-nT-1)r J

(1)

6 W. Keller, Proc. Natl. Acad. Sci. U.S.A. 72, 4876 (1975). 7 M. Shure and J. Vinograd, Cell 8, 215 (1976). 8 D. E. Depew and J. C. Wang, Proc. Natl. Acad. Sci. U.S.A. 72, 4275 (1975). 9 D. E. Pulleyblank, M. Shure, D. Tang, J. Vinograd, and H.-P. Vosberg, Proc. Natl. Acad. Sci. U.S.A. 72, 4280 (1975). 10 R. Bowater, E Aboul-Ela, and D. M. Lilley, Methods Enzymol. 212, 105 (1992). I I C.-H. Lee, H. Mizusawa, and T. Kakefuda, Proc. Natl. Acad. Sci. U.S.A. 78, 2838 (1981). 12 G. Scatchard, Ann. N.Y. Acad. Sci. 51, 660 (1949). 13 D. M. Crothers, Biopolymers 6, 575 (1968). 14 W. Mfiller and D. M. Crothers, J. Mol. Biol. 35, 251 (1968). 15 j. D. McGhee and P. H. Von Hippel, J. Mol. Biol. 86, 469 (1974).

[3]

MEASUREMENT OF BINDING AND UNWINDING

55

where CF is the free ligand concentration, 1"is the ratio of bound ligand to base pairs, Ka is the intrinsic association constant under the chosen conditions, and n is the number of sites (normally base pairs) covered by one molecule of bound ligand. Equation (1) simplifies to linear form at low binding when r << 1/n: r/CF = Ka[1 - (2n - 1)r]

(2)

Traditionally, r has been calculated optically via the measured apparent ligand extinction coefficient difference between free and bound states, Ae ap. With increasing DNA concentration, Aeap approaches Ae, and the fraction of bound ligand is A e a p / A e . With CB = (AEap/A6)CT, CB

r = -

CN

-

A6apCT

(3)

AeCN

where CB, CT, and CN are, respectively, the concentrations of bound ligand, total ligand, and total DNA binding sites, either base pairs or specific sites. Substitution of Eq. (3) into Eq. (2) and expansion to first order in (Ae - Aeap) yields the linear equation 16: AEap Aeap = A e -

Ka[CN - ( 2 n

- I)CT]

(4)

Here, a plot of Aeap versus A £ a p / [ C N - (2n - 1)CT] intercepts the ordinate at Ae as CN approaches infinity. The slope of the extrapolation yields Ka directly as its negative inverse. It is Eq. (4) that we have by analogy brought to bear on the problem of unwinding angle determination. Here A,gap and Ae are replaced by, respectively, ~)ap and 4~, the apparent unwinding elicited by incomplete binding of ligand where r << 1In and the unwinding angle of the ligand itself at extrapolated infinite CN. Hence Eq. (4) can be rewritten to describe the binding equilibrium of ligand to DNA by monitoring apparent unwinding, thereby yielding the structural parameter q~ rather than the optical descriptor Ae:

q~.p Ckap = ck - K~[CN - (2n - 1)Cr]

(5)

Thus, a plot o f q~apversus 4~ap/[CN -- (2n - 1)CT] has slope -Ka andy axis intercept q~ under the temperature and ionic conditions of the Topo I relaxation reaction. The concentration CN in Eq. (5) has alternate interpretations. With a nonspecific drug such as ethidium, CN is the total concentration of base pairs. In such cases the neighbor exclusion model applies, with n = 2 for ethidium. For drugs of known 16V. A. Bloomfield, D. M. Crothers, and I. T. Tinoco, Jr., in "Nucleic Acids: Structures, Properties and Functions," p. 552. University Science Books, Sausalito, California, 2000.

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BIOPHYSICALAPPROACHES

[31

sequence specificity, CN is the total concentration of specific binding sites, and n = 1 (assuming that the specific sites do not overlap). The method described here for determining ~b and Ka applies to low degrees of binding, and so is not suited for determining the neighbor exclusion parameter n. However, once ¢ is known, the degree of unwinding can be measured at any degree of binding, and the number of ligands bound can be calculated from the total unwinding divided by the unwinding per ligand, 4. The resulting binding isotherm can be used to determine n by fitting to Eq. (1). P r o c e d u r e in G e n e r a l The underlying principle of the present method is to run multiple Topo I relaxation reactions, each containing identical absolute amounts of DNA and ligand (and therefore identical DNA-to-ligand ratios), but carried out in different total reaction volumes. Increasing total reaction volume in a reaction series while maintaining constant the absolute molar amount of DNA and ligand leads, of course, to a decrease in concentration for both ligand and DNA. This decrease shifts the binding equilibrium toward an unbound state and the magnitude of this shift over the course of volume increase is governed by the intrinsic binding of the ligand, Ka, under the particular reaction conditions. It therefore makes sense that products of reactions run at higher volumes manifest less overall unwinding relative to those of reactions run at lower volumes, because the extent of unwinding effected by a particular intercalating ligand is directly proportional to the amount of that ligand that is bound to the DNA. That the base pair-to-ligand ratio remains unchanged (and high) throughout the dilution series ensures that i" remains low as required by the mathematical approximations given above. Reactions are treated and resolved as described below to determine ALkc, the Boltzmann center of mass of integrated topoisomer band intensity representing the average most populous topoisomer. For a given distribution of topoisomeric reaction products, ~bapis determined as follows: 360ND(ALkc - ALk °)

~ap ~

NL

(6)

where ALkc and ALk ° are the Boltzmann centers of topoisomer band intensity of, respectively, a reaction with ligand and a control reaction containing no ligand (this difference is A ALkc). ND and NL represent, respectively, the actual numbers of ccDNA and ligand molecules in a particular reaction. The ligand is presumed to be randomly distributed throughout all the ccDNA molecules present in the reaction mixture. The factor of 360 represents 360 ° per ALk in the ccDNA, so ~bapcarries the expected units of degrees. With qSapknown, Eq. (5) can be used to obtain ~b and Ka.

[3]

MEASUREMENTOF BINDINGAND UNWINDING

57

P r o c e d u r e in Detail We now describe the particular reaction conditions we have found to generate reproducible unwinding angles and association constants. Specific reaction schemes are given for the common intercalators daunomycin (DA) and ethidium bromide (EB), and it is hoped that these will allow the reader to standardize equipment and methodology to what has worked for us before, extending the method to binders of unknown q9 and/Ca. We have found that pUC19 plasmid works well as substrate DNA for the present method, and routinely use this DNA as isolated from transformed JM101 Escherichia coli and purified by the Qiagen (Valencia, CA) protocol. After purification, the DNA concentration ofpUC19 (2686 bp in length) is commonly adjusted to between 1.5 and 2 mM in base pairs, using a molar extinction coefficient of e260 13,200 M (bp) -I cm -I. DNA is stored at - 7 0 ° in a standard solution of 10 mM Tris-HC1 buffer at pH 7.5, and is thawed slowly at 4 ° prior to starting reactions in order to avoid unwanted nicking of ccDNA, ccDNA can typically be considered usable for the present method if it is in at least 95% supercoiled form as measured by one-dimensional agarose gel electrophoresis. It is a good idea to check this periodically, especially if a plasmid stock is more than 2 months old and/or has been freeze-thawed multiple times. Standard drug solutions of DA and EB are typically maintained in double-distilled water (ddH20) at the respective concentrations of approximately 60/zM (using an e480 of 11,500 M-l cm-l) and approximately 35 # M (using an E480of 5600 M -~ cm-I). We have found it fully satisfactory to purchase Topo I enzyme from GIBCO-BRL (Gaithersburg, MD) and use this directly as delivered in a concentration of 12 units/#l in 1 × Topo I buffer containing 50 mM Tris-HC1 (pH 7.5), 50 mM KC1, 10 mM MgC12, 0.5 mM dithiothreitol (DTT), 100 # M EDTA, and bovine serum albumin (BSA, 30 ~g/ml). We also keep a stock of 10 × Topo I buffer on hand for use in supplementing Topo I reactions of higher volume so that solution conditions remain constant in all reactions regardless of volume. =

Setup of Reactions Using Daunomycin as Intercalator To ensure uniform DNA aliquoting into all reactions, a 176-#1 premix containing 23.9 #1 of pUC19 ccDNA at 1.98 mM in base pairs, 22 #1 of 10 × Topo I buffer, and 130.1 #1 of ddHzO is mixed by light vortexing in an Eppendorf tube. This is intended to provide ample premix for 15-#1 aliquots into each of eight reactions in a series as well as two additional aliquots of 14 #1 each for verifying the DNA concentration actually going into each reaction. Checking this DNA concentration by spectrophotometry, using the above-described extinction coefficient, yields a DNA concentration in the premix of 346.8 # M in base pairs going into the reactions. However, the precise DNA concentration in the premix going into the reactions is not in itself important. Of much greater

58

BIOPHYSICALAPPROACHES

[3]

importance is to know exactly what this concentration is in each reaction volume, because it is used later to calculate ND, the number of plasmid molecules per reaction. This parameter is of vital importance for the calculation of ~bav, and hence for the construction of extrapolation plots according to Eq. (5) as explained above. Reactions are set up for DA according to the left-hand side of Table I. For reaction volumes of 22, 46, 86, and 166 /zl, polymerase chain reaction (PCR) tubes of 0.5-ml volume are used, whereas standard Eppendorf tubes of 2.0-ml volume are used for reaction volumes of 326, 646, 1006, and 1506 #1. Given the high density of the Topo I enzyme solution as obtained from GIBCO-BRL, it is important for complete reaction of all ccDNA that the reactions be gently mixed before being set aside for extended reaction incubation. Reaction conditions and workup are detailed further below.

Setup of Reactions Using Ethidium Bromide as Intercalator In general, the preparatory steps for reactions using EB as an unwinder are as described above for DA. Specific differences in a representative experimental run are as follows: 23.9/11 of pUC19 DNA at 1.98 mM in base pairs, 22/~1 of 10 × Topo I reaction buffer, and 130.1 #1 of ddH20 are combined to yield a premix solution of 176-#1 volume, sufficient for aliquots of 16 #1 each into each of eight unwinding reactions and two further aliquots of 16 #1 each for checking the concentration by spectrophotometry. According to this check, the DNA concentration of our premix is 302.9 #M in base pairs. Reactions are constructed with EB as an unwinder according to the left-hand side of Table II.

General Comments Regarding Construction of Reactions Note in Tables I and II that as the ligand and DNA concentrations decrease with increasing total reaction volume, the ratios of DNA-to-ligand concentration remain constant throughout. On the basis of the known concentrations of ligand and premix DNA solutions entering the reactions, it is possible to calculate the values of No and NL, the respective numbers of actual plasmid and ligand molecules in each reaction. According to the concentrations depicted in Table I, reactions using DA as an unwinding ligand contain 1.165 × 1012 plasmid molecules (No) and 1.463 × 1014 DA molecules per reaction (NL), implying a global value of 125.6 molecules of DA per plasmid. Similarly, Table II implies that each reaction using EB as an unwinding ligand contains 1.086 × 1012 plasmid molecules per reaction (ND) and 8.206 × 1013 EB molecules per reaction (NL), giving a global value of 75.56 molecules of EB per plasmid. The equations as explained above presuppose that for each reaction, whatever the volume, all drug present in a given reaction is treated as being bound. It is this assumption that allows calculation of ~bapfor each reaction according to Eq. (6)

II

© b.,

<~ <~

<

I t l l l l I I

e~

z 1 Z

0 Z > <

£

Z

II o Z

_z Z <

4 o

Z <

0

[.. o

x~

I

1

I

I

I

I I I I l l l

0 ,< .a

I I I I I I I

[-

z <~ I I I I 0 m

g.

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z

Z z Z <

r,

4

©

e~ Z <

$ b-

m

0

e~

[3]

MEASUREMENTOF BINDINGAND UNWINDING

61

above, given a knowledge of the difference ALkc-ALk ° (AALkc), NL, and No. The calculation of AALkc itself is described in further detail below. Because of the importance of an exact knowledge of ligand and DNA concentrations for reliable and reproducible calculation of ~ and Ka, we have found it necessary to calibrate all micropipetting apparati prior to the crucial steps of (1) premix preparation, (2) spectrophotometrically checking premix concentration, (3) aliquoting premix into reactions, and (4) aliquoting drug/ligand into reactions. Calibration of these apparati is performed at each preparation step by determining the exact apparatus setting required to deliver the water weight corresponding to the volume desired. Although this procedure may seem somewhat tedious, the sensitivity of the method to even small shifts in the ligand-to-DNA ratio, one of the method's advantages, has in our experience required such precautions; miscalculation of volumes by as little as 2.5% (0.1 /~1 on 4 ttl in added ligand volume, for instance) leads to measurable skewing of the result. Reactions and Workup Unwinding reactions are run in 50 mM Tris-HC1 (pH 7.5), 50 mM KC1, l0 mM MgCI2, 0.5 mM DTT, 100 #M EDTA and BSA (30 #g/ml) (1 × Topo I buffer) for 4 hr at 37 °. After this, samples are extracted with an equal volume of phenol-CHC13 (1 : 1) buffered to pH 7.5 with 50 mM Tris-HC1 to remove protein and ligand. The phases are partitioned and the aqueous phase from each sample is concentrated to a total volume of approximately 10 #1, using Microcon and Centricon centrifugal concentrators (Amicon, Danvers, MA). Small (about 0.5-#1) aliquots are removed for preliminary analysis by one-dimensional l% (w/v) agarose gel electrophoresis to check whether all unwinding reactions work properly (i.e., whether all ccDNA is acted on by Topo I) prior to commencing the more time-consuming two-dimensional electrophoretic analysis. Control reactions lacking unwinding ligand are identically reacted, extracted, and concentrated as those containing ligand. Concentrated reactions not immediately subjected to analysis by two-dimensional electrophoresis are stored at - 7 0 ° until analysis is carried out. Analysis by Two-Dimensional Electrophoresis Two-dimensional gel casting units (each 17 x 17 cm) are constructed by hand on the basis of established protocols. 1° Two glass runners (1 × 17 cm each) are attached lengthwise on opposite sides of a single glass base plate (17 x 17 cm), using silicone-based Dow Coming (midland, MI) aquarium sealant. During electrophoresis, these parallel glass runners are kept in an orientation parallel to the electric current to ensure perpendicularity between subsequent electrophoretic dimensions.

62

BIOPHYSICALAPPROACHES

[3]

Gels are cast with 250 ml of 1% (w/v) agarose in 80 mM TBE (80 mM Trisborate, 1 mM EDTA) to a final gel thickness of approximately 8 mm. A single loading well, 1 mm in diameter, is made in a comer of each gel by suspending an appropriately dimensioned peg into the gel while cooling. The first dimension is run for 20 hr in 1 liter of 80 mM TBE running buffer at 1.9 V/cm. After the first dimension has been run, the gel is rotated 90 ° in its casting tray, chloroquine is added in a small volume to a final concentration of 1.1 #g/ml, and the gel is soaked with light agitation for 1 hr. Electrophoresis in the second dimension then proceeds at 1.9 V/cm for an additional 18 hr. V i s u a l i z a t i o n a n d A n a l y s i s of T w o - D i m e n s i o n a l Electrophoresis Results After electrophoresis, gels are soaked in an EB solution of at least 1 ttg/ml for 2 hr, destained in at least two changes of water over 2 hr, and photographed on a UV transilluminator with black-and-white Polaroid film. As the total amount of DNA corresponding to a given topoisomer embedded in the gel is sometimes small, especially at the extremes of Boltzmann distribution (see Fig. 2), we have found it necessary to use as high concentrations of EB as possible in the staining solution. In this way, we are able to maximize the otherwise faint contrast in these regions during visualization and photography. Photographs obtained in this way are scanned with a Hewlett-Packard (Palo Alto, CA) Scan Jet 3P and the photonegatives of individual topoisomer bands are quantitated with NIH Image software. For a given Boltzmann distribution of topoisomer band intensity in a gel, we typically measure the intensity of each topoisomer individually by positioning an electronically delimited measurement region such that the entire topoisomer area lies within this region. This measurement region is typically extended to be several times longer than the respective topoisomer bands being measured in order to encompass ample regions of surrounding baseline area containing no topoisomers. For this reason, it is important to run two-dimensional electrophoresis long enough to fully resolve a given topoisomer from its neighbors. Baselines are typically constructed by sampling several gel regions devoid of any topoisomers with the same measurement region described above and then averaging the result. This baseline is then subtracted from each topoisomer intensity peak in subsequent analysis. We used KaleidaGraph software (Synergy Software, Reading, PA) to analyze resulting data sets for each region of topoisomer intensity, although any graphic analysis program can be used for this purpose. The integrated intensities for each topoisomer are then plotted to give a Boltzmann distribution from which ALkc is calculated by fitting to I = ]M e x p [ - w ( A L k -- ALkc)], where ] and ]M are, respectively, integrated band intensity and maximum band intensity at the peak of the topoisomer distribution, and w is the width of the distribution. Figure 2 shows an

[3]

63

MEASUREMENT OF BINDING AND UNWINDING

I

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1 :

~

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..tJ.. "O

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0.6

,"

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-4

I

,'; o° ,_.1,.~'°1~ • I

-2

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,

91~o

:

I , "A--...I

,

2

1~ I *l',

4

zXl_k FIG. 2. Example calculation of ~bapaccording to Eq. (6). EB-stained gels are shown (top) with their corresponding Boltzmann distributions (bottom). The results shown are from reactions containing (left, a) and lacking (right, ©) the unwinding ligand DA, and values for ALk and electrophoretic dimensions are indicated on and next to the gels, respectively. ALkc is -0.951 for the distribution from the reaction containing DA, and the maximum of the ligand-free control reaction, ALkc°, is +1.49. This implies a AALkc of --2.44. With NL = 1.46 x 1014 ligands reaction and ND = 1.17 × 10 j2 ccDNA/reaction, and assuming that all ligand present in the reaction is bound to the ccDNA, Eq. (6) gives (Pap = - 7 . 0 3 °. This value of 4~ao then gives rise to a single point in an extrapolated binding isotherm plotted according to Eq. (5) as shown in Fig. 3.

example of plotted topoisomeric intensities for reaction product distributions from two reactions, one with and one without ligand. Under the conditions described herein, control reactions containing DNA but no unwinding ligand repeatedly give topoisomeric product distributions centering on ALkc° = + 1.5 superhelical turns, and this value is used in all calculations of A ALkc. An example of how one arrives at a value of AALL:, and thus at ~bap, for a given unwinding reaction containing unwinding ligand is shown in Fig. 2. The extrapolation plots obtained with ~bap, CN, CT and n from the above-described reaction series using DA and EB are shown in Fig. 3.

64

BIOPHYSICAL APPROACHES I

I

'

'

[3]

%

%

-5 -10

~

•

*aP_15 -20 -25 -3O -35

I

-2

I

d

t

I

-1.5

I

B

J

I

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i

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I

i

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-1 -0.5 ~ap/(CN- (2n-1)Cr)

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FIG.3. Binding isotherms for DA (C], n = 4) and EB (O, n = 2). Unwinding data obtained as shown in Fig. 2 are plotted according to Eq. (5). Interceptionof the ordinate at extrapolated infiniteCN yields ~bdirectly, and the negative inverse of the slope yields K a. An additional hypothetical isotherm for DA (O, n = 6) has been included to show that the intercept value of ~bdoes not depend on the value of n used. Valuesfor n = 4 for DA and n = 2 for EB are accepted as standard for these ligands in the literature.14.15

Nonuniform binding of EB during gel staining might be expected to introduce artifacts in observed band intensity that could skew the calculated value of ALkc. Both the superhelical state of the ccDNA as well as the absolute amount of ccDNA in a given band might be expected to cause such nonuniformity in EB binding. Two sets of control experiments are performed in order to investigate these possibilities. In one, identical amounts of ccDNA intercalated by varying amounts of EB are completely relaxed and treated as described above so that the product ccDNAs span a range of superhelicities. These products are resolved by one-dimensional agarose gel electrophoresis and visualized by EB staining. For the range of superhelicities relevant for the present method, the observed intensity (quantitated as described above) shows no systematic dependence on ALk. In another control, varying absolute amounts of ccDNA homogeneous in A L k are resolved by one-dimensional agarose gel electrophoresis and visualized by EB staining. Quantitation of total topoisomer band intensity shows a linear dependence of intensity on the amount of ccDNA up to 1.2 /zg of ccDNA, after which point the intensity saturates. Because the present method involves no twodimensional topoisomer band containing more than 0 . 6 / z g of ccDNA, all quantitations of band intensity are well within the linear range required for reliable calculations.

[3]

MEASUREMENT OF BINDING AND UNWINDING

65

General Comments Regarding Method Past methods for determining ~bhave required knowledge of either Lk or v, the maximum bound mole fraction of ligand per base pair. The present assay for the simultaneous determination of ~band/Ca differs in this respect in that it presupposes no prior knowledge of either Lk or v, employing instead sufficiently high CN-to-CT ratios such that extrapolation to infinite CN is possible. At this infinity point, the ligand is assumed to be essentially completely bound to the DNA, a scenario in which ~ap = ~b. The prerequisites for the present method are as follows: (1) that the/Ca of the ligand-DNA complex be sufficiently large to warrant the assumption of near complete ligand binding at high Cy, (2) that the size (in base pairs) of the ccDNA used in the relaxation reactions be known; and (3) that an estimate of the number of potential binding sites along a molecule of the ccDNA be used. Consideration of Eq. (5) reveals the need for an estimate of n, the number of potential binding sites covered by one molecule of bound ligand. For intercalating ligands such as DA and EB, which show only minimal sequence selectivity in their binding to bulk DNA, each base pair in the ccDNA may be seen as making an equal contribution to the number of total viable binding sites. Under these conditions potential binding sites overlap one another and the neighbor exclusion model13- Is applies, with n greater than unity [notice that substitution of n = 1 into Eq. (1) regenerates the original Scatchard binding isotherm,12 in which independent potential binding sites do not overlap]. If the serially diluted reaction series is run at a sufficiently high DNA-to-ligand ratio, then Ca- << Cy and the choice of n in Eq. (5) has little effect on the resulting plot. For less potent unwinders, it may be necessary to run reactions of higher ligand concentration in order to generate a quantifiable A ALl%. In these situations, the CT/CN quotient rises and the choice of n becomes more important. However, even here, n affects only the slope of the line in Eq. (5) (the source of/Ca), and has no bearing on the intercept of the line with the ordinate, from which the unwinding angle is determined. Given a reasonable estimate of the number of potential binding sites along the ccDNA, this method is therefore capable of generating a good measure of q~ regardless of the chosen n. For exceedingly weak unwinders, a detectable AALI% is sometimes possible only by addition of excess ligand such that all potential binding sites remain saturated even at low CN. At this point, Eq. (5) becomes invalid. The limit of the applicability of the method is thus reached as the Cv/CN ratio approaches unity. The choices of n = 2 for EB and n - 4 for DA in the methodology shown above are based on the literature.~7,~8 When relaxing ccDNA with Topo I in the absence of ligand, theory predicts that the average most populous topoisomer should center on the fully relaxed species 17 H.-W. WU, N. Dattagupta, M. Hogan, and D. M. Crothers, Biochemistry 19, 626 (1980). J8 j. B. Chaires, N. Dattagupta, and D. M. Crothers, Biochemistry 22, 284 (1983).

66

BIOPHYSICALAPPROACHES

[3]

for which ALk ° = (Fig. 1A). Yet the value for ALk ° used in the above-described calculations is not zero, but ÷ 1.5. This shift of ÷ 1.5 superhelical turns away from the expected zero results from the fact that the enzymatic relaxation reaction is carried out at higher ionic strength than the subsequent two-dimensional agarose gel analysis, and the DNA helical repeat depends on ionic strength. However, the writhe that results from this twist change has no effect on the calculation of AALkc, because both ligand-free and ligand-containing samples are reacted and treated identically. Because superhelicity of ccDNA partially depends on ionic strength, it is of paramount importance to ensure identical salt conditions for all samples during reaction. To this end, supplementing samples of higher volume with appropriate additional reaction buffer is essential. It is then merely a matter of convention which topoisomer is chosen to use as a reference to compare control and ligand-containing samples. For this, the Boltzmann, center of topoisomer band intensity is the most obvious and most accurately calculable point of reference for the determination of A ALkc. It is therefore to be expected that changing salt and/or temperature conditions from those described herein will result in the control value of ALk ° being other than +1.5 for pUC19 DNA. Of course, choosing another ccDNA species as substrate for the unwinding reactions would be expected to yield a different value of ALk ° altogether, even for ionic and temperature conditions identical to those given above. Typical Problems Encountered Topoisomer Bands in Two-Dimensional Gels Too Faint to Measure

Faint bands represented a recurring problem in the development of the method. We found that this problem has two sources. One cause of faint bands is the use of ccDNA that is too large. Because the ccDNA concentration is spectrophotometrically determined with respect to base pairs in the premix, a given base pair concentration will imply fewer actual ccDNA molecules for a large plasmid than for a small one. Furthermore, as the size of ccDNA increases, so too will the number of topoisomers generated from this ccDNA, resulting in a scenario for large ccDNA in which fewer actual plasmid molecules are distributed over more topoisomer bands. This reduces the amount of ccDNA present in a given band, effectively attenuating the intensity of each band beyond a level compensable even by staining with high EB concentrations. A second cause of faint bands is circular DNA that is not intact, that is, DNA that has a single-stranded nick. Such DNA is not capable of forming topoisomers because there is free rotation at the nick. Nicked molecules are readily distinguished by their electrophoretic mobility, but if present in large amount, they reduce the amount of DNA in the topoisomer distribution, reducing the quantitative reliability

[3]

MEASUREMENTOF BINDINGAND UNWINDING

67

of the measurements. DNA solutions prone to such deleterious nicking include those that have been subjected to multiple freeze-thaw cycles, that have been stored too long in liquid phase, and/or that are too large.

Inadequate Resolution of Topoisomer Bands Inadequate resolution might be the result of several factors. Because of its high degree of conformational freedom, ccDNA that is too large (i.e., plasmids with too many base pairs) will yield more topoisomers in a Boltzmann distribution as shown in Fig. 2 than smaller plasmids. As the number of topoisomeric bands increases, the ability of a given agarose gel to resolve them adequately naturally decreases. For this reason, we have found pBR322 (4362 bp) too large to adequately resolve into discrete topoisomer bands that can be individually quantitated. On the other hand, care must be taken not to decrease the size of the ccDNA used to such an extent that there no longer exist enough topoisomer bands to construct a statistically meaningful Boltzmann distribution after quantitation; that is, such that the Boltzmann distribution becomes "digital" in character. The ideal substrate is of a size that avoids both extremes. As stated above, we have found that pUC 19 DNA yields a sufficient number of topoisomer bands for constructing meaningful Boltzmann distributions, while not giving rise to so many topoisomer bands that these are not separable from one another on agarose gels. Gels in which the agarose density is too low tend to be incapable of resolving topoisomer bands sharply enough to render their intensities discretely measureable. On the other hand, the agarose density must not be increased so much that inordinate running times and/or brittleness result. We have found that an agarose density of about 1% yields topoisomer bands that are discretely quantifiable. Although the running times associated with this agarose density are indeed somewhat long (18-20 hr per dimension), this is not active bench time requiring constant attention and, if planned well and run in parallel on multiple two-dimensional apparatuses, allow a full series consisting of eight reactions to be analyzed in several days.

DNA Incompletely Relaxed after Topoisomerase I Reactions The persistence of highly supercoiled ccDNA after reaction with Topo I is most commonly caused by inhomogeneity of the reaction mixture with respect to Topo I enzyme, leading to insufficient contact between enzyme and ccDNA. This can be alleviated by proper but careful mixing of the reaction mixture containing enzyme prior to reaction. Care, however, must be exercised such that mixing is performed gently (i.e., no vigorous mixing), because rigorous conditions will inactivate the Topo I enzyme and/or nick the ccDNA, rendering it useless for the purposes of the assay. For reactions in which the total volumes are large enough (i.e., those of at least 300 #1), repeated gentle inversion of the reaction tube back and forth was found to be the best way of distributing the dense enzyme solution uniformly

68

BIOPHYSICALAPPROACHES

[4]

throughout the reaction mixture. For smaller reactions, gentle pipetting of the solution up and down was found to be sufficient to mix the enzyme while retaining its integrity as well as that of the DNA. Another cause of incomplete relaxation of DNA can be related to the ligand itself, the unwinding properties of which are to be measured. Certain DNA intercalators act as poisons to Topo I, preventing the latter from relaxing supercoiled DNA. Obviously, such a ligand cannot be assayed with the present method using Topo I. An alternative to Topo I is topoisomerase II (Topo II), a homodimer enzyme that can effect the same unwinding of ccDNA as Topo I, albeit via a different mechanism. Many compounds that inhibit the topology-altering characteristics of one of the two enzymes will not poison the other. However, Topo II requires reaction conditions that differ from those of Topo I. Given the dependence of superhelical state on such conditions, use of Topo II in place of Topo I therefore necessitates the running of ligand-free controls under Topo lI conditions in order to obtain a meaningful value for ~ L k °. However, if at all possible, reactions should as a general rule be run with Topo I rather than with Topo II, because the former tends to be more robust and yields more reproducible results.

[4] L i n e a r

and

Drug-Nucleic

Circular

Dichroism

of

Acid Complexes

By MAGDALENAERIKSSONand BENGTNORD1EN Introduction Polarized light spectroscopy offers possibilities to quickly characterize nucleic acids and their complexes with bound proteins or drug molecules, using a relatively small amount of sample. Linear dichroism (LD) provides structure information in terms of relative orientation between the bound drug molecule and the DNA molecular long axis, and also about the effects of ligand binding on the host. Circular dichroism (CD) may contribute additional structural details about such complexes. LD can be used to sensitively measure the directions of light-absorbing transition moments of individual chromophores. Specifically, linear dichroism can be used to probe angles between transition moments of DNA bases relative to the orientation axis, which in flow or electric field-oriented DNA coincides with the helix axis. The induced CD (ICD) of the same transition may provide information about molecular orientation relative to the surrounding nucleobases. A ligand chromophore bound to a long DNA molecule oriented by hydrodynamic flow or in an electric field gives rise to an LD signal that itself constitutes

METHODS IN ENZYMOLOGY. VOL. 340

Copyright 2(X)I by Academic Press All rights of reproduction in any |brm reserved. (1(176-687W00 $35,00

68

BIOPHYSICALAPPROACHES

[4]

throughout the reaction mixture. For smaller reactions, gentle pipetting of the solution up and down was found to be sufficient to mix the enzyme while retaining its integrity as well as that of the DNA. Another cause of incomplete relaxation of DNA can be related to the ligand itself, the unwinding properties of which are to be measured. Certain DNA intercalators act as poisons to Topo I, preventing the latter from relaxing supercoiled DNA. Obviously, such a ligand cannot be assayed with the present method using Topo I. An alternative to Topo I is topoisomerase II (Topo II), a homodimer enzyme that can effect the same unwinding of ccDNA as Topo I, albeit via a different mechanism. Many compounds that inhibit the topology-altering characteristics of one of the two enzymes will not poison the other. However, Topo II requires reaction conditions that differ from those of Topo I. Given the dependence of superhelical state on such conditions, use of Topo II in place of Topo I therefore necessitates the running of ligand-free controls under Topo lI conditions in order to obtain a meaningful value for ~ L k °. However, if at all possible, reactions should as a general rule be run with Topo I rather than with Topo II, because the former tends to be more robust and yields more reproducible results.

[4] L i n e a r

and

Drug-Nucleic

Circular

Dichroism

of

Acid Complexes

By MAGDALENAERIKSSONand BENGTNORD1EN Introduction Polarized light spectroscopy offers possibilities to quickly characterize nucleic acids and their complexes with bound proteins or drug molecules, using a relatively small amount of sample. Linear dichroism (LD) provides structure information in terms of relative orientation between the bound drug molecule and the DNA molecular long axis, and also about the effects of ligand binding on the host. Circular dichroism (CD) may contribute additional structural details about such complexes. LD can be used to sensitively measure the directions of light-absorbing transition moments of individual chromophores. Specifically, linear dichroism can be used to probe angles between transition moments of DNA bases relative to the orientation axis, which in flow or electric field-oriented DNA coincides with the helix axis. The induced CD (ICD) of the same transition may provide information about molecular orientation relative to the surrounding nucleobases. A ligand chromophore bound to a long DNA molecule oriented by hydrodynamic flow or in an electric field gives rise to an LD signal that itself constitutes

METHODS IN ENZYMOLOGY. VOL. 340

Copyright 2(X)I by Academic Press All rights of reproduction in any |brm reserved. (1(176-687W00 $35,00

[4]

LD AND CD OF DRUG-NUCLEICACID COMPLEXES

69

proof of the existence of an associative interaction. Small molecules tumble freely in solution and are isotropically (randomly) oriented and will, therefore, in contrast to DNA-bound molecules, not give rise to any LD signal in their absorption region. The presence of detectable LD thus means that the ligand is bound to the oriented DNA, while the sign and amplitude of the LD signal contain information about the nature of the predominant binding mode. Use of complementary techniques, such as CD and fluorescence spectroscopy, may allow further conclusions regarding the structure and dynamics of DNA-ligand complexes. Nonchiral drug molecules (molecules that lack handedness and thereby optical activity) have no CD signal. However, when bound to DNA nonchiral molecules often give rise to CD spectra because of induced CD (ICD) resulting from the chiral surrounding of the molecule. Similar to an LD spectrum, an ICD signal observed in the absorption band of a nonchiral ligand is direct evidence of interaction with DNA, and also contains information about the drug-DNA interaction. Changes in the intrinsic CD of DNA or of the ligand itself (in case it is chiral) indicate that a change has occurred, possibly but not necessarily of a structural nature, and are more difficult to interpret. Historically, induced optical activity, observable as a CD signal in the absorption band of nonchiral dyes interacting with biopolymers, has played an important role as a diagnostic tool in the study of drug binding to proteins and nucleic acids. The ICD signal in the spectral region where the drug absorbs light is typically weaker than that of the DNA molecule. Therefore the sample concentration may have to be higher or the optical path length increased for detection of ICD in the drug. This particularly applies to intercalating molecules, for which the ICD tends to be weaker than for groove binding ligands. Here we emphasize the practical use of LD in combination with CD, in particular ICD, for distinguishing between different types of DNA-drug interactions, such as intercalation and major or minor groove binding. We also discuss how to determine the orientation and location of the ligand within its binding site, and consider the occurrence of ligands bound as dimers. For in-depth descriptions of LD and CD, please refer to earlier review articles, 1 or to books. 2-4 Methodology

Basic Mechanism of Absorption of Polarized Light A transition between two molecular energy states can be induced by electromagnetic radiation provided that there exists a molecular electric or magnetic dipole moment, resulting from specific movement of electrons, with which the electric or magnetic component of the radiation field can interact. I B. Nord6n, M. Kubista, and T. Kurucsev, Q. Rev. Biophys. 25, 51 (1992). 2 N. Berova, K. Nakanishi, and R. W. Woody, "Circular Dichroism--Principles and Applications." Wiley-VCH, New York, 2000.

70

BIOPHYSICALAPPROACHES

[41

Linear Dichroism Linear dichroism, or absorption anisotropy, arises if the light-absorbing molecules are preferentially oriented relative to a laboratory (macroscopic) frame of reference. In contrast to CD, which depends on both electric and magnetic interactions, LD is a purely electric phenomenon. Linear dichroism (LD) is measured in absorption units and is quantitatively defined as the differential absorption: LD = All - A±

(1)

where All and A± refer to the absorption measured with light linearly (plane) polarized parallel and perpendicular, respectively, to a reference direction in the laboratory (by convention the orientation axis; Fig. 1A). The polarized light absorption is proportional to cos2ot, where a is the angle between the molecular electric transition moment and the electric field polarization direction of the light. For quantitative interpretation the LD is generally normalized by division by the absorption measured for the same sample in an isotropic state, that is, in absence of flow or electric field, to form the reduced linear dichroism, LD r, at each wavelength: LDr()0 = LD(2.)/Aiso()0

(2)

DNA Orientation Methods Hydrodynamic Flow. Extended molecules, such as DNA, are readily oriented by hydrodynamic flow. The Couette cell is a device designed for this purpose, consisting of two concentric quartz cylinders. The diameter of the inner cylinder is typically 3 cm, and the two cylinders are separated by a 0.5-mm gap into which the sample solution (2 ml) is inserted (Fig. 2). As one of the cylinders (inner or outer) rotates, typically by 1000 revolutions per minute (rpm), a flow that is characterized be a shear gradient of around 3000 sec-1 results. The flow may align DNA or polynucleotide molecules as short as a few hundred base pairs, sufficient for LD measurements to be conducted. The shear gradient, G, measured in sec I is calculated as G = Jr D (rpm)/60 d, where D is the diameter of the cylinder, rpm denotes the rotation speed, and d is the distance of the annular gap. Stiff molecules, such as single- or double-stranded DNA in complex with the protein RecA, are efficiently orientated in the Couette cell by rotating the cylinder at 20 rpm. Such slow flow achieves selective orientation of RecA-bound DNA molecules, because uncomplexed DNA is virtually unaffected by such low gradients. LD can thus be used to measure the amount of complex formed and can, for example, be used to monitor binding kinetics. 3 A. Rodger and B. Nord6n, "Circular Dichroism and Linear Dichroism." Oxford University Press, London, 1997. 4 j. Michl and E. W. Thulstrup, "Spectroscopy with Polarized Light. Solute Alignment by Photoselection, in Liquid, Polymers, and Membranes." VCH, Weinheim, Germany, 1995.

[4] (A)

L D AND C D OF DRUG-NUCLEIC ACID COMPLEXES

Plane polarized light

Oriented molecule

71

Absorption spectrum A

II

A LD = A l l - A± LD

"V LD spectrum

(B)

Circularly polarized light

Chirai molecule

CD spectrum CD

-k_/ CD = A k - AR FIG. 1. Principles of LD (A) and CD (B) measurements.

~

k

72

BIOPHYSICALAPPROACHES

[41

3_. I

FIG.2. Schematicrepresentationof Couette cell. As an alternative means of achieving orientation by flow, the DNA solution may be pumped through a capillary or an infrared (IR) sandwich cell. In a cylindrical capillary the flow adopts uniaxial symmetry and thus the LD can be obtained from the difference in absorption, in presence (AI0 and in absence (Aiso) of flow, measured with light polarized parallel with the flow lines: AA = All - Aiso. LD = All -- A± is then calculated using the general expression for uniaxial samples: Aiso= (1/3)(All + 2A±)

(3)

LD = (3/2)(AII - A~so)

(4)

Thus, for a uniaxial sample:

A stopped-flow apparatus with a polarizer inserted into the light beam can also be used to accomplish flow that is adequate for measuring LD, and can be used to gain information about orientation dynamics and reaction kinetics. There are, however, drawbacks; only short transients may be recorded because equilibrium quickly is reestablished (compared with the steady state orientation of the Couette method) and the relatively large amounts of sample that are required. Humid Polyvinyl Alcohol Film. A film made of polyvinyl alcohol (PVA) is a suitable matrix for aligning shorter DNA fragments (20-1000 base pairs). PVA, when equilibrated to a high level of humidity, becomes elastic and can be stretched while retaining a relatively large amount of water. The solute (DNA) is added to a

[4]

LD AND CD OF DRUG-NUCLEICACID COMPLEXES

73

5-10% solution of PVA in buffer prepared by dissolving commercially available PVA powder in cold water, heating while stirring vigorously, and then cooling. The film is cast on a horizontal glass plate and most of the water is allowed to evaporate, which takes a day or two at ambient temperature. The dry film is mounted in a stretching device and allowed to equilibrate in a humid chamber for a few hours prior to stretching and conducting measurements. Electrophoretic Orientation. DNA can become oriented through migration through a gel matrix, induced by application of an electric field. The mesh provided by the gel, typically agarose, can achieve a high degree of alignment of long molecules, such as T2 phage DNA, through mechanical interactions with the gel fibers. The time dependence of the LD signal contains information about the dynamic motion of the DNA molecule migrating through the gel, changing configuration between compact coils and stretched states, and has explained the mechanism behind pulsed-field electrophoresis. 5 Electrophoretic orientation can also be used to study the effect on helix flexibility by the interaction of drugs. 6 Electric Field. A classic method of orienting DNA is to a apply a high voltage ( 1 kV) across two parallel electrodes, typically separated by 2 mm, in a l-cm quartz cuvette. The change in absorption, measured with light polarized parallel with the electric field, is observed on an oscilloscope as a function of time. For evaluation of ~xA = All -- Also in terms of LD, the same equation [Eq. (4)] as for uniaxial flow in a capillary may be used. Because of heat dissipation only short electric field pulses (1 msec) can be used and although the orientation can be high (S = 0.5 or more for long DNA), the method is tedious as it requires the recording of many single transients. Electric orientation also has the disadvantage of being restricted to low salt conditions (< 10 mM NaCI). Determination o f Electric Transition Moments. Utilization of LD for structure studies requires knowledge of how the electric transition dipole moments are directed within the molecule of interest. The electric transition moments of three drug molecules and one DNA base are indicated in Fig. 3. Transition moment directions of a drug molecule may be determined experimentally by measuring LD of the drug alone. This is done by dissolving the molecule in a plastic film that can be stretched to obtain a high degree of alignment of the drug. For water-soluble molecules the following procedure can be applied. First, dissolve the drug in a 10% mixture of polyvinyl alcohol and water. Pour the mixture onto a horizontal glass plate and let the water evaporate until a thin (0.1-mm) film remains. Second, align the molecules in the film by stretching the film in one direction (elongating the film some three to five times its original size) after heating it to approximately 80 °, for example, in the hot air from a hair dryer. Third, measure the absorption after inserting a polarizer in the light beam preceding the sample film. Record two spectra, one 5 B. Nordrn, C. Elvingson, M. Jonsson, and B. Akerman,Q. Rev. Biophys. 24, 103 (1991). 6 K. Gisself~ilt,P. Lincoln, B. Nordrn, and M. Jonsson,J. Phys. Chem. B 104, 3651 (2000).

74

BIOPHYSICALAPPROACHES

[4]

with the polarizer parallel and one with the polarizer perpendicular to the stretch direction. If the absorption of a certain absorption band is strong when the light is polarized parallel and weak (close to vanishing) when polarized orthogonal to the sample orientation axis, the transition moment for that absorption band is directed parallel to the long axis of the dye. This experiment illustrates the physical role of a transition moment as an oriented antenna through which the molecule absorbs light of a particular wavelength and how light of specific polarization directions is absorbed. A large value of LD r (typically near the upper theoretical limit, -t-3, for a uniaxial sample) would indicate that the dye molecules are nearly perfectly oriented within the film. For molecules that do not have an obvious long axis, a number of orientation parameters (five of them in the general case and three for a planar molecule) must be determined. A general theoretical description of the treatment of LD data for planar molecules in stretched films is given in Appendix I. The electric transition moments may also be calculated theoretically. There are at

(A)

300-350

nm

N

2 8 0 nm

2,7-dlazapyrene

(B)

450-500

300 nm

k ~ /

nm

k . ~

ethidium FIG. 3. Electric transition dipole moments in (A) a symmetric chromophore (2,7-diazapyrene), (B) an unsymmetric planar DNA intercalator (ethidium), (C) a minor groove binder (DAPI), and (D) the DNA base adenine.

[4]

LD AND CD OF DRUG-NUCLEICACID COMPLEXES

75

present several software packages available for performing quantum chemical calculations that include options for obtaining transition moment directions, oscillator strengths, and wavelengths of molecular absorption bands. Circular Dichroism The principle of measuring CD is schematically illustrated in Fig. lB. The occurrence of a CD spectrum requires that a molecule possess, in addition to an electric transition moment, a magnetic transition moment. This makes CD spectroscopy, in comparison with LD, a conceptually more complicated method that is not as straightforward to interpret. The amplitude of the CD signal is proportional to the scalar product between the electric and magnetic transition moments. It is not practically possible to determine magnetic transition moments experimentally, so in cases when a molecule possesses an inherent magnetic transition moment one must resort to theoretical calculations. Most of the common small drug molecules, however, lack intrinsic magnetic moments but still give rise to CD spectra. Their

(c) NH2 M.,

~

/'--~./'~

34o nm

NH2

4',6-diamidinophenyl-2-indole

(D) .NH2 N

N

248 nm

adenine FL~. 3. (Co.tinued)

(DAPI)

76

BIOPHYSICALAPPROACHES

[4]

CD signals are instead caused by intermolecular interactions, typically between electric transition moments of the ligand and its host, and are therefore relatively easy to handle. Optical activity (and circular dichroism) can arise only if there are nonvanishing parallel components of electric and magnetic transition moments that interact to generate helical paths for the mobile electrons within of the molecule. If such electrons are excited by circularly polarized light of appropriate wavelength, resonance will occur, which can be seen as a CD signal. The ligands that will be considered here are nonchiral (optically inactive) molecules that on interaction with DNA become optically active. The system of nonchiral drugs and DNA may acquire a magnetic transition moment that has a component parallel to an electric transition moment and thereby gives rise to a CD signal. We note that the interacting molecular groups or species giving rise to ICD through this mechanism are at Van der Waals distances (3-10 A). In its simplest form ICD arises from the coupling of electric transition moments of the DNA and of the ligand. Knowledge of the magnitude and direction of host and guest electric transition moments is therefore essential for extracting structural information from such induced optical activity. Changes in the CD spectrum of DNA as a result of complexation can often be ascribed to alterations in the DNA structure. The reverse, a significant redistribution among different conformational species of the ligand induced by its interaction with DNA, may lead to an excess of one chiral ligand form and thereby affect its apparent ICD spectrum. Such effects have been observed with the inversion-labile trigonal iron complex [Fe( 1,10-phenanthroline)3] 2+.7 DNA-Ligand Binding. The negatively charged phosphate groups along the backbone of DNA may attract positively charged ligands, such as many drugs and DNA-binding domains of proteins. However, their influence on the preferred binding modes of ligands is generally considered to be of secondary importance. What type of DNA binding mode is most favored by a ligand is rather governed by nonionic interactions. Three types of ligand binding to duplex DNA in B conformation are generally recognized: intercalation between base pairs, minor groove binding, and major groove binding, as schematically shown in Fig. 4. Intercalation. Intercalation, the insertion of a planar molecule, such as an aromatic drug molecule, between consecutive base pairs of DNA, is accompanied by distinct changes in the local DNA structure. The base pairs surrounding the intercalator have to separate by around 3.4 A to allow space for the drug. Such separation leads to increased DNA contour length, which generally improves its orientation in flow [increased S in Eq. (5)]. Furthermore, the DNA helix must locally unwind from its unperturbed helical twist of 36 ° to about 10° in order to accommodate the ligand without breaking the phosphate-sugar backbone. Intercalation of a light-absorbing drug molecule can readily be established by linear 7 T. Hard and B. Nord6n,Biopolymers25, 1209 (1986).

[4]

LD AND CD OF DRUG-NUCLEICACID COMPLEXES

(A)

(B)

Intercalation

77

(C)

Minor groove binding

Major groove binding

FIG. 4. Schematic illustration of DNA binding modes. (A) Drug intercalation, (B) minor groove binding drug, and (C) major groove binding protein.

dichroism, because the intercalated molecules are close to coplanar with the DNA base pairs. Intercalative binding occurs with site exclusion: occupation of a site prevents intercalation into immediate neighboring sites. 8'9 This simplifies the situation in the sense that, because of the relatively large distance between their transition moments, the so-called exciton interactions (see below) between drug molecules at different intercalation sites may be neglected. Groove Binding. The two grooves of DNA in B conformation are significantly different. The major groove is approximately 9 • deep and 12 ,~ wide whereas the minor groove is approximately 8/~ deep and 6/~ wide. The narrow shape of the minor groove provides a good match for several groove binding aromatic ligands. However, G-C base pairs are sterically less accessible in the minor groove because of the exocyclic amino group of guanine protruding into the minor groove. Table I lists examples of DNA binding molecules along with their spectral characteristics and DNA binding properties. I n t e r p r e t a t i o n of S p e c t r a

Linear Dichroism Intercalative binding can be demonstrated using LD to show that the molecular plane of the ligand is parallel to the DNA base pairs. The reduced linear dichroism, obtained by dividing the LD spectrum by the isotropic absorption counterpart, A,

8 R. W. Armstrong, T. Kurucsev, and U. E Strauss, J. Am. Chem. Soc. 92, 3174 (1970). 9 j. D. McGhee and P. H. von Hippel, J. Mol. Biol. 86, 469 (1974).

78

BIOPHYSICAL APPROACHES

.I

!

[4]

.I.~-.I .I=== .~

iiiiii

:i:i i:!i.-! i i

.I %

._i~

i~~~~.o ~.-

i £

==

Z

i i i zm

i

@

z

~

< <

~z

i

.II i

.I

i" I'~ -~"=

g

i=-

~

.I

~ 0 ~

£()

~E

i;

~

~

i

i

[4]

LD AND CD OF DRUG-NUCLEIC ACID COMPLEXES

79

can be written as ~ LDr()~) = LD()~)/A()~) = S(3/2)(3 cos ~2 _ 1)

(5)

where o! is the angle between the transition moment of the chromophore and the orientation axis (see Fig. 5a). The orientation parameter, S, describes the degree of molecular alignment, assuming the value 1.0 for perfectly aligned molecules and 0 for an isotropic sample. S can be evaluated from the value of LD/A in the DNA absorption region, at 260 nm, setting the effective orientation of the base pair transition moments, OfDNA,to 86 ° for mixed-sequence B-form DNA. ~0 Once S has been determined, the same orientation factor can be used in Eq. (5) for calculating the average angle between the transition moment direction of the ligand and the helix axis, o~liga,d, using the LD/A measured in the ligand absorption region. This treatment of LD data is applicable to intercalating as well as groove binding DNA ligands. Similar analysis can also be used for characterizing the conformations of various polynucleotides and their responses to different solvent conditions.

Circular Dichroism Intercalators. An explicit expression for the induced CD of an intercalated molecule can be derived (Appendix II), assuming an idealized intercalation complex with a planar ligand symmetrically inserted between two DNA base pairs with all three planes perpendicular to the DNA helix axis. Furthermore, if it may be assumed that intercalation occurs randomly among equivalent intercalation sites we arrive at a model in which the ICD of the ligand can be described by 1l, 12 RL = (1 -- 2cos 2 y)Df(VL, 1)i)

(6)

where RL is the ICD (more accurately the rotational strength) and D is the dipole strength (absorption intensity) of the ligand transition moment observed at frequency VL, coupling with the transition moments of DNA at frequency vi. The angle y is the angle between the transition moment of the ligand and the pseudo dyad axis (defined in Fig. 5B). The sign and amplitude of the f u n c t i o n f are determined solely by the properties of DNA and its magnitude is close to constant if the ligand transition occurs at a wavelength distant from the DNA absorption. From empirical observations 11 as well as from theoretical calculations 13 it has been concluded that the sign o f f is negative. It thus follows that the sign of ICD in a ligand with a transition moment located in the center of the intercalation site

l0 y. Matsuoka and B. Nordrn, Biopolymers 22, 1731 (1982). II p. E. Schipper, B. Nord6n, and E Tjemeld, Chem. Phys. Lett. 70, 17 (1980). I~ B. Nordrn and E Tjerneld, Biopolymers21, 1713 (1982). 13R. Lyng,T. Hard, and B. Nordrn, Biopolymers 26, 1327 (1987).

(A)

(a) lower base-pair

upper base-pair

pseudo dyad axis (C)

DNA'/ Helix axis

FIG. 5. Definition of parameters used in equations. (A) LD; (B) intercalator ICD; (C) groove-binding drug ICD.

[4]

L D AND C D OF DRUG-NUCLEIC ACID COMPLEXES

81

will be positive for transitions parallel to the pseudo dyad axis (y = 0) and negative for transitions perpendicular to it (y = 90°). From theoretical considerations a number of conclusions may be drawn. 13-15 First, themagnitudeoflCDforanintercalatoris small, generally Ae < 10M-Jcm -I at the maximum of the ICD spectrum observed. Second, ICD is dependent both on the orientation of the transition moment inside the intercalation site, y, and on the lateral displacement of the intercalator relative to the helix axis. Third, ICD of the intercalated drug depends on the type of base pairs forming the intercalation site, for example, whether the site is a 5'-purine-3'-pyrimidine or a 5'-pyrimidine-3'-purine step. For example, an intercalator centered on the helix axis of [poly(dG-dC)]2 with its long axis perpendicular to the pseudo dyad axis should have positive 1CD in both the 5'-G-3'-C and the 5'-C-3'-G steps. However, if its long axis is parallel to the pseudo dyad axis the ICD should be positive in the 5'-G-3'-C step and negative in the 5'-C-3'-G step, which may result in partial cancellation of the signal. Whereas ICD of a centrally located ligand may be interpreted in terms of its orientation, described by the angle y, it appears at present not possible to determine the exact position of an intercalator in its binding site from the amplitude and sign of ICD. The use of calculations thus seems limited to assessments of whether a particular interaction geometry is compatible with experimental data. Groove Binding. The magnitude and sign of the ICD of groove-bound ligands depend on the position and orientation of the ligand as well as on the nature of the binding site. For a minor groove-bound ligand with the transition moment oriented along the groove of B-form DNA, forming an angle near 45 ° with the bases, the ICD is expected to be positive. The ICD observed in the absorption band of a groove-bound drug is a result of interactions between the transition moments of the drug and the adjacent DNA bases. An approximate estimate of the CD signal can be obtained as CD ~ sin 2fl [(1/)~DNA - - 1/)~drug)-IF-2]

(7)

where/3 is the angle between the transition moment of the drug and the plane of the closest base pair (measured counterclockwise, as indicated in Fig. 5C). , ~ D N A and ~,drug are the wavelengths of the absorption maxima of the DNA bases and drug, respectively, and r is the distance between the drug and the base pair. Equation (7) is a simplification based on the assumption that all contributing transitions of adjacent bases can be represented by one effective transition moment positioned in the plane of the base pair closest in space and along a direction perpendicular to the line connecting the base pair and the drug as illustrated in Fig. 5C.

14R. Lyng, A. Rodger, and B. Nord6n, Biopolymers31, 1709 (1991). t5 R. Lyng, A. Rodger, and B. Nord6n, Biopolymers32, 1201 (1992).

82

BIOPHYSICALAPPROACHES

[4]

Drug-Dimer Interactions Many DNA binding drugs include aromatic ring systems and would thus have a tendency to form dimers in aqueous solution, were it not prohibited by repulsion between positive charges. Association with the negatively charged DNA neutralizes such repulsions and therefore, at high drug-DNA concentration ratios, effects of drug dimers can be observed. Drugs bound as dimers generally exhibit features that drastically differ from those of the monomers. The CD spectrum appears with characteristic exciton features, one positive and one negative peak around the absorption maximum. The intensities and the order of the two peaks are determined by the relative orientation of the drug chromophores. Also, the LD spectrum of a drug dimer may display features that originate from exciton coupling of the drug chromophores. The effects of dimer binding to DNA are illustrated by the example of cyanine dyes below, and a more detailed description of the analysis of exciton effects in LD and CD is given in Appendix III.

Calculations Analysis of theoretical calculations of ICD for groove binding compared with intercalation is complicated by the large number of potential binding sites. However, generally the ICD of a groove-bound ligand is one to two orders of magnitude larger than the ICD of an intercalated ligand. Accordingly, a large experimental ICD, well exceeding Ae = 10M -1 cm -l at the maximum of the ICD spectrum, is incompatible with intercalative binding.

Experimental Results LD is a useful technique for measuring the angle between the DNA helix axis and the transition moment of a groove-bound ligand. For a minor groove-bound ligand the molecular long axis typically forms an angle of about 45 ° relative to the helix axis. If the transition moment of the ligand significantly deviates from the molecular long axis a different angle ~ in Eq. (5) may result. Furthermore, a ligand bound in the major groove may adopt a wide range of orientations, which may result in an overall near-random orientation that does not give a measurable LD signal. Another general characteristic of groove-bound molecules is, as discussed above, a relatively strong ICD signal compared with intercalated ligands. Illustrative Examples of Linear Dichroism and Circular Dichroism Applications

Linear Dichroism for Discrimination between Intercalation and Groove Binding As an example illustrating the use of LD for discriminating between groove binding and intercalation we first discuss the binding of the molecule

[4]

LD AND CD OF DRUG-NUCLEICACID COMPLEXES

83

4',6-diamidino-2-phenylindole(DAPI) to DNA and synthetic polynucleotides. The long-axis polarized transition moment of DAPI (Fig. 3C) bound to mixed sequence DNA gives rise to a positive LD signal of about 350 nm, as shown in Fig. 6. Using Eq. (5) and the LD r value measured at 260 nm (-0.18), inserting ~ = 86 °, a value for the orientation factor S is obtained. The value of S (0.124) is then used in Eq. (5) in combination with the LD r measured for DAPI at 360 nm (0.09), which gives the angle ~ for DAPI of 45 °. This angle is consistent with binding of DAPI inserted in an edgewise manner into the minor groove of an undisturbed B-DNA duplex, as has been observed in a crystal structure of DAPI bound to a dodecamer. 16 Similar spectra are obtained when DNA of mixed sequence is replaced by poly(dA-dT). These results suggest that DAPI preferentially binds to AT-rich regions in DNA, as DAPI binding to AT-less sequences gives rise to markedly different LD spectra. As shown in Fig. 6B, the LD of DAPI bound to poly(dG-dC) is negative in the 350-nm absorption band. The different binding properties observed with different polynucleotides have been ascribed to steric blocking by the exocyclic amino group of guanines in the minor groove of poly(dG-dC), which is unfavorable for minor groove binding, and instead intercalative binding occurs. Substitution of the guanine for inosine bases (inosine is a guanine base without amino groups) restores the affinity of DAPI for binding to the minor groove, and changes the LD signal from negative to positive. 17 These conclusions based on LD data are further supported by CD experiments with mixed sequence DNA, synthetic polynucleotides, and oligonucleotides that have at least three or four AT base pairs. In all cases strong positive ICD at the DAPI absorption band is observed, also in agreement with minor groove binding. Monitoring Binding Rearrangements by Linear Dichroism LD is a particularly useful technique for detecting structural rearrangements in molecular interactions, for two reasons. First, the relatively quick data collection allows detection of events that occur on the time scale of seconds. Second, LD is highly sensitive to changes in the orientation of molecular chromophores. Structural effects may be seen to result from, for example, addition of drug to DNA, or addition of salt (titration). One example is the time course of binding of the electrostatically neutral anticancer drug cisplatin, cis-[Pt(NH3)2Cl2], to DNA. As a first step, prior to interaction with DNA, hydrolysis is believed to occur, resulting in the positively charged aquo species cis-[Pt(NH3)2(H20)2] 2+, which binds electrostatically to DNA, possibly by intercalation. Both of the associated water molecules are slowly (on a time scale of minutes) replaced by covalent bonds to nitrogens, primarily to guanine bases. One drug molecule can bind two adjacent DNA bases, and thereby force DNA to bend. The kink thus formed is believed to block transcription. 16Y. A. Larsen, D. S. Goodsell, D. Cascio, K. Grzeskowial,and R. E. Dickerson,J. Biomol. Struct. Dyn. 7, 477 (1989). t7 U. Sehlstedt, S. K. Kim, and B. Nord6n,J. Am. Chem. Soc. 115, 12258 (1993).

(A)

10 A

×5O i=\

8

0 LD

x 50

0.6

0.2 0 --0.2

-0.6

-1 0.2

I v

I

I

I

LD r

0.12 I I

0.04 0 -0.04

-0.12

-0.2 200

240

I 280 320 Wavelength

I 360

4~nm

FIG. 6. Absorption (A), linear dichroism (LD), and reduced linear dichroism (LD r) spectra of DAPI binding to DNA or polynucleotides. (A) DAPI-DNA complexes at concentration ratios of 0.01 (- - - ) and 0.05 (--). (B) Top: DAPI ( - - - ) , DAPI-poly(dG-dC) (--), and DAPl-poly(dG-dmSC) (.- .) complexes after subtraction of absorption from free polynucleotides; center and bottom: DAPl-poly (dG-dC) (--) and DAPI-poly(dG-dmSC) (...) complexes at DAPI-to-nucleotide concentration ratios of 0.10. A and LD were measured with optical path lengths of 10 and 1 mm, respectively.

84

(s) s~ ,

\,

0.3

L) t-

\

"\

.

",

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.

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340

380

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260

300

420

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-100

£3

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260

I

300

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t

340

W a v e l e n g t h (nm) FIG. 6.

(Continued)

85

I

380

i

I 420

86

BIOPHYSICALAPPROACHES

[41

LD has been used to monitor a slow conformational change in DNA caused by the model compound cis-[Pt(2,2'-bipyridyl)2(H20)2] 2+, which undergoes a transformation from intercalative binding to a covalent complex. Addition of the platinum complex to flow-oriented DNA results in an LD signal in the wavelength region 300-320 nm, where a transition directed along the molecular long axis of bipyridyl is responsible for absorption, as shown in Fig. 7.18 The LD signal is initially negative, with an amplitude that is consistent with intercalation, but shifts slowly to positive, suggesting a radical change of binding geometry. At the same time the LD signal of the DNA bases decreases markedly. These observations may be explained by intercalative binding that slowly switches into a covalent complex, as two water molecules are replaced by coordination of the platinum to DNA bases. The positive LD signal implies that the angle ot be smaller than 55 °, which agrees well with the ligand being coordinated to two adjacent bases on the same DNA strand, as found in a crystal structure of a related complex. 19 The decrease in the LD of the DNA bases shows that the change into a covalently bound complex is accompanied by a bend or other radical change of the DNA structure that disturbs the orientation of the DNA molecule. The initial negative LD signal was also used to assess the amount of drug bound in a series of varying concentrations, and to estimate a binding constant (Kb = 105 M-I), and binding density, which was found consistent with nearest neighbor exclusion for the initial intercalative binding. Another example of how LD can be used to monitor the time course of binding to DNA is the interaction of the carcinogen N-acetoxy-N-2-acetylaminofluorene (AAAF) with DNA. 2° The initial negative LD of the all in-plane polarized transition of the fluorene chromophore is slowly replaced by positive peaks as the carcinogen binds covalently to the DNA bases. It has therefore been concluded that AAAF, similar to the platinum complex, changes from initial intercalative binding to an orientation closer to parallel with the helix axis in the covalent adduct.

DNA Binding Cyanine Dyes Pseudoisocyanine (1,1'-diethyl-2,2'-cyanine, PIC) at low concentration binds to DNA by intercalation and gives rise to a negative peak in the LD spectrum at its absorption maximum, 540 nm, as indicated in Table I. 21 At intermediate PIC-to-DNA concentration ratios a positive peak appears at a shorter wavelength (490 nm) as a result of exciton interactions between closely bound PIC molecules. Further increase of the drug: DNA ratio results in polymerization of the 18B. Nordgn,FEBS Lett. 94, 204 (1978). 19R M. Takahara,A. C. Rosenzweig,C. A. Frederick, and S. J. Lippard,Nature (London) 377, 649 (1995). 20B. Nord6n,Biophys. Chem. 8, 385 (1978). 2J B. Norddnand F. Tjerneld, Biophys. Chem. 6, 31 (1977).

[4]

LD

AND

CD

OF DRUG-NUCLEIC ACID COMPLEXES

87

(A) A/cm

+1 0

-1 -2 -3 -4 -5 I

I

I

250

300

350

I

nm

(B)

A/cm 1

~~"'"..".

0

/,

5

2

.

•

5

~

.':",":" ~=~

0

LDxlO /cm -2 -4

'..~..",,o.i I

I

I

250

300

350

I

nm

FIG. 7. (A) Absorption (A) and linear dichroism (LD) spectra of DNA (3.7 10-4 M nucleotide) immediately after addition of [Pt(bipyridyl)2(H20) 2]2+at concentrations as indicated in units of 10-5 M. (B) Absorption (A) and linear dichroism (LD) spectra recorded 30 hr after addition of metal complex.

88

BIOPHYSICAL APPROACHES

[41

DNA-associated PIC, which in turn gives rise to a positive CD band at 553 nm. Similarly, as PIC is added to DNA the LD spectrum changes from negative, indicative of intercalation at low concentration ratios, to include a strong positive peak at 553 nm arising from a polymerized form of PIC (Table I). Site-Directed Linear Dichpvism

A complication often encountered in biological systems is spectral overlap among the different chromophores present. This can make it difficult to interpret LD data, but may be circumvented by strategic replacements of light-absorbing amino acids, which may allow measurement of the orientation of specific residues. This technique, termed site-directed linear dichroism, 22 was first developed for structural characterization of complexes between the Escherichia coli recombination protein RecA and DNA, fibrous structures not amenable to high-resolution studies by nuclear magnetic resource (NMR) or X-ray crystallography methods. The RecA protein binds to double-stranded DNA in the presence of an ATPlike cofactor (the nonhydrolyzable ATPvS), forming a helical fiber, for which the helical pitch and flow orientation properties [the orientation factor S in Eq. (5)] have been studied by small-angle neutron scattering. 23 From the intrinsic LD of DNA it was inferred that the nucleobases in this complex preferentially align with their planes perpendicular to the helix axis, just like in regular duplex DNA, despite a substantial increase in the contour length; DNA becomes elongated by some 50%. However, the LD spectrum arising from RecA absorption has contributions from several UV-absorbing residues of the protein and cannot be related to the orientation of any specific residue in the fibrous complex. To selectively record LD signals of individual residues a mutant of RecA was constructed, in which one of the two light-absorbant tryptophan residues present in wild-type RecA was replaced by UV-transparent threonine residue. The mutant was confirmed to be biologically active and to exhibit binding kinetics and overall hydrodynamic properties, when binding to DNA, that were equivalent to the unmodified (wildtype) form of the protein. Figure 8 shows the differential molar absorption AA = mmutant DNA--Awild-typeDNA with spectral features corresponding to the absorption of one stoichiometric equivalent of tryptophan. Also shown is the differential LD, ALD = LDmutantDNA- - LDwild-t y p e DNA,which analogously corresponds to the LD spectrum of the single replaced tryptophan residue. The LD spectrum is positive at the wavelengths of two of its near-UV in-plane transition moments (about 280 and 265 nm), and negative at shorter wavelengths (about 225 nm), where the absorption of another transition dominates (see Fig. 8). The corresponding differential

22 p. Hagmar, B. Nord6n, D. Baty, M. Chattier, and M. Takahashi, J. Mol. Biol. 226, 1193 (1992). 23 B. Nord6n, C. Elvingson, M. Kubista, B. Sjtiberg, H. Ryberg, M. Ryberg, K. Mortensen, and M. Takahashi, J. Mol. Biol. 226, 1175 (1992).

15

e

l

!i

0

P, .D

<

B -- t r a n s i t i o n

moments N 47 o

/

"Lo

(b)

2 x3 /.¢o,-

0

"~

.....

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--:

--4

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I

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-0-5

e

I 250

I 500

I 350

W0velength (nm) FIG. 8. (A) Differential absorption (AA), (B) differential linear dichroism (ALD), and (C) differential reduced linear dichroism (ALD/AA) of wild-type versus mutated RecA bound to double-stranded DNA (--) at a 3 : 1 ratio of DNA base pair : RecA, and differential linear dichroism of RecA bound to single-stranded DNA [poly(deA)] at a 3 : 1 ratio of DNA base : RecA (- - -), Transition moments in tryptophan chromophore: Lb (280 nm), La (265 nm), and B (225 nm).

90

BIOPHYSICAL APPROACHES

[4]

reduced dichroism can be obtained as ALD/AA. Assuming that the directions of the transition moments of 3-methylindole are applicable to the tryptophan chromophore, the orientation of the transition moments absorbing both at 265 and 225 nm can be determined, and were found to be 29 4- 5 ° (265-nm transition) and 61 4- 2 ° (225-nm transition), respectively, for Trp-291 of RecA bound to DNA. These angles define the orientation of the plane of the substituted tryptophan in the RecA-DNA complex. Several residues of RecA, such as phenylalanines and tyrosine, have been substituted for tryptophan, to give site-directed LD spectra of tryptophan in different positions within the protein. The results have been used to build a three-dimensional model of the RecA-DNA complex, based on the crystal structure of the protein alone, allowing detection of a certain conformational change, which may be induced by binding to the nucleic acid and the ATP cofactor.

(PNA)2 DNA Triplex Formation Monitored by Linear Dichtvism The synthetic DNA analog peptide nucleic acid (PNA) forms stable and specific complexes with natural nucleic acids by forming Watson-Crick complementary base pairs. In the PNA molecule the phosphate-sugar backbone has been replaced by pseudopeptide units that lack electrostatic charge, as well as chirality, and are structurally more flexible than DNA. PNA strands containing only thymine residues, PNA-Ts, bind strongly to poly adenine strands, forming highly stable triplexes through Watson-Crick and Hoogsteen hydrogen bonds. Such triplex formation was first observed using CD spectroscopy to monitor titration of poly(dA) with PNA- T8.24 The CD signal at 260 nm increases linearly as PNA-T8 is added to a stoichiometric T : A base ratio of 2 : 1 (Fig. 9A). From corresponding LD measurements it was seen that the orientation of the PNA-DNA complex increases slowly at low PNA : DNA concentration ratios, but grows drastically as the PNA : DNA ratio approached 2 : 1 (Fig. 9B). This LD result shows that the PNA 8-mers preferentially form stretches of triplex with the DNA strand, leaving flexible naked regions of poly(dA-dT), rather than as a first step forming a continuous duplex, to which PNA binds in a second step to complete the triplex. lnducedHandednessinNonchiraIPeptide-NucleicAcidDuplex. A PNA strand can base pair with another PNA strand of Watson-Crick complementary sequence to form a stable duplex. If the PNA duplex adopts a helical structure, given the lack of chirality of the PNA molecule, an equal mixture of right- and left-handed helices would result, and their respective CD spectra would exactly cancel and give no detectable signal. 25 However, in short PNA duplexes where one of the strands has a D-lysine attached to its C terminus, a CD spectrum can be observed in the 24 S. K. Kim, P. Nielsen, M. Egholm, O. Buchardt, R. H. Berg, and B. Nord6n, .1. Am. Chem. Soc. 115, 6477 (1993). z5 M. Eriksson and R E. Nielsen, Nat. Struct. Biol. 3, 410 (1996).

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LD AND CD OF DRUG-NUCLEIC ACID COMPLEXES

91

i I Q

I

CD 0

%°°

N

\

.C

\ 0 C~

,

I

I 'T

'7.

(ore} 0 7

I

I

I

I

o °

~ ~

(Bep tu) 0 1 ~

el

Z

n

-

e~ .~ .= .=

o

N~

~ ~.~-

N~ r...) <

u~

92

BIOPHYSICAL APPROACHES

[4]

6

2

-

-4

-

D,L~A

....

L-Lys-TG

....

D-Lys-TG

-'6

210 220 230 240 250 260 270 280 290 300 310 320 Wavelength {rim)

FIG. 10. CD spectra of the 10-mer PNA H-AGTGATACTAC-(L/D)-Lys-NH2 (thick/thin curves) and H-GTAGATCAGT-(L/D)-Lys-NH2 (dotted/broken curves).

nucleobase absorption band, evidence of helical stacking of the bases. Exchange for an L-lysine results in the mirror image CD spectrum, indicating that the handedness of the helix is reversed (Fig. 10). The amplitude of the CD spectrum increases as the length of the PNA duplex grows, reaching a limit of 10-12 base pairs, after which the conformation appears no longer to be governed from the amino acid attached to the terminus of one of the PNA strands. 26 Conclusions LD and CD, like other optical techniques, offer possibilities for rapid characterization of nucleic acid complexes using relatively small amounts of sample. The information that may be gained varies with the level of sophistication at which the measurements are carried out and interpreted. We briefly summarize here some practical applications of increasing complexity. 1. Detection of Interaction LD can be used to detect binding of a ligand to flow-oriented DNA (or RNA) as a signal in the absorption region of the ligand. Such LD is proof that the ligand indeed interacts with the polynucleotide. Correspondingly, an ICD signal seen in the absorption band of a nonchiral ligand demonstrates interaction with a chiral species (DNA or RNA), but less sensitively than LD. 26E Wittung, M. Eriksson, R. Lyng, P. Nielsen, and B. Nord6n, J. Am. Chem. Soc. 117, 10167 (1995).

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LD AND CD OF DRUG-NUCLEIC ACID COMPLEXES

93

2. Detection o[ Drug-Drug Interactions A weak ICD signal observed in the absorption band, and of the same shape as the absorption spectrum, seen from a nonchiral ligand bound to DNA, is characteristic of the ligand binding as a monomer. This ICD signal is gradually replaced by a strong bisignate exciton CD feature if ligand dimers (or higher order complexes) are formed.

3. Structure Information from Linear Dichroism Linear dichroism can provide insights regarding the orientation of molecules bound to flow-oriented DNA. Intercalators may clearly be distinguished from groove binders and the effects of ligands on the DNA, such as bends or kinks, or elongation may be assessed. Similarly, the impact of proteins bound to DNA may be estimated from LD, possibly by strategic substitution of bases by analogs that allow specific detection. Furthermore, a technique specially developed for structure studies of DNAbound protein complexes that are not amenable to crystallography or NMR methods is the so-called site-directed linear dichroism. By including mutations that direct the light-absorbing properties to selected residues in the protein, the differential LD spectra obtained from mutant and wild-type protein can be used to conclude the orientation of the chosen residue. From experiments carried out on a set of systematically designed mutants, a model of the protein-nucleic acid complex structure may be produced.

4. Structure Information from Circular Dichroism The sign of the induced CD (ICD) of an intercalator depends on its orientation inside the intercalation pocket. In contrast to intercalators, the ICD of ligands bound in the minor groove of B-DNA is strong and always positive, provided the observed transition moments are polarized along the groove. The CD spectrum of the nucleic acid itself, or of a DNA- or RNA-bound protein, is sensitive to conformational changes that result from the interaction. The complexity of CD effects, however, does not allow interpretations in structural terms as directly as LD techniques. Appendix I

Linear Dichroism We here give an outline for quantitative evaluation of transition moment directions from stretched film measurements. Unlike the flow orientation accomplished in the Couette cell, the orientation in a stretched polymer film can be treated as uniaxial. Uniaxial means that the orientation is symmetric around one axis (the stretch direction). Equation (I. 1) applies to the general case of reduced LD of a

94

BIOPHYSICAL APPROACHES

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uniaxial sample27: LDr(~.) = 3[Szzez()V) + SyyeyO0 + SxxexOQ]/[g,~(iv) + ey()V) + ex(*k)]

(I. 1)

where ez(X), ey()V), and e~(~) are the extinction coefficients (in units of M -1 cm J) for linearly polarized light, hypothetically measured with the plane of polarization (the electric field of light) parallel with the z, y, and x axes, respectively, of an orthogonal coordinate system attached to the molecule (as explained below, this coordinate system may not be arbitrarily chosen). Szz, Syy, and S,x are the so-called order parameters for the molecular axes z, y, and x. Their sum is zero, so in fact there are only two independent orientation parameters. If the molecule behaves like a cylindrical rod with its long axis, z, oriented along the stretch direction, then S= = 1, and Syy = Sxx = --0.5. However, generally molecules do not behave in this idealized fashion. For example, 2,7-diazapyrene (Fig. 3A) shows roughly the orientation pattern S= = 0.3, Syy = 0, and Sxx = - 0 . 3 , where z is the long axis, y is the in-plane short axis, and x is the axis perpendicular to the plane of the molecule. This indicates the convention for assigning the three coordinate axes in molecules that possess high symmetry, the case we consider first. The interpretation of an LD spectrum is done in three steps. 1. Assign tentatively the wavelength, X., where of the spectrum displays the highest value of LD r as corresponding to a transition polarized parallel to the long axis, z, in the molecule and assume that its absorption does not overlap with any differently polarized intensity [i.e., ey()~,.) and ex(~.z) are both zero]. Equation (I. 1) is thus simplified to LDr(X:) = 3S.:, which yields S=. 2. Now we consider the minimum value of LD r, possibly negative, and use 2,7-diazapyrene (Fig. 3A) as an example. If the point of minimum corresponds to a band with weak isotropic absorption that shows strongly negative LD r, typically 1, we are probably dealing with an out-of-plane polarized n --+ zr* transition. This provides us with a value of Sxx according to LDr()vx) = 3&x. However, generally we are dealing with only in-plane polarized rr --+ Jr* transitions, so the minimum LD r, which is positive or only weakly negative, instead provides us with Syy according to LDr(~y) = 3Syy. In case the absorption bands overlap completely, one must apply a trial-and-error iterative procedure or simulate the overlapping bands, again using Eq. (I.1). 3. Once Szz, Syy, and Sxx have been determined (and in the case of a symmetric molecule, also the transition moment directions assigned--they can only be directed along either of the three axes x, y, or z) we may use Eq. (I. 1) to determine the shape of the polarized spectra, e_.O-) and ey(~.), in the molecular frame, and then also the degree of overlap between the two spectra e.()v) + ey(X). The -

27 B. Nord6n, Appl. Spectrosc. Rev. 14, 157 (1978).

[4]

LD AND CD OF DRUG-NUCLEIC ACID COMPLEXES

95

sum of ezO.) and E'y(~) should be equal to the isotropic absorption according to Aiso = dc[ez()O + ey(~.)]/3, where d is the optical path length and c is the concentration. From Eq (I. 1) we thus obtain ez(~.) = Aiso[LDr(~.)

-

3Syy]/(Szz - Syy)dc

ey(L) = Aiso[LDr()Q - 3Sz~]/(Syy - S~z)dc In case the molecule is planar but lacks other elements of symmetry, the situation is more complicated because the orientation axis, z, of the molecule is not obvious from symmetry. If only in-plane (xy-plane) transitions exist and their absorptions do not overlap, Eq. (I. 1) can be rewritten as LDr(~-) = 3(Szz cos 2 0i +

Syy sin 2 0i)

(1.2)

Where Oi is the angle between transition moment i and the orientation axis z in the molecular plane. The angle Oi may be obtained by trial and error, the largest value of LD r corresponding to the smallest angle (in case the transition is polarized parallel with the z axis Oi = 0 °, and when perpendicular Oi = 90 °, which also determines the values of Szz and Syy, respectively). However, generally transitions overlap, and in such cases LDr()Q in Eq. (I.2) must be replaced by (LDr)i, the specific reduced linear dichroism of transition i. The total (experimental) LD r, which includes overlap between the absorptions of different transitions, is analyzed by the following expression: LDtrot(X) = Zei(X)(LDr)i/Eei(X)

(1.3)

Alternative to measuring LD one may measure All and A± separately. LD r may then be determined by trial and error, using (LDF)i = 3 ( d i - 1 ) / ( d i + 2), where di is the coefficient that makes a specific spectral feature, corresponding uniquely to transition i, vanish by linear combination, Art - di A±. 4

A p p e n d i x II

Induced Circular Dichroism We here present a theoretical framework for induced CD in a nonchiral chromophore arising from interaction of its electric dipole 'allowed transitions with the electric dipole allowed transitions of a chirai host. CD involves, in addition to an electric transition moment in the molecule, also a magnetic transition moment. The magnitude of the induced molecular circular dichroism is measured as its rotatory strength, R. Interaction between two identical transition moments of equal energies (the degenerate case) will cause a characteristic circular dichroism pattern consisting of one positive and one negative contribution of equal rotatory

96

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strengths that are proportional to the energy of the transition. This is, for example, the case of a drug dimer bound to DNA, the topic of Appendix III. The CD spectrum observed in this instance has a characteristic bisignate shape, resulting from the superposition of the positive and negative contributions, with an energy difference that increases with the magnitude of the interaction energy, V, between the two transition moments. Such spectra are often referred to as exciton CD spectra. By contrast to the bisignate exciton CD, the shape of the circular dichroism spectrum induced in a nonchiral guest molecule is in general identical to the corresponding apsorption spectrum, with the proviso that it may be either positive or negative. The rotatory strength is proportional to a pairwise sum of contributions of scalar triple products: R L = ( 1 / 4 c ) ~ ]~Ca.kCb,lrkl(/~Oa k X ~t~Obl)

(II.1)

with each term providing the contribution of a given pair of transitions to the overall rotational strength. Here c is the velocity of light,/Z0a k and/Z0b 1 are the electric transition dipole moments for transitions 0 --+ a and 0 --+ b in the interacting chromophores k and l, respectively, and rkl is the radius vector from k to 1. In firstorder perturbation theory the weighting factor Ca,kCb, I is proportional to Va.k;b,I, the subunit interaction potential. For many practical purposes, such as when rkj is large compared with bond lengths, the ICD of a transition 0 --+ b of the ligand L can be approximated as R(bL) = [VLVi/2zreoh(v 2 -

U2)]y~,Wa,k;b,LrkL(#0ak X #0(bL))

(II.2)

where V = (/Z0ak • /Z0bL)/r~L -- 3(/~0a k " rkL)(/Z0b L • rkL)/r~L

(II.3)

An important property, incorporated in Eq. (II.2), is the dependence of ICD on where in the spectrum the ligand transition appears, as given by the factor VLVi/(v 2 -- V2). Thus the magnitude of CD induced in a ligand transition should decrease approximately as the inverse of the separation in frequency from the position of the effective inducing DNA transition, (v 2 - v2) 1 From Eq. (II.2) it follows that the outcome of calculations will not be strongly affected by replacement of the unknown vacuum-UV transition of the DNA bases by an isotropic sum of background oscillators. Under special circumstances when the ligand and DNA transition frequencies are far apart (rE << I)i) it may even be possible to calculate ligand ICD by assuming that the ligand transition moment couples with just one suitable defined effective DNA transition (one vi), which is assumed in the model behind Eq. (6).

[4]

LD AND CD OF DRUG-NUCLEICACID COMPLEXES

97

A p p e n d i x III Exciton Circular and Lineal" Dichroism When drug molecules bind to DNA as dimers (or oligomers) their absorption spectra are generally different from their spectra as monomers, as a result of exciton coupling. From the CD and LD exciton spectra one may deduce information about the geometry of such dimers. If the dimer is chiral, which is generally the case when formed onto a chiral template like DNA, the CD spectrum will display a characteristic bisignate "couplet" consisting of one positive and one negative CD band of equal intensity positioned at each side of the absorption maximum of the monomer. The sign and magnitude of the CD (the rotational strength R) are given by R

=

]A excit°n • m excit°n =

CrAB • (/~A × /-tB)

(III. 1)

where ~ e x c i t o n and m excit°n a r e , respectively, the effective electric and magnetic transition moments of the dimer resulting from interactions of the individual transition moments /~A and/zB of the two chromophore units A and B of the dimer. C is a constant, and tAB is a vector connecting the two chromophore units A and B. In Eq. (Ill. 1) the "dot" means scalar product and may be replaced by cosine of the angle that tAB forms with a vector perpendicular to the plane of the vectors /~A and/ZB, and the "cross" may be replaced by sinus of the angle between the latter two vectors. The energy splitting between the two exciton bands is given by the Coulombic interaction between the two transition dipoles/~A and/~B: V : ( ~ A " /AB)/r3B -- 3(P'A ' rAB)(~B • r A B ) / r 5 B

(III.2)

The interaction energy, V, thus decreases with the third power of the distance between the chromophores and generally the exciton effect of dimers separated by more than l0 ,~ can be neglected. Equations (III.1) and (III.2) both contain information about how the two monomer units are arranged relative to each other. Equation (III.1) also, by the sign of the sinus dependence, contains information about the absolute configuration of the chiral dimer. Linear dichroism may also provide structure information about dimers. The angles between the helix axis and the two exciton transition moments may be calculated using Eq. (5). These exciton transitions are obtained from the vector sum and vector difference between the transition moments of the two monomers: #exciton in-phase = [1/(21/2)]{/_tA + /_tB } #exciton out-of-phase = [ 1/(21/2)]

{/.tA

-- ]-tB }

(III.3) (IliA)

The CD and LD spectra from a DNA-bound dimer may be qualitatively analyzed by exciton theory, without complicated calculations, as follows. As a hypothetical example let us assume that one drug unit be intercalated with its long axis parallel

98

BIOPHYSICALAPPROACHES

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to the long axis of the intercalation pocket and a second drug is located in the minor groove, pointing at an angle of 45 ° to the long-axis of the first drug unit (this corresponds to the geometry in Fig. 5C, substituting the DNA base pair in Fig. 5C by an intercalated ligand). The transition moment is assumed to be directed along the long axis of the drug (as indicated by the arrows in Fig. 5C). The analysis is then performed as follows. 1. Draw transition m o m e n t s (ILLA and/~B) of each drug unit as single-headed arrows with the heads arbitrarily oriented. Let them, for example, be directed closer to parallel than antiparallel to each other. According to Eq. (III.2) this corresponds to repulsive orientation, that is, V > 0 (sin 45 ° > 0). If the arrows instead are antiparallel, that is, forming an angle of 135 °, we have an attractive interaction (sin 135 ° < 0). Repulsive orientation corresponds to the exciton couplet seen at the high energy end of the spectrum, that is, the band at the short-wavelength side of the absorption maximum of the monomer drug absorption peak. For right-handed DNA this component should display a positive CD and a negative LD, which may be deduced as follows. 2. LD is determined by the resultant vector of the two vectors (~A + ~B) in Eq. (III.3), in this case a vector bisecting the angle between the two drug units, that is, symmetrically in between 90 and 45 ° to the helix axis. We thus obtain the angle 67.5 °, and according to Eq. (5) we thus conclude a negative LD for the short-wavelength band. For the case of oppositely directed arrows--the attractive coupling, that is, the exciton band at longer wavelength than the monomer--one instead finds the vector (]..La -- ~ B ) oriented at 157.5 ° relative to the helix axis, and thus displaying a positive LD signal. 3. To deduce the sign of the CD, we again consider the resultants of the two /ZA and/~B vectros, and start with the in-phase interaction observed at high energy, lt~A -]- jt~B. CD is positive if any component of the magnetic moment created by circulation of charge around this direction is parallel, and negative if the magnetic moment is antiparallel to this direction. With the anticipated arrangement in the groove of a right-handed DNA molecule, we find that/~A and/~a will create an anticlockwise rotation of charge around the vector ~A +/ZB. Using the right hand and pointing with the index finger in the direction of the current, the thumb will point in the direction of the magnetic moment. Because/ZA + ~B and m are thus antiparallel, the CD signal will be negative. Correspondingly, for the out-of-phase component, (/~A --/~B) and m will be parallel, so the CD at the long-wavelength side will show positive intensity. So with the dimer bound to right-handed DNA in the fashion assumed, a bisignate exciton CD spectrum should appear having a negative peak at short wavelength and a positive peak at long wavelength.

[5]

RAPID SCREENING OF LIGAND-DNA BINDING

99

[5] Rapid Screening of Structurally Selective Ligand Binding to Nucleic Acids B y JINSONG REN and JONATHAN B. CHAIRES

Introduction DNA is polymorphic, and exists in a variety of secondary and tertiary structures that are dictated by both sequence and environmental conditions.1 Unique DNA structures represent potential targets for small molecules, and provide a promising new avenue for drug development. 2,3 Small molecules that selectively bind to a particular structure could interfere with whatever biological function involves that structure. For example, multistranded triplex and tetraplex nucleic acid structures have attracted considerable attention as targets for small molecule. 2 6 Attempts to rationally design small molecules that bind selectively to a particular structure are hampered by the lack of a rapid and convenient assay for structural selectivity. Typically, thermal denaturation studies monitored by UV absorbance are used, but such an assay is both time consuming and incapable of screening a large number of different structures simultaneously. In addition, melting curves in the presence of bound ligand are often complex and multiphasic, 7-9 making the analysis and interpretation of data difficult. Alternatively, binding constants for the interaction of ligands with the target structure can be measured and compared with standard duplex DNA, but such measurements are even more time consuming and tedious than are those involved in obtaining melting curves. To circumvent these difficulties, we have devised a novel competition dialysis assay for rapidly screening ligand binding to a variety of nucleic acid structures) °- 12 Practical details of the assay are described here, along with examples of experimental data.

I S. Neidle, "Oxford Handbook of Nucleic Acid Structure." Oxford University Press, New York, 1999. 2 j. L. Mergny and C. Helene, Nat. Med. 4, 1366 (1998). 3 T. C. Jenkins, Curl: Med. Chem. 7, 99 (2000). 4 j. L. Mergny,G. Duval-Valentin,C. H. Nguyen, L. Perrouault,B. Faucon, M. Rougee, T. MontenayGarestier, and E. Bisagni, Science 256, 1681 (1992). 5 L. H. Hurley and E L. Boyd, Trends Pharmacol. Sci. 9, 402 (1988). 6 L. H. Hurley, J. Med. Chem. 32, 2027 (1989). 7 L. Cai, L. Chen, S. Raghavan, R. Ratliff, R. Moyzis, and A. Rich, Nucleic Acids Res. 26, 4696 (1998). 8 D. M. Crothers, Biopolymers 10, 2147 (1971). 9 j. D. McGhee, Biopolymers 15, 1345 (1976). l0 j. Ren and J. B. Chaires, Biochemistry 38, 16067 (1999). 11j. Ren and J. B. Chaires, J. Am. Chem. Soc. 122, 424 (2000). 12j. Ren, C. Bailly, and J. B. Chaires, FEBS Lett. 470, 355 (2000).

METHODS 1N ENZYMOLOGY, VOL. 340

Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. 0076-6879/00 $35.00

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Principle of Method and Historical Background The competition dialysis assay is based on the fundamental thermodynamic principle of equilibrium dialysis.13 The approach is easily summarized. A macromolecule solution is placed inside a semipermeable dialysis membrane with pore sizes that allow small molecules to pass through, but which retain the large macromolecules. The dialysis unit is then suspended in a solution containing small molecule ligand. At equilibrium, the chemical potential of the free ligand must be equal inside and outside the dialysis unit, and any excess ligand on the macromolecule side of the membrane may be attributed to binding to the macromolecule. Equilibrium dialysis has been widely used to measure the binding of small molecules or ions to macromolecules to provide the primary data for the determination of accurate binding constants.13.14 Muller and Crothers first introduced an important variant of the equilibrium dialysis experiment, the competition dialysis procedure.15 That procedure was first used to evaluate the base specificity of drug-DNA interactions. The procedure utilized a three-part dialysis chamber, in which two dialysis membranes separated the central chamber from two outside chambers. Two DNA samples of identical concentration but of differing base composition were placed in each of the two outer chambers. A ligand solution was placed in the central chamber, and allowed to equilibrate among the three chambers. If the ligand bound selectively to AT or GC base pairs, more ligand would accumulate in the chamber containing the DNA with the base composition containing the highest percentage of the preferred base. Rather detailed quantitative inferences concerning the nature of the preferred binding site are possible by this method using simple probability concepts.15'16 This version of the competition dialysis method unfortunately saw little widespread application, and was supplanted by footprinting methods that provide a higher resolution glimpse into sequence selective binding. Competition dialysis was, however, adapted to provide a tool for probing structural selective ligand binding to nucleic acids. Becker and Dervan used the approach to show selective binding of bis(methidum)spermine to a D N A - R N A hybrid. 17 Chaires and coworkers used the method as a probe for ligand binding to left- and right-handed DNAJ s'19 More recently, the method was used to probe ligand binding to duplex and triplex DNA. 2° The assay described here is a simple and straightforward extension of the Crothers competition dialysis method. Instead of a three-chambered dialysis 13K. E. vanHolde, W. C. Johnson, and R S. Ho, "Principles of Physical Biochemistry."Prentice Hall, Upper Saddle River, New Jersey, 1998. 14I. M. Klotz, "Ligand Receptor Energetics." John Wiley & Sons, New York, 1997. 15W. Muller and D. M. Crothers, Eur. ,l. Biochem. 54, 267 (1975). 16j. B. Chaires, in "Advances in DNA Sequence Specific Agents," Vol. l, pp. 3-23. (L. H. Hurley, ed.). JAI Press, Greenwich, Connecticut, 1992. 17M. M. Becker and E B. Dervan, J. Am. Chem. Soc. 101, 3664 (1979).

[5]

RAPID SCREENING OF L I G A N D - - D N A BINDING

10 1

lyzer g DNA

I#M Free tigand

Stir bar FIG. 1. Schematic diagram of the experimental arrangement for the competition dialysis assay. Disposable dialysis units (either Spectrum DispoDialyzers or Pierce Slide-A-Lyzers) are used to contain the desired nucleic acid structures. Structures are dialyzed against a common test ligand solution to reach equilibrium. The amount of ligand bound to each structure is then measured by absorbance spectroscopy or fluorescence.

apparatus, disposable dialysis units are used to contain a wide variety of nucleic acid structures at identical concentrations. These units are simply placed into a beaker containing ligand solution, as illustrated in Fig. 1. At equilibrium, the free ligand concentration is identical throughout the system, and preferential binding by a particular structure leads to a greater accumulation of total ligand within that dialysis unit. Structural selectivity can be easily measured by measuring total ligand concentration within each dialysis unit. The challenge to be met is to find a suitable buffer in which the nucleic acid structures of interest are all stable. In the first-generation assay, l° 13 structures were found to be stable in a simple phosphate buffer containing 200 mM NaCI. The assay was expanded to include 19 structures in the second-generation assay l~ using the same buffer conditions. Description of these structures and a detailed protocol for the assay follow.

18 j. B. Chaires, Biochemistry 24, 7479 (1985). 19 S. Satyanarayana, J. C. Dabrowiak, and J. B. Chaires, Biochemistry 32, 2573 (1993). 2(I I. Haq, J. E. Ladbury, B. Z. Chowdhry, and T. C. Jenkins, J. Ant. Chem. Soc. 118, 10693 (1996).

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Experimental Method

Overview of Assay The competition dialysis assay is straightforward and simple. First, stock solutions of the structures of interest are prepared and characterized. Second, the assay is set up for the study of a ligand of interest by filling disposable dialysis units with solutions of the various structures at identical concentrations, placing these in a common ligand solution, and allowing the system to reach dialysis equilibrium. Third, concentrations of total ligand within each dialysis unit are determined by absorbance or fluorescence spectroscopy (or any other convenient analytical tool). Finally, results are graphed for analysis and interpretation.

Materials and Sample Preparation Dialysis Units. The first-generation assay used 1.0- or 0.5-ml DispoDialyzer units (Spectrum Medical Industries, Rancho Domingneg, CA; www.spectrumlabs. com) of the desired molecular weight cutoff (MWCO), usually 3500 MWCO. Subsequently, a less expensive and overall more suitable alternative was found, Pierce Slide-A-Lyzer MINI dialyzer units (Pierce, Rockford, IL, www.piercenet.com). Pierce proved to be helpful and cooperative in providing technical information about their product, and is the recommended source for dialysis units for the assay. The quality of each minidialysis unit is evaluated before experiments by dialysis against water to ensure against leaks. In that test, 200 #1 of water is added to each dialysis unit, which are then dialyzed against water for 24 hr. The volume of solution within each dialysis unit is then measured. Units with excess volumes of water that exceed the original 200/zl should be discarded. Dialysis units may be reused several times after careful rinsing with buffer solution. Nucleic Acids. The nucleic acid structures used in the current version of the competition dialysis assay are shown schematically in Fig. 2 and listed in Table I. All samples are dissolved in BPES buffer, consisting of 6 mM NazHPO4, 2 mM NaHzPO4, 1 mM NazEDTA, 0.185 M NaC1 (pH 7.0). Samples of DNA from ClostHdium perfringens, calf. thymus, and Micrococcus lysodeikticus are purchased from Sigma (St. Louis, MO), and are sonicated, phenol extracted, and purified as previously described. 21 Poly(dA), poly(dT), poly(U), poly(dC), poly(dA)-poly(dT), poly(dA-dT), and poly(dG-dC) are purchased from Pharmacia Biotech (Piscataway, N J). Poly(rA) and poly(A)-poly(U) are purchased from Sigma. Deoxyoligonucleotides 5'-T2G20T2, 5'-G10TaGI0, and 5'-AG3(TTAG3)3 are purchased from Research Genetics (Huntsville, AL). Synthetic single-stranded and duplex polynucleotides are used without further purification. The poly(rA)-poly(dT) DNA-RNA hybrid is prepared by mixing 21j. B. Chaires, N. Dattagupta,and D. M. Crothers,Biochemistry21, 3933 (1982).

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1

103

Form

Samples

Single-strand

poly dA, poly dT poly A, poly U

Duplex

natural DNA, Z-DNA RNA, DNA:RNA hybrid, polynucleotides,

Triplex

poly dA:[poly dTL (DNA) polyA:[poly U]2 (RNA)

Tetraplex

[T2G2oT]. A(G3I-IA)3G3 [G,oT4G,o]. [Poly dC].

(Parallel) (Folded) (G-wire) (i-motif)

FIG. 2. Schematic of structures used in the current generation of the competition dialysis assay.

poly(rA) and poly(dT) in a 1 : 1 molar ratio, heating to 90 °, and slowly cooling to room temperature. The DNA and RNA triplex forms are prepared by mixing poly(dA)-poly(dT) with poly(dT) [or poly(A)-poly(U) with poly(U)] in a 1:1 molar ratio, heating to 90 °, and slowly cooling to room temperature. Tetraplex DNA and i-motif DNA are prepared by heating the oligonucleotide 5'-T2G20T2, 5'-Gl0TaGl0, 5'-A(G3TTA)3G3, or poly(dC) to 90 ° for 2 min, slowly cooling to room temperature and then equilibrating for 48 hr at 4 ° before use. Left-handed Z-DNA is prepared by bromination of poly(dG-dC) as previously describedY Concentration Determinations. Concentrations of nucleic acid samples are determined by UV absorbance measurements, using the extinction coefficients and absorption maxima listed in Table I. Stock solutions of nucleic acids are prepared and maintained at 75/zM concentration, using the monomeric unit of each polynucleotide as the concentration standard: nucleotides (nt) for singlestranded forms, base pairs (bp) for duplex forms, triplets for triplex forms, and quartets for tetraplex forms. The extinction coefficients listed in Table I all refer to these concentration standards. Sample Quality Control. All nucleic acid preparations are characterized by circular dichroism, using a JASCO (Tokyo, Japan) J500A instrument, and by thermal 22 A. Moiler, A. Nordheim, S. A. Kozlowski, D. J. Patel, and A. Rich, Biochemistry 23, 54 (1984).

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TABLE I NUCLEIC ACID CONFORMATION AND SAMPLES USED IN COMPETITION DIALYSIS EXPERIMENTSa

Conformation

DNA/oligonucleotide

Single-strandpurine

Poly (dA) Poly(A) Single-strandpyrimidine Poly(dT) Poly(U) Duplex DNA CIostridium po~[Hngens (31% GC) Calf thymus (42% GC)

DNA-RNAh y b r i d Duplex RNA Z-DNAb Triplex DNA Triplex RNA Tetraplex DNA 1 Tetraplex DNA 2 Tetraplex DNA 3 i-motif

X(nm) E (M Icm I)

T~,~('C)

%H

---

257 258 264 260 260

8,600 9,800 8,520 9,350 12,476

-82.5

1.24 1.29 --1.28

Miclvcoccus lysodeikticus

260 260

12,824 13,846

85.5 > 100

1.36 1.16

(72% GC) Poly(dA)-poly(dT) Poly(dA-dT) Poly(dG-dC) Poly(rA)-poly(dT) Poly(A)-poly(U) Br-poly(dG-dC) Poly(dA)-[poly(dT)]2 Poly(A)[poly(U)]2 (5'-T2G20T2)4 5'-A(G3TTA)3G~ 5'-(Gl0T4GI0)x [Poly(dC)]4

260 262 254 260 260 254 260 260 260 260 260 274

12,000 13,200 16,800 12,460 14,280 16,060 17,200 17,840 39,267 73,000 39,400 27,200

75.2 67.8 > 100 70.9 62.5 > 100 42.5 62.5 89.0 66.8 89.0 46.9; 55.0

1.58 1.53 1.49 1.54 1.59 1.67 1.06 1.04 1.02 1.38

'~x, Wavelength;e, molar extinctioncoefficient;Tin,meltingtemperature;%H, hyperchromicity. ~'A.Moiler,A. Nordheim, S. J. Kozloski,and A. Rich, Biochemistry23, 56 (1984).

denaturation monitored by UV absorbance. Denaturation studies are done with a Varian (Palo Alto, CA) Cary 3E spectrophotometer equipped with a Peltier temperature control unit. Charateristic melting temperatures and hyperchromicity values are listed in Table I. Spectra and melting curves for all samples are available as supplementary materials to published articles.l°' 11

Competition Dialysis Protocol For each competition dialysis assay, 400 ml of the dialysate solution containing 1 /zM test ligand concentration is placed into a beaker. A volume of 180 #1 (at 75 # M monomeric unit) of each of the DNA samples listed in Table I is pipetted into a separate Slide-A-Lyzer MINI dialysis unit with 7000 M W C O membrane. All 19 dialysis units are placed in a MINI dialysis flotation device (Pierce) and then the whole unit is placed in the beaker containing the dialysate solution. The beaker is covered with Parafilm, wrapped in foil, and allowed to equilibrate with continuous stirring for 24 hr at room temperature (20-22°). At the end of the

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equilibration period, DNA samples are carefully collected in the comer of the MINI dialysis unit and transferred to microcentrifuge tubes. The DNA samples are then made to a final concentration of 1% (w/v) sodium dodecyl sulfate (SDS) by the addition of appropriate volumes of a 10% (w/v) stock solution. The total concentration of drug (Ct) with each MINI dialysis unit is then determined spectrophotometrically, using wavelengths and extinction appropriate for each ligand. An appropriate correction for the slight dilution of the same resulting from the addition of the stock SDS solution is made. The free ligand concentration (C0 is determined spectrophotometrically with an aliquot of the dialysate solution, although its concentration usually does not vary appreciably from the initial 1 #M concentration. The amount of bound drug is determined as Cb = Ct - C f . Data are plotted as a bar graph, using Origin software (version 5.1 ; Microcal, Northampton, MA) for plotting and analysis. In the initial experiments, and on occasion thereafter, the competition dialysis method is checked by a detailed mass-balance accounting of total ligand, using the procedures described in detail by Klotz.]4

Results and Interpretation Figure 3 shows results obtained with the standard intercalator ethidium. Resuits from both the first-generation (Fig. 3A) and second-generation (Fig. 3B) assays are shown. Additional structures were included in the second-generation assay. These results compare binding to 19 different structures. No binding is observed to any single-stranded DNA or RNA forms [poly(A), poly(U), poly(dA), and poly(dT)]. Duplex DNA, both from natural sources or synthetic polynucleotides,

M tyso DNA CT DNA

M lyso DNA CT DNA C perf DNA polydT polydA polyU polyA

C perl DNA poly dA poly dT 0

3

6

[Bound]~M

9

o

3

6

[Bound]pM

FIG.3. Resultsobtainedforthe simpleintercalatorethidium,usingthe first-generationassay(A) that included 13differentnucleicacid structures,or the expandedsecond-generationassay(B) that included the 19 structures listed in TableI.

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-3

0

C ¢- Cr0NA b'--b

S

4

[5]

•

3

4J

Cb.

FIG. 4. Alternate graphical representations of experimental data. Data obtained by the secondgeneration assay for ethidium (Fig. 3B) are replotted in alternate forms. (A) The difference in the amount bound relative to the amount bound to duplex calf thymus (CT) DNA is shown. (B) The difference relative to the arithmetic mean bound to all structural forms is shown.

binds ethidium well. Poly(dA)-poly(dT), however, binds ethidium only weakly, a result that arises from the unusual structure adopted by the polymer in solution. 23 25 RNA [poly(A)-poly(U)] and the DNA-RNA hybrid (poly(rA)-poly(dT) show the most ethidium binding, a finding consistent with more laborious titration binding studies. 26 Left-handed Z-DNA shows only weak binding, although ethidium can aliosterically convert the polynucleotide to an intercalated right-handed form at higher-ligand concentrations, z7 DNA and RNA triplex forms bind ethidium about as well as duplex forms, consistent with equilibrium binding studies. 28 Finally, ethidium binds weakly to G-tetraplex structures, and not at all to the unusual i-motif. The results of Fig. 3 provide a rapid comparison of ethidium binding to 19 different structures, and provide a sound basis for selecting particular structures for more detailed biophysical or biochemical studies. Figure 4 shows two of many possible alternate graphical representations of competition dialysis data, using the results obtained with ethidium in the secondgeneration assay (Fig. 3B). In Fig. 4A, the difference in the amount of ethidium bound to a particular structure relative to that bound to calf thymus DNA is shown. 23 S. M. He, Y. C. Sun, S. L. Wei, C. B. Wu, E U. Wang, S. D. Wang, and J. X. Xie, Yao Hsueh Hsueh Pao 28, 859 (1993). 24 j. E. Herrera and J. B. Chaires, Biochemistry 28, 1993 (1989). 25 S. S. Chan, K. J. Breslauer, M. E. Hogan, O. J. Kessler, R. H. Austin, J. Ojemann, J. M. Passner, and N. C. Wiles, Biochemistry 29, 6161 (1990). 26 j. L. Bresloff and D. M. Crothers, Biochemistry 20, 3547 (1981). 27 G. T. Walker, M. P. Stone, and T. R. Krugh, Biochemistry 24, 7462 (1985). 28 p. V. Scaria and R. H. Shafer, J. Biol. Chem. 266, 5417 (1991).

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Calf thymus DNA represents a standard duplex DNA, so this plot emphasizes differences relative to that standard. Positive values of the difference in the amount bound clearly indicate preferential binding relative to calf thymus DNA, while negative values indicate poorer binding. Figure 4B shows data plotted as the difference from the mean value of binding to all structural forms. In that plot, the difference is relative to the simple arithmetic mean, with positive values indicative of preferential binding. Other manipulations of the data are possible and may be desirable on occasion. Perhaps most useful is calculation of the apparent binding constant, Kapp, which is simply given by the equation Cb Kapp -- Cf x [NA]total

where Cb and Cf are the bound and free ligand concentrations, respectively, and [NA]total is the total nucleic acid concentration. Typically, in our experimental setup, Cf = 1 # M and [NA]total = 75/.zM. Cb is the experimentally measured amount of ligand bound to each form. This treatment assumes that potential binding sites are in excess, so possible neighbor exclusion effects can be neglected. The units of Kapp would refer to the monomeric unit (nucleotide, base pair, etc.) of each polymer. Values of Kapp determined from competition dialysis were found to be in excellent agreement with binding constants obtained from more complete titration studies.12 The primary utility and the power of the competition dialysis assay are illustrated in Fig. 5, in which the structural selectivity of four different compounds is compared. The compounds are 9-aminoacridine, quinacrine, methylene blue, and telomerase inhibitor V (Fig. 5D). At a glance, the distinct structural preferences of these compounds are evident. 9-Aminoacridine (Fig. 5A) is a well-known intercalator, and shows the expected binding to duplex DNA structures. It has a strong preference, however, for triplex DNA and binds well to the parallel-stranded, "G-wire" tetraplex structure (tetraplex 3). Quinacrine (Fig. 5B) shows an even stronger preference for triplex DNA. Methylene blue (Fig. 5C) shows a strong preference for multistranded DNA structures, and binds well to all tetraplex forms and to triplex DNA. Finally, telomerase inhibitor V shows a similar preference for multistranded structures, but in contrast shows appreciable binding to certain duplex forms, poly(dA-dT) and the DNA-RNA hybrid. Telomerase inhibitor V is one of the few compounds assayed that binds appreciably to single-stranded forms, in this case poly(A). The few examples shown here are the tip of a rather large iceberg of experimental results. We have efficiently and rapidly screened more than 150 compounds to date by the competition dialysis assay. The results obtained with these compounds provide a comprehensive view of structural selectivity from which principles will emerge that should serve to guide the rational design process for

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~ CI~

2

4

[Bound] g M

s

e

v

CH~ NHCI~(CH2)3N(C2HJz OCH3 -x~

o

3

s

O

3

IS [Bound]

v

9

9

12

12

g M

FIG. 5. Competition dialysis results obtained with (A) 9-aminoacridine, (B) quinacrine, (C) methylene blue, and (D) telomerase inhibitor V (2,6-bis[3-(N-piperidino)propionamido]anthracene9,10-dione; Calbiochem, La Jolla, CA).

new small molecules that can target specific structures. So far, detailed results on several compounds have been published,l°-12 and descriptions of many more are in preparation. We hope that the description of the protocol presented here will allow other laboratories to implement the method as a routine part of their studies of drug-nucleic acid interactions. Acknowledgment This work was supported by grant CA35635 from the National Cancer Institute.

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toward high-order nucleic acid system. The purpose of this chapter has been (1) to illustrate the importance and power of calorimetry as a biophysical tool, and (2) to outline certain practical considerations in the use of ITC and DSC methods for DNA triplex/tetraplex stability and binding studies. It must be stressed that, if used properly and within limits, calorimetry represents a powerful and versatile array of techniques that can yield great insight into the energetics of biomolecular stability and binding processes. The genuine strength of calorimetry becomes apparent when it is used in conjunction with parallel techniques such as UV-visible/CD/fluorescence spectrophotometry, NMR spectroscopy, X-ray crystallography, and stopped-flow kinetic methods. Such a multitechnique approach to studies of DNA-drug interactions (or indeed any binding interaction) can provide a comprehensive and cohesive picture for the underlying biomolecular events. Acknowledgments We thank Ms. Maria Halla (Universityof Greenwich) for conducting all the DSC experiments with the G2 tetraplex. Financial support for the authors' laboratories is generouslyprovidedby the Universityof Sheffield(to I.H.)and YorkshireCancerResearch(to T.C.J.)and Universityof Greenwich (to BZC). I.H. is an EPSRC AdvancedResearchFellow.

[7] Volume Changes Accompanying Interaction of Ligands with Nucleic Acids By LUIS A. MARKY, DONALD W. KUPKE, and BESIK I. KANKIA Introduction The biological function of nucleic acids is carried out through interactions with other molecules, specifically with proteins. To completely understand the expression of genes, it is imperative to obtain a complete description of the structure, thermodynamics, and kinetics of these interacting systems. In the specific case of the energetics of protein-nucleic acid complex formation, few thermodynamic investigations have been carried out because of the limited solubility of proteins. Alternatively, the energetic contributions of the specific molecular interactions observed in the structure of the protein-nucleic acid complex can be obtained by measuring complete thermodynamic profiles of the interaction of small molecules with nucleic acids. At the same time, some of these small molecules, called ligands or "drugs," interact with nucleic acids to prevent replication and transcription. The extent of their biological activity depends on several factors, such as binding affinity, specificity, and binding mode. The literature is full of reports on the molecular

METHODS IN ENZYMOLOGY, VOL. 340

Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. 0076-6879/00 $35.00

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forces involved in these types of association reactions, but little is known of the hydration effects that accompany these reactions or the association of two smaller solutes. We are finding answers to the following questions: how does water hydrate a small or large solute? What are the physical properties of water around charged, polar, or nonpolar groups? The measurement of the volume change, AV, for association reactions with a proper interpretation usually yields information about these hydration effects. Water plays a fundamental role in the stability of the secondary and tertiary structures of proteins and nucleic acids. 1.2 In particular, the nucleic acid molecule is well hydrated and its hydration depends on the nucleic acid conformation and composition.l'3 7 The hydration properties provide a rationale for the conformational plasticity (A, B, or Z) of DNA; for example, slight changes of the water activity can shift the equilibrium to favor one duplex conformation over others.I At the present time, the measurement of hydration changes has been achieved with the development of high-sensitivity density techniques, which make it possible to determine the change in volume of samples containing 1 mg or less of solute. 8'9 However, the interpretation of results remains sometimes difficult because of the existence of two types of water that are hydrating a particular macromolecule or complex: water of electrostriction around charged and polar groups and hydrophobic water around nonpolar or organic groups. For example, a release of water molecules is expected to be measured in the formation of a simple complex AB from the reaction of its component A and B solutes and to a first approximation, which is due to the overlap of hydration shells of the solutes in the complex. In biochemical systems at constant temperature and pressure, the volume expansions and contractions attending reactive chemical changes reflect the average change in the molar volume of water in the vicinity of the participating solutes. The solutes themselves ordinarily do not possess the propensity to change the solution volume to the same order of magnitude (a relatively large net difference in the amount of void spaces within globular macromolecules perhaps being an exception). Liquid water by virtue of its quadrupolar nature [and possessing a large dipole moment of 1.85 Debye (D) units, or more in the liquid phase] has the capacity to change its molar volume dramatically. The open, quasi-tetrahedral structure of bulk water I W. Saenger, "Principles of Nucleic Acid Structure." Springer-Verlag, New York, 1984. 2 C. Tanford, Adv. Protein Chem. 23, 121 (1968). 3 M.-J. B. Tunis and J. E. Hearst, Biopolymers 6, 1325 (1968). 4 L. A. Marky and D. W. Kupke, Biochemistry 28, 9982 (1989). 5 E. Westhof, Annu. Rev. Phys. Chem. 17, 125 (1988). 6 V. A. Buckin, B. I. Kankiya, N. V. Bulichov, A. V. Lebedev, V. P. Gukovsky, A. P. Sarvazyan, and A. R. Williams, Nature (London) 340, 321 (1989). 7 W. A. Buckin, B. I. Kankiya, A. P. Sarvazyan, and H. Uedaira, Nucleic Acids Res. 17, 4189 (1989). 8 G. T. Gillies and D. W. Kupke, Rev. Sci. lnstrum. 59, 307 (1988). 9 D. W. Kupke and J. W. Beams, Methods Enzymol. 26, 74 (1972).

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is some 50% larger in molar volume than if the individual molecules were packed in a noninteracting hexagonal array.l° Accordingly, ion-water dipole interactions are likely to be the major sources of biochemical volume changes (at constant temperature and pressure). The intense electric field close to an ion induces a local collapse of the bulk water structure by compressing the adjacent water dipoles at pressures corresponding to several thousand atmospheres, conditions that may even reduce the molar volume of water to "-~12 ml/mol, lO This compression, which depends on the charge and radius of an ion and on the local dielectric gradient, is known colloquially as electrostriction, j°'l] Hence, neutralization reactions, formation of ion pairs (salt bridges), and the coordination of metal ions result in water decompressions that can give rise to relatively large expansions at millimolar reactant concentrations. 12 The changes in molar volume of water molecules interacting over much shorter effective distances (dipole~lipole, hydrophobic-dipole, partial charge-dipole, etc.) are less well understood. According to electrostriction theory, we would expect these compressions to be comparatively small. Also, it is not clear whether the interaction of organic groups with water can in some cases cause a small expansion relative to bulk water. ]3 The volume effect of hydrophobic moieties on charged solutes is known to augment the contractions 14'15; however, unless these moieties undergo chemical changes during mixing of aqueous phases, the difference in volume from this source should contribute only a marginal effect. The electrostriction effect for hydrogen bonding in nucleic acid base pairing, between the partially charged nitrogen and oxygen donor/acceptor atoms, is predicted to be small because the contraction is approximately proportional to the square of the charge density. 16 Moreover, the volume change attending the loss of hydrogen-bonding water to these sites prior to the formation of a duplex would tend to compensate for the putative contraction accompanying hydrogen bond formation between the bases. It appears, therefore, that an uptake of water molecules is observed in the formation of a nucleic acid duplex, from the mixing of its complementary strands. 17,18This is because the resulting volume change is interpreted to reflect the compensating effects that take place between changes in electrostriction of the duplex water quadrupoles and changes of hydrophobically bound water of the single strands. 10B. E. Conway,"Ionic Hydration in Chemistry and Biophysics."Elsevier, New York, 1981. ] I E J. Millero, in "Waterand AqueousSolutions"(R. A. Home, ed.), p. 519. Wiley-lnterscience,New York, 1972. 12D. W. Kupke and B. S. Shanck,,I. Phys. Chem. 93, 2101 (1989). 13E H. Stillinger, Science 209, 451 (1980). 14E. J. King,J. Phys. Chem. 73, 1220 (1969). ]5 j. E. Desnoyers,Phys. Chem. Liq. 7, 63 (1977). 1rE J. Millero, Chem. Rev. 71,147 (1971). 17K. Zieba, T. M. Chu, D. W. Kupke, and L. A. Marky, Biochemistly 30, 8018 (1991). 18L. A. Marky and D. W. Kupke, Methods Enzymol. 323, 419 (2000).

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In this chapter, we initially provide a brief theoretical and experimental description of current techniques for the measurement of volume changes; we then illustrate the applicability of this approach for the interaction of ligands with nucleic acids. We provide specific examples from data obtained in our laboratories; these include ligands with different physical binding modes, the contribution of nucleic acid conformation, bent deoxyoligonucleotides, and the formation of oligomer duplexes with chemically attached ligands. M e a s u r i n g V o l u m e C h a n g e for A s s o c i a t i o n R e a c t i o n There are several techniques that are currently in use for the measurement of the volume change, 2xV, for an association reaction. Model-dependent techniques are based on the dependence of the equilibrium constant on hydrostatic pressure; model-independent techniques include direct measurements by dilatometry and indirect measurements of the mass and density of reactants and products of a particular reaction.

Dependence of K on Pressure The volume change is determined by the following relationship: A V = (O A G /

Op)T, substitution of AG(---- -RTlnK) yields AV = -RT(O In K/Op)T, where R is the gas constant and T is the temperature. Thus, the volume change is obtained from the dependence of the equilibrium constant, K, on hydrostatic pressure and may be monitored by optical techniques. The measurement involves the placement of a sample solution in a quartz cell contained in an optical high-pressure cell. The concentration of reactants and products is measured optically, using standard absorption or fluorescence techniques, as a function of hydrostatic pressure in the range of 1 to 3000 atm ( t 0 . 1 - 3 0 0 MPa). In this method, it is assumed that the AV is independent of pressure, that is, (OAV/Op)T equal to zero. This is equivalent to an optical measurement of model-dependent enthalpies, A HuH, using the van't Hoff equation AH~n = RT2(OIn K/OT)p, in which it is assumed that AH~n is independent of temperature or ACp is zero.

Dilatometry Dilatometry is a straightforward method for A V measurements. In dilatometry, ~ 6 ml of solution of each reactant is placed in each arm of an inverted "Y" glass tube with arms widely separated; a water-immiscible liquid, for example, heptane, proved to be inert, is placed above the reactants to separate them initially and fill up the entire tube. This tube is closed with a capillary tube and the level of the organic liquid is within the linear scale of the capillary in millimeters. The whole tube is placed in a water bath with a temperature control of 4- 0.001 ° and the height of the capillary is read after reaching temperature equilibrium. Then, the reactants

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are mixed by moving the dilatometer sideways without removing it from the bath. The height of the liquid in the capillary is read again and the difference in this height, negative or positive, is proportional to the extent of the volume contraction or expansion of the particular reaction. The main disadvantage of this technique is the large amount of materials needed for each independent measurement.

Indirect Methods The AV value is obtained from the measurements of the mass and the equilibrium density of solutions before and after mixing. The masses are obtained with a microbalance, and the densities are obtained with a magnetic suspension densimeter or with an Anton Paar (Graz Austria) densimeter. The observed change in volume, Av, on adding reactant A to reactant B is given by A1) = Pproducts -- 1)reactants = mmix/Pmix -- (mA/PA -}- mB/PB)

(1)

where m is the mass in grams and p is the density of the solution in grams per milliliter. The sum of the two terms within parentheses gives the initial volume before mixing. The value of Av in milliliters is then normalized per mole of the limiting reagent to give AV. Our laboratories use this procedure extensively; therefore, we describe in the following sections the techniques in use for the density measurements.

Magnetic Suspension Densimetry The instrument used in this study is an improved version of the earlier design for biochemical purposes by Senterl9and has been previously described, s This instrument requires only 100 #1 per measurement, and its sensitivity is such that it can deliver volume differences with a precision of less than half a nanoliter. This densimeter is calibrated with aqueous KC1 solutions of known density. The density of each sample is obtained by relating the measured voltage to the straightline calibration equation of voltage versus density. This is because the electrical current required to stabilize the tiny permanent magnet jacketed at a present height below the meniscus is directly proportional to the density of the surrounding fluid. 9 According to Eq. (1), small weighing errors have no appreciable effect on A v and the three density values, while independent, need not be of high absolute accuracy because it is their differences that are required for A v. The density is measured with a precision of 5 × 10 - 6 g/ml. The temperature is always kept within 4-0.001 °. In these experiments, the concentration of nucleic acid samples (in phosphate) ranges from 2 to 3 mM; thus, the effect of solute-solute interactions on the volume property is deemed to be negligible. 2° Usually, equal volumes of solutions of each reactant 19j. p. Senter, Rev.Sci. lnstrum. 40, 334 (1969). 20R. S. Berry, S. A. Rice, and J. Ross, "Physical Chemistry." Oxford University Press, New York, 2000.

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are mixed to give about 300 #1 of the final solution in 0.4-ml polyethylene tubes to prevent evaporation and to allow for rinsings, and the mean value of three readings is taken for A v. Three of these solutions are normally studied. The molar volume change is obtained by dividing A v by the number of moles of the limiting reagent. Oscillating M e t h o d

In the oscillating method the density is measured with an Anton Paar densimeter (DMA 60) with two microcells (DMA 602). Each glass cell is a U-shaped tube with a volume of 0.2 ml, and is electromagnetically forced into oscillation. To reduce the influence of temperature fluctuations, a differential method is used in which two identical cells are connected to a common thermostat. The sample cell is filled with the sample solution while the reference cell is filled with solvent of equal composition. The density is obtained by measuring the period of oscillation, T, of the solution (or gas), using the relationship p = A ( T 2 - B ) , where A and B are constants determined from a calibration curve. This instrument is calibrated by using the densities and periods of water and air, or solutions of known densities as in the case of the magnetic suspension densimeter. The precision in the density is 4- 5 × 10 6 g/ml. The investigator should be cautious in using this instrument for density determinations of viscous macromolecule solutions; it is best to reduce the viscosity of these solutions by sonication. M o l e c u l a r I n t e r p r e t a t i o n of V o l u m e C h a n g e To understand the physical significance of the volume change of an association reaction, it is best to discuss initially the apparent molar volume, qbV, of a solute molecule. This is defined as the volume difference of a solution relative to that of the solvent 11,21: • V = ( V - n l V l ) / n 2 = Vm -]-

AVh

(2)

where V is the volume of a solution that contains nl mole of the solvent (water) and n2 moles of solute; VI is the molar volume of pure solvent and is in turn equal to the sum of two terms, that is, the fight-hand side of Eq. (2), which includes Vm (the intrinsic molar volume of the solute that is inaccessible to the surrounding solvent) and AVh (the hydration contribution), which is the volume change of water around the solute molecule as a result of solute-water interactions, and the void volume between the solute molecule and the surrounding water (the so-called thermal volume because it is due to thermally induced molecular vibrations). 22'23 21 D. W. Kupke, in "Physical Principles and Techniques of Protein Chemistry" (S. J. Leach, ed.), p. 1. Academic Press, New York, 1973. 22 S. Terasawa, H. Itsuki, and S. Arakawa, J. Phys. Chem. 79, 2345 (1975). 23 D. E Kharakoz, J. Sol. Chem. 21, 569 (1992).

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In general, the majority of solutes have positive Vmvalues and negative AVh values. For most solutes, the sum of the magnitudes of the intrinsic and thermal volumes is greater than the magnitude of AVh, which leads to positive values of the apparent volume.11,24,25 The few exceptions are some metal ions with negative ~ V values because of their small size and strong hydration contribution.1 l The volume changes accompanying the binding of a ligand molecule to a DNA molecule can be expressed in the following way: AV = AVm + AAVh, where AVm is the change in the intrinsic volume of both DNA and ligand (normally negligible) and A A Vh is the hydration contributions of the interacting molecules, DNA and ligand molecules. The AVm of association reactions involving nucleic acids without conformational changes is normally considered equal to zero, that is, the intrinsic volume of a nucleic acid molecule remains the same. Therefore, A V is simply equal to the volume changes due to the rearrangements of water molecules in the hydration shells of the interacting molecules. Because hydrating water has a lower molar volume than bulk water at room temperature, a positive value of AV indicates an expansion of the system or release of water molecules from hydration shells while a negative value of A V is indicative of a system contraction or uptake of water molecules. Another parameter that can be estimated quantitatively from A V measurements is the actual number of water molecules involved in a reaction, but only if the type of the participating water is known. In most cases, this analysis is complicated because during DNA-ligand interaction different types of atomic groups are involved, including charged, polar, and nonpolar groups. These groups have characteristic hydration states involving two types of water molecules that have different molar volumes. However, a lower estimate of the number of water molecules, N, participating in a particular association reaction can be obtained from the following relationship: N = A A V h / ( V E - Vw); where VE and Vw are the molar volumes of electrostricted and bulk water, respectively. In using this equation, the apparent molar volume of electrostricted water is considered equal to 15.5 ml/mo116,18 and the assumption is made that the apparent molar volume of structural water is higher than this value. Volume Effects on Interaction of Ligands with DNA a s F u n c t i o n of B i n d i n g M o d e A small ligand interacts with a nucleic acid double helix in different physical binding modes, and by physicochemical attachment. Some of these binding 24 H. Hoiland, in "Thermodynamic Data for Biochemistry and Biotechnology" (H.-J. Hinz, ed.), p. 17. Springer-Verlag, Berlin, 1986. 25 H. Durchschlag, in "Thermodynamic Data for Biochemistry and Biotechnology" (H.-J. Hinz, ed.), p. 45. Springer-Verlag, Berlin, 1986.

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modes includes the following: outside the helix, where the complex is stabilized by electrostatic interactions between the positive charges of the ligand and the negatively charged phosphate groups of the nucleic acid; by intercalation of planar aromatic rings between adjacent base pairs, which are stabilized mainly by van der Waals and electrostatic interactions; and by geometric complementarity with the grooves, the complex is stabilized by a combination of electrostatic, van der Waals, and hydrogen-bonding interactions. Typical examples include the electrostatic binding of Mg 2÷ ions, the intercalators ethidium and propidium, and the minor groove ligand netropsin. In this section, we discuss the volume effects that take place in the binding of these ligands to calf thymus DNA (CTDNA). In addition, we show the volume effects for the interaction of actinomycin D with CT-DNA, which is an example of both intercalative and minor groove binder. Electrostatic Interaction o f m g 2+ Ions

Mg 2+ plays essential roles in biology by stabilizing the secondary structure of DNA, the tertiary folding of RNA molecules, and protein-nucleic acid complexes. 1,26 In general, metal ions are well hydrated, particularly small cations such as Mg2+. 11'27 DNA molecules are also strongly hydrated I and it has been shown that among the DNA conformations the B form has the strongest hydration. 1,28 Therefore, the interaction of Mg 2+ with DNA should result in strong hydration effects if their hydration layers overlap. Consistent with this expectation, we obtained a AV of 22 ml/mol of bound Mg 2+ to CT-DNA (see Table I). The positive value indicates an overall release of water molecules, which is consistent with earlier measurements of 27 ml/mol at lower ionic strengths. 29'3° This comparison at different ionic strengths also confirms the electrostatic nature of Mg 2+ binding. Because in the binding of 1 mol of Mg 2+ there is a release of 2 mol of Na + ions, the resulting AV value also includes the hydration contribution for this exchange of Na + with Mg 2+ ions in the ionic atmosphere of DNA. 31 However, DNA-condensed sodium ions keep their hydration layers intact and the hydration contribution from the release of Na + may be considered negligible. X-Ray structural data of MgZ+-oligonucleotide crystals have indicated that Mg 2+ ion binds in the major groove of DNA, keeping intact its first hydration layer. 32 Previous results from our laboratory have shown that Mg 2+ binding is somewhat specific to 26V. K. Misra and D. E. Draper, Biopolymers 48, 113 (1999). 27E J. Millero, G. K. Ward, E K. Lepple, and E. V. Hoff,J. Phys. Chem. 78, 1636 (1974). 28D. Rentzeperis, D. W. Kupke, and L. A. Marky,Biopolymers 33, 117 (1993). 29R. M. Clement, J. Sturm, and M. P. Daune, Biopo(ymers 12, 405 (1973). 3oB. I. Kankia, Biophys. Chem. 84, 227 (2000). 31 V. A. Buckin, H. Tran, V. Morozov,and L. A. Marky,J. Am. Chem. Soc. 118, 7033 (1996). 32L. McFail-Isom,X. Shui, and L. D. Williams, Biochemistry 37, 17105 (1998).

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DNA by forming inner-sphere complexes with dG. dC base pairs, 33 justifying the bending of GC sequences induced by Mg 2+. We arrive at the conclusion that the measured dehydration effects correspond to a partial removal of water molecules from the hydration shells of both Mg 2+ and DNA atomic groups. Furthermore, we estimate a removal of approximately nine water molecules in this interaction by dividing AV ( = 22 ml/mol) by the overall molar compression of electrostricted water of 2.5 ml/mol obtained earlier. 18,27 Intemalative Binding o f Ethidium and Propidium

Ethidium is a classic intercalator that has been investigated extensively and is commonly used in biotechnology as a fluorescent dye to characterize nucleic acid molecules in gel electrophoresis experiments. This charged molecule has a planar phenanthroline ring that intercalates into DNA base pairs with a neighbor exclusion parameter of 0.5 base pair, while propidium, a similar molecule, possesses an extra charged side chain (see structures in Fig. 1). Table I lists the resulting AVvalues for the interaction of these ligands with DNA at two different [ligand]/[DNA] ratios in the intercalative mode. These ratios correspond to two different slopes of the heat (and ultrasonic velocity) versus ligand-DNA titrations curves prior to saturation of DNA by intercalation (data not shown). All A V values are negative, corresponding to an uptake of water molecules; the values for ethidium are consistent with earlier dilatometric measurements. 34 However, two features should be noted about our volume effects: at the smaller degree of binding, the uptake of water molecules for each drug is larger; and propidium shows a larger uptake of water molecules. It is difficult to interpret unambiguously the resulting volume effects of intercalation. Several hydration contributions are involved in the formation of the intercalative complex. Some of these contributions include (1) the release of water due to electrostatic interactions of the positive charges of the ligands and the negative phosphates of DNA, (2) the hydrophobic release of water from the ligands prior to intercalation, and (3) the hydrophobic uptake of water after the intercalation of the phenanthroline ring due to the unwinding and lengthening of DNA, which also gives a higher exposure of organic groups to the solvent. The net effect is that the uptake of water of the last contribution overrides the release of water of the first two contributions. This is consistent with the higher uptake of water molecules at the smaller degree of binding, in which larger changes of the DNA structure takes place, and with the presence of heat capacity effects that have been measured in the binding of these ligands to CT-DNA. 35'36 33W.A. Buckin,B. I. Kankiya,D. Rentzeperis,and L. A. Marky,J. Am. Chem. Soc. 116,9423 (1994). 34E Delben,E Quadrifoglio,V. Giancotti, and V. Crescenzi,Biopolymers 21, 331 (1982). 35j. Ren, T. C. Jenkins, and J. B. Chaires, Biochemistry 39, 8439 (2000). 36A. M. Soto, C. Dziowgo,and L. A. Marky, in "12th Annual Gibbs Conferenceon Biothermodynamics," Book of Abstracts, p. 98. Carbondale, IL, 1998.

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H2N~~~

?=.:,

(~

NH2

H

C2H5

MeVal I Sar

MeVal "-7 I Sar

o

Pro

Pro

o

D-Val

D-Val

I

I Thr , NH I CO

I

I'--Thr ,

NH I CO [r~N'~~

'

NH2 N~'N('CH2)3N+(C2H5)H3 2C

Propidium

I

I

~

~

Ethidium

I

2

[71

-,~'NH2 H2N NH HN

NH2

CH3

,Y'N CH3

CH3

Actinomycin D

H2N

O

"~'O

CH3

Netropsin

FIG. 1. Chemical structures of ethidium, propidium, actinomycin D, and netropsin.

The overall uptake of water molecules is higher for propidium and this may be due to its extra quaternary amino charged group. If it is assumed that both ligands form similar structures with DNA, the differential AV is --9 ml/mol, which corresponds to an uptake of about four water molecules. Minor Groove Binding A classic groove binder is the dicationic oligopeptide netropsin (see Fig. 1). Netropsin binds with high affinity and high specificity to the minor groove of 4-5 dA- dT base pairs of DNA (for references see Marky and Kupke4); it actually penetrates down to the floor of this groove. This complex is stabilized by electrostatic, van der Waals interactions of the ligand with the sugar-phosphate backbone, and by hydrogen bonding between the amide protons of netropsin with N-3 of adenine and 0-2 of thymine facing the floor of the minor groove. Binding affinities drop

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TABLE I VOLUME EFFECTS ACCOMPANYINGINTERACTIONOF LIGANDS WITHCALF THYMUSDNAa Ligand Mg 2+ Ethidium Propidium Netropsin Actinomycin D

AV (ml/mol) 22 - 3l - 12I' -40 -21 h 1 16

Binding mode Electrostatic Intercalation (0.20) Intercalation (0.50) Intercalation (0.18) Intercalation (0.38) Minor groove Intercalation and grooves

"All values determined at 20 ° in 10 mM sodium HEPES, pH 7.5. Values in parentheses are the final [ligand]/[DNA] ratios used in measurements. t, These AV values have been measured at the indicated [ligand]/[DNA] ratios but corrected for the initial binding of ligands.

quite significantly with the increase in GC content of the DNA 37 because the NH2 of guanine blocks by steric hindrance the deep penetration of the ligand in the minor groove. In the case of CT-DNA, which has "-~40% GC content, netropsin binds to this polymer with a binding affinity of ~105. The observed volume effect is close to zero (see Table I) and consistent with netropsin lying in or partially penetrating the minor groove. The release of water molecules, from the formation of ion pairs, cancels the immobilization of water by the partial exposure of the netropsin and/or van der Waals interactions (see below for the volume effects of the binding of netropsin to oligomers containing AT sequences). Intercalation and Minor Groove Binding Actinomycin D is an antiobiotic commonly used clinically in the treatment of rhabdomyosarcoma and Wilms' tumor in children (for references see JaresErijman e t a/.38). It consists of a planar phenoxazone chromophore and two pentapeptide cyclic rings of similar sequence (see Fig. 1). The pharmacological action of actinomycin D is generally attributed to its tight and specific binding to DNA, which results in the inhibition of transcription elongation by blocking the RNA polymerase. Binding to DNA takes place by intercalation of the phenoxazone chromophore and by the formation of hydrogen bonds between specific groups 37 D. Rentzeperis, L. A. Marky, and D. W. Kupke, Biopolymers 32, 1065 (1992). 38 E. A. Jares-Erijman, R. Klement, R. Machinek, R. M. Wadkins, L. A. Marky, B. I. Kankia, and T. M. Jovin, Nucleosides Nucleotides 16, 661 (1997).

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of the pentapeptide cyclic rings and the nucleic bases facing the minor groove of the DNA duplex. 39 This ligand is not charged; therefore, the hydration effects would be attributed to the uptake of water from the dissociation of actimomycin D aggregates prior to binding and from the intercalation event, and to the release of water from hydrogen bonding and from the physical placement of the hydrophobic peptide chains of the ligand on the minor groove of DNA. Inspection of Table I shows a positive AV, or release of water. To a first approximation, this can be explained as a cancellation of the opposing contributions from the dissociation of aggregates and the intercalation event with the physical placement of the peptide groups. What remains is the hydrogen-bonding term, which allows for the formation of tighter complexes and the release of more water molecules. The volume effect for the formation of one hydrogen bond between polar atomic groups has been estimated previously to be ~ 4 ml/mol. 23 Dividing our AV of 16 ml/mol by this number presents the formation of four hydrogen bonds between actinomycin D molecule and CT-DNA. I n t e r c a l a t i o n of E t h i d i u m a n d P r o p i d i u m w i t h RNA P o l y n u c l e o t i d e s To include the role of nucleic acid conformation, we reported the volume effects for the interaction of ethidium and propidium with poly(rA), poly(rU) and poly(rA), and the results are compared with those presented for CT-DNA. A DNA helix adopts the B conformation and poly(rA)-poly(rU) adopts the A conformation, whereas poly(rA) forms a single-stranded helix (due to the stacking of adenines) in the A-like conformation. Table II lists the resulting AV values for the intercalation of these ligands with poly(rA) • poly(rU) and at two different [ligand]/[DNA] ratios. Similar to the results with DNA, the AV values are negative and indicate an uptake of water molecules. Also, at the smaller degree of binding, the uptake of water molecules for each drug is higher and propidium shows the most uptake of water molecules. All the hydration contributions mentioned above for the intercalative binding to DNA hold for RNA. Relative to CT-DNA and at the lower degree of binding, the water uptake of ethidium and propidium binding with RNA is somewhat smaller, whereas it is marginally larger at the higher degree of binding. The lower uptake of water may be explained in terms of the larger helical radius for RNA, which induces a better shielding of the phenanthroline ring from the solvent. This explanation does not take into account the smaller hydration of the free RNA polynucleotide and any contributions from differences in the nucleotide sequence. The AV values for ethidium and propidium binding to poly(rA) are both positive (see Table II). This means that the hemiintercalation of the ligands, between 39 T. R. Krugh, Proc. Natl. Acad. Sci. U.S.A. 69, 1911 (1972).

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TABLE I1 VOLUME EFFECTS ACCOMPANYINGINTERACTIONOF LIGANDSWITHPOLYRIBONUCLEOTIDESa Ligand

AV (ml/mol)

Binding mode

Poly(rA)opoly(rU) Ethidium Propidium

-24 - 15~ -31 - 2 6 t'

Intercalation Intercalation Intercalation Intercalation

(0.25) (0.52) (0.20) (0.45)

Poly(rA) Ethidium Propidium

13 17

Hemintercalation Hemintercalation

a All values determined at 20 ° in 10 mM sodium HEPES, pH 7.5. Values in parentheses are the final [ligand]/[DNA] ratios used in the measurements. b These AV values have been measured at the indicated [ligand]/[DNA] ratios but corrected for the initial binding of ligands.

two adjacent adenines, is accompanied by a release of water molecules. The singlestranded poly(rA) helix does not have grooves (which are normally well hydrated in the duplex state) and the phenanthroline ring is partially buried; therefore, the volume effect reflects primarily the water release of electrostatic interactions between negatively charged phosphates of the nucleic acid and the positively charged atomic groups of the ligands. B i n d i n g of N e t r o p s i n to M o d e l B e n t S e q u e n c e s We studied the interaction of netropsin with a set of isomeric oligonucleotide duplexes, [d(GAaTaC)]2 and [d(GTaA4C)]2; each oligomer binds two netropsin molecules and end-to-end ligation of the first oligomer forms curved polymers, which run in gel electrophoresis experiments with a slower mobility. The comparison of the resulting volume effects yielded the type of water involved in bent sequences. We obtained volume changes by mixing netropsin and oligomer duplex in two different ways: to form 1 : 1 ligand-oligomer complexes followed by the addition of a second equivalent of ligand, and to form directly the 2 : 1 complexes; both methods yielded similar results shown in Table III. Binding of netropsin to the first site of each oligomer generated a volume contraction whereas a volume expansion is observed in the binding to the second site. The magnitude of these volume effects was higher with the GAaT4C oligomer than with its GT4AaC isomer. Thus, at this salt concentration of 0.1 M, we infer that a net hydration accompanies the

162

BIOPHYSICALAPPROACHES

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TABLE III VOLUME EFFECTS ACCOMPANYINGINTERACTIONOF NETROPSIN WITH DNA OLIGOMERSa Duplex

AV (ml/mol)

[d(GA4T4C)12

- 100(first site) 37(second site) -63(saturated complex) -83(first site) 15(second site) -68(saturated complex)

[d(GT4A4C)]2

All values determined at 20" in 10 mM sodium phosphate buffer, 100 mM NaC1, 0.1 mM Na2EDTA, pH 7.

total binding to these oligomer duplexes as reflected by the apparent compression of water molecules. The [d(GA4T4C)]2 is slightly more hydrated by 5 ml/mol of duplex and indicates that this net hydration depends on the DNA sequence and the type of site of the oligomer duplex. To obtain a better understanding of the observed differences, we have dissected the overall volume profiles into the individual contributions for netropsin binding to the first and second site, respectively, and have taken the differences for each site, using the formation of the netropsin[d(GT4AaC)]2 as a reference state. The resulting AAV parameters are 17 ml/mol for the first site and 22 ml/mol for the second site. A similar procedure for the dissection of the AAG ° terms obtained from melting experiments (data not shown) yielded a marginal AAG ° of 0.4 kcal/mol for each binding site of netropsin. ~8 These results indicate that in the sequential binding of netropsin, there is an initial increase in hydration due to a hydrophobic or structural effect (opposite signs of AAV and AAG°), followed by a dehydration event due to electrostriction effects (similar signs of AAV and AAG°). ~8 Analysis of the electrophoretic mobility of the free and netropsin-bound oligomer duplexes shows that the mobility of the free [d(GAaT4C)]2 is lower than that of the free [d(GT4A4C)]2; the 1 : 1 netropsinoligomer complexes ran with similar mobilities whereas for the 2 : 1 complexes the trend is reversed. These results strongly indicate that binding of the first netropsin straightens the curved [d(GA4T4C)]2 duplex, yielding an important correlation with our volumetric studies in that the straightening of this oligomer invokes the participation of structural water. F o r m a t i o n of O l i g o m e r D u p l e x e s w i t h C o v a l e n t l y A t t a c h e d L i g a n d s One way to study the hydration properties of covalently attached moieties to DNA is by comparing the AV values for the formation of oligomer duplexes,

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163

by mixing its component single strands with and without the covalently attached moieties. In this section, we present results for two sets of oligonucleotides: the inclusion of two enantiomers of benzo[a]pyrene diol epoxide, (BPDE), (+)-BPDE and (-)-BPDE, with d(CCATCGCTACC)/d(GGTAGCGATGG) at the central Gforming DNA adducts; and the covalent attachment of cisplatin, cis-[Pt(NH3)2Cl2], to the central G G step of d(CTCTGGTCTC)/d(GAGACCAGAG). The volume effects accompanying the formation of a nucleic acid, from mixing its complementary strands, are interpreted to reflect changes in the electrostriction and/or hydrophobicity of water dipoles, which are immobilized by each of the participating species. For instance, the single strands have the bases more exposed to the solvent and immobilize more structural water. The base pairing and base pair stacks of the duplex bring the bases away from the solvent, yielding a duplex with a higher charge density that interacts with counterions more effectively and immobilizes more electrostricted water. The net result will depend on these two effects, which tend to compensate for each other. Furthermore, in the comparison of volume changes for each of these two sets we are assuming that the modified strands make a similar contribution to AV; this may be true because the complementary strands for a particular set have similar chemical compositions. Benzo[a]pyrene Diol Epoxide Adducts

BPDE molecules are metabolized in vivo and bind covalently to DNA. Their biological activities are significantly dependent on their stereochemical configurations. For instance, (+)-BPDE is strongly tumorigenic, whereas ( - ) - B P D E is not. 4°'41 All AV values were measured with a magnetic suspension densimeter. The formation of each duplex is accompanied by an uptake of water molecules as seen in the negative AV values of Table IV. The main observation is that the uptake of water for the unmodified duplex and the ( - ) - B P D E duplex is the same, whereas the water uptake is about 50% larger for the (+)-BPDE duplex. 42 Relative to the unmodified duplex, we obtained A AV values of 8 ml/mol of duplex for the ( - ) - B P D E duplex and - 6 5 ml/mol of duplex for the (+)-BPDE duplex. The modified duplexes are less stable, A AG ° of "-~5kcal/mol, but the overall counterion uptake is similar for all three duplexes, 40.8 mol of Na + per mole of duplex. The opposite signs of these thermodynamic parameters for the (+)-BPDE duplex reveal a high ordering of structural water whereas the similarity of the signs for the ( - ) - B P D E duplex indicate a marginal uptake of electrostricted water. The nuclear magnetic resonance (NMR) structures have indicated that these adducts lie in the minor groove of this duplex with different orientations: the (+)-anti-BPDE toward 4oA. H. Conney,Cancer Res. 42, 4875 (1982). 41p. Brookesand M. R. Osborne,Carcinogenesis 3, 1223 (1982). 42L. A. Marky,D. Rentzeperis,N. E Luneva,M. Cosman,N. E. Geacintov,and D. W. Kupke,J. Am. Chem. Soc. 118, 3810 (1996).

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TABLE IV VOLUME EFFECTS ACCOMPANYINGFORMATION OF DUPLEXES WITH CHEMICAL ALTERATIONS a Duplex

AV(ml/mol)

Inclusion of BPDE moieties h Unmodified With (-)-BPDE With (+)-BPDE

- 144 - 136 -209

Inclusion of cisplatin' Unmodified With cisplatin

19 - 35

a All measurements determined at 20 °. ~' Inclusion of BPDE moieties in 20 mM sodium phosphate buffer, 100 mM NaC1, pH 7. c Inclusion of cisplatin in 10 mM Na-HEPES, pH 7.5.

the 5' end of the unmodified strand and the ( - ) - B P D E toward the 3' end of the unmodified strand. 43 The sequence of this duplex is symmetric with respect to the modified base pair at the center, and therefore the (+)-BPDE is exposing more organic groups to the solvent. In addition, end-to-end ligation of (+)-BPDE duplexes yields bent structures 44 similar to the [d(GA4T4C)]2 duplex, and accordingly we conclude that bent duplex structures have a higher ordering of hydrophobic water.

Cisplatin The antitumor activity of cisplatin, a widely used drug, is attributed to its covalent interaction with DNA. The major cisplatin adducts are with GG or AG intrastrand cross-links. We used the Anton Paar densimeter to measure the volume changes accompanying the formation of duplexes. Unmodified duplex formation is accompanied by a marginal release of water molecules, whereas the cisplatin duplex reveals a significant uptake of water molecules (see Table IV); X-ray 45 and NMR 46 studies of d(CCTCTGGTCTCC)]d(GGAGACCAGAGG) with a cisplatin attached to the G G position demonstrated that the adduct causes the guanine bases to roll toward one another by 49 °, leading to an overall helix bend angle of 78 ° . We 43 C. De los Santos, M. Cosman, B. E. Hingerty, V. Ibanez, L. A. Margulis, N. E. Geacintov, S. Broyde, and D. J. Patel, Biochemistry 31, 5245 (1992). 44 B. Mar, Ph.D. Dissertation, New York University, 1994. 45 p. M. Takahara, A. C. Rosenzweig, C. A. Frederick, and S. J. Lippard, Nature (London) 377, 649 (1995). 46 A. Gelasco and S. J. Lippard, Biothemistry 37, 9230 (1998).

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investigated here a shorter version of this duplex, without one dC- dG base pair at either end, and it is expected to have a similar bend at the modified base pair. We have done an analysis similar to that of the previous set of duplexes and obtained, relative to the unmodified duplex, a AAV value of --54 ml/mol of duplex for the cisplatin duplex and a AAG ° of "-~3kcal/mol. 46 The opposite signs of AAV and AAG ° reveal that the bent cisplatin duplex is immobilizing structural water. Acknowledgment This work was supported by grant GM42223 (L.A.M.) from the National Institutes of Health.

47 B. 1. Kankia and L. A. Marky, unpublished data (2000).

[8] Calculating Sequence-Dependent Melting Stability of Duplex DNA Oligomers and Multiplex Sequence Analysis by Graphs By ALBERT

S. BENIGHT, PETR PANt~OSKA, RICHARD OWCZARZY,

PETER M . VALLONE, JAROSLAV NESETI},IL, a n d PETER V. RICCELLI

Introduction DNA duplex formation by annealing or hybridization of two complementary single strands is a central reaction in many important cellular events and such fundamental biological processes as replication, transcription, and recombination. In vitro binding and melting studies have shown, for short duplex DNA oligomers of 12 to 22 base pairs, that thermodynamic stability of flanking sequences is inversely related to the binding strength of several DNA restriction enzymes and site-specific binding ligands.l-3 Because DNA hybridization is sequence dependent, this basic reaction also underlies many research and commercial diagnostic applications. 4-6

1 A. S. Benight, E J. Gallo, T. M. Paner, K. D. Bishop, B. D. Faldesz, and M. J. Lane, Adv. Biophys. Chem. 55, 1 (1995). 2 p. V. Riccelli, P. M. Vallone, I. Kashin, B. D. Faldasz, M. J. Lane, and A. S. Benight, Biochemistry 38, 11197 (1999). 3 p. M. Vallone and A. S. Benight, Biochemiso'y 39, 7835 (2000). 4 E. A. Wagar, J. Clin. Lab. Anal. 10, 312 (1996). 5 j. G. Wetmur, Crit. Rev. Biochem. Mol. Biol. 26, 227 (1991). 6 K. B. Mullis, Ann. Biol. Clin. 48, 579 (1990).

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investigated here a shorter version of this duplex, without one dC- dG base pair at either end, and it is expected to have a similar bend at the modified base pair. We have done an analysis similar to that of the previous set of duplexes and obtained, relative to the unmodified duplex, a AAV value of --54 ml/mol of duplex for the cisplatin duplex and a AAG ° of "-~3kcal/mol. 46 The opposite signs of AAV and AAG ° reveal that the bent cisplatin duplex is immobilizing structural water. Acknowledgment This work was supported by grant GM42223 (L.A.M.) from the National Institutes of Health.

47 B. 1. Kankia and L. A. Marky, unpublished data (2000).

[8] Calculating Sequence-Dependent Melting Stability of Duplex DNA Oligomers and Multiplex Sequence Analysis by Graphs By ALBERT

S. BENIGHT, PETR PANt~OSKA, RICHARD OWCZARZY,

PETER M . VALLONE, JAROSLAV NESETI},IL, a n d PETER V. RICCELLI

Introduction DNA duplex formation by annealing or hybridization of two complementary single strands is a central reaction in many important cellular events and such fundamental biological processes as replication, transcription, and recombination. In vitro binding and melting studies have shown, for short duplex DNA oligomers of 12 to 22 base pairs, that thermodynamic stability of flanking sequences is inversely related to the binding strength of several DNA restriction enzymes and site-specific binding ligands.l-3 Because DNA hybridization is sequence dependent, this basic reaction also underlies many research and commercial diagnostic applications. 4-6

1 A. S. Benight, E J. Gallo, T. M. Paner, K. D. Bishop, B. D. Faldesz, and M. J. Lane, Adv. Biophys. Chem. 55, 1 (1995). 2 p. V. Riccelli, P. M. Vallone, I. Kashin, B. D. Faldasz, M. J. Lane, and A. S. Benight, Biochemistry 38, 11197 (1999). 3 p. M. Vallone and A. S. Benight, Biochemiso'y 39, 7835 (2000). 4 E. A. Wagar, J. Clin. Lab. Anal. 10, 312 (1996). 5 j. G. Wetmur, Crit. Rev. Biochem. Mol. Biol. 26, 227 (1991). 6 K. B. Mullis, Ann. Biol. Clin. 48, 579 (1990).

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Many assays utilize sequence-specific hybridization as the basis for recognition of nucleic acids. The functional utility of these assays relies heavily on the fidelity of DNA sequence-dependent hybridization and stability of the hybridized products. A number of practical applications employ hybridization between relatively short duplex regions. Stability of the hybridized products is determined by the sequence dependence of the transition temperatures (Tin values) of the duplex complexes. To optimize probe and primer sequence design 5'7 it is of particular necessity to be able to accurately predict the Tm values of DNA oligomer duplexes under a given set of conditions (ionic strength and strand concentration) from their base pair sequencesf In addition, an understanding of solution hybridization and DNA sequence-dependent thermodynamic stability has direct practical applications in the design of multiplex reactions in which many desired hybridizations are required to occur with high fidelity over a narrow temperature range, with all the required strands present in the same reaction mixture. These sequences are said to be "isothermal," in that they have the same Tm (or at least differ over a narrow range of T,,~ values). DNA microarrays are being designed or contemplated that will have thousands to millions of different DNA sequences on them. 9 For these microarrays to be useful all sequences will be required to hybridize precisely to their complement strand, and to no other. For these reasons there is widespread interest in analytical calculations of sequence-dependent stability of duplex DNA. A significant challenge in multiplex reaction design is the selection of isothermal sequences. An obvious approach to solving this problem might involve brute force calculation of the stabilities of all sequences, and then grouping them according to their calculated thermodynamic stability. We present a graph theory approach that can be employed to analyze and classify sequence families. This representation of sequences provides an efficient means for grouping them according to their graph topologies. Using these graph methods it is not necessary to calculate the stability of every individual sequence, group sequences in isothermal sets according to their calculated stabilities, and then count the sequences in each group. Rather, it is possible to classify sequences according to their graphical representation in such a way that all sequences within a class, represented by the same graph, can readily be counted, and by definition have the same sequence composition (base pairs and nearest neighbors) and thus must have identical calculated thermostabilities. The presentation is in two parts. In the first part, the nearest-neighbor (n-n) model for characterizing DNA sequence-dependent stability is reviewed. A set of n-n parameters is presented and the use of these parameters to calculate sequencedependent stability of short linear DNA oligomers is demonstrated. In the second 7 W. Rychlik, W. J. Spencer, and R. E. Rhoads, Nucleic Acids Res. 18, 6409 (1990). s R. Owczarzy, P. M. Vallone, E J. Gallo, T. M. Paner, M. J. Lane, and A. S. Benight, Biopolymers (Nucleic AcidSci.) 44, 217 (1997). R. J. Sapolsky, L. Hsie, A. Berno, G. Ghandour, M. Mittmann, and J. B. Fan, Genet. Anal. 14, 187 (1999).

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part, application of graph theory to sequence analysis is described. Essential elements of the theory are presented and necessary mathematical relationships are recalled. Finally, several illustrative examples are presented. Review of A n a l y t i c a l M e t h o d s a n d R e s u l t s In this section the basic model of melting short duplex DNA is reviewed. Key parameters are identified. The models that consider DNA sequence dependence are also reviewed, and a set of n-n parameters evaluated from melting studies of DNA dumbbells is presented. Use of the n-n stability values to calculate the thermodynamic parameters and Tm values of two short DNA duplex oligomers is demonstrated.

Two-State Melting Theory Melting of short duplex DNAs (<25 base pairs) is assumed to occur in an allor-none, or two-state, manner. That is, two complementary strands that combine to form a duplex exist either in the nonbonded single-strand or fully intact duplex state, with no partially melted intermediates. The simple two-state equilibrium for the formation of the duplex, D, by association of two single strands, S1 and $2, with equilibrium constant, KD, is written KD Sl + S 2 ~ - D

(1)

KD depends on the various molecular degrees of freedom of the duplex and singlestrand species. In terms of statistical thermodynamics KD can be expressed as the product of the ratios of the statistical weights of the internal and external degrees of freedom of the duplex and single-strand states. 8' 10-13 KD = [Zint(D)Zext(D)]/[Zint(Sl)Zint(S2)Zext(Sl)Zext(S2)]

(2)

The statistical weight ratio for the internal degrees of freedom accounts for the thermodynamic differences between the duplex state, Zint(D ), and single-strand states, Zint(Sl) and Zint(S2), for the concentration-independent part of duplex formation. This ratio is defined as Kint ~ Zint(D)/[Zint(S1)Zint(S2)]

(3)

l0 D. Poland and H. A. Scheraga, in "Theory of Helix-Coil Transitions in Biopolymers: Statistical Mechanical Theory of Order-Disorder Transitions in Biological Macromolecules." Academic Press New York, 1970. I l A. S. Benight, Y. Wang, M. Amaratunga, R Chattopadhyaya, J. Henderson, S. Hanlon, and S. Ikuta, Biochemistry 28, 3323 (1989). 12 A. S. Benight and R. M. Wartell, Biopolymers 22, 1409 (1983). 13 A. S. Benight, R. M. Wartell, and D. K. Howell, Nature (London) 289, 203 (1981).

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where Kin t is the equilibrium constant for changes in the molecular internal degrees of freedom associated with establishment of the forces (hydrogen bonding and stacking, salt binding and release, solvation and solvent rearrangement, which conceivably could also be affected by the external degrees of freedom) required for duplex formation. Kint for the sequence is defined in terms of the duplex melting transition enthalpy, AHduple×, and entropy, ASduplex: AHduplex - T ASduplex = - R T

In Kint

(4)

The thermodynamic quantities AHduplex and ASduplex are calculated directly from the base pair sequence, using any of the available n-n stability values 8,14,15 as described below, and are assumed to be temperature independent. The temperature independence for AHduplexand ASduplexin the above-described analysis relies on the assumption that there is zero difference between the heat capacities of the duplex and single strands, that is, ACp = 0. In fact, several authors have presented experimental and analytical arguments for a nonzero ACp between duplex and single-strand states.16-2 J It was also noted that measurements of this difference by differential scanning calorimetry (DSC) could be difficult. 2°'21 If ACp ¢ 0, the values of AHduplex and ASduplexmay not be entirely independent of temperature. Depending on the strength of the temperature dependence, thermodynamic parameters evaluated from analysis of melting transitions could be different (even inaccurate) at temperatures far removed from the transition region. The statistical weight ratio for the external degrees of freedom of the duplex and single strands considers the concentration dependence and configurational, orientational, and other factors associated with changes in the external (translational and rotational) degrees of freedom when single strands anneal to form a duplex. The quantity fl has been defined 8' 10-13 as Z e x t ( D ) / [ Z e x t ( S l ) Z e x t ( S 2 ) ] =--/~

AHnuc - TASnuc = - R T

(5) In 13

(6)

Different types of annealing between two single strands to form a duplex might be envisioned. For example, there is the common case in which complementary 14j. SantaLucia,Jr., Proc. Natl. Acad. Sci. U.S.A. 95, 1460 (1998). 15R. Owczarzy,R M. Vallone, R. E Goldstein, and A. S. Benight, Biopolymers (Nucleic" Acids Sei.) 52, 29 (1999). 16j. A. Holbrook, M. W. Capp, R. M. Saecker~and M. T. Record, Jr., Biochemistry 38, 8409 (1999). 17T. V. Chalikian, J. V61ker, G. E. Plum, and K. J. Breslauer, Proc. Natl. Acad. Sci. U.S.A. 96, 7853 (1999). 18I. Rouzina and V. A. Bloomfield,Biophys. J. 77, 3242 (1999). 19I. Rouzina and V. A. Bloomfield,Biophys. J. 77, 3252 (1999). 2oI. Jelesarov, C. Crane-Robinson,and R L. Privalov,J. Mol. Biol. 294, 981 (1999). 21 R L. Privalov,I. Jelesarov, C. M. Reid, A. I. Dragan, and C. Crane-Robinson,J. MoL Biol. 294, 997 (1999).

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169

single strands free in solution undergo diffusion-controlled encounters and anneal. Another case of interest might be one in which annealing of two complementary single strands occurs, but only one of the strands is free in solution and the other is attached to a solid support or surface, and therefore considerably restricted. It is expected that the form of fl could be different in formal treatments of these two cases. An analytical basis for treating annealing on microarray surfaces and solid supports has been attempted, 22 but experimental data verifying these theories are sparse. As more data are collected for annealing reactions performed on microarrays, our understanding of the process will be refined and, if appropriate, different forms of/3 will be tested and evaluated. In the context considered here,/3 is referred to as the nucleation (or initiation) parameter. From Eqs. (2), (3), and (5), KD = Kint/3

(7)

At any given temperature, the net fraction of intact base pairs, 0net, can be expressed in terms of the external and internal degrees of freedom of the duplex and single strands 1°-13 as 0net = 0ext0int

(8)

w h e r e 0ex t is the fraction of strands with at least one intact base pair, and 0in t is the

fraction of intact base pairs on duplex strands having at least one base pair. 1°-13 If the melting transition is two state, and the fully intact duplex and completely dissociated single strands are the only states that are populated by every strand throughout the entire melting transition, then 0int = 1 and 0net = 0ext. The total concentration of strands, CT, is given by Cv = [S i ] + [$2] + 2[D]. The expression for 0ex t in terms of Ko and CT is 0ext = [1 + uCTKD -- (1 + 20tCTKD)O'5]/~CTKD

(9)

For the case in which the strands are different (Sl # $2), c~ = 4; if the two strands of the duplex are the same ($1 = $2), c~ = 1. A plot of 0ext versus temperature provides the theoretical melting curve that is comparable to the baselinecorrected, experimentally measured optical melting curve. 8

Order-Dependent Sequence-Specific Interactions in Duplex DNA The model of thermodynamic stability of duplex DNA considers the important sequence-dependent forces, that keep the double helix together, to arise from two rather obvious components. These are hydrogen bonding between complementary base pairs (H-bonding), and nearest neighbor (n-n) stacking interactions with neighboring base pairs along and across the helix axis. In our approach to 22 g. Chan, D. J. Graves, and S. E. Mckenzie,Biophys.J. 69, 2243 (1995).

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treating sequence-specific interactions of increasing order in DNA, each level of order of interaction is evaluated sequentially. As described in detail elsewhere, 15 the procedure started from singlets (single base pair interactions), was expanded to doublets (n-n base pair interactions), and then further extended to include triplet (n-n-n base pair) sequence-specific interactions. The study of melting short duplex DNA oligomers as a means to evaluate sequence-dependent thermodynamic stability of DNA was pioneered by Breslauer and Marky in the early 1 9 8 0 s . 23-25 In 1986 they reported the first set of n-n sequence-dependent stability parameters for DNA evaluated from melting curves of short DNA oligomers and long repeating DNA polymers. 24 Since then a number of new and improved n-n sets, evaluated from melting experiments of short DNA duplex oligomers, DNA polymers, and DNA dumbbells, have appeared. 14,15,26,27 To date, the n-n model provides the most quantitative approach to providing accurate quantitative predictions of sequence-dependent DNA stability. Over the past 20 years there have been at least 13 different reports on the use of n-n stability parameters to calculate duplex thermodynamic stability. 14'15'24,26-36 Although impressive advances have been made, and new, improved, unified n-n sets have appeared, 14'15 current confidence is not yet absolute regarding our ability to accurately calculate the Tm of any short duplex DNA, given its sequence, concentration, and solvent environment. 8 There are several potential reasons for this: (1) deviations from two-state melting, (2) lack of understanding of the ionic strength dependence and related sequence effects on melting, (3) ACp not equal to zero, (4) long range and secondary effects of various sequence contexts (ends, length, sequence distributions, etc). Although our understanding has progressed considerably there is still much to be learned regarding sequence dependent thermodynamic stability of short duplex DNA.

23 L. A. Marky and K. J. Breslauer, Biopolymers 26, 1601 (1987). 24 K. J. Breslauer, R. Frank, H. B16cker, and L. A. Marky, Proc. Natl. Acad. Sci. U.S.A. 83, 3646 (1986). 25 L. A. Marky and K. J. Breslauer, Biopolymers 21, 2185 (1982). 26 M. J. Doktycz, R. E Goldstein, T. M. Paner, E J. Gallo, and A. S. Benight, Biopolymers 32, 849 (1992). 27 S. G. Delcourt and R. D. Blake, J. Biol. Chem. 266, 15160 (1991). 28 O. Gotoh and Y. Tagashira, Biopolymers 20, 1033 (1981). 29 R. L. Ornstein and J. R. Fresco, Biopolymers 22, 1979 (1983). 3o A. V. Vologodskii, B. R. Amirikyan, Y. L. Lyubchenko, and M. D. Frank-Kamenetskii, J. BiomoL Struct. Dynam. 2, 131 (1984). 31R. M. Wartell and A. S. Benight, Phys. Rep. 126, 67 (1985). 32 M. J. Aida, Theor. Biol. 130, 327 (1988). 33 C. R. McCampbell, R. M. Wartell, and R. R. Plaskon, Biopolymers 28, 1745 (1989). 34 p. Otto, J. Mol. Struct. 188, 277 (1989). 35 j. SantaLucia, H. Allawi, and P. Seneviratne, Biochemistry 35, 3555 (1996). 36 N. Sugimoto, S.-I. Nakano, M. Yoneyama, and K.-J. Honda, Nucleic Acids Res. 24, 4501 (1996).

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DNA SEQUENCE-DEPENDENTSTABILITYAND GRAPHS

17 1

Nearest-Neighbor Model

There are 16 possible different n-n base pairs, and hence 16 possible n-n interactions. Because of the antiparallel structure of duplex DNA, 6 of these possible stacks are degenerate, and only 10 of the 16 possible are unique and distinguishable. These unique n-n stacks, designated 5'-MN-3', are A A = TT, AT, TA, CA = TG, GT = AC, CT -----AG, GA = TC, CG, and GG -- CC, GC. There are 10 unique thermodynamic quantities (enthalpies, entropies, etc.) associated with melting each of the 10 unique n-n doublets that, depending on assumptions about the ends, can be evaluated from melting studies of an appropriate set of molecules. In addition, there are, in principle, four different sequence-dependent end interactions. Representing an end as E, the n-n end interactions are EA = TE, ET = AE, EG --- CE, and EC = GE. If these end interactions are assumed to be zero, then for an appropriate set of molecules, one in which all 10 n-n sequences are sufficiently represented, 10 linearly independent equations relating n-n sequence content and measured experimental parameters can be constructed. To solve for the required 10 unknowns (the n-n interactions), there must be 10 linearly independent equations. Because 10 linearly independent equations for the 10 unknowns exist, a unique solution for each of the 10 n-n base pair stacking interactions can be obtained (subject to the initial assumption regarding the ends). 37 Alternatively, if the end interactions are not assumed to be zero, but instead assumed to be the same, with a single, sequence-independent value, then 11 linearly independent equations can be written. In this case, a unique solution can be found for the 10 n-n sequence interactions and 1 end interaction. However, if the four additional n-n sequence-dependent end interactions are not assumed to be equivalent, or zero, there are 14 unknowns (the 10 n-n sequence interactions and the 4 end interactions) that should be evaluated. Unfortunately, for this situation there are only 12 linearly independent equations that can be written, 8,15'37 and a unique solution for all 14 unknowns cannot be obtained. Instead, only 12 linearly independent combinations of the n-n sequence interactions, and the ends, can be constructed and solved for. The linearly independent linear combinations can be used directly to calculate sequence-dependent stability.l's' 15,26,37 Values for these combinations provide direct insight into the relative contributions of different n-n sequence combinations and sequence-dependent end interactions. Generally, in treatments of n-n sequence dependence, the ends have been ignored or assumed to be the same and absorbed into a single nucleation parameter. 34"35'38 From published melting data on short duplex DNA oligomers, we evaluated 12 unique linearly independent n-n combinations. 8 Of these 12 linear combinations, 2 included contributions from only the n-n sequence-dependent end

37R. E Goldstein and A. S. Benight,Biopolymers 32, 1679 (1992). 38K. J. Breslauer, Methods MoL Biol. 26, 347 (1994).

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interactions. Results showed that although the fitted values of the enthalpies and entropies obtained were different, the free energies at 20 ° for both combinations were essentially identical. Precisely the same result would be obtained if a single end interaction were assumed. This eleventh unknown has been referred to by some as a nucleation parameter. 34,35'3s However, as pointed out, s the fact that a single eleventh parameter can be fit within the errors is not due to the existence of an eleventh unique parameter that is independent of n-n interactions per se. Rather, it is because the free energies of the end interactions are equivalent. In short, considering explicitly the n-n dependence of ends, there are 12 linearly independent linear combinations of n-n interactions. When values of these combinations were evaluated, the free energies of the two combinations that included end interactions were the same. As a consequence there are only 11 unique n-n parameters. Singlet and Doublet Formats'

n-n sequence dependence has been modeled in two ways. Both approaches facilitate empirical evaluations of thermodynamic stability parameters. These are referred to as the singlet and doublet formats. 1,8,J5 In the singlet format, contributions from H-bonding and stacking interactions are considered separately. 1.8,15,31 From melting data, plots of Tm versus %G-C are constructed that provide a means to evaluate the average H-bond contributions. 15,26,31 Deviations from the average due to n-n sequence dependence define the n-n interactions. In the doublet format the entire n-n interaction (H-bonding and stacking) is considered in a single parameter. Detailed descriptions of these formats and how they are equated have been reported.~,8,15,26 Calculated results from the singlet and doublet formats are numerically equivalent so long as the appropriate correction factor for the end base pairs is employed] Brief summaries of the singlet and doublet formats follow. In the singlet format, because contributions to DNA thermodynamic stability are apportioned into two parts, that is, H-bonding and n-n stacking, a unique n-n-dependent thermodynamic parameter can be assigned to every individual base pair along a DNA sequence. The primary component of this parameter includes average effects of Na + ionic strength on base pair H-bonding, and accounts for the difference in hydrogen bonding strength between A. T and G. C base pairs. In addition to the H-bonding between complementary base pairs on opposite strands, sequence-dependent stacking interactions with neighboring base pairs on either side are also considered and assigned thermodynamic parameter values. For example, in the singlet format, the free-energy change in forming base pair i, AGi, depends on the type of base pair i (A. T or G. C) and n-n stacking interactions with neighboring base pairs i - 1 and i + 1, and is given by AGi = ASbp(Ti -- T)

(10)

with the n-n-dependent effective melting temperature of base pair i = TH-B q-(~Hi l,i q"~Hi.i+l)/2ASbp

(11)

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173

TABLE I NEAREST NEIGHBOR-DEPENDENT STABILITY PARAMETERS

SHflfno ~ H ~Tn/AT

25 mM

55 mM

85 mM --76.4 92.0 --155.5

115 mM

?JH~/TA 6H~-~/TT

--39.3 124.0 --97.1

--56.5 105.2 --116.1

3 H~Cn/GT

--338.5

--125.0

--3.2

50.3

~H~/T G ~H~-~/TC 6H~/c T ,~H~/CG

210.8

--58.9

--214.8

--240.0

--366.2 509.6

-- 163.9 225.6

--41.1 84.4

--0.4 20.1

567.6 --882.6 261.8

81.1 --408.6 206.8

--214.6 --188.6 188.9

--313.1 --95.4 172.0

A G AT/AT n-n n-n AGTA/TA A G n-n AA/TT

--399.3 --236.0

--557.0 --395.3

--654.4 --486.0

-712.0 --556.6

--457.1

-616.6

--733.5

--787.4

AG~~/GT

-1228.2

-1128.1

-1068.9

-1058.8

A G~~/TG A G n-n GA/TC A G n-n AG/CT A G n-n CG/CG

--678.9 -1255.9 -380.1 --851.7

- 1062.0 -1167.0 -777.5 --1424.6

- 1280.5 -1106.8 -981.3 -1767.9

- 1349.1 -1109.5 -1089.0 --1899.6

AG~~/GC

--2301.9 -1157.5

--1914.3 --1298.9

--1741.9 -1364.4

--1681.9 -1414.5

~H~cn/GC n-n ~HGG/CC

--80.2 75.2 --155.6

AG~)"h

A G n-n GG/CC

a The deviations of the average enthalpy of stacking due to nearest-neighbor interactions. Units are calories per mole. h Nearest-neighbor doublet free-energies at 37°C. Units are calories per mole.

TH-B( = TATor T6c) is the average melting temperature of either an A. T (T. A) or G- C (C. G)-type base pair and includes effects of the H-bonding strength and the average of all 10 types of n-n interactions. The explicit n-n sequence dependence is carried in the 3Hg,i±l terms in Eq. (11), which are actually deviations from the average n-n stacking enthalpy, specific for each type of n-n stack. They can take on 10 different values. The values of ~Hi,i± 1 evaluated in four Na + environments are reported in Table I. The parameters listed in Table I were evaluated by assuming that n-n interactions for the end base pairs are like any other (end interactions are assumed to be zero) and that n-n sequence dependence in entirely enthalpic. The entropy of base pair melting, ASbp, is assumed to be essentially independent of sequence, s In the doublet format, n-n sequence dependence is considered to arise from the cumulative contributions of the H-bonds and n-n stacking interactions associated

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with a doublet of two base pairs. The individual contributions of H-bonding and n-n stacking are not separately parsed as in the single format. Transition enthalpies, AHMN, and entropies, ASMN, for each of the 10 possible 5'-MN-3' n-n doublets have been reported. 24'35 From the reported values of AHMN and ASMN, assuming no heat capacity difference between duplex and single strand states (ACp = 0), the free energy of each MN doublet, AGMN, can be determined as a function of temperature, T, according to AGMN = AHMN -- TASMN

(12)

Some investigators have reported individual sequence-dependent entropy values, ASMN, while others have assumed a single sequence-independent value for all doublets.t4'24 Although the evidence for a sequence-independent entropy is fairly strong, the actual sequence-independent values that have been reported are different, with ASMN = mSbp = -24.85(4-1.74) cal K -1 tool i bp l probably more appropriate for long DNA polymers and DNA dumbbells (15,25), and mSbp = - 2 2 . 4 c a l K -j mol ~bp -1 reported to be more appropriate for short DNA oligomers. 14 Empirically derived corrections for Na + environment and duplex length were also reported. 14 If ASMN = ASbp then, by analogy to Eq. (10), AGMN

= ASbp(TMN --

(13)

T)

where TMN is the effective melting temperature of doublet MN. The singlet and doublet formats can be equated by considering that TMN is the melting temperature for the doublet consisting of neighboring base pairs M and N. These base pairs have individual melting temperatures TM and TN equal to TAT or Tac, and a contribution from the stacking interaction between them. This stacking interaction is written as the deviation due to n-n stacking, aTMN, of TMN from the average melting temperatures of base pairs M and N, that is, aTMN = TMN -- (TM + TN)/2

(14)

6TMN = aHMN/ASMN

(15)

Again, assuming ASMN = ASbp and substituting these expressions in Eq. (13) the free energy of each n-n doublet can be determined according to AGMN = ASbp[(TM --}-TN)/2 +

~HMN/ASbp

--

T]

(16)

The set of doublet parameters determined using Eq. (16) and the singlet values in the upper part of Table I are tabulated in the lower part of Table I. These are provided because often it is easier to use doublet values to calculate sequencedependent stability. Results using either singlet or doublet values are numerically equivalent so long as the correction factor (described below) is employed.

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175

For any DNA sequence the total free energy of melting can be calculated with the reported singlet free energy values, AGT(singlet) = y ~ AGi

(17a)

i

or doublet values, AGT(doublet) = Z

AGMN

(17b)

MN

However, as calculated in Eqs. (17a) and (17b), AGT(singlet) is not numerically equivalent to AGT(doublet). Because of the averaging in Eq. (14) used to convert n-n singlet values to doublets, the two summed expressions in Eqs. (17a) and (17b) are not numerically equivalent. For an N base pair DNA, the correction factor required for numerical equivalence is (1)

AGcor = AGx(singlet) - AGx(doublet) = ASbp[(TI q- TN)/2 -- T]

(18)

The use of these equations and the values in Table I to calculate DNA thermodynamic stability is demonstrated below.

Calculating Transition Temperature The central observable parameter obtained from melting curve experiments is the transition temperature, Tin. This important parameter is defined as the temperature at which 0ext = 0.5 on plots of 0ext versus temperature. From Eqs. (4) and (6) the following relationship between Tm and C-r can be defineds'23 as

1~Tin = (R/AHT)ln(CT/ot) + AST/AHT

(19)

where AHT = AOduplex ~- AOnuc and AST = ASduplex+ ASnuc. Equation (19) is the familiar van't Hoff expression routinely employed to evaluate AHT and ASx from plots of 1/Tm as a function of ln(Cx/ot). 23 Rearranging Eq. (19) yields the two-state expression for Tm in terms of the respective thermodynamic quantities and total strand concentration, Tm = (AHduplex q- AHnuc)/[R ln(Cx/~) --I-ASduplex + ASnuc]

(20)

This expression can be employed to calculate Tm values of duplex sequences as a function of strand concentration, which are then compared directly with results obtained from experimental measurements. Calculations of Tm require input values for AHduplex, AHnuc, ASduplex, ASnuc, and CT. As described above, AHduplex and ASduplex are calculated according to the base pair sequence. The nucleation parameters have been the subject of some discussion and nonzero values of both ASnnc and A H.uc have been reported.S' 14,35,36,38 We employ an empirically determined form of the nucleation parameter in which AS, uc = 0 and AH,~c is a function of

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duplex length and %G-C. 8 The form we used improved average predictions of Tm values of short duplex DNA oligomers. 8

Empirical Corrections for Na + Dependence For oligomeric duplexes shorter than about 14 base pairs, the change in Tm with salt concentration, OTm/Olog[Na+], depends primarily on the duplex length N up to [Na +] about 0.4 M. 36 In fact, it was suggested 39 that the derivative can be crudely approximated by a value of N + 2, and that sequence identity and composition exert only secondary effects. Several empirically derived correction factors that account for the dependence of oligomer stability on Na + have been reported. 14,35 To scale calculated Tm values between 115 mM and 1.0 M Na + we reported the correction factor, 8 AT~ = Tin(0.115) - Tm(1.0) = (8.91)f~c - 11.34

(21)

Empirically derived equations for converting melting free energies for polymers and oligomers in different Na + environments were also reported.14

Using Nearest-Neighbor Parameters to Calculate Duplex Stability In the sample calculations that follow, base pairs on the ends are assumed to behave just as any other base pair and any n-n-dependent end interactions are assumed to be zero. This is precisely the assumption under which the 10 n-n parameters in Table I were evaluated. The n-n sequence-dependent transition enthalpy of the duplex is written in terms of the hydrogen-bonding component, AHH-bond, which depends only on the number of A-T (T-A) and G-C (C-G) base pairs, and the n-n interaction component, AHn-n. The duplex transition enthalpy is then determined according to AHduplex ~- AHH-bond -k- AHn_n =

ASbp[NATTAT q- NGcTGc] -~- y ~ . Nij(~Hij) ij (22)

where NAT and NGC are the numbers of A. T or G. C-type base pairs in the duplex sequence. The average melting temperatures of A- T or G. C base pairs are given by TAT or TGC. These values are readily obtained as a function of [Na+], using the following equations derived from the Frank-Kamenetskii relationships4°: TAT = 3 5 5 . 5 5 + 7.95 ln[Na +]

(23a)

TG¢ = 391.55 + 4.89 ln[Na +]

(23b)

The summed term on the right-hand side in Eq. (22) includes the n-n sequence dependence. Nij is the number of times the n-n doublet ij (ij = 1-10) occurs in 39 H. T. Allawi and J. SantaLucia, Biochemistry36, 1058 l (1997). 40 M. D. Frank-Kamenetskii, Biopolymers10, 2623 (1971).

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the duplex sequence, and ~nij is the deviation from the average n-n dependent enthalpy for sequence doublet, ij. The entropy change of base pair melting, ASbp, is assumed to be independent of sequence and depends only on [Na +] (slightly) and duplex length. 14 In the calculations below we use the reported value for DNA oligomers of - 2 2 . 4 cal K-1 mol -~ bp 1.~4In the following examples we do not use the suggested corrections of ASbp for salt and length. Thus, the total transition entropy of the duplex is simply ASduplex =

ASbp[NAT -1- NGC]

(24)

The duplex melting transition free energy AGduplex(T) = AHduplex - T ASduplex. In terms of the above quantities, ~Gduplex(T) = ASbp]NAT(TAT -- T) -k- NGC(TGc -- T)] +

~ij Nij (~Gij)

(25)

It bears mentioning that the aforementioned quantities were evaluated assuming ACp ¢ 0. Currently, there are no well-accepted or tested corrections to account for the potential temperature dependence of the relevant thermodynamic parameters. It has been suggested that any correction required due to ACp ¢ 0 may be largely independent of sequence. Iv Thus, two points should be noted for thermodynamic parameters calculated assuming ACp = 0, if such is actually not the case: (1) errors in thermodynamic values will be largest when extrapolated to temperatures far from Tm outside the transition region, and (2) even though calculated thermodynamic parameters for individual sequences may not be as quantitative as desired, relative differences in calculated stability for different DNA duplexes should be reliable and useful.

Specific Example for Two Sequences In this section two 10-base pair duplex sequences are considered in two different Na + environments. The n-n parameters in Table I are used to calculate their melting transition thermodynamic parameters and Tm values under a given set of conditions. For the first example, consider the 10-mer duplex sequence 5'-A-T-TA-T-G-G-G-G-C-3' in 115 mM Na +. From Eqs. (23a) and (23b), in 0.115 M Na +, TAT = 338.36 K (65.21 °) and Tcc = 380.97 K (107.82°). There are five A. T and five G. C base pairs, thus NAT = NGC = 5. The explicit terms of Eq. (22) are given by the following, for the H-bonding components: AHH-bond = ASbp[NAT(TAT) + NGc(TGc)] = -22.4[5(338.36) + 5(380.97)] = - 80,565.0 cal/mol The n-n sequence-dependent contributions to the stability depend on the numbers of the particular n-n doublets present. For the given sequence these

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are NAA/TT ~---1, NAG/CT ~---0, NAT/AT= 2, NAC/GT = 0, NGA/TC = 0, NGG/CC = 3, NGC/GC = 1, NTA/TA ~---1, NTG/CA ---~1, and NCG/CG = 0. From these numbers and the n-n parameters reported in Table I, the n-n-dependent contribution is given by

Z NijSHij -~ I(6HAA/TT) +

2(~HAT/AT) ÷ 3(~HGG/CC)

ij

+ I(3HGc/Gc) + I(3HTA/TA) + I(6HTG/CA) = --155.6 + 2(--80.2) + 3(172.0) + --95.4 + 75.2 + --2.40 = --60.2 cal/mol Thus, in this case the n-n dependence contributes only - 6 0 . 2 cal/mol to the total transition enthalpy. Assuming ACp = 0, the free energy of melting at 37 °, AGduplex(37°), is

AGduplex(37 °) = AHduplex -- 310.15(Nbv ASbp ) = -80,625.16 - 310.15(10)(-22.4) = - 11,151.6 cal/mol The value is also obtained with the doublet values in Table I and Eqs. (17b) and (18). That is,

AGT(doublet) = Z

AGMN = I(AGAA/TT) ÷ 2(AGAT/AT) ÷ 3(AGGG/CC)

MN

÷ I(AGGc/GC) ÷ I(AGTA/TA) + I(AGTG/CA) = 1(-787.4) + 2 ( - 7 1 2 . 0 ) + 3(-1414.5) + 1(-1681.9) + 1(-556.6)+ I(-1349.1) = - 10,042.5 cal/mol Applying the correction in Eq. (18), AGduplex(37 °) = AGT(doublet) + ASbp[(TAT ÷ TGC)/2 -- T] = -10,042.5 + -22.4(359.67 - 310.15) = - 11,151.6 cal/mol The Tm is calculated from Eq. (20) with AHduplex and ASduplex = NbpASbp, given above. For a reasonable prediction of Tin the contribution of duplex nucleation must also be considered. Nucleation accounts for unfavorable interactions between complementary single strands and other barriers that must be overcome in order for duplex formation to occur. Slightly different values for the nucleation parameters have been reported by authors of different n-n sets. 8' ~4,35,36,3s Our strictly empirical nucleation parameter is assumed to be entirely enthalpic, (ASnuc = 0), 8 AHnuc = 7654.71 - 3469.93(fGc) -- 186.5 I(N) cal/mol

(26)

DNA SEQUENCE-DEPENDENT STABILITY AND GRAPHS

[8]

179

Thus, asfGc and N increase, the enthalpic cost of duplex nucleation decreases. For the test sequence of interest with fGc = 0.5, AHnuc = 7654.71 - 3469.93(0.5) 1865.1 = 4054.65 cal/mol. If the total strand concentration, CT, is assumed to be 10 -6 M, the estimated Tm of the above sequence in 115 mM Na + is obtained by substitution of the appropriate quantities in Eq. (20), that is, T m = (-80,625.16 ÷ 4054.65)/[1.987 ln(10-6/4) + - 2 2 4 + 0] = 301.21 K = 28.06 ° Thus, under the stated conditions the sequence is predicted to have a Tm = 28.1 °. In the second example, consider the sequence 5'-A-G-T-A-T-G-A-C-G-T-3' in 55 mM Na +. This sequence has six A . T and four G. C base pairs, thus NAT = 6, NGC = 4, andfGc = 0.4. From Eqs. (23a) and (23b), in 55 mM Na +, TA.T = 332.49K and TG.C = 377.37K and ASbp = - 2 2 . 4 cal K -t mo1-1. Again, the hydrogen-bonding contribution from Eq. (22) is

AHH_bond =

ASbp[NATTAT q- NGCTGC]

= -22.4[6(332.49) + 4(377.37)] = -78,499.0 cal/mol For this sequence the following n-n doublets are present: AG/CT, TA/TA, AT/AT, TG/CA, GA/TC, and CG/CG and three GT/AC doublets. Thus, the numbers of different types of doublets in the sequence are NAT/AT = 1, NTA/TA = 1, NAA/TT = 0, NAC/6T = 3, NCA/TG -- 1, NTC/GA = 1, NCT/AG = 1, NCG/CG = 1, NGC/GC = 0, and NG6/CC = 0. Therefore, the n-n dependent contribution is

AHn-n =

Z N i j S H i j = I(~HAT/AT) + I(~HTA/TA) + 3(~HAc/GT) q- I(~HTc/GA) ij + I(~HcT/AG) q- I(SHcA/TG) -q- I(~HcG/GC)

With the appropriate values from Table I,

AH,_, = Z Nij3Hij = ij

1(-56.5) + 1(105.2) + 3 ( - 1 2 5 ) + 1(-163.9)

+ 1(225.6) + 1(-58.9) + (81.1) = - 2 4 2 . 4 cal tool -l With the above values for AHH-bond and AHn_n, the total transition enthalpy is given by AH~uplex = --78,499 + (--242.4) = -78,741.4 cal mo1-1 and the free energy at 37 ° is AGduplex(37 °) = --78,741.4 -- 310.15(10)(--22.40) = --9267.8 cal mol -I

180

BIOPHYSICAL APPROACHES

[81

Again, this value can also be obtained with the appropriate doublet parameters in Table I. That is, AGT(doublet) = ~

AGMN = I(AGAT/AT) + I(AGTA/TA) + 3(AGAc/6T)

MN

+ I(~XGTc/6A) +

I(AGcT/A6) + I(AGcA/T6) + 1(AG¢6/c6)

= 1(--557.0) + 1(-395.3) + 3(-1128.1) + 1(--1167.0) + ( - - 7 7 7 . 5 ) + 1(--1062)+ 1(--1424.6) = - 8 7 6 7 . 7 cal/mol and with the correction in Eq. (18), AGduple×(37 °) = AGT(doublet) + ASbp[(TAT -- T] = --8767.7 + -22.4(332.49 -- 310.15) = - 9 2 6 7 . 8 cal/mol According to Eq. (26), AHnuc for this sequence is AHnuc -----7654.71 -- 3469.93(0.4) -- 1865.1 = 4401.6 cal/mol Thus, from Eq. (20), at CT = 1 #M, the calculated Tm for this sequence in 55 mM Na ÷ is Tm = (-78,741.4 + 4401.6)/[1.987 1n(10-6/4) - 224 + 0] = 292.44K = 19.29 ° We have briefly reviewed essential features of the melting of short DNA oligomers. It should be noted that the analytical methods reviewed and described here pertain strictly to homogeneous populations of molecules. That is, every duplex formed is the same and formed from a set of only two complementary single strands (or a set of the same self-complementary strands). This distinction is made to point out the many potential differences in the annealing behavior of two complementary strands in solutions, versus the situation for multiplex reactions such as those that occur on DNA microarrays, where not only are many different strands present in the same solution, but also one of the complementary single strands is restricted by attachment to a solid support. The differences between annealing in solution and to a solid support are not known. Thus, it remains uncertain what meaning, if any, predictions made from solution thermodynamic parameters have for microarray annealing.

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181

R e p r e s e n t a t i o n s of S e q u e n c e s a s G r a p h s Graph theory has been applied to gain new insights into a number of important problems in physics, biology, and chemistry. 41'42 Many proposed and successful applications have been reported. 43 Here we demonstrate the use of graph theory for sequence analysis. In particular we exploit the properties of Eulerian graphs 43 to characterize and group sequences according to their H-bond and n-n components, which in turn constitute the basis for their calculated thermodynamic stabilities. For many applications involving multiplex hybridization reactions it is desirable to have many different sequences present having the same Tm (or at least that vary over a narrow range of Tm values). Such sequences are referred to as isothermal sequences. In general, the problem is to determine, for sequences of N base pairs, how many subsets of isothermal sequences exist out of the entire set of all 4 N sequences possible, and how many sequences are in each subset. This can be done by brute force, calculating the stabilities of all 4 N sequences using the H-bond and n-n parameters described in the previous section, and then sorting the sequences according to their predicted thermodynamic stability, and counting the sequences in each sorted set. An alternative approach, presented in the sections that follow, uses graph representations of DNA sequences as an efficient method for generating groups of isothermal sequences.

Graph Representation of DNA Sequence In this treatment the sequence of one strand is taken to represent the duplex considered. DNA sequences are represented in the form of the matrix shown in Fig. 1. In this matrix, termed the adjacency matrix 43 as described below, the diagonal elements correspond to the numbers of different types of individual bases in the sequences. The off-diagonal elements are the numbers of different types of n-n doublets in the sequences. In effect, the H-bonding contributions to stability appear along the main diagonal, while off-diagonal elements are the n-n contributions. The fifth column (E) considers the end interactions, where an end is designated by E, as done earlier. Only one of the EX (XE) matrix elements will be nonzero in the fifth row and column, depending on identities of the bases at the ends. To better visualize this representation, consider the two 10-base sequences analyzed as examples in the previous section. These sequences and their matrix forms are shown in Fig. 2a. In the parlance of graph theory these matrices are referred to as the adjacency matrices of graphs, with five vertices labeled A, C, G, T, and E, connected by oriented edges. 43 The graphs for the two sequences with the adjacency 41 M. S. Waterman, "Introduction to Computational Biology: Maps, Sequences and Genomes." Chapman & Hall, Boca Raton, Florida, 1995. 42 p. Pan6o~ka, V. Janota, and T. A. Keiderling, Anal. Biochem. 267, 72 (1999). 43 j. Matou~ek and J. Ne~effil, "Invitation to Discrete Mathematics." Oxford University Press, New York, 1998.

182

BIOPHYSICAL APPROACHES A

C

G

T

E

#A

AC

AG

AT

AE

C

CA

#C

CG

CT

CE

G

GA GC

#G

GT

GE

A

T

TA

TC

TG

#T

TE

E

EA

EC

EG

ET

1

[8]

FIG. I. General form of the adjacency matrix used as the descriptor of DNA sequence topology. Matrix elements on the main diagonal (#X) are the numbers of A, C, G, and T bases in the represented sequence. Off-diagonal elements (XY) are the numbers of doublets in the order indicated. For example, if the off-diagonal matrix element CA = 3, the sequence that is represented has the sequence motif: 5'E ~ . . . ~ C A ~ . . . ~ C A ~ . . . ~ C A ~ . . . ~ E - 3 ' , that is, three CA doublets. The fifth column E represents the 5' and 3' ends. For example, the sequence represented with elements EC = 1 and GE = 1, and other elements EX and XE = 0, has the ends: 5'-C~...~G-3'.

matrices defined in Fig. 2a are shown in Fig. 2b. Edges extend from the vertex corresponding to the row base in the adjacency matrix, to the vertex corresponding to the column base in the matrix. The number of oriented edges connecting particular vertices in the graph is the number of particular n-n doublets present a) 5'-A-T-T-A-T-G-G-G-G-C-3'

5'-A-G-T-~T-G-A-C-G-T-3'

A C G T E

A C G T E

A

2

0

0

2

0

A

3

1

1

1

C

0

1

0

0

1

C

0

1

1

0

0 0

G

0

1

4

0

0

G

1

0

3

2

0

T

1

0

1

3

0

T

I

0

1

3

1

E

1

0

0

0

1

E

1

0

0

0

1

b) 5'-A-G-T-A-T-G-A-C-G-T-3'

5'-A-T-T-A-T-G-G-G-G-C-3'

!

0

FIG. 2. Adjacency matrices and Eulerian graphs for two lO-mer sequences. (a) The adjacency matrices (as described in Fig. 1) for the specific DNA sequences shown. (b) Oriented Eulerian graphs that represent the set of sequences associated with the parent 10-mer sequences. These graphs are defined by the adjacency matrices in (a).

[8]

D N A SEQUENCE-DEPENDENT

STABILITYAND GRAPHS

183

in the sequence. For example, for the sequence on the left, there is no edge between vertex A and vertex C because there are no 5'-AC-3' doublets present in the sequence. In the graph for the sequence on the left, the doublet on the 3' end is CE and there is an edge between the C and E vertices of the graph. The loops leading from the G vertex represent multiple occurrences of the 5'-GG-3' n-n doublet. That there are three loops indicates that there are three 5'-GG-3' doublets. Finally, two edges leading from the A vertex to the T vertex represent the occurrence of two 5'-AT-3' doublets. The sequence can be traced in the graph by starting at vertex E and moving along the edges through each vertex and ending at vertex E. The graphs in Fig. 2b readily reveal differences in the topologies of the two sequences. The graph for the sequence on the left contains several loops (edges that exit and enter the same vertex) about the T and G vertices. In contrast, the graph for the sequence on the right has no loops. The topological difference between the sequences and their graphs is also evident from their adjacency matrices. For the sequence on the right, each diagonal element is equal to the sum of off-diagonal elements in the corresponding row and column. For example, the diagonal element for the third row and column is 3. This is obtained by adding the off-diagonal elements in column 3 (1 + 1 + 1 = 3) or row 3 (1 + 2 = 3). In contrast, for analysis of the sequence on the left, the adjacency matrix [A] must be decomposed into two submatrices, that is, [A] = [Al] + [A2], as shown in Fig. 3. The diagonal elements of the second submatrix, [A2], in Fig. 3 are the number of n-n doublets in the sequence for which both bases are the same. These numbers also correspond to the numbers of loops at each vertex in the graph. In the first submatrix, [All, the sum of the off-diagonal elements in each row and column is equal to the diagonal element for that row and column. This is the same property displayed by the full adjacency matrix for the sequence on the right. It turns out that any adjacency matrix, [A], can be decomposed into submatrices [A i] and [A2]. 43 In this decomposition the diagonal elements in [A2] are the differences between the diagonal elements of [A] and the sum of the off-diagonal elements in the corresponding rows and columns of [A]. Submatrix [Aj] is obtained by [A] A

C

G

= T

[All

E

A 2 0 0 2 0 C 0 1 0 0 1

A

=

C

U

+ T

E

A 2 0 0 2 0 C 0 1 0 0 1

[A2] A

+

C

G

T

E

A 0 0 0 0 0 C O O 0 0 0

G

0

1

4

0

0

G

0

1

l

0

0

G

0

0

3

0

T

1

0

1

3

0

T

1

0

1

2

0

T

0

0

0

1

0 0

E

1

0

0

0

1

E

1

0

0

0

1

E

0

0

0

0

0

FIG. 3. Decomposition of the adjacency matrix. The adjacency matrix for the sequence on the left in Fig. 2 (5'-A-T-T-A-T-G-G-G-G-C-3') is decomposed into submatrices [AI] and [A2]. As described in text, this decomposition is required in order to enumerate the number of sequences, N(G), that are predicted to be isothermal with the parent sequence. The form for N(G) is given in Eq. (28) in text.

184

BIOPHYSICALAPPROACHES

[8]

replacing the diagonal elements of [A] by the sum of the off-diagonal row or column elements of [A]. This decomposition separates the simple counting of bases of a given type X into two equivalent steps. Counting can be performed by using the base on the left or right of X, making the distinction of whether the base is different from (matrix [A 1]) or identical to (matrix [A2]) X. For example, the total number of C bases in any sequence is also equal to the number of 5'-YC-3' or 5'-CY-3' n-n doublets (Y = A, T, G, or E) plus the number of 5'-CC-3' doublets. This decomposition of the [A] matrix is essential for determining whether there are multiple sequences corresponding to the same adjacency matrix [A], and how many such sequences there are. Any adjacency matrix [A] defines a unique combination of H-bond and n-n interactions in the sequences it represents. Therefore, if there are multiple DNA sequences with a given adjacency matrix [A], they are, according to their calculated thermodynamic stability, by definition isothermal. The sequences can be identified by realizing that any matrix [A] can be represented by a special type of oriented graph. A synopsis of the construction of the graph for the sequence on the left in Fig. 2 is shown in Fig. 4. First, the linear sequence at the top is circularized by bringing the ends together and then folded as shown. Each vertex of the graph is generated by merging all bases of a given type into a point (vertex A, C, T, G, or E) and preserving the covalent links between bases as the edges. This type of graph can be drawn by tracing along every edge only once through every vertex with a single, noninterrupted stroke of the pen. Such a graph is called a Eulerian graph and the trail traced through it is called an Eulerian trail. 43 For every Eulerian graph there is at least one Eulerian trail. 43 More importantly, for our purposes of selecting groups of isothermal sequences, this mathematical representation of DNA sequences is quite useful for the following reasons. In most cases for any given Eulerian graph there is more than one Eulerian trail starting and ending at vertex E. Each of these trails represents a different linear sequence with exactly the same H-bond and n-n sequence composition and length. For example, for the graph in Fig. 2b, there are two possible trails that lead to two different sequences. These are the sequence: 5'-A-T-T-A-T-G-G-G-G-C-3'(the original) and 5'-A-T-AT-T-G-G-G-G-C-3'. The topology of the graph is unchanged no matter which path is taken. Thus, both sequences have the same adjacency matrix, and therefore by definition have the same number of individual bases and n-n doublets, and perforce are predicted to be isothermal. The above examples can be generalized for any DNA sequence. The adjacency matrix of a Eulerian graph can represent many different sequences, and is thermodynamically invariant for the sequences associated with it. Each class of sequences is represented mathematically by the set of all possible Eulerian trails in a given Eulerian graph defined by [A]. By taking this course, our problem of grouping sequences according to their thermodynamic stability has been converted to a problem of enumerating the Eulerian trails starting and ending at vertex E in

[8]

D N A SEQUENCE-DEPENDENT STABILITY AND GRAPHS a

EAT

T

1 85

b

ATe,

0

d

G

G

C

c

!

! e

f

FIG. 4. Schematic of graph representation of a DNA sequence. Sequential steps (a)-(f) in the construction of the Eulerian graph for the left sequence in Fig. 2 are diagrammed. (a) The linear sequence 5'-A-T-T-A-T-G-G-G-G-C-3' is circularized by merging the end E vertices into a single vertex. (b) Vertices for the same type of bases are sequentially merged into a single vertex with graph edges representing covalent links between the merged bases as follows: (c) merging of the A vertices (bases), (d) merging of the G vertices (bases), and (e) merging of the T vertices (bases). (f) The merged vertices are represented by a single corresponding vertex with loops as shown. This graph representation of the sequence contains information about sequence length (the number of edges), sequence composition (number of edges entering and leaving vertices), sequence context, and nearestneighbor doublets (number of edges and loops at vertices). What is not preserved is the specific order of the bases in the original sequence. Thus, all other sequences compatible with the restrictions preserved in the graph have the same length, composition, and nearest-neighbor doublets, and consequently the same calculated thermodynamic stability. the E u l e r i a n g r a p h r e p r e s e n t i n g the s a m e class o f D N A s e q u e n c e s . Fortunately, a g e n e r a l s o l u t i o n o f this p r o b l e m h a s b e e n derived. 44

Mathematical Relationships from Graph Theory Here, the b a s i c m a t h e m a t i c a l n o m e n c l a t u r e o f g r a p h t h e o r y a n d n e c e s s a r y p e r t i n e n t results are r e v i e w e d . L e t G = (V,E) b e a n E u l e r i a n g r a p h t h a t r e p r e s e n t s 44 p. Panro~ka, V. Janota, and Jaroslav Ne~etfil, in "Contemporary Trends in Discrete Mathematics" (R. L. Graham, J. Kratochvfl, J. Ne~effil, and E S. Roberts, eds). American Mathematical Society, Providence, Rhode Island, 1999.

186

BIOPHYSICAL APPROACHES

[8]

a class of DNA sequences of a given length. Vertices connected by an edge e E E (e is an element of the edge set, see below) are denoted by the oriented pair (X,Y) representing the doublet 5'-XY-3' in the sequence, where X and Y represent one of the four bases A, T, C, and G or the end, E. Let vi be a member of the vertex set V = {vl . . . . . v,,}, and let ei be a member of the edge set E = {el . . . . . era}. For a given vertex vi the number of edges e/that exit vi are denoted by d + (vi), and the number of edges that enter vi by d- (vi). These are actually the numbers of 5'-XY-3' and 5'-YX-3' n-n doublets (X,Y = A, T, G, C, E). Let the number of loops (the edges [vi, vi] in our notation) be denoted by m(vi). The m(vi) are the diagonal elements of submatrix [A2] and represent the number of 5'-XX-3' n-n doublets in the sequence. Similarly, the number of edges connecting different vertices vi and vj with i # j, are given by m(vi, Vj). Consider two important matrices associated with an Eulerian graph G. These are the adjacency matrix [A(G)] = [aij], described above and defined by aij = m(vi, v j), and the Kirchoff(or combinatorial Laplacian) matrix [L(G)] = [Sij]. 43'44 Elements of the latter are defined by

Sij = - m ( v i , v j), Sii = ~

i # j

(27a)

m(vi, Vj) = d+(vi) -- m(vi)

(27b)

i(.i)

The nonsymmetric elements in Eq. (27a) are just the number of edges connecting different vertices, i and j, i # j. The symmetric elements in Eq. (27b) are the number of edges that exit vertices minus the loops. Elements of matrix [L(G)] are obtained from the submatrix [Ad by changing the signs of the of-diagonal elements. The number of Eulerian trails that begin and end at vl (E is selected as this vertex), N(G), can be computed according to 43

N(G) = det(LEE)

d+(vl)! [ ' I ( d + ( v i ) - 1)! i#l l-I(aij) ! I-I m(vi )! i,j i

(28)

Matrix LEE is the submatrix of [L(G)] obtained by omitting the last row and column (which correspond to the sequence ends) from [L(G)]. With our choice of vl = E, it follows that d+(vl) = d+(E) = 1, and the expression in Eq. (28) is simplified.

Finding Set of Eulerian Trails The validity of Eq. (28) is based on the algorithm for finding all Eulerian trails in a given Eulerian graph. The algorithm is discussed here to provide the basic meaning of the terms in Eq. (28). The algorithm comprises two steps. A spanning tree is a graph with edges that leave only vertices. 43 The network of these leaving edges connects all vertices in the graph. The first step involves

[8]

DNA SEQUENCE-DEPENDENTSTABILITYAND GRAPHS

187

finding all subgraphs of G that are spanning trees "rooted" (or starting) at vertex E. This ensures there will always be a path back along the tree edges from any vertex.43 The terms in det(LEe) enumerate the number of trees in a given Eulerian graph G. 41 In the second step, starting at vertex E, a search is made for a path to another vertex through an edge that is not part of the tree. An edge that is part of the tree is used only if there is no other way to leave the vertex. This procedure is repeated consecutively for every vertex visited (each base in the sequence) and ends necessarily when the path generated by it is an Eulerian trail in the graph. 43 Given the graph G, the number of possibilities encountered in the second step is determined by the second term in Eq. (28). Thus, the total number of sequences is the product of a term reflecting the restrictions on the number of tree subgraphs of G, introduced by the sequence mapped into the graph topology, and a term dependent on the length and n-n topology of the sequence mapped on the number of edges incident with each vertex.43 By definition, loops do not influence the number of spanning trees in a given graph. Therefore, the submatrix [Ai] is used to define det(LEE). Even so, loops do represent covalent connections between adjacent bases in the sequence, and should be included in evaluations of the combinatorial complexity of possible DNA sequences associated with a given graph G. The second term of Eq. (28) requires the matrix elements of the complete adjacency matrix [A].

Finding Number of lsothermal Sequences The problem is as follows: given a DNA sequence and its graph, find the number of all other DNA sequences associated with that graph. As described above, this is the number of sequences in a set that will be calculated to be isothermal. In this example, the numbers of sequences associated with the two sample sequences shown in Fig. 2 are determined. Consider the sequence on the right-hand side first. The corresponding graph has no loops. Therefore, matrix [A] can be used directly to evaluate the required quantities, because [A2] = 0 and [A] = [Ad. It follows from Eq. (28) that the number of Eulerian trails (sequences) is 3

N(G) = det

0 -1 -1

-1 0 0

-1 3 -1

-i 1

1!.(3-1)!.(1-1)!.(3-1)!.(3-1)! 1!1!1!1!1!2!1!1!0!

8 =7.-=28 2 Thus, there are 28 sequences isothermal with the original sequence. The graph for the left-hand sequence has loops• Therefore, decomposition of the adjacency matrix [A] into submatrices [A1] and [A2], as shown in Fig. 3, is required in order to determine the individual terms in Eq. (28). From matrix [All

188

BIOPHYSICAL APPROACHES

[8]

we obtain for the first term,

det[(LeE)l] = det

and for the second term, d+(Vl)! - 1-I(d+(vi)i:fil

1)!

2

0

0

0 0 -1

1 -1 0

0 1 -1

I

1!. ( 2 - 1)!- (1 - 1)!. ( 4 - 1)! • (3 - 1)!

1-I(aif)" l-Im(vi)! i,j

il

=2

(2!). (3!. 1!)

i

Therefore, the number of sequences is N(G) = 2 x 1 = 2. Thus, for the left-hand sequence, there are only two members in the set of isothermal sequences. This reduction in the number of isothermal sequences for the left-hand sequence compared with the right-hand sequence reflects the effect of restrictions on the primary sequence topology of the right-hand sequence brought about by the requirement of having no contiguous stretches of G and T bases. Such sequence requirements minimize the combinatorial possibilities of generating predicted isothermal sequences.

Numbers of Isothermal Sequences with Different Sequence Restrictions As a second example of the application of these techniques, the use of the graph representation to consider the effects of a priori sequence restrictions (e.g., no contiguous bases of a given type) is demonstrated. Consider the graph in Fig. 5a, which represents a set of isothermal DNA duplexes with 16 base pairs. The adjacency matrix decomposed into submatrices [A1] and [A2] for this graph is shown in Fig. 5b. According to Eq. (28),

det[(LEe)l] = det

and d+(Vl)! • I-I(d+(vi) i¢1

- 1)!

]-I(aij) ! " [-I m(vi)! i,.]"

E3°-2i1 0 -2 0

2 0 -2

2304 -- 72 (2!) 5

0 3 -1

~

=10

N(G) = 10 × 72 = 720

i

Thus, the graph in Fig. 5 corresponds to a class of 720 isothermal sequences. The graph in Fig. 6 represents again a set of isothermal 16-mer sequences with the same sequence content as the sequences represented in Fig. 5, except sequences containing 5'-GGG-3', 5'-CC-3', and 5'-TTT-3' base stretches were not allowed. In terms of their graph representations, the effect of this restriction can be obtained

[8]

DNA SEQUENCE-DEPENDENTSTABILITYAND GRAPHS

189

a)

b) [A]

=

[A1]

+

[A2]

A

C

G

T

E

A

C

G

T

E

A

C

G

T

E

A

3

0

2

1

0

A

3

0

2

1

0

A

0

0

0

0

0

C

0

3

0

1

1

C

0

2

0

1

1

C

0

1

0

0

0

G

2

0

5

1

0

G

2

0

3

1

0

G

0

0

2

0

0

T

0

2

1

5

0

T

0

2

1

3

0

T

0

0

0

2

0

E

1

0

0

0

1

E

1

0

0

0

1

E

0

0

0

0

0

=

+

FIG. 5. Graph representation of a set of sequences predicted to be isothermal. (a) Eulerian graph representation of a set of 16-mer DNA sequences composed of 18.75% A and C bases and 31.25% G and T bases. (b) The adjacency matrix [A] defined by the sequence graph in (a) and decomposition of [A] into submatrices [A]] and [A2] required to determine N(G) [from Eq. (28) in text], the number of sequences calculated to be isothermal.

by transformation of the graph in Fig. 5 to that in Fig. 6. Elimination of GGGG, CC, and TTT stretches necessarily causes elimination of the loops from the graph in Fig. 5. Any pair of vertices in the original graph in Fig. 5 that contain a loop can be connected by two antiparallel edges that replace the two loops. This can be done without compromising the graph representation of sequences with the same sequence content and length. In this way the graph in Fig. 5 was transformed into the graph in Fig. 6. The number of 16-mer isothermal sequences associated with this new graph is

N(G) =

\~--

/ = 2016

This is a factor of 2.8 times more than the number of sequences in the restricted parent sequences associated with the graphs in Fig. 5.

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FIG. 6. Graph representation of another set of sequences predicted to be isothermal. Eulerian graph representation of a set of 16-mer DNA sequences composed of 18.75% A and C bases and 31.25% G and T bases. These sequences have the same composition as those represented by the graph in Fig. 5, except 5 ' ~ . . . ~ G G G G ~ . . . ~ 3 ' , 5 ' ~ . . . ~ C C ~ . . . ~ 3 ' , and 5 ' ~ . . . ~ T T T ~ . . . ~ 3 ' sequence motifs are not allowed. As described in text, by invoking these restrictions, the graph in Fig. 5 was transformed to the graph shown here. These sequence restrictions increase the number of sequences with the given sequence composition predicted to be isothermal to 2016, compared with the 720 sequences predicted to be isothermal represented by the graph in Fig. 5.

Analysis of Sequences of Blocks The graph techniques can be directly adapted for the analysis of DNA sequences that are assembled not from individual bases, but instead from a limited number of oligomeric blocks. For example, the graph in Fig. 7 is a representation of one isothermal set from the 57 = 78,125 possible 28-base DNA sequences assembled from different combinations of 7 blocks drawn from the 5 tetramers ATGT (1), AGAT (2), TGAT (3), GATT (4), and AATG (5). In the graph representation, vertices represent sequence blocks instead of individual bases. The number of isothermal block sequences associated with the graph in Fig. 7 is derived from the adjacency matrix for the graph also shown in Fig. 7. This matrix has det(LEE) = 2. From Eq. (28) the number of isothermal sequences is calculated to be N(G) = 2(1/1) = 2. Thus, the resulting two isothermal block sequences are

5'-ATGT-AGAT-TGAT-GATT-ATGT-GATT-AATG-3' 5'-ATGT-GATT-ATGT-AGAT-TGAT-GATT-AATG-3' As another example of the effect of sequence restrictions on the number of isothermal sequences, consider that some of the block sequences from the complete set of 78,125 possible will not contain all tetramers from the block set. Figure 8 shows an example of an adjacency matrix and corresponding graph for such a restricted case. That is, of the five block sequences considered above, the

[8]

D N A SEQUENCE-DEPENDENT STABILITY AND GRAPHS

191

(a)

(b) 1 2 345 1

2

1

0

2 3 4 5 E

0 0 1 0 1

1 1 0 1 002 0 0 0 0

t

E 0

0

0 0 0 1 0 0 1 0 0 1 1 0 0 1

FIG. 7. Graph representation of a set of sequences, predicted to be isothermal, that are constructed from sequence blocks. (a) The graph of the 28-mer sequences constructed from the five four-base blocks with the sequence shown (the vertices). Identical oligomeric blocks in the sequence merge in the graph vertices and covalent attachments between blocks are represented by the oriented edges. The H-bonding and nearest-neighbor interactions within each block are preserved, while new contributions are generated at block boundaries represented by the graph edges. As for sequences composed of monomer base blocks, this representation provides for the enumeration of block sequences predicted to be isothermal. (b) The adjacency matrix associated with the Eulerian graph (a). Row and column elements represent sequence blocks, that is, 1 = ATGT, 2 = AGAT, 3 = TGAT, 4 = GATT, 5 = AATG. The 5' and 3' ends of the block sequence are represented by nonzero elements in the corresponding E row and column of the matrix. All sequences associated with the graph in (a) have the same terminal blocks, 5' -ATGT~... and ...~AATG-3'. Diagonal elements are the numbers of blocks of a given type in the sequence. Off-diagonal elements are the numbers of interblock boundaries of a given type.

A A T G (4) a n d G A T T (5) b l o c k s e q u e n c e s are not allowed. T h u s , the p r o b l e m is to find the set o f 2 8 - m e r s e q u e n c e s , c o m p o s e d o f o n l y the A T G T (1), A G A T (2), a n d T G A T (3) t e t r a m e r blocks, that are p r e d i c t e d to b e i s o t h e r m a l . F o r this situation, d e t e r m i n a t i o n o f the n u m b e r o f i s o t h e r m a l s e q u e n c e s is b a s e d solely o n the a d j a c e n c y m a t r i x o f the c o n n e c t e d s u b g r a p h o f G. T h e s u b g r a p h in Fig. 8 is chara c t e r i z e d b y the a d j a c e n c y m a t r i x s h o w n in Fig. 8. T h i s m a t r i x has det(LEE) = 4 a n d f r o m Eq. (28) the n u m b e r o f i s o t h e r m a l s e q u e n c e s is c a l c u l a t e d to b e N ( G ) = 4(2/2) = 4. T h e s a m e p r o c e s s c a n b e a p p l i e d to the situation in w h i c h the r e s t r i c t i o n s are o n i n d i v i d u a l bases. F o r e x a m p l e , c o n s i d e r the s e q u e n c e s c o n s i s t i n g o f A, T, G, a n d E b a s e s only. T h e effect o f this r e s t r i c t i o n o n the c a l c u l a t e d n u m b e r o f i s o t h e r m a l s e q u e n c e s c a n b e f o u n d f r o m the r e s u l t a n t s u b g r a p h s ( h a v i n g o n l y A,

192

BIOPHYSICAL APPROACHES

[8]

(a)

(b) l 2 3 E 1

3

2

l

0

2 I 2 1 0 3 t 0 2 1 E 1 0 0 l FIG. 8. Graph representation for a subset of the possible 28-mer block sequences predicted to be isothermal. (a) Graph for the 28-mer block sequences that can be constructed from three of the five tetramer blocks used to construct the graph in Fig. 7. These sequences contain blocks 1 (ATGT), 2 (AGAT), and 3 (TGAT) of Fig. 7, but do not contain blocks 4 (GATT) and 5 (AATG). (b) The adjacency matrix associated with the graph in (a). The sequences represented have the same 5'- and Y-terminal blocks, that is, 5'-ATGT~... and ...~TGAT-Y.

T, and G vertices) of the complete graphs (not shown). Thus, only four vertices in the graph are used to enumerate the number of isothermal sequences. Summary and Conclusions The analytical methods for characterizing DNA sequence-dependent thermodynamic stability have been reviewed. A set of n-n sequence stability parameters is presented. Examples in which these values are used to calculate the thermodynamic stability of short duplex DNA oligomers are presented. The problem of determining sets of isothermal sequences is addressed by representing DNA sequences as graphs. Representing DNA sequences by a graph descriptor with special mathematical properties minimizes the computational difficulty of determining the number of DNA sequences with identical predicted thermodynamic stability. This is achieved by replacement of a whole set of sequences by a single representative. Applications of this concept were demonstrated for sequences assembled from individual bases and sequences assembled from oligomeric blocks.

[9]

THERMALDENATURATION

193

[9] T h e r m a l Denaturation as Tool to S t u d y DNA-Ligand Interactions By CHARLESH. SPINK and SUSAN E. WELLMAN Introduction Proteins often associate with DNA via binding to sequence-specific sites on the duplex chain. However, a variety of tigands, including proteins, drugs, cationic peptides, and lipids, can be nonspecific binders that have only weak preference for sequence. These ligands associate with the DNA via combinations of electrostatic, hydrogen-bonding, or hydrophobic interactions that are relatively independent of the base sequence. This nonspecific association leads to DNA-ligand complexes whose stabilities are affected not only by inherent DNA-ligand interactions, but also by statistical factors, such as neighbor exclusion, or by cooperative effects between binding ligands. The association of these ligands with DNA can have dramatic effects on the helix-coil transition in the DNA, leading to either increases or decreases in the transition temperature. Both stabilizing interactions with duplex form and destabilizing effects due to ligand association with single-strand polynucleotide are possible. Thus, to analyze the thermodynamic consequences of thermal melting in the presence of ligand binding to DNA, consideration must be given to the effects of ligands on both the helix and coil forms. It is to this fundamental thermodynamic problem that this chapter is directed. The basic question is, "is it possible to come up with a self-consistent model for the melting of DNA in the presence of nonspecific ligand binding such that thermodynamic information on the binding process can be derived from experimental thermal transition curves?" Relatively few attempts have been made to analyze this question, but perhaps the most successful and rigorous approach was published in 1976 by James McGhee. i Using an Ising model approximation, 2 combined with calculations that are based on Lifson's sequence-generating functions, 3 this method provides a basis for analyzing DNA melting under circumstances encountered in a variety of DNA-ligand systems. It is our intention in this chapter to examine the development of ideas relating to the analysis of thermal melting curves in the presence of ligands, and then present the theoretical background to the McGhee method and its application to DNA melting. Finally, we present a few cases in which the McGhee method has been applied with some degree of success. We hope to show that in the appropriate I j. D. McGhee, Biopolymers 15, 1345 (1976). 2 D. Poland and H. A. Scheraga, " T h e o r y o f H e l i x - C o i l Transitions in Biopolymers.'" A c a d e m i c Press. New York, 1970. 3 S. Lifson, J. Chem. Phys. 40, 3706 (1964).

METHODS IN ENZYMOLOGY,VOL. 340

Copyright(~)2001 by AcademicPress All rights of reproductionin any form reserved. 0076-6879/00$35.00

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systems, analysis of DNA melting in the presence of ligands can help elucidate thermodynamic properties of the binding processes involved. Background The basic problem that must be confronted in developing models for the prediction of melting behavior of DNA in the presence of ligands is to find an appropriate binding theory for association of ligands with DNA. The classic approach developed by Scatcbard4 provides a way for determining binding constants and site size from a plot of the amount of bound ligand per mole of lattice sites versus the free ligand concentration (the famous Scatchard plot). This treatment, while working quite well for protein-ligand binding, is limited to cases in which ligand binds to only one repeating unit of the lattice of sites. Problems arise when the ligand binds to two or more residues along the lattice, a common situation in binding of proteins or drugs to DNA. Attempts to deal with this problem 5-8 have met with somewhat varying success in the treatment of experimental data. A comprehensive treatment, which includes not only multiple site binding, but also neighbor exclusion and cooperativity between ligands, has been developed by McGhee and von Hippel. 9 This method provides equations in closed form that can be used in curve-fitting procedures, and provides a more detailed analysis of the characteristics of the binding process. To treat melting transitions for DNA with bound ligands, however, the theory must include evaluation of the extent of binding to the coil form of the polynucleotide. One of the first attempts at analyzing nucleic acid melting in the presence of binding ligands was made by Crothers. 1° On the basis of a straightforward thermodynamic approach that develops relationships between the shift in melting temperature and activity (concentration) of ligand, equations were developed for cases involving binding of small molecules to DNA polymers. Situations with varying degrees of saturation of the DNA chain, or when the ligand binds only weakly to the coil form, were analyzed. Also considered were examples of binding of small molecules to oligonucleotides, for which the thermal transition is between ligand-bound helical form and unbound coil form. Finally, the case of association of oligomers with polymeric nucleotides to form double or triple helical structures was presented. The equations that were developed for these examples allowed the plotting of experimental data to extract thermodynamic data for the thermal 4G. Scatchard,Ann: N.Y. Acad. Sci. 51,660 (1949). 5 S. A. Latt and H. A. Sober,Biochemistry 6, 3293 (1967). 6 D. M. Crothers, Biopolymers 6, 575 (1968). 7j. A. Schellman,Israel J. Chem. 12, 219 (1974). 8A. S. Zasedatelev, G. V. Gurskii, and M. V. Volenshtein,Mol. Biol. (Russ.) 5, 245 (1971). 9j. D. McGheeand P. H. von Hippel,J. Mol. Biol. 86, 469 (1974). 10D. M. Crothers,Biopolymers 10, 2147 (197l).

[9]

THERMAL DENATURATION

~'=

195

ligand coil

cooperativity Melting of DNA FIG. 1. Melting of DNA in the presence of ligands that can bind to either helix or coil regions.

transitions and in some cases the binding parameters. However, the approach is restricted to small molecules that do not interact with multiple sites and are noncooperative. Other theoretical strategies have been developed for dealing with helix-coil transitions in the presence of small ligands, for example, ions. Bond et al.11 have used the framework of preferential interactions to analyze the melting of duplex and triplex helices as a function of salt concentration, but the application of this method is designed to determine the effects of ionic environment on the stability of the polynucleotides. These theoretical models have little relevance to the binding of drugs, large proteins, or even cationic lipids, where multisite binding and cooperativity are likely. In the literature of the past 25 years or so, there have been virtually no theoretical papers, other than that of McGhee,~ dedicated to the rigorous analysis of the problem of the melting of DNA in the presence of binding ligands, which takes into account the coverage of large numbers of base pairs, and includes cooperativity both in the helix and coil form. For this reason we would like to give a cursory overview of the basis for the model, and the thermodynamic information that can be obtained by application of the method to analysis of DNA melting. McGhee Model In the model proposed by McGhee, homogeneous DNA is considered during melting as consisting of regions of helix, coil, helix with ligand bound, and coil with ligand bound, with some ligands free in solution (see Fig. 1). Bound ligands that are nearest neighbors can interact, so that cooperativity becomes a parameter in the model. The bound ligand on the helical duplex is allowed to cover nh contiguous base pairs, which makes these bases inaccessible to other ligands. An 11 j. p. Bond, C. E Anderson, and M. T. Record, Jr., Biophys. J. 67, 825 (1994).

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association constant, Kh, defines the strength of binding of the ligand to a stretch of helix base pairs, and a cooperativity parameter, COh,is defined as the equilibrium constant for moving isolated ligand into contact with a nearest neighbor. Similar parameters are defined for binding to the coil form of the polynucleotide, that is, Kc, no, and coc. In addition to these binding parameters, it is necessary to define parameters that characterize the melting of the DNA. The polynucleotide is modeled to be infinitely long, and is treated according to the nearest neighbor Ising model. 2 Two parameters, s, the equilibrium constant for forming a base pair, and or, the nucleation parameter, are needed in order to define the helix-coil transition. The equilibrium constant, s, varies with temperature according to the van't Hoff equation, so that the enthalpy of melting of the DNA is required in calculations of the melting curve. The rest of the problem is then to define the partition functions for the system and work out methods for obtaining melting curves from the solutions to the partition function equations. The details of the method for obtaining theoretical melting curves are presented in the original article by McGhee.l The basic approach drew on the work of Lifson, 3 who worked out an estimate of the system partition function using sequence-generating functions. Lifson considered a polymer consisting of N units, each unit of which could exist in one of several states. The microscopic states of the polymer were described in terms of lengths of alternating sequences of different types. Lifson reasoned that for a polymer of N units, a quantity xl could be defined such that

for sufficiently large N, where Z is the system partition function. Lifson further defined sequence-generating functions, U(x), for sequences of different types in the form oo

U(x) = ~ , uix

i

(2)

i--1

where ui is the contribution to the statistical weight of a microscopic state made by a sequence of a particular type of length i. Because in Eq. (2) the summation is from 1 to ~ , the equation is a power series that converges to a closed form. 3 A matrix M, was then defined, the elements of which represent all possible types of sequence-generating functions, and has the property that the largest root of the equation 11 - M ( x ) l

= 0

(3)

is xl. McGhee I applied the approach of Lifson to the helix-coil transition of DNA in the presence of a ligand that binds to DNA. The statistical weights, ui, in Eq. (2) are defined with the coil base pair as reference, the statistical weight of

[9]

THERMALDENATURATION

197

which is defined as one. The statistical weight of the helix base pair is s, the equilibrium constant for forming a helix base pair from a coil base pair. The statistical weight of the helix-ligand complex relative to the coil base pair is SnKhL,where Kh is the intrinsic association constant of ligand for a stretch of nh base pairs, and L is the free ligand concentration. For the coil-ligand complex the statistical weight is KcL, where Kc is the association constant of the ligand for a stretch of nc bases in the coil form. As mentioned above, ligand-ligand cooperativity is accounted for through the parameters Wh and Wc, for the helix and coil interactions, respectively. The nucleation parameter, or, can be considered the statistical weight of a boundary between helix and coil regions. Using these statistical weights, McGhee then wrote sequence-generating functions for each of the four possible configurations of the DNA: a sequence of one or more base pairs, a sequence of coil bases, a sequence of base pairs bound by ligand, and a sequence of coil bases bound by ligand. The power series limits of the sequence-generating functions were used to construct the matrix, M, and the function f(x) = I1 - M(x)l = 0 was evaluated. For a particular configuration with an associated statistical weight the average number of bases in that configuration can be determined. For example, the average number of base pairs that are helical (without bound ligand), Y, with the statistical weight of the base pair equal to s, is -

-

Y-

OlnZ 0 Ins

(4)

To find the average fraction of base pairs in this configuration, the average would be divided by N, and recalling that Z ~ x~, the average fraction becomes Y

0 lnxl

- N - Olns

(5)

The quantities of interest, for example, the fraction of base pairs in a particular configuration Y, with statistical weight y, can also be obtained by differentiation of Eq. (2), f(x) = 0, used to define xl:

N

Of(x)/O lnxl Of(x)/O Ins

(6)

where the derivatives are evaluated at xt. McGhee I wrote a computer program that solved for xl at specific values of s, or, nh, Kh, n~, Ko the cooperativity parameters (Oh and we, and the free ligand concentration, L. In a later version McGhee also included temperature dependence of the K values, provided the AH values for the association equilibria are available. From this information the fraction helix, fraction coil, fraction bound ligand, etc., at each temperature can be determined over a specified range. Thus, the entire melting curve can be calculated for a set of parameters. To relate the calculated curves to experimental conditions, the DNA concentration, value of Tm for the free DNA,

198

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AHm, the enthalpy of melting of the DNA, and an estimate of cr for the transition must be specified. The presently available computer program offers the option of whether the user wants the total or free ligand concentration to remain constant during the transition, and the output returns the fraction helix, fraction coil, and concentration of free ligand at each temperature over the specified temperature range for the transition. Before looking at some specific applications for which experimental data are available for comparison with the model, it is informative to examine what the theory predicts about the effects of the magnitudes of some of the parameters on the characteristics of the thermal transition curves. What follows is a summary of the extensive analysis of the model presented by McGhee in the original article. When a ligand binds to helical DNA the free energy per base pair is lowered, and thus the Tm for the transition is raised. This free energy of binding, AGb, is made up of two contributions, an intrinsic binding free energy, AGint, which is related to the basic interactions responsible for the binding process, and a mixing free energy, AGmix, which is proportional to the number of different ways the ligand can be arranged on the helix. 1 Thus, for binding of Bh sites of size nh on a helix Of Nh total sites, the binding free energy is AGb = BhAGint -[- AGmix

(7)

Assuming no cooperativity, the intrinsic free energy, AGint, = - R T ln(KhL), and the mixing free energy, AGmix, = - R T ln(~), where f2 is a combinatorial term related to the number of different arrangements of the ligand on the helix. ~2 =

(Nh -- Bhnh + Bh)! (Nh -- Bhnh)!Bh!

(8)

Using these basic definitions and the calculations developed by McGhee] it is possible to make predictions about how the melting temperature and shape of the transition curve will be affected by ligand binding to the helix. Because similar equations apply to binding to the coil form, it is possible to calculate the effects that ligands will have on destabilizing the helix due to binding to the coil form. Below we summarize some of the consequences of ligand binding to the helix and coil forms separately, and then consider the case in which binding to both forms of the polynucleotide occurs.

Effects of Ligand Size Keeping the intrinsic free energy, KhL, constant, the effect of ligand size, nh or no affects the mixing free energy. For helix binders, the smaller the size of the ligand, the larger the number of ways the ligand can be arranged on the helix, and thus Tm will be increased to a greater extent for a small than for a larger ligand. For example, with nh -----30, the melting temperature of a specific transition would be increased by 2 °, whereas for the smallest ligand, nh = 1, the Tm is increased by

[91

THERMALDENATURATION

199

about 19 °. For coil binders (helix destabilizers), Tm will be decreased accordingly, although because of the statistical problem that when DNA melts two coils are produced, the effects on Tm will be about double for a given value of n and KL.

Effect of lntrinsic Binding Constant and Free Ligand Activity Because the intrinsic free energy depends on both K and L, the effects of both on the melting temperature will be in the same direction. At a given free ligand concentration, increasing K will cause more extensive binding to the helix form. This greater degree of saturation of sites will lead to fewer ways to rearrange ligands along the chain, so AGmix will be less, but the intrinsic free energy becomes increasingly negative, leading to increases in Tin. At higher values of Kh the effects on Tm will be less for larger ligands, because at higher levels of saturation with large ligands the number of gaps between bound ligands becomes smaller than the ligand size. The effect of free ligand concentration on the melting temperature will be the same as that of K; that is, as L increases at a given value of K, melting temperature will increase correspondingly, and the increases will be more pronounced for smaller ligands. As mentioned above, for coil binding there will be destabilization of the helix (decreasing Tm) corresponding to increases in K or L, and again for smaller ligands the effects will be more dramatic.

Effects of Ligand-Ligand Cooperativity The above-described calculations were done assuming that there was no cooperativity in the binding of the ligands. If oJh is increased from the noncooperative value of 1, the melting temperature of the helix increases. This effect is due to the fact that cooperativity effectively increases the intrinsic binding constant from Kh to Khcoh, and thus there is an added free energy of stabilization of the helix. Cooperativity effects are greater for smaller ligands because there are more potential ligand-ligand contacts than ligand-DNA contacts in this case. An interesting point made by McGhee I is that with increasing COhthe slopes of the melting curves become greater; that is, the transitions are steeper around Tm. Also, from the calculations using sequence-generating functions, it is possible to determine the fraction of unbound helical regions, which increase with increased cooperativity in ligand binding. Both of these effects are caused by the ligands now binding in longer contiguous stretches, leaving parts of the helix clear of bound ligands. Cooperativity in coil binding is again destabilizing of the helix, and thus Tm will decrease more with cooperative binding than without, and the smaller ligands have a greater effect.

Titrations with Constant Total Ligand Concentration The discussions above have been for conditions in which the free ligand concentration in the solution is constant. A more realistic situation is for a typical ligand titration, in which melting points are determined for a series of solutions

200

BIOPHYSICAL APPROACHES

[91

of fixed DNA concentration with increasing total ligand. In the computer program developed by McGhee an iteration cycle is added to the calculations such that Lfree + Lbound = Ltot is determined, and thus for a given DNA concentration Lfree and Lbound are known for each point in the transition. It is informative to see the melting curves generated by McGhee I for this situation (see Fig. 2). The data are generated for total DNA concentration of 5 x 10 -5 M base pairs, with total ligand concentrations increasing from 5 x 10 - 7 to 1 x 10-SM. Figure 2 shows cases for

I 1.0

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0.0 1,0

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b'

c'

~ f

("

o

0

¢..

d

._o "~ 0.5

;

i

d

LL

0.0

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!

,

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I

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30

40

50

60

70

8O

....

Temperature, (°C) FIG. 2. Calculated melting curves of DNA at a series of total ligand concentrations, ranging from 10 to 200% saturation (curves f-a) of either helix or coil forms. (Coil curves are the primed letters.) Site size was fixed at 10 bp or 10 bases in the coil form. Ligand binding constants are K = I06M I (top); K = 108M -1 (middle); K = 101°M - j (bottom). DNA-only curve is the dashed line. [Taken from J. D. McGhee, Biopolymers 15, 1345 (1976), with permission.]

[9]

THERMALDENATURATION

201

Kh at three different magnitudes: 106, 108, and 101° M -1 . The ligand is assumed to cover 10 base pairs, and no cooperativity is allowed in this case, nor is there ligand binding to the coil form. Shown in Fig. 2 are curves for the destabilizing binding to the coil form under similar conditions, except that no binding to the helix is allowed. The behavior of the melting curves is as expected (discussed above); increasing the total ligand concentration increases Tin, as more of the helix becomes saturated with ligand, and for increases in Kh the melting temperature also increases. An interesting feature of these curves is the biphasic character that develops when Kh is large, and as the total ligand increases. The sizes of the two phases approximately reflect the ligand-to-DNA ratio. This biphasic behavior, which was predicted by Crothers, l° is a consequence of the strong binding to the helix form, leading to the situation that as the DNA melts, liberated ligand can rebind to regions of unmelted helix. Ultimately, the helix has no unbound regions remaining, and the melting of the second phase corresponds to the unraveling of the high-temperature, totally ligand-bound phase. This theoretically predicted biphasic melting has been observed in several cases of ligand binding to DNA (see below). Cases in Which Both Helix and Coil Bind Ligand

Although there may be examples of ligands that bind exclusively to helix or exclusively to coil, in many cases the ligand binds to some degree to both helix and coil forms of the DNA. The effects on the melting curves of the DNA under these circumstances will be complicated by the fact that the two influences have opposite effects on Tin: ligand-coil binding leads to destabilization of the helix, whereas ligand-helix interactions cause stabilization. Thus, the exact shift in the melting temperature will depend on the relative magnitudes of KhL compared with KcL. In the absence of cooperativity and for small ligands it can be shown that the condition that leads to no shift in Tm (compensating effects) is for KcL ~- (KhL) 1/2. For larger ligands, for example, nh = nc = 10, compensating shifts occur for KcL = 0.3(KhL) 1/2. In situations in which there is cooperativity and in which the site sizes differ, it is possible to estimate that again compensation occurs at roughly square root dependence, that is, K~Lo~c = (KhLOJh) I/2. These relationships are only approximate, so the best way to deal with these situations is to calculate the curves with best estimates of the parameters. This brings us to looking at actual applications of the McGhee melting theory. Although few examples have been analyzed, we present some work for which the McGhee model works well in describing melting in the presence of binding ligands. A p p l i c a t i o n s of M c G h e e Model of Melting The major problem in calculating melting transitions for DNA, using the McGhee methodJ is in finding reasonable estimates for the large number of

202

BIOPHYSICALAPPROACHES

[91

parameters required. Recall that for a complete description of the ligand-DNA system, values OfKh, nh, COh,Kc, nc, toe, and properties of the melting transition for the native DNA without ligand are needed. The latter properties include Tm, A H m , and the nucleation parameter, ~. The program available now also includes provision for allowing for temperature dependence of the two equilibrium constants from knowledge of the enthalpies of binding to both helix and coil form. With so many parameters it is virtually impossible to use any kind of least-squares algorithm to determine all the parameters. Thus, to use the model, it is useful to have values or estimates for as many of the parameters as is possible. Values of Tin, AHm, and are either available or can be determined from a melting transition in the absence of ligand. Often values for Kh and nh for binding to the helix are available, and if the binding is cooperative, there are procedures for getting accurate values for the cooperativity parameter, Wh.9 It is often possible to estimate site size by using molecular models of the ligand, although this is not easy with protein ligands. Thus, the number of parameters that need to be varied in order to fit the model to data can be reduced to two or three, and often these are parameters relating to binding to the coil state. Below are some examples of work that has used the McGhee model for the analysis of thermal melting of DNA in the presence of different types of ligands.

Melting of Polyld(AT)] in Presence of Netropsin Netropsin is a drug that is a minor groove binder and its interaction with DNA has been studied extensively, z2,13 McGhee reported melting experiments for poly[d(AT)] in the presence of netropsin, and applied his model for melting to this system. 1 In the analysis presented the netropsin was assumed, on the basis of previous work, 12 to bind noncooperatively to the helix, and not at all to the coil form. Thus, to calculate the melting curves, only nh and Kh needed to be adjusted to fit the experimental data. Figure 3 shows the calculated and experimental curves for melting of poly[d(AT)] at seven different levels of netropsin binding, ranging, in drug: base pair ratio from 1 : 36.6 to 1 : 1.9. Using g = 0.0001, AHm = 8 kcal/mol, and T m = 27 ° under the conditions employed, the best values for the adjustable parameters turn out to be nh : 4 and Kh : 5 x 108M -l for netropsin binding to the helix. These values compare well with previously determined parameters by direct measurement of n and K, 3 and about 5 × 106, respectively, for conditions with somewhat higher salt content in the medium. As is especially clear from curve d in Fig. 3, there is biphasic character in the melting transition. This behavior is most likely due to reequilibration of the ligand during melting to unbound sites on the DNA, leading ultimately to binding saturation, and a second phase that is predominantly due to melting of netropsin-DNA complex. Curve g in Fig. 3 is essentially at 200% saturation, and it is interesting that the slope of the melting

12 R. M. Wartell, J. E. Larson, and R. D. Wells, J. Biol. Chem. 101, 6719 (1974). 13 C. Zimmer, Plvg. Nucleic Acid Res. Mol. Biol. 15, 285 (1975).

[9]

THERMAL DENATURATION I

I

[

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I

I

a

203 I

b

e

/

de

I

1

g

1.0 Q

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(O t- 0.5 o

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20

30

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40

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80

I

I

90

100

Temperature, (°C) FIG. 3. Comparison of experimental and calculated melting curves for poly(dA-dT) - poly(dT-dA) in the presence of netropsin. Points are experimental and curves are calculated for different drug:base pair ratios: (a)0; (b) 1:36.6; (c) 1 : 14.9; (d) 1:7.6; (e) 1:5.1; (f) 1:3.7; (g) 1: 1.9. Curves are generated with a site size of 4 and a helix binding constant of 5 × 108. [Taken from J. D. McGhee, Biopolymers 15, 1345 (1976), with permission.]

curve is similar to the DNA melt in the absence of ligand, curve a. This curve should correspond to the melting of pure netropsin-DNA complex. The analysis of netropsin binding by the McGhee model seems to be quite consistent with previous work, and lends support to the use of this melting model to obtain thermodynamic information about the binding process.

Melting of DNA in Presence of Cationic Lipids The thermodynamics of binding of cationic lipids to DNA have been studied in detail because of interest in the condensed states of DNA that result from aggregation of the lipid complexes. 14-17Many of the condensed structures have potential application as vehicles for DNA delivery in gene therapy applications.18-21 Thermal melting transitions of Escherichia coli DNA in the presence of hexadecyltrimethylammonium bromide (HTAB) were studied as a part of a broader investigation of the thermodynamics of the cationic ligand-binding process.t4 In this study the binding parameters for the helical DNA were independently measured, the Kh and AHh for binding of HTAB to helix being determined from isothermal titration calorimetry experiments. These binding experiments could also be used 14 C. H. Spink and J. B. Chaires, J. Am. Chem. Soc. 119, 10920 (1997). 15 S. M. Mel'nikov, V. G. Sergeyev, and K. Yoshikawa, J. Am. Chem. Soc. 117, 9951 (1995). 16 K. Shirahama, K. Takashima, and N. Takisawa, Bull. Chem. Soc. Jpn. 60, 43 (1987). 17 K. Hayakawa, J. P. Sanaterre, and J. C. T. Kosaak, Biophys. Chem. 17, 175 (1983). 18 p. L. Feigner, Adv. Drug Delivery Res. $, 163 (1990). 19 N. Z h u , D. Ligget, Y. Liu, and R. Debbs, Science 261,209 (1993). 20 y. Xu and E C. Szoka, Biochemistry 35, 5616 (1996). 2l X. Gao and L. Huang, Bioehemistrv 35, 1027 (1996).

204

BIOPHYSICALAPPROACHES

[9]

to estimate values for n h and COhby casting the results in a form that could be analyzed by the McGhee-von Hippel isotherm. 9 The results for helical binding are as follows: Kh = 1.5(4-0.3) x 103M-I; COh= 60 4- 10;

nh = 1.4 4- 0.3;

AHh = 3200 4- 300J/mol

These numbers reflect the characteristics of binding of a positive alkylammonium ion to phosphate sites on the DNA. The small nh is expected for simple ion exchange of the lipidic ion for sodium ion territorially bound to the helix. The relatively high cooperativity is due to the tendency of the 16-carbon alkyl groups to associate hydrophobically as binding density increases. The small values of Kh and AHh for binding to the helix again suggest that the association is territorial rather than specific site binding, and the enthalpy is likely a consequence of hydrophobic association, similar to that observed for detergent association reactions. 14 There are no data for binding to the coil form, although some reasonable estimates for some of the coil parameters can be made. First, it is unlikely that nc would be very different from unity, because the helical binding site size is near one. With reduced charge density in the coil form it is not expected that site size would increase. Also, because the phosphate sites are farther apart in the denatured state, cooperativity should be reduced significantly. Thus, in trying to model the melting transitions for E. coli in the presence of HTAB, the parameters Kc and AHc for the coil form become the main variables to adjust. Figure 4 shows the melting 0.06

0.06

0.04

I--

0

0.06

0.03

0.02

////\\\to,12

/

0.0I

0.00

o -0.01 100

Temperature FIG. 4. UV melting transitions in the presence of HTAB of 0.24 mM base pairs of E. coli DNA, plotted as the derivative of absorbance at 260 nm. Numbers correspond to the millimolar concentration of HTAB in the solution. [Taken from C. H. Spink and J. B. Chaires, J. Am. Chem. Soc. 119, 10920 (1997), with permission.]

[9]

THERMALDENATURATION

205

transitions for solutions of HTAB with E. coli DNA, plotted as the derivative of the absorbance at 260 nm as a function of temperature. The derivative is presented to clearly show the unique biphasic melting observed for this system. Note that the stoichiometric ratio of HTAB to DNA phosphate sites does not exceed 0.6, because beyond this ratio the lipid-DNA complex condenses and precipitates out of solution. To model the melting transition, data for the DNA alone were used to evaluate Tm and or, and the AHm was taken from calorimetric measurements of the melting enthalpy 14. From the data available then, the initial parameters that were used to model the melting curves were as follows: nh = 1,

Kh = 1 5 0 0 M - I , ~oc=l,

Oah = 60,

~r=0.0025,

AHh = 3200cal/mol,

T m = 7 3 °,

nc = 1,

AHm=33.5kJ/mol

The missing parameters, AHc and Kc, were then varied to obtain an optimal fit with the experimental melting transitions. Once some sort of convergence was obtained, then the other parameters were also allowed to vary, not only to try to improve the fit, but also to obtain some idea of the sensitivity of the fits to the data to the variation in parameter values. Figure 5 shows the optimal fits of the

1.0!I -0

N

E

.~ 0 •~ IJ_

i

i d

I

:

""..... . " ," ' e

:"..""

f

0.8

""

o.6 0.4

o.2 0.0

40 1.0 "o 0.8

i

I

60 i

~

i

I

I

80

100

i

a "'b

...=

]

,,,

0.6

E .o 0.4 ~

LL

0.2 0.0

40

60 80 Temperature

1O0

FIG. 5. Experimental (--) and calculated ( . . -) thermal melting curves for E. coli D N A of 0.24 mM concentration in the presence of varying amounts of HTAB: (a) 0; (b) 0.09; (c) 0.15; (d) 0.06; (e) 0.12; and (f) 0.18 mM. Parameters for calculations are in Table 1. [Taken from C. H. Spink and J. B. Chaires, J. Am. Chem. Soc. 119, 10920 (1997), with permission.]

206

BIOPHYSICALAPPROACHES

[91

TABLE I THERMODYNAMIC PARAMETERS FOR MELTING OF DNA IN PRESENCE OF CETYLTRIMETHYLAMMONIUM BROMIDEa CTAB (mM)

nh

nc

Kh

Kc

AHh

AHc

COb

oJ~.

0.06 0.09 0.12 0.15 h 0.18 h

1.30 1.10 1.06 1. t 0 1.07

1 1 t 1 1

1,500 1,500 1,800 1,500 1,500

10,000 10,250 15,000 14,000 16,000

3.3 3.3 3.3 3.3 3.3

-4.2 -3.3 -3.8 -4.6 -3.8

65 70 60 60 60

1 1 I 1 1

Summary Error range ''

1.1 ± 0.2

1 ± 0.2

1,500 ± 200

13,100 ± 2,000

3.3 ± 0.8

-3.8 -4- 1.5

63 ± 20

1 ± 0.5

a The parameters provide the best fit to the experimental melting curves for the following DNA melting properties in the absence of CTAB: Cpi = 0.24 mM; Tm = 73.0°; AHm = 33.5 kJ/mol; ~7 = 0.0025. Units o f K are M - I and of A H are kilojoules per mole. h In the 0.15 and 0.18 mM CTAB solutions slightly better fits were obtained with A H m set at 35.6 kJ/mol, and cr = 0.0034. ' Error range was estimated by variation in each parameter that caused a 2 c deviation from the best fit curve at fraction melted equal to 0.4.

model to the data, and Table I shows the best values for the parameters for the optimal fit. Given the rather complex nature of the melting transitions, the calculated values seem to agree well with experimental curves. To illustrate the sensitivity of the fits to the choice of parameters, Fig. 6 shows curves for the optimal fits along with the effects on the calculated curves of varying specific parameters. This establishes a range of values for the parameters that give results that are within the experimental uncertainties of the data. It was found that the model is quite sensitive to values of n and oa, with as small as 20% changes in these parameters causing 4-5 ° shifts in the temperatures. Although there is some cross-dependence of parameters, particularly between K~ and AHc, it was found that agreement with the experimental curves restricted the range of Kc to a rather narrow range (8-15 x 103 M-I), which in turn confines the range in the binding enthalpy to - 3 . 8 to - 4 . 2 kJ/mol. Table I summarizes uncertainties estimated for the parameters on the basis of variations in the parameters that keep agreement between experimental and calculated curves to within 2 ° when measured at 0.4 fraction melted. The agreement between calculated and experimental curves is good and, given the fact that five different melting curves with rather different melting profiles are involved in the fitting procedure, raises confidence in the values used for the resolved parameters. Mel'nikov et al.iS have measured the binding of HTAB to helical T4 DNA by a potentiometric method and find values of Kh of 1200 M- I and cooperativity parameter, COb,equal to about 80. These values are in good

[9]

THERMAL DENATURATION •

1.0 Variation of ttH. (fixed value 800 caIImole)

"O

AHh

0.6

i

.

= 400 c a ~ . . . "

~::~J'..:'~/' ""

Variation of K c (best fit

//

40

•

L

= 9000 M"1)

.

;-~"

:i;iiii;! K~= 8500M / " I~ = 9500 M" 4

0.4 1

""~H. = 1200 cal/mole

0.2[ 0.01

5'0 =

"Variation of AH¢

8'0

7'0

=

•

(best fit

i

8'0 •

i

= 500 cal/mole)

9'0 •

t

loo

;0

40

•

-

,

0'0 •

7'0

,

,

AHc =1000c a l ~

0.4

~

0.2 0.0

8'0

.. c ' "

Temperature,

,

•

i

•

~=55

0.81

t%=65

o.4 I

..'" ,~H¢= -500 cal/rnole

6'0'7'0

.

,00

= 60)

0.8

,..:.-' _ - / , . . ,..,,:-'" ,""

11) 0.6

9'0

;0

1.01

~ 7 " -

Variation of o~n (best fit

0.8

,o

i

/,

~

0.2

.

0.8 1

0 . 0 - -

LL

J

o.6 I

0.4

1.0

.

1.0 I

0.8

LL

t

207

0.2~ o.01

8'0

9'0

loo

40

5'0

8'0

7'0

8'0

9'0

Temperature

FIG. 6. Effects of varying parameters on the calculated melting transitions for E. coli DNA. Solid curves are the experimental melting curve, and dotted curves are calculated. Parameter values are indicated in the individual panels. [Taken from C. H. Spink and J. B. Chaires, J. Am. Chem. Soc. 119, 10920 (1997), with permission].

agreement with the numbers reported above. The benefit of examining the melting curves and using the McGhee model is that thermodynamic data for binding to the coil form can be resolved. Because it would not be possible to analyze the melting transitions without taking into account coil binding in this case, there is confidence that the coil parameters make significant contributions to the overall behavior in transformation from helix to coil. The thermodynamic parameters for both the helix and coil forms are interesting from the point of view that although the binding to helix is more favorable by a factor of about 7 when including the cooperative nature of the helix association, both helix and coil binding involve small enthalpic contributions to the free energy. 14 Thus, binding to both the helix and coil involve substantial entropic contributions that most likely arise from hydrophobic interactions. In the case of helix binding the cooperative hydrophobic association between ligand alkyl groups is probably the contributing factor in the

loo

208

BIOPHYSICALAPPROACHES

[9]

large entropic effect. For the coil form hydrophobic interaction of the exposed bases with the alkyl groups of HTAB would seem a likely source of the entropic contribution to binding of the ligand. It would be difficult to sort out these effects without having estimates of the coil parameters obtained from the McGhee analysis. Melting of Polyld(AT)] . poly[d(TA)] in Presence of Trivalent Cations Plum and Bloomfield22 applied the method of McGhee to studying the binding of three trivalent cations, spermidine, a methylated spermidine derivative, and hexaammine cobalt(III), to poly[d(AT)], poly[d(TA)]. The impetus for the study was the idea that small oligomeric cations, such as spermidine, may influence the nonspecific binding of proteins to DNA by reducing charge density. Also, these small multicharged cations can interact with single-stranded DNA, which alters the stability of the duplex and thus may mediate protein interactions with both helix and coil conformations. In applying the McGhee method to these cases noncooperative binding was assumed for both helix and coil forms, and the binding constants were assumed to be temperature independent. The parameters for melting of DNA in the absence of ligands, Tm and ~r, were determined from experimental melting curves, whereas AHm was taken from a published calorimetric value. (Tin = 35 °, AHm = 7.1 kcal/mol, and cr = 5 x 10-5 provided the best description for melting in the absence of ligands.) In the presence of ligands curves were calculated and compared with experimental curves, using a trial-and-error approach. For spermidine and hexaammine cobalt(III), for which there are binding data from equilibrium dialysis experiments, the resulting parameters were in reasonably good agreement. The Kh and nh values for spermidine were found to be 9 x 105 M -1 and 2.2 base pairs, respectively, compared with 5.7 x 105 M -1 and 3 base pairs from the equilibrium dialysis data. The helix parameters determined for the methylated spermidine derivative were similar to those for spermidine. The three trivalent cations had similar affinities for double-stranded DNA (1-9 × 105 M -l) and similar site sizes (1.4-2.2 base pairs). All three ligands bound to single-stranded DNA, but differences in affinities for the coil form were pronounced, with spermidine binding most tightly (1.3 × 10 4 M - l , site size of 4.5 bases) and hexaammine cobalt(III) most weakly (600 M -1, site size of 6 bases). This observation of different affinities for single-stranded DNA was especially interesting in light of the fact that the three ligands have identical charges. An analysis of the charge spacing in the single-strand form, based on counterion condensation theory, suggests that spermidine and hexaammine cobalt(III) prevent some unraveling of the coil, producing smaller separation between phosphate charges. 22 G. E. Plum and V. A. Bloomfield, Biopolymers 29, 13 (1990).

[9]

THERMALDENATURATION

209

Melting Studies of DNA in Presence of i l l Histories The method of McGhee was used to analyze the binding of two H1 histone proteins to DNA. 23 The H1 (or linker) histones are involved in formation and maintenance of chromatin structure in eukaryotic cells. The interactions between H1 histones and DNA are thought to be primarily, if not entirely, electrostatic. Application of the McGhee approach was successful only when using DNA homocopolymers. Data obtained using heterogeneous sequences of DNA were consistent with the idea that H1 histones exhibit sequence selectivity to at least some degree, and therefore do not necessarily bind to heterogeneous DNA as if it were a lattice of overlapping, equivalent sites. Several homocopolymers were used in this study of histone binding. Values of Tm, the temperature at the midpoint of the helix-coil transition, were determined from DNA-only curves, and published values of AHm, the enthalpy of melting per base pair, were used for the homocopolymers. Values of a, the nucleation parameter, were determined empirically. For two of the DNA polymers, poly(dGdA)- poly(dC-dT) and poly(dC-dA), poly(dG-dT), in calculations of the DNAonly melting curves, a larger value was used below the Tm than above (5 or 2 x 10 -5 and 5 or 1 x 10 -4, respectively). For poly(dA-dT) - poly(dT-dA) the value used for cr was 1 × 10 -4 for the DNA-only transition. The values of cr in the presence of H1 histones were also varied. In the presence of the protein the same value was used throughout the transition for poly(dG-dA) • poly(dC-dT) (5 x 10 -4) and for poly(dC-dA), poly(dG-dT) (5 x 10-5). For poly(dA-dT), poly(dT-dA) the values for ~r were 1-5 x 10 - 4 in the presence of the protein. The rationale for these adjustments is that the probability that a single-stranded base is contiguous with a base pair, which is reflected in the value of a, is altered as ligand occupancy of the DNA lattice increases. For the H1 histone binding a simple model was used for calculations, that of noncooperative binding to helix only (Kc = 0), with a AH for binding of zero. Curves were calculated and compared with experimental curves, with the best correspondence being determined by trial and error. For one of the H1 histone variants, Hit, curves that corresponded well to the experimental curves were calculated by using values of 10-12 base pairs for nh and 108-109 for Kh (see Fig. 7). For the variant H1 °, the values used in calculated curves were 20-30 base pairs and 101°-1014 for Kh. Unfortunately, there are few, if any, estimates of these parameters for the H1 histones in the literature. Consistent results for binding site sizes, 15-25 base pairs, have been obtained using heterogeneous DNA (N. M. Mamoon, Y. Song, and S. E. Wellman, unpublished results, 2000). Binding of HI histones to heterogeneous DNA is not described by the McGhee-von Hippel neighbor exclusion model, 9 and so comparable affinity constants are not available. 23S. E. Wellman,Y. Song, and N. M. Mamoon,Biochemisn3, 38, 13112 (1999).

210 1.0

BIOPHYSICALAPPROACHES

[9]

"

0.8

oOO°°°°°

"8 0.6

ed

,'0 i s ~'0 ~'5 6'° g5 ,;'

/

7'o ;5

8'0 is' 9'0 9'~ ~'~ 6'° 6'5'7'0 ~5 6'0 ;~'9'0 Temperature

FIG.7. Calculated and experimentalmelting curves for three different DNAs in the presence of Hit histone ligand. (A) Poly(dA-dT).poly(dT-dA); (B) poly(dC-dA),poly(dG-dT); (C) poly(dGdA). poly(dC-dT).See text for parameterdetails. [Takenfrom Wellmanet al. 38, 13112 (1999), with permission.].

Attempts were made to apply more complex models for the histone binding to DNA. Curves were calculated that included binding to both helix and coil regions, but the effect of incorporating nonzero values for Kc could be offset by an increased value for Kh. Similarly, using nonzero values for the enthalpy of binding could be offset by altering values of Kh. These results clearly demonstrate that because values of many of the parameters are correlated, this method is not well suited to elucidating details of binding when information is lacking for some of the needed parameters. The McGhee approach is most useful for systems in which characteristics of binding are well understood and for which there are good estimates of at least some of the binding parameters. In the case ofhistone binding to homocopolymers the approach was nevertheless useful. The data unambiguously demonstrate that affinities of H 1 histones for single-stranded DNA are considerably weaker than those for double-stranded DNA. This analysis was also useful for comparison of binding characteristics. The two variant proteins are clearly different from one another (different apparent intrinsic affinities and different site sizes), and both proteins have higher affinity for one of the homocopolymers, poly(dAdT). poly(dT-dA), than for the other two.

Conclusions As the illustrations given above demonstrate, the application of the McGhee analysis of thermal melting curves for ligand binding to DNA can provide thermodynamic information about the binding process. The information that becomes accessible makes the experimental melting transitions more useful, and thus provides

[9]

THERMALDENATURATION

21 1

an additional tool for examining the characteristics of the binding process. The model helps to support ideas about ligand binding to DNA that are difficult to obtain by other methods, particularly with regard to the influences of binding on the coil form. The need for so many parameters in the analysis has drawbacks, but on the other hand, these resolved parameters can be important in describing the nature of ligand association reactions. The relative magnitudes of binding parameters for a series of related compounds can be used to order the effects that the ligands exhibit, and make sense out of the effects of structure on the binding properties. The main drawback of the model is the large number of parameters required. Perhaps the best situation is in systems for which some binding data are available by other experimental methods. If several of the parameters are known from independent experimental measurement, then the fitting problem becomes much simpler, and more confidence can be had in the resolved parameters. It is possible to make use of molecular modeling for smaller ligands to make initial estimates of the site size parameters. Binding constants and cooperativity effects can be evaluated by independent measurements in some cases, and often judgments can be made about coil state binding on the basis of correlations with other systems that are similar. Trying to reach a minimum in the deviations of the experimental and calculated melting curves is difficult without holding some parameters fixed. However, with independent data for three or four parameters convergence can be rapid because of the sensitivity of the fitting to small variations in parameters. This sensitivity to parameter values limits the range of numbers that will work in reaching convergence, an advantage in trying to sort out the best values for the binding parameters. Nonetheless, even with the above caveats, because melting experiments are relatively easy to do, applying the McGhee model to DNA melting for ligands such as drugs, cationic lipids, or small proteins can give valuable information about the association of the ligands with DNA.

Availability of Computer Program With the permission of James McGhee the authors of this article can make a compiled ForTran program available to users who might be interested in using the program to analyze melting curves. An uncompiled version is also available for those who might want to make alterations or fit the procedure into existing software. Acknowledgments The authors express appreciation to Dr. James McGhee for providing the original programsfor doing these calculations, and for his permission to distribute the program to others. We also thank Dr. Jonathan B. Chaires for providinginspiration to pursue the questions and problems relating to the analysis of DNA melting by these methods.

212

BIOPHYSICALAPPROACHES

[10] K i n e t i c s o f B i n d i n g Studied by Stopped-Flow

[ 1 O]

of Hoechst Dyes to DNA Fluorescence Techniques

B y SOPHIA Y. BREUSEGEM, FRANK G . LOONT1ENS, PETER REGENFUSS, a n d ROBERT M . CLEGG

Introduction The dye Hoechst 33258, termed p-OH Hoechst in this article, is a bisbenzimidazole derivative with phenolic and N-methylpiperazine groups at opposite ends (Fig. 1A). The dye binds with high affinity and specificity to (A-T)-rich sequences in the minor groove of B-DNA. 1,2 In aqueous solution, the free dye molecule is only weakly fluorescent, whereas its complex with A-T sequences is brightly fluorescent, making it an excellent chromosome stain in fluorescence microscopy. The dye-DNA structures determined both by X-ray crystallography3-6 and solution nuclear magnetic resonance (NMR) 7'8 show the molecular details of the complex formed and make the p-OH Hoechst-DNA interaction an excellent model system with which to study strong and sequence-specific binding in the minor groove of DNA. To gain insight into the binding mechanism we have complemented the structural information with the kinetics of complex formation and dissociation. When we originally investigated the kinetics of p-OH Hoechst binding to the strongest binding sites on calf thymus DNA, using stopped-flow experiments,9 we obtained a kinetically defined association constant gkin that was two to three orders of magnitude larger than the equilibrium association constant Ka found in the literature. These unexpected kinetic results motivated us to revise the published equilibrium binding data with some polymeric DNAs. 1° In addition, we determined the affinity of the dye for small, well-defined binding sites in oligomeric duplexes. Ii I C. Zimmer and U. Wahnert, Prog. Biophys. Mol. Biol. 47, 31 (1986). 2 S. Frau, J. Bernardou, and B. Meunier, Bull. Soc. Chim. Fr 133, 1053 (1996). 3 E E. Pjura, K. Grzeskowiak, and R. E. Dickerson, J. Mol. Biol. 197, 257 (1987). 4 M.-K. Teng, N. Usman, C. A. Frederick, and A. H.-J. Wang, Nucleic Acids Res. 16, 2671 (1988). 5 j. R. Quintana, A. A. Lipanov, and R. E. Dickerson, Biochemistry 30, 10294 (1991). 6 M. A. Carrondo, M. Coll, J. Aymami, A. H.-J. Wang, G. A. van der Marel, J. H. van Boom, and A. Rich, Biochemistry 28, 7849 (1989). 7 j. A. Parkinson, J. Barber, K. T. Douglas, J. Rosamond, and D. Sharpies, Biochemistry 29, 10181 (1990). 8 A. Fede, A. Labhardt, W. Bannwarth, and W. Leupin, Biochemistry 30, 11377 (1991). 9 p. Regenfuss, E G. Loontiens, and R. M. Clegg, Arch. Int. Physiol. Biochem. 97, B113 (1989). l0 F. G. Loontiens, P. Regenfuss, A. Zechel, L. Dumortier, and R. M. Clegg, Biochemistry 29, 9029 (1990). 11 F. G. Loontiens, L. W. McLaughlin, S. Diekmann, and R. M. Clegg, Biochemistry 29, 182 ( 1991 ).

METHODSIN ENZYMOLOGY.VOL.340

Copyright© 2001 by AcademicPress All rightsof reproductionin any formreserved. 0()76-6879/0(I$35.00

[ 10]

HOECHSTDYE BINDINGKINETICS

213

A

/

H3CH ' I V

H

p-OH (Hoechst 33258): R= ~ m-OCH3, p-OH:

O

H

R = --~OH k

OMe m-OH:

R= OH

m-OH, p-OCH3:

R =~ O M e OH OH

bis-m-OH:

R= - ~ OH

B

5'-CGCGAATTCGCGTT

III III III III II If II II Ill III IIIIII 3'-GCGCTTAAGCGC T FIG. 1. Structure of dyes and DNA hairpins. (A) Structure of p-OH Hoechst and the Hoechst dye derivatives p-OH, m-OCH3 Hoechst; m-OH Hoechst; m-OH, p-OCH3 Hoechst; and bis-m-OH Hoechst. (B) AATF DNA hairpin. The AATr binding site for the dye is highlighted in boldface. The other sequences investigated were incorporated within a similar hairpin structure.

Fluorescence titrations at equilibrium with nanomolar dye concentrations confirmed our kinetic data, in particular with a Ka of 7 x 108 M-1 for p-OH Hoechst binding to the AATT site at 6°, two orders of magnitude higher than reported in the earlier literature. These results in turn motivated us to look at the minor groove binding of p-OH Hoechst in more detail, using both fluorescence titrations at equilibrium and kinetic experiments, and modifying both the DNA sequence and the dye structure. The DNA sequence and structure can be modified in several ways. Equilibrium titrations with oligomeric duplexes containing modified exocyclic base substituents were published earlier} 1 Here we discuss the kinetics of the binding of p-OH Hoechst to several polymeric DNAs and to (A]T)4 sites incorporated in five DNA

214

BIOPHYSICALAPPROACHES

[ 1 O]

hairpin structure as shown in Fig. 1B. Variation in the possible DNA contacts or in their arrangement could have large effects on the dye affinity and its association and/or dissociation rate. The dye molecule was also modified by introducing p- and m-hydroxy and methoxy groups in its phenyl moiety (Fig. 1A). Again, the availability of different hydrogen bond donor and acceptor groups in the dye molecule could modulate its affinity and kinetics. In particular, NMR and molecular modeling had predicted additional hydrogen bonding between an m-OH substituent and the exocyclic amino group of guanine at position 4 of the d(CGCGAATTCGCG)2 duplex when bound at its central AATT site. 12 For all the Hoechst dye-hairpin complexes we studied, the association rate parameter ko, is comparable (between 0.68 x 108 and 2.86 × 108 M 1 sec 1), suggesting a diffusion-controlled association reaction in which the dye molecule has immediate access to the binding site in the DNA minor groove. In contrast, the dissociation rate parameter koffvaries over four orders of magnitude (between 0.012 and 96 sec -I) and determines the relative affinity of the Hoechst dye molecules for the (A]T)4 site. The poorer-binding sequences with a TpA step (TATA and TTAA) have a kofffor p-OH Hoechst that is about 200 times faster than with the strongest AATT binding site. The slowest koffrate found (0.012 sec -1) is for the tight binding of the bis-m-OH Hoechst derivative to the AATT sequence. In this case the longer lifetime of the complex can be correlated with the greater number of interactions in and near the minor groove, involving hydrogen bonds with its two m-OH groups. Practical Aspects

Instrumentation and Data Analysis In the stopped-flow 13 instrument ]SLM-Aminco (Thermo Spectronic, Rochester, NY) Milli-stopped-flow reactor], excitation is at 366 nm [Zeiss (Thornwood, NY) PMQII monochromator], using a 150-W Hamamatsu (Tokyo, Japan) L2482 Hg/Xe lamp, stabilized by an Oriel (Stratford, CT) 68805 power supply. Fluorescence emission is measured through a Schott (Iserlohn, Germany) KV418 cutoff filter in front of an EMI-Thorn 9558 photomultiplier (DE Technologies, La Madera, NM). This output is divided by the output of an IP28 reference photomultiplier and the ratio is fed into the data acquisition system [National Instruments (Austin, TX) Lab-NB 12-bit A/D converter with a 62.5-kHz maximum sampling rate]. Four hundred to 2000 data points are collected per trace. The pneumatic drive is operated at 4 to 5 atm and delivers 45/zl of each of the reactants through the 32-#1 (10 × 2 mm) observation cell. The dead time is estimated as 2 to 3 msec. All experiments are done at 20°under pseudo-first-order conditions with 12 j. A. Parkinson, S. E. S. Ebrahimi, J. H. McKie, and K. T. Douglas, Biochemistry 33, 8442 (1994). 13 Q. H. Gibson, Methods Enzymol. 16, 187 (1969).

[10]

HOECHSTDYE BINDINGKINETICS

215

DNA sites in excess; the buffer after mixing is 50 mM Tris, 100 mM NaC1 (pH 7.5). For a given concentration, up to 25 traces are averaged (LabVIEW; National Instruments) and the change in fluorescence is analyzed as the sum of a maximum of three exponentials [LabVIEW; KaleidaGraph (Synergy Software, Reading, PA)].

Sample Preparation Milli-Q water (Millipore, Bedford, MA) is used and care is taken to avoid contamination with dust particles. Buffers are kept in minimal contact with plastic surfaces (containers, pipette tips) to maintain a low buffer blank. Glassware is cleaned with Nochromix (Godax Laboratories, Tacoma Park, MD). Low-buffer blanks are obtained with NaC1 ("Fractopur") and Tris (GR buffer substance) from Merck (Rahway, NJ). Buffer and HC1 solutions are decanted fresh at least once each day. DNA is dissolved in 10 mM Tris, 10 mM NaC1, 0.1 mM EDTA (pH 7.5). Calf thymus DNA (sodium salt, type V; Sigma, St. Louis, MO) is sonicated. Poly[d(A-T)], poly(dA) • poly(dT) (both from Boehringer Mannheim, Indianapolis, IN), poly[d(A-5BrU)] (Pharmacia, Piscataway, NJ), the AATT DNA hairpin (Midland, Midland, TX; 6260 1.98 x t05 hairpin M I cm-1), and the TAAT, ATAT, TATA, and TTAA DNA, hairpins (MWG, Bayern, Germany) are annealed before use with concentrations determined in 50 mM Tris, 100 mM NaCI (pH 7.5). Hoechst 33258 (termedp-OH Hoechst here; e338 = 4.2 x 104 M-I cm-1 at pH 7.5) is a gift from H. Loewe (Frankfurt, Germany) and the p- and m-phenyl-substituted Hoechst derivatives (Fig. 1A), synthesized as reported, 14 are a gift from K. Douglas (Manchester, England). Each dye is dissolved in water (1-2 mg/ml) and its concentration is determined in a polystyrene cuvette (OD33s about 0.7; 50 mM Tris, 100 mM NaC1, pH 7.5) after mixing with a silanized glass stirrer. Before mixing in the stopped-flow instrument, DNA is diluted into 100 mM Tris, 100 mM NaC1 while a Hoechst stock solution is diluted three times successively (factors about 1 : 100) with 5 mM HC1, 100 mM NaC1. For each given concentration pair, 2 ml of DNA and 2 ml of dye solution are prepared fresh in dust-free polystyrene vials and used to rinse and fill the reservoir syringes immediately before the reaction. =

Experimental Precautions When Working with Hoechst Dyes 1. Adsorption of a diluted Hoechst solution to surfaces is prevented in 5 mM HC1 or, with neutral dye solutions, by silanizing glass, quartz, and metal surfaces with a 2% dimethyldichlorosilane solution in 1,1,1-trichloroethane (BDH, Poole, UK). Diluted solutions of the dye are always kept for less than 10 hr in polystyrene vials and contact with Parafilm, pipette tips, and polyethylene tubing is avoided. 2. At concentrations of 1 /~M and above, the dye seems to form aggregates of defined stoichiometry in the minor groove of polymeric DNAs, and can also bind 14S. E. S. Ebrahimi,M. C. Bibby,K. R. Fox,and K. T. Douglas,AnticancerDrugDes. 10, 463 (1995).

216

BIOPHYSICALAPPROACHES

[ 10]

and aggregate at the DNA phosphates.l° These complex binding events prevent an unambiguous determination of Ka for a Hoechst dye from saturating binding curves using DNA polymers. In contrast, with an isolated AATT site the dye forms a 1:1 complex, independent of the concentration used. j5-17 3. To determine reliably the high Ka values (Tables I-III) under nonstoichiometric conditions, low dye concentrations ( ~ 1 nM) must be used. DNA concentrations are also relatively small, and to prevent melting of a duplex at room temperature we use DNA hairpin structures as shown in Fig. 1B. K i n e t i c s of p - O H H o e c h s t B i n d i n g to P o l y m e r i c DNAs We studied the kinetics of p-OH Hoechst binding to calf thymus DNA, poly[d(A-T)], poly(dA) • poly(dT), and poly[d(A-5BrU)]. In all experiments, low concentrations of p-OH Hoechst are used (1 to 5 nM) with the DNA concentration (in sites, see legend of Table I) in 10- to 300-fold excess (pseudo-first-order conditions). These conditions offer several advantages: (1) only strong, site-specific binding is observed (no unspecific binding or stoichiometric aggregation of the dye on a polymer); (2) the binding to a polymer follows a simple independent-site binding model (statistically excluded site binding does not occur); (3) relatively low DNA concentrations can be used; and (4) the reaction is sufficiently slow to be accessible to the stopped-flow technique. All stopped-flow curves obtained at low occupancy of the polymeric DNAs are well described by a monoexponential increase in fluorescence. Plots of the observed reaction rate constants kobs v e r s u s DNA concentration are linear, as expected for a simple binding mechanism, with the association rate parameter ko, corresponding to the slope and the dissociation rate parameter koffcorresponding to the intercept. As an example, the results forp-OH Hoechst binding to poly[d(A-T)] are shown in Fig. 2. In Table I we compare the kon values for the different polymeric DNAs with their equilibrium constants Ka obtained from fluorescence titrations. 10 The intercepts of the reaction rate-versus-concentration plots are not accurate and the corresponding koff values can be estimated better from the tabulated Ka and kon values. The lowest kon value (6.1 x 107 M l sec-I) is for poly(dA)- poly(dT), possibly reflecting the decreased accessibility to the narrow minor groove.18 With poly[d(A-T)], kon (1.15 x 108 M 1 sec-1) is comparable to the value found for the well-defined smaller ( A ] T ) 4 site (see Tables II and III). The highest ko, values are for poly[d(A-5BrU)l (kon = 5.2 x 10s M -1 sec -1) and calf thymus DNA (kon = 1.4 x 10 9 M 1 s e c - 1 for an estimated probability of a high-affinity binding site 15 I. Haq, J. E. Ladbury, B. Z. Chowdhry, T. C. Jenkins, and J. B. Chaires, J. Mol. Biol. 271,244 (1997). 16 C. E. Bostock-Smith and M. S. Searle, Nucleic Acids Res. 27, 1619 (1999). 17 S. Y. Breusegem, E G. Loontiens, and R. M. Clegg, Biophys. J. 76, A128 (1999). 18 H. C. M. Nelson, J. T. Finch, F. L. Bonaventura, and A. Klug, Nature (London) 330, 221 (1987).

[ 1 O]

HOECHST DYE BINDING KINETICS

217

TABLE I ASSOCIATIONCONSTANTSANDASSOCIATIONRATEPARAMETERSFOR BINDINGOFp-OH HOECHSTTO POLYMERICDNAS AT 20° DNAa Calf thymus DNA Poly[d(A-5BrU)] Poly[d(A-T)] Poly(dA). poly(dT)

Ka (M-l) h 7× 8x 1.7 × 2.4 ×

108 109 108 108

kon (M 1 sec - I ) , (1.4 4. 0.1)× (5.2 4- 0.1) x (1.15 4. 0.02) × (6.1 i 0 . 2 ) ×

109

108 108 107

'~All DNA concentrations are expressed as sites and the extent of a binding site is defined as N = 2n- 1, with n the number of elements covered or excluded from binding at lattice saturation [J. D. McGhee and P. H. von Hippel, J. Mol. Biol. 86, 469 (1974)]; N was estimated as 80 base pairs for calf thymus DNA and as 10 base pairs for poly[d(A-T)] [K G. Loontiens, P. Regenfuss, A. Zechel, L. Dumortier, and R. M. Clegg, Biochemistry 29, 9029 (1990)], a value also assumed for poly[d(A-5BrU)] and poly(dA) - poly(dT). t' Ka values were determined by fluorescence equilibrium titrations of I nM p-OH Hoechst with excess DNA. Our unpublished data (2000), except for calf thymus DNA [EG. Loontiens, P. Regenfuss, A. Zechel, L. Dumortier, and R. M. Clegg, Biochemistry 29, 9029 (1990)]. c kon was obtained from a linear plot of kob~ versus DNA concentration, maximally (here in base pairs for comparison) 3 tiM with calf thymus DNA, 4 tiM with poly[d(A-5BrU)], 15 tiM with poly[d(A-T)] as in Fig. 2, and 5 tiM with poly(dA) - poly(dT). The errors are for the linear fit to the data as for the inset in Fig. 2.

occurring once in 80 base pairs). The high ko, values for poly[d(A-T)], p o l y [ d ( A 5BrU)], and calf thymus D N A suggest a binding association step that is controlled by diffusion, in which the dye m o l e c u l e has i m m e d i a t e access to the m i n o r g r o o v e on collision.

Kinetics of p-OH Hoechst and Four p- or m-Phenyl-Substituted OH and OCHs Hoechst Derivatives Binding to Isolated AATr Site Association Kinetics

Similar to the binding o f p - O H Hoechst to the p o l y m e r i c D N A s , binding of p - O H Hoechst and four p - and m-substituted O H and OCH3 H o e c h s t derivatives (Fig. 1A) to the A A T T site is well described by a m o n o e x p o n e n t i a l increase in fluorescence. This is illustrated in Fig. 3 for b i s - m - O H Hoechst, the best ligand in this study. Figure 4 shows the plots o f ko~s versus concentration for the five H o e c h s t molecules. The ko, values obtained f r o m the slopes in Fig. 4 are given in Table II and are similar for the five dye m o l e c u l e s (between 2.00 × 108 and

218

BIOPHYSICAL

[ 1 O]

APPROACHES

1000 800 ~D

"~

2001

600,

,

I

'7

100

o 400

0

200

I./

0 1 10 6 [poly[d(A-T)] sites] (M)

0

t

_801

,

0

,

5

i

~

i

,

, - -

10 15 Time (ms)

20

25

FIG. 2. Binding kinetics of p-OH Hoechst and poly[d(A-T)]. The stopped-flow fluorescence trace is the average of 24 reactions for binding of 5 nM p-OH Hoechst with a large excess of poly[d(A-T)] (1.44 #M in sites). The data are fitted by a single exponential with a time constant corresponding to 1/kobs = (6.13 -4- 0.08) msec. The residuals are represented in the bottom graph. The inset shows the linear plot of kobsversus DNA site concentration with a slope corresponding to kon = (1,15 ± 0.02) × 108 M -I sec -1.

TABLE II ASSOCIATIONCONSTANTSAND REACTIONRATE PARAMETERSFOR BINDINGOF HOECHST DERIVATIVESTO AATT DNA HAIRPINAT 20 ° Hoechst phenyl substitution Bis-m-OH m-OH, p-OCH3 m-OH m-OCH3, p-OH p-OH

10-8 Ka~ (M -l) 190 71 38 9.3 5.2

± 4± 4±

50 6 3 0.4 0.2

10-Sko~ (M -1 sec -I) 2.16 2.86 2.00 2.64 2.46

-4- 0.04 4- 0.09 4- 0.07 4- 0.08 ± 0.05

koffc (sec I) 0.012 0.0342 0.0510 0.270 0.419

± 0.002 4- 0.0006 -4- 0.0002 ± 0.003 ± 0.005

10-8 Kkind (M-l) 180 84 39 9.8 5.9

i 30 :~ 4 ± 2 ~ 0.4 5:0.2

~Ka values were determined by fluorescence equilibrium titrations of 0.7 to 1.6 nM p-OH Hoechst with excess DNA (our unpublished data, 2000). The errors are for the fit to a single titration curve. /, From the plots in Fig. 4. The errors are for the linear fit to the data. "Displacement kinetics as shown in Fig. 5, using poly[d(A-5BrU)] as a quenching scavenger of free Hoechst dye. The errors are for the monoexponential fit to the averaged reaction trace. d/(kin = kon/koff for a simple bimolecular association.

[10]

HOECHSTDYE BINDINGKINETICS I

I

I

I

I

I

219

600

~ 400 200

0 40 .

0

.

.

100

.

200 Time (ms)

.

300

400

FIG. 3. Association kinetics of bis-m-OH Hoechst and the AATP site. The trace is the average of 24 reactions for binding of 2 nM bis-m-OH Hoechst, the best ligand in this study, with 78.5 nM AATT hairpin. The single exponential fit corresponds to 1/kob~ = (61.8 ± 0.8) msec. The residuals for monoexponential fitting are shown in the bottom graph.

2.86 × 108 M -1 sec-I); these values are between those for poly[d(A-T)] and poly[d(A-5BrU)] in Table I. The intercepts in Fig. 4 are small, with an error that is too large to reliably estimate the corresponding koff, e.g., 1.6 4- 0.6 for p-OH Hoechst. We therefore determined these values by an independent method as described in the next section. Dissociation Kinetics

Poly[d(A-5BrU)] has interesting properties that can be exploited to determine the ko~ for Hoechst-DNA complexes. The binding of p-OH Hoechst to poly[d(A-5BrU)] is tight 19 (Ka = 8 x 109 M -1) and fast (kon = 5.2 x 108 M -1 sec- l; Table I), and the complex formed has a greatly reduced fluorescence intensity (14 times smaller2°) compared with the complex with poly[d(A-T)] or an (A/T)4 site. Therefore this halogenated synthetic polymer is a rapid and specific quenching scavenger of Hoechst dye molecules and will displace bound dye molecules from their weaker complexes at the AATT site. Mixing a preformed complex between a Hoechst dye and the AATr site with a large excess of poly[d(A-5BrU)], either in the stopped-flow apparatus (for the rapidly dissociating complex of p-OH Hoechst 19 S. Y. Breusegem, E G. Loontiens, and R. M. Clegg, Biophys. J. 78, 305A (2000). 20 T. Hard, P. Fan, and D. R. Kearns, Photochem. Photobiol. 51, 77 (1990).

220

BIOPHYSICALAPPROACHES

[ 10]

40 ~. 30

m-OCH3, p-OH _

~ 20 r~

10 0

i

t

50

,

,

40

- + - m-OH' p-OCH3] " bis-m-OH I

30

~>- m-OH

I

A

-JJ o

~-

~ 20 10 0 0

5 10-8 1 10-7 [AATT DNA hairpin] (M)

1.5 10-7

FIG, 4. Determination of kon for the binding of the Hoechst derivatives to the AATT hairpin. The kob~ values correspond to the time constants for monoexponential fitting of the stopped-flow traces using the AATT hairpin in excess over 2 nM dye (as in Fig. 3). (A) p-OH substituted Hoechst derivatives; (B) m-OH substituted derivatives. The kon values are obtained from the slope of the linear fits and are collected in Table I1.

and AATT) or by manual mixing (for the more tightly binding derivatives), allows determination of the kofffor the highly fluorescent complex. It was verified that the fastest koffvalue obtained is independent of the concentration of poly[d(A-5BrU)] when the latter is used in 25- to 75-fold excess. Figure 5 shows kinetic traces obtained for the dissociation of the five Hoechst dye-AATT complexes with poly[d(A-5BrU)]. The corresponding koffvalues are given in Table II. The least stable dye complex, p-OH Hoechst-AATT, determined at a concentration of 2.7 nM, has the shortest lifetime, corresponding to the largest koff value (0.419 sec-1). The koff is slowest for bis-m-OH Hoechst (0.012 sec-I), the most strongly binding ligand. Determining the dissociation kinetics of Hoechst dyes using poly[d(A-5BrU)] is simpler than the two following alternative approaches tested with p-OH Hoechst bound at the A A T T site:

[ 10]

HOECHSTDYE BINDINGKINETICS 1500

A

1250

p-OH,

o

--

221

0

10 Time (s)

5

1800

'

15 '

20 B

1500

~ 1200

90(3

L 1()0

2;0

300

Time (s) FIG. 5. Determination of koft- for the AATT hairpin-Hoechst dye derivative complexes, using poly[d(A-5BrU)]. The preformed and strongly fluorescent complex (mixture of equal analytical concentrations, resulting in a final concentration of 2.7 to 4 nM complex) was rapidly mixed with an excess of the specific fluorescence quencher poly[d(A-5BrU)] at a final concentration between 100 and 300 nM sites. (A) Fastest two dissociation curves for p-OH Hoechst and for p-OH, m-OCH3 Hoechst. For p-OH Hoechst the stopped-flow technique was used and 17 traces were averaged. For the p-OH, m-OCH3 derivative, the preformed complex in a 1 x 1 x 4.5 cm cuvette was manually mixed with a concentrated solution of poly[d(A-5BrU)] and the trace is the average of four experiments. (B) Three m-OH derivatives and manual mixing with poly[d(A-5BrU)]. For the m-OH derivative the average of four traces is represented, the slow dissociation of the m-OH, p-OCH3, and bis-m-OH derivatives is represented as a single trace. The smooth curves through the data are monoexponential fits with inverse time constants corresponding to the/;off values given in Table II.

1. Competitive Hoechst dissociation kinetics using netropsin depended on the concentration of this putative competitor. 2. Disrupting the DNA structure with 1% sodium dodecyl sulfate (SDS) resulted in biexponential kinetics with time constants that differ by a factor of about 10 and with comparable amplitudes, suggesting an additional slower process that depends on the SDS.

222

BIOPHYSICALA P P R O A C H E S

[ 1 O]

Comparison of Kinetics with Equilibrium Data Our kinetic results show that for binding to the AATT site koffdecreases in the series p-OH > p-OH, m-OCH3 > m-OH > m-OH, p-OCH3 > bis-m-OH whereas kon is nearly constant for the five dyes. The values of kon and koefdetermine a kinetically defined association constant Kki n = kon/koffthat, within experimental error, is identical to the value of Ka determined at equilibrium by fluorescence titrations (Table II). The internal consistency of the Ka, ko,, and koffvalues, obtained by three independent experiments (equilibrium titrations and stopped-flow association and dissociation reactions), for the five dyes, is strong experimental evidence for a simple binding mechanism, in which one type of complex is formed in a diffusioncontrolled association step and the affinity is determined by the dissociation rate. However, in spite of this good agreement, the kinetics are more complex than a single binding step, as we show below.

Effect of m-OH and Bis-m-OH Phenyl Substitution in Hoechst Dye The increase in K a for the AATT site in the DNA hairpin in the series p-OH < p-OH, m-OCH3 < m-OH < m-OH, p-OCH3 < bis-m-OH is the same as we have found for poly[d(A-T)] at equilibrium (our unpublished data, 2000); however, the Ka is greater for binding of a given dye to an isolated site than for poly[d(A-T)]. This observation with the DNA hairpin is consistent with the possibility of additional hydrogen bonding by the m-OH group with 2-0 of C-25 and 2-NH2 of G-4 in the minor groove.12 However, the observation with poly[d(A-T)] indicates that other factors are responsible for the increased affinity of an m-OH derivative compared with a p-OH derivative, because the hydrogen bond with the 2-NH2 group is not possible in poly[d(A-T)]. When going from p-OH to m-OH Hoechst, there is an increase in binding energy of 1.1 kcal for the AATT site and of 0.5 kcal for the poly[d(A-T)] site. Rather than direct contact with the base substituents in the neighboring G-C pair, this seems to involve hydrogen bonding with two water molecules that bridge base substituents and a ribosidic ring oxygen on binding an m-OH Hoechst derivative. 21 When going from m-OH to bis-m-OH Hoechst there is an additional and equal gain of 1 kcal on binding both to the AATT hairpin and to the poly[d(A-T)] site. One of the m-OH groups in bis-m-OH points out of the groove and can, in the duplex, make hydrogen bonds with three water molecules that connect base substituents, phosphate oxygen and ribosidic oxygen [see the crystal structures NDB X96024 and X96025 in the Nucleic Acid Database (NDB)]. Therefore, the decrease in koff, and consequently the increase in Ka for m-OH and particularly for bis-m-OH, might result from a stronger integration of the polar ends of the molecules within the structured water network in the DNA groove. 2~G. R. Clark,C. J. Squire, E. J. Gray,W. Leupin,and S. Neidle,NucleicAcidsRes.24, 4882 (1996).

[ 1 0]

HOECHST DYE BINDING KINETICS 120

,

,

223

,

100 ~

o= 8 0

From top to bottom:

60

o 40 20 fl v0

///

TATA

~

ATAT AATT TAAT

_

I

[

I

0.05

0.l Time(s)

0.15

0.2

FIG.6. Comparison of the association kinetics of p-OH Hoechst and five (A/T)4 sites in DNA hairpin structures, from top to bottom: TTAA,TATA,ATAT,AATT,and TAAT.The data were obtained on mixing of 2 or 10 nM p-OH Hoechst with about 100 nM DNA (sites), the higher dye concentration being required for sufficientcomplex formation with TTAA and TATA.From top to bottom the time constant 1/kobsof the corresponding monoexponential fit increases as indicated and the number of traces averaged is given in parentheses: TTAA, 8.4 msec, (21); TATA, 12.9 msec (13), ATAT,32.3 msec (23); AATT,38.4 msec (14); TAAT,43.1 msec (21). This process brings the phosphoribose chains closer around the dye molecule, increases the van der Waals contacts, and increases the hydrophobic interactions, which have surprisingly been suggested as the dominant contribution to the binding energy. 15 The Ka value for bis-m-OH Hoechst binding to AATT (1.90 × 101° M - 1) corresponds to a binding energy of 14,0 kcal m o l - l ; this is only 1.7 kcal m o l - I less than for binding of the restriction enzyme EcoRI to its GAATTC sitefl 2 This observation is noteworthy, and emphasizes that to realize strong binding to DNA, or to achieve stringent specificity, it is not necessary to have a large molecule such as a protein molecule with large interaction surface areas and many points of interaction.

Kinetics of p-OH Hoechst Binding to Five Different (A/T)4 S e q u e n c e s Pseudo-First-Order Kinetics In an additional effort to better understand the specific binding of p - O H Hoechst to defined (A/T) sequences, we studied its binding to different (A/T)4 sequences: AATT, TAAT, ATAT, TATA, and TTAA. Comparison of the stopped-flow traces for the five sequences at about equal concentrations (Fig. 6) with the equilibrium 22 D. R. Lesser, M. R. Kurpiewski,and L. Jen-Jacobson,Science 250, 776 (1990).

224

BIOPHYSICAL APPROACHES

[ 10]

TABLE III BINDING PARAMETERS FOR p-OH HOECHST AND DIFFERENT (A]T)4 SITES AT 20 ° 5'--> 3'

10 g Kab

Sequence ~

(M-I)

AATT TAAT ATAT TATA TTAA

10 -7 (1/C50) "

10 8kond

koff d

(M-1 sec-I)

(sec t)

5.2 ± 0.2 2.78 ± 0.85 2.47 ± 0.03 (1) (1) 0.271 ± 0.003 0.051 ± 0.(121 1.57 ± 0.03 (0.0518) (0.0183) 0.240 ± 0.002 0.022 ± 0,014 1.26 ± 0.002 (0.0459) (0.0079) 0.026 ± 0.001 0.008 ± (I.004 0.68 ± 0.01 (O.0O5O) (0.0028) 0.030 ± 0.001 0.007 ± 0.005 1.93 ± 0.07 (0.0058) (0.0024)

10 g Kkine

(M-~)

0.419 ± 0.005

5.9 ± 0.1

7.0 ± 0.4

0.22 ± 0.02

10.5 ± 0.3

0.12 ± 0.04

66 ± 3

0.010 ± 0.001

96 ± 4

0.020 ± 0.003

a The 5' --->3' sequences are constructed in DNA hairpin structures as in Fig, lB. h Ka values were determined by fluorescence equilibrium titrations of 1.6 nM p-OH Hoechst with excess DNA (our unpublished data, 2000). The errors are for the fit to the single titration curve. The relative values of Ka (relative to the AATT sequence) are given in parentheses, c Data from Abu-Daya etal. [footprinting experiments; A. Abu-Daya, P. M. Brown, and K. R. Fox, Nucleic Acids Res. 23, 3385 ( 1995)1. d The kon and koff values are from the plots in Fig. 7, except for the koff for AATT (taken from Fig. 5). The errors are for the linear fit to the data. e Kki,1 = kon/koff for a simple bimolecular association.

constants s u m m a r i z e d in Table III shows that the kinetic traces b e c o m e slower as the binding to the different sequences b e c o m e s stronger. This difference in the reaction rate is almost exclusively due to the dissociation rate parameter koff. Again, the stopped-flow reactions were fitted as m o n o e x p o n e n t i a l s and the plots o f kobs as a function of concentration are linear for the five sequences (Fig. 7). B e c a u s e the koff for p - O H H o e c h s t and the TAAT, ATAT, TATA, and T T A A sequences are larger than for AATT, they could be determined f r o m the intercept in the concentration plots. The koff ['or the A A T T site was obtained by displacement kinetics with p o l y [ d ( A - 5 B r U ) ] as described above. The ko, values are similar (0.68 x 108 to 2.47 x 108 M 1 s e c - I ) , and are also similar to the kon values for the derivatives in Table II. In contrast, the koft" values for the sequences differ by a factor of almost 200 and define the affinity of p - O H Hoechst for the different sequences. Thus, as o p p o s e d to binding of p - O H H o e c h s t to p o l y m e r i c D N A s (Table I), binding of a H o e c h s t dye to the A A T T site (Table II), as w e l l as to the four additional (A/T)4 sites (Table III), is characterized by a ko, value that is independent of the dye derivative or the (A/T)4 sequence. The affinity is determined exclusively by the koff or the lifetime of the c o m p l e x that is formed. This important conclusion m i g h t apply generally to other peripheral binding processes, including regulatory

[ 1O]

HOECHSTDYEBINDINGKINETICS

225

40 ,-, 30

A

20 ,_~o

A

~'~ 10 / D ~ ~1~"

~ J

300

• AATT TAAT ' ATAT I

1 10 -7 2 10 -7 [(A/T)4 site] (M) ' /~

B

zoo

100 I

0

5 10.7 [(A/T)4 site] (M)

I

1 10.6

FIG. 7. Determination of konand koffforp-OH Hoechst binding to five (A/T)4 sites in a DNA hairpin. The kobsvalues correspond to the time constants for monoexponential fitting of the stopped-flow traces using the DNA hairpins in excess over 2 or 10 nMp-OH Hoechst (as in Fig. 6). (A) Strongest binding sequences AATT, TAAT, and ATAT;(B) weaker binding sequences TATA and TTAA.

proteins and enzymes. The specificitycomes from the dissociation rate parameter and the shape of the ligand. Comparison o f Kinetics with Equilibrium Data As for the derivatives, the kinetic association constant Kki, = kon/koff is identical or close to the corresponding K~ value obtained by equilibrium fluorescence titrations.19 T h e s e stopped-flow and equilibrium data confirm and refine the footprinting data o f A b u - D a y a et al. 23

23 A. Abu-Daya, E M. Brown, and K. R. Fox, NucleicAcids Res. 23, 3385 (1995).

226

BIOPHYSICALAPPROACHES

[ 1 0]

Effect o f Base-Pair Roll and Minor-Groove Width on p - O H Hoechst Affinity

The highest affinity of p-OH Hoechst for the AATT sequence seems to correlate with the zero base pair roll and reduced minor groove width (as small as 3.1 ]k) at the central AATT sequence in the d(CGCGAATTCGCG)2 duplex, probably due to the binding of monovalent cations in the bottom of the groove.24 This contrasts with TATA as the poorest affinity site: in solution, the [d(CGCTATACGCG)]2 sequence shows an increased roll (17.2 °) at the two TpA steps that widens the minor groove to an overall width of 7.2 A. 25 Likewise, the single and central TpA step in CGATTAATCG has a local roll of 9.4 ~' and corresponds to a minor groove width of about 8 ]~26 Although narrowing of the minor groove at the AATT sequence correlates with the affinity, it is puzzling that sequencedependent tightening of the minor groove in the AATT sequence does not reduce its value of kon that is the largest of the five sequences investigated (see the next section).

More C o m p l e x Kinetics: T w o - S t e p B i n d i n g M o d e l s We have seen above that the progress curves of p-OH Hoechst binding to polymeric DNAs and to defined oligonucleotides can be represented well by a single exponential. Analysis and interpretation of the concentration-dependent rate data are then straightforward, corresponding to a single-step bimolecular reaction mechanism, with the binding step being almost diffusion controlled and the dissociation rate determining the affinity of the dye for the DNA target. In addition, the reaction rate parameters obtained from this simple kinetic analysis agree well with the equilibrium titration data. However, it is reasonable to wonder how a molecule that fits so snugly into the minor groove can bind with a kinetic association constant that is close to diffusion controlled, especially considering the extended nature of the ligand. In addition, we know that there are specific attachment sites between the dye and the DNA that require a precise positioning of the dye in the groove. For every collision with the DNA the dye cannot be oriented correctly for insertion into the helix groove, and we might suspect that the geometry and size of the minor groove would slow down the association step. Indeed, this is the case, as we show, and it is likely that this conclusion applies to other dye molecules that fit precisely into relatively deep sites in the DNA molecule.

24X. Shui, C. C. Sines, L. McFail-Isom,D. VanDerveer,and L. D. Williams, Biochemistry 37, 16877 (1998). 25J.-W. Cheng,S.-H. Chou,M. Salazar, and B. R. Reid, J. Mol. Biol. 228, 118 (1992). 26j. R. Quintana, K. Grzeskowiak,K. Yanagi,and R. E. Dickerson,J. Mol. Biol. 225, 379 (1992).

[10]

HOECHST DYE BINDING KINETICS I

I

227

i

100 8O

~= 60

g 40 !

2o

A 1

I

I

+

I

o

'

I

I

I

d

,,,

eO

~a

2F

0

+ It~

0.1

] '

i

0.2 T i m e (s)

I

0.3

i

i

0.4

FIG. 8. Association kinetics of p-OH Hoechst and the AATT hairpin with improved signal-tonoise ratio. (A) Average of 20 traces (2000 points per trace) for binding of 8 nM p-OH Hoechst to 52 nM AATT hairpin and its single-exponential fit (corresponding to r = 98.6 msec). (B), (C), and (D) compare the residuals for different fits: (B) monoexponential (R = 0.99869); (C) biexponential (rfast = 45.7 msec and rslow = 113.3 msec, R = 0.99952); (D) triexponential (r~ = 11.3 msec, t2 67.7 msec, and r3 = 2.0 msec, R = 0.99958). =

ExperimentalApproach To detect any deviations from single-exponential behavior we have increased the signal-to-noise levels of the kinetic data acquisition by using higher dye and D N A concentrations. We have done this deliberately with the well-defined D N A oligomers so as not to confuse any deviations with polymeric effects. We have also m a i n t a i n e d the pseudo-first-order conditions with the D N A hairpin in excess over the dye. Figure 8A shows an improved kinetic trace for p - O H Hoechst b i n d i n g to the A A T T hairpin and its single-exponential fit; the residuals of the fit are shown in Fig. 8B. The systematic deviation in the residuals seen in Fig. 8B is found with most

228

BIOPHYSICAL APPROACHES 0

i

[ 101

i

7°' 1 obs,i

60

[]

obs,ii [

•

,-, 50 40 "~° 30 20

plateau value

10

~

~ I

0

~

~

0 4

I

5 10 -8 1 10 -7 1.5 10 7 [AATT D N A hairpin] (M)

2 10 -7

FIG. 9. Concentration plot for binding of p-OH Hoechst to the AATT hairpin, For each DNA concentration about 20 stopped-flow traces (500 points per trace) were averaged and fitted by a double exponential. The linear fit through the inverse fast time constants has a slope of 2.7 × 108 M - I sec -1 and an intercept equal to 13 sec 1.

of the DNA concentrations and with all the DNA hairpin sites in this study. The deviation is small, as we would expect because the previous one-exponential curves already represented the data well. However, this systematic deviation indicates that a single exponential is not an adequate representation of the progress curves.

Biexponential Association Kinetics Fitting the data with two exponentials results in a significant improvement in the residuals for most of the concentrations the residuals now approach a random deviation around zero (Fig. 8C). The fitting parameters obtained show relatively large errors, as we expected considering the previous satisfactory one-exponential fits, and the plots of the inverse time constants ( 1/rfast = kobs,i and 1/ rslow = kobs,ii) versus the DNA concentration, for p-OH Hoechst binding to AATT, show relatively large scatter (Fig. 9). However, the concentration plot in Fig. 9 suggests that the kobs, i values increase approximately linearly with DNA concentration while the kobs,ii values increase slowly and eventually reach a plateau value.

Model 1: Bimolecular Association followed by lsomerization Because we observe two exponential processes there are at least two welldefined kinetic steps for the reaction. Several two-step reactions 27'28 are consistent with the behavior seen in Fig. 9. We can, for example, consider a classic binding

[ 10]

HOECHST DYE BINDING KINETICS

229

model consisting of a bimolecular association reaction followed by a conformational change of the complex: k+l

k+2

DNA+dye ~C~-D k I

k-2

(i)

with Ki = k+l/k-l and K2 ----k+2/k 2. Under first-order conditions, with the DNA sites in large excess over the dye, and assuming that the bimolecular reaction is rapid compared with the rate of the second reaction, kobs.i and ko6s.ii for the above-described binding mechanism are described by 28 kobs,i = k I + k+I(DNA0)

kobs,ii =

k+zKl (DNA0) KI(DNA0) + 1 + k-2

(2) (3)

with (DNA0) the total DNA concentration. A linear fit of the kobs,i in Fig. 9 gives a slope k+l -- 2.7 × 108M -I sec -I, close to the association rate parameter obtained from the single-exponential analysis (2.47 × 108M-Isec-1). The intercept k_l = 13 sec -I is not so well defined and is about 10 times larger than the value from the single-exponential analysis. These kinetic parameters define Ki ----k+l/k_j = 2.1 × 107M -l for the first bimolecular reaction step, significantly smaller than Kkin obtained in the single-exponential analysis or Ka obtained from the equilibrium titrations (see Table III, Ka = 25K1). From the plot of kobs,ii v e r s u s the total DNA concentration (Fig. 9) we can deduce that we are already in the concentration region where kobsd i increases only slowly and has almost reached its plateau value of k+2 + k 2 estimated as 15 sec- J. Relatively high DNA concentrations were used, much higher than the dye concentration (8 nM), in order to be under pseudo-first-order conditions. These concentrations are too high for estimating the reverse rate parameter k_2 from the intercept. However, we can calculate the effective equilibrium constant of this reaction model to be Ka --- KI(1 + K2), assuming that we are observing both bound states as one state in the equilibrium experiments. Because Ka ----25KI (see above), we expect that K2 = k + z / k - 2 = 24. Because we estimated that k+2 q-- k - 2 = 15 s e c -1 w e can solve for k+2 and k-2. The result is k+2 = 14.4sec -I and k 2 = 0.6sec -1. The latter value of k_2 compares well with the overall dissociation rate of p-OH Hoechst from the AATT site measured with poly[d(A-5BrU)] (0.419 sec -1) and is also close to the intercept of the linear concentration plot for the single-exponential fit (1.6 sec-1). The analysis of our kinetic data according to the above-described two-step mechanism [Eq. (1)] contributes considerably to our understanding of the 27 C. F. Bernasconi, "Relaxation Kinetics," Chapter 3, Table 3.1. Academic Press, New York, 1976. 28 A. Van Landschoot, E G. Loontiens, R. M. Clegg, and T. P. Jovin, Era: J. Biochem. 103, 313 (1980).

230

BIOPHYSICALAPPROACHES

[ 10]

mechanism of the reaction. Specifically, it shows why we can measure an association rate that is close to diffusion controlled, assuming a single-step binding mechanism. The p-OH Hoechst molecule collides with the DNA molecule (in a diffusion-controlled step), and there is at least one additional step following this collision in which the dye molecule snuggles into the minor groove and makes its specific contacts with the DNA. The dye molecule cannot dissociate from the DNA until this second monomolecular step is reversed, requiring the dye to find its way out of the enclosed and restrictive environment of the deep, narrow minor groove. This is a slow process that involves the specific interactions between the dye and molecular groups in the DNA minor groove, and, as we discussed above, its rate determines the affinity of the dye for the particular DNA site. The process may also involve dynamics of the DNA molecule itself, an intriguing possibility that must await further investigations. Note that the kinetic constants k-i = 13 sec -I and k+2 = 14.4sec I are approximately equal, so that when we calculate the ratio of k+l and the dissociation rate measured competitively with poly[d(A-5BrU)]) we obtain a value that agrees well with the equilibrium association constant found by accurate fluorescence titration experiments. This means that the dye molecule in the initial, "outside" bound state has about equal probability to dissociate completely from the DNA molecule into the solution or to enter the enclosed state of the minor groove. Model 2: Two Single-Step Associations

The binding-plus-isomerization model described above is in agreement with high-resolution (1.5-A) X-ray diffraction data that suggest that a Hoechst dye, bound in a particular orientation, can "slide" over the AATT binding site over a distance of 3 ~.29 However, in addition, the X-ray data suggest that the dye can bind in two opposite orientations that partially overlap and mutually exclude each other. This requires the dye to dissociate from the site before reentering, a process that is not included in the above-described two-step binding-plus-isomerization mechanism with only one d y e - D N A complex species. Therefore we consider an alternative model that is kinetically equivalent but mechanistically different, consisting of two single-step associations forming two complexes that mutually exclude each other: k+~

DNA + dye ~- C k ~

k+b

and

DNA + dye ~ D

(4)

k-t,

with KA = k+a/k-a and KB = k+b/k b. Both alternative mechanisms are difficult to distinguish. 3° For the two competing single-step associations the inverse fast and slow time constants (1/rf~t = kobs,i 29C. J. Squire, L. J. Baker,G. R. Clark, R. E Martin, and J. White,Nucleic Acids Res. 28, 1252(2000). 30R. D. Viale, J. Theor. Biol. 31,501 (1971).

[ 1 O]

HOECHSTDYE BINDINGKINETICS

231

and 1/Vslow = kobs.ii) are described by kobs.i = k_a + k+a(DNA0)

kobs.ii

=

k+b(DNA0) KA(DNA0) + 1

Jr- k_ b

(5) (6)

The faster reaction has the same properties as in the first mechanism [Eq. (1)], giving KA = 2.1 x 107M -l . Assuming that both C and D contribute in our equilibrium fluorescence titrations, the overall association constant Ka (5.2 x 108 M - l ) equals KA + KB, which characterizes the slower reaction as forming the tightest complex with KB = 5.0 x 10SM -l. The plateau value in the kobs,ii plot (Fig. 9) is n o w k - b q- (k+b/Ka) = 15 sec 1. Neither the slope k+b n o r the intercept k_ b can be determined from our plot of kobs,ii v e r s u s the DNA concentration (Fig. 9); however, for the complexes C and D, k - a and k - b are expected to be different. We therefore also reexamined the dissociation experiments with poly[d(A5BrU)] under conditions of improved signal-to-noise ratio, and data for the p-OH Hoechst-AATT complex are shown in Fig. 10. The dissociation curves are fitted significantly better with two (Fig. 10C) rather than with one exponential (Fig. 10B). However, the fitted rates are close to each other and differ by a factor of 2.6 only, possibly reflecting a small difference between the values of k-a and k-b. This is an indication of multiple bound states of the dye molecule in the DNA minor groove, each state having its distinctive overall dissociation rate.

Triexponential Association Kinetics As a further note, we mention that in some cases a two-exponential fit of the association kinetic traces still shows systematic deviations that are reduced by fitting with three exponentials (Fig. 8D). The deviation from biexponential behavior is small and we cannot reliably deconvolute another process from the data. Perhaps by varying the experimental conditions (different temperatures, pressures, ionic strength) we will be able to decide whether this is a real effect. At this level of accuracy care is needed not to be confused by small artifacts, even if they are systematic. The instrumentation is stable, but there could be small nonlinearities that are difficult to expose unless one can reproduce the exact conditions of the progress curve. Conclusions In this chapter we have given the reader a working example of a kinetic analysis involving a ligand that binds strongly to specific sequences of DNA. The Hoechst dye-DNA interaction is an informative model system for the study of the highly A/T-specific minor groove binding to DNA. Titrations at equilibrium allow us to determine the high association constants. However, the determination of binding

232

BIOPHYSICALAPPROACHES

[ 10]

A 80 ca

60 o

40

20

~

I

,a 2LI/

'

'

=21

.

.

.

.

B

~

Time (s) FIG. 10. Dissociation kinetics of the p-OH Hoechst-AATT complex determined by displacement with poly[d(A-5BrU)]. (A) Average of 15 kinetic traces (500 points per trace) for a mixture of 10 nM Hoechst 33258 and 10 nM AATT hairpin, forming 6.5 nM complex, rapidly mixed with 210 nM poly[d(A-5BrU)] using stopped flow. For the sake of clarity no simulation is superposed on the data. (B) Residuals for a monoexponential fit (1/kobs = 2.1 sec). (C) Residuals for a biexponential fit (rl = 1.5 sec, r2 = 3.9 sec).

kinetics considerably augments our understanding of the factors responsible for the high affinity. Unless the data are accurate the stopped-flow curves are well described by single exponentials and the kinetic analysis suggests a simple, single-step binding mechanism. The association rate is almost diffusion controlled and the specificity of the dye comes solely from the reverse rate parameter, possibly determined by the number of dye-DNA interactions in the minor groove that must be broken for the dye molecule to escape. When more interactions become involved, as with the strongest ligand in this study (bis-m-OH), the time for the ligand to dissociate becomes longer, and its affinity increases. Similar principles might well apply to other DNA-binding drugs as well as to the docking of enzymes and regulatory proteins on to DNA.

[ 10]

HOECHST DYE BINDING KINETICS

233

We have discussed details of a two-exponential analysis in order to demonstrate the complexities that can be expected when dye molecules interact strongly with specific DNA sequences. Such expanded kinetic models [Eqs. (1) and (4)] became necessary after increasing the signal-to-noise levels of the kinetic experiments, because the one-exponential analysis was no longer adequate to fit the data. The first of the two-step models [Eq. (1)] has allowed us to understand how a molecule that fits so snugly in the minor groove can still behave as though the association step is approximately diffusion controlled. In addition, we have shown that the affinities of p-OH Hoechst for different sequences are reflected in the dissociation rates--not in the association steps. The two-step mechanisms we have discussed are commonly applied to simple chemical reactions, as well as to complex biomacromolecular reactions. Multistep mechanisms for macromolecular reactions reveal the concept of conformational changes initiated by binding events that are so prevalent in biology. In this case, the step(s) following the initial binding event is probably related to the nestling of the loosely bound dye molecule into the enclosed, confined binding pocket of the DNA minor groove. This reaction step probably also requires small dynamic conformational adjustments of the DNA molecule. The second of the two-step models [Eq. (4)] is kinetically equivalent to the first, but requires that the dye completely dissociate from the site to occupy it again, possibly in a different orientation. Importantly, high-resolution X-ray diffraction data 29 corroborate our two models. It is necessary to remember that there are certainly other mechanisms that can account for the data. Kinetic experiments can never prove the exclusive validity of one particular model, and binding of p-OH Hoechst could be more complex than a two-step mechanism. However, combining equilibrium experiments with a variety of kinetic experiments can provide the rationale for understanding and quantifying fundamental steps of the binding mechanism, and sheds light on the general physical principles that produce highly selective and extremely strong interactions involving complex biological macromolecules, such as these dyeDNA interactions. Acknowledgments This work was initiated with the support of (N)FWO and (B)OZF (to EG.L.). We thank Kenneth T. Douglas and Seyed E. Ebrahimi for their gift of the Hoechst derivatives and Etienne Wouters and Eddy Van Outryve for their help in constructing and optimizing the equipment.

234

BIOPHYSICALAPPROACHES

[1 11

[ 1 1 ] Scanning Force Microscopy of Nucleic Acid Complexes By PETER T. LILLEHEIand LAWRENCE A.

BOTTOMLEY

Introduction Small ligands bind to DNA by (1) intercalation between adjacent base pairs, (2) binding within the major or minor grooves, (3) nonclassic modes] or (4) a combination of these modes. Definitive assays for mode of binding are threedimensional (3D) structure determination by X-ray diffraction or nuclear magnetic resonance (NMR) spectroscopy. These labor-intensive techniques are applicable only to binding studies of short DNA fragments because of difficulties in obtaining diffraction-quality crystals or interpretation of complicated chemical shift data. Their use is often precluded by lack of site specificity, rapid exchange, or multiple binding modes. Assays suitable for long DNA fragments involve viscometry, sedimentation, and linear and circular dichroism. These methods are reliable when ligands bind by conventional intercalative or minor groove modes, 2 but can be confounded by mixed and nonclassic modes. All traditional assays require large quantities of material. Whereas traditional techniques average over population and time, microscopic imaging techniques enable structural determinations of complexes from single molecules that are effectively frozen in time and space. Evidence of the mode and site of drug binding is, in principle, directly attainable from image analysis of a small number of nucleic acid molecules. Optical microscopy has insufficient spatial resolution for visualization of most drug-DNA complexes. The elaborate sample preparation techniques required for electron microscopy have precluded its use as an assay for drug binding. Scanning force microscopy (SFM, also known as atomic force microscopy) enables direct visualization of single DNA strands. A tip, fabricated onto the end of a readily deformable, rectangular or triangular-shaped cantilever, is positioned near or on the surface of the sample to be imaged. The tip and sample are moved with respect to one another and the cantilever bends in response to surface topography. Laser light is reflected off the cantilever onto a position-sensitive photodetector to track cantilever deflection. An image is produced by plotting cantilever deflection as a function of the position of the tip on the sample. SFM has advantages over optical and electron microscopy for imaging nucleic acids. Images can be acquired in vacuum, under liquid, or in air at ambient, 1 L. A. Lipscomb, K X. Zhou, S. R. Presnell, R. J. Woo, M. E. Peek, R. R. Plaskon, and L. D. Williams, Biothemistry 35, 2818 (1996). 2 D. Suh and J. B. Chaires, Bioorg. MeN. Chem. 3, 723 (1995).

METHODSIN ENZYMOLOGY,VOL.340

Copyright@ 2001 by AcademicPress All rightsof reproduclion in any lbrm reserved. 0076-6879/00 $35.00

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235

cryogenic, or elevated temperatures. Sample preparation is minimal. Addition of a stain or contrast agent, replica formation, or application of a conductive coating is not required. As a result, SFM is inherently a higher resolution technique. SFM has limitations. Images of nucleic acids can be obtained only when the molecule is physisorbed or chemisorbed to an atomically flat substrate. Image features are dependent on tip geometry and structural details are limited only to the portions of the molecule in direct interaction with the tip. Analysis and interpretation of images of nucleic acids require (1) assessment of tip shape and diameter as well as how these parameters may have changed during the course of imaging, (2) consideration of elastic and inelastic sample deformation, and (3) knowledge of the sources and treatment of image artifacts. Even with these limitations, the capability of acquiring subnanometer scale images of single molecules in vitro renders SFM superior to other forms of microscopy for determining nucleic acid structures.

Instrumental Considerations Imaging Modes Three modes are used to image nucleic acids: contact, noncontact, and intermittent contact imaging. Schematics of each are given in Fig. 1. Each has advantages and limitations. Contact Mode. As the name implies, with the contact imaging mode the tip is kept in contact with the sample. Contact mode images are obtained by plotting cantilever deflection as a function of the xy coordinates of the tip on the sample. Distortion of soft surfaces (or compliant molecules adsorbed on them) is commonplace with this imaging mode because of variation in the vertical force exerted by the tip on the sample. To image under fixed vertical load, a feedback mechanism is required. As the cantilever bends in response to surface topography, the scanner is retracted or extended to return cantilever deflection to its original value. Images are obtained by plotting the z piezovoltage as a function of the xy coordinates of the tip on the sample. Because the movements of the scanner are calibrated to subangstrom dimensions, accurate topographic images are obtained with properly tuned scanner feedback control. High-quality images of DNA can be acquired in just a few minutes with this imaging mode. Minimal vertical loading must be used to eliminate sample deformation or damage and tip wear. In the absence of strong adsorbate-substrate interaction, molecules will be readily displaced with this imaging method. When samples are imaged in air under ambient humidity, a thin film of water forms on the sample and tip surface. Contact between these water layers produces increased vertical and lateral forces that cause severe artifacts in the image. 3 Whenever possible, contact mode imaging should either be performed under a dry atmosphere or under fluid.

236

BIOPHYSICALAPPROACHES

a

I

11 11

%

FIG. 1. Schematic of the tip-surface interaction during (a) contact mode, (b) noncontact mode, and, (c) intermittent contact mode imaging.

Noncontact Mode. The noncontact imaging mode maintains the sample and probe tip in close proximity while scanning. The cantilever oscillates at its resonant frequency and any change in the forces acting on the cantilever (magnetic, electrostatic, and/or van der Waals forces) will be manifested as a shift in resonant frequency. Topographical information is obtained by monitoring resonant frequency shifts as a function of the xy coordinates of the tip above the sample. Tip wear and sample deformation or damage are eliminated in noncontact mode imaging. Because the forces are often small, tip-sample separation must be minimized. This mode is best suited for imaging under vacuum, in which van der Waals interactions are not dampened by intervening liquid or by adsorbed gases on the surface of the tip and sample. 3 T. Thundat, R. J. Warmack, D. E Allison, L. A. Bottomley, A. Lourenco, and T. L. Ferrell, J. Vac. Sci. Technol. A. 10, 630 (1992).

[1 1]

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Intermittent Contact Mode. Presently, most biological samples are imaged us-

ing intermittent contact. In this mode, the cantilever is oscillated at its natural resonance or driven at some selected frequency. Separation of the sample and tip is reduced until the oscillation of the cantilever is dampened. Images are acquired by tracking the shift in either amplitude or phase of oscillation as a function of the xy coordinates of the tip above the sample. Maps of sample topography are produced by recording shifts in amplitude whereas maps of sample compliance are produced by recording shifts in phase. Resolution is enhanced with intermittent contact imaging compared with contact imaging because the reduced tip wear allows for imaging with sharper and more delicate probe tips. Sample and tip damage is reduced because the tip is not shearing the surface; rather, its motion is always normal to the sample. Sticking of the tip to the surface is minimized because tip momentum is sufficient to overcome surface adhesion forces. The method by which the cantilever is oscillated varies among microscope manufacturers. Digital Instruments (Santa Barbara, CA) uses TappingMode whereas Molecular Imaging (Phoenix, AZ) uses MAG mode. In TappingMode, a piezoelectric oscillator is placed in contact with the cantilever holder and the entire cantilever assembly is oscillated at a desired frequency.4 The MAG mode oscillates a cantilever, coated with a magnetic material, with an induction coil placed beneath the sample. The oscillating magnetic field emanating from the coil drives the cantilever.5 Cantilever oscillation amplitudes are reduced compared with TappingMode. The relative advantages and limitations of each method are the subject of an ongoing debate in which we chose not to engage. In the context of this chapter, we shall treat both modes as equivalent. Tip and Scanner Calibration Cantilevers, tips, and the scanner are critical components of the microscope. Commercially available rectangular and V-shaped cantilevers are appropriate for nucleic acid imaging. The latter provides lower mechanical resistance to vertical deflection and high resistance to lateral torsion. 6 The size, shape, and material composition of an SFM tip have important consequences on the imaging of nucleic acids. Because the resultant scanning probe microscopy (SPM) image is always some convolution between the tip shape and image topography, tip artifacts will always be intrinsic to the technique.

4 D. A. Waiters, J. P. Cleveland, N. H. Thomson, P. K. Hansma, M. A. Wendman, G. Gurley, and V. Elings, Rev. Sci. lnstru. 67, 3583 (1996). 5 W. H. Han, S. M. Lindsay, and T. W. Jing, Appl. Phys. Lett. 69, 4111 (1996). 6 R. Howland and L. Benatar, in "A Practical Guide to Scanning Probe Microscopy." Park Scientific Instruments, Sunnyvale, California, 1996.

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[ 1 1]

Even though only the end of an SFM probe tip interacts with the molecule, the size of the tip dramatically affects the quality of images of nucleic acids. Tip irregularities away from the end are of importance only when imaging larger structures such as nucleosomes of chromatin. 7 For simplicity, the shape of the end of the tip is routinely assumed to be spherical when estimating actual feature widths from the apparent widths seen in micrographs. 8 The relationship between the observed diameter, Dobserved, and tip radius, Rtip, is given in Eq. (1). Dobserved =

4

-I/2 (Rtiprfeature) -

(1)

Equation (1) is applicable only when the radius of the feature, rfeature, is less than or equal to the radius of tip. When Rti p
[1 1]

SFM OF NUCLEICACID COMPLEXES

239

vertical quarters. Deformation of the tube occurs when a strain is induced in the cylinder through appropriately applied voltages. This strain causes the cylinder to bend, thus moving the sample (or tip) laterally. An electrode attached to the center of the tube provides for extension or contraction of the tube vertically, thereby moving the sample (or tip) up or down. Bending of the scanner tube in the horizontal direction causes a bowing artifact in large-scale images.15 Bowing occurs because as the scanner bends, the probe tip moves out of the plane of the sample. The scanner must then contract to maintain a contact cantilever deflection angle, resulting in an image that appears to be curved. SFM images of single nucleic acid molecules are often "flattened" through use of graphics software provided with the microscope, even when fiat substrates are used. Image resolution is also related to scanner performance, size of the scan area, and the specifications of the analog-to-digital converter (ADC) used for converting the analog photodiode detector signal to a digital signal. Lateral resolution is determined by the scan domain and the number of samples per scan line whereas vertical resolution is determined by the scanner's maximum z displacement, the fraction sampled by the ADC, and the number of bits of the ADC. For example, when a 1.0-#m 2 image is acquired with 512 samples per line and scan lines per image, the lateral resolution is 1.95 nm (distance tip traverses divided by the number of samples per line). If this image is acquired with a scanner with 6.0-#m vertical displacement sampled over this entire z range by a 16-bit ADC, the vertical resolution is 0.9 A (z displacement divided by number of digitization bits). Tip geometry impacts only the apparent feature width, not its height. Calibration of the scanner is required for accurate measurement of nucleic acid contour or persistence lengths. Because lead zirconium titanate piezoelectric materials exhibit intrinsic nonlinearity, hysteresis, creep, aging, and cross-coupling effects, frequent calibration of the scanner is recommended) 6 Calibration standards are readily available, reducing relative errors in length measurements of single molecules below 3%. Colloidal gold can be used as an internal standard when imaging biomolecules. Colloidal gold particles have multiple uses as three-dimensional, incompressible, monodisperse, and spherical SFM imaging standards as well as particles that can be exploited to characterize scanning tip geometry. 17.~s Biological molecules are "soft" and require lower forces for successful imaging. ~9,20The total force exerted by the tip is composed of the force resulting 1_5j. p. Starink and T. M. Jovin, Surf Sci. 359, 291 (1996). 16 R. Durselen, U. Grunewald, and W. Preuss, Scanning 17, 91 (1995). 17 j. Vesenka, S. Manne, R. Giberson, T. Marsh, and E. Henderson, Biophys. J. 65, 992 (1993). 18 j. W. Carlson, B. J. Godfrey, and S. G. Sligar, Langmuir 15, 3086 (1999). 19 T. E. Schaffer, J. P. Cleveland, E Ohnesorge, D. A. Waiters, and P. K. Hansma, J. Appl. Phys. 80, 3622 1996. 20 C. A. Putman, K. O. Vanderwerf, B. G. Degrooth, N. E Vanhulst, and J. Greve, Appl. Phys. Lett. 64, 2454 1994.

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from deflection of the cantilever, adhesive forces, and capillary forces. Typically, when imaging nucleic acids, forces less than 10 nN are required for high-quality micrographs. Capillary forces between water layers found on the tip and sample surfaces in ambient air exceed this value. These forces can be minimized by performing the experiment under low-humidity conditions or under fluid. Imaging Drug-DNA Complexes

Preparing Substrates For SFM imaging, substrates should be atomically flat, exhibit strong affinity for the molecule to be imaged, and have minimal attraction to the tip. Commonly used atomically flat substrates are mica, silicon, and thin films of gold evaporated onto a rigid support (mica, glass, or silicon). Mica is a two-dimensional material that, when cleaved along crystallographic planes, presents an atomically smooth surface over several hundred square microns. Silicon wafers, polished to subnanometer roughness levels, afford an even larger area for imaging plasmid DNA. Evaporated gold films are also used as substrates, but grain size markedly depends on the method of preparation. 21 Atomically flat domains of several hundred square microns are possible when the films are prepared by the template stripping method. 22 Selection of the substrate impacts the immobilization mode(s) available. Established protocols for immobilization of nucleic acids involve either electrostatic immobilization. 23,24 or covalent interaction25 28 between the molecule and the surface. Electrostatic immobilization is quick and requires no modifications to the nucleic acid. Covalent attachment pins the nucleic acid to the surface at known points on the molecule and affords a stronger attachment to the surface. Cleavage of mica leaves a surface that readily adsorbs divalent cations. The resulting surface attracts the negatively charged phosphate groups on nucleic acids. This strategy is applicable to imaging under buffer29-3j and in air. 23

21 j. A. DeRose, T. Thundat, L. A. Nagahara, and S. M. Lindsay, Surf Sci. 256, 102 (1991). 22 R Wagner, M. Hegner, H. J. Guntherodt, and G. Semenza, Langmuir 11, 3867 (1995). 23 j. Vesenka, M. Guthold, C. L. Tang, D. Keller, E. Delaine, and C. Bustamante, Ultramicroscopy 42-44, 1243 (1992). 24 y. L. Lyubchenko, A. A. Gall, L. S. Shlyakhtenko, R. E. Harrington, B. L. Jacobs, P. 1. Oden, and S. M. Lindsay, J. Biomol. Struct. Dyn. 10, 589 (1992). 25 A. J. Leavitt, L. A. Wenzler, J. M. Williamsand, and T. R Beebe, J. Phys. Chem. 98, 8742 0994). 26 C. Bamdad, Biophys. J. 75, 1997 (1998). 27 E Wagner, M. Hegner, E Kemen, E Zangg, and G. Semenza, Biophys. J. 70, 2052 (1996). 28 L. A. Chrisey, G. U. Lee, and C. E. O'Ferrall, Nucleic Acids Res. 24, 3031 (1996). 29 H. G. Hansma and D. E. Laney, Biophys. J. 70, 1933 (1996). 3o M. Fritz, M. Radmacher, J. R Cleveland, M. W. Allersma, R. J. Stewart, R. Gieselmann, E Janmey, C. E Schmidt, and E K. Hansma, Langmuir l l , 3529 (1995). 31 C. Bustamante, C. Rivetti, and D. J. Keller, Curt. Opin. Struct. Biol. 7, 709 (1997).

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Imaging under buffer requires adjustment of solution pH to maintain negatively charged DNA and careful choice of divalent cation. Hansma and co-workers 29,30 have shown Ni 2+, Co 2+, or Zn 2+ is needed to promote binding of DNA to mica surfaces for imaging under buffer. Individual molecules can be adsorbed and released by varying the concentration of these cations in the buffer. Imaging in air requires the presence of a divalent cation in the DNA loading solution and a means to remove unwarranted deposits of buffer salts. The latter is achieved either by copious rinsing of the DNA-laden mica surface or by using volatile buffers that will not leave a salt residue (e.g., ammonium acetate). 32 We typically use a loading sample containing DNA at a concentration between 0.1 and 10 #g/ml in 200 mM ammonium acetate, 5 mM MgC12, adjusted to pH 7.0 with NaOH, in 18-Mr2 water. A 10- to 50-/zl aliquot is incubated on the mica surface for 10 to 60 min, depending on the concentration of DNA and the desired number of molecules per square micrometer. After incubation the mica is dipped in water, a 1 : 1 (v/v) mixture of water and ethanol, and then anhydrous ethanol to wash away any excess salt deposits from the buffers. Excess liquid is "wicked" away with a Kimwipe and the disk is blown dry with clean compressed chlorofluorocarbon gas or dry N2 directed normal to the disk surface. The sample is stored in a desiccator until imaged under low-humidity conditions (dry He or N2 atmosphere). Electrostatic immobilization of DNA to other substrates requires that these surfaces be chemically modified. The most popular method uses 3-aminopropyltriethoxysilane (APTES) to create an electropositive layer on mica, silicon, or glass. 24 Vapor transfer is preferred over direct contact of APTES-containing solutions to minimize the formation of multilayers on the surface. An advantage of this method is that APTES-derivatized surfaces can be stored in the dry state for long periods of time. Loading of the DNA on this surface is achieved by the same protocol as described; however, buffers need not contain divalent cations. Nucleic acids can be covalently attached to gold surfaces. 25-2s DNA molecules with either thiolated phosphates 25 or pendant thiol groups 26 will spontaneously from Au-S bonds with the surface. Alternatively, gold, mica, silicon, or glass surfaces can be modified with heterobifunctional linkers reactive to DNA. Commonly used heterobifunctional linkers contain thiol or silane moieties at one end and succinimidyl, carboxylic acid, or maleimide moieties at the other end. 27'2s Thiolated linkers react with gold surfaces whereas silanated linkers react with oxide functionality on mica, silicon, and glass. Pendant amino groups on the DNA react with succinimidyl or carboxylic acid moieties [in the presence of 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide] forming peptide linkages. Pendant thiol groups react with maleimide moieties forming thioether linkages. DNA has also been linked to mica with a psoralen-terminated alkylsilane. 33 With this 32 E. Droz, M. Taborelli, T. N. Wells, and R Descounts, Biophys. J. 65, 1180 (1993). 33 L. S. Shlyakhtenko, A. A. Gall, J. J. Weimer, D. D. Hawn, and Y. L. Lyubchenko, Biophys. J. 77, 568 (1999).

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heterobifunctional linker, the DNA molecule is held on the surface by intercalation (via hydrogen bonding and Jr-stacking interactions). This approach suggests that site-specific attachment of DNA to surfaces may be possible with monolayers possessing functionality that binds within the grooves of double-stranded DNA (dsDNA). Assayforlntercalation. Intercalators are flat, polyaromatic molecules that bind to duplex DNA by insertion between the base pairs without breaking Watson-Crick hydrogen bonds. Each intercalator lengthens the DNA by an amount equivalent to the van der Waals thickness of the intercalating moiety and unwinds the double helix. Although groove-binding ligands may cause changes in the tertiary structure of the nucleic acid (e.g., bending), none are known to lengthen the molecule. SFM provides a straightforward means for determining lengths of nucleic acids with high precision. Coury and co-workers have established SFM as a direct, rapid, and unambiguous assay for mode of binding of conventional and nonclassic ligands to DNA. 34-36 The assay directly measures the lengths of individual DNA molecules before and after incubation with DNA-binding ligands. DNA lengthening on ligand binding provides direct evidence of intercalation. Implicit in this assay is the assumption that the length and conformation of immobilized nucleic acid molecules are commensurate with their length and conformation in solution. Convincing evidence in support of this assumption has been obtained by these and other workers. 34'35'37 Measured lengths of electrostatically immobilized, isolated DNA molecules are, within experimental error, equal to the expected length for B-DNA. Molecular lengths are correlated with ethanol concentration in the DNA loading solution. Increasing ethanol concentration results in shorter molecular lengths, consistent with the well-established ethanol-induced DNA conformational change. At ethanol concentrations greater than 30% (v/v), measured lengths are equal to the expected length for A-DNA. 35'37 Interestingly, once immobilized on the surface, molecular lengths are fixed and no significant changes are observed over the course of several days or after rinsing in ethanol. Contour lengths can be determined by cumulative addition of line segment lengths, drawn along the molecule with off-line analysis software. 38'39 As apparent

34j. E. Coury, L. McFail-lsom,L. D. Williams, and L. A. Bottomley,Proc. Natl. Acad. Sci. U.S.A. 93, 12283 (1996). 35j. E. Coury, J. R. Anderson, L. McFaiMsom, L. D. Williams, and L. A. Bottomley,J. Ant. Chem. Soc. 119, 3792 (1997). 36D. T. Breslin, J. E. Coury, J. R. Anderson, L. McFail-isom, L. D. Williams, L. A. Bottomley,and G. B. Schuster, J. Am. Chem. Soc. 119, 5043 (1997). 37y. Fang, T. S. Spisz, and J. H. Hoh, Nucleic Acids Res. 27, 1943 (1999). 38 y. Fang, T. S. Spisz, T. Wiltshire, N. P. D'Costa, I. N. Bankman, R. H. Reeves, and J. H. Hoh, AnaL Chem. 70, 2124 (1998). 39T. S. Spisz, Y. Fang, C. K. Seymour,R. H. Reeves, J. H. Hoh, and I. N. Bankman, Med. Biol. Eng. Comput. 36, 667 (1998).

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widths of the molecules increase, bends in the molecule become obscured and can no longer be accurately measured, thus contributing to greater deviation from molecule to molecule. Thus, DNA molecules with ambiguous topology should be excluded. The contribution of tip shape and diameter to the contour length measurement depends on the length of the strand. Tip diameter has a minimal impact on measurements of long DNA fragments (e.g., linearized plasmids). For example, when imaging a linearized plasmid containing 10,305 bp with a 20-nm tip radius, tip effects contribute only ",~18 nm (or 0.5%) of the measured DNA length. However, when the expected contour length of the DNA strand is equal to or less than the diameter of the tip, measured lengths must be corrected for tip effects. This is illustrated in Fig. 2. To identify an intercalator by using the Coury assay, 34 a series of solutions containing the ligand and DNA in varying concentration ratios are prepared,

FIG. 2. Topographic images of DNA on mica, illustrating the impact of tip shape on feature dimension. (a) Linearized pBBHb in B-DNA confimaation. Bar: 500 nm. The contour length of the molecule is 3500 nm. (b) Two short dsDNA molecules (20 bp each) joined by a disulfide linkage, that is, [5'-S-(CH2)6-GCGAT(A)IoACTGG-3']. Bar: Apparent length of 50 nm.

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244

[ 11]

5500

5000

E e-

.ff e- 4 5 0 0 .J

< Z 0

~ ...... ~.q/

4000

j

.D.

.........

. ..........

O . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

, . - ~ .....

i1."

~,;

3500

I

I

I

I

[

I00

200

300

400

500

Intercalator Concentration,

FIG. 3. Plot illustrating the impact of exclusion number on the length of a completely saturated intercalator-DNA complex. Lengths were computed on the basis of 10,300 bp for the dsDNA, 0.34 nm per binding event, and a binding affinityof 5.0 x 104. n = 2 (--), 3 (<>),4 (©), or 5 (0).

applied to atomically flat substrates, and imaged. Images of isolated molecules are obtained for each solution and at several locations on the substrate. Contour lengths are determined, collected, and plotted as a function of total ligand concentration. If the ligand intercalates, molecular lengths should increase with ligand concentration. The length at saturation provides a simple means to determine the exclusion number for the intercalator. If, at saturation, a ligand occupies every potential site (every dinucleotide step), the length of B-DNA is twice that of the unintercalated molecule. If, at saturation, a ligand occupies only every other dinucleotide step, the length of B-DNA is 50% longer than that of the unitercalated molecule. Thus, the exclusion number, n, can be computed from the saturation length, Lsat, and the measured length obtained in the absence of intercalator, L0, according to Eq. (2): n -- -

L0

-

Lsat --

L0

(2)

The impact of exclusion number in plots of measured DNA length versus ligand concentration is shown in Fig. 3. Noninteger values for the exclusion number are

[ 1 1]

SFM OF NUCLEIC ACID COMPLEXES

245

5500 ] I

_

~

-

-

-

-

~

~

............

,~. . . . . . . v . : :

:v:::

5000~ c ~ , ~ - ' " ' " ~ . " " ....• ---".v.v~;::.".,,, .............. ..... .--------.-v'. -.e ............... ~," ,...,.% . . . . . . . . . ~¢~'¢..0'

4000

3500 i

i 0

50

,

,

,

100

150

200

Intercalator Concentration, ixM FIG. 4. Plot illustrating the impact of binding affinity on the curvature of a length of DNA versus intercalator concentration. Lengths were computed on the basis of 10,300 bp for the dsDNA, 0.34 nm per binding event, and an exclusion number of 2. Binding affinity of 5.0 x 104 (--), 7.5 × 10a (O), 1.0 × 105 (O), 2.5 x 105 (O), and 5.0 × 105 (O).

possible with this method. Noninteger exclusion numbers indicate either sequencespecific binding 4° or multiple configurations existing along the DNA molecule. 41 Multiple configurations originate from out-of-plane substituents on the intercalator blocking entrance to other binding sites, adjacent or not. Random filling of binding sites creates gaps along the DNA that result in apparent noninteger exclusion numbers. The binding affinity of an intercalator can be determined from the rising portion of the measured DNA length-versus-ligand concentration plot. Figure 4 shows the impact of increasing intercalator binding affinity on the curvature of DNA lengthversus-intercalator concentration traces. Binding affinity of an intercalator, K is defined as [occupied intercalation sites] K = (3) ]unoccupied intercalation sites] [free drug] The fractional increase in DNA length at a given ligand concentration indicates the fraction of intercalation sites occupied. These concentrations can be explicitly 40 j. j. Correia and J. B. Chaires, Methods Enzymol. 240, 593 (1994). 41 j. D. McGhee and P. H. yon Hippel, J. MoL Biol. 86, 469 (1974).

246

BIOPHYSICALAPPROACHES

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related to measured lengths by the expressions [Occupied intercalation sites] = ( ~ - ~ - ) occ pie interca a ion

[DNA]

(4)

=

[Freedrug]=Cf=llo-(LaL'~))[DNA]]

(6)

where L is the length of intercalated DNA molecule, Cf is the free drug concentration, I0 is total intercalator concentration, B is the number of base pairs per DNA molecule, and a is the lengthening per intercalation event. Direct substitution of these equations into Eq. (3) results in a function that is transcendental with respect to L. Solving this equation for I0 yields Eq. (4), which can be used to extract binding parameters from SFM length data. I°:K[B_

( g _ ~ ) ] +[DNA]

T

(7)

L0, [DNA], and B are, in most circumstances, precisely known experimental constants. Nonlinear least-squares analysis may be applied to compute values of K, n, and a from experimental data. Alternatively, the number of parameters to be computed may be reduced by setting a = 3.4 A, the van der Waals thickness of most intercalators. Whenever conformational changes in DNA are possible, caution should be exercised in assigning this variable as a constant. Note that of the parameters n and a are strongly correlated, using nonlinear least-squares fitting routines. Binding affinities determined in this way are comparable to values determined by conventional assays. SFM length data can be satisfactorily fit by Scatchard models 42 as well as more sophisticated models incorporating cooperativity, size exclusionf and binding heterogeneity.43 The Scatchard model is 1" --

Cf

:

(8)

K ( N - r )

where r is the number of ligands bound per base pair and N is the number of binding sites available per base pair (i.e., 1/exclusion number = The number of ligands per base pair can be computed from microscopic measurements according to Eq. (8):

1/n).

r

--

L -L0

aB

(9)

42 G. Scatchard, Ann. N.Y. Acad. Sci. 51, 660 (1949). 43 j. B. Chaires, in "Advances in DNA Sequences Specific Agents," (L. H. Hurley, ed.), p. 3. JAI Press, Greenwich, Connecticut, 1992.

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Empirical values of K and n are obtained by nonlinear least-squares fitting of the model to data. McGhee and von Hippel have put forth more complicated models incorporating cooperative interactions between ligands. 41 Implicit in either approach is the assumption that the independent variable r is precisely known. This assumption may not be valid when r is computed from SFM lengthening data. Uncertainties in length measurements propagate in computed values of r and Cf; this is especially significant at low ligand loading levels. Site oflntercalation. Coury and co-workers 44 have also demonstrated the utility of SFM for determining the site of intercalation. Biotinylated psoralen was intercalated into a linearized plasmid and the binding site was marked by reaction of the biotin moiety with streptavidin bound to a colloidal gold bead. Streptavidin alone can be used because the biotin-streptavidin conjugate is readily visualized in SFM images of DNA. Alternative labeling strategies now available include reaction of pendant amino or thiol termini on the intercalator with C60 or gold nanoclusters possessing succinimidyl or thiol surface functionality. 45 Assay f o r Groove Binding

Parts of each base pair are exposed in the two distinct grooves of doublehelical DNA. Thus, drugs that bind in either the major or minor groove do so with a high degree of sequence specificity. As yet, no SFM-based assay has been developed that provides a direct measure of ligand binding in the major or minor groove. Commercially available SFM tips are presently too broad to provide direct visualize the major or minor groove of the double helix. Such images require tips with diameters approximating the dimensions of the grooves. Until angstrom-sized tips become available, visualization of groove-bound ligands is limited to either changes in DNA strand height (contrast) and width, or to labeled ligands. Hansma et al. have observed differences in curvature between uncomplexed DNA and DNA bound with distamycin, a well-known minor groove binder. 46 To our knowledge, this is the only published study involving minor groove binding. Binding in the minor groove can be assessed by a competitive binding approach, 34-36 For example, ethidium will compete with a drug that binds exclusively in the minor groove. On intercalation, the ethyl and phenyl substituents of ethidium are resident in and block the DNA minor groove, whereas the major groove remains sterically unencumbered. Thus, when the ratio of these competing ligands is systematically varied, the length of the DNA complex should scale with ethidium concentration but be significantly less than that observed in the absence of the minor groove-binding ligand. Note that samples must be prepared and imaged 44j. E. Coury,L. McFail-lsom,S. Presnell, L. D. Williams,and L. A. Bottomley,,I. Vac. Sci. Technol. A 13, 1746(1995). 4.5K. Sugawara,S. Tanaka,and H. Nakamura,Anal. Chem. 67, 299 (1995). 46 H. G. Hansma,K. A. Browne, M. Bezailla,and T. C. Bruice. Biochemisow 33, 8436 (1994).

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in the same manner employed for intercalator-DNA complexes alone. The major groove-binding mode can be identified through competitive binding experiments as well. Ditercalinium, an intercalator with substituents that obstruct the major but not the minor groove, competes with major groove binders. Nogalamycin, an intercalator with substituents in both grooves, will compete with all groove-binding ligands. Several workers have acquired images of oligonucleotides bound in the major groove.4%49 Hansma et al. observed triple-stranded DNA structures in images of poly(dA) • poly(dT) and poly(dC), poly(dG).47 These structures were twice as high as double-stranded DNA and the same width. Cherny and co-workers4s presented SFM images of a G,A-containing triplex-forming oligomer conjugated to a long plasmid DNA. Triplex regions were 40.4 nm higher than the corresponding duplex regions. Pfannschmidt and co-workers49 carried out site-specific labeling of closed circular DNA by using triplex-forming oligonucleotides containing a reactive psoralen at the 5' end and a biotin at the 3' end. The probes were directed to different target sites on plasmid DNA and photocross-linked to the target to increase stability. The covalent adduct DNAs were visualized by SFM, using avidin or streptavidin as protein tags for the biotin group on the oligonucleotide probes. Height measurements in the region of the triplex were consistent with those observed by Hansma et al. and by Chemy et al. Imaging Protein-DNA Complexes Analysis of the structure of protein-nucleic acid complexes is an important area of application of SFM. Protein-DNA complexes are prepared and immobilized electrostatically onto mica substrates by the protocol described above for immobilizing drug-DNA complexes. Structural details of the protein-nucleic acid complex are obtained from analysis of images acquired in air or under buffer. For example, SFM images acquired by Cary et al. 5° revealed sequence-independent DNA looping by Ku protein and the DNA-dependent protein kinase, two proteins responsible for DNA repair and genetic regulation. Images by Pang et al. 51 showed that Ku protein binds predominantly to the ends of double-stranded linear DNA, does not bind to circular plasmids, and joins two ends of DNA together. These observations suggest that Ku protein may play a role in physically orienting DNA 47 H. G. Hansma, I. Revenko, K. Kim, and D. E. Laney, Nucleic Acids Res. 24, 713 (1996). 48 D. ][. Cherny, A. Fourcade, E Svinarchuk, R E. Nielsen, C. Malvy, and E. Delain, Biophys. J. 74, 1015 (1998), 49 C. Pfannschrnidt, A. Schaper, G. Heim, T. M. Jovin, and J. Langowski, Nucleic' Acids Res. 24, 1702 (1996). 5o R. B. Cary, S. R. Peterson, J. Wang, D. G. Bear, E. M. Bradbury, and D. J. Chen, Proc. Natl. Sci. U.S.A. 94, 4267 (1997). 51 D. Pang, S. Yoo, W. S. Dynan, M. Jung, and A. Dritschilo, Cancer Res. 57, 1412 (1997).

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for ligation by binding the ends of adjacent DNA molecules. Valle et al. 52 studied the interaction of DNA with bacteriophage 429 connector. SFM images revealed that the circular DNA-protein complex bends the strands to a mean angle of 132 ° and that the DNA binds to the outer side of the protein. Garcia and co-workers 53 determined that M . H h a I , a cytosine C5 DNA methyltransferase, causes only a 2 ° bend on binding its recognition site. In contrast, images of the M . E c o R I , an adenine N6 DNA methyltransferase, shows an average bend angle of ~ 5 2 °. This distortion of DNA conformation by M . E c o R I is important for sequence-specific binding. Formation of protein-DNA complexes can also be used to mark specific sites on the nucleic acid. For example, Allison and co-workers 54'55 used mutant E c o R I endonuclease binding to specific nucleotide sequences to create distinctive features in images of plasmids. Binding of the mutant endonuclease to specific nucleotide sequences produced distinctive features in images of plasmids. By measuring contour length distances between these features, physical maps of individual cosmid DNAs were produced by direct SFM imaging with an accuracy of better than 1%. This method is faster and more accurate when compared with conventional electrophoretic mapping methods. Klinov and co-workers 56 performed high-resolution mapping of individual plasmids and cosmids, using RNA probes specific for long terminal repeats within these DNA. The RNA probes formed so-called R-loops that, when stabilized by glyoxal, were readily imaged after chemisorption of the conjugate onto Mg2+-modified mica. R-loop positions were accurate to 0.5% of the cosmid length. SFM imaging of protein-DNA complexes has been used to gain insight into their function. For example, the repression of transcription of two overlapping promoters of the g a l operon in E s c h e r i c h i a coli requires Gal repressor (GalR) and the histone-like protein HU. Lyubchenko et al. 57 imaged Gal-DNA complexes with proteins and found that GalR mediated DNA looping in which HU plays an obligatory role by helping GalR tetramerization. Nettikadan and co-workers 58 investigated transcription factor AP2 binding to DNA. SFM images proved that protein-binding sites can be mapped over a few kilobases of target DNA and 52 M. Valle, J. M. Valpuesta, J. L. Carrascosa, J. Tamayo,and R. J. Garcia, J. Struct, BioL 116, 390 (1996). 53 R. A. Garcia, C. J. Bustamante, and N. O. Reich, Proc. Natl. Acad. Sci. U.S.A. 93, 7618 (1996). 54 D. P. Allison, P. S. Kerper, M. J. Doktycz,T. Thundat, P. Modrich, F. W. Larimer, D. K. Johnson, P. R. Hoyt, M. L. Mucenski, and R. J. Warmack, Genomics 41,379 (1997). 55p. R. Hoyt, M. J. Doktycz,P. Modrich, R. J. Warmack,and D. P. Allison, Ultramicroscopy 82, 237 (2000). 56D. V. Klinov, I. V. Lagutina, V. V. Prokhorov,T. Neretina, P. P. Khil, Y. B. Lebedev, D. I. Cherny, V. V. Demin, and E. D. Sverdlov,Nucleic Acids Res. 26, 4603 (1998). 57y. L. Lyubchenko,L. S. Shlyakhtenko,T. Aki, and S. Adhya,Nucleic Acids Res. 25, 873 (1997). 58 S. Nettikadan, F. Tokumasu, and K. Takeyasu,Biochem. Biophys. Res. Commun. 226, 645 (1996).

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that multimerization state of DNA-binding proteins can be determined simply by measuring the sizes of proteins bound of the DNA. Rippe et al. 59 used SFM to study transcriptional activation of E. coli RNA polymerase ¢r54 at the glnA promoter by the constitutive mutant of the nitrogen regulatory protein C. DNA-protein complexes were deposited on mica and imaged in air. By choosing appropriate conditions, the structure of various intermediates in the transcription process could be visualized and analyzed. Smith et al. 6° investigated the complexes formed between purified poly(A)-binding protein and poly(A) RNA, using SFM. Poly(A)-binding protein is an RNA-binding protein that binds specifically to the poly(A) tail of mRNAs in eukaryotes. SFM images revealed that the protein formed variable-size complexes bound lengthwise along the RNA. Poly(A) RNA alone appeared to contain a knoblike structure that largely disappeared once the protein was bound. Margeat and colleagues 61 visualized the protein-protein and protein-DNA complexes involved in transcriptional regulation by the trp repressor (TR). Plasmid fragments bearing the natural operators trpEDCBA and trpR, as well as nonspecific fragments, were deposited onto mica in the presence of varying concentrations of TR and imaged. Specific and nonspecific complexes of TR with DNA are found, as well as free TR assemblies directly deposited onto the mica surface, Their findings suggest protein-protein interactions serve a role in transcriptional regulation by the trp repressor. Perhaps the most exciting application of SFM lies in the real-time visualization of protein-DNA complexation. The major challenge is to find imaging conditions that promote adsorption of protein-DNA complexes onto atomically fiat substrates without destabilizing or inhibiting movement of the complex. Once these conditions have been found, reconstruction of images acquired over time provides a straightforward means for analyzing motion in biological systems. Thomson and co-workers 62 prepared recombinant RNA polymerase containing histidine tags (His-RNAP) on the C terminus and immobilized them onto ultraflat gold via a mixed monolayer of alkanethiols. Specific binding of this molecule to a 42-base circular single-stranded D N A template was confirmed by in situ SFM images showing the production of huge RNA transcripts. Bustamante and co-workers 63'64 obtained tapping-mode SFM images that demonstrated the diffusion of E. coli RNA polymerase along DNA. Direct evidence 59K. Rippe, M. Guthold,E H. von Hippel, and C. J. Bustamante,Mol. Biol. 270, 125 (1997). 60B. L. Smith,D. R. Gallie, H. Le, and P. K. Hansma,J. Struct. Biol. 119, 109 (1997). 61E. Margeat,C. Le Grimellec,and C. A. Royer,Biophys. J. 75, 2712 (1998). 62N. H. Thomson,B. L. Smith, N. Almqvist,L. Schmitt, M. Kashlev,E. T. Kool,and P. K. Hansma, Biophys. J, 76, 1024 (1999), 63E. T. Kool, P. K. Hansma, M. Kashlev, S. Kasas, N. H. Thomson, B. L. Smith, H. G. Hansma, X. Zhu, M. Guthold, and C. Bustamante,Biochemistry 36, 461 (1997). 64C. Bustamante,M. Guthold, X. Zhu, and G. Yang,J. Biol. Chem. 274, 16665 (1999).

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of facilitated targeting of RNA polymerase by intersegment transfer and possibly hopping (intradomain association and dissociation) was obtained for the first time. Ternary intermediates, in which RNA polymerase appears to be simultaneously bound to two DNA segments, were directly observed during intersegment transfer events. In some image sets, transfer events were preceded and followed by sliding processes. Their results provide additional insight into the mechanism by which RNA polmerase searches for the promoter, a key step in transcription. Their highresolution images and analysis provided the first direct evidence of individual complexes involved in transcription. Summary SFM is a viable and effective method for determining the mode of binding, the extent of binding, and the site of binding of intercalators to nucleic acids. Establishing the presence of a groove-bound ligand can be achieved either by competitive binding experiments with a well-defined intercalator (minor groove) or by changes in apparent contrast {major groove). In our opinion, SFM has an important role in resolving the structural polymorphisms for small molecule-DNA complexes. Application of these assays in the study of polyintercalator molecules is currently underway in our laboratory. SFM is an important, new tool in the study of protein-DNA complexes. New insights into the structure and function of these complexes are enabled by real-time visualization. Currently the temporal resolution of the SPM limits the degree to which definitive rate data can be determined. Several binding and unbinding events could take place in the time it takes to acquire one image. New developments in SFM technology will allow faster scanning and will improve the temporal resolution of so-called SFM movies. To this end, the Hansma group is developing small cantilevers 65 and improved optical deflection systems 66 to enable intermittent imaging at scanning rates of 1.7 sec per image. These improvements will enable SFM visualization of complex biological processes as they occur, one molecule at a time. Acknowledgments Financial supportfrom the ONR-sponsoredGeorgiaTech MolecularDesign Institute is gratefully acknowledged.We thankthe followingindividualsfor their insights, ingenuity,and involvementin the developmentof SFM assaysfor drug-DNAbinding: JosephCoury,JaimieAndersen,LorenWilliams, Lori McFail-Isom,and Brad Chaires. 65M. B. Viani, T. E. Schaffer, G. T, Paloczi, L. I. Pietrasanta, B. L. Smith, J. B. Thompson, M. Richter, M. Rief, H. E. Gaub,K. W. PLaxco,A. N. Cleland, H. G. Hansma,and P. K. Hansma,Rev. Sci. lnstr. 70, 4300 (1999). 66T. E. Schafferand P. K. Hansma,J. Appl. Phys. 84, 4661 (1998).

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[12] Nuclear Magnetic Resonance Studies of Drug-DNA Complexes in Solution By ANDREWN. LANE Introduction There is a pleasing logic about targeting DNA to regulate cellular function, whether to modulate the amount of expression of a particular gene, or to interfere with DNA replication. Gene targeting automatically bypasses two amplification steps (transcription and translation), so that the concentration of drug required can potentially be low, assuming that high affinity can be attained. The affinity problem can be circumvented by forming covalent bonds, and there are numerous DNA-directed drugs in this class, such as mustard alkylators, cisplatin, and others I (see [17] in this volumela). However, the other side of the coin is specificity, not only toward particular DNA sequences, but also against other cellular components, especially proteins. To optimize both specificity and affinity, it is necessary to understand the conformational features of the interacting components as well as the mechanism and the thermodynamics. Hence structural studies are an essential complement to thermodynamic, kinetic and functional analysis of drug-DNA interactions. Of the structural methods, nuclear magnetic resonance (NMR) may be particularly well suited to the DNA problem as it is possible to work with any desired DNA sequence (not all DNA sequences crystallize in the B-form) in aqueous solution. NMR can be used in both its structural and its spectroscopic modes. The latter is especially useful for mapping binding sites by titration, and for studies of the dynamics involved in the interaction. Multiple conformations and degrees of hydration, which are needed for a molecular understanding of the thermodynamics of drug-DNA interactions, can also be studied by NMR. The goals are to determine the modes of binding and their stoichiometry, the changes in conformation and dynamics of the DNA and the ligand on binding, and the influence of hydration on the complex. In this chapter, the essential requirements for an NMR investigation of the conformation and dynamics of DNA and interaction with drugs are outlined, with specific examples demonstrating the utility of this approach for systems of different complexity, ranging from simple groove binders that do not distort the DNA, intercalators that do distort the DNA, to the major groove-binding oligonucleotides. l B. Koberle, J. R. Masters, J. A. Hartley, and R. D. Wood, Curr. Biol. 9, 273 (1999). ~a C. J. M c D a r k , R J. M c H u g h , N. J. Tilby, K. A. Dremaldi, and J. A. Hartley, Methods Enzymol. 340, [17] 2001 (this volume).

METHODS IN ENZYMOLOGY,VOL.341)

Copyright© 2001 by AcademicPress All rights of reproductionin any form reserved. 0076-6879/00$35.00

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Spectroscopy

The basic procedure for NMR analysis of d r u g - D N A interactions is the same regardless of the particular system, and involves assignment of the signals of the DNA and drug, both free and in the complex. As the methods for assigning the 1H NMR spectra of DNA and DNA-drug complexes have been reviewed extensively, 2-5 they are not reviewed further here. However, increasing use is being made of heteronuclear NMR as hardware continues to improve, and also as biochemical and chemical methods become available for complete or partial isotopic enrichment with 13C and 15N. J3C spectra are readily assigned by correlation with the JH spectrum, using heteronuclear single quantum coherence (HSQC), which is sufficiently sensitive to work well at natural abundance. The 31p NMR spectra can be similarly assigned, and if the scalar coupling is retained, the value of ~IPH3' can be determined from the spectrum (and see below). NMR spectroscopy is an information-rich method, and many of the spectral parameters can be interpreted directly in terms of conformation or dynamics. As with any kind of spectroscopy, each peak (or resonance) represents a transition between a ground and an excited state. The frequency of the resonance (the chemical shift) is in principle different for each magnetically active nucleus in the molecule. This arises mainly from a combination of the details of chemical bonding and electronegativity of atoms, and to the local magnetic fields generated by the electrons in the bonding network. It is this sensitivity to chemical environment that makes NMR such a powerful form of spectroscopy. Chemical Shifts

As IH, 13C, l~N, and 31p are all present in nucleic acids, there is a large number of signals present that can report on structure, dynamics, and the effects of ligand binding. The chemical shift is itself difficult to interpret in a quantitative sense. Nevertheless, there are useful general conclusions that can be drawn from specific chemical shift values, or changes due to binding of a ligand. A good correlation between 31p NMR shift and conformation about e and ~" has been demonstrated6; a substantial downfield shift in 31p NMR correlates well with conversion from the usual BI conformation @, ~" =- 180, - 6 0 ) to the less stable BII conformation (e, ~" = - 6 0 , 180), which is often found in mismatched DNA and RNA molecules, 7-9 2 B. R. Reid, Q. Rev. Biophys. 20, 1 (1987). 3 B. A. Borgias, M. Gochin, D. J. Kerwood,and T. L. James, Prog. Nuel. Magn. Reson. Spectrose. 22, 83 (1990). 4 M. S. Searle, Prog. Nucl. Magn. Resort. Spectrosc. 25, 403 (1983). 5 j. Feigon, K. M. Koshlap, and E W. Smith, Methods Enzymol. 261, 225 (1995). 6 D. G. Gorenstein, Methods Enzymol. 211, 254 (1992). 7 A. N. Lane, S. R. Martin, S. Ebel, and T. Brown, Biochemistry 31, 12087 (1992). 8 S.-H. Chou, J.-W. Cheng, and B. R. Reid, J. Mol. Biol. 288, 138 (1992).

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and also in intercalated drug-DNA complexes. 4 However, upfield shifts are also found under some circumstances, which presumably have a different origin. Ring currents from an aromatic ligand can be interpreted in general terms because the nature of ring current shifts is reasonably well understood. I°, Jl The direction and magnitude of the shift can be related to the position of the nucleus with respect to the ring current field of the ligand, 12 provided there are no other effects from conformational changes that can affect the DNA resonances, which are largely determined by the ring currents of the adjacent bases. This is likely to be more straightforward for groove-binding drugs than for intercalators, which certainly alter the conformation of the DNA (see below). In principle, the magnitude of the ring current shift could be incorporated into the structure refinement process in the same way as are many other experimental parameters. ~3 Such calculations are likely to increase in importance as computer programs become more sophisticated. Chemical shift changes due to binding are either environmental and/or conformational; they are nevertheless convenient for monitoring a titration and thereby determining the end points. The chemical shift titration is perhaps the simplest NMR experiment that provides detailed information about stoichiometry of binding, kinetics, and location of the binding site. Assuming that the resonances in the free DNA and ligand have been properly assigned, the titration may produce changes in shifts of a subset of resonances of both components. The affected resonances may move continuously from the initial frequency to a final (saturated) frequency (the last exchange limit), remain at the original frequency but decrease in intensity (slow exchange limit), or both shift and broaden while the fractional saturation is <1 (fast intermediate exchange). At some point saturation should be achieved, from which a limit to the binding stoichiometry can be determined. This will be accurate if the concentration of both components is >Kd, which is the usual situation for NMR unless rather weakly binding ligands are being studied (see below). Otherwise, the value of Kd can itself be determined. As the initial and final chemical shifts can be measured unambiguously, the change in shift can be converted to a frequency difference, which sets a time scale for the interaction kinetics. Thus, if slow exchange conditions are observed for all resonances over the entire titration, then the dissociation rate constant koff must be at least an order of magnitude smaller than the smallest observed shift difference. As rather small frequency differences can be measured (e.g., <5 Hz), and generally the shift perturbation decreases with increasing distance from the binding site, this can be

9 E Legault and A. Pardi, J. Magn. Reson. 103B, 82 (1994). l0 C. Giessner-Prette and B. Pullman, Q. Rev. Biophys. 20, 113 (1987). t l D. A. Case, Curl: Opin. Struct. Biol. 8, 624 (1998). 12 j. A, Parkinson, J. Barner, K. T. Douglas, J. Rosamund, and D. Sharpies, Biochemistry 29, 10181 (1990). 13 G. Cornilescu, F. Delaglio, and A. Bax, J. Biomol. Nucl. Magn. Reson. 13, 289 (1999).

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fairly stringent. As a rule of thumb, ligands that bind with a Kd value < 1 /zM tend to show mainly slow exchange behavior. For single-step binding kinetics, the dissociation rate constant is expected to be about 108 Kd sec -l. For Kd in the range of 1 to 50/zM, koffmay be in the range of 100 to 5000 sec -1 and so, for moderate shift differences, give rise to fast intermediate exchange. Under these conditions, the line width as well as the frequency of the averaged peak vary throughout the titration, with the maximum line broadening observed at 33% saturation. If Kd is known independently, the dependence of the line width (which equals R2/sr) on the degree of saturation can be analyzed to provide the binding rate constants. The rate constants can also be determined by careful analysis of the spin-locking field strength dependence of Ti p. 14 Rip = R°p + 4:rr2pa(1 -

pa)Av2k/(og~ + k 2)

(1)

where R°p is the relaxation rate constant in the absence of exchange, pa is the mole traction of state A, Av is the frequency difference (in hertz), k is the exchange rate constant = kon[L] ÷ koff, and oJI is the spin-lock field strength. In the absence of a spin-lock field, Eq. (1) gives the spin-spin relaxation rate constant in the presence of exchange. Because Av can be determined accurately from the end points of the titration, then measurements of R~p at different spin-lock field strengths immediately gives k, and if carried out at different ligand concentrations, both k and Kd = koff/koncan be determined. Accurate measurement of line widths requires resolved resonances and fiat baselines. The influence of magnetic inhomogeneity also needs to be accounted for. Modern shim systems can usually reduce the magnetic field inhomogeneity to about l to 1.5 Hz for iH (nonspinning sample), and most DNA line widths are substantially broader than this (e.g., a purine H8 will have a line width of about 4 Hz at a correlation time of 3 nsec). The inhomogeneity can be controlled by subtracting the observed line width of a resonance whose intrinsic line width is much smaller than the residual inhomogeneity, such as the formate resonance. Alternatively, a Carr-Purcell-Meiboom-Gill (CPMG) spin--echo experiment with composite 180 ° pulses gives an accurate value for T2 .14 Tip experiments can be easily done using the continuous wave (CW) spin-lock method 14"15 with a continuous calibrated Bi pulse from the transmitter. The main difficulty with the Tip experiment is limitations on the B1 field that can be tolerated by the probe. This depends not only on the power, but also on the duration, of the pulse. Unfortunately, as the spinlock field strength is increased, the Tip value increases, requiring longer spin-lock times in the experiment. This means significantly higher heating, which in turn can affect the peak position (exchanging systems are usually rather temperature sensitive), and the precise temperature of the experiments. As activation energies 14 D. G. Davis, M. E. Perlman, and R. E. London, J. Magn. Reson. 104B, 266 (1994). I5 R. Freeman and H. D. W. Hill. J. Magn. Reson. 55, 1985 (1971).

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can be large, this may mean that the exact temperature for calculating the rate constant is not fully determined. Equation (1) is correct on resonance. For small values of Bl, there can be substantial offset effects in the tilted field. Fortunately as the end points are known, the degree of the tilted field can be calculated for any col, and accounted for [Eq. (2)]. Rip = RI sin e ~ + Re c o s 2 ~

(2)

where ~ is the tilt angle, equal to cos -1 [~ox/(co~ + Aco2)1/2], Aco is the offset of resonance from the carrier frequency, and col is the radio frequency field strength.14 In addition to thermodynamic and kinetic information, the titration curve also provides some information about the location of the binding site(s). When the chemical shift changes (i.e., -4- ligand) are plotted against the DNA sequence, it is often found that the shift changes cluster around a small number of bases pairs, and show maximum shift differences flanked by decreasing values. The maximum differences generally correlate well with the location of the binding site. This is in part because DNA is extremely flexible and conformational distortions are not propagated far along the duplex (typically only the nearest and next nearest neighbors are affected), and ring current effects from the bases are comparatively short range (also nearest and next nearest neighbors). It is not always possible to follow the entire titration for all resonances; the end points therefore often need to be obtained by reassigning the resonances in the complex using the full battery of two-dimensional techniques. Chemical shifts of N and NH are rather sensitive to hydrogen bonding, and to the nature of the acceptor or donor atom. Thus, in lSN-labeled oligonucleotides, the chemical shift of the nitrogen is a good indicator of whether it is involved in a hydrogen bond, and the relative strength of the hydrogen bond.16 13C shifts are also useful indicators of type of atom [e.g., adenine (Ade) C2] and are sensitive to changes in aromaticity such as on protonation of aromatic rings (e.g., at the N3 of cytosine in DNA triplexes; see below). 17

Referencing For chemical shifts to be useful in an absolute sense, and also when titrations or thermal studies are to be carried out, it is essential to have an appropriate internal shift references. For DNA, 2,2'-dimethylsilapentane 5-sulfonate (DSS) is a good standard, because being negatively charged, it does not interact significantly with DNA. It is water soluble, and usually its NMR, spectrum does not interfere with DNA resonances. It is simultaneously a good proton and Z3Cinternal reference, and

16 B. L. Gaffney, P. P. Kung, C. Wang, and R. A. Jones, J. Am. Chem. Soc. 117, 1228 l (1995). 17 J.-L. Asensio, T. Brown, and A. N. Lane, Nucleic Acids Res. 26, 3677 (1998).

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the methyl resonance is independent o f p H in the range 4 < pH < 10. Once a good internal reference is available, other nuclei can be straightforwardly referenced to the proton spectrum, using the gyromagnetic ratio. This is routinely used for 13C and 15N in protein studies, 18 and can also be done in the same manner for 31p. This is especially important as the primary standard for 31p is external 85% phosphoric acid. Coupling Constants

Coupling between magnetic nuclei through the covalent network gives rise to splitting of resonances. The magnitude of the splitting, J, is called the scalar coupling constant and is independent of the magnetic field strength. The three-bond coupling constant, especially between protons, is useful because of its dependence on the dihedral angle. Thus in an HCCH fragment, the coupling constant 3JHH depends on whether the two protons are gauche or trans. This is particularly useful for analyzing the conformation of the sugar rings in both DNA and RNA. Because the sugars are substituted furanoses, there is a limit to the number of conformational states that can be accessed, and the number of parameters needed to describe the conformation. If drug binding causes a change in the sugar conformation, it should be reflected in the sums of coupling constants E v , Y~2,, and ~2',. The three-bond coupling constant is related to the dihedral angle q~ by the Karplus equation: J = A cos(0) + B cos(~b) + C

(3)

where A, B, and C are constants determined by parameterization against a database of known nucleotide structures. The dependence of 3J1,2,, 3Ji,2,, , etc., on the pseudorotation phase angle (P) and maximum amplitude (q~m) have been worked out in detail, including electronegativity corrections. ~9'2° These expressions can be used either in tabular or graphical form, or simply be programmed to allow optimization of structure against the experimental coupling constants. The analysis of three-bond coupling constants requires resolved resonances, and line widths narrower than the coupling constant to be measured. Even then, the recovery of coupling constants can be difficult, and may require significant computational effort. 21'22 For larger oligonucleotides, it appears that dipolar cross-correlation 18D. S. Wishart, C. G. Bigam, J. Yao, E Abildgaard, H. D. Dyson, E. Oldfield, J. L. Markley, and B. D. Sykes,J. BiomoL Nucl. Magn. Reson. 6, 135 (1995). 19j. van Wijk, B. D. Huckriede, J. H. lppel, and C. Altona,Methods Enzymol. 211,286 (1992). 2oS. S. Wijmenga, M. M. W. Mooren, and C. W. Hilbers, "NMR ofMacromolecules: A Practical Approach" (G. C. K. Roberts, ed.), pp. 217-288. IRL Press, Oxford, 1993. 21 U. Schmitz, G. Zon, and T. L, James, Biochemistry 29, 2537 (1990). 22U. Schmitz, I. Sethson, W. M. Egan, and T, L. James, J. Mol. Biol. 227, 510 (1992).

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effects can be quite severe, and make it difficult to recover individual coupling constants. 23'24 However, sums of coupling constants, measured from the separation of the outermost components of the multiplet, do not suffer this limitation, and if three such sums can be measured, then a reasonable description of the sugar puckering can be determined, especially when supplemented with some accurate nuclear overhauses effect (NOE) information about H l ' - H 4 ' distances. 25 A proper treatment of coupling constants is an important aspect of conformational analysis 2s,26 and heteronuclear couplings in labeled DNAs are likely to make such measurements of sugar conformation even more reliable. 27 The sugars can exist in a number of conformations separated by comparatively low energy barriers. The energy wells of each state (roughly corresponding to C2'endo or"S" and C3'-endo or "N") are themselves quite broad. This implies that the sugar conformations constantly fluctuate about a range of related conformations (i.e., the S or N states), and occasionally interconvert between different conformers (e.g., N and S). In general, the S states are more highly populated than the N states in duplex DNA, although more equal distributions are sometimes found in DNA. RNA hybrid duplexes, and the Hoogsteen strand of parallel triple helices (see below). Each conformation requires at least two parameters to describe it fully (a pseudorotation phase angle and a maximum amplitude or, alternatively, any two of the ring torsion angles). Hence, two coupling constants or two sums of coupling constants are sufficient to determine a single conformation. However, if two conformations are present, as is likely given the energetics of sugar puckering, then at least five parameters are needed to specify the equilibrium properties (two structural parameters for each conformation, and an equilibrium constant). Clearly this in general is not possible. If one state dominates, then the minor species can often be modeled as a C3'-endo state, so that only three parameters are needed to specify the system (two conformational parameters plus the equilibrium constant). The coupling constants average in a straightforward manner, as the interconversion is much faster than the difference in coupling constants (fast exchange). The question usually is whether a dynamic equilibrium of two states (e.g., N and S) is a better representation than an intermediate conformation (e.g., O4'-endo). In principle, these can be distinguished by accurate coupling constants and critical NOEs (H I ' - H 4 ' and H2"-H4'). However, the data are not always of sufficient quality to give an absolutely unambiguous answer. Where sufficient high-quality data have been obtained, they support the dynamic equilibrium model, as do molecular 23 L. Zhu, B. R. Reid, M. Kennedy, and G. E Drobny, J. Magn. Resort. I l i A , 195 (1994). 24 T. J. Norwood, J. Magn. Reson. I14A, 92 (1995). 25 M. R. Conte, C. J. Bauer, and A. N. Lane, J. Biomol. Nucl. Magn. Resort. 7, 190 (1996). 26 U. Schmitz and T. L. James, Methods Enzymol. 261, 3 (1995). 27 j. H. Ippel, S. S. Wijmenga, R. de Jong, H. A. Heus, C. W. Hilbers, E. de Vroom, G. A. van der Marel, and J. H. van Boom, Magn. Resort. Chem. 34, S156 (1996).

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modeling with realistic force fields and measured dynamics. In terms of structure calculations, the single conformation versus the dynamic model are different. The assumption underlying the single conformation model is that the experimental data represent a single set of parameters, with any averaging restricted to local smallamplitude motions. In the dynamic model the observed data are the result of an averaging process over an ensemble of conformations. This is a much more difficult problem, and in general the experimental data available are insufficient to specify the entire ensemble in detail. Numerous approaches to this problem have been proposed, with varying degrees of s u c c e s s . 26'28-34 (and see below). However, a major advantage of ensemble calculations is that they relate more closely to the actual behavior of DNA in solution, and with the thermodynamics of ligand binding. Analyzing coupling constants and NOE data together for sugar conformations can be done exhaustively as a grid search on the (up to five) parameters needed to describe a two-state conformational equilibrium. In essence, each parameter is allowed to vary between lower and upper bounds, such as P~, q~m, and ~, and as finely graded as desired. The NOEs must also be weighted properly. The proper weighting depends on the time scale of the averaging; 1" 6 averaging is usually assumed in practice. An error function is calculated at each step and stored, and at the end of the calculation (or as a running estimate), the minimal value of E is found. A commonly used error function is E : Z(Jc

- J 0 ) 2 q-

~~(Nc - N 0 ) 2

(4)

where Jc and J0 are the calculated and observed coupling constants (or sums), and No, and No are the corresponding NOE values. This corresponds to the best agreement with the data. As there are likely to be numerous conformations that agree with the data within the experimental error, all possible solutions should be listed, so that a range or variance of each parameter can be estimated. With good quality data, the values of Ps and f~ can be determined with reasonable accuracy and precision (about 4- 20 ° and + 0.05, respectively), whereas the maximal amplitude is often not well determined. Note also that a fairly wide range of Ps ( 4- 20 °) is not only a consequence of the form of the functional dependence of the coupling constant on P in the S range, but is also representative of the broad energy well in this region of conformational 28 A. M. J. J. Bonvin, R. Boelens, and R. Kaptein, J. Biomol. Nucl. Magn. Reson. 4, 143 (1994). 29 A. M. J. J. Bonvin and A. T. Briinger, J. Mol. Biol. 250, 80 (1995). 30 A. M. J. J. Bonvin and A. T. Briinger, J. Biomol. Nucl. Magn. Reson. 7, 72 (1996). 3t C. Gonzalez, W. Stec, M. A. Reynolds, and T. L. James, Biochemistry 34, 4969 (1995). 32 N. B. Ulyanov, U. Schmitz, A. Kumar, and T. L. James, Biophys. J. 68, 13 (1995). 33 j. I. Gyi, A. N. Lane, G. L. Conn, and T. Brown, Biochemistry 37, 73 (1998). 34 A. N. Lane, in "Molecular Modeling of Nucleic Acids" (N. B. Leontis and J. SantaLucia, eds.), pp. 106--121. ACS Symposium Series 682. American Chemical Society, Washington, D.C., 1998.

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space. Regardless of the accuracy of the coupling constants, quoting P values to several decimal places makes no physical sense. If drug binding alters the equilibrium distribution by more than 0.05 and/or the optimal value of P~ by more than about 25-30 °, then coupling constants can be used to analyze this drug-dependent change in detail. The influence of drug binding on sugar conformations can be assessed in an unbiased manner in this way, as changes in coupling constants will be consistent either with changes in actual conformations, or the distribution of N and S pucker states. The three-bond coupling constants H 3 ' - P and H5'/H5"-P are also valuable, along with shift information, for providing information about e and t~ in the backbone, which may be sensitive to ligand binding, especially for intercalation (see below). One-bond CH coupling constants have been shown to be sensitive to conformation. For example, CI'-H correlates with the glycosyl angle, 27,35 and in the aromatic rings, the value depends on the charge state of the ring [e.g., in cytosine (Cyt) N3H+]. ~7 One-bond coupling constants are straightforward to measure in F2-coupled HSQC spectra at natural abundance. Relaxation Times and Dynamics The NMR experiment relies on creating nonequilibrium spin states. There is a restoring force to return the system to equilibrium. In the absence of extensive dipole~lipole interactions, each spin relaxes exponentially toward the equilibrium state, and the time constant for this process is a relaxation time. Usually two different relaxation times are defined, which relate to the restoration of z magnetization (spin-lattice relaxation) or the Joss of phase coherence in the x-y plane (spin-spin relaxation). The relaxation rate constants R1, R2, and NOE together provide rich information about motions of vectors within the DNA molecule (or bound drug). For aliphatic carbon atoms and protons, the relaxation is dominated by the dipolar interactions, and R1 =

fl /r6[ j ( Ao)) -~- 3J(~ox) + 6J (cox + wH)]

R2 = fi/r6[4J(O) + J ( A ~ ) + 3J(cox) + 6 J ( w x + co~) + 6J(WH)] NOE = 1 + ~ / y x a / R j c~ = fl/r6[6J(cox + coil) -- J(Aco)]

(5) (6) (7a) (7b)

/3 is a constant that depends on the magnetic properties of the interacting nuclei, and has the value 56.92 for proton-proton relaxation if r is in angstroms (~) and the correlation time r is in nanoseconds. For X-H relaxation, the value of/3 is reduced by the ratio (VX/YH)2 SO forX = 13C,/3 = 3.56, forX = 15N,/3 = 0.56

35D. B. Davies,M. MacCross,and S. S. Danyluk,J. Chem. Soc. Chem. Commun. 536 (1984).

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and forX = 31p, ~ = 9.33. The spectral densities J(o9) are related to the motional properties of the X - H vector. A common formalism that accounts both for overall rotation of the molecule and for internal motions is 36 J(og) = S 2 j ( o ) , l'0) q- (1 - S2)j((.o, "re)

(8)

J((.o) = 1"/(1 + o921"2)

(9)

S 2 is the order parameter that describes the amplitude of the internal motion(s) and can vary between 1 (rigid body) and 0 (complete reorientational freedom). re is a correlation time that describes the time scale of the internal motions, and is usually poorly determined. 1-0 is the overall correlation time for rotation of the DNA duplex. As DNA duplexes are not spherical, this expression may need to be modified. 37 However, for short duplexes (about 10 bp), the spherical approximation is adequate, and measurement of all three relaxation rate constants [Eqs. (5)-(7)] allows all three parameters (1-0, S 2, and re) to be determined for each X-H vector. These relaxation rate constants are interpretable for heteronuclear (X-H) vectors, where the distance r and source of relaxation (i.e., the attached proton) are known. It is not sensible for IH relaxation, which in general will be multi exponential owing to a variety of relaxing nearby protons all at different distances. The definition of R j is then problematic. 3s However, the cross-relaxation rate constant for pairs of protons having a known and fixed separation can give useful information about overall rotation of the DNA duplex,37which can be compared with information obtained from 31p and 13C NMR relaxation. For 31p and aromatic 13C, chemical shift anisotropy (CSA) is also an effective relaxation mechanism, and the CSA must be added to the total relaxation. Relaxation by CSA contributes to R 1 and R 2 as RI(CSA) = (133.33 x 10 6)X2(.o2J(o)x) R2(CSA) = (22.22 x

lO-6)Z2w214J(O) + 3J(wx)]

(10) (11)

where X2 is the effective CSA.

31p relaxation in phosphodiesters is dominated by the CSA at magnetic field strength of 11.7 T and above because X ~ - 1 7 0 ppm and rap p ,~ 2.1 ]k. 39 Hence only RI and R 2 are useful parameters, which can be analyzed to determine both the magnitude of the CSA of each phosphodiester, as well as the local correlation time and order paramter. Generally, 31p NMR gives a good estimate of the global rotational correlation time of the nucleic acid. Changes in backbone dynamics and conformation from the CSA have been monitored both for groove binders 4° 36 G. Lipari and A. Szabo, J. Am. Chem. Soc. 104, 45469 (1982). 37 A. N. Lane, Methods Enzymol. 26l, 413 (1995). 38 A. N. Lane and T. Fulcher, J. Magn. Reson. B 107, 34 (1995). 39 M. J. Forster and A. N. Lane, Eur. Biophys. J. 18, 347 (1990). 40 A. N. Lane, T. C. Jenkins, T. Brown, and S. Neidle, Biochemistry 30, 1372 (1991).

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and intercalators. 4 13CNMR is better suited to measuring internal dynamics of the DNA, and how they are affected by drugs. 13C relaxation is dominated by the dipolar interaction with the attached proton for aliphatic carbons, and by both dipolar and C S A mechanisms for the aromatic carbons. 41 The latter therefore require multiple magnetic field strengths to separate the two mechanisms [CSA varies with the square of the field strength; cf. Eqs. (10) and (11)]. In the simplest analysis, the global correlation times [two for an anisotropic rotor such as double-stranded DNA (dsDNA)], an order parameter and an internal motion correlation time for each C - H vector can be measured. For the C2' atoms, the two vectors can interact (crosscorrelation), which can give rise to additional motional information. 41 43 Thus, it would be possible to measure changes in order parameters due to drug binding if the changes are sufficiently large. Such changes can be related to the contribution high-frequency motions make to the entropy change of binding. 44 Although there are now several 13C N M R studies of oligonucleotides that give a consistent picture of the internal dynamics, 3v'41,45 little has been done on d r u g - D N A complexes. This is clearly an area that is ripe for exploitation now that specifically labeled DNAs can be made. 46-4s 31p CSA and the global correlation time can be determined at 11.75 T or higher, using just R2 and R1 with proton decoupling. r = ( 1/2wp)[(6Rz/R1

) - 7] 0.5

CSA = [(R2 - 0 . 5 R 1 ) / ( 8 8 . 8 8 × 10-6)~O2pr]°5

(12) (13)

where r is in nanoseconds and CSA is in parts per million. This is correct for a rigid body showing no exchange (or other) contribution to R2. Rapid, low-amplitude fluctuations have little influence on the analysis, and it appears that in B-DNA the fluctuation of the CSA is colinear with the principal axis of the diffusion tensor, 39 and therefore does not contribute to the relaxation. However, exchange contributions greatly affect R2 (see above), and must be accounted for. As both fast intermediate exchange and CSA vary with the square of the field strength, B0 dependence is not always helpful. However, varying the temperature is useful, as for macromolecules the correlation time and therefore the relaxation rate constants should show a predictable monotonic dependence on temperature

41 C. Kojima, A. Ono, M. Kainosho, and T, L. James, J. MaSh. Reson. 135, 310 (1998). 42 V. A. Daragan and K. H. Mayo, Pros. Nucl. Magn. Reson. Spectrosc. 31, 61 (1997). 43 j. Engelke and H. Rtiterjans,J. Biomol. Nucl. Mash. Reson. 11, 165 (1998). 44 D. Yang and L. E. Kay,J. Mol. Biol. 263, 369 (1996). 45 E N. Borer, S. R. Laplante, A. Kumar, N. Zanatta, A. Martin, A, Hakkinen, and G. C. Levy, Biochemistry 33, 2441 (1994). 46 S. Tare, Y. Kubo, A. Ono, and M. Kainosho,J. Am. Chem. Soc. 117, 7277 (1995). 47 D. P. Zimmer and D. M. Crothers, Proc. Natl. Acad. Sci. U.S.A. 92, 3091 (1995). 48 j. M. Louis, R. G. Martin, G. M. CIore, and A. M. Gronenborn,J. Biol. Chem. 273, 2374 (1998).

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through the viscosity of the solvent. 49 Paramagnetic broadening also needs to be eliminated, necessitating careful preparation of the sample with exhaustive dialysis against chelating agents such as EDTA and ultra low-conductivity water (> 18 Mf2-cm). 13C relaxation measurements can be made at natural abundance, which apart from the low sensitivity has some advantages (e.g., n o 13C-13C coupling), and because proton detection can be used, sensitivity is no longer such an issue. Indeed, the line width can be obtained by HSQC if the indirect dimension is sufficiently well digitized, and the protons are rigorously decoupled during the tj evolution period, otherwise cross-correlation effects become severe. 5° The C8 resonance will have a line width of about 4 Hz at 3 nsec, so a reasonably accurate estimate of the line width from HSQC would need residual contributions from decoupling artifacts and digitization to be <0.4 Hz. With 13C enrichment, experiments are much more sensitive, and the additional information potentially available makes it likely that such experiments will become more frequent in the future, if significant changes are observed in drug binding. For small drugs, the effect of binding on the correlation times should be small unless it induces dimerization or aggregation of the DNA. However, large changes in structure would be expected to be reflected in changes in hydrodynamic properties. For example, substantial bending of DNA, for example, a 45 ° kink, will greatly affect its rotational diffusion constant, which can be measured accurately by ~3C NMR, and modeled using the bead formulation of Garcia de la Torre. 51'52 The rotational diffusion can also be compared with translational diffusion determined by the pulsed field-gradient spin-echo experiment. 53'54 This gives long-range information from the NMR experiment that is nol accessible from NOEs and coupling constants, and is generally not determined by traditional structure calculations. A different aspect of dynamics can be obtained by measuring the exchange rate of imino protons, which reports single-base pair fluctuations that lead to rupture of the Watson-Crick hydrogen bond. The fluctuation rate constant (opening) is on the order 500 sec -I, differing by about 3-fold between GC and AT base pairs, and varies also according to structure. The rate constant is much smaller in dA. dT tracts.55 A drug that perturbs the hydrogen bonding would be expected to affect the base pair-opening rate constant. It is important to ensure that the exchange kinetics faithfully reflect the opening kinetics, and not the chemical exchange reaction from the open state. This is best achieved with high concentrations of catalysts such as 49 A. J. Birchall and A. N. Lane, Eul: Biophys. J. 19, 73 (1990). 50 j. Boyd, U. Hommel, and I. D. Campbel, Chem. Phys. Lett. 175, 477 (1990). 5J M. M. Tirado and J. Garcia de la Torre, J. Chem. Phys. 71, 2581 (1979). 5~ M. M. Tirado and J. Garcia de la Torre, J. Chem. Phys. 73, 1986 (1980). 53 S. J. Gibbs and C. S. Johnson, Jr., J. Magn. Reson. 93, 395 (1991). 54 j. E. Tanner, J. Chem. Phys. 52, 2523 (1970). 55 M. GuEron and J.-L. Leroy, Methods" Enzymol. 261, 383 (1995).

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Tris or ammonia, at a pH above the pK value (about 8 for Tris, and about 9 for ammonium). As drugs usually stablize the duplex form over the strand states, the opening rate constant (or equilibrium constant) can be expected to be influenced by drug binding.

Cross-Relaxation: Nuclear Overhauser Effect From the point of view of conformational analysis, the NOE is probably the single most important parameter. Because of the strong (r -6) dependence of the cross-relaxation rate constant on distance, measurement of the NOE, or better its time dependence, can give quite stringent distance constraints between many pairs of protons. However, to interpret an NOE it is necessary to be able to account for all the contributions to the magnetization. A single-point NOE is just one sampling point on a complex time course, so the conversion to a distance is not straightforward. An absolute calibration of NOEs requires knowing also the rotational correlation time(s) of the system, the extent of internal motions and conformational averaging, the spin diffusion pathways, and in the case of complexes, the contribution of chemical exchange to relaxation processes. All these are at least in principle measurable, and there are algorithms that more or less allow the various problems to be treated, albeit with different degrees of rigor. 26"4°'56 Thus, if one wishes to know the subtle changes in conformation and dynamics due to ligand binding, a rather sophisticated analysis is required, and sufficient data must be obtained. The NOE also provides the most definitive evidence for the location of a binding site, as it involves direct contact between ligand and DNA protons. For a rigid ligand, a fairly small number of NOEs is sufficient to define its orientation within the binding site, and simple docking algorithms are adequate for mapping out the interactions. Where the ligand is flexible, and/or there are substantial conformational changes on forming the complex, many more NOEs (intraligand and ligand-DNA) are needed to define the interaction surface accurately. For a tight drug-DNA complex with a defined stoichiomet~% the treatment of the NOE spectroscopy (NOESY) spectrum is the same as for the free DNA, the only difference being that there are now additional NOEs (i.e., drug-DNA and drug-drug). For weak complexes, where the drug is in exchange between free and bound states, it is still possible to analyze the data, as in the transferred NOE experiment. 57'58 It is essential, however, to ensure that the DNA~lrug interactions (spin diffusion pathways) are properly accounted for as otherwise serious errors in structure can arise.

56 C, B. Post, R. P. Meadows, and D. G. Gorenstein, J. Am. Chem. Soc. 112, 6796 (1990). 57 F. Ni, Prog. Nucl. Magn. Resort. Spectrosc. 26, 517 (1994). 58 H. N. B. Moseley, E. B. Curto, and N. Rama Krishna, J. Magn. Reson. 108B, 243 (1995).

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For high sensitivity, long mixing times are appropriate for assignments, which can be aided by the substantial spin diffusion that always occurs in macromolecules. However, the term long is relative, and really needs to be defined with respect to rotational correlation times. This is because for macromolecules, the crossrelaxation rate constants ~r are directly proportional to the rotational correlation time of the molecule (or complex), as also are the intrinsic spin-lattice relaxation rate constants. The latter include all the cross-relaxation and leakage longitudinal relaxation processes, which compete with direct magnetization transfer between a pair of spins. In two-dimensional NOESY, the initial perturbation of the spin system creates a specific nonequilibrium state, which then relaxes back to equilibrium; the final state (at sufficiently long mixing times) is the equilibrium state, so that the energy input "diffuses" away from the source and is ultimately lost to the sample as heat. Thus there is a time at which the net magnetization is at a maximum (in magnitude). The time at which this maximum occurs is inversely proportional to the correlation time, and is therefore strongly affected by molecular size and temperature. Many structural protocols make use of distances, which are derived from the initial rate approximation, that is, NOE(t) = ~rtm. This can require short mixing times, where the signal-to-noise (S/N) ratio for all but the strongest NOEs is poor. It is generally preferable to record NOESY spectra at several mixing times, and analyze the time courses either directly against a model 4o or iteratively to the relaxation matrix and then the distances. 26'56 In both methods, a good choice of mixing time is essential, and at least three should be considered as the minimum. The intrinsic longitudinal relaxation rate constant determines the maximum NOE under otherwise perfect conditions, and corresponds to the initial rate of recovery of a selectively excited spin. Because protons are in magnetic contact with several other nearby spins, they do not relax exponentially, but in fact as a sum of exponentials (equal to the number of interacting spins). The initial rate of recovery of a nonselectively excited spin system is much slower than in the selective case, and the system as a whole tends to approach equilibrium rather slowly. However, as the NOE is proportional to the magnetization at the beginning of the experiment, an insufficient relaxation time reduces the initial magnetization and therefore the NOE intensity. Thus, for a spin system in which the effective Tl values all differ, either one must wait for complete thermal equilibrium to be reestablished after each pulse (many seconds), 27 or make the appropriate correction for the saturation factors. 38,59

Hydration For many DNA-ligand interactions, there is a change in the number of water molecules bound to the drug and the DNA on forming the complex, which is related 59S.-G. Kim and B. R. Reid,Biochemistry31, 12103 (1992).

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to the solvent-accessible surface area, SASA (see [6] in this volume59a). Thus it would be desirable to be able to monitor the changes in hydration of the DNA due to ligand binding. It is possible to examine the dipolar interaction between the protons of water and DNA. 6°-64 As the dipolar interaction depends both on the distance between protons and the local correlation time, it is possible in principle to detect not only the water contact site (i.e., from the actual NOE between water and DNA), but also to give an indication of the binding lifetime of the water molecules. To a first approximation, the lifetime can be assessed from the sign of the NOE. The NOE is negative for long correlation times (>0.3 nsec at 500 MHz) and positive for short correlation times (and therefore zero for a specific correlation time). The correlation time and the frequency at which the NOE is zero can be experimentally adjusted by using different spectrometer frequencies, for example, between 500 and 800 MHz, and by changing the temperature, which changes not only correlation times but also the chemical shift of water itself. This latter point is helpful where the water resonance overlaps solute protons (e.g., H3' of purine residues). Thus, if the sign of an NOE changes over a small change in spectrometer frequency or temperature range, then the effective local correlation time must be close to l/w. The local correlation time for the water-solute interaction is a function not only of the rotational correlation time of the DNA (a few nanoseconds), but also depends on internal motion of the water within the complex, and on the exchange rate with bulk water. This is limited by diffusion, which sets a lower limit to the lifetime of a "bound" water molecule; in the absence of any significant interaction with solute, the lifetime would be on the order 50 to 100 psec. If this were to determine the local magnetic correlation time, the NOE would be positive. Because the absolute magnitude of the NOE depends also on r -6, the NOE alone cannot be used to determine both r and r. However, although the cross-relaxation rate constant in the rotating frame also depends on 1"-6, it is determined by a different mix of spectra density functions, such that it is always positive; a negative peak in a rotating-frame overhauser enhancement spectroscopy (ROESY) experiment is either due to spin diffusion, or to chemical exchange. If the latter two contributions can be ruled out or avoided expeimentally, the ratio of the NOE to the ROE depends only on spectral density functions, and not on distances. or(L) = j ~ / r 6 1 6 J ( 2 o g ) - J(O)]

(14)

or(R) =- fl / r6[ 3 j (o)) + 2J(O)]

(15)

cr(L)/cr(R) = [6J(2~o) - J(O)]/[3J(~o) + 2J(O)]

59aI. Haq, B. Z. Chowdhry,and T. C. Jenkins, Methods Enzymol. 340, [6] 2001 (this volume). 60E. Liepinsh, G. Otting, and K. Wiithrich, Nucleic' Acids Res. 20, 6549 (1992). 61 E. Liepinsh, W. Leupin, and G. Otting, Nucleic Acids Res. 22, 2249 (1994). 62M. G. Kubinec and D. E. Wemmer,J. Am. Chem. Soc. 114, 8739 (1992). 63S. A. Fawthrop, J.-C. Yang, and J. Fisher, Nucleic Acids Res. 21, 4860 (1993). 64 A. N. Lane, T. C. Jenkins, and T. A. Frenkiel, Biochim. Biophys. Acta 1350, 205 (1997).

[121

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267

where a(L), a(R) are the cross-relaxation rate constants in the laboratory and rotating frames, respectively. For a rigid sphere,

J(w) = r/(1

+ o)2"t'2)

(16)

where r is the rotational correlation time. For large (macromolecular) correlation times, the cross-relaxation rate constant is dominated by J(0), and specifically a (L) and ~ (R) differ by a factor of 2. At short correlation times, both rate constants approach zero, and at some intermediate time, the NOE is zero [i.e., when J(2o)) = j(0)]. Thus the ratio r varies from - 0 . 5 to 1, with the zero crossing at r = 1.12/o). This corresponds to 0.36 nsec at 11.75 T, 0.3 nsec at 14.1 T, and 0.22 nsec at 18.8 T. Here, however, we are interested in the correlation time of the water-solute H-H vector, which requires a motional model. The simplest commonly used model is that the water-solute correlation time is dominated by the fluctuation of the water proton-solute proton vector because of translational and simultaneous rotational motion of the water, that is, exchange. 65 The correlation time then largely reflects the lifetime of the "bound" water molecule (sometimes called the residence time). The limits on determining residence times are between about 0.1 and 2 nsec. Residence times of less than about 0.5 nsec imply water that is not bound in a thermodynamic sense, as the effective dissociation constant is > 1 M (as the water concentration is about 55 M, this means >98% occupancy at Ka = 1 M). Typically, a relatively small number of water-DNA contacts is observed, some in the major groove and others in the minor groove. This is largely a technological problem related to resolution and the fact that there are comparatively few protons available to report interactions with water. It is not possible to measure an NOE between water and ring nitrogen or phosphate atoms because of substantial CSA relaxation. Equations (14)-(16) imply that the cross-relaxation rate constant is measured in both the laboratory and rotating frames. Usually, however, a single mixing times or perhaps two mixing times are used for each experiment, and the ratio of the peak intensities is taken as a measure of r. However, this requires that the spectra are also accurately scaled, which is difficult with independent experiments of different types. It is therefore convenient to use an internal reference. A convenient one is the cross-relaxation between Cyt H5 and Cyt H6, for which the distance is known and fixed. The ratio of the cross-peak intensity (actually area in cross-section through the water frequency) to that of the standard scales the desired ratio in Eq. (16) by exactly 2, because for the cytosine the correlation time is in the slow tumbling limit, and J(0) >> J(2o)). Taking the ratio also reduces some of the error associated with longitudinal relaxation, allowing longer mixing times to be used than would 65 G. Otting, E. Liepinsh, and K. Wtithrich, Science 254, 974 (1991).

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otherwise be the case. Because cross-relaxation in the rotating frame is up to twice as fast as in the laboratory frame, it is usual to use mixing times in that ratio (which would then give approximately equal intensities if other relaxation processes are the same). For moderate-sized oligonucleotides at 5 to 1@, where the rotational correlation time is < 10 nsec, mixing times of 50 msec (NOESY) and 25 msec (ROESY) seem to give good results. 64 Another major difficulty, and one that can seriously affect quantitative aspects of this kind of experiment, is that effective T1 relaxation is much faster than expected because of radiation damping, which becomes increasingly efficient as the magnetic field strength is increased. To avoid this problem in NOESY, a flip-back kind of experiment should be u s e d . 66 Once the cross-peaks have been identified and quantified, it is necessary to account for other artifacts that can influence the analysis. One is that there may be protons resonating at the same frequency as the water, and give rise to an ordinary NOE (e.g., H3' to H8). Although this is unlikely to be significant for C2'-endo sugars and short mixing times, the effect can often be ruled out by changing the temperature, as the water frequency is rather strongly temperature dependent compared with solute protons. A much more severe problem arises from exchangeable protons. A proton that exchanges with solvent during the mixing time gives rise to a cross-peak. In NOESY, this will be the same sign as the NOEs, although such peaks are usually also much broader. In ROESY, the exchange cross-peak is of opposite sign to direct ROE peaks (but in ROESY, spin diffusion changes the sign of the cross-peak). Thus exchanging protons can be readily identified. However, if a proton exchanges from, say, an NH2 group to water, and the NH2 is also close to a nonexchangeable proton, then there will be an indirect interaction between water and the target proton as far as magnetization transfer is concerned. Unfortunately, this exchangemediated transfer will have a normal ROE sign, and cannot be distinguished from a direct ROE except by careful mixing of time dependence studies and calculations based on known structures. Often potential water-solute cross-peaks are ignored if there is an exchangeable proton within 5 • of the target proton. This may be unnecessarily restrictive; if the exchange rate constant can be estimated from the exchange peak with the water, then the geometry and simple simulation of time courses can help discriminate between genuine direct NOE (ROE) and exchangemediated effects. These latter effects tend to become important in drug-DNA c o m p l e x e s . 66,67

Spectrometer Requirements For titrations, typically one needs about 0.5 ml at 0.25-0.5 mM DNA (and therefore > 5 - 1 0 mM ligand solution) in a buffer that does not contain protonated 66 G. Otting, Prog. Nucl. Magn. Reson. Spectrosc. 31, 259 (1998). 67 H. E. L. Williams and M. S. Searle, J. Mol. Biol. 290, 699 (1999).

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salts. Sodium phosphate plus KC1 is a good general salt system that buffers close to pH 7, and is essentially independent of temperature. At lower pH values, deuterated acetate can be used, but should not be lyophilized as acetic acid escapes. Titrations can be done in both H20 and D20 buffers. For He0, it is necessary to suppress the strong solvent signal; the pulsed gradient-based techniques such as Watergate 68 work well, and avoid the problems of saturation transfer that occurs with presaturation, as well as giving much better suppression. Titrations need to be carried out to greater than the stoichiometric point. For weakly soluble drugs, one can prepare a series of solutions of drug-DNA complexes by adding the DNA to the dilute drug (possibly in a nonaqueous solvent). The aromatic drugs are not always soluble in water without extensive aggregation. This creates a problem for binding and stoichiometry at NMR concentrations and also for determining the reference ligand shifts in the absence of DNA. For structural analysis, a 1 : 1 complex can usually be prepared by adding excess ligand, and resolving on a small desalting column. 4° This works only for tightly binding ligands. Solvent suppression techniques, and proton-detected X experiments require pulsed-field gradient capability (now common with modern spectrometers). The gradients are used for dephasing signals and coherence selection. HSQC experiments can easily be run at natural abundance (13C) at millimolar DNA-drug concentrations. Large z gradients (> 50 G/cm) can also be used for measuring the translation diffusion coefficient with the pulsed field gradient spin-echo method, 53'54 which can be valuable for investigating the hydrodynamic properties of the DNA and the DNA--drug complex (cf. bending). For this application, the gradient must be carefully calibrated using, for example, doped HOD and also against systems having known translation diffusion coefficients. The gradient linearity also needs to be checked; it may be necessary to restrict the length of the sample, using Shigemi tubes (we routinely use 16 to 18 mm for this purpose). In general, an inverse 5-mm probe can be used as the standard for modern spectrometers. If aggregation is a problem, more dilute solutions can be studied with minimal sensitivity cost, using 8-ram-diameter tubes (also available as Shigemi tubes, using about 1 ml). With a dilution of 3-fold, there is a small reduction in sensitivity, but may be sufficient to reduce dimerization to a manageable level. 69 The sample requirements are also reduced by using the increased sensitivity associated with 18.8-T spectrometers, which allow titrations to be carried out at 100 # M or less, and good quality two-dimensional spectra can be acquired at 25 # M with 8-mm tubes or a cryoprobe. For quantifying peak areas or volumes, it is essential to produce spectra that have fiat baselines or base planes, are free of ridges and spikes, and are adequately digitized. For measuring coupling constants from double quantum filtered correlation spectroscopy (DQF-COSY) or primitive 68 M. Piotto, V. Saudek, and V. Sklenar, J. Biomol. Struct. 2, 661 (1992). 69 J.-L. Asensio, T. Brown, and A. N. Lane, Struct,re 7, 1 (1999).

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[ 1 21

exclusive correlation spectroscopy (PE-COSY) spectra, digitization and resolution are important, as also is sensitivity. Poorly digitized, noisy spectra do not generally give reliable coupling constants or peak volumes. 25 For these purposes, high fields and long acquisition times are essential. In NOESY spectral peak volumes are also sensitive to saturation effects from rapid recycle times. Either long relaxation delays must be used (based on measured T1 values), or careful normalization of the peak volumes is required if fitting to the peak volumes is to be attempted. 59 Calculation Strategies The analysis of the three bond coupling constants will show whether the DNA and drug are predominantly to be found in a single conformer, or whether multiple state averaging has to be considered. If the former approach is chosen (and justified by the experimental data to hand), then the calculation strategy becomes very simple, and the same as for unliganded DNA. 26 Simulated annealing protocols and restrained molecular dynamics (rMD) with and without time averaged restraints (TARs) 26'7°'71 will be effective in refining the structure, from which many important conclusions can be derived about the nature of the interacting groups. A non-TAR structure can give misleading results about the extent of the fluctuations and how they are sampled compared with the free DNA, and therefore cause an important component of the overall affinity, and possibly specificity, to be missed. It is also clear that the free and ligand-bound structures and dynamics must be compared for any conclusions about affinity or stability to have any justification whatsoever. It is usually obvious where the binding site is from a cursory analysis of patterns of chemical shift perturbations and intermolecular NOEs (see below). The latter usually also indicate the preferred orientation of the drug in the binding pocket. However, other information may indicate that there are two possible orientations, or multiple overlapping binding sites, in which case the structure analysis becomes complicated; some way is needed to assign the NOEs to different possible states. Under these conditions, forcing a unique structure on the data could be misleading when relating it to specificity and affinity. There is no alternative to doing higher level structure calculations, using one of the ensemble methods or, at the very least, MD-TAR. The dynamics simulations also provide valuable information about flexibility of the system on the subnanosecond time scale, which can be compared with the experimental relaxation data. Order parameters extracted from 13C relaxation data can be compared with those calculated from the autocorrelation functions constructed from the dynamics trajectory. For preference, the system should be simulated in H20, as this generally gives superior results, and is in any case essential for calculating nanosecond dynamics of 70T. E. Torda, R. M. Scheek, and W. E van Gunsteren,Chem. Phys. Lett. 157, 2890 (1989). 71 U. Schmitz,N. B. Ulyanov,A. Kumar,and T. L. James,J. Mol. Biol. 234, 373 (1993).

[ 1 2]

NMR STUDIESOF DRUG-DNA COMPLEXES

271

/sNH 2 X H2N

X: -N=N-NH-

berenii

-O-(CH2)3-O-

propamidine

-O-(CH2)s-O"

pentamidine

FIG. 1. Chemical structure of bisamidine compounds.

the systems. Furthermore, it also allows one to analyze the positions and persistence of individual hydration sites, which can be compared with the (probably limited) information from the NMR data. Taken together, the structure, conformational flexibility, dynamics, and hydration data can be used for more detailed calculations of drug binding (such as SASA; see [6] in this volume59a), and for parsing the free energy relationships. Specific E x a m p l e s

Minor Groove Binding by Bisamidinium Compounds Members of the bisamidinium class of compounds (Fig. 1) are simple and cytotoxic. They bind to the minor groove of DNA with moderate affinity (l-10 # M range) and with a preference for AT-rich sites (Table I). 72 Extensive correlations of affinity and potency are available, and there are several X-ray and NMR structures of these drugs bound to DNA. 73-79 Titrations of the DNA with ligand always showed fast or fast intermediate exchange, implying moderate affinity (in agreement with footprinting and optical methods). Figure 2 shows NOESY spectra of the d(CGCAAATTTGCG)2 : propamidine complex in H20, and titration shifts for selected DNA resonances. The titration shifts (Fig. 2B) show that the binding site is in the AT-rich central region. Furthermore, the symmetry and extent of the shift profile suggest an extended, asymmetric

72 C. Zimmer and U. Wahnert, Ppvg. Biophys. Mol. Biol. 47, 31 (1986). 73 K. J. Edwards, T. C. Jenkins, and S. Neidle, Biochemistry 31, 7104 (1992). 74 Y. C+ Jenkins, D. G. Brown, S. Neidle, and A. N. Lane, Eur J. Biochem. 213, 1175 (1993). 75 T. C. Jenkins and A. N, Lane, Biochim. Biophys. Acta 1350, 189 (1997). 76 M. R. Conte, T. C. Jenkins, and A. N. Lane, Eur. J. Biochem. 229, 433 (1995). 77 S. Hu, K. Weisz, T. L. James, and R. H. Shafer, Eu~: J. Biochem. 204+ 31 (1992). 78 K. R. FOX,C. E. Sansom, and M. E G. Stevens, FEBSLett. 266, 150 (1990). 79 C. A. Laughton, T. C. Jenkins, K. R. Fox, and S. Neidle, Nucleic Acids Res. 18, 4479 (1990).

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TABLE I DRUG-DNAINTERACTIONS:NMR STUDIES Ligand

Footprint (bp)

Kd (298 K) (/zM)

Site (NMR)

Berenil

3.5

Propamidine Pentamidine Hoechst 33258

4 4.5 4-5

1-2 > I0 4 ~5 >5 0.01

AAT ATG TAA AAT AATT AAATT

4

2

YpG

11

2

Minor groove

lntercalator

Nogalamycin Major groove

Triplex 1l-mer

AGAAGAAAGGA

binding site such that the drug can bind in both orientations with essentially equal affinity, and that exchange between the binding modes is fast on the NMR shift time scale. The shift profile indicates an effective binding site of about 4 bp, in reasonable agreement with the DNase I footprint. As Table ! shows, the shift profile generally is a good indicator of the binding site size for this class of ligands. NOEs between the ligand protons and DNA protons (Fig. 2A) were exclusively to the minor groove (AdeC2H and H I ' plus some H4' and H5'/H5"), and to the bases expected from the shift profile. This confirmed the location of the binding site in the minor groove. The chemical shifts of the ligand also change on binding to the minor groove (Table II). In general, the downfield shifts increase with increasing site size, except for pentamidine, which can effectively bind only to the longest AT tract. Docking and energy minimization showed that the binding site was 3 base-pairs for berenil, and not symmetrically displaced with respect to the pseudodyad in AATT and AAATTT tracts. Similar results were obtained for berenil, propamidine, and pentamidine acting on AATT and AAATTT. The main differences were that berenil can occupy more than one site in the extended AT tracts. For propamidine, essentially all resonances were in fast exchange. The largest shift difference was about 0.5 ppm (250 Hz at 11.5 T), implying an exchange rate constant of at least 2000 sec -1. However, berenil binding showed intermediate exchange behavior. The line width is smallest in the free DNA, increased during the titration, and finally narrowed again on reaching saturation. The variation of the line width with drug concentration was analyzed according to the fast exchange equations: LW - LW ° = 47rpa(1 - p a ) A v 2 / k

(17)

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N M R STUDIES OF DRUG-DNA COMPLEXES

273

A PNH 2

P3,5

P 2,6

4,5" =

G

- r 7 H l ' / 91b • °

5.5-

~

e

Q

O

6.5 Q

7.5

o

1

,0*0

8.5 ¸ I , i 9 1 8.7

,

i 8.3

' I7'9I ' ' ' '7'5

7.1

6.7

6.3

chemical shift/ppm

B 0.4

B 0.2 E

-0.2

[]

[] []

-0.4

-0.6

C

G

HI' H2 H4'

C

A

A

A

T

T

T

G

C

G

FIG. 2. Interaction of d(CGCAAATTTGCG)2 with propamidine. (A) NOESY spectrum of the DNA-propamidine complex recorded in HeO at 277 K. P, ligand propamidine. (B) Titration shifts; &(complex) - S(DNA) for selected protons. [Reproduced with permission of Elsevier from A. N. Lane, T. C. Jenkins, and T. A. Frenkiel, Biochim. Biophys. Acta 1350, 205 (1997).1

where Pa is the m o l e fraction o f the unliganded D N A , A v is the frequency difference in hertz, determined from the free and fully saturated shift values, and k is the rate constant = kl[L] ÷ kl. As Kd was known, the on and off rate constants could be determined, koff = 500 sec -1, and kon = 5 × 108 M - I sec - l ,

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BIOPHYSICAL APPROACHES

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TABLE II SHIFTS OF BISAMIDINECOMPOUNDS BINDING TO DIFFERENT TARGETS IH chemical shift (ppm)

Ligand Berenil Propamidine

Pentamidine

Proton

Free

A2Ta

A2T2h

A3T3'

2, 6 3, 5 2, 6 3, 5

7.67 7.84 7.16 7.78 4.34 2.33 8.63 8.30 7.14 7.77 4.21 1.90 1.66

7.68 7.87 7.29 7.87 4.38 2.36 --------

7.68 7.90 7.42 8.00 4.48 2.41 8.78 8.63 7.31 8.05 4.47 1.93 1.70

7.97 8.18 7.52 8.10 4,56 2.48 8,95 8,85 7.21 7.95 4.33 1.87 1.61

NH2a NH2b 2, 6 3, 5 c~ fi y

a d(GCAATGAGCG),d(CGCTCATTGC). t, d(CGCGAATTCGCG)2. c d(CGCAAATTTGCG)2.

which is close to that expected for a diffusion limited reaction, indicating minimal reorganization of the initial d r u g - D N A complex (i.e., a preorganized site). For other ligands that bind less tightly, the dissociation rate constant was substantially larger, suggesting that the affinity for these ligands is determined by the off rate constant. The ligands showed only small effects on coupling constants, glycosyl torsion angles determined from NOE build-up curves, and 3tp N M R spectra, which is consistent with minimal perturbation of the D N A structure on forming the complexes, 74,75 and agrees with the binding kinetics (see above). This was confirmed by analyzing the structures of the DNA with and without ligand (Fig. 3); the structures were essentially identical within the experimental error and the limitations of the calculations. In the asymmetric D N A fragment, it was shown that there is some tolerance of a GC base pair by both berenil and propamidine, 76 which agrees with the observed low sequence selectivity (about 10-fold). 78,79 The structures all showed insignificant DNA distortion, and that the ligands make numerous good van der Waals contacts with the walls of the minor groove. The amidinium groups are able to make electrostatic interaction with the phosphodiester backbone. The larger ligands may bind their optimal target more weakly than berenil because the latter is preformed in the correct conformation, whereas the flexible

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NMR STUDIESOF DRUG-DNA COMPLEXES

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FIG.3. Solution structure of propamidine:d(CGCAAA'lTI'GCG)2.The structure was determined with rMD (Laneet al.64). The boundligand is knownin black, viewedend-onfrom the minorgroove.

linker in propamidine and pentamidine gives an unfavorable entropic contribution to binding. d(CGCAAATTTGCG)2 and d(CGCGAATTCGCG)2 both showed the expected water of hydration in the minor groove adjacent to the Ade residues; all AdeC2H showed negative NOEs to water. On binding propamidine, some water molecules were displaced, but others remain associated with the DNA, and probably also with the drug molecules. Thus, the amount of water displaced by drug binding is less than might be estimated simply by displacement of the minor groove hydration. Potential energy calculations give the correct position for the binding

276

BIOPHYSICALAPPROACHES

[ 12]

OH (HaC)2HN+~,.,.~

"r-o

o

Vy% # OH

O

OH

.~

0-1"-1-o4 .

H 3 C O ~

OCH3 C

G

A

1"

FIG. 4. Structureof nogalamycinand its complexwith DNA. Top:Nogalamycinstructure. Bottom: Structure of nogalamycinintercalated into the TpG/ApCsite. 67

site, and rank order the affinities for alternative sites. 75 A detailed parsing of the free energy has been carried out by Jenkins and co-workers (see [6] in this volume59a).

Intercalation by Nogalamycin The anthracycline antibiotic nogalamycin (Fig. 4) binds preferentially to YpG steps of DNA by intercalation, with the sugar in the minor groove. In contrast to the minor groove binders, the intercalators induce major local rearrangements and distortion of the DNA, which may be paid for by intrinsic binding energy; intercalators generally have rather low affinity for DNA. The structure of DNAnogalamycin complexes has been solved by both X-ray diffraction and NMR. The NMR spectrum is characterized by large changes due to the intercalation, in both 1H and 31E Indeed, the large (1-ppm) downfield shift of the YpG is characteristic of a B[-to-Bii transition. This is a way for the DNA to unwind, as intercalation

[12]

NMR STUDIESOF DRUG-DNA COMPLEXES

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requires separation of the two base pairs. The CSA of the affected phosphate increases, as one might expect as the symmetry and therefore bonding about the phosphorus atom are changed. However, the new conformation appears to be quite rigid by 31p relaxation.4 The structure of phosphodiester backbone necessitates a large change in the backbone torsion angles, which is primarily achieved by the B~-to-Bli transition. However, some other local backbone torsions alter to accommodate the intercalator. Thus, in the 1 : 1 complex with d(ATGCAT) • d(TACGTA), nogalamycin intercalated at the TpG/ApC step. The T moved primarily into the N domain of sugar conformations, whereas the other sugars remained mainly in the S domain. The CpA step, however, showed the transition to the BH conformation (e, ~" = - 6 0 , 180), with a concomitant change in G(A) to 180 ° (trans) and a compensating change in el(A) from g - to g+ (crankshaft motion). Other residues showed minimal changes in local conformations. These data show that the binding is notably asymmetric across the two affected base pairs, 67 reflecting the nonsymmetric structure of the drug, and the lifting of the degeneracy of the self-complementary DNA duplex. To accommodate the aromatic ring, the helical twist is now only about 20 °, compared with the usual 36 ° in B-DNA, and the helical rise is now 7 A, reflecting the sandwich of the aromatic intercalator adding about 3.4 ]k. In addition, other helical parameters for the 2 base pairs forming the intercalation site are altered, and even in restrained MD, showed substantial fluctuations about the average values. Larger fluctuations would be expected in a free dynamics trajectory or using the MD-TAR protocol. 26'7° Thus the local distortion of the DNA by the intercalator is severe, and must cost substantial potential energy. However, the destacking is presumably compensated for by making new van der Waals and electrostatic interactions with the intercalating ring system. Despite the severe local distortion imposed by the intercalator, the remainder of the molecule is normal B-DNA, and the effects do not propagate far along the double helix. This is in accord with many other observations of the inherent flexibility of the DNA backbone, that major distortions are rapidly damped out by the large number of degrees of freedom in the DNA duplex. 7'34 The hydration of the nogalamycin-DNA complex was also examined by NMR methods and simulation, 67 which could be compared with information from the X-ray structures. A remarkable feature is the incomplete agreement among the methods. Some of this is likely to be due to the inherent limitations of each method; NMR measures a time-dependent water residence and can detect only waters that persist for longer than about 100 psec close to a solute proton (i.e., no water associated with the phosphates can be detected by NMR), whereas X-ray and simulation detect mainly population averages. X-Ray crystallographic refinement methods and resolution have been shown to affect the estimate of water, and there are certainly limitations in the treatment of water potential in MD simulations. Nevertheless, significantly bound water molecules (residence times >0.5 nsec at 15°) were detected in the major groove, and indications of hydration of the bound

278

BIOPHYSICALAPPROACHES

[ 121

drug were obtained. The simulations showed some interesting correlations with the experimental data, especially regarding the hydration of the bound drug, and suggested that there may be water-mediated interactions between drug side chains and the DNA.

Major Groove Binding: Triplex-FormingOligonucleotides In addition to minor groove recognition and intercalation, it is possible to recognize the major groove with oligonucleotides, making use of Hoogsteen hydrogen bonding to a polypurine stretch. The most promising avenue in this area of antigene appears to be the parallel motif, despite the need to protonate the cytosine at N3. High affinity and specificity have been achieved with modified bases, such as including a positive charge on the base, and adding propanyl chains at the 5 position. To date the only full and detailed conformational information about DNA triplexes has come from NMR spectroscopy. There are now several structures of the parallel motif, and the basic observations are the same; the minor groove of the underlying duplex is compressed, and the helix axis is displaced about 2 A. The sugars in the Hoogsteen strand, especially the protonated cytosines, appear to be less S state than usual, consistent with the notion that the most stable triplex is one in which the Hoogsteen oligonucleotide has C3'-endo sugars. 5'8°'81 This is shown by the relative stability of triplexes containing a DNA third strand and an RNA third strand. 8° 82 The presence of the positive charge on the protonated cytosine residues was verified by 13C NMR spectroscopy, which showed that both the chemical shifts and the one-bond C H coupling constant (C5 and C6) reflect the different electron density of CH + and C (Fig. 5). 17'69 The charge also appears to alter the local conformation of the purine residues, and was associated with a transition between y(g+) and y(t) of the purine nucleotide 5' to the CH + residue. The value of y was determined from a combination of coupling constants at H4' and NOEs and unusual shifts for H3' and sequential NOE. In the usual g+ rotamer, the H4' as a small coupling to H3' (S sugar) and two equal, small coupling to H5' and H5". The sum of the coupling is less than 12 Hz. However, for the t conformer, one of the couplings is large (12 Hz) and the other is small, leading to a splitting of the H4' resonance. This can be seen readily in cross-section of an NOESY through the H3'-H4', with high digital resolution along F2. In addition, in the DQF-COSY, the appearance of a strong H4'-H5'/H5" cross-peak and of NOEs between H4', and H5'/H5" also proves the presence of g - or t (Fig. 6). The NOEs in turn allow a stereospecific assignment to be made. In all cases that we have examined, the presence of y (t) is accompained by an unusual up field shift of H3', which shows an NOE to so R. F. Macaya, P. Schultze, and J. Feigon, J. Ant, Chem. Soc. 114, 781 (1992). Sl J.-L. Asensio, R. Carr, T. Brown, and A. N. Lane, J. Am. Chem. Soc. 121, 11063 (1999). 82 H. Han and E B. Dervan, Proc. Natl. Acad. Sci, U.S.A. 90, 3806 (1993).

[12]

NMR STUDIES OF DRUG-DNA COMPLEXES

279

H6/H8 E

136_

G3 o..4-qp~o Gqo ~ o

Q.

140_ = 0

•~

144 _

61 0

148-

"¢: 152 O (,3 e~ 156 -

C8

~ C9+C10

C6

~ C15+C16

o

otO~

od AC2

83 7:8

C2

7;5 7.~ 6~9

1H Chemical Shift (ppm)

FIG. 5. HSQC spectrum CH +. 13C-IH HSQC spectrum of d(AAGGAA) - d(TTCCTT), d(TTC + C+TT) recorded at 14.1 T and 30 °, showing the C8, C6 and C2 regions. The protonated carbons (15 and 16) have significantly different shifts and nJcH from C9 and C10. ]Reproduced with permission from J.-L. Asensio, T. Brown, and A. N. Lane, Nuclei(' Acids Res. 26, 3677 (1998). Oxford University Press.]

H4' HS' HS"

~-81

_

~

= r

o

H4' HS' HS"

381

~J

oo

& .

~

HS' HS"

46t

48:

H3'

" r~ 5 . 8 5.96.0" G.1-

• 4.5S

. GS H4' O.Z4'<~ Hz 0

'l

(~

6~

. 4.45

,

. 4.35

,

. 4.25

Chemical Shift (ppm)

~ < I ~

A3 H4'

.

G2 H4' Z4'<10 HZ

~

HI'

~4.4 e.-T~-]

G.3-

w

, 4.50

,

n 4.40

,

n 4.30

i

I

4.20

Chemical Shift (ppm)

FIG. 6. Determination of the backbone torsion ?" in the DNA triple helix d(AGAAGA).d (TCTTCT) - d(TC+TTC+T). Top left: H4t-H3 ' region of an NOESY spectrum. Top right: H3'-H4' region of a DQF-COSY spectrum. Bottom: H 4 ' - H I ' region of the NOESY spectrum. The A3 and A4 H4' shifts, splitting, and interactions with H5'/H5 I1 indicate/(t) for these residues, whereas all others are 7(g+). [Reproduced with permission of Oxford University Press, from J.-L. Asensio, T. Brown, and A. N. Lane, Nucleic Acids Res. 26, 3677 (1998).]

280

BIOPHYSICALAPPROACHES

[ 12]

the preceding nucleotide. The t conformer requires a compensating change in c~, which is always found during the structure calculations if the phosphate backbone torsion angles are not restrained. The presence of F (t) alters the course of the phosphodiester backbone, so that the major groove of the Waston-Crick duplex is no longer divided into two grooves in the fixed ratio 2 : 1, but into a continuously varying ratio. 69,81 This has implications for the binding of drugs to either of these grooves. The local structures of the various triplexes that have been examined do provide a plausible mechanical explanation for some of the measured thermodynamic properties. However, there is much less dynamics information and solvation 83 than is available for DNA duplexes. Most of this kind of information has been obtained from simulations s4 However, to date the simulations have not reproduced all the conformational features of triplexes that have been detected by NMR experiments. Future Developments Increasing magnetic field strength affords greater spectral dispersion and sensitivity, allowing the study of more complex systems, or current systems at much higher dilution (as well as with improved resolution). An additional benefit of ultrahigh field strength is magnetic ordering. Because nucleic acids contain linear stacks of aromatic rings, which are themselves strongly magnetically anisotropic, long polymers have a high degree of anisotropy in the magnetic susceptibility, and therefore orient in a magnetic field. As the degree of orientation is proportional to the square of the field strength, 85 the newest magnets (at 18.8 T and higher) induce significant ordering, and measurable dipolar splittings in 13C and IH.86 The dipolar splitting is proportional to the gyromagnetic ratio divided by r 3, so for a given orientation, the relative splittings are 1 for N-H (r = 1.02 2k), 2.05 for C-H (r = 1.09 A), and 0.72 for H-H at 2.45 ]k. In the ordered state the dipolar couplings do not average to zero, so the residual dipolar coupling can provide information about the orientation of the particular dipole vector relative to the principal axis of the susceptibility tensor. In DNA this axis is expected to be essentially colinear with this helix axis, and for duplexes and quadriplexes the symmetry should be axial. The residual dipolar coupling gives a straightforward way of determining long-range features, such as bending, that cannot be accurately measured from NOEs and scalar coupling constants.

83 I. Radhakrishnan and D. J. Patel, Biochemistry 33, 11405 (1994). 84 G. C. Shields, C. A. Laughton, and M. Orozco, J. Am. Chem. Soc. 119, 7463 (1997). 85 j. R. Tolman, J. M. Flanagan, M. A. Kennedy, and J. H. Prestegard, Proe. Natl. Acad. Sci. U.S.A. 92, 9279 (1995). 86 H. C. Kung, K, Y. Wang, I. Goljer, and P. H. Bolton, J. Magn. Reson. 109B, 323 (1995).

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Isotopic labeling has long been fairly straightforward for RNA using in vitro transcription, 87'88 although with limitations on selective labeling. There are now chemical and biochemical methods for labeling DNA 46-48 that will greatly improve resolution and open up the possibility of detailed dynamic analysis. 41'89 Accurate determination of the rotational diffusion tensor, especially in conjuction with dipolar coupling, will allow gross morphological drug-induced changes to be monitored, as well as accurate change in order parameters for a wide range of vectors, which can be converted into changes in entropy.44 It has been shown that the hydrogen bonds in nucleic acids have a substantial covalent character, which is manifested by the surprisingly large scalar coupling between donor and acceptor nitrogen of2J = 6 to 8 H z . 90'91 These couplings have been observed in both DNA triplexes and quadriplexes, and provide unambiguous and direct identification of the donor and acceptor atoms in hydrogen bonds. 90'91 This is easy to measure by HSQC, and shows which nitrogen atoms are involved in a hydrogen bond. In principle, drug-DNA interactions could exploit this, and even intermolecular hydrogen bonding could be observed. Improved hardware and methodologies will allow closer ties to be made to other experimental data, such as from thermodynamics and kinetics, via the intermediary simulation approach. The possibilities for improved drug design based on this kind of interaction seem especially propitious. Acknowledgments This work was supported by the Neal Radnew Trust and the Medical Research Council of the U.K. I thank my colleagues Dr. Terry Jenkins and Dr. Tom Frenkiel for helpful discussions.

87 R. T. Batey, M. lnada, E. Jujawinski, J. D. Puglisi, and J. R. Williamson, Nucleic Acids Res. 20, 4515 (1992). 88 E. P. Nikonowicz, A. Sirr, P. Legault, E M. Jucker, L. M. Baer, and A. Pardi, Nucleic Acids Res. 20, 4507 (1992). 89 E Gaudin, E Paquet, L. Chanteloup, J.-M. Beau, N. T. Thuong, and G. Lancelot, J. Biomol. Nucl. Magn. Resort. 5, 49 (1995). 9o A. J. Dingley, J. E. Masse, R. D. Peterson, M. Barfield, J. Feigon, and S. J. Grzesiek, J. Am. Chem. Soc. 121, 6019 (1999). 91 K. Pervushin, A. Ono, C. Fernandez, T. Szyperski, M. Kainosho, and K. Wiithrich, Proc. Natl. Acad. Sci. U.S.A. 95, 14157 (1998).

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[13] X-Ray Crystallography of DNA-Drug Complexes By MARY ELIZABETHPEEK and LORENDEAN WILLIAMS Introduction Macromolecular X-ray crystallography is the quintessential method for determining three-dimensional structures of biological macromolecules to atomic resolution. Several textbooks and reviews provide excellent practical and experimental treatments of protein crystallography.l-3 DNA crystallography has the same fundamental theoretical and experimental underpinnings as protein crystallography. As in a protein crystallography experiment, DNA crystallography is performed by the following steps: (1) crystal growth; (2) X-ray diffraction data collection and reduction; (3) phase determination by molecular replacement, multiple isomorphous replacement (MIR), or multiple wavelength anomalous diffraction (MAD); and (4) refinement. However, DNA crystallography differs somewhat from protein crystallography in methods of crystal growth, data collection and reduction, and phase determination. Those distinctions are the primary focus of this chapter. Crystal Growth To initiate crystal growth, a solution is slowly brought to supersaturation. 4 Small aggregates act as nuclei, allowing crystal growth to commence. One impediment to growth of high-quality crystals is the prescriptive difference in optimum solution conditions for nucleation and for growth. Optimum crystallization conditions favor conformationally stable and homogeneous populations and promote specific intermolecular interactions. DNA crystallization, in our view, is best understood by analogy with DNA condensation into toroids. DNA toroids are less ordered than crystals but more ordered than aggregates. The conditions that promote nucleation and growth of crystals can be anticipated by those required for nucleation and growth of toroids, with the caveat that condensation involves DNA polymers whereas crystallization involves DNA oligonucleotides. As reviewed by Bloomfield,5 DNA condenses in the presence of polyamines such as spermine and spermidine, or trivalent inorganic cations I D. E. McRee, "Practical Protein Crystallography."Academic Press, New York, 1999. 2 j. Drenth, "Principles of Protein X-Ray Crystallography." Springer-Verlag, New York (1999). 3 C. W. Carter, Jr. and R. M. Sweet, eds., Methods Enzymol. 276 and 277 (1997). 4 A. McPherson, "Preparation and Analysis of Protein Crystals." Robert E. Krieger Publishing, Malabar, Florida, 1989. 5 V. A. Bloomfield, Biopolymers 44, 269 (1997).

METHODS IN ENZYMOLOGY,VOL 340

Copyright© 2001 by AcademicPress All rightsof reproductionin any form reserved. 0076-6879]00$35.00

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X - R A Y CRYSTALLOGRAPHY OF D R U G - D N A COMPLEXES

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such as cobalt hexaamine. DNA also condenses in the presence of both alcohols and divalent cations. The majority of DNA structures contained within the Nucleic Acid Structural Database (NDB) 6 were obtained from crystals grown from solution conditions favorable to DNA condensation. To find solution conditions favorable for crystal growth, a multiparameter search must be performed. As described in a previous review of DNA crystallography, 7 the parameters to be varied in a search for DNA crystallization conditions include (1) metal ions and polyamines, (2) type and concentration of DNA ligands, (3) alcohol and buffer, (4) pH, (5) temperature, and (6) "precipitating agents." In our laboratory, pH, temperature, and precipitating agents are not generally early search parameters for growing DNA or DNA-drug crystals. Differences between DNA and protein crystal growth arise from the polyanionic nature of DNA and the dependence of DNA conformation and stability on cations, as opposed to a less ionic character of most proteins. Cations are the first and generally most important parameter varied during searches for DNA or DNAdrug crystallization conditions in our laboratory. Our initial screen is invariably a spermine-versus-magnesium grid, with all other components held fixed. The initial ratio of DNA to ligand is fixed at 1.1 tool of ligand per 1.0 mol of DNA-binding site. Once magnesium and spermine concentrations are optimized, a fine search of the DNA-ligand ratio is performed. In addition to cations and polyamines, certain alcohols and buffers also appear to be important for crystallization of DNA and DNA-drug complexes. The NDB presently contains 402 DNA crystal structures that lack protein. Slightly more than one-half (208) of those crystals were grown from solutions containing both methylpentanediol (MPD) and Mg 2+. By contrast, few DNA-protein crystals, only 13 of 346 structures contained in the NDB, have been obtained from solutions containing MPD. Of DNA crystal structures that lack protein, 65% were obtained from solutions containing cacodylate buffer. Only 7% of DNA-protein crystals were obtained from solutions containing cacodylate. pH plays a different role in nucleic acid crystallization than in protein crystallization. It is not feasible to crystallize DNA under conditions near the isoelectric point. Unlike proteins, DNA does not contain functional groups that change ionization state around physiological pH. Important exceptions to this rule are encountered when cytosine is protonated within non-Watson-Crick base-pairing schemes, such as hemiprotonated C-C base pairs, 8'9 and Hoogsteen base pairs, l° 6 H. M. Berman, C. Zardecki, and J. Westbrook, Acta Crystallogr. Sect. D 54, 1095 (1998). 7 A. H.-J. Wang and Y.-G. GaG, Methods 1, 91 (1990). 8 C. H. Kang, I. Berger, C. Lockshin, R. Rafliff, R. Moyzis, and A. Rich, Proc. Natl. Acad. Sci. U.S.A. 91, t 1636 (1994). 9 L. Chen, L. Cai, X. Zhang, and A. Rich, Biochemistry 33, 13540 (1994). l0 G. J. Quigley, G. Ughetto, G. A. van der Marel, J. H. van Boom, A. H.-J. Wang, and A. Rich, Science 232, 1255 (1986).

284

BIOPHYSICALAPPROACHES

[ 131

and when DNA ligands contain functional groups that change ionization state near physiological pH. It is not possible to unambiguously eliminate tradition or other biases in determining the origins of differential patterns in protein and DNA crystallization conditions. However, protein crystallization conditions in general, even DNA-protein crystallization conditions, can provide poor models for DNA crystallization. X-Ray Data Collection and Reduction The ultimate goal of the X-ray diffraction experiment is to determine the electron density, p (x, y, z), for each atom in the macromolecule according to Eq. (1).

p(x, y , z ) = ( l / V ) E l F ( h k l ) l e x p [ - 2 a v i ( h x

+ky+lz)+iethkl]

(1)

hkl

where V is the unit cell volume; ]F(hkl)] is the structure factor amplitude; hkl is the Miller index; x, y, and z are the real space coordinates; and e~h,l is the relative phase of reflection hkl. In an X-ray diffraction experiment, one measures intensities, [l(hkl)], of many thousands of "reflections" using a charge-coupled device (CCD) camera or an imaging plate. Intensities are convened to structure factor amplitudes []F(hkl)[] by l (hkl) = ]F(hkl)] 2.

Dynamic Range Success in the solution of a structure by MIR or MAD, and the accuracy of a final refined model, are critically dependent on the accuracy of IF(hkl)l. It is inherently more difficult to accurately determine [F(hkl)l from a DNA crystal than from a protein crystal because the dynamic range in IF(hkl)l from DNA crystals can be significantly greater than from protein crystals. Moreover, certain packing arrangements within DNA crystals cause greater dynamic range than other packing arrangements. Specifically the dynamic range of IF(hkl)L is greater for DNA crystals with end-to-end stacking (i.e., with pseudo-infinite helical axis) than for other packing arrangements such as end-to-groove packing (i.e., with skewed helical axes). This trend in dynamic range is illustrated in Table 111-13 and Fig. 1, where [F(hkl)] from three crystals of the same quality (1.4-]k resolution) are compared. DNA with a pseudo-infinite helical axis is compared with DNA with skewed helical axes, and with a globular protein (containing c~ helix plus [3 sheet). To quantitate dynamic range, the reflections of these data sets were independently sorted by I I G. A. Leonard, T. Brown, and W. N. Hunter, Eur. J. Biochem. 204, 69 (1992). 12 X. Shui, L. McFail-Isom, G. G. Hu, and L. D. Williams, Biochemistry 37, 8341 (1998). 13 G. Xiao, M. Tordova, J. Jagadeesh, A. C. Drohat, J. T. Stivers, and G. L. Gilliland, Proteins 35, 13 (1999).

[l 3]

X - R A Y CRYSTALLOGRAPHY OF D R U G - D N A COMPLEXES

285

TABLE I DYNAMIC RANGE IN X-RAY INTENSITY DATA

Crystal

Identifier

Resolution (~,)

Space group

TGTACA-4'epiadriamycin

NDB entry DDF035

1.4

P41212

CGCGAAT TCGCG Escherichia coli uracil DNA glycosylase

NDB entry BD1084 PDB entry 4eug

1.4

P212i2]

1.4

P21212]

Characterization Intercalated B-DNA, pseudo-infinite helical axis B-DNA, skewed helical axes a helix and [3 sheet

Fmed(hkl)/ Frn~x(hkl)

Ref.

0.38

11

0.17

12

0.13

13

amplitude and F(hkl), and the median and maximum amplitudes (IF(hkl)lmed and IF(hkl)lm,x) were identified. The ratio IF(hkl)lmed/IF(hkl)lmax varies with dynamic range. As shown in Table I. IF(hkl)lmed/IF(hkl)lm,× for the pseudoinfinite helical axis DNA is more than two times greater than that for the skewed helical axes DNA, and nearly three times greater than that for the globular protein. This trend is illustrated in detail in Fig. 1. If weak IF(hkl)l is defined as those that are less than IF(hkl)[ re.x/20, then ~30% are weak in a globular protein, --~42% are

80-

i/pseudo_infinite i helical axis DNA

60

~-40-

~i

~ skewed he cal 20-

0

! \ ~,N~/protein 0

-

2'0 4'0 IF(hkl)l/IF(hk/)lmax(%)

-

6'0

FIG. 1. Dynamic range of X-ray diffraction data from crystals of a DNA complex (TGTACA-4'epiadriamycin) with a pseudo-infinite axis, a DNA fragment d(CGCGAATTCGCG) with a skewed helical axes, and a globular protein (Escherichia coli uracil DNA glycosylase). The graph shows plots of frequency of observation versus amplitude, normalized to the greatest amplitude in that data set. To obtain frequency of observation, IF(hkl)l were sorted into bins of IF(hkl)l > O.051F(hkl)lmax, O.051F(hkl)lmax > IF(hkl)l > O.lOIF(hkl)lmax, O. lOIF(hkl)lmax > IF(hkl)l > O.151F(hkl)[max, etc.

286

BIOPHYSICAL APPROACHES

b

[ 13]

"

1016

\ 4~

- elongat~

S

FIG. 2. Precession photograph of a crystal of the DNA-porphyrin complex CGATCG-CuTMPyP4 [Cu(II)meso-(4-N-tetramethylpyridyl)porphyrin,NDB entry DDF060].I6 This 6 ° screened precession photograph, taken with Ni-filtered Cu radiation, recorded the hOI layer. In this crystal (a = 39.49 ~, c = 56.15 ~, c~ = 90 °, ~ = 120, space group P 6122) the pseudo-helical axis is nearly parallel to the crystallographic c axis, causing extremely large values of IF(1 0 16)1. Anisotropy in crystalline order causes the diffraction pattern to fade out at a lower angle along the a* axis than along the c* axis. The elongated cross is caused by rotational disorder about the helical axis. The 6-fold screw axis along the c axis is indicated by systematic absences along the c* axis.

weak in the skewed helical axis DNA, and ~80% are weak in the pseudo-infinite helical axis DNA. The effects of a pseudo-infinite helical axis of DNA on the diffraction pattern can been observed directly in the precession photograph shown in Fig. 2. In this case (a DNA-porphyrin complex), the pseudo-infinite helical axis is directed nearly

[13]

X-RAY CRYSTALLOGRAPHYOF DRUG-DNA COMPLEXES

li

"

0

d,

I

.........

0.0 0.5

287

X

7.2 Al [3.6A t -0.5

FIG. 3. Patterson map from a crystal of duplex [d(CGTACG)]2 bound to the bisintercalator D232 (a = 28.24 A, b = 28.24 ,&, c = 72.74 ~, e~ = 90.0,13 = 90.0 ~, 3' = 120.0~) .14 The pseudo-infinite helical axis is parallel to the crystallographic c axis.

along the crystallographic c axis. Extremely intense reflections with Miller indices 1 0 16, 1 0 -16, - 1 0 16, and - 1 0 - 1 6 are observable in the photograph. The origin of differences in dynamic ranges of IF(hkl)l from globular proteins and DNA crystals can be understood in part from a Patterson analysis. In a globular protein crystal, interatomic vectors generally have random lengths, directions, and origins. By contrast, a B-DNA or intercalated B-DNA crystal will yield a large number of"stacking vectors" with conserved lengths ('--3.4 A, 6.8 ,~, 10.2 A, etc.), directions (along the helical axis), and semiconserved origins (from the planes of base pairs). When the B-DNA is organized in an end-to-end fashion in the crystal (with a pseudo-infinite helical axis), then all the stacking vectors are nearly aligned. Therefore a DNA Patterson map can contain intense peaks spaced by 43.4 A. A Patterson map from a bisintercalated DNA crystal (NDB entry DD0018)14 with a pseudo-infinite helical axis is shown in Fig. 3. A Fourier decomposition of the electron density within a B-DNA or intercalated DNA crystal gives large amplitude waves with wavelengths of ~3.4 ~, ~6.8 ]k, ~ 10.2 ~, etc. For example, F(hkl)m.,~× in the data set collected by Hunter and co-workers 1~ on TGTACA-4'-epiadriamycin (Table I) has the Miller index 0 0 16. Dividing the length of the crystallographic c axis (52.39 ~) by l = 16 gives a d spacing for this reflection of 3.27 A. The Miller plane normal is parallel to t4 X. Shui, M. E. Peek, L. A. Lipscomb, Q. Gao, C. Ogata, B. R Roques, C. Garbay-Jaureguiberry, and L. D. Williams, Curl: Med. Chem. 7, 59 (2000).

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the c* axis. The spacing and direction of this intense reflection are consistent with base-base and base-intercalator stacking, and alignment of the helical axis along the crystallographic c axis. The second and third ranked F(hkl) in this data set have Miller indices l 1 16 and 0 2 16 (3.25-A d spacing with Miller plane normals within 15° of the c axis). Thus the Fourier decomposition of the electron density in this crystal contains three stacking waves of similar wavelength and direction. Those stacking waves have amplitudes nearly 40-fold greater than the median intensity in the data set and dominate the diffraction pattern. Their positions are tightly clustered in reciprocal space.

Data Collection Strategy One of the goals of most diffraction experiments is to collect data to the highest resolution possible. The highest resolution data have the lowest intensities, requiring long collection times, intense X-ray beams, and sensitive detectors. It is often impossible to collect weak high-resolution data simultaneously with strong stacking reflections. One part of the strategy employed in our laboratory is to collect stacking intensities independently from high-resolution data by collecting multiple data sets on the same crystal. The stacking intensities are collected with short exposures, low amperage on a rotating anode, or an attenuated the beam at a synchrotron. The short- and long-exposure data sets are scaled and merged. It is best to exclude the stacking reflections from the scaling, for example, by merging data from 20 to 2.6 A (short exposure) with data from 3.1 to 1.4 ]k (long exposure). In this case the overlap, used for scaling, is 3.1 to 2.6/~ and would not contain the ~3.4-2k stacking reflections. A second part of our strategy is to identify the stacking reflections prior to initiating data collection, and to set the collection parameters to ensure accurate measurement of their intensities. Detector overload and peak overlap must be avoided by empirical adjustment of exposure time and crystalto-detector distance. The stacking reflections are in close proximity in reciprocal space, and are generally broader than other reflections.

Crystallographic Anisotropy Elongated flexible molecules that lack lateral structural hooks exhibit characteristic types of crystalline disorder. In the precession photograph in Fig. 2, the diffraction pattern of a DNA-porphyrin complex is seen to extend out to the edge of the photograph along c*, but to fade out at lower resolution along a*. Thus the diffraction pattern is anisotropic, indicating that the disorder within the crystal is anisotropic. The DNA is more highly ordered along the helical axis than along the perpendicular to the helical axis. These systematic errors in the data can be attenuated, once the refinement is near completion, by anisotropic or local scaling

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of observed-to-calculated data. One cost of these corrections is that they naturally introduce additional parameters to the refinement. Additional information about disorder within the DNA-porphyrin crystal is provided by the faint elongated cross pattern, stretching between the intense stacking reflections (Fig. 2, "X"). This elongated cross is reminiscent of a DNA fiber diffraction pattern. Our interpretation of the origin of the elongated cross in Fig. 2 is that a fraction of the DNA within the crystal is disordered by random rotation about the helical axis, just as in a fiber. Structure Determination

Molecular Replacement Determination of helical axis orientation by Patterson analysis can be combined with symmetry information to simplify structure solution by molecular replacement. For example, the Patterson map and strong 0 1 16 reflection indicate that the helical axis of the CGTACG-porphyrin complex is nearly parallel to the c axis. Simple volume calculations and spectroscopic analysis of a dissolved crystal indicate that the asymmetric unit contains one strand of DNA plus one porphyrin molecule. If the DNA forms a duplex, then the duplex must be centered on a crystallographic 2-fold axis. The DNA-porphyrin complex must be centered on one of the two crystallographic 2-fold axes (in space group p61(5)22). Two possible orientations, which differ by 180 °, are possible on each 2-fold axis. Thus the molecular replacement search is limited to four one-dimensional translations. However, the success of molecular replacement always depends on an accurate search model. The CGTACG-porphyrin structure contains an unanticipated flipped-out base. Therefore molecular replacement failed in this case, even though location and orientation of the complex were correctly anticipated.

Multiple Isomorphous Replacement and Multiple Wavelength Anomalous Diffraction Structure determination by MIR or MAD requires substitution with heavy atoms. Derivatives can be generated by de novo crystal growth of modified DNA. Cytosine and uracil can be substituted with bromine or iodine at the C5 position.14 16 A search of the NDB indicates 30 bromine-substituted DNA fragments have been crystallized. Guanines are effective targets for soaking, for example, by accepting platinum at the N7 position. ~6 J5C. L. Kielkopf,K. E. Erkkila, B. E Hudson,J. K. Barton, and D. C. Rees,Nat. Struct.Biol. 7, 117 (20OO). 16L. A. Lipscomb,E X. Zhou,S. R. Presnell, R. J. Woo,M. E. Peek,R. R. Plaskon,and L. D. Williams, Biochemistry35, 2818 (1996).

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Summary Here we have stressed important differences between protein and DNA crystallography. Crystal growth and data collection methodologies are not directly transferable between the two subfields. In addition, we note that analysis of symmetry and packing of DNA crystals can be useful and a uniquely aesthetic exercise. Acknowledgments This work was supported by the National Science Foundation (Grant M C B - 9 9 7 6 4 9 8 ) and the A m e r i c a n C a n c e r Society (Grant R P G - 9 5 - 1 1 6 - 0 3 - G M C ) .

[14] Molecular Modeling of Drug-DNA Complexes: An Update By J O H N

O. TRENT

Introduction This is an exciting time in the field of molecular modeling of DNA and DNA :ligand structures. There have been several advances in the field, such as second-generation force fields and the implementation of the particle mesh Ewald (PME) methods that more accurately reproduce experimental structures. When combined with the ever-increasing speed of computational resources (supercomputers and, in particular, parallel processor workstations), this places molecular simulations of biologically relevant systems at the fingertips of researchers. We now are able to investigate systems that previously were not practical, either because of less reliable force fields or computational expense. This is not to say that the discipline of molecular modeling has fully matured, as we are only at the second generation of force fields. However, with the increasing reliability, physical size of simulated DNA and proteins, and length of simulations, more researchers are turning toward simulations to aid them in the rationalization and prediction of experimental data, and in the design of specific experiments. The application of molecular modeling (and structural biology as a whole) to nucleic acids is, and will continue to be, increasingly important in future, particularly in light of the complete mapping of the human genome. The ability to target a particular gene or control region of genetic DNA in a sequence-specific fashion is one of the biggest challenges in drug design today. The strategy of inhibiting the transcription or translation of a specific gene has several advantages. These include low drug concentration needed (in theory, only a few molecules of a gene-targeting

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Summary Here we have stressed important differences between protein and DNA crystallography. Crystal growth and data collection methodologies are not directly transferable between the two subfields. In addition, we note that analysis of symmetry and packing of DNA crystals can be useful and a uniquely aesthetic exercise. Acknowledgments This work was supported by the National Science Foundation (Grant M C B - 9 9 7 6 4 9 8 ) and the A m e r i c a n C a n c e r Society (Grant R P G - 9 5 - 1 1 6 - 0 3 - G M C ) .

[14] Molecular Modeling of Drug-DNA Complexes: An Update By J O H N

O. TRENT

Introduction This is an exciting time in the field of molecular modeling of DNA and DNA :ligand structures. There have been several advances in the field, such as second-generation force fields and the implementation of the particle mesh Ewald (PME) methods that more accurately reproduce experimental structures. When combined with the ever-increasing speed of computational resources (supercomputers and, in particular, parallel processor workstations), this places molecular simulations of biologically relevant systems at the fingertips of researchers. We now are able to investigate systems that previously were not practical, either because of less reliable force fields or computational expense. This is not to say that the discipline of molecular modeling has fully matured, as we are only at the second generation of force fields. However, with the increasing reliability, physical size of simulated DNA and proteins, and length of simulations, more researchers are turning toward simulations to aid them in the rationalization and prediction of experimental data, and in the design of specific experiments. The application of molecular modeling (and structural biology as a whole) to nucleic acids is, and will continue to be, increasingly important in future, particularly in light of the complete mapping of the human genome. The ability to target a particular gene or control region of genetic DNA in a sequence-specific fashion is one of the biggest challenges in drug design today. The strategy of inhibiting the transcription or translation of a specific gene has several advantages. These include low drug concentration needed (in theory, only a few molecules of a gene-targeting

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drug w o u l d be required per cell), the r e m o v a l of nonspecific side effects (observed for m o s t current c h e m o t h e r a p e u t i c drugs), and the ability to investigate the effect o f "turning o f f " one or m o r e genes. There are several D N A - b a s e d approaches that are currently under investigation to do this, such as the antigene l-5 (targeting transcription by triplex formation), antisense 4'6'7 (targeting translation), and aptamer s-~3 (posttranslational, targeting particular proteins) approaches. The antisense and antigene approaches use the natural base c o m p l e m e n t a r i t y of base pairs and base triplets, respectively, to target relatively long regions (typically 2 0 - 3 0 bases) o f R N A or D N A . In theory it is possible to use synthetic organic ligands as an alternative to targeting specific gene sequences, and several researchers ~4-19 are investigating this approach. This o f course relies on the ability to design sequencespecific ultratight D N A - b i n d i n g ligands. Our understanding o f the c o n s e q u e n c e s o f specific genetic manipulation is s o m e w h a t limited at this stage, but is likely to expand with the advent of new technologies such as G e n e Chip arrays 2° and proteomics.2

J K. R. Fox, Curr. Med. Chem. 7, 17 (2000). 2 K. M. Vasquez and J. H. Wilson, Trends Biochem. Sci. 23, 4 (1998). 3 H. G. Kim, J. F. Reddoch, C. Mayfield, S. Ebbinghaus, N. Vigneswaran, S. Thomas, D. E. Jones, and D. M. Miller, Bioehemistry 37, 2299 (1998). 4 p. C. vander Vliet and J. D, Gralla, eds., Biochim. Biophys. Acta Gene Struet. Express. 1489(1) (1999). This issue is dedicated to oligonucleotide therapies and has several excellent reviews. 5 p. p. Chan and P. M. Glazer, J. Mol. Med. 75, 267 (1997). 6 S. Agrawal and E. R. Kandimalla, Mol. Med. Today 6, 72 (2000). 7 S. T. Crooke, Methods Enzymol. 313, 3 (2000). s S. A. Robertson, K. Harada, A. D. Frankel, and D. E. Wemmer, Biochemistry 39, 946 (2000). 9 I, Smimov and R. H. Shafer, Biochemistry 39, 1462 (2000). I0 D. E. Huizenga and J. W. Szostak, Biochemistry 34, 656 (1995). 11 L. Gold, J. BioI.Chem. 270, 13581 (1995). 12 p. j. Bates, J. B. Kahlon, S. D. Thomas, J. O. Trent, and D. M. Miller, J. Biol. Chem. 274, 26369 (1999). 13 M. Koizumi and R. R. Breaker, Biochemistry 39, 8983 (2000). 14N. R. Wurtz and P. B. Dervan, Chem. Biol. 7, 153 (2000). ~5p. B. Dervan and R. W. Burli, Curr. Opin. Chem. Biol. 3, 688 (1999). 16C. A. Hawkins, R. P. d. Clairac, R. N. Dominey, E. E. Baird, S. White, P. B. Dervan, and D. E. Wemmer, J. Am. Chem. Soc. 122, 5235 (2000). 17 R. L. d. Clairac, B. H. Geierstanger, M. Mrksich, P. B. Dervan, and D. E. Wemmer, J. Am. Chem. Soc. 119, 7909 (1997). 18j. B. Chaires, E Leng, T. Przewloka, I. Fokt, Y. H. Ling, R. Perez-Soler, and W. Priebe, J. Med. Chem. 40, 261 (1997). 19B. Martin, A. Vaquero, W. Priebe, and J. Portugal, Nucleic Acids Res. 27, 3402 (1999). 20 U. Scherf, D. T. Ross, M. Waltham, L. H. Smith, J. K. Lee, L. Tanabe, K. W. Kohn, W. C. Reinhold, T. G. Myers, D. T. Andrews, D. A. Scudiero, M. B. Eisen, E. A. Sausville, Y. Pommier, D. Botstein, P. O. Brown, and J. N. Weinstein, Nat. Genet. 24, 236 (2000). Comment in Nat. Genet. 24, 208 (2000). 21 A. A. Alaiya, B. Franzen, G. Auer, and S. Linder, Electropholesis 21, 1210 (2000).

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It is apparent for most biologically relevant systems that a multidisciplinary approach, including biochemistry, bioinformatics, biophysics, chemical synthesis, molecular biology, and structural biology [nuclear magnetic resonance (NMR), X-ray crystallography, molecular modeling], is needed to fully elucidate the system of interest. This review covers the advances since 1995 in DNA and DNA : ligand molecular modeling. There are several excellent reviews 22-25 that cover the development and theoretical aspects of molecular modeling, and this review briefly summarizes these advancements and focuses on the applications of these to biologically relevant systems. Molecular Modeling

Introduction Molecular modeling is a broad term that encompasses ab initio quantum mechanical calculations, semiempirical calculations, and empirical calculations (charge-dependent molecular mechanics force fields). These techniques can be used to study the three-dimensional structure, dynamics, and properties of a molecule of interest. The force field approach has been traditionally used for DNA calculations of any size (more than a few bases) because of the computational expense of the ab initio or semiempirical approaches. However, the increasing CPU power of workstations and the advent of hybrid methods such as quantum mechanics/molecular mechanics (QM/MM) 26'27 now make it possible to treat a region of interest with a high level of theory while considering the surrounding shell of the system at a less computationally intensive level. Molecular modeling, particularly molecular mechanics and dynamics, are highly complementary to macromolecular NMR and X-ray crystallography. A combined approach is desirable and, where the parent experimental structure is available, the simulation can be partially validated 28 by reproducing the structure (assuming the structure is correct). The system of interest can then be investigated with more confidence.

Advances in Molecular Modeling of DNA Issues of molecular motion and flexibility are expected to be a crucial component in developing a fundamental understanding of the relationship between DNA structure and function in molecular biophysics and genetics. Diverse

22 D. Beveridge and K. Campbell, Curl: Opin. Struct. Biol. 10, 182 (2000). 23 T. E. Cheatham and P. Kollman, Annu. Rev. Phys. Chem. 51,435 (2000). 24 T. E. Cheatham, B. R. Brooks, and P. A. Kollman, in "Current Protocols in Nucleic Acid Chemistry," p. 7.5.1. John Wiley & Sons, New York, 1999. 25 T. E. Cheatham and B. R. Brooks, Theor. Chem. Acc. 99, 279 (1998).

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biophysical methods can be applied, but no single experimental technique is capable of full elucidation of the dynamical structure of DNA in a solvent environment. Molecular dynamics simulation can, in principle, provide a complete theoretical description of DNA structure and motions, and is thus a valuable independent means of developing models and interpreting experimental data. Furthermore, molecular dynamics force fields now play a key supplementary role in the determination of structures by iterative refinement procedures in crystallography and NMR spectroscopy. Both X-ray crystallography and NMR rely on fitting experimental data [electron density, or nuclear Overhauser effect restraints (NOEs), respectively] to an empirical force field. It should be noted that if the experimental data has low-quality regions, then the structures are dependent on the inherent force field to a greater extent. It is becoming increasingly popular for NMR investigators to use "complete system" protocols with full periodic box solvation and the particle mesh Ewald (PME) methods to improve the structure refinement (see Duplex DNA : Ligand Complexes, below). This seems a logical approach as the NOEs are determined in the solution phase, and so the best approximation of the solution phase should be used for the structure refinement. Molecular dynamics simulations have proved to be an excellent adjunct to experimental techniques for the determination of protein structures. 29 However, it is only the advances in force fields and the accurate representation of long-range effects that make molecular dynamics extremely powerful for the study of DNA structures. The two most significant advances in modeling of DNA have been the development of second-generation force fields and the effective treatments of longrange electrostatic interactions. Three second-generation all-atom force fields 3°-32 specifically designed for simulations including explicit solvent were developed in 1995. Two, AMBER 31 and CHARMM, 32'33 have been updated and extensively applied to DNA simulations. Discussions of the relative merits of these force fields are available 22,23,28 and in some instances similar effects are being reported. In both cases the force fields are now based on ab initio calculations with derived parameters.

26 Q. Cui and M. Karplus, J. Phys. Chem. B 104, 3721 (2000). 27 B. Hernandez, E J. Luque, and M. Orozco, J. Comput. Aided Mol. Des. 14, 329 (2000). 28 W. E van Gunsteren and A. E. Mark, J. Chem. Phys. 108, 6109 (1998). 29 p. A. Kollman, Acc. Chem. Res. 29, 461 (1996). 3o M. Levitt, M. Hirshberg, R. Sharon, and V. Daggett, Comput. Phys. Commun. 91, 215 (1995). 3JW. D. Cornell, E Cieplak, C. I. Bayly, I. R. Gould, K. M. Merz, D. M. Ferguson, D. C. Spellmeyer, T. Fox, J. W. Caldwell, and R A. Kollman, J. Am. Chem. Soc. 117, 5179 (1995). http://www.amber.ucsf.edu, and references therein. 32 A. D. MacKerell, J. Wiorkiewicz-Kuczera, and M. Karplus, J. Am. Chem, Soc. 117, 11946 (1995). 33 B. R. Brooks, R. E. Bruccoleri, B. D. Olafson, D. J. States, S. Swaminathan, and M. Karplus, J. Cornp. Chem. 4, 187 (1983). http://yuri.harvard, edu/.

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Use of the Ewald methods in the treatment of long-range electrostatic interactions has become popular with efficient implementations of the particleparticle mesh Ewald, 34'35 and the particle mesh Ewald 36-38 methods (see Sagui and Darden 39 for a review). Molecular dynamics simulations using the PME method show significant improvement in the dynamic stability of simulated DNA, and stable multiple nanosecond trajectories of explicitly solvated [using periodic boundary conditions (PBC)] DNA are now routine using PME methods. However, possible artifacts 4°,4j from this treatment may occur if the system is not neutrally charged or exhibits large effective charge separation, although a net correction term has been reported 42 to remove artifacts of energy for systems with a net charge. Also the "flying ice cube" 43 is a potential problem with periodic velocity rescaling, which can be alleviated by periodically removing the translational and rotational motion or reassigning the velocities. Finally, the treatment of low dielectric solvents has been investigated4°'41 with potential disturbing aspects. The popular choice for simulations includes a second-generation force field with a PME implementation and PBC, and these have been shown to reproduce experimental observations (see below). There has been a resurgence in the use of implicit solvation techniques, as these are computationally less intensive and greater simulation times are available. Originally implemented by Still et a1.,44 the generalized Born/solvent accessibility (GB/SA) method, with appropriate parameters, can reproduce hydration free energies, 45'46 and the development of GB parameters compatible with AMBER is included in AMBER 6.0. However, the implicit solvation methods do not include specific ion and water association interactions. DNA Duph, x Simulation

Some of the first solution-phase simulations47-51 showed that nanosecond simulations were stable and reproduced a B-form DNA, instead of the intermediate A 34R. W. Hockney and J. W. Eastwood, "'Computer Simulation Using Particles." McGraw-Hill, New York, 1981. 35 B. A. Luty, 1. G. Tironi,and W. F. van Gunsteren.,/. Chem. Phys. 103, 3014 (1995). 3f~T. A. Darden, D. M. York, and L. G. Pedersen,.1. Chem. Phys. 98, 10089(1993). 3vU. Essmann,L. Perera, M. L. Berkowitz,T. A. Darden, H. Lee, and L. G. Pedersen,.1. Chem. Phys. 103, 8577 (1995). :~sH. G. Petersen,J. Chem. Phys. 103, 3668 (1995). 3'~C. Sagui and T. A. Darden,Aimu. Rev. Biophys. Biomol. StI'I~('L 28, 155 (1999). 4OR H. Hunenbergerand J. A. McCammon,Biophys. Chem. 78, 69 (1999). 41 R H. Hunenbergerand J. A. McCammon,J. Chem. Phys. 110, 1856(1999). 42 S. Bogusz,T. E. Chealham,and B. Brooks,.I. Chem. Phys. 109, 7070 (1998). 4:~S. C. Harvey,R. K. Z. Tan,and T. E. Cheathmn,.I. Comput. Chem. 19, 726 (1998). 44 W. C. Still, A. Tempczyk,R. C. Hawley,and T. Hendrickson,.I. Am. Chem. Soc. 112, 6127 (1990). 45 B. Jayaram, D. Sprous, and D. L. Beveridge,.1. Phys. Chem. B 102, 9571 (1998). 46 B. N. Dominyand C. L. Brooks,J. Phys. Chem. B 103, 3765 (1999).

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and B forms from previous simulations. This was extended 52'53 with the AMBER second generation force field by Comell et al. 31 on a larger time scale (>5 nsec) with varied Na + ion distributions, using PME methods. These simulations suggested that the force field reproduced B-type DNA extremely well. The fact that the simulation was closer to canonical B-DNA than the crystal structure does not make the simulation less accurate; rather, the crystal structure had contributing crystal packing effects. It is more appropriate to compare NMR solution structures with solvated simulations and in this case there was good agreement with the NMR structure 54 from Lane's group. However, few helical parameters were available for comparison. Improvements in the deficiencies such as unwinding of the DNA, average sugar pucker, and )~ angles in the Cornell force field have been made. 55 C o n f o r m a t i o n a l Transitions

The stable dynamics simulations enabled Cheatham et al. to examine conformational transitions, such as the A- to B-form transition. Under low-salt conditions, using the Cornell et al. 31 force field, the A- to B-form transition occurred in 500 psec. 56 It also was shown 57 that models of structures that experimentally stabilize in the A form, such as RNA : RNA and RNA : DNA duplexes, were stabilized in the A form by the Cornell et al. 31 force field. In addition, the phosphoramidatemodified DNA, which has a known preference for the A form, was observed to undergo a B- to A-form transition in the simulation. 58 Cheatham et al. 59 and Beveridge's group 6° have extended these simulations by modeling experimental conditions known to stabilize A-from DNA, such as high ethanol content, and found stabilization of the A-form DNA. Langley has developed the BMS force field for nucleic acids that shows the B- to A-form DNA transitions in 75% ethanol, and also reproduces this under high-salt conditions (4 M NaC1). Under ionic conditions 47 Y. E. Cheatham, J. L. Miller, T. Fox, T. A. Darden, and P. A. Kollman, J. Am. Chem. Soc. 117, 4193 (1995). 48 D. A. Zichi, ,l. Am. Chem. Soc. 117, 2957 (1995). 49 S. Weerasinghe, P. E. Smith, V. Mohan, Y.-K. Cheng, and B. M. Pettitt, J. Am. Chem. Soc. 117, 2147 (1995). 5o S. Weerasinghe, P. E. Smith, and B. M. Pettitt, Biochemistry 34, 16269 (1995). 51L. Yang, S. Weerasinghe, E E. Smith, and B. M. Pettitt, Biophys. J. 69, 1519 (1995). 52 M. A. Young, B. Jayaram, and D. L. Beveridge, .l. Am. Chem. Soc. 119, 59 (1997). 53 M. A. Young, G. Ravishanker, and D. L. Beveridge, Biophys. J. 73, 2313 (1997). 54 A. Lane, T. C. Jenkins, T. Brown, and S. Neidle, Biochemiso 3, 30, 1372 (1991). 55 T. E. Cheatham, E Cieplak, and E A. Kollman, J. Biomol. Struct. Dyn. 16, 845 (1999). 56 T. E. Cheatham and E A. Kollman, J. Mol. Biol. 259, 434 (1996). 57 Y. E. Cheatham and P. A. Kollman, J. Am. Chem. Soc. 119, 4805 (1997). 58 p. Cieplak, I. T. E. Cheatham, and E A. Kollman, J. Am. Chem. Soc. 119, 6722 (1997). 59 T. E. Cheatham, M. E Crowley, T. Fox, and E A. Kollman, Proc. Natl. Acad. Sci. U.S.A. 94, 9626 (1997). 6o D. Sprous, M. A. Young, and D. L. Beveridge, J. Phys. Chem. B 102, 4658 (1998).

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known to stabilize A-form DNA the Cornell e t a l . 31 force field also showed spontaneous transition of B-form to A-form DNA. 6j The A- to B-form transition also has been observed for triplex simulations 62 using A M B E R (PBC and PME conditions). The conformation of duplexes involving peptide nucleic acid (PNA) also has been investigated using A M B E R and C H A R M M . Molecular dynamics simulations of the PNA : DNA and PNA : RNA duplexes, using A M B E R (PBC and PME), by Soliva et al. 63 demonstrated the reproduction of PNA : DNA and PNA : RNA NMR structures. They found that the radius of convergence was wide, although it was noted that convergence can fail if the starting model is severely in error. Sen and Nilsson 64 used C H A R M M (PBC) to investigate the PNA : PNA duplex, PNA : DNA antiparallel duplex, PNA : DNA parallel duplex, and DNA : DNA duplex and reported excellent agreement for the PNA : PNA duplex with the crystallographic structure and good agreement of the PNA : DNA antiparallel duplex with the N M R structure. Overall this is encouraging, as the second-generation force fields are beginning to reproduce experimental structures with experimental conditions. Further improvements are likely to extend the experimental conditions that can be reproduced, for example, the B- to Z-form and Z- to B-form transitions, which are currently under investigation. 65"66 DNA Hydration and Ion Interactions

There has been great interest in the hydration and ion interactions of DNA stemming largely from the Williams 67 high-resolution (1.4-]k) crystal structure, which clearly shows the detailed spine of hydration of the minor groove extending past the first solvation shell. Several groups 52'53'57-59'61,68-75 have shown that molecular dynamics simulations can accurately reproduce several features of DNA hydration 61 T. E. Cheatham and E A. Kollman, Structure 5, 1297 (1997). 62 G. C. Shields, C. A. Laughton, and M. Orozco, J. Am. Chem. Soc. 119, 7463 (1997). 63 R. Soliva, E. Sherer, E J. Luque, C. A. Laughton, and M. Orozco, J. Am. Chem. Soc. 122, 5997 (2000). 64 S. Sen and L. Nilsson, ,l. Am. Chem. Soc. 120, 619 (1998). 65 j. O. Trent, unpublished data, Gridrun program (available from the author). 66 T. E. Cheatham, personal communication (2000). 67 L. D. Williams, Biochemistry 37, 8341 (1998). 68T. E. Cheatham, J. Srinivasan, D. A. Case, and R A. Kollman, J. Biomol. Struct. Dyn. 16, 265 (1998). 69 y. Duan, R Wilkosz, M. Crowley, and J. M. Rosenberg,J. Mol. Biol. 272, 553 (1997). 7o M. Feig and B. M. Pettitt, J. Mol. Biol. 286, 1075 (1999). 7t M. A. Young and D. L. Beveridge,J. Mol. Biol. 281, 675 (1998). 72 D. R. Langley, J. Biomol. Struct. Dyn. 3, 487 (1998). 73 M. Feig and B. M. Pettitt, J. Phys. Chem. B 101, 7361 (1997). 74 M. Feig and B. M. Pettitt, Biophys. J. 75, 134 (1998). 75 D. R. Langley,J. Biomol. Struct. Dyn. 16, 1366 (1999).

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including the spine of hydration in A tracts, hydration in the major groove, the cones of hydration around phosphates, and the differences in the hydration patterns for A-form DNA and RNA. Specific ionic interactions also are being reproduced by simulations, 52'53'57"59' 61,71,72,75,76 such as the ion association in the minor groove of B-form DNA and in the major groove of A-form DNA. Moreover, molecular dynamics reproduced specific effects on the stabilization of the structure that are consistent with experimental data, such as enhanced bending of the A tracts, 71'77the B- to A-form transitions, 61'72 and differences in the behavior of specific ions. 76'78 Again, the ability to reproduce structures consistent with experimental conditions is highly encouraging. Poisson-Boltzmann calculations have been used to look at multivalent ions and the B- to Z-form transitions. 79 A review by Pack et aL so details Monte Carlo and Poisson-Boltzmann approaches to examine divalent cations and the electrostatic potential surrounding DNA. Modeling of DNA : Ligand Complexes Several examples of modeling DNA:ligand complexes and multistranded DNA have been chosen, with particular emphasis on the utilization of the advances mentioned above. Duplex DNA : Ligand Complexes. Several groups have been applying explicit water and PME methods to aid in the refinement of NMR data. Williams and Searle 81 have studied the structure, dynamics, and hydration of nogalamycin with d(ATGCAT)2 by using NMR and molecular dynamics and show that many sequence-dependent structural features observed in X-ray crystal structures lie within the range of dynamic fluctuations observed in the molecular dynamics simulations. The Searle and Laughton groups also have examined s2 the DNA minor groove recognition by a tetrahydropyrimidinium analog of Hoechst 33258 by NMR and molecular dynamics studies of the complex with d(GGTAATTACC)2, and the molecular recognition and dynamic equilibrium between a pentacyclic acridinium salt and DNA sequences in a combined approach of optical spectroscopic techniques, NMR, and molecular dynamics 83 (AMBER, PBC, and PME). The ability to calculate thermodynamic parameters for DNA : ligand complexes allows direct comparison with experiments. The absolute free energy of association 76 A. D. MacKerell, J. Phys. Chem. B 101,646 (1997). 77 C. E. Bostock-Smith, C. A. Laughton, and M. S. Searle, Biochem. J. 342, 125 (1999). 78 A. P. Lyubartsev and A. Laaksonen, J. Biomol. Struct. Dyn. 16, 579 (1998). 79 M. Gueron, J. Demaret, and M. Filoche, Biophys. J. 78, 1070 (2000). 80 G. R. Pack, L. Wong, and G. Lamm, Biopolymelw 49, 575 (1999). 81 H. E. Williams and M. S. Searle, J. Mol. Biol. 290, 699 (1999). 82 C. E. Bostock-Smith, C. A. Laughton, and M. S. Searle, Nucleic Acids Res. 26, 1660 (1998). 83 C. E. Bostock-Smith, E. Gimenez-Arnau, S. Missailidis, C. A. Laughton, M. E Stevens, and M. S. Searle, Biochemisoy 38, 6723 (1999).

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of netropsin with the Dickerson dodecamer has been determined with the AMBER suite of programs by Singh and Kollman 84 and a fully detailed atomic model for both solute and solvent. The calculated free energy of - 10.3 kcal/mol is in good agreement with the experimentally determined value of - 1 1.5 kcal/mol, and the number of water molecules found to diffuse into the uncomplexed DNA was equal to the number of water molecules in the binding site of the uncomplexed crystal structure. The free energy difference between thymine and difluorotoluene in DNA duplex has been determined by molecular dynamics and thermodynamic integration and was found 85 to be in good agreement with experimental values. This was extended to make predictions about the effect of the difluorotoluene substitution in triplex DNA. Triplex DNA Complexes. The simulation of triplex DNA is well established with reports from Laughton and co-workers62'86 and Pettitt,49'5°'87 among others. It has been shown 62 for the TAT parallel triplex structure, by using molecular dynamics (AMBER with PME), that the A-, B-, and P-form (PNA-like) starting models converge to a single B-form structure that is consistent with experimental evidence. We have observed similar convergence effects for the RNA third strand: DNA triplexes (see Triplex Simulation of Antigene Approach, below). Molecular dynamics simulations with AMBER have been used 88 to investigate the PNA. DNA : PNA system and convergence of starting models also has been demonstrated. It is well known that certain ligands have a preference for triplex DNA over other forms of DNA, and a new dialysis assay (see [5] in this volume TM) enables the direct comparison of binding preferences of ligands for different forms of duplex, triplex, and quadruplex DNA. Qualitative modeling 89 of intercalating mono- and disubstituted amidoanthraquinone regioisomers has rationalized the relative affinities determined by DNase I footprinting. Mithramycin and distamycin also have been investigated for triplex stability by gel mobility shifts, nuclease and chemical hypersensitivity, two-dimensional gel topological analyses, triplex-specific antibody-binding studies, and qualitative modeling. 9° These minor groove binders have preferences for GC (A-type DNA) and AT (B-type DNA), respectively. Modeling reproduced the experimental observation that mithramycin 84 S. B. Singh and E A. Kollman, J. Am. Chem. Soc. 121, 3267 (1999). 85 E. Cubero, C. A. Laughton, E J. Luque, and M. Orozco, J. Am. Chem. Sot., in press (2000). 86 R. Soliva, C. A. Laughton, F. J. Luque, and M. Orozco, J. Am~ Chem. Soc. 120, 11226 (1998). 87 Y.-K. Cheng and B. M. Pettitt, Biopolymers 35, 457 (1995). 88 G. C. Shields, C. A. Laughton, and M. Orozco, J. Am. Chem. Soc. 120, 5895 (1998). 88a j. Ren and J. B. Chaires, Methods Enzymol. 340, [5] 2001 (this volume). 89 M. D. Keppler, M. A. Read, E J. Perry, J. O. Trent, T. C. Jenkins, A. E Reszka, S. Neidle, and K. R. Fox, Era: J. Biochem. 263, 817 (1999). 9o N. Vigneswaran, J, Thayaparan, J. Knops, J. O. Trent, D. M. Miller, and W. Zacharias, Biol. Chem., in press (2001).

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destabilizes triplex and also inhibits triplex formation, whereas distamycin stabilizes triplex and facilitates triplex formation. Q u a d r u p l e x D N A Complexes. Molecular dynamics simulations have reproduced the high-resolution crystallographic structure and NMR structures of several quadruplex species. The parallel d(G4)4 quadruplex simulation 91 including the central core Na + ions reproduces the crystal structure extremely well [ < 1-]k root mean square deviation (RMSd)], and it was observed that the Na + ions are important for stability. The antiparallel quadruplex d(G4T4G4)2 simulation 91 also compares will with the NMR structure. A comparison 92 of the in vacuo and fully solvated (AMBER, PME) NMR refinement of the antiparallel d(G3T4G3)2 quadruplex indicated that explicit solvent treatment led to better R factors. An NMR investigation 93 into the determinants of DNA quadruplex structures and potassium binding, using C H A R M M (implicit solvation), showed that potassium is needed for the chair-type but not for the basket-type quadruplex. Ligands that bind to quadruplexes have been of interest 94 as potential telomerase inhibitors. The amidoanthraquinone series has been reported 95 to have inhibitory activity and qualitative modeling shows that "end pasting" is one possible binding mode for the human telomere quadruplex. More detailed simulations 96 using explicit water and the consistent force field (CFF) supported this, although other binding sites were not examined. The porphyrin class of telomerase inhibitors has been examined 97 by isothermal titration calorimetry (ITC), spectrophotometry, and fully solvated molecular dynamics (AMBER and PME), and this clearly showed intercalation in a number of quadruplexes. Protocols for Molecular Modeling of DNA: Ligand Complexes When contemplating a molecular modeling simulation the first priority is to determine the fundamental question to be answered and to decide which (if any) method is the most appropriate to achieve this. There is little point in performing a simulation that cannot, by the experimental design and limitations of the algorithms, provide the answer or insight into the fundamental question. 91 N. Spackova,I. Berger, and J. Sponer,J. Am. Chem. Soc. 121, 5519 (1999). 92G. D. Strahan, M. A. Keniry, and R. H. Shafter, Biophys. J. 75, 868 (1998). 93W.M. Marathias and P. H. Bolton,Biochemistry 38, 4355 (1999). 94D. Sun, B. Thompson, B. E. Cathers, M. Salazar, S. M. Kerwin, J. O. Trent. T. C. Jenkins, and S. Neidle, J. Med. Chem. 40, 2113 (1997). 95p. j. Perry, A. P. Reszka, A. A. Wood,M. A. Read, S. M. Gowan, H. S. Dosanjh, J. O. Trent, T. C. Jenkins, L. R. Kelland, and S. Neidle,J. Med. Chem. 41, 4873 (1998). 96M. A. Read, A. A. Wood, J. R. Harrison, S. M. Gowan, L. R. Kelland, H. S. Dosanjh, and S. Neidle, J. Med. Chem. 42, 4538 (1999). 97I. Haq, J. O. Trent, B. Z. Chowdhry,and T. C. Jenkins, J. Am. Chem. Soc. 121, 1768 (1999).

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M o l e c u l a r D y n a m i c s Simulations

It should be emphasized that no two systems are alike and that most protocols must be adapted to every situation. Simulations should be constantly monitored to check progress. Five components are required to performing molecular dynamics simulations, namely, model building, charge generation, mechanics and dynamics, visualization, and analysis. Various software packages are available that perform one or more of these functions. The increasing abundance of more user-friendly modeling programs is enabling a greater number of researchers to utilize molecular modeling. However, extreme care should be taken and a "black box" approach should never be used. Investigators should understand in detail the "default" parameters and should tailor the simulation to meet their needs in accordance with the literature on simulations. The software used in this chapter is Macromodel, 9s GAMESS, 99 the AMBER suite, 3j GRASP, 1°° Insight II, and Curves. I°1 There are three basic steps in the overall simulation: the generation of starting models, the actual simulation, and the evaluation of the production run. Generation o f Starting Models D N A Duplex Generation. Generation of the DNA starting model (or models) can be done by a variety of software programs, such as NUCGEN, 3j NAB, 1°2 and Macromodel, 98 which can generate different forms (A-form, B-form, Z-form) of duplex DNA. The Protein Database 1°3 (PDB) and the Nucleic Acid Database TM (NDB) are invaluable resources for obtaining experimentally determined structures. Care should be taken with the addition of hydrogen atoms to experimentally derived crystal and software-generated DNA structures, as some software can place them in nonoptimal positions. Also, atom names and the format of the PDB file often need to be changed when generating structures in one program and using another for the simulation because, although there is a standard PDB format, historical variations are still common. The file conversion program Babel 1°5 is

98E Mohamadi, N. G. Richards, W. C. Guida, R. Liskamp, M. Lipton, C. Caufield, G, Chang, T. Hendricksen,and W. C. Still,J. Comp. Chem. 11,440 (1990). http://www.schrodinger.com. 99M. W. Schmidt, K. K. Baldridge,J. A. Boatz, S. T. Elbert, M. S. Gordon,J. J. Jensen, S. Koseki, N. Matsunaga,K. A. Nguyen,S. Su, T. L. Windus,M. Dupuis,and J. A. Montgomery,.I. Comp. Chem. 14, 1347 (1993). http://www.msg.ameslab.gov/gamess/gamess.html. lo0 A. Nicholls, K. Sharp, and B. Honig,Proteins Struct. Funct. Genet. 11, 281 (1991). http://trantor. bioc.columbia.edu/grasp/. 101R. Lavery and H. Sklenar, J. Biomol. Struct. Dyn. 6, 63 (1998). http://www.ibpc.fr/ UPR9080/curindex.html. 1olT. Macke and D. A. Case, in "Molecular Modelingof Nucleic Acids" (N. B. Leontes and J. S. Lucia, eds.), p. 379. AmericanChemical Society, Washington,D.C., 1998. 1o3H. M. Berman,J. Westbrook,Z. Feng,G. Gilliland,T. N. Bhat, H. Weissig,I. N. Shindyalov,and R E. Bourne,Nucleic Acids Res. 28, 235 (2000). 1o4H. M. Berman,W. K. Olson,D. L. Beveridge,J. Westbrook, A. Gelbin,T. Demeny,S. H. Hsieh, A. R. Srinivasan,and B. Schneider,Biophys. J. 63, 751 (1992).

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useful in the manipulation of different file formats, although most modeling software will now export PDB files. A program that converts nonstandard PDB files to AMBER-style PDB files with atom renaming (based only on identifying the element of each individual atom) is available from the author. Generation of Triplex DNA. There are a number of NMR triplex structures in the PDB and NDB that can be modified for the generation of the particular triplex sequence and motif (parallel or antiparallel) of interest. The most notable X-ray crystal structure is that of Rhee et al., 1°6 which has an overlap region of three consecutive base triplets consisting of C+-GC, BU-ABU (BU, 5-bromouracil), and C+-GC. Alternatively, the fiber diffraction structures of parallel and antiparallel triplexes are available 1°7 109 and a simple algorithm can be used to generate the triplex of interest. However, care should be taken as the triplex structure generated by this method has enforced symmetry. Additional equilibrium periods should be utilized to relax the symmetry prior to using these structures as starting models. Generation of Quadruplex DNA. The generation of parallel and antiparallel quadruplex is based on crystallographic and NMR structures, particularly the high-resolution (0.75-~) parallel d(TGGGGT)4 structure of Phillips et al.ll° and the antiparallel human telomere d[AG3(T2AG3)3] NMR structure of Wang and Patel.I I1 There are a number of different motifs available in the PDB and NDB and, as the structures of interest are usually short (2-4 base quartets), these structures can easily be modified for use. DNA Ligand Generation. Generation of the starting model for each DNA ligand is variable. Many different programs can be used to generate the initial threedimensional model of the ligand. The three-dimensional coordinates may be available from the Cambridge Structure Database, j J2 or the structure of the DNA : ligand complex may already have been reported. More commonly, when dealing with a new ligand, it will have to be optimized and parameterized in some way. The geometry can be optimized at a level suited to the overall calculation. Many of the van der Waals, bond, angle, torsion, and improper torsion parameters can be derived by analogy. A particular force field may already be parameterized for the ligand, such Jo5 p. Waiters and M. Stahl, ftp://ccl.osc.edu/pub/chemistry/software/UNIXfoabel/. 106 S. Rhee, Z. Han, K. Liu, H. T. Miles, and D. R. Davies, Biochemistry 38, 16810 (1999). 1o7 G. Raghunathan, H. T. Miles, and V. Sasisekharan, Biochemistry 32, 455 (1993). 108 G. Raghunathan, H. T. Miles, and V. Sasisekharan, Biopolymers 36, 333 (1995). 109 E B. Howard, H. T. Miles, K. Liu, J. Frazier, G. Raghunathan, and V. Sasisekharan, Biochemistry 31, 10671 (1992). lm K. Phillips, M. Orozco, E Q. Luque, S. J. Teat, W. Clegg, and B. Luisi, to be published (PDB code 1DJ4). IlL y. Wang and D. J. Patel, Structure 1,263 (1993). 112 E H. Allen, S. Bellard, M. D. Brice, B. A. Cartright, A. Doubleday, H. Higgs, T. Hummelink, B. G. Hummerlink-Peters, O. Kennard, W. D. S. Motherwell, J. R. Rodgers, and D. G. Watson, Acta Crystallogr B 35, 2331 (1979),

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as MM2, and can be used to provide a reasonable starting model. If conformational flexibility is present, a grid search by torsion driving (one to three torsion angles) or Monte Carlo searches (greater than three torsion angles) can be used to fully explore the conformational space of the ligand. Other conformational techniques can be employed such as pure methods JBW, ADF, TMC, and SD or mixed methods jl3 ADF/JBW, SD/TMC (original MCSD method), SD/TMC, SD/JBW, and SD/ADF/JBW. It is important to get close to the "global" minima as the ligand may undergo a conformational change on binding to DNA and this energy penalty is included in the overall energy of the complex. The starting models can be used for ab initio optimization and analogous parameters may be incorporated into the force field, although great care must be taken as the format of force fields may be different in the terms they use. The worst case scenario is that full parameterization using ab initio calculations is required, although this is faster than in the past. The level of effort that should be expended is related to the scientific question being asked. (For a tutorial on parameter development see Kollman's tutorial, http://www.amber.ucsf.edu/amber/newparams.html. The calculation of accurate partial charges uses an ab initio program, such as Gaussian 98 or GAMESS 99 (this can be done with parallel processors). The accurate representation of electrostatic interactions is crucial for a force field intended for application to biological molecules, particularly as DNA is a highly charged polyanionic molecule. The charges also should be consistent with the force field. When using the A M B E R force field at least the 6-31G* basis set should be used, and restricted electrostatic potential (RESP) j 14,~15 charges are advisable. DNA : Ligand Complex Generation. The starting position(s) of the ligand in the D N A : l i g a n d complex is a crucial element. Ideally a similar structure has been solved and coordinates can be superimposed to produce a reasonable starting model. However, obviously not all systems of interest have been solved, and although force fields have improved and can find stable and experimentally derived DNA conformations from starting models that are reasonably distant from the final structure, the DNA : ligand starting complex must be chosen with care. There are two pitfalls here; the first is that the conformations become locked into a local minima so that it cannot move over local energy barriers, and second, user bias is incorporated so that the final structure derived is that which was considered most likely by the investigator. Ideally the ligand should be placed near the DNA and allowed to "find" the most stable bound complex. However, even if this were possible, it would require simulations that are on a much greater time scale than are currently accessible. There are a number of ways to overcome this situation. l J3 SD, stochastic dynamics; ADF, all degrees of freedom Metropolis Monte Carlo; TMC, torsional Monte Carlo; JBW,jumping between wells importance-samplingMonte Carlo. 114C. I. Bayly, P. Cieplak, W. D. Comell, and P. A. Mollman,J. Phys. Chem. 97, 10269 (1993). 115W. D. Cornell, E Cieplak, C. I. Bayly, and E A. Kollman,J. Am. Chem. Soc. 115, 9620 (1993).

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Simulated annealing, Monte Carlo searches, and docking calculations can be used to obtain starting structures. Often the ligand is in a known class of DNA binder, such as an intercalator or a groove binder, which reduces the uncertainty of binding location. Also, additional experimental evidence, such as DNase I footprinting and sequence preference techniques such as REPSA, 116 can provide information that is useful in generation of the complex. Generation o f DNA : lntercalator Starting Models. The intercalation site can usually be obtained and modified from similar structures from the PDB or NDB for duplex DNA. A known intercalator step can be incorporated into the sequence of interest. Alternatively, an intercalation step can be built by placing the intercalation site bases at a similar rise and helical twist as known intercalation sites. The phosphodiester (or other) backbone can be inserted and optimized while constraining the rest of the DNA. The optimization should follow a protocol similar to that used for the overall simulation and include minimization and dynamics combined with solvent treatment to arrive at a reasonable structure. Constraints can be relaxed for portions of the DNA; however, the rise should not be, as the intercalation site will collapse without the ligand present. We have developed a simple approach (Gridrun 65) to generate starting structures for new intercalator complexes, as we have found that standard docking programs tend not to reproduce the intercalated ligand location. The only way to sample all possible binding sites is to do a grid search of all possible positions of the ligand in the intercalation site. The grid search in Gridrun translates the ligand in an 8 × 8 ~ (any size can be used) grid on the xz plane (with the helical axis of the DNA aligned to the y axis). In combination with this, full rotation in 15 ° increments (any increment can be used) rotates the ligand around the y axis. This typically generates 2025 structures, although this can be reduced, as the translation and rotation increments can be modified to a user-specified range if additional information is known and combined with chemical or structural intuition. Molecular mechanics minimization using the implicit GB/SA solvation approximation and AMBER94 force field within Macromodel 7.098 (modified to include AMBER charges from the input file) is run on all structures (this can be done with parallel processors) and an energy window of 50 kcal is used to keep possible starting structures. Incorporation of a heavy atom position comparison during the minimization reduces the actual number of models fully minimized, thus decreasing the actual computation time. The region surrounding the intercalation site can be excluded from the calculation by using a frozen shell that further reduces the computational expense. Additional conformational freedom can be included in this approach, such as torsion driving of any substituents on the planer aromatic intercalation component. Monte Carlo searches also can be included for a large

116E Hardenbol,J. C. Wang,and M. W. Van Dyke,Bioconiug. Chem. 8, 617 (1997).

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number of torsion angles of the substituent, although this rapidly becomes computationally expensive. The grid search can be enhanced by molecular dynamics for relaxation of the intercalation complex if a reduced number of starting models is used (for computation expense). In fact, any conformational space search techniques available in Macromode198 can be used. The results of this type of search depend on the quality of the intercalation site. There are a number of other, more sophisticated approaches available for this, but the author has found that this is relatively efficient and removes any doubt about sampling conformational space. Obviously, at this time an explicit solvation method would be time consuming and computationally expensive for a grid search. A selection of the lowest energy models is then used for explicit solvation simulations, using AMBER. 3t As with all starting models, care should be taken in the selection, as this is an approximate method and the user does not want to exclude reasonable models. We have found that explicit solvation is an essential factor as water is an integral structural component of DNA. This method has reproduced the daunomycin intercalation position and has been applied successfully to Z-DNA intercalation, triplex intercalation, and quadruplex intercalation. Generation of Minor Groove-Binding Ligand Starting Models. In general, groove-binding ligand starting models can be generated on the basis of known structures as they typically do not greatly distort DNA and are AT specific, although some prefer GC regions. Some classes of ligands cause a conformation change in the DNA, such as mithramycin-induced B- to A-form transition. A method for the generation of minor groove-binding ligand starting models, similar to that used for the generation of intercalation site starting models, has been implemented by the author. To find the optimal binding site for the ligand, a long sequence containing multiple binding specificities is used and the ligand is simply translated and rotated around the helical axis in the minor groove, and evaluated by molecular mechanics minimization using the implicit GB/SA solvation approximation and AMBER94 force field within Macromodel. 98 This method has reproduced the binding sites of berenil, pentamidine, distamycin, netropsin, and Hoechst 33258, and has been successfully applied to both duplex and triplex minor groove-binding ligands. 9°

Molecular Dynamics Simulation Molecular dynamics simulation has two components: the equilibrium phase and the production phase. There are many different opinions about how to carry out the equilibrium and production phases, but only our chosen method is presented here. In fully solvated systems the equilibrium period is essential for the reorganization and relaxation of the solvent and counterions before the production phase to reduce potentially destabilizing van der Waals and electrostatic interactions. In some cases, if the starting model is not optimal, the DNA : ligand complex should be equilibrated in vacuo prior to addition of solvent, to remove any potentially

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bad contacts. Equilibrating the solvent with the DNA:ligand complex initially restrained reduces artifacts from the interactions of the nonoptimized solvent with the complex. The gradual removal of restraints allows the system to slowly interact fully. The equilibrium period is dependent on each simulation and is typically on the order of 30-100 psec. However, some systems may require longer equilibrium periods; the section Triplex Simulation of Antigene Approach (below) shows an equilibrium period for triplex DNA that required 155 psec, and ionic interactions may require even much longer periods. The systems should be monitored by individual terms of the force field such as bond, angle, dihedral, van der Waals, and electrostatics for anomalies, as well as density, pressure, potential energy, and temperature to establish whether equilibrium has been reached. The production phase is run for the time period required, typically 5001000 psec, and can be done in a variety of ways. This protocol uses the Sander program of AMBER 31 at constant pressure with the PME summation and a time step of 1.5 fsec. The time step is usually varied between 1 and 2 fsec. A shorter time step may be required if coordinate resetting is too large and causes the simulation to stop. Typically the bond lengths or only the hydrogen atom bond lengths are fixed, using the SHAKE method. The equilibrium and production phases should be constantly monitored and not just checked at completion, as every system is different and a number of things can happen that require intervention. Often, molecular simulations are iterative and the process may require modification from the generation of starting models to the production phase.

Evaluation of Simulation Evaluation of the molecular dynamics trajectory will depend on what fundamental question is being asked. In simple cases, if a stable structure for comparison is required, simple time averaging over snapshots near the end of the production run may be sufficient. To compare two structures use of the root mean square deviations (RMSd) of coordinates is common. It should be noted that the average structure is precisely that, an average, and is analogous to crystal structures and NMR average structures. Detailed dynamics may be concealed in average structures. If the dynamics of the system are of interest, monitoring a particular region or specific interactions over the whole production phase is informative, and a variety of parameters may be calculated, such as order parameters from correlation functions, to compare with experimentally derived NMR parameters. There is a wealth of information in the trajectory regarding DNA-ligand interactions and dynamics (nanosecond and subnanosecond range) of the DNA, ligand, DNA : ligand complex, solvent, and counterion interactions with the DNA and ligand. Any three-dimensional quantity can be measured such as hydrogen bonding, van der Waals interactions, and solvent density. There are a number of parameters specific for DNA structure that can be used for comparison. These can be divided into five groups: the global property (helical twist); the backbone torsion angles

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or, r , y, 6, e; and (; the furanose sugar conformation; the glycosidic torsion angle X; and the base helical parameters. The backbone torsion angles are indicative of the form of DNA. The pseudorotation of the furanose ring defines the sugar pucker amplitude and the pseudorotation phase angles also are indicative of the form of DNA, and a population of conformations is not uncommon. The glycosidic angle )~ defines the orientation of the sugar to the base. The helical parameters are defined for the axis (x displacement, y displacement, and inclination), base pair (slide, shift, rise, twist, roll, and tilt), and bases (stagger, stretch, shear, opening, buckle, and propeller twist). The two common programs used for helicoidal parameters are Curves l°l and NEWHELIX, 117 although more recent programs are more mathematically consistent I Js (see Lu et al)19 for a review of nucleic acid analysis programs). Care should be taken when comparing helicoidal parameters that are calculated by different programs, as different definitions of the reference frame may be used. Typically three simulations are needed: the DNA : ligand complex, DNA only, and the ligand only. The energy of binding can be calculated in two ways. If the same number of water molecules was added (manual manipulation of the topology file) to different conformations of the DNA : ligand complexes, a direct comparison of energies is possible. Or more commonly, the systems are stripped of water and minimized, and the simple equation of Ebinding = Ecomplex--EDNA -Eligand is used, although this does not directly take into consideration the hydration energetic component. Average structures distort the water positions as they are moving rapidly compared with the complex. Movies of the trajectories can be made using Moil-view 12° or VMD TM and some imaging may be necessary (see http://www.amber.ucsf.edu/amber/tutorial/ polyA-polyT/analysis.html for guidelines on this). Hydration may be investigated by looking at the radial distribution of water around a region of interest, using Carnal. 31 Also, the water density can be calculated by Watden 122 in order to identify regions with high-density or long-lived water molecules. General Molecular Simulation Protocol

GENERATION OF STARTING MODELS 1. Creation of a three-dimensional model of the ligand: A b initio partial charge calculation (at least at the 6-31 G* basis set level) of the ligand with restricted 117R. E. Dickerson, Nucleic Acids Res. 17, 1797 (1989). 118M. S. Babcock,E. E Pednault, and W. K. Olson,J. Mol. Biol. 237, 125 (1994). 1L9X. J. Lu, M, S. Babcock,and W. K. Olson, J. Biomol. Struct. Dyn. 16, 833 (1999). 120C. Simmerling,R. Elber, and J. Zhang, in "Modellingof BiomolecularStructure and Mechanisms" (A. Pullman, ed.), p. 241. Kluwer, Amsterdam,The Netherlands, 1995. 121W. Humphrey,A. Dalke, and K. Schulten,J. Mol. Graphics 14, 33 (1996), 122C. A. Laughton,Watden,a programto calculate waterdensities from AMBERmoleculardynamics trajectories.

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electrostatic potential charge fit (RESP) and parameterization of the force field (if required) 2. Generation of starting models of DNA complex (Insight II, Macromodel,98 PDB, 103 NDB, 104 NUCGEN ,31 NAB 102) 3. Minimization of starting model in v a c u o with GB/SA solvation 4. Counterion addition: This can be done in at least three ways: (1) counterions added at the most electronegative positions, calculated by solving the PoissonBoltzmann equation (DELPHI123); (2) counterions added as per the rules in the Edit or LEAP program of AMBER; or (3) counterions randomly placed inside the periodic box 5. Periodic box of TIP3P water molecules added, typically 10-A box around the complex (~3500 water molecules) MOLECULARSIMULATION:EQUILIBRIUMPHASE 6. Initial equilibrium of the solvent: a. Minimization, holding DNA complex fixed [100 kcal (mol ./~)-1 restraints, 10,000-step steepest descents] b. Molecular dynamics at 100 K, holding DNA complex fixed [100 kcal (mol- A)-1 restraints, 1.5-fsec time step, 25-psec dynamics] c. Minimization, holding DNA complex fixed [100 kcal (mol./~)-1 restraints, 10,000-step steepest descents] d. Minimization of total system (10,000-step steepest descents) 7. Equilibrium of system: a. Molecular dynamics at 100 K, holding DNA complex fixed [100 kcal (mol. ~)-J restraints, 1.5-fsec time step, 25-psec dynamics] b. Molecular dynamics at 300 K, holding DNA complex fixed [100 kcal (mol. ]k)-I restraints, 1.5-fsec time step, 25-psec dynamics] c. Molecular dynamics at 300 K, gradually reducing the restraints on DNA complex over 100 psec [starting at 100 kcal (mol. ~)-1 restraints going to 0 kcal (mol. A) -I, 1.5-fsec time step, 80-psec dynamics) MOLECULARSIMULATION:PRODUCTIONPHASE 8. Production run: Unrestrained molecular dynamics at 300 K for 1 nsec, using the AMBER98 force field in the isothermal isobaric ensemble (P = 1 atm, T = 300 K), using periodic boundary conditions and the PME algorithm. A 1.5-fsec time step was used with all bond distances frozen using SHAKE for the 1-nsec production run 123B. Honigand A. Nicholls,Science 268, 1144(1995). http://trantor.bioc.columbia.edu/delphi/.

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EVALUATION 9. Sampling of production run and analysis: Time-averaged structures (50 snapshots sampled in the last 50 psec of the production trajectory) with further minimization (10,000 steps steepest descents), helical parameter structural analysis using Curves lm (helical parameters), trajectory analysis for energetics, hydration, and dynamic parameters 10. Calculation of binding energies from the DNA, ligand, and DNA : ligand simulations: Direct comparison of two or more conformations of the ligand in the DNA : ligand complexes E x a m p l e s of M o l e c u l a r M o d e l i n g of Biologically R e l e v a n t D u p l e x , Triplex, and Quadruplex Systems Four examples of modeling biologically relevant DNA : ligand complexes follow, representing duplex, triplex, and quadruplex DNA. The duplex example shows a new DNA ligand that allosterically converts B-form DNA to Z-form DNA, which could lead to a new class of DNA ligands with a novel mechanism. The first triplex example shows the structure of the RNA and 2'-modified RNA : DNA triplex system (and an A-form to B-form conformational transition) and is involved in the structure-based drug design of antigene agents. The second triplex example deals with triplex stabilization and shows how stereochemically and energetically reasonable models can be generated when structural data are unavailable. The quadruplex example examines the stoichiometry of ligands binding to quadruplex systems, as the binding site of these ligands has not been fully elucidated, and is part of a structure-based drug design effort for the development of telomerase inihibitors. Duplex Intercalation in Z-DNA

The binding interactions of (-)-daunorubicin (WP900) (Fig. 1) a newly synthesized enantiomer of the anticancer drug (+)-daunorubicin, with right- and lefthanded DNA have been studied TM quantitatively by equilibrium dialysis, fluorescence spectroscopy, and circular dichroism. (+)-Daunorubicin binds selectively to right-handed DNA, whereas the enantiomeric WP900 ligand binds selectively to left-handed DNA. Further, binding of the enantiomeric pair to DNA is clearly chirally selective and each of the enantiomers was found to act as an aliosteric effector of DNA conformation.124 Molecular dynamics studies using the AMBER suite of programs 3~ resulted in a stereochemically feasible model for the intercalation of WP900 into left-handed Z-DNA. 124X. Qu, J. O. Trent,I. Fokt, W. Priebe, and J. B. Chaires, Proc. Natl. Acad. Sci. U.S.A. 9, 12032 (2000).

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A

309

B

FIG. 1. Structures of (+)-daunorubicin (A) and (-)-daunorubicin, WP900 (B).

The strong preference of (+)-daunorubicin for right-handed, B-form DNA is thought to arise from the precise fit of the daunosamine moiety into the minor groove. 125 High-resolution crystal structures of daunorubicin complexed to DNA shows that the angle of the daunosamine relative to the intercalated chromophore is such that it aligns with the minor groove, allowing a favorable stereochemical fit. 126 Because there are no crystal structures of WP900 bound to Z-DNA, computer simulations using molecular dynamics were applied to build a reasonable model of the complex. Figure 2 shows WP900 intercalated into left-handed DNA. The model is energetically and stereochemically feasible, and provides a detailed view of a left-handed DNA intercalation complex. Simulations of all possible combinations of daunorubicin enantiomers and DNA forms yielded only two stable complexes, one with daunorubicin bound to B-form DNA and another with WP900 bound to Z-form DNA. The B-form DNA : WP900 and Z-form DNA : daunorubicin complexes were not stable and resulted in disruption of duplex stability and loss of intercalation. Such ligand-induced disruptions of the disfavored B- and Z-form duplexes with WP900 and daunorubicin, respectively, may contribute to the allosteric effect by facilitating the backbone conformation change. The simulated B-form DNA: daunorubicin complex (not shown) resembles the published crystal structure.126 The Z-form DNA : WP900 complex (Fig. 2) shows that the AMBER95 force field is able to reproduce the left-handed DNA, as the structure is conserved under the salt conditions used, and the intercalation site is stable. The models reproduce the experimental observations the WP900 preferentially binds to Z-form DNA and daunorubicin preferentially binds to B-form DNA. The preference of WP900 for Z-form DNA arises from the orientation and shape complementarity of the D-daunosamine and the minor J25 j. B. Chaires, J. Biol. Chem. 261, 8899 (1986). 126 A. H. Wang, G. Ughetto, G. J. Quigley, and A. Rich, Biochemistry 26, 1152 (1987).

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FIG.2. Energy-minimizedaveragemodelof the left-handedZ-DNA-WP900intercalationcomplex formed with the d(CGCGCGCG)2duplex. groove. In contrast, the L-daunosamine group of daunorubicin has steric clashes with the minor groove and backbone of Z-DNA, which causes a positional change of the intercalated chromophore that decreases the overall integrity of the binding site. Analogous effects are observed for WP900 and the disfavored B-form DNA. Intercalation of WP900 into Z-DNA is the likely binding mode, a supposition that is supported by our computer simulations using molecular dynamics (Fig. 2), where a stereochemically reasonable WtX)00 : Z-DNA intercalation complex can be formed. Because the same methods accurately reproduce the structure of the

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31 1

B-DNA : daunorubicin intercalation complex, for which high-resolution X-ray data are available, the model derived from molecular dynamics is plausible. We note that a variety of computer simulations have been performed previously in attempts to obtain models for a Z-DNA intercalation complex.127' 12s The methods employed in our studies represent significant improvements over these earlier attempts for several reasons. First, a greatly improved AMBER force field with the PME summation method has been used. Second, our simulations include both solvent and ions, whereas earlier studies modeled structures in vacuo. Finally, reliable highresolution structural data for our system were available as starting models for at least the B family of structures used in our simulations. The average structure obtained from the molecular dynamics simulation for the daunorubicin : B-DNA intercalation complex is consistent with, and essentially reproduces, existing structural data. J26 The Z-DNA simulations are in ~imilar close agreement with existing structural data ~°4 for Z-DNA for the nonintercalated region of the complex. No structural data exist for a Z-DNA intercalation complex. Modeling Protocol. The starting model for daunorubicin was taken from the literature j26 and was inverted to create the WP900 enantiomer. Parameterization and partial charges were obtained from ab initio calculations at the 6-31 G* level with the GAMESS 99 package, using the restrained electrostatic potential fit routine. 114,115 The geometry of the resulting daunorubicin closely resembled the published crystal structure. 126Starting models for the right-handed B and left-handed Z forms of d(CGCGCGCG)2 were created with the standard conformations provided in the Macromodel program. 98 The intercalation site (highlighted in bold face) for the B-form DNA was built by inserting a standard B-form phosphate backbone intercalation site conformation. 126 The Z-form intercalation site (highlighted in boldface) was built by separating the bases of the Z-form of d(CGCGCGCG)2 by 6.8 A with reduction of the helical twist and inserting phosphate backbones, which were subsequently optimized before the intercalator was added. Daunorubicin and wPg00 ligand molecules were then inserted into the generated intercalation site by superimposing the DNA : daunorubicin crystal structure, and four models were used for simulations: B-forms of DNA : daunorubicin and DNA : WP900, and the corresponding Z-forms of DNA : daunorubicin and DNA : WP900. The intercalation sites were optimized by molecular mechanics and dynamics with restraints of 100 kcal (tool - A) i applied to the nonintercalation site bases using the implicit solvation within Macromodel. The optimal positioning of each intercalator was determined by a grid search with translation of 8 × 8 A and rotation ~ 9 0 (using 1 ]~ and 15~ resolution steps). The resulting structures were used as starting structures for the explicit solvation calculations. [27 M. Prabhakaran and S. C. Harvey, Biopolymers 27, 1239 (1988). 128 G. Gupta, M. M. Dhingra, and R. H. Sarma, .I. Biomol. Struct. Dyn. l, 97 (1983).

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Model structures were hydrated using standard A M B E R 5.031 rules in a 10-~ box of TIP3P waters. Sodium cations for the B-form complexes were added, using the Edit placement routine, and chloride anions were added randomly for charge neutrality. The Na + and C1- counterions for the Z-form complexes were added randomly to simulate a neutral 3 M aqueous NaC1 solution. The systems were heated slowly to 300 K and equilibrated carefully during 50 psec with gradual decrease of restraints on the DNA and ligand from 100 to 10 kcal (mol. ~ ) - I . Further 400- and 600-psec periods of restrained molecular dynamics were used for the B-form and Z-form complexes, respectively, to ensure complete solvent equilibrium. Molecular dynamics simulations using the AMBER95 force field were performed in the isothermal isobaric ensemble (P = 1 atm, T = 300 K) with the A M B E R 5.0 program, 31 using periodic boundary conditions and the PME algorithm. A 2-fsec time step was used with all bond distances frozen by using SHAKE. After heating and equilibration, molecular dynamics production runs of 800 psec were used to derive average structures for the complexes taken from 50 snapshots accumulated in the last 50 psec with subsequent minimization. Triplex Simulation o f Antigene Approach

The c-myc gene plays an important role in the regulation of cellular proliferation and differentiation. The c-myc protooncogene encodes a DNA-binding phosphoprotein129 131 that plays an important regulatory role in cell proliferation and differentiation, and is overexpressed in many types of cancer. The c-myc gene product appears to play an important role in the final common pathway regulating cellular proliferation, which makes it an ideal target for transcriptional inhibition. The Miller group has shown that DNA-binding drugs and triplex-forming oligonucleotides inhibit transcription by preventing the binding of regulatory proteins that are necessary for transcriptional activity.132'133 They have developed the molecular, cellular, and whole animal model systems with which to effectively screen for synthetic compounds that inhibit expression of target genes in a sequence-specific manner. They have paid particular attention to factors that influence the intracellular stability of triplex-forming oligonucleotides and that allow formation of triplex DNA by a broader variety of sequences. 134 129E O. Donner, I. G. Wilke, and K. Moelling,Nature (London) 296, 262 (1982). 130M. D. Cole, Cell GrowthDiffer. 65, 715 (1991). J31 K. Kelly and U. Siebenlist, J. Biol. Chem. 263, 4828 (1998). 132V. W. Campbell, D. Davin, S. Thomas, D. Jones, J. Roesel, R. Tran-PaUerson, C. A. Mayfield, B. Rodu, D. M. Miller, and R. A. Hiramoto,Am. J. Med. Sci. 307, 167 (1994). 133D. Chaudharyand D. M. Miller, Biochemistry 34, 3438 (1995). 134N. Vigneswaran, C. A. Mayfield, B. Rodu, R. James, H. G. Kim, and D. M. Miller, Biochemistry 35, 1106 (1996).

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There has been much interest in triplex approaches to antigene therapy. This is primarily due to the potential for durable sequence-specific inhibition of gene expression, as demonstrated by reports showing effective in vitro and cell culture inhibition of transcription of several genes by triplex-forming oligonucleotides. A contributing factor is the Food and Drug Administration (FDA) approval of an antisense oligonucleotide for use in acquired immunodeficiency syndrome (AIDS) patients with cytomegalovirus (CMV), demonstrating that oligonucleotides can be used safely as therapeutic agents. Several approaches to triplex formation and stabilization are being used, including modified oligonucleotides, 135 137 DNA binders, and oligonucleotide-DNA binder conjugates ~38,139that stabilize triplex formation. Use of the pyrimidine motif (pyrimidine third-strand hydrogen bonded via Hoogsteen bonds parallel to the purine strand of the duplex) has not been examined in detail as the cytosine in the third strand must be protonated. One article 14° has shown that the pH requirement of the pyrimidine motif may not be as problematic as previously thought. The NMR experiments unequivocally showed that the pKa of the protonated cytosine residue is higher than physiologic pH for internal positions. The advantage of the pyrimidine motif over the purine motif is that the purine triplex-forming oligonucleotide is always G-rich and, under physiologic concentrations of monovalent cations, especially K + and Na +, the third strand can self-associate and form dimers and tetramers in the presence of these ions, thereby reducing the effective concentration of the third strand available for triplex formation. There are problems associated with all oligonucleotide-based therapies, such as reagent delivery, cellular uptake, and stability (both thermodynamic and enzymatic). Because of these issues, and in-depth understanding of triplex binding is critical in optimizing triplex-forming oligonucleotide design. Nuclease sensitivity of phosphodiester oligonucleotides is a major problem hindering their use as gene-specific compounds in vivo. RNA oligonucleotides with modifications at the T-position of the ribose sugar are more nuclease resistant than unmodified DNA oligonucleotides and are able to form stable triplexes with duplex DNA.141 It has been shown that the most stable RNA :DNA triplexes are

135 j. Michel, G. Gueguen, J. Vercauteren, and S. Moreau, Tetrahedron 53, 8457 (1997). 136 E J. Bates, C. A. Laughton, T. C. Jenkins, D. C. Capaldi, P. D. Roselt, C. B. Reese, and S. Neidle, Nucleic Acids Res. 24, 4176 (1996). 137 S. A. Cassidy, E Slickers, J. O. Trent, D. C. Capaldi, E D. Roselt, C. B. Reese, S. Neidle, and K. R. Fox, Nucleic Acids Res. 25, 4891 (1997). 138 j. Robles and L. W. McLaughlin, J. Am. Chem. Soc. II9, 6014 (1997). 139 A. E Faruqi, S. H. Krawczyk, M. D. Matteucci, and P. M. Glazer, Nucleic Acids Res. 25, 633 (1997). 140 j. L. Asensio, A. N. Lane, J. Dhesi, S. Bergqvist, and T. Brown, J. Mol. Biol. 275, 8 l 1 (1998). 141 A. M. Iribarren, B. S. Sproat, E Neuner, I. Sulston, U. Ryder, and A. L. Lamond, Ptw'. Natl. Acad. Sci. U.S.A. 87, 7747 (1990).

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2'-O-methyl-RNA third strand bound to duplex DNA. 142 The unmodified RNA third strand bound to duplex DNA is less stable that the T-modified RNA but is more stable than the parent DNA triplex. An NMR structure of an intramolecular RNA third strand [r(UCUCUCUU)] : DNA duplex linked by hexakis(ethylene glycol) has been reported by Feigon's group. 143 Another relevant structure has been reported by Lane's group/44 This is of an intramolecular RNA third strand [r(UCUUC)] : DNA duplex linked by hexakis(ethylene glycol), and an intramolecular 2'-O-methyl-RNA third strand [r(UCUUC)] : DNA duplex linked by hexakis(ethylene glycol). However, the structures from the two groups are not in complete agreement with respect to the sugar conformations, base inclination, and backbone parameters. As these are the only two NMR structures deposited, there is an obvious need for more structural information to define the RNA and modified RNA : DNA triplex system. A molecular modeling report 62 used the AMBER force field to calculate 1-nsec molecular dynamics trajectories for the TAT pyrimidine triplex, starting from the A- and B-forms (analogous to A- and B-DNA conformation) and the P-form (analogous to the peptide nucleic acid structure, which can be loosely described as an exaggerated A-form). The structures converged to a single structure that was consistent with the B-form proposed by NMR-derived structures from Patel's group.145'146 This study further demonstrated that high-level simulations can model the A- to B-form transition, as the starting models were in the A, B, and P forms. We have applied molecular dynamics simulations to determine the structure of the RNA and 2'-modifed RNA triplex-forming third strand targeted to the P2 promoter of the protooncogene c-myc. As there was no structure available when this work commenced, we implemented a similar protocol as Shields et al. 62 That is, we started with three models for the sequence r(UUUCUUC) • d(CGAAAGAAGTTTTCTTCTTTCG), the A-form triplex (all three strands in the A form), the B-form triplex (all three strands in the B form), and the AB form (duplex DNA in the B form and the triplex third strand in the A form) (Fig. 3). The target triplex was selected for several reasons: the duplex DNA is part of the murine c-myc P2 promoter 5'-d(AAAGAAG); the target duplex has no contiguous guanine bases, which would require destabilizing contiguous protonated cytosine bases in the third strand; a hairpin duplex decreases the possible number of species in solution

142E J. Bates, J. E Reddock, S. D. Thomas, V. Guarcello, A. Arrow, R. Dale, and D. M. Miller, unpublished results (2000). 143 C. H. Goffredgen, E Shultze, and J. Feigon, J. Am. Chem. Soc. 120, 4281 (1998). 144 j. L. Asensio, R. Carr, T. Brown, and A. N. Lane, J. Am. Chem. Soc. 121, 11063 (1999). 145 I. Radhakrishnan, X. Gao, C. de los Santos, D. Live, and D. J. Patel, Biochemistry 30, 9022 (1991). 146 I. Radbakrishnan and D. J. Patel, Structure 2, 395 (1994).

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A-form

AB-form

315

B-form

FIG.3. Startingmodels for the moleculardynamicsstudies of the RNA : DNA triplex systems. and aids in the assignment of the resonance signals of the duplex. The T4 loop was designed from the literature, 147 and dCG was added to the 5' site to give enhanced stability by avoiding AT fraying and to give information on the triplex-duplex boundary. The nanosecond trajectories, after 155 psec of equilibrium, for the three starting models converge to a single AB-hybrid form (distinct from the starting AB form) (Fig. 4, Tables I and II). This shows again that the modeling protocol has allowed transition of the A-form DNA duplex to the more stable B-form DNA duplex (the AB-form triplex has an underlying B-form duplex). The Feigon structure t43 generally compares well with the modeled structure, with subtle differences in the sugar conformations, base inclination, and backbone parameters. The converged model is in much closer agreement with the Lane structure. 144 It should be noted that the two most recent RNA third-strand structures are not the same in the detailed analysis and there may be some sequence dependence. It has been shown that the most stable RNA : DNA triplexes are T-O-methylmodified RNA third strand to duplex DNA. 142 A surface plasmon resonance technique (BIAcore, Uppsala, Sweden) also has been used 142 to examine the real-time association and dissociation between a 24-mer triplex-forming oligonucleotide and a 30-bp hairpin duplex representing the murine c - m y c P2 promoter. These experiments showed that both the association and dissociation of the triplex were slower for a 2'-O-methyl-RNA third strand compared with its RNA analog. We undertook a similar modeling approach as with the RNA : DNA triplex for the T-O-methylRNA : DNA triplex. 147Z. Kuklenyik,S. Yao,and L. G. Marzilli,Eur. J. Biochem. 236, 960 (1996).

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FIG.4. The final converged average AB-form structure for the RNA : DNA triplex systems. Starting from the A, AB, and B forms, the A and AB forms converged to a single AB-form structure. However, the B form did not converge and the base triplets were disrupted because of the unfavorable steric clash of the T-O-methyl group with the backbone hindering the B-to-A transition. It should be noted that the ends of the triplex regions frayed, which is consistent with DNase I footprinting data 142 of the longer RNA triplex formed at the c-myc P2 promoter sequence. The calculated order of stability for the modeled triplexes is DNA triplex < RNA : DNA triplex <2'-O-methyl-RNA : DNA triplex, which accords with BIAcore data that the rate of dissociation of the 24-mer triplex-forming oligonucleotide is DNA > RNA > 2'-O-methyl-RNA. The RNA : DNA and 2'-O-methylR N A : D N A triplexes were similar in overall helical parameters. However, the hydration patterns were different, although the regions of high water density reproduced the spine of hydration in the duplex minor groove in both triplex models.

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TABLE I GROOVE WIDTHSOF MODELED STARTINGAND FINAL STRUCTURESFOR RNA : DNA TRIPLEX a

Starting structureh

Production dynamics structure

DNA-RNA form

Major

Minor

Major

Minor

A form AB form B form TAT DNA triplex a Feigon RNA-DNA triplex e

10.5 12.5 15.2

10.5 6.2 6.7

18.8 18.2 18.4 18.9 18.2

6.2 6.1 6.2 6.8 6.2

Time of production run" (psec) 1000 1000 1000 1000

Major and minor groove widths (for the duplex DNA of the triplex) measured in angstroms (A) using phosphotus atoms (curves; R. Lavery and H. Sklenar, ,I. Biomol. Struct. Dyn. 6, 63 ( 1988); http://www.ibpc.fr/UPR9080/Curindex.html). l, Generated from the literature. c After equilibrium period of 155 psec with warming from 0 to 300 K and gradual removal of restraints on the DNA and RNA to allow full equilibrium of the counterions and solvent. ~/From G. C. Shields, C. A. Laughton, and M. Orozco, J. Am. Chem. Soc. 119, 7463 (1997). e From C. H. Gotfredgen, P. Shultze, and J. Feigon, J. Am. Chem. Soc. 120, 4281 (1998).

TABLE II PREDOMINANTSUGAR CONFORMATIONSOF DNA AND RNA STRANDSa FOR MODELED STARTINGAND FINAL STRUCTURESOF RNA : DNA TRIPLEX Starting structure DNA-RNA form A form AB form B form Feigon grouph Lane group~'

Final structure

Purine

Pyrimidine

RNA third strand

C3t-endo C2~-endo C21-endo

C3t-endo C2~-endo CT-endo

C3'-endo C3'-endo CT-endo

Pyrimidine

RNA third strand

Production run time (psec)

Cl'-exo Cl'-exo Cl%xo

Cl'-exo Cl'-exo Cl~-exo

C3'-endo C3'-endo C3'-endo

1000 1000 1000

Cll-exo

Cl%xo/O'endo Cl'-exo

C31-endo

Purine

Cl'-exo

C3%ndo

a Purine is GAAAGAAG, pyrimidine is CTTCTTTC of the hairpin d(CGAAAGAAGTTrTCTI'CTI'TCG). h From C. H. Gotfredgen, P. Shultze, and J. Feigon, J. Am. Chem. Soc. 120, 4281 (1998), average sugar puckers; the sugar puckers vary in this structure and seem to be sequence dependent. "From J. L. Asensio, R. Carr, T. Brown, and A. N. Lane, J. Am. Chem. Soc. 121, 11063 (1999).

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FIG. 5. The final converged average AB-form structure for the 2'-O-methyI-RNA: DNA triplex system. The regions of high water density are shown as white cnntours. The water density shows that there are few long-lived water molecules in the R N A - p u r i n e groove, but the water is ordered in the 2'-O-methyl-RNA-purine groove (Fig. 5). The increased stability and ordered hydration of the T-O-methylRNA : DNA triplex may explain the BIAcore data showing that 2'-O-methyl-RNA triplex-forming oligonucleotide associated more slowly than the RNA analog, as the solvation shell of the duplex would have to rearrange to an ordered state on binding of the third strand, thus taking longer for association to occur, and similarly the dissociation may be slower because of the disruption of the ordered environment. Modeling Protocol. The 2'-O-methylguanine, uracil, and protonated cytosine bases built in Macromodel were optimized with the 6-31G* basis set, using G A M E S S , 99 and the RESP j 14,115 charges were derived. Standard RNA bases from

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AMBER 31 were used for r(G). The triplex in A fornl, 148 B form, I°7 and AB hybrid form l°s were generated according to the literature. The T4-1oop conformation was optimized by a Monte Carlo search on rotatable torsion angles within Macromodel, 98 using the AMBER94 force field and GB/SA solvation. Simulations were run for the A, B, and AB forms of both the RNA and 2'-O-methyl-RNA triplexes, B-form duplex, B-form DNA triplex, RNA third strand, and 2'-O-methylRNA third strand, using the methods below. Models were hydrated in a 10-~ box of TIP3P waters according to standard AMBER31rules; Na + cations were added by CION, and then CI- counterions were placed randomly for charge neutrality. Box sizes were adjusted to include 3667 waters for each model. Simulations were performed in the isothermal isobaric ensemble (300 K, 1 atm) with the AMBER 5.0 program 31 and parameters from parm96.dat, using periodic boundary conditions and the PME algorithm. Molecular dynamics simulations used the Sander routine ( 1.5-fsec time step), with SHAKE to freeze all bonds involving hydrogen. Initial equilibrium for 155 psec following the general protocol above (Triplex Simulation of Antigene Approach) was performed. The production runs were 1 nsec in length and average structures for each complex (taken from 50 snapshots accumulated in the last 50 psec) were obtained. The waters were stripped from the average structures that were subsequently fully minimized to give the final structures. Helical parameter analysis used Curves 5.1.101 Triplex Intercalation

A current investigation into new DNA ligands is an example of how multidisciplinary investigations can provide insight into triplex intercalation. In collaboration with J. B. Chaires (see [ 1] in this volume 148a ), molecular modeling is rationalizing data from the dialysis assay. By screening compounds through the assay it was found that ~- and/3-naphtholflavone preferentially bind to TAT parallel triplex, and do not appreciably bind to duplex DNA. The o!-naphtholflavone binds 4-fold tighter to triplex than does the fl isomer. The molecular models reproduce these data and rationalize this new class of triplex intercalator. It is assumed that the ligand binds as a triplex intercalator, as there is no duplex binding and the minor grooves of duplex and triplex are similar (although not identical). It is highly unlikely that an aromatic compound would bind in the major groove, and the planar aromatic portion of the molecule seems likely to intercalate. The simulations show that there are three structures that are sterically and energetically reasonable for c~-naphtholflavone and three structures for/~-naptholflavone (Fig. 6). The difference in binding free energies is being investigated. 148S. Arnott, P. J. Bond, E. Selsing, and P. J. C. Smith,Nucleic Acids Res. l l, 4141 (1976). 14Saj. B. Chaires,Methods Enzymol. 340, [1 ] 2001 (this volume).

320

[ 141

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A

13

12

I1 FIG.6. The sterically and energeticallyreasonable models for (A~2) ot-naphtholflavoneand (D-F) /3-naphtholflavonebound to the pyrimidine d(Ts) •d(As)d(T8) triplex. ~- and fl-Naphtholflavoneare shown in black, with only the d(Ts), d(As)d(T4)triplet shown for clarity.

Modeling Protocol. The intercalation site was built with standard DNA intercalation site backbones 126 and inserted into the B-form TAT triplex model [d(Ts)-d(As)d(Ts)] and the backbones were optimized by molecular mechanics and dynamics (minimization, dynamics equilibrium period of 50 psec, production period of 300 psec, with the average structure from the last 25 psec using 25 snapshots, being minimized and used subsequently) using Macromode198 with AMBER94 and GB/SA solvation. Gridrun 65 was used to identify possible binding sites for the oe- and /3-naphtholflavone in the intercalation site. As a - and /3-naphtholflavone are asymmetric, a 180 ° rotation around the z axis was added. Four and three positions were identified for ~- and/3-naphtholflavone, respectively, within 10 kcal. These were fully solvated in a 10-]k box of TIP3P water and the general simulation protocol (Triplex Simulation of Antigene Approach) was used with 85-psec equilibrium and 1-nsec production runs with a 2-fsec time step. Quadruplex Intercalation Porphyrin [H2TMPyP: 5,10,15,20-tetrakis( 1-methyl-4-pyridyl)-21H-porphine] binding to DNA sequences from the Oxytricha and human telomeres and a thrombin-binding aptamer have been examined 97 by isothermal titration calorimetry (ITC) and spectrophotometry under conditions that favor self-assembly to their respective intermolecular or intramolecular quadruplex structures. Analysis of the ITC and optical data reveals (1) saturating porphyrin/quadruplex binding stoichiometries of 1 : 1, 2 : 1, and 3 : 1 for d(G2T2GzTGTG2TzG2), d(AG3[T2AG3]3), and [d(T4G4)]4, respectively, involving near-equivalent sites, (2) weak binding of

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only (0.3-2) x 105 M-1 per site, and (3) no evidence of stepwise complexation in solutions containing K + ions. This stoichiometry is maintained in Na + solutions, if the quadruplex is stable, but nondegenerate sites are implicated for the 2 : 1 and 3 : 1 complexes, where the first porphyrin binds with 20- to 40-fold greater affinity than any subsequent ligand(s). Importantly, the stoichiometries correspond to the number of intervals between successive G-tetrad planes in each quadruplex. The results indicate binding by threading intercalation at each closely similar GpG site, possibly without invoking neighbor exclusion for adjacent sites, rather than through either external electrostatic processes or end-pasted stacking modes. This mechanism is supported by dynamic molecular modeling simulations with two DNA quadruplexes, which show that stable intercalated complexes can be realized. A plausible model is developed that accounts for the occupation of adjacent sites, where unfavorable interligand contacts are avoided by phased asymmetric positioning of the porphyrin molecules within each G-tetrad intercalation pocket. Molecular dynamics studies of intercalated complexes support the conclusions from the biophysical data for binding. Two distinct DNA systems were selected to examine possible intercalative modes for the HzTMPyP ligand, involving (1) 1 : 1 binding to the folded intramolecular d(GzTzGzTGTG2T2G2) quadruplex, and (2) 1 : 1 and 3 : 1 binding to the intermolecular tetrameric [d(TGGGGT)]4 quadruplex. The latter parallel system was used to enable a comparison with the G4 structure, as structural data are unavailable for an intermolecular quadruplex with blunt-end strands (see modeling protocol, above). Energy minimization resulted in stable structures for all complexes examined with the DNA quadruplexes. A major finding is that the planar HzTMPyP ligand molecule can be intercalated within the [d(TGGGGT)]4 and d(G2T2G2TGTG2T2G2) structures to stabilize the complex through favorable 7r-stacked interactions between aromatic residues, as for "classic" duplex intercalation, but without significant disruption of the guanine tetrads. Figure 7A shows the stable 1 : 1 core-intercalated complex with the [d(TG4T)4]4 quadruplex, 5'-TGG*GGT, after refinement as described. Examination of the model shows that the ligand adopts a parallel alignment between the successive tetrad planes with the molecule displaced asymmetrically from the DNA helical axis. The ligand is offset so that two of the noncoplanar 1-methylpyridinium groups effectively protrude into adjacent groove conduits of the quadruplex. As a consequence, the two other substituent rings do no extend as far into their respective DNA grooves and are drawn toward the intercalation site, thereby inducing a minor perturbation of the local tetrad planarity. This result suggests that the extended planar ring system of the porphyrin is actually smaller than a G-tetrad and hence cannot effect a simultaneous equivalent rr-stabilization of all four guanine bases in the array. Sliding-type rearrangement or dynamic relocation of the porphyrin would be expected to stabilize the entire tetrad in each plane, although this was not seen in the production-run time scale (400 psec) used for our simulations.

322

BIOPHYSICALAPPROACHES

[ 141 St

5!

T

T

G

G G

G

G

G

G G

T ,

T

FIG. 7. Modeled structures for possible intercalation of the [d(TG4T)]4 tetraplex with (A) 5'TGG*GGT (1:1 binding) and (B) 51-TG*G*G*GT (3:1 binding), where each asterisk denotes an H2TMPyP ligand. The complexes are viewed by looking toward the G-tetrad stack with the porphyrin(s) shown in space-fill mode. Note the axial offset of the porphyrin relative to the long helical axis in the 1 : 1 core-intercalated complex, and the alternating pattern of ligand displacements in the 3 : l complex.

The binding process requires a change of local DNA conformation to accommodate the porphyrin, so that the phosphate backbone is linearized relative to that of the B-type DNA host and the tetrad planes implicated are rotated to a more superimposable alignment (Fig. 7B). However, such conformational effects are not propagated extensively, as the overall integrity of the quadruplex is retained and the structure remains stable throughout the dynamic simulation. The core-intercalated 5'-TGG*GGT model complex is well behaved and remains stable after inclusion of further porphyrin molecules between the adjacent stacked G-tetrads to achieve higher stoichiometric ratios. Figure 7B shows the 3 : 1 complex 5'-TG*G*G*GT after minimization, showing the axial or slipped displacement of each HzTMPyP ligand and the alternating groove protrusion for each successive ligand. Such asymmetric binding results in an effective removal of interligand contacts and is suggested to diminish the influence of neighbor exclusion on binding stoichiometry and/or increased ligand toad. The structure for the 3 : 1 complex illustrates the binding-induced linearization afforded to the DNA backbone and confirms and effective coplanarity and retention of stacking for both the ligand and G-tetrad systems. Further molecular modeling is in progress to examine alternative end-stacked or end-pasted arrangements (e.g., 5'-T*GGGGT and 5'-TGGGG*), and to establish the influence of flanking 5' and 3' bases on the stability of the complexes.

[ 14]

MOLECULAR MODELING OF D R U G - D N A COMPLEXES

323

More extensive simulations were carried out for the three possible 1 : 1 intercalated d(GGTTGGTGTGGTTGG)-H2TMPyPcomplexes to probe site-dependent binding effects. The averaged trajectory energies after molecular dynamics equilibration showed that the 5'-G*GTT model for intercalation of the core G-tract is more stable than either alternative end-pasted 5'-*GGTT or intercalated ("endstacked") 5'-GG*TT models involving the different loop regions of the host G2 quadruplex. The time-averaged structure for each complex was stripped of water and counterions, and then subjected to a full refinement procedure (see modeling protocol, above). Comparison of energies for the minimized structures revealed that the 5'-G*GTT complex (Fig. 8B) is considerably more favorable than the 5'-*GGTT (Fig. 8A) or 5'-GG*TT (Fig. 8C) complexes, with relative energies of 0, + 140, and +250 kcal mol- 1, respectively. This ranking order reflects differences in re-overlap for the G-tetrad and porphyrin ring systems, disturbance of favorable 5'/3'-end stacking from the loop thymines, and disruption of the guanine quartets by the porphyrin substituents due to asymmetric positioning of the ligand (see above). Thus, the H2TMPyP molecule in the 5'-GG*TT complex (Fig. 8C) moved considerably from the initial docked position to give a structure where zr-overlap is achieved with only two guanines in the adjacent tetrad. Such a lateral displacement results in spear rather than full intercalation in the TT loop region, and hence a poorer binding energy. In contrast, significantly greater Jr-overlap is achieved for the 5'-*GGTT complex involving the less sterically crowded TGT loop region of the structure. Taken together, these modeling results support the conclusions from the ITC and optical data, and demonstrate that intercalation of both inter- and intramolecular quadruplexes can result in stable DNA-porphyrin complexes. Further studies are required to assess the effects of increased ligand load (i.e., higher stoichiometry) on the site specificity and structural integrity. To this end, we examined the stabilization of the folded G3 human quadruplex by a series of isomeric anthraquinones, using a less extended modeling protocol.95 This study culminated in a model for complexation with the ligand intercalated at the 5'-A*G step that accounts for the 1 : 1 binding stoichiometry established using ITC. Given that we find a 2 : 1 stoichiometry for porphyrin binding to the same G3 structure in both K + and Na + buffer, it is likely that factors such as sequence context and ligand complexion may govern the site specificity and/or ultimate stoichiometry. This conclusion is supported by the distinct stoichiometries established from an NMR binding study of a perylene ligand to intermolecular quadruplexes with blunt-ended or capped G-tract strands, resulting in 5'-TTAGGG* and 5'-(T)TAGGG*TT(A) complexes, respectively.149 The bulky substituents attached to the planar chromophore of this ligand would be predicted to influence any binding mechanism where groove accommodation is compromised. 149 O. Y. Fedoroff, M. Salazar, H. Han, V. V. Chemeris, S. M. Kerwin, and L. H. Hurley, Biochemistry 37, 12367 (1998).

324

BIOPHYSICALAPPROACHES

[ 14]

M o l e c u l a r P r o t o c o l . Both intramolecular (G2) and intermolecular DNA quadruplex systems were selected for molecular modeling and energy minimization. Starting models were built with DNA coordinates reported for the chair-type folded NMR structure15° of d(GzTzG2TGTG2T2G2) and the crystal structurelSl of [d(TG4T)]4 (PDB 103 entries 1QDF and 244D). The intermolecular parallel quadruplex was used to construct G4-type models as no structure is available for a bluntended DNA system. As the 5' and 3' thymine bases are probably involved in a zr-stacked stabilization 151,152 of the terminal G-tetrads in the [d(TG4T)]4 system, these flanking bases were included in the 5'-TGGGGT models, using standard B-DNA geometries. Intercalation sites were incorporated for each quadruplex by using phosphodiester backbone parameters taken from published structures for intercalated DNA duplex-ligand complexes. A constructed model for the HzTMPyP tetracation was optimized by the A M B E R 5.0 program, 31 with appropriate forcefield parameters and partial charges obtained from a b initio calculations at the 6-31G* level with the GAMESS package 99; the restrained electrostatic potential fit routine j14"115 was used. The geometry of the resulting porphyrin closely resembled the published crystal structure.153 On the basis of the binding results obtained, 97 two porphyrin-G4 models were evaluated to examine possible intercalative modes: (1) the I : 1 complex TGG*GGT with a single, centrally positioned ligand (denoted by an asterisk) within the G-tract core, and (2) the 3 : 1 complex TG*G*G*GT, where a ligand is accommodated between each pair of adjacent tetrad planes. In contrast, models for three distinct 1 : 1 complexes were evaluated for the folded G2 structure to compare the relative site stabilities for core- and loop-intercalated or "end-pasted" modes, denoted as 5'-*GGTT,5'-G*GTT, and 5'-GG*TT (Fig. 8). Each H2TMPyP ligand molecule was positioned in starting models to effect greatest re-overlap between the aromatic rings of the ligand and adjacent G-tetrad(s) with each DNA, but with minimal contact to bases in either loop region for the G2 complexes. Models were hydrated in a 10-A box of TIP3P waters according to standard rules; Na + cations were added, and then C I - counterions were placed randomly for charge neutrality. Box sizes were adjusted to include 2790 waters for each model. Simulations were performed in the isothermal isobaric ensemble (300 K, I atm) with the A M B E R 5.0 program 31 and AMBER95 force field, using periodic boundary conditions and the particle mesh Ewald algorithm. Molecular dynamics simulations used the Sander routine (1.5-fsec time step), with SHAKE to freeze all

15°V. M. Marathias, K. Y. Wang, S. Kumar, T. Q. Pham, S. Swaminathan, and P. H. Bolton,J. Mol. Biol. 260, 378 (1996). 151G. Laughlan, A. I. Murchie, D. G. Norman. M. H. Moore,E C. Moody,D. M. Lilley, and B. Luisi, Science 265, 520 (1994). 152K. Phillips, Z. Dauter, A. I. H. Murchie, D. M. J. Lilley,and B. Luisi,J. Mol. Biol. 273, 171 (1997). 153K. G. Ford, L. H. Pearl, and S. Neidle, Nucleic Acids Res. 15, 6553 (1987).

[14]

M O L E C U L AMODELING R OF DRUG-DNA COMPLEXES

325

i

/

e-

~

i

~",.. i"..:: X .~

'4 4~.

e.,

m ,...= .~ ~

-

t¢3

.~

0

.

=

0 .r.° ....~Z/b--• m

~

[~

'; ~ .__,8~

i ..... X

~"... \ ,~_j_~

",,;~.~ y

.,.-,

,.... ~

~.~ . ~

.-

~

~.;~ ~ .r, .~

--I-

/

l'-

!--..

X

"~

~-i-~-~-\

N =0 "a ~ A =~

d~

om'-

°

A~

["

~

o

326

BIOPHYSICALAPPROACHES

[ 14]

bonds involving hydrogen. All calculations were performed on a Silicon Graphics Origin 200 server and an NPACI Cray T3E (San Diego Supercomputer Center, San Diego, CA). The general modeling protocol (Triplex Simulation of Antigene Approach, above) with 150-psec equilibrium and additional 200-psec hydrogenbonded restraints [initially 20 kcal (mol. A)- l, with gradual linear removal] was used to maintain the G-tetrad conformation and to prevent artifacts from the choice of starting model. After equilibration, production runs of 400 psec were used to obtain an average structure for each complex (taken from 100 snapshots accumulated in the last 100 psec), which was then fully minimized to give the final structure. Conclusions The advances in molecular modeling have been significant. We are able now to reproduce several forms of DNA in various applicable experimental conditions and observe similar hydration and ion associations in some systems. It is hoped that this is just the beginning for reliable simulations that can rationalize experimental observations and then can be used to predict and design new experiments. With increasing confidence in the methods and results it will enable the application of molecular modeling to many biologically relevant problems. Acknowledgments The author thanks his collaborators on DNA projects for their contributions and multidisciplinary talents: Paula J. Bates, Jonathon B. Chaires, Terence C. Jenkins, Donald M. Miller, Waldemar Priebe, Nadarajah Vigneswaren, and Wolfgang Zacharias. The author is grateful to Paula J. Bates and Diane Konzen for reading this chapter and for helpful suggestions, to Thomas Cheatham III for supplying preprints of his work, and also to Nick Peiper and Nina Mfihr for typing references.

[1 5]

PNA TARGETINGOF DOUBLE-STRANDEDDNA

329

[ 15] Peptide Nucleic Acid Targeting of Double-Stranded DNA B y PETER E. NIELSEN

Introduction The DNA mimic PNA (peptide nucleic acid) was originally conceived and designed as a DNA major groove-binding ligand mimicking triple helix-forming oligonucleotides. 1 Chemically, PNA is composed of a polyamide (pseudo-peptide) backbone consisting of N-(2-aminoethyl)glycine units to which nucleobase acetic acid ligands are connected to the glycine nitrogen via amide bonds 1-3 (Fig. 1). Therefore in a chemical sense PNA oligomers are much more closely related to peptides and proteins than they are to oligonucleotides.4 Structurally, however, PNA oligomers are good mimics of DNA or RNA and they form tight helical duplexes with sequence complementary DNA, RNA (or PNA) oligomers. 5 v These duplexes are generally more stable than the corresponding natural nucleotide complexes. Interestingly, duplexes containing mostly purines in the PNA strand are much more stable than the "mirror" duplexes having the most purines in the DNA strand. 8 Indeed, an empirical formula for calculation of the thermal stability (Tin) of a P N A - D N A duplex based on its sequence and the Tm calculated for the corresponding D N A - D N A duplex has been derived9: Tm,pred = 20.79 + 0.83Tm,nnDNA -- 26.13fpyr q- 0.44L in which, Tm,,nDNAis the Tm as calculated by using a nearest neighbor model for the corresponding D N A - D N A duplex, applying AH ° and AS ° values as described by SantaLucia et al. 1° and without taking end effects into account, fpyr denotes the fractional pyrimidine content, and L is the PNA sequence length in bases. The above-mentioned asymmetry is reflected by thefpyr factor in the equation.

I R E. Nielsen,M. Egholm, R. H. Berg, and O. Buchardt,Science 254, 1497 (1991). 2 M. Egholm, O. Buchardt, P. E. Nielsen,and R. H. Berg,J. Am. Chem. Soc. 114, 1895 (1992). 3 B. Hyrupand P. E. Nielsen,Bioorg. Biomed. Chem. 4, 5 (1996). 4 p. E. Nielsen,Acc. Chem. Res. 32, 624 (1999). 5 M. Egholm, O. Buchardt,L. Christensen,C. Behrens, S. M. Freier, D. A. Driver, R. H. Berg, S. K. Kim, B. Nord6n, and P. E. Nielsen,Nature (London) 365, 556 (1993). 6 K. K. Jensen,H. t0rum,P. E. Nielsen,and B. Norden,Biochemistry 36, 5072 (1997). 7 p. Wittung,P. E. Nielsen,O. Buchardt,M. Egholm,and B. Nord6n,Nature (London) 368, 561 (1994). 8 p. E. Nielsenand L. Christensen,J. Am. Chem. Soc. 118, 2287 (1996). 9 U. Giesen,W. Kleider,C. Berding,A. Geiger, H. Orum, and P. E. Nielsen,Nucleic" Acids Res. 26, 5004 (1998). l0 j. SantaLucia,H. T. Allawi,and P. A. Seneviratne,Biochemistry 35, 3555 (1995).

METHODSINENZYMOLOGY,VOL.340

Copyright~3~2001byAcademicPress Allrightsofreproductioninanyformreserved. 0076-6879/00$35.00

330

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[ 15]

B

0

B O

"'"

0

0.~/

(5'-)

0-

o

(~o.~/

o...

DNA B

B

B

Lro Lro Lro

v,- terminal', '~'

"0

0

0

PNA FIG. 1. Chemical structures of DNA and PNA. B, One of the four bases (adenine, cytosine, guanine, thymine).

However, complexes formed by homopyrimidine PNA oligomers and sequence-complementary homopurine DNA (or RNA) oligomers are far more stable than would be predicted from the preceding equation. In fact, they are more stable than even the most stable duplexes. This is due to the formation of PNA2-DNA triplexesfl These are fully analogous to traditional DNA triplexes, and similarly show highest stability in an antiparallel PNA-DNA Watson-Crick configuration and a parallel PNA-DNA Hoogsteen configuration.l l Four different binding modes on targeting double-stranded DNA with PNAs have been described (Fig. 2). These include conventional triplex binding in the major groove of the DNA double helix (Fig. 2A) and triplex invasion in which two preferably antiparallel PNA oligomers invade and open this DNA double helix through formation of an internal PNA-DNA-PNA Watson~Crick-Hoogsteen triplex (Figs. 2B and 3). Duplex invasion through an internal PNA-DNA WatsonCrick duplex (Fig. 2C) has also been observed under certain conditions, and if special modified pseudo-complementary bases, such as the diaminopurine-thiouracil base pair (see Fig. 5), are employed in the PNA oligomers, binding to the target

1l M. Egholm, L. Christensen, K. Dueholm, O. Buchardt, J. Coull, and P. E. Nielsen, Nucleic Acids Res. 23, 217 (1995).

[ 15]

PNA TARGETING OF DOUBLE-STRANDED D N A

331

m

l.."

N m

Ell mien men mien mien mimm mimmm mm

mien

4

men

D Triplex

Triplex Invasion

(A)

(B)

Duplex Invasion

(C)

I Double Duplex Invasion

(D)

FIG. 2. (A-D) Schematic drawing of the binding modes on targeting double-stranded DNA by PNA. The ladder symbolizes the DNA double helix, and the PNA oligomer is drawn in boldface.

by double duplex invasion is possible (Fig. 2D). Not surprisingly, these complexes form under different conditions and have different properties. Triplex B i n d i n g Although PNAs were originally conceived as major groove triple helix-binding reagents, this binding mode does not seem to yield high-affinity complexes with PNA oligomers. "Conventional" PNA-DNA2 triplexes have been observed in a few cases. ]2,13 They seem to require a high cytosine content and are less stable than the corresponding triplex invasion complexes. However, not much information about such PNA-DNA2 triplexes is yet available. Triplex I n v a s i o n It came as a major surprise that the preferred binding mode of homopyrimidine PNA oligomers when targeting double-stranded DNA turned out to be a novel triplex invasion complex. ], ]4 The relative readiness by which these complexes are formed, combined with their remarkable stability and their potentially exciting utility in molecular biology and medical applications, have made these complexes 12 D. Praseuth, M. Grigoriev, A. L. Guieysse, L. L. Pritchard, A. Harel-Bellan, E E. Nielsen, and C. Helene, Biochim. Biophys. Acta 1309, 226 (1996). 13 E Wittung, R E. Nielsen, and B. Norden, Biochemistry 36, 7973 (1997). 14 p. E. Nielsen, M. Egholm, and O. Buchardt, J. Mol. Recognit. 7, 165 (1994).

332

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[ 15]

CH 3 CH 3 L ~

j

__~

)~0, O

,O,

H,

N

..,N

N

!

T A 1,1

"~,.--N Off

c+

H

-,. H

//

II

"0"

..-" ; .,,-"

j"

1-I

G

I

N ~ N %

c

Watson-Crick

FIG. 3. Chemical structures of T-A-T and C+-G-C triplets formed via Watson-Crick and Hoogsteen base pairing.

the subject of intensive study. 15-2° A triplex invasion complex requires two homopyrimidine PNA strands binding to the homopurine strand of the DNA target. The one PNA strand recognizes the target by Watson-Crick base pairing whereas the other relies on Hoogsteen base pairing. Thus the complex is based on conventional T-A-T and C+-G-C tripletsl4 (Fig. 3). Consequently, for guanine-containing targets strong binding requires low pH for efficient protonation of cytosine in full

15 D. Y. Chemy, B. R Belotserkovskii, M. D. Frank-Kamenetskii, M. Egholm, O. Buchardt, R. H. Berg, and P. E. Nielsen, Proc. Natl. Acad. Sci. U.S.A. 90, 1667-1670. 16j.-H. Kim, K.-H. Kim, N. E. M011egaard, P. E. Nielsen, and H. S. Koo, Nucleic Acids Res. 27, 2842-2847. 17T. Bentin and P. E. Nielsen, in "Triple Helix Forming Oligonucleotides" (C. Malvy, A. Barel-Bellan, and L. Pritchard, eds.), pp. 245-255. Kluwer Academic Publishers, Dordrecht, The Netherlands. 18H. J. Larsen and P. E. Nielsen, in "Peptide Nucleic Acids: Protocols and Applications" (IF'.E. Nielsen and M. Egholm, eds.), pp. 221-240. Horizon Scientific Press, NorfoLk, UK. 19 p. Wittung, P. E. Nielsen, and B. Nordrn, J. Am. Chem. Soc. 118, 7049 (1996). 20 H. Kuhn, V. V. Demidov, P. E. Nielsen, and M. D. Frank-Kamenetskii, J. Mol. Biol. 286, 1337 (1999).

[15]

PNA TARGETINGOF DOUBLE-STRANDEDDNA H

333

H

H,, "0"" I

.,,,," "-,~',~ i

H

G FIG. 4. Chemical structures of diaminopurine-thymine, diaminopurine-thiouracil, adeninethymine, and adenine-thiouracil base pairs, illustrating the steric clash in the diaminopurine-thiouracil pair, which is not present in the adenine-thiouracil pair.

analogy to triplex-forming oligonucleotides. It was realized early on that antiparallel binding is preferred for the Watson-Crick PNA-DNA duplex, 5' 11 whereas parallel binding is preferred for the Hoogsteen-bound PNA.I 1 Thus the construction of"bifunctional" bis-PNAs joining both PNA strands by continuous synthesis was logical. 11'2x Such bis-PNAs were further improved by replacing cytosines in the Hoogsteen strand by pseudoisocytosine ("J"), which is a pH independent mimic of N3-protonated cytosine (Fig. 4), 11 and by including positively charged units, such as lysines, either in the linker between the two PNA strands or (better) at the carboxy and/or the amino-terminal end of the bis-PNA. 22 Positively charged PNAs bind to their double-stranded DNA (dsDNA) targets with greatly increased on rates and correspondingly enhanced efficiency.22 Although no systematic studies have yet been performed on the influence of PNA length and target sequence composition, it has been found that bis-PNAs as short as heptamers (2 × 7) form stable triplex invasion complexes with their cognate dsDNA target.Z3(Bis)decamers have been most extensively studied and shown to exhibit extraordinary stability once formed. Half-lives of these complexes have not been accurately measured, but are counted in (several) days at 37°. 21 Therefore under normal experimental conditions binding of PNA (decamers) to dsDNA targets is kinetically controlled and does not reach equilibrium. Consequently, it is neither productive nor really relevant to describe the DNA binding of these PNAs by an affinity constant (Ka = kon/kon), but rather by the relative on rate (kon) instead. 24 It might be feared that this type of binding could result in poor 21 M. C. Griffith, L. M. Risen, M. J. Greig, E. A. Lesnik, K. G. Sprankle, R. H. Griffey, J. S. Kiely, and S. M. Freier, J. Am. Chem. Soc. 117, 831 (1995). 22 H. Kuhn, V. Demidov, M. D. Frank-Kamenetskii, and P. E. Nielsen, Nucleic Acids Res. 26, 582 (1997). 23 A. G. Veselkov, V. V. Demidov, P. E. Nielsen, and M. Frank-Kamenetskii, Nucleic Acids Res. 24, 2483 (1996).

334

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

[ 1 5]

sequence discrimination, but the results have shown that this is also kinetically controlled and may be as large as 1000-fold for a single mismatch in a 10-mer target. 22 Duplex Invasion Although PNA-DNA duplexes are slightly more stable than the corresponding DNA-DNA duplexes for most sequences, PNAs are generally not able to invade a double-stranded DNA target, because of the competition from the displaced strand. However, G-rich, homopurine PNA oligomers form duplexes with complementary deoxyoligonucleotides of especially high thermal stability. For instance, the PNADNA duplex of a 10-mer (A4GzAGAG) PNA has a Tmof 70 °, which is similar to that of a triplex-invading 10-mer homopyrimidine PNA. Indeed, this homopurine PNA was found to form a duplex invasion complex, 8 but the stability of this complex was considerably less than that of a triplex invasion complex. Some stabilization was, however, observed on exchanging all the adenines with diaminopurines, which gave a Tm increase of about 15°. 25 These results clearly show that only a little further stabilization is necessary to observe duplex invasion with PNA oligomers. Alternatively, this binding mode may be possible if the DNA double helix is destabilized as seen in cruciform structures, a6 or perhaps by moderate to high negative supercoiling. It has been established that triplex-invasive binding can be greatly accelerated (even 100-fold) by negative supercoiling. 27 Double Duplex Invasion A duplex invasion complex may also be dramatically stabilized by targeting the displaced DNA, forming a double duplex invasion complex (Fig. 2D). 28 Naturally, in this binding mode the two PNA oligomers are sequence complementary, and if composed of natural bases, the two PNAs would "quench" each other by forming the more stable PNA-PNA duplex. However, it is possible to create pseudo-complementary PNAs by employing the sterically compromised diaminopurine-thiouracil base pair instead of the natural adenine-thymine base pair (Fig. 5). Such pseudo-complementary PNAs retain full binding to sequence complementary DNA, but because of the steric interference of the diaminopurinethiouracil base pair, they bind poorly if at all to each other. At this stage only a pseudo-complementary A-T base pair has been developed. Consequently, efficient

24g. g. Demidov,M. V.Yavnilovich,B. R Belotserkovskii,M. D. Frank-Kamenetskii,and P.E. Nielsen, Proc. Natl. Acad. Sci. U.S.A. 92, 2637 (1995). 25G. Haaima,H. E Hansen,L. Christensen, O. Dahl, and R E. Nielsen,Nucleic Acids Res. 25, 4639 (1997). 26T. Ishiharaand D. R. Corey,J. Am. Chem. Soc. 121, 2012 (1999). 27T. Bentin and P. E. Nielsen,Biochemistry 35, 8863 (1996). 28j. Lohse,O. Dahl, and P. E. Nielsen, Proc. Natl. Acad. Sci. U.S.A. 96, 11804 (1999).

[15]

PNA TARGETINGOF DOUBLE-STRANDEDDNA

335

OH3 H~ ,...H"/

.o0

',~

H....N/H. - ' " ' 0 " ~ ' 1 /N.

N--

,..~1~ ,.:H.X" /Thymine "

,.-i:-., ; I -'"~N ~ IN,,..-ff :~.~.... T~ouraci, H

H

Diaminopurine

Diaminopurine

H.~ ,IH"'"O~ N L - H / N y N" " ~ f "1~".... N.,.-"'~Nj~..H

0

H.~.N.... H"""""O'~'~ i

Thymine

I

Adenine

'~ I~.--., Thi~uracil

N-- -,~.,--""

~

N "~'- ~'q'IM" ./'~, ~

I

'"

II

{

~

". "_ ."

~

."

....

Adenine

FIG.5. Chemical structure of a pseudoisocytosine(qJiC)-G-Ctriplet, showinghow the ~iC base is able to form two hydrogenbondsto the guaninein the Hoogsteenmode.

double duplex invasion by PNAs requires a target having at least 50% AT content. 28 Although no thorough kinetic and stability studies have yet been reported on PNA double duplex invasion complexes, they appear to behave quite similarly to triplex invasion complexes in terms of slow rate of formation and sensitivity to elevated ionic strength. However, once formed these complexes also have high stability, zs "Functional" Invasion Complexes A variety of biological effects of PNA (triplex) invasion complexes have been discovered and the unusual and tight binding have also inspired the development of various molecular biology methodologies. In contrast to oligonucleotide triple helix complexes, PNA triplex invasion complexes cause elongation arrest of transcribing phage, bacterial, and eukaryotic RNA polymerases, and the effect is (not surprisingly) much more pronounced when the PNA is bound to the template strand. 29-31 In fact, for phage polymerases 29E E. Nielsen,M. Egholm, and O. Buchardt, Gene 149, 139 (1994). 3oN. J. Peffer, J. C. Hanvey,J. E. Bisi, S. A. Thomson,C. E Hassman,S. A. Noble,and L. E. Babiss, Proc. Natl. Acad. Sci. U.S.A. 90, 10648 (1993). 31 N. E. Mollegaard,O. Buchardt,M. Egholm,and E E. Nielsen,Proc. Natl. Acad. Sci. U.S.A. 91, 3892 (1994).

336

C H E M I C A AND L MOLECULAR BIOLOGICAL APPROACHES

[1 5]

almost no effect is observed when the PNA is bound to the nontemplate strand. 29 Furthermore, both triplex and double duplex invasion complexes efficiently inhibit the binding of proteins, such as RNA polymerase, transcription factors, or restriction enzymes and methylases that bind to a DNA site overlapping or proximal to the PNA target. 32 34 Most interestingly, it has also been found that RNA polymerase may recognize a PNA triplex invasion complex as a promoter, and start transcription by using the single-stranded loop as the template. 35 Thus PNAs may function as artificial transcription factors, using their dsDNA targets as PNA-inducible promoters. Therefore, PNAs have the potential to be developed into efficient antigene (or even gene-activating) therapeutic drugs. However, apart from the nontrivial issue of cellular uptake, which is not discussed here, it is far from clear whether PNA once present in the cell nucleus will actually be able to invade the duplex DNA in chromatin at a rate that can result in a sufficiently high target site occupancy to elicit a biological effect. However, even a slow on rate will, because of the extremely high stability (slow off rate) of the complex, eventually result in complex accumulation. Indeed, a 10-fold increase in mutation rate at a PNA site has been reported on treating mouse cells ex vivo with a bis-PNA aimed at this target, 35 and evidence of PNA-directed transcriptional activation in cells in culture was also presented. 36 Apart from their biological significance, these results strongly suggest that PNA invasion complexes may indeed form in intact living cells. Furthermore, the results may be rationalized on the basis of in vitro results showing that negative supercoiling may enhance PNA binding by as much as 200-fold and that any process that facilitates DNA unwinding and opening, such as the transcription process itself, 37 another DNA-opening ligand, 38 or DNA cruciforms 26 catalyzed invasive PNA binding. Indeed, it appears that under certain conditions of D N A constraint (supercoiling) and structure (cruciform) even simple duplex invasion complexes may be sufficiently stabilized to elicit biological effects. Thus a series of quite puzzling results indicating that simple duplex-forming PNAs may show antigene transcription inhibition in cell culture e x vivo 39'40 could perhaps be explained on the basis of target structure and DNA topology in vivo. However, more evidence is required.

32E E. Nielsen, M. Egholm, R. H. Berg, and O. Buchardt, Nucleie Acids Res. 21, 197 (1993). 33 K. I. Izvolsky, V. V. Demidov, N. O. Bukanov, and M. D. Frank-Kamenetskii,Nucleic: Acids Res. 26, 5011 (1998). 34T. A. Vickers, M. C. Griffieth, K. Ramasamy, L. M. Risen, and S. M. Freier, Nucleic Acids Res. 23, 3003 0995). 35A. E Faruqi, M. Egholm, and E M. Glazer, Proc. Natl. Acad. Sci. U.S.A. 95, 1398 (1998). 36G. Wang, X. Xu, B. Pace, D. A. Dean, R M. Glazer, E Chan, S. R. Goodman, and I. Shokolenko, Nucleic Acids Res. 27, 2806 (1999). 37 H. J. Larsen and R E. Nielsen, Nucleic Acids Res. 24, 458 (1996). 38 A. Kurakin, H. J. Larsen, and E E. Nielsen, Chem. Biol. 5, 81 (1998). 39 L. C. Boffa, R L. Morris, E. M. Carpaneto, M. Louissaint, and V. G. Allfrey, J. Biol. Chem. 271, 13228 (1996).

[1 5]

PNA TARGETING OF DOUBLE-STRANDED D N A

337

A variety of molecular biology and diagnostic techniques have also been developed by exploiting the unique properties of PNA invasion complexes. The high sequence specificity and high stability of PNA triplex invasion complexes have been exploited to develop an Achilles heel DNA cleavage method that allows sequence methylase/restriction enzyme-assisted single-site cleavage of large D N A molecules, such as yeast chromosomes. 33 This technology was expanded to include targets for double duplex invasive pseudo-complementary PNAs. 41 Two triplex-invading PNAs binding to closely positioned homopurine/ homopyrimidine targets have been used as "openers" to form a PD-loop, in which an oligonucleotide is hybridized to the DNA in between the PNA targets. 42 This principle has been successfully employed to capture specific DNA fragments, using a biotinylated oligonucleotide, 43 as the basis of a D N A quantification method, 44 and as a means to form interesting, topologically linked earring or padlock DNA structures. 45 Finally, PNA invasion complexes can be used to tag plasmid D N A in transfection applications. Using a fluorescent PNA, the plasmid can be traced after transfection, 46 and the PNA may also be used to tag the plasmid D N A with ligands that target it to the nucleus using a nuclear localization signal (NLS) peptide 47 or a ligand that targets the plasmid to a certain cell type. 48 Such techniques could find wide applications in gene therapy.

Methods for Studying DNA Binding

Protein Nucleic Acid Double-Stranded

Standard methods used for investigating l i g a n d - D N A interactions, such as gel mobility shift assays and DNase I footprinting, can be employed to study P N A dsDNA binding. 49 However, to establish that an invasion complex is present, it is

40 L. C. Boffa, E. M. Carpaneto, M. R. Mariani, M. Louissaint, and V. G. Allfrey, Oncol. Res. 9, 41 (1997). 41 K. I. Izvolsky, V. V. Demidov, P. E. Nielsen, and M. D. Frank-Kamenetskii, Biochemistry 35, 10908

(2OOO). 42 N. O. Bukanov, V. V. Demidov, P. E. Nielsen, and M. D. Frank-Kamenetskii,Proc. Natl. Acad. Sci. U.S.A. 95, 5516 (1998). 43 V. Demidov, N. O. Bukanov, and M. D. Frank-Kamenetskii,Curr. Issues Mol. Biol. 2, 31 (2000). 44 N. E. Broude, V. V. Demidov, H. Kuhn, J. Gorenstein, H. Pulyaeva, P. Volkovitsky,A. K. Drukier, and M. D. Frank-Kamenetskii,J. Biomol. Struct. Dyn. 17, 237 (1999). 45 H. Kuhn, V. V. Demidov, and M. D. Frank-Kamenetskii,Angew. Chem. Int. Ed. 38, 1446 (1999). 46 O. Zelphati, X. Liang, C. Nguyen, S. Barlow, S. Sheng, Z. Shao, and P. L. Feigner, BioTeehniques 28, 304-308, 310, 312-314, 316 (2000). 47 L. J. Brand6n, A. J. Mohamed, and C. I. E. Smith, Nat. Biotechnol. 17, 784 (1999). 48 K. W. Liang, E. P. Hoffman, and L. Huang, Mol. Ther. 1,236 (2000). 49 H. J. Larsen and R E. Nielsen, in "Peptide Nucleic Acids: Protocols and Applications" (R E. Nielsen and M. Egholm, eds.), pp. 221-240. Horizon Scientific Press, Norfolk, UK, 1999.

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CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[ 15]

most convenient to use methods that are able to probe the presence of a singlestranded DNA loop or at least unpaired bases. Probably the easiest and most informative method--provided unpaired (unstacked) thymines, are present--is permanganate oxidation of the 5-6 double bond of thymine. Subsequent alkaline (piperidine) treatment produces strand scissions at reactive thymines. Unstacked thymines, such as those in a single-stranded loop, or in DNA junctions, react much more efficiently with permanganate than thymines in double-stranded DNA and are easily identified semiquantitively by this method. 14'27 Alternatively, nuclease S1 or mung bean nuclease can be used to detect single-stranded DNA regions, but this requires a change of buffer. 1'49'5° Unpaired adenines may with relatively low sensitivity be detected by probing with diethyl pyrocarbonate (DEPC)/ Analogously to the probing of guanine N7 participation in DNA triple helices, dimethyl sulfate (DMS) is also a specific probe for guanine Hoogsteen binding in PNA2DNA triplexes. 14 Finally, PNA oligomers conjugated to a DNA-cleaving moiety, such as EDTA/ Fe 51 porphyrin, 52 Gly-His-His/Cu+, 53 or an anthraquinone, 54 may be used for affinity cleavage experiments. D e s i g n o f P r o t e i n Nucleic Acids for D o u b l e - S t r a n d e d DNA T a r g e t i n g As outlined above, four different binding mechanisms may be exploited when targeting double-stranded DNA with PNA oligomers. In terms of stability the following general ranking is expected for "comparable" complexes: triplex invasion > double duplex invasion >> triplex > duplex invasion. However, whereas the invasion complexes have a slow rate of formation, which is further decreased by increasing ionic strength, major groove triplexes form much faster, but virtually nothing is known about their properties in general at this stage. Therefore, if possible, the best advice is to target a homopurine site of 8-10 bases in length, using a bis-PNA (Fig. 6). The complex loses stability below about 8 bases and sequence specificity above about 10-12 bases. It is also advisable to include three or four positive charges, for example, in the form of lysines, in order to increase the binding rate. Most probably the increased cationic character of the PNA ensures a high local concentration of the PNA in close proximity to the 50 V. V. Demidov, M. V. Yavnilovich, and M. D. Frank-Kamenetskii, Biophys. J. 72, 2763 (1997). 51 j. Lohse, C. Hui, S. H. SOnnichsen, and P. E. Nielsen, in "DNA and RNA Cleavers and Chemotherapy of Cancer and Viral Diseases" (B. Meunier, ed.), pp. 133-141. NATO ASI Series. Kluwer Academic Publishers, Dordrecht, The Netherlands, 1996. .52 p. Bigey, S. H. S0nnichsen, P. E. Nielsen, and B. Meunier, Bioconjug. Chem. 3, 267 (1997). 53 M. Footer, M. Egholm, S. Kron, J. M. Coull, and P. Matsudaira, Biochemistry 35, 10673 (1996). 54 B. Armitage, T. Koch, H. Frydenlund, H. Orum, H. G. Batz, and G. B. Schuster, Nucleic Acids Res. 25, 4674 (1997).

[15]

PNA TARGETINGOF DOUBLE-STRANDEDDNA

339

H-Lys-Lys-TTTTJJTJTJ ~ . 5- AAAAGGAGAG ~-- 3 ' H2N- Lys- TTTTCCTCTC ~ FIG. 6. Schematic drawing of a bis-PNA binding to its purine target. The bis-PNA is constructed with G-recognizing pseudoisocytosine (J) in the parallel Hoogsteen strand, and the Watson-Crick strand is binding in an antiparallel orientation. The linker is conveniently constructed from three 8amino-3,6-dioxaoctanoic acid units [-NH-(CH2)2-O-(CH2)2-O-(CH2)-CO-]. H-, The amino terminal of the PNA; H2N-, the amidated carboxyl terminal.

DNA, thereby increasing the probability of invading the dynamically breathing and therefore transiently open DNA helix. 27'37'38 Finally, pseudoisocytosine should be used in the Hoogsteen strand to avoid pH sensitivity of the binding. On the other hand, if binding is possible at acidic pH, the presence of cytosines in the Hoogsteen strand will increase both binding rate and sequence discrimination. 2° If homopurine targets are not available, the double duplex invasion principle is an obvious option, although not yet thoroughly explored and characterized. In this case the target (about 10 bp) should contain at least 50% AT base pairs. In principle, it should also be possible to target nonhomopurine targets by triplex-forming bis-PNAs if novel bases are employed that recognize cytosine or thymine in the "Hoogsteen mode" [such as the E-base 55 (Fig. 7)]. Many attempts to construct such bases have been reported in a DNA as well as a PNA context, but the results have been rather disappointing56; no efficient candidates have yet been identified. Finally, under certain conditions not yet fully understood simple duplex invasion complexes may form and be sufficiently stable to yield biological effects, but this area needs much further research before it can be recommended as a reliable method.

° ...... ....... H

.,.,..'1~

" - - _ _ - ~ N''H''" T

l

N~'~,,

N'%"~N

A

0

FIG. 7. Chemical structure of a putative triplet formed by binding of the E-base to the T-A base pair.

55 A. B. Eldrup, O. Dahl, and E E. Nielsen, J. Am. Chem. Soc. 119, 11116 (1997). 56 K. R. Fox, Curr. Med. Chem. 7, 17 (2000).

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CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[ 1 6]

Concluding Remarks The results obtained so far clearly show that PNA is an interesting and useful reagent for sequence-specific targeting of double-stranded DNA. It is also clear that we are far from understanding all the relevant aspects, both as regards binding modes and mechanism of this methodology. This is particularly true when considering effects and possible (medical) applications in live cells and animals. Continued studies and development of peptide nucleic acids should yield both a deeper understanding of bimolecular recognition principles and also contribute to various areas of molecular biology, gene therapeutic drug development, and genetic diagnostics. Acknowledgment This work was supported by the Lundbeck Foundation.

[16]

Drug

By

Interaction

with

Triple-Helical

CHRISTOPHE ESCUD[~, THI~RESE GARESTIER,

and

Nucleic

Acids

JIAN-SHENG SUN

Introduction Triple-helical DNA was first discovered more than 40 years ago. Renewed interest in this structure developed at the end of the 1980s because of two discoveries. First, sequence-specific recognition of double-helical DNA can be achieved by short oligonucleotides forming a short intermolecular triple helix.l'2 This recognition can be used to design tools for manipulating double-stranded DNA in vitro as well as for interfering with DNA-related processes in vivo. Numerous studies have now shown that the so-called triplex-forming oligonucleotides (TFOs) can be used to cleave or chemically modify a DNA target, 3 and that TFOs can interfere with binding of transcription factors, transcription elongation, or DNA repair. 4 Moreover, it has been shown that DNA can adopt an intramolecular triple-helical structure, also called H-DNA, under certain conditions. 5 The formation of this structure has been shown in bacteria and may play a role in the 1 T. Le Doan, L. Perrouault, D. Praseuth, N. Habhoub, J.-L. Decout, N. T. Thuong, J. Lhomme, and C. H61~ne, Nucleic Acids Res. 15, 7749 (1987). 2 H. E. Moser and P. B. Dervan, Science 238, 645 (1987). 3 N. T. Thuong and C. H61~ne, Angew. Chem. Int. Ed. 32, 666 (1993). 4 C. Giovannangeli and C. H6l~ne, Antisense Nucleic Acid Drug Dev. 7, 413 (1997). 5 R. D. Wells, D. A. Collier, J. C. Hanvey, M. Shimizu, and F. Wohlrab, FASEB J. 2, 2939 (1988).

METHODSIN ENZYMOLOGY,VOL.340

Copyright© 2001by AcademicPress All rightsof reproductionin any formreserved. 0076-6879/00 $35.1,)0

340

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[ 1 6]

Concluding Remarks The results obtained so far clearly show that PNA is an interesting and useful reagent for sequence-specific targeting of double-stranded DNA. It is also clear that we are far from understanding all the relevant aspects, both as regards binding modes and mechanism of this methodology. This is particularly true when considering effects and possible (medical) applications in live cells and animals. Continued studies and development of peptide nucleic acids should yield both a deeper understanding of bimolecular recognition principles and also contribute to various areas of molecular biology, gene therapeutic drug development, and genetic diagnostics. Acknowledgment This work was supported by the Lundbeck Foundation.

[16]

Drug

By

Interaction

with

Triple-Helical

CHRISTOPHE ESCUD[~, THI~RESE GARESTIER,

and

Nucleic

Acids

JIAN-SHENG SUN

Introduction Triple-helical DNA was first discovered more than 40 years ago. Renewed interest in this structure developed at the end of the 1980s because of two discoveries. First, sequence-specific recognition of double-helical DNA can be achieved by short oligonucleotides forming a short intermolecular triple helix.l'2 This recognition can be used to design tools for manipulating double-stranded DNA in vitro as well as for interfering with DNA-related processes in vivo. Numerous studies have now shown that the so-called triplex-forming oligonucleotides (TFOs) can be used to cleave or chemically modify a DNA target, 3 and that TFOs can interfere with binding of transcription factors, transcription elongation, or DNA repair. 4 Moreover, it has been shown that DNA can adopt an intramolecular triple-helical structure, also called H-DNA, under certain conditions. 5 The formation of this structure has been shown in bacteria and may play a role in the 1 T. Le Doan, L. Perrouault, D. Praseuth, N. Habhoub, J.-L. Decout, N. T. Thuong, J. Lhomme, and C. H61~ne, Nucleic Acids Res. 15, 7749 (1987). 2 H. E. Moser and P. B. Dervan, Science 238, 645 (1987). 3 N. T. Thuong and C. H61~ne, Angew. Chem. Int. Ed. 32, 666 (1993). 4 C. Giovannangeli and C. H6l~ne, Antisense Nucleic Acid Drug Dev. 7, 413 (1997). 5 R. D. Wells, D. A. Collier, J. C. Hanvey, M. Shimizu, and F. Wohlrab, FASEB J. 2, 2939 (1988).

METHODSIN ENZYMOLOGY,VOL.340

Copyright© 2001by AcademicPress All rightsof reproductionin any formreserved. 0076-6879/00 $35.1,)0

[16]

DRUG INTERACTIONWITHTRIPLE-HELICALNAs

341

regulation of some eukaryotic genes (see Mirkin and Frank-Kamenetskii 6 for a review). One limitation to the possible use of TFOs as therapeutic agents or the possible biological role of H-DNA is the low stability of some commonly studied triplexes under physiological conditions. This problem may be overcome by the action of proteins that can specifically bind triple-helical DNA, and thus would be able to stabilize them. 7-9 Alternatively, low molecular weight compounds that can bind triplex DNA more tightly than duplex DNA would also have the ability to enhance triplex stability. If their target is an intramolecular triplex, whose formation may interfere with the regulation of gene expression, these molecules may directly affect the gene expression profile. Therefore, such molecules constitute potential drugs, either in conjunction with TFOs or by themselves as structure-specific DNA-binding compounds. In this chapter, after a brief description of the characteristics of triple-helical DNA, we describe the physicochemical aspects of drug binding to triplex DNA, with special emphasis on the different methods used to study the binding of low molecular weight compounds to triplex DNA, through a review of examples published in the literature. We then discuss some methodological and general considerations about targeting triplehelical nucleic acids with drugs, and present the biological applications of such drugs. C h a r a c t e r i s t i c s of T r i p l e - H e l i c a l DNA The structure of triple-helical DNA has been extensively studied (see Radhakrishnan and Patel l° for a review). A triplex-forming oligonucleotide binds into the major groove of double-stranded DNA (dsDNA) at an oligopyrimidine • oligopurine sequence. Specific recognition is achieved by formation of hydrogen bonds between bases in the third strand and purine bases in the target sequence (Fig. 1) (see Sun et al. 11 for more details). Pyrimidine-containing oligonucleotides bind in a parallel orientation with respect to the oligopurine sequence by formation of Hoogsteen hydrogen bonds between a T or a protonated C in the third strand and a T. A or a C. G base pair in the duplex target, respectively. Oligonucleotides containing purines can also bind double-helical DNA. G can form two reverse Hoogsteen hydrogen bonds with a C. G base pair and A or T can do so with a T. A base pair. In this configuration, binding of the o!igonucleotide occurs in an antiparallel orientation. Some (G, T)-containing oligonucleotides may also bind parallel

6 S. M. Mirkin and M. D. Frank-Kamenetskii, Annu. Rev. Biophys. Biomol. Struct. 23, 541 (1994). 7 A. L. Guieysse, D. Praseuth, and C. H61~ne, J. MoL Biol. 267, 289 (1997). 8 M. Musso, L. D. Nelson, and M. W. Van Dyke, Biochemistry 37, 3086 (1998). 9 L. D. Nelson, M. Musso, and M. W. Van Dyke, J. Biol. Chem. 275, 5573 (2000). l0 I. Radhakrishnan and D. J. Patel, Biochemistry 33, 11405 (1994). 11 j. S. Sun, T. Garestier, and C. H61~ne, Curr. Opin. Struct. Biol. 6, 327 (1996).

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CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

Pyrimidine 3'

[1 g]

motif

51

~'-W-C W...~

-~--C-H

C..->

<--W-H "

5' 3' 5'

3 ~ 5'

~"

w-c Purine-rich

motif

5'

--'H-"/ 3 q

3' 5'

d

H

g

I1

i

j

N----H---.~

.'

\

,o....._.( H

FIG. 1. Structure of triple helices and examples of base triplets. Left: The structure of triple helices is shown. The white ribbon is called the Watson strand (W) and contains an oligopyrimidine sequence, the gray ribbon is called the Crick strand (C) and contains an oligopurine sequence, whereas the black ribbon is the third strand, also called the Hoogsteen strand (H), which can bind in a parallel orientation for the pyrimidine motif (top) or in an antiparallel orientation for the purine-rich motif (bottom). Right: The structure of various base triplets is shown, with the names given to the different grooves of the triple helix: Watson-Crick (W~S) between the Watson and Crick strands, Watson-Hoogsteen (W-H) between the Watson and Hoogsteen strands, and Crick-Hoogsteen (C-H) between the Crick and Hoogsteen strands.

to the oligopurine sequence. However, the different geometries of the T . A*T and C . G*G base triplets in the Hoogsteen pattern require that there be few A p G and G p A steps in the oligopurine sequence. In triplex DNA, the major groove of the duplex is occupied by the third strand, resulting in the formation of two new grooves, which are called the W a t s o n - H o o g s t e e n and C r i c k - H o o g s t e e n grooves (see Fig. 1). C o m p a r e d with duplex DNA, the helical parameters are slightly modified, and the base triplets form an increased stacking area. The hydration pattern

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DRUG INTERACTIONWITH TRIPLE-HELICALNAs

343

within the different grooves is specific, and the electrostatic potential is affected by the presence of an additional negative charge for each base platform and the occasional presence of protonated cytosines. These specific features may be recognized by nucleic acid-binding agents. Experimental

Study of Drug Binding to Triplex DNA

As early as 1990 it was demonstrated that the junction between triplex and duplex represents a strong site for intercalation of an ellipticine derivative. 12 This intercalation resulted in DNA distortion, as detected by hypersensitivity to chemical probes.13 Evidence of drug binding to a triple-helical structure was provided for the intercalator ethidium bromide.14' 15 Subsequently, many different compounds, including many large polycyclic aromatic molecules, which are believed to be potential intercalators and known groove-binding agents, have been studied for their ability to bind triplex DNA. The first compound that binds much more tightly to triplex DNA than duplex DNA, a benzo[e]pyridoindole derivative (BePI, 1), was described in 1992.16 The structures of this molecule and of several other triplex binding agents (2-15) are shown in Fig. 2. They will be described in greater detail below. Triple helix formation itself can be detected by thermal denaturation experiments, electrophoretic mobility shift assays, DNase I or chemical footprinting, or inhibition of restriction enzyme cleavage (see Franqois et al. 17 for a review of methods used to study triple-helical DNA). By extension, the effects of various ligands on triplex formation and stability can be studied by these methods. Detailed thermodynamic or structural data about triplex DNA-ligand complexes requires. in-depth investigation using more sophisticated biophysical methods. T h e r m a l Denaturation E x p e r i m e n t s

Thermal denaturation is a commonly used technique for investigating triplehelical DNA formation. In general, it is possible to observe a biphasic melting curve in which the transition at the lower temperature corresponds to the dissociation of the triplex into a duplex and a third strand, and the transition at the higher temperature corresponds to the melting of the duplex into single strands.

t2 L. Perrouault, U. Asseline, C. Rivalle, N. T. Thuong, E. Bisagni, C. Giovannangeli,T. Le Doan, and C. H616ne,Nature (London) 344, 358 (1990). 13D. A. Collier, J. L. Mergny, N. T. Thuong, and C. H616ne,NucleieAcids Res. 19, 4219 (1991). 14p. V. Scaria and R. H. Shafer, J. Biol. Chem. 266, 5417 (1991). t5 j. L. Mergny, D. Collier, M. Roug6e,T. Montenay-Garestier,and C. H61~ne,Nucleic Acids Res. 19, 1521 (1991). 16j. L. Mergny,G. Duval-Valen,tin,C. H. Nguyen,L. Perrouault, B. Faucon, M. Roug6e,T. MontenayGarestier, E. Bisagni, and C. H616ne,Science 256, 1681 (1992). 17j. C. Franqois, J. Lacoste, L. Lacroix, and J. L. Mergny,Methods Enzymol. 313, 74 (2000).

344

[16]

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

R

~"~ OCH3 R

R

CH3 ~l 1 - -

~"/Y" OCH3

CH3

~L,~..,~.OCH 3 3

/..~_.~OCH3

FI

R

N+--

v

_4

v

-OCHa

5

OCH3 ~OCH3 CH30- v

R 7

R

"~ ~ CH3

"~ N ~ CH3

8

~ R

9

R O R1

lO

. ~ ''OCH3

2

~

R6

O

11

12

14

15

O--k

H3CO~

O OCH3 13

NH2

FIG. 2. Chemical structures of the triplex-binding agents 1-15. The names and references are indicated in the chapter. For the sake of clarity, the structure of alkylamine side chains has been omitted. For 1, 2, 3, 4, 5, 6, and 9, R = NH(CH2)3NH3 +. For 7, R = NH(CH2)2NH+(CH3)2 . For 10, R = (CH2)3NH+(CH3)2. For II and 12, R = NHCO(CH2)2NH+(CH3)2.

Specific binding of a drug to triplex D N A results in an increase of the melting temperature for the triplex. For example, it has been shown that a triple helix formed by a 14-mer oligonucleotide, which dissociates from the duplex at 18 ° in the absence of any ligand, can have its melting temperature increased to 38 ° in the presence of 1 5 / z M BePI (1), 16 tO 62 ° in the presence of a 15 # M concentration of the dibenzophenanthroline 10,18 and up to more than 70 ° in the presence of

18 O. Baudoin, C. Marchand, M.-E Teulade-Fichou, J.-E Vigneron, J. S. Sun, T. Garestier, C. Hrlrne, and J. M. Lehn, Chem. Eur. J. 4, 1504 (1998).

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DRUG INTERACTIONWITH TRIPLE-HELICALNAs

345

benzo[f]quino[3,4-b]quinoxaline (BQQ 6).19 It should be noted that this temperature is higher than the melting temperature of the 26-bp duplex DNA used as a target under the same conditions. In such cases, be aware that the absence of biphasic melting does not reflect the absence of triplex formation. Triple-stranded homopolymers have often been used for screening the triplex-stabilizing properties of nucleic acid-binding agents. These include the naphthylquinoline derivative 7, where two bicyclic rings are joined by a flexible link, 2° and several commercially available polyaromatic compounds, such as coralyne (8) 21 or various other related derivatives. 22 When homopolymers are used, ligand redistribution on melting of triplex DNA can lead to multiphasic melting curves, the interpretation of which can be difficult. Fluorescence Methods

Binding of a chromophore to nucleic acids can induce changes in the absorption and emission spectra of the dye. For example, the fluorescence of ethidium bromide is enhanced on binding to duplex and triplex DNA,15 whereas that of BePI (1) and BgPI (2) is quenched. 23 The quenching is higher in triplex DNA compared with duplex DNA. This may result from differences in the binding geometry between the two structures, such that a stronger quenching effect is observed in the ligand-triplex complexes. The quenching effect can be used to study binding of drug to triplexes, using Scatchard plots or the McGhee and van Hippel model. For example, in the case of coralyne, binding to triplex DNA induces up to 70% quenching. Examination of binding curves allowed Lee and colleagues to measure a binding constant of 1.5 x 106 M -I for T. A*T triplets in a pH 8.0 buffer containing 2 mM Mg2+. 21 Fluorescence can also be quenched in the presence of acrylamide or negatively charged molecules such as [Fe(CN)6] 4-. This quenching is usually abolished when the dye is intercalated between DNA base pairs. Such a phenomenon has been observed for ethidium bromide in the presence of triplehelical DNA. 14 The observation of fluorescence energy transfer also provides an indication for an intercalative mode of binding.15' J6 The average number of bases that transfer excitation energy to the bound molecule can be calculated by adapting the method of analysis described by Rayner et al. 2a This number, which provide

19C. Escudr, C. H. Nguyen, S. Kukreti, Y. Janin, J. Sun, E. Bisagni, T. Garestier, and C. Hrlbne, Proc. Natl. Acad. Sci. U.S.A. 95, 3591 (1998). 20W. D. Wilson, E A. Tanious, S. Mizan, S. Yao,A. S. Kiselyov,G. Zon, and L. Strekowski,Biochemistry 32, 10614 (1993). 2t j. S. Lee, L. J. P. Latimer, and K. J. Hampel,Biochemistry 32, 5591 (1993). 22L. J. P. Latimer, N. Payton,G. Forsyth,and J. S. Lee, Biochem. Cell. Biol. 73, 11 (1995). 23D. S. Pilch, M. T. Martin, C. H. Nguyen,J. S. Sun, E. Bisagni, T. Garestier, and C. Hrl~ne, J. Am. Chem. Soc. 115, 9942 (1993). 24D. M. Rayner, A. G. Szabo, R. O. Loutfy, and R. W. Yip,J. Phys. Chem. 84, 289 (1980).

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CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[16]

a measure of energy transfer efficiency, was found to be higher (4.8) in a BePItriplex complex than in a BePI-duplex complex (2.5). 25 The greater efficiency of energy transfer in the former complex may indicate that this complex involves better stacking interactions. Circular Dichroism

Circular dichroism (CD) is another commonly used technique for the study of nucleic acids and their complexes with ligands. Changes in the CD spectra of both the nucleic acid and the bound ligand can be monitored. The binding of numerous classic groove binders to triplex DNA has been investigated by this method, including netropsin, 26'27 distamycin, 28 Hoechst 33258, 29 berenil, 3°'31 and 4',6-diamidino-2-phenylindole (DAPI, 14). 32 Titration of a nucleic acid with the ligand allows measurement of the binding affinity, using the Epstein theory. It is generally found that these groove-binding drugs bind to duplex DNA with a higher affinity than to triplex DNA, resulting in the destabilization of triplex DNA. For example, the apparent KD of netropsin for triplex DNA was 107 M-1 whereas it was > 109 M-1 for duplex DNA. 27 This technique also allows some structural investigation. The CD spectrum of a ligand bound to double-stranded DNA can differ from that of the same ligand bound to triple-stranded DNA. Such a phenomenon, which has been observed with Hoechst 29 or berenil, 3° but not with netropsin, suggests that the conformations of the ligand are slightly different when it is bound to the duplex or the triplex. Linear Dichroism

Linear dichroism (LD) can been used to study the binding of polyaromatic compounds to homopolymeric helical structures. For BePI, BgPI, the benzo[f]pyrido[4,3-b]quinoxaline (BfPQ) derivative (9), and BQQ (6), the similar magnitude of reduced LD of the electronic transitions of the polynucleotide bases and of the bound ligands suggests an orientation of the plane of the fused-ring systems perpendicular to the helix axis, which is indicative of intercalation. 33 This technique

25 D. S. Pilch, M. J. Waring, J. S. Sun, M. RougEe, C. H. Nguyen, E. Bisagni, T. Garestier, and C. H61bne, J. Mol. Biol. 232, 926 (1993). 26 y. W. Park and K. J. Breslauer, Proc. Natl. Acad. Sci. U.S.A. 89, 6653 (1992). 27 M. Durand, N. T. Thuong, and J. C. Maurizot, J. Biol. Chem. 267, 24394 (1992). 28 M. Durand and J. C. Maurizot, Biochemistry 35, 9133 (1996). 29 M. Durand, N. T. Thuong, and J. C. Maurizot, Biochimie 76, 181 (1994). 30 M. Durand, N. T. Thuong, and J. C. Maurizot, J. BiomoL Struct. Dyn. 11, 1191 (1994). 31 D. S. Pilch, M. A. Kirolos, and K. J. Breslauer, Biochemistry 34, 16107 (1995). 32 D. S. Pilch and K. J. Breslauer, Proc. Natl. Acad. Sci. U.S.A. 91, 9332 (1994). 33 S. K. Kim, J. S. Sun, T. Garestier, C. Hfl~ne, E. Bisagni, A. Rodger, and B. Norden, Biopolymers 42, 101 (1997).

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has also shown that various complexes of ruthenium can intercalate into triplehelical DNA and stabilize it. 34

Nuclear Magnetic Resonance Spectroscopy Only two studies have employed nuclear magnetic resonance (NMR) to explore ligand binding to triplex DNA. In the first, NMR was used to monitor the solvent exchange properties of the exchangeable protons of the ligand. This technique allowed measurement of self-association constants for both the BePI (1) and BgPI (2) derivatives, 23 but did not provide high-resolution structural information. The binding of coralyne was also investigated by using short intramolecular triplex DNA with different sequences. 35 The intercalative binding mode was confirmed. However, at NMR concentrations, and even in the presence of an excess of DNA, two different binding sites were observed: a primary site between two T-A*T triplets, and a neighboring site with a lower affinity. These complexes led to two sets of overlapping spectra, which made the precise assignment of proton resonances impossible.

Hydrodynamic Methods Viscometric titration of closed circular duplex DNA is commonly used to determine whether a compound intercalates into double-helical DNA. Intercalating agents usually induce the unwinding of supercoiled plasmids and the lengthening of linear helical structures, which are reflected by an increase in the viscosity of the samples. Several polyaromatic molecules that stabilize triple helices bind duplex DNA by intercalation, as evidenced by unwinding of supercoiled plasmid DNA. 36 In the case of BePI, an increase in the relative contour lengths of both double- and triple-stranded homopolymer structures [i.e., poly(dA) • poly(dT) and poly(dA) • [poly(dT)]2] has been demonstrated. 25

Competition Dialysis Competition dialysis techniques provide a direct method to obtain the DNAbinding preferences of a candidate ligand. For example, this method has been used to show that the 1,4-disubstituted anthraquinone 11 binds preferentially to homopolymeric duplexes whereas the 2,6-derivative 12 favors binding to triplexes, 37 An improved competition dialysis procedure has been developed that allows quick 34 S.-D. Choi, M.-S. Kim, S. K. Kim, E Lincoln, E. Tuite, and B. Norden, Biochemistry 36, 214 (1997). 35 A. A. Moraru-Allen, S. Cassidy, J. L. Asensio Alvarez, K. R. Fox, T. Brown, and A. N. Lane, Nucleic Acids Res. 25, 1890 (1997). 36 C. Marchand, C. Bailly, C. H. Nguyen, E, Bisagni, T. Garestier, C. H61~ne, and M. J. Waring, Biochemistry 35, 5022 (1996). 37 I. Haq, J. E. Ladbury, B. Z. Chowdhry, and T. C. Jenkins, J. Am. Chem. Soc. 118, 10693 (t996).

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and efficient comparison of the relative affinity of DNA-binding agents for different structural motifs, such as triplex DNA, i-DNA, tetraplex DNA, or Z-DNA and various duplex DNA samples with different base contents. 38 In this method, equal volumes of solutions of different DNA structures at the same concentration are dialyzed against a common solution containing the ligand being studied. After dialysis, the amount of ligand bound to a particular DNA structure is measured by spectrophotometry. The amount of bound ligand is proportional to the association constant for binding of the ligand to this DNA structure. This study has confirmed the structural preference for triplex DNA (represented by a poly(dA) • [poly(dT)]2) of previously studied DNA-binding agents, such as BePI or coralyne, and made possible the discovery of the strong triplex selectivity of some less well-characterized ligands, such as berberine (13) or a novel 3,3'-diethyloxadicarbocyanine (15). 39 Further experiments are required to apply this technique to triplexes with different sequences. Calorimetric Methods

Isothermal titration calorimetry (ITC) provides the only means to determine directly the molar calorimetric binding enthalpy, which can be used to determine the equilibrium binding constant for a bimolecular DNA-ligand interaction. This method has shown that preferential binding of the disubstituted amidoanthraquinone derivative 12 to triplex DNA is enthalpically driven whereas binding of 11 to duplex DNA is entropically driven. 37 This technique has also suggested that the stoichiometry of binding of netropsin to a triplex would be 2 : 1.4° One molecule of netropsin would bind in the Watson-Crick groove, and another would bind in the Watson-Hoogsteen groove. The assignment of this new binding site was suggested by the fact that the AH for binding of netropsin to this second site was exothermic. This is expected for binding of a ligand in a hydrophobic groove by a mechanism involving mostly van der Waals interactions. Gel-Shift Assays

Triplex formation can be detected by gel-shift methods. Binding of the third strand to a radiolabeled duplex results in the formation of a complex with a slower migration rate. Low molecular weight drugs are too small to induce a significant mobility shift when binding to triplex DNA. However, the formation of a triple helix that does not form in the absence of ligand can be detected, and an apparent dissociation constant can be estimated by the concentration of third strand required 38j. Ren and J. B. Chaires, Biochemiso'y 38, 16067 (1999). 39j. Ren and J. B. Chaires, J. Am. Chem. Soc. 122, 424 (2000). 40 D. Rentzeperis and L. A. Marky,J. Am. Chem. Soc. 117, 5423 (1995).

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for 50% triplex formation. Such a method has shown that BeP141 and various polyamine analogs 42 can stabilize triple helices with a GT antiparallel third strand. In the former case, triplex formation could be detected at 4 °, but not at room temperature, although the melting temperature (measured by thermal denaturation experiments) was as high as 51 °. This is probably due to too short a lifetime of the triplex-drug complex at room temperature, which results in dissociation of the complex during gel migration.

Footprinting Methods Footprinting methods provide a powerful tool for investigating the sequence specificity of DNA-binding agents. DNase I, dimethyl sulfate (DMS), diethyl pyrocarbonate (DEPC), osmium tetroxide, EDTA-Fe II, and methidiumpropyl-EDTA (MPE) have been used to probe triplex DNA alone or complexed with triplexbinding agents. Cleavage by DNase I, DMS, or MPE is inhibited by triplex formation itself. In the presence of a specific triplex-binding agent, the concentration of third strand required to produce a footprint is lower than that required in the absence of ligand. For example, DNase I and MPE-Fe I1 footprinting reveal that the binding constant for a 27-mer oligonucleotide is enhanced by a factor of 10 in the presence of the BfPQ derivative 9. 36 As another example, the concentration of a 10-mer oligonucleotide required for 50% inhibition of cleavage by DNase I can be reduced by at least a factor of 100 (i.e., from more than 50 to 0.5 #M) in the presence of a 10 # M concentration of the naphthylquinoline derivative 7. 43 DMS footprinting has been used to show that basic oligopeptides can stabilize triplehelical DNA. 44 It should be mentioned that footprinting methods can also detect structural changes induced by the ligands. For example, an enhanced cleavage of the oligopyrimidine target strand at a specific site, surrounded by footprints on both sides, has been reported when the binding of a GT antiparallel third strand was investigated by DNase I footprinting in the presence of BePI. 41 Enhanced cleavage has also been described at the 5'-thymine of 5'-TTC sequences, when probing a triple helix with a pyrimidine third strand in the presence of BPQ and using MPE as cleaving agent. 36

Molecular Modeling In the absence of high-resolution structural information, quantitative structureactivity relationship (QSAR) studies and/or molecular modeling can be used to 41 C. Escudr, J. S. Sun, C. H. Nguyen,E. Bisagni, T. Garestier, and C. Hrl~ne, Biochemistry35, 5735 (1996). 42 M. Musso, T. Thomas, A. Shirahata, L. H. Sigal, M. W. VanDyke,and T. J. Thomas, Biochemistry 36, 1441 (1997). 43 S. A. Cassidy, L. Strekowski, W. D. Wilson, and K. R. Fox, Biochemistry33, 15338 (1994). 44W.N. Potaman and R. R. Sinden, Biochemistry34, 14885 (1995).

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understand the critical chemical moieties required for preferential binding to triplex DNA. This has been done for related compounds in the benzopyridoindole family, where molecular modeling has provided some insights into possible mechanisms responsible for the different efficiency of the compounds by proposing a model for intercalation of these compounds into triplex DNA. 45 According to this model, the side chain of BePI (1) lies in the Watson-Hoogsteen groove whereas the side chain of BgPI (2) lies in the Watson-Crick groove. A similar model was built for the benzo[f]pyrido[3,4-b]quinoxaline derivative 3. This model suggested that stacking interactions with base triplets could be improved by adding a new ring to the aromatic system. The resulting rational design of a new compound based on this model gave a very efficient triplex-binding agent, BQQ (6). ~9 The same strategy applied to BePI (1) and BgPI (2) led to the synthesis of the new benzoindoloquinoline derivatives 4 and 5. 46 Another study provided a rationale for the different effects of the disubstituted anthraquinone derivatives 11 and 12 on triple helices.47 Molecular modeling suggested that 12 can intercalate with one side chain located in the Watson-Crick groove whereas the other lies in the WatsonHoogsteen groove. The threading intercalation provides good stacking interactions between the chromophore and base triplets. For 11, however, the relative position of the two side chains does not accommodate both intercalation and binding of the third strand. As a result, the triple helix is destabilized. Smface Plasmon Resonance

Surface plasmon resonance has been used to study the kinetics of a pyrimidinemotif triple-helix formation in the presence and in the absence of BPI ligands. Under given conditions, the presence of 6 # M BePI (1) or BgPI (2) accelerates the association rate constant by 27- and 23-fold, respectively, as compared with that of the TFO alone. The dissociation rate constant was decreased by a factor of 1.2 and 4.5, respectively. Therefore, these triplex-specific ligands enhance the binding constant of TFO by 32- and 104-fold, respectively (Sun et al., unpublished data, 1999). The main contribution of BPI ligands to triplex formation is evidently to the association phase. This implies that BPI ligands likely act on the nucleation step by stabilization of the initiation fragment. BgPI derivatives exhibited a larger effect on the dissociation (and thus increased the lifetime of the triplex) than BePI. This can be explained by a better stacking interaction of BgPI with base triplets than BePI. It should be pointed out that the lifetime of the triplex is relevant when the triple-stranded structure must interfere with transcription or other biological 45C. Escud6, C. H. Nguyen,J. L. Mergny,J. S. Sun, E. Bisagni, T. Garestier,and C. H61bne,J. Am. Chem. Soc. 117, 10212 (1995). 46C. H. Nguyen,C. Marchand, S. Delage,J. S. Sun, T. Garestier.C. HEl~ne,and E. Bisagni,J. Am. Chem. Soc. 120, 2501 (1998). 47K. R. Fox, E Polucci, T. C. Jenkins, and S. Neidle, Piw'. Natl. Acad. Sci. U.S.A. 92, 7887 (1995).

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process. Kinetic knowledge of the effect of triplex-specific ligands on triple-helix formation provides insight into their mechanisms of action, and helps to design even better ligands. Targeting Triple-Helical Nucleic Acids with Drugs

Choice of Experimental System Physicochemical methods are generally applicable to homopolymeric nucleic acid structures, mostly poly(dA) • [poly(dT)]2 or poly(dAG) • [poly(dTC)]2 in the case of triple helices. Such experiments are useful for determining the binding mode, observing the specific features of intercalative versus groove-binding compounds, and for assessing cooperative binding. Although they can reveal triplexstabilizing properties, they are poorly predictive of the capability of these compounds to stabilize short triplex sequences containing C. G*C + base triplets at low concentration. For example, coralyne was suggested to stabilize the poly(dAG) • [poly(dTC)]2 triple helix efficiently, but further experiments using short oligonucleotides showed that this effect was weak. 35 The choice of the sequence is also important. Ethidium bromide was shown to stabilize triple helices made o f T . A*T base triplets, but destabilized triple helices containing C • G*C + base triplets. 15This was attributed to electrostatic repulsion between ethidum bromide and the protonated cytosines. Except for hydrodynamic methods and linear dichroism, most physicochemical techniques can be applied to short oligonucleotides, which opens the possibility of exploring various sequence contexts. The use of intramolecular triple helices artificially enhances the stability of triple-helical DNA and ensures the formation of a correctly folded and stoichiometric structure, which can be an interesting advantage, especially in the design of NMR experiments.

Choice of Experimental Conditions Triple helix formation is known to be strongly dependent on pH and ionic conditions. Experimental results have shown that solution conditions also have an important influence on the ability of drugs to bind triplex DNA. Indeed, electrostatic interactions are likely to provide an important contribution to DNA binding. Aliphatic nitrogen atoms, often found on the side chains of intercalating compounds, are generally protonated below pH 8.0. The pKa of aromatic nitrogen atoms can be much lower. For example, the pKa of the aromatic N-10 atom of BePI (1) was measured by recording the absorption spectrum at different pH. A pKa of 6 was measured for the free ligand, which was increased to 8 when the ligand was bound to triple-helical DNA. 25 Binding to DNA can therefore stabilize the form with the highest affinity for DNA, that is, the protonated form. The naphthylquinoline derivative 7 was shown to stabilize a triple helix with a GA third strand only at acidic pH. This was likely due to a requirement for protonation of

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the chromophore. It has also been reported that berenil, a minor groove-binding drug, can stabilize triple-helical DNA at low ionic strength, but destabilizes it at higher ionic strength. 3°

Binding Mode Study The binding mode of various triplex binding agents has been studied using techniques such as those described above. As these are all indirect methods, the binding mode is generally deduced by using several different techniques. Both intercalative and groove-binding mode have been observed for drug binding to triple-helical DNA. The binding mode is usually the same for duplex and triplex DNA, although small differences in the structure of the nucleic acid~trug complex are suggested by circular dichroism for some groove-binding agents, and by linear dichroism for intercalators. DAPI (14) is the only agent that may bind in the minor groove of duplex DNA and intercalate into triplex DNA. 48 NMR has not provided high-resolution structural information about drug-triplex DNA complexes. This may be due to the difficulty of obtaining a triple-helical structure with only one binding site for the ligand. The concentrations used for NMR are usually much higher than the binding affinity for secondary binding sites. Crystals of triplex DNA diffract only at low resolution. Attempts to crystallize triple helical DNA in the presence of triplex-stabilizing agents have not been successful so far.

Structural Specificity Compounds with high selectivity for triplex DNA versus duplex DNA have now been described. However, an in-depth investigation using, for example, the competition dialysis method described above may reveal that such compounds also bind to other types of multistranded nucleic acid structures, for example, tetraplex DNA. The search for compounds specific for triplex DNA is therefore a complicated task. Moreover, different types of triple helices (with a Hoogsteen or a reverse Hoogsteen configuration) can be targeted by a particular drug. The same ligand can stabilize both types of structures (as, e.g., BePI), but the more efficient ligands are not the same for both types of structure. It should be mentioned that triple-stranded structures containing both RNA and DNA strands may also be targeted by drugs. Specific stabilization of such structures has been reported. For example, in the presence of the minor groove binders berenil or DAPI, the poly(rA). [poly(dT)]2 and poly(dT). [poly(rA)]2 triple helices could be formed whereas they did not form in the absence of ligand. 32 Specific and efficient stabilization of RNA-containing triple-helical structures has also been observed with 48 Z. Xu, D. Pilch, A. Srinivasan, W. Olson, N. Geacintov, and K. Breslauer, Bioorg. Med. Chem. 5, 1137 (1997).

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BePI (Escud6 et al., unpublished results, 1994). These results suggest that unpredicted triple-stranded structures involving both RNA and DNA may form within living cells in the presence of some drugs.

Binding Site Study Some qualitative information about the sequence preference of a drug may be gleaned by comparing the effects of the drug on different sequences, using thermal denaturation or footprinting experiments. For example, it has been shown that stabilization by BePI was reduced when T- A*T triplets were substituted by C. G*C + triplets. 16 Also, the naphthylquinoline derivative stabilizes the binding of a T6C 2 oligonucleotide, but not a Y2C 6 oligonucleotide.49 These results have been used to infer that the likely intercalation site for the pyrimidine motif is between T- A*T triplets. Indeed, intercalation does occur between T- A*T triplets, but some experiments have suggested that binding between T. A*T and C. G*C + triplets may be possible, especially at high drug concentration. 35 Few studies have addressed the binding of ligands to triple helices with purine-containing third strands. BePI has been shown to considerably enhance triple helix formation with a GT antiparallel third strand, leading to high stability under physiological conditions. 41 In another study, coralyne was shown to stabilize triple helices made of T. A*T triplets with the third strand in an antiparallel orientation. 35 All these results suggest that different sites within different structural contexts of triple-helical DNA may be targeted by drugs. These sites are still to be characterized. Footprinting methods allow the observation of specific cleavage patterns in the presence of triplex-binding agents (see Footprinting Methods, above). However, the results do not allow accurate mapping of the binding sites for these ligands. Thus, further investigations using established high-resolution structural methods are required to provide accurate information about binding sites.

Designing New Ligands for Triple-Helical DNA Most of the compounds that have been shown to stabilize triple-helical DNA efficiently are positively charged intercalators. The efficiency of triplex stabilization is the result of higher binding affinity for the triplex than for the duplex. This structural specificity can be achieved by the shape of a polyaromatic ring system, but also by electrostatic interactions and steric constraints. In particular, the nature and the position of side chains are likely to have an important influence on triplex-stabilizing properties. Rational design based on experimental studies has led to the synthesis of new compounds that bind to triplex DNA with high 49S. A. Cassidy,L. Strekowski,and K. R, Fox,NucleicAcqdsRes. 24, 4133 (1996).

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affinity and specificity. 19 New strategies are emerging. For example, a dimer of the naphthylquinoline derivative 7 has been designed, synthesized, and studied9 Although the bisintercalation of this compound in triple helices was not conclusively demonstrated, it was shown to stabilize DNA triplexes much more effectively and at much lower concentrations than the monomer. T r i p l e x - B i n d i n g A g e n t s in M o l e c u l a r a n d C e l l u l a r Biology

Improving Antigene Strategy Initial efforts to find triplex-binding agents were motivated by the need to form stable triple helices under physiological conditions. Indeed, several triplex-binding agents have been shown to improve the stability of triple helices. Triplex-forming oligonucleotides can be used to compete with the binding of transcription factors, or to inhibit transcription initiation or elongation by RNA polymerase. Most studies of triplex-stabilizing agents were carried out with pyrimidine-rich third strand. Although efficient stabilization can be obtained, the stabilized triple helices are still pH dependent. Nevertheless, BePI has been shown to improve the TFO-mediated inhibition of in vitro transcription elongation in two different systems.16'51 Another study has suggested that the use of a triplex-stabilizing polyamine enhanced the TFO-mediated inhibition of the c-myc oncogene expression in MCF-7 breast cancer cells, but the evidence of a triplex-based mechanism was lacking in this case. 52 Further in vivo studies are required to establish the potential of triplex-binding agents as antigene enhancers. These studies will also have to deal with the potential toxicity of the compounds. Another interest in triplex-binding drugs stems from their potential to enhance triplex formation on sequences that have interruptions in the target oligopurine • oligopyrimidine sequence. The relative binding strength of different oligonucleotide substitutions could be affected, although the least destabilizing triplets have been shown to be the same in the absence and presence of ligand in two different studies) TM The combined use of a triplex-binding agent and a modified base analog, for example, 3-nitropyrrole, can promote discrimination for a specific base pair inversion, s4 It should be kept in mind that although the use of triplex-stabilizing agents will broaden the range of potential triplextargeted sites, they will also promote the formation of complexes at nontargeted sites. 50 M. Keppler, O. Zegrocka, L. Strekowski, and K. R. Fox, FEBS Lett. 447, 223 (1999). 51 C. Giovannangeli, L. Perrouault, C. Escud6, N. Thuong, and C. H61~ne, Biochemistry 35, 10539 (1996). 52 T. J. Thomas, C. A. Faaland, M. A. Gallo, and T. Thomas, Nucleic Acids Res. 23, 3594 (1995). 53 S. P. Chandler, L. Strekowski, W. D. Wilson, and K. R. Fox, Biochemistry 34, 7234 (1995). 54 S. Kukreti, J. S. Sun, D. Loakes, D. Brown, C. Nguyen, E. Bisagni, T. Garestier, and C. H61~ne, Nucleic Acids Res. 26, 2179 (1998).

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Triplex-Binding Agents and H-DNA It is known that H-DNA can form naturally within supercoiled double-stranded DNA, and formation of this structure has been demonstrated in living bacteria (see Mirkin and Frank-Kamenetskii6). Only one study has addressed the question of the effect of triplex-binding agents on intramolecular triple helices. 55 A mirror repeat sequence was inserted into a plasmid between the Escherichia coli/~-lactamase gene, which confers resistance to the antibiotic ampicillin, and its promoter. This plasmid also carries the resistance gene for the antibiotic tetracycline. It was shown that bacteria transformed with the plasmid containing the mirror repeat sequence grew more slowly than those transformed with the unmodified plasmid in the presence of ampicillin. Both types of cells could grow at the same rate in the presence of tetracycline. Thus, transcription of the/%lactamase gene was apparently inhibited by the mirror repeat sequence, whereas the plasmid could replicate normally. The addition of the triplex-stabilizing agent BePI was shown to further reduce the growth rate in the presence of ampicillin selectively. Chemical probes (chloroacetaldehyde) showed that BePI stabilized the H-DNA structure induced by supercoiling in the plasmid. These experiments suggest that mirror sequences that are able to form H-DNA structures could act as gene regulators, and that regulation of these genes could be influenced by triplex-stabilizing agents. Many sequences that are likely to be able to adopt an H-DNA structure exist in eukaryotic genomes, and many of them are located in gene promoter regions. If these structures do play a functional role, drugs binding to them may serve as tools for investigating this role and may even constitute potential drugs able to interfere with specific biological processes. This has also been postulated in the case of drugs binding to tetraplex DNA. ~6 Some of the triplex-stabilizing agents described above, for example, BePI and coralyne, have cytotoxic properties. Stabilization of intramolecular triple-helical DNA might represent a new mechanism of action for these drugs. Other substances, for example, mithramycin, have been shown to destabilize intermolecular triplexes by binding strongly to duplex DNA. It has been suggested that this antibiotic may inhibit intramolecular triplex formation within a sequence in the c-myc gene. 57 Irreversible Modifications: Triplex-Cleaving Agents Agents that could inflict irreversible damage on triplex DNA would be helpful first as a structural probe for the complexes formed by these agents and the nucleic

55G. Duval-Valentin,T. Debizemont,M. Takasugi,J. L. Mergny,E. Bisagni, and C. H61~ne,J. Mol. Biol. 247, 847 (1995). 56j. L. Mergnyand C. H61bne,Nat. Med. 4, 1366 (1998). 57N. Vigneswaran,C. A. Mayfield,B. Rodu, R. James, H. G. Kim, and D, M. Miller, Biochemistry 35, 1106 (1996).

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acids, and second as a probe for detecting the presence and studying the role of triplex DNA within a biological context. In the presence of iron, a reducing agent, and molecular oxygen, EDTA-Fe H can generate hydroxyl radicals through the Fenton reaction. The covalent attachment of EDTA to a TFO has been used to induce specific cleavage of the targeted double-stranded DNA. 2 EDTA has been covalently attached to various triplex intercalators in our laboratory, namely BePI (1), BgPI (2), and BQQ (6). 58,59 Experiments have been conducted with different double-stranded DNAs containing a putative target for triplex formation, in the presence and absence of third strand. It was shown that strong DNA cleavage was promoted by BgPI-EDTA on both strands within the target site, whereas with BePI-EDTA a slightly enhanced cleavage of the oligopyrimidine strand and a footprint in the oligopurine strand were observed. DNA cleavage was observed all along the triplex sequence, with peaks corresponding to more intense cleavage centered around T-A*T stretches. This suggests that there are various sites for intercalation within the triplex region and that these compounds may exhibit some preferential binding to T. A'T-rich sequences. BQQ-EDTA exhibited the same behavior as BgPI-EDTA, except that it was much more specific than BgPI-EDTA, that is, nonspecific cleavage of double-stranded DNA was reduced. These results are in agreement with results of melting temperature experiments. 19 The observed difference in the pattern and efficiency of cleavage can be explained by the structure of the intercalation complex. Efficient cleavage of both strands is observed when the cleaving moiety is located in the Watson-Crick groove, as with BgPI-EDTA and BQQ-EDTA, whereas with BePI-EDTA hydroxyl radicals must diffuse from the Watson-Hoogsteen groove, an inefficient process that results in some weak cleavage of the more accessible strand, the oligopyrimidine Watson strand (see Fig. 3). These results therefore furnish experimental verification for the model that was proposed on the basis of QSAR and molecular modeling studies. 45 The selectivity of the BQQ-EDTA conjugate for a triplex structure was sufficiently high to induce oligonucleotide-directed DNA cleavage at a single site on a 2718-bp plasmid DNA. Experiments aimed at using this compound to cleave intramolecular triplex DNA in a supercoiled plasmid are under way. More recently, a dibenzophenanthroline derivative, 10, which had been shown to stabilize triple helices,t8 was shown to induce cleavage close to the site of binding of a TFO on irradiation at 360 nm. This cleavage occurred only in the presence of the TFO under conditions favorable for triplex formation. The mechanism of this triplex-mediated photoinduced cleavage is under investigation. 6°

58C. Marchand,C. H. Nguyen,B. Ward,J.-S. Sun,E. Bisagni,T. Garestier, and C. H~16ne,Chem. Eur J. 6, 1559(2000). 59R. Zain, C. Marchand,J.-S. Sun,C. H. Nguyen,E. Bisagni,T. Garestier, and C. H616ne,Chem. Biol. 6, 771 (1999). 6oD. Perrin, et al., in preparation (2001).

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k

k

FIG. 3. A scheme for the cleavage of triple-helical DNA by EDTA-ligand conjugates. The intercalating ligand is shown in gray, with the EDTA-Fe 11moiety represented by a star shape. It generates hydroxyl radicals, which can diffuse over short distances (dashed arrows). Small dark arrows point to the CI' and C4' atoms of deoxyribose. The arrows are located on the side from which the hydrogen atoms (HI' and H4 I) are accessible. Left: The EDTA-Fe II moiety is located in the Watson-Hoogsteen groove, and the HI' and H4' atoms are not readily accessible to hydroxyl radicals, especially that of the Crick strand. Right: The EDTA-Fe n moiety is located in the Watson-Crick groove, and the HI' and H4 f atoms of both the Watson and Crick strands are readily accessible to hydroxyl radicals.

Conclusion Ligands that are selective for triplex DNA can be used to promote triple helix formation, but their interest is much broader as they represent a unique class of structure-specific ligands that could be used as tools to study the biological relevance of triple-stranded structures, or potentially as drugs acting on structurerelated gene regulation. Both high-resolution structural information and thermodynamic as well as kinetic studies still need to be pursued to yield a better understanding of how drugs bind specifically to triplex DNA. Nevertheless, the progress that has been made in characterizing triplex-ligand complexes has led to the design of some specific and efficient triplex-targeted drugs.

Acknowledgments The authors thank all the scientists of the Laboratoire de Biophysique who contributed to the study of triplex-stabilizing agents, and gratefully acknowledge the chemists at the Institut Curie and the Coll6ge de France who synthesized some of the compounds described in this chapter.

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[ 17] M e a s u r e m e n t of Covalent D r u g - D N A I n t e r a c t i o n s at the Nucleotide Level in Cells at P h a r m a c o l o g i c a l l y Relevant Doses By CLAIREJ. MCGURK, PETERJ. MCHUGH, MICHAELJ. TILBY, KEITH A. GRIMALDI,and JOHN A. HARTLEY Introduction Many clinically important anticancer drugs, such as those from the alkylating agent and platinum classes, act by forming covalent adducts with DNA. The majority of these adducts are formed with a base specificity and often show limited sequence specificity. For example, cisplatin [cis-diamminedichloroplatinum II] binds mainly the N7 positions of guanine nucleotides in the major groove of DNA and forms mostly intrastrand cross-links at GG and AG. More recently, novel drugs that show a high degree of sequence-selective binding and alkylation in the minor groove of DNA have emerged based on natural product structures. Attempts to rationally design agents that will target DNA damage to unique sequences are ongoing. Sequence specificity has mainly been determined by examining the lesions produced on isolated DNA. Although important, such analyses are limited. Nucleotidelevel mapping of adducts in cells indicates whether binding to a particular sequence is conserved, because sequence context, methylation, and cellular factors such as nucleosomes and DNA-binding proteins influence drug-DNA interactions. Ligation-mediated polymerase chain reaction (LM-PCR) is the sensitive technique that was first used to map nucleotide-level DNA damage in single-copy genes in mammalian cells after exposure to UV. 1'2 Lesions are cleaved with enzymes or chemical reagents to create strand breaks to which a double-stranded oligonucleotide linker is ligated. The molecules, of varying lengths determined by the location of the lesion, are exponentially amplified, separated on sequencing gels, and detected by hybridization with a fragment-specific probe. For many drug-DNA adducts, however, there are no effective cleavage agents available. Single-strand ligation PCR (sslig-PCR) 3-6 was developed, which relies on the blockage of Taq DNA polymerase at covalent lesions and therefore can be I R R. Mueller and B. Wold, Science 246, 780 (1989). 2 G. P. Pfeifer and A. D. Riggs, Genes Dev. 5, 1102 (1991). 3 K. A. Grimaldi, S. R. McAdam, R. L. Souhami, and J. A. Hartley, Nucleic Acids Res. 22, 2311 (1994). 4 K. A. Grimaldi, S. R. McAdam, and J. A. Hartley, Single-strand ligation PCR for detection of DNA adducts, in "Technologies for Detection of DNA Damage and Mutations" (G. R Pfeifer, ed.). Plenum Press, New York, 1996.

METHODS IN ENZYMOLOGY.VOL. 340

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used to detect most d r u g - D N A adducts. An initial linear PCR step samples each target molecule many times. Ligation of a single-stranded oligonucleotide with T4 RNA ligase facilitates exponential PCR, followed by a third round of linear PCR with a labeled primer. Terminal transferase-dependent PCR (TD-PCR), developed later, is a related technique that has been used to detect cyclobutane pyrimidine dimers 7 and mycotoxin aflatoxin B l addu cts.8 Like sslig-PCR, it involves initial linear PCR and relies on a DNA polymerase blockage at the sites of lesions. However, an additional step utilizes terminal deoxynucleotidyltransferase (TdT) to add ribo-guanines onto the 3' termini. In combination with a double-stranded linker possessing a cytosine overhang, this provides an efficient substrate for T4 DNA ligase, a more efficient enzyme than T4 RNA ligase in this context. The ligated products are amplified and detected in the same way as for LM-PCR. Currently the major drawback to studying nucleotide level damage is the relatively high doses of damaging agent required to generate detectable levels of lesions. The study of drug-DNA interactions at these doses, although generating important comparative information, may not reflect the pharmacological situation. F o r example, the distribution of lesions may be distorted, many biological responses may be impaired or saturated, or the cells may initiate apoptosis. In particular, studies of the repair of lesions at the nucleotide level are difficult at such high drug doses. Antibodies to O6-ethylguanine, 9'1° thymine glycols, I1 and benzo[a]pyrenediol epoxide (BPDE)-guanine adductsl2,13 have been used successfully to purify fragments of DNA for use in PCR, and have provided the sensitivity to detect adducts at the gene level at relatively low doses. In this chapter we describe how the specificity of DNA damage antibodies can be combined with the sensitivity of genomic sequencing methods to study the lesions produced by DNA-damaging drugs at pharmacologically relevant doses at the nucleotide level in single-copy genes in mammalian cells. A modified method adapted from both the TD-PCR and sslig-PCR procedures is described (see Fig. 1), which can be used to study any covalent DNA lesions that

5 K. A. Grimaldi and J. A. Hartley, Methods Mol. Biol. 90, 157 (1997). 6 K. A. Grimaldi, S. R. McAdam,and J. A. Hartley,Methods Mol. Biol. 113, 241 (1999). 7 j. Komuraand A. D. Riggs, Nucleic Acids Res. 26, 1807 (1998). 8 M. F. Denissenko, T. B. Koudriakova, L. Smith, T. R. O'Connor, A. D. Riggs. and G. P. Pfeifer, Oncogene 17, 3007 (1998). 9 K. Hochleitner,J. Thomale, A. Y. Nikitin, and M. E Rajewsky,Nucleic Acids Res. 19, 4467 ( 1991). l0 j. Thomale, K. Hochleitner, and M. E Rajewsky,J. Biol. Chem. 269, 1681 (1994). I l S. A. Leadon and D. A. Lawrence,J. Biol. Chem. 267, 23175 (1992). 12M. E Denissenko, S. Venkatachalam,E. E Yamasaki, and A. A. Wani,Nucleic Acids Res. 22, 2351 (1994). J3 M. E Denissenko, S. Venkatachalam,Y. Ma, and A. A. Wani, Biotechniques 21, 187 (1996).

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Damaged Genomic DNA, Cut with Appropriate Restriction Enzyme(s) /

Linear PCR using [ Biotinylated Primer I.B

Capture on Streptavidin Dynabeads

......

~O

Ribonucleotide Tailing by Terminal DeoxynueleotidylTransferase

rGrGrG Ligation of Double-Stranded Linker [ by T4 DNA Ligase [

C C C[ [

cccl

IrGrGrG Exponential PCR with Upper Oligo Primer [ and Primer 2

CC(~ . . . . . . . . . . . .

[rGrGrG

~ 2 PCR with Radiolabeled Primer 3

I

cccl

............

Run DNA on Sequencing Gel

C C C[

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prevent the progression of a DNA polymerase. An antibody-dependent purification method is also described that can be used in combination with the modified PCR method to study any lesions to which an antibody is available or can be raised. This combined method significantly improves the sensitivity and quality of the results. As an example of the method, the sequence specificity of binding of cisplatin to a single-copy gene sequence in intact cells is illustrated. Method

DNA-Damaging Agent Treatments PCRs are carried out on damaged DNA, undamaged control DNA, and also Maxam and Gilbert-treated DNA to provide a sequence reference. During MaxamGilbert sequencing, 143/zg of unlabeled DNA is chemically cleaved at base-specific sites and subsequently processed by the PCR technique to produce accurate sequencing lanes that will directly correspond to the lanes containing DNA that has been damaged. The method is applicable to any type of cell, including freshly isolated lymphocytes or cell preparations from solid tissue. For comparison with DNA extracted from drug-exposed intact cells, isolated genomic DNA is treated with the drug and subjected to the same analyses. Treatment of Isolated DNA. Genomic DNA is isolated as described below. The drug incubation time with naked DNA will depend on the reactivity of the drug. For many drugs, 1 hr is sufficient. However, when using cisplatin (David Bull Laboratories, Mulgrave, Australia) we have found that increasing incubation times up to 12 hr generates increasing amounts of damage to template DNA. 1. Incubate 3 # g of DNA in 1.5-ml tubes with DNA-damaging agent at 37 ° for appropriate times in I x Teoa [10x Teoa is 250 mM triethanolamine (pH 7.2), 10 mM EDTA] in a total volume of 50 #1. Teoa should be stored at 4 °. 14 M. Maxam and W. Gilbert, Methods"Enzymol. 65, 499 (1980).

FIG. 1. Outline of the PCR procedure. An intrastrand cisplatin adduct in the starting DNA is indicated by a small triangle. Ten cycles of linear amplification of damaged DNA with a 5'-biotinylated primer produce a family of single-stranded molecules, the Y ends of which are defined by the positions at which Taq polymerase has stopped at DNA lesions. The products are captured on streptavidin-coated magnetic Dynabeads and washed to remove any excess genomic DNA. Terminal deoxynucleotidyltransferase adds three ribo-G molecules to the 3' ends of the amplified DNA fragments. A double-stranded linker with a complementary Y overhang of three C's is ligated to the ribo-G tail, facilitating exponential amplification. A final round of PCR with a radiolabeled nested primer enables the products to be visualized when run on a sequencing gel.

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2. Add 50/zl of 0.6 M sodium acetate "drug stop" solution (pH 5.2) and 3 volumes of 95% (v/v) ethanol (-20°). Vortex. 3. Place the samples in a dry ice bath for 10 min and centrifuge for 15 min at 13,000g at 4 °. 4. Wash the DNA pellet with 1 ml of 70% (v/v) ethanol (room temperature) by gently inverting the tube, centrifuging for 5 min at 11,600 g at room temperature, and removing the supernatant. Repeat the wash and dry the DNA pellet under vacuum. 5. Resuspend the DNA in 10/xl of distilled water.

Treatment of Cells. The time of incubation of drug with cells will again depend on the reactivity of the drug. For example, with cisplatin lesions can be detected within a few hours, but with carboplatin 24 hr or more is needed because of its slower reactivity. Some drugs, including cisplatin, are able to bind the proteins present in fetal calf serum. Therefore, when using cells in culture it is advisable at low doses to carry out the drug incubations in serum-free medium to achieve optimal drug binding to DNA. For long incubation times, however, it may be necessary to include fetal calf serum in the drug incubation to ensure cell survival. Treatment of Suspension Culture Cells 1. Count the cells and resuspend at a density of 2 x 106/ml in serum-free tissue culture medium. 2. Add the required amount of drug (diluted in serum-free tissue culture medium) to the wells of six-well fiat-bottomed tissue culture plates. 3. Add tissue culture medium to make the volume up to 1.5 ml. 4. Add 1.5 ml of cell suspension (3 x 1 0 6 cells), mix well, and incubate at 37 ° for the appropriate time. 5. Transfer the cells to 2 x 1.5-ml tubes and centrifuge at 3000g for 5 min at 4 °. 6. Wash out the wells with 1 ml of serum-free tissue culture medium. 7. Remove the supernatant from 2 × 1.5-ml tubes. Resuspend and combine both cell pellets in 1 ml of medium from the wash (step 6) into one 1.5-ml tube. Discard the other 1.5-ml tube. 8. Centrifuge at 3000g for 5 min at 4 °. 9. Remove the supernatant. The cell pellet may be stored at - 2 0 ° at this point until DNA isolation.

Treatment of Adherent Cells 1. Grow cells almost to confluence in the wells of six-well flat-bottomed tissue culture plates.

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2. Dilute the required amount of drug in 2 ml of serum-free tissue culture medium and add to the cells. 3. Incubate at 37 ° for the appropriate time. 4. Remove the drug medium and gently wash the cells three times with 1 ml of fresh tissue culture medium. [Some cells may become detached from the wells. If so, place into 1.5-ml tubes and centrifuge at 3000g for 5 min at 4 °, remove the supernatant, and add to cells harvested from wells by trypsinization (step 5).] 5. Harvest the cells by trypsinization, centrifuge at 3000g for 5 min at 4 °, and remove the supernatant. The cell pellet may be stored at - 2 0 ° at this point until DNA isolation.

Preparation of DNA 1. Isolate DNA by conventional procedures. The isolated DNA must be of a sufficiently high quality for use in PCR. Contaminating proteins may inhibit the progression of Taq polymerase and generate excess background; therefore additional phenol-chloroform and proteinase K steps may be implemented to increase the quality of the DNA) 5 We have used a kit (Wizard Genomic DNA Purification Kit; Promega, Southampton, UK) that generates a relatively high yield of DNA, 15-30/.zg from 3 × 106 cells. We have obtained equivalent results from DNA by using this kit and DNA that has been further purified with phenol--chloroform and proteinase K. However, these extra protein purification steps may be necessary with some DNA isolation procedures and particular cell lines. An RNase incubation step should be included in the protocol. RNase is necessary, especially if DNA is to be measured spectrophotometrically, as excess RNA could contribute to the total measurement of DNA in the sample and thus could lead to an overestimate of the DNA, creating inconsistency between samples. This is particularly important when quantitating lanes and making comparisons, such as in repair experiments. 2. Digest the DNA with a restriction enzyme that cuts at a site between 100 and a few hundred bases downstream of the binding site of the biotinylated primer that is to be used to create a natural stop site in the first PCR step. 3. Check the DNA concentration fluorimetrically. The measurement of DNA by fluorimeter is extremely accurate, especially when measuring small quantities of DNA. 16 For analyzing single-copy genes we have found that 3 #g of DNA produces the optimal signal. If insufficient material is available (e.g., from clinical samples), 15j. Sambrook,E. E Fritsch, and T. Maniatis, "MolecularCloning: A LaboratoryManual," 2nd Ed. Cold Spring HarborLaboratory,Cold Spring Harbor,New York, 1989. t6 D. Kowalski,Anal.Biochem.93, 346 (1979).

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then it is possible to use less DNA and add cycles during the second-round exponential PCR step to compensate. However, nanogram quantities of DNA can be used if studying multiple-copy sequences, such as mitochondrial DNA or DNA from organisms with smaller genomes (e.g., yeast), and still achieve equivalent results.

Polymerase Chain Reaction The following PCR procedure, outlined in Fig. 1, is adapted from both the terminal transferase-dependent PCR (TD-PCR) procedure 7 and the single-strand ligation PCR (sslig-PCR) technique. 3-6 Biotinylated primers are used in the linear PCR step to facilitate purification of amplified DNA fragments from genomic DNA; streptavidin-Dynabeads (Dynal Biotech, Oslo, Norway) prevent nonspecific amplification in further PCR steps. Lengthy ethanol precipitation steps are replaced by simple washes of Dynabeads, thus reducing time required and the possibility of intersample variation. Time-consuming electroblotting and hybridization are replaced with a third round of PCR using a labeled, nested primer. We have found that Taq polymerase produces more discrete bands compared with Vent (exo-) (New England BioLabs, Hitchin, UK). Vent produces no amplification product because of interference with biotin. MgCI2 concentrations for linear and exponential PCR steps must be determined for each set of primers individually; optimization experiments are usually carried out on drug-treated isolated DNA. Gelatin is included in the first-round "linear" PCR to improve specificity, which is important when using a single primer to target a single-copy gene. The conditions shown here are for analyzing the nontranscribed strand of a coding region of the human c-jun gene using primers (JT1, JT2, and JT3) as previously describedJ 7 Oligonucleotides. Oligonucleotides are obtained from MWG (Ebersberg, Germany). All oligonucleotides are stored in small aliquots at - 2 0 °. The double-stranded oligonucleotide linker used is linker y as previously described. 7 Lower Oligo: 5'-AATTCAGATCTCCCGGGTCACCGC This oligonucleotide should be gel or high-performance liquid chromatography (HPLC) purified. It must also be 5' phosphorylated and possess a T-terminal amine group to block self-ligation. The amine should be incorporated at synthesis, but phosphorylation is more complete if carried out after purification. Upper Oligo: 5'-GCGGTGACCCGGGAGATCTGAATTCCC 17 y. Tu, S. Tornaletti, and G. E Pfeifer, E M B O J. 15, 675 (1996).

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This oligonucleotide is complementary to Lower Oligo except for an additional three C's at the 3' terminus. It should also be gel or HPLC purified. Three nested oligonucleotide primers, each 16 to 28 nucleotides long, should be used per reaction. The sequences of the oligonucleotides will depend on the region of interest to be studied. The first oligonucleotide primer should have biotin incorporated at the 5' end (this modification should be incorporated at synthesis). The Tm of the oligonucleotide primers used is calculated according to the manufacturer equation (MWG): Tm

=

69.3 + (0.41 × %GC) - (650/n)

where %GC is the percent GC content and n is the length of the primer. The Tm of the first oligonucleotide primer should usually be between 50 and 60 °. The Tm of oligonucleotide primers 2 and 3 usually falls between 60 and 70 ° and should ideally be as close to the Tm of Upper Oligo as possible (71 °). The sequences of the primers used in the following example are outlined below and are as previously described 17: JT-1.B: 5'-biotin-ACCGGTGCGAGCGAAG

JT-2: 5'-CGTCCTTCTTCTCTTGCGTGGCTCT JT-3: 5'-GGCTCTCCGCCGCCTTCTGGTCTTT

5' Phosphorylation of Lower Oligonucleotide of Linker. Oligonucleotides are phosphorylated with T4 polynucleotide kinase, using kits (GIBCO-BRL, Paisley, UK) with forward reaction buffer. 1. Prepare 200-pmol/#l (200/zM) stocks of upper and lower oligonucleotides with distilled water. 2. To phosphorylate the 5' terminus of the lower oligonucleotide, incubate in a 1.5-ml tube: Lower oligonucleotide (200 pmol]/zl) 11.1 #1 1x Forward reaction buffer [as provided with enzyme; 20/zl 5x is 350 mM Tris-HC1 (pH 7.6), 50 mM MgC12, 500 mM KC1, 5 mM 2-mercaptoethanol] Distilled water 62.9 #1 ATP (100 raM) 1 #1 T4 polynucleotide kinase (10 units/#l) 5 #1 3. Incubate at 37 ° for 2 hr and then heat inactivate the kinase at 65 ° for 20 min. 4. To the phosphorylated lower primers add 11.1 #1 of the upper oligonucleotide (200 pmol/#l). 5. Heat this mixture to 95 ° for 3 rain and gradually cool to 4 ° by placing the sample in a 70 ° removable hot block for 1 min and then placing the hot block into

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a refrigerator. When cool, the linker is stored at - 2 0 ° until ready for use in the ligation reaction.

First-Round Linear Polymerase Chain Reaction. The single biotinylated primer used in the first round of PCR determines in which region and on what strand damage is measured (i.e., a primer complementary to the transcribed strand detects damage on the transcribed strand). All the reagents for the PCR steps are stored at - 2 0 ° (except MgCI2 and gelatin, which are stored at 4°), defrosted if necessary, and kept on ice until needed except Taq polymerase, which is kept at - 2 0 ° until needed. Amplifications are carried out in a thermal cycler with a heated lid (MJ PTC -100: MJ Research, Watertown, MA) so that it is not necessary to use mineral oil. 1. Place 3 ~g of template DNA (or beads-antibody-DNA mixture if using immunoprecipitated DNA) in a 0.5-ml tube. Bring the volume to 10 #1 with distilled water. 2. Prepare a PCR cocktail mix in a 1.5-ml tube, using the volumes given below after multiplying by the number of samples. 10x Reaction buffer [as provided with enzyme; 10x is 4.0/~1 100 mM Tris-HCl (pH 9.0 at 25°), 500 mM KC1, and 1% (v]v) Triton X- 100] MgCI2 (25 mM) 1.6 #1 Gelatin (0.2%, w/v) 4.0 #1 Distilled water 10.4/zl Mixture containing a 2.5 mM concentration of each dNTP 4.0 #1 (Amersham Pharmacia Biotech, Little Chalfont, UK) Oligonucleotide primer 1 (0.6 pmol/#l) 1.0/~1 Place 25.0 #1 of the PCR cocktail mix into each of the sample tubes; mix well. 3. In a separate 1.5-ml tube, dilute Taq DNA polymerase (5 units/ttl; Promega) to 0.2 units//~l in a total volume of 5/~1 per sample, using distilled water. Add 5 #1 (1 unit) of diluted Taq DNA polymerase to each 0.5-ml tube; tap gently to mix. 4. Denature at 94 ° for 5 min, and then cycle 10 times at 94 ° for 1 min, 56 ° for 3 min, and 74 ~ for 2 min. For complete extension of all products, incubate at 74 ° for 5 rain. Denature at 95 ° for 2 min and then cool to 4 °.

Capture of Biotinylated Polymerase Chain Reaction Products 1. Transfer streptavidin-coated magnetic beads (Dynabeads M-280; Dynal Biotech) to 1.5-ml tubes. Use 5/zl (10/zg//zi) per sample plus 5 #l extra. 2. Sediment the Dynabeads in a magnetic rack (e.g., Dynal MPC-E) for 30 sec and remove the supematant. 3. Wash the Dynabeads twice by removing the tubes from the magnetic rack, resuspending the Dynabeads in 200 #1 of 1 x WBB [1 x washing and binding

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buffer; 10 mM Tris-HC1 (pH 7.7), 1 mM EDTA, 2 M NaC1], replacing the tubes in the magnetic rack to sediment the beads, removing the supernatant, and repeating. 4. Resuspend the Dynabeads in 40 #1 of 1 x WBB per sample. 5. Transfer 40-#1 aliquots to fresh 1.5-ml tubes. Sediment the Dynabeads and remove the supernatant from each tube. Remove the 1.5-ml tubes from the magnetic rack. 6. To each 0.5-ml tube containing 40 #1 of products from the first-round PCR add 10 #1 of 5 x WBB [5 x washing and binding buffer; 50 mM Tris-HC1 (pH 7.7), 5 mM EDTA, 10 M NaC1]. Transfer this 50-#1 mixture to each of the 1.5-ml tubes containing prepared Dynabeads. 7. Incubate at 37 ° for 30 rain, agitating occasionally to resuspend the Dynabeads. 8. Wash the Dynabeads three times with 200 #1 ofTE [ 10 mM Tris-HC1 (pH 7.6), 1 mM EDTA] (see step 3). 9. ResuspendtheDynabeadsin50/zlofdistilledwaterandcentrifugeat 11,600g for 10 sec.

TerminalDeoxynucleotidyltransferaseRibo-Tailing. Ribo-G's are added to the 3' ends of the single-stranded products from linear PCR. The majority of the ends will possess three ribo-G's. 7 All the reagents for ribo-tailing and ligation are stored at - 2 0 ° [except polyethylene glycol 8000 (PEG), which is stored at 4°], defrosted if necessary, and kept on ice until needed except for TdT and DNA ligase, which are kept at - 2 0 ° until needed. 1. Place 1.5-ml tubes containing Dynabeads bound to biotinylated PCR products into a magnetic rack and remove the supernatant. 2. Resuspend the Dynabeads in 10/~1 of 0.1 x TE [1 mM Tris-HC1 (pH 7.6), 0.1 mM EDTA]. 3. Prepare a ribo-tail mix in a 1.5-ml tube, using the volumes given below after multiplying by the number of samples. 5 × TdT buffer [as provided with enzyme; 0.5 M potassium 4.00/zl cacodylate (pH 7.2), 10 mM COC12, and 1 mM dithiothreito! (DTT)] rGTP (10 mM; Promega) 4.00 #1 Distilled water 1.33 #1 TdT (15 units/#l; GIBCO-BRL) 0.67 #1 4. Place 10.0 #1 of the ribo-tail mix into the tubes with the Dynabeads and pipette to mix. 5. Incubate at 37 ° for 15 min with occasional agitation. 6. Add 180 #1 of TE [ 10 mM Tris-HC1 (pH 7.6), 1 mM EDTA], place the tubes in a magnetic rack, and remove the supernatant. Wash the Dynabeads again with 200 #I of TE (twice).

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7. Resuspend the Dynabeads in 15/zl of 0.1 × TE [ 1 mM Tris-HCl (pH 7.6), 0.1 mM EDTA].

LigationofDouble-StrandedLinker. The families of single-stranded molecules that possess rGTP tails are ligated to the double-stranded linker. The doublestranded linker has a 3-C single-stranded overhang that is complementary to the three ribo-G tails. The ligation reaction contains PEG, which concentrates the reagents and keeps the Dynabeads partially suspended, as is important when incubating for long periods. 1. Prepare a ligation mix in a 1.5-ml tube, using the volumes given below after multiplying by the number of samples. Ligation buffer [622 mM Tris-HCl (pH 7.6), 126 mM MgC12, 2.7 #1 126 mM DTT, and 12.6 mM ATP] Prepared double-stranded linker (20/zM; see above) 3.0/zl PEG (50%, w/v) 9.1 #1 T4 DNA ligase (20 units//zl; Promega) 0.2/zl 2. Place 15.0 #1 of the ligation mix into the tubes with the Dynabeads from the ribo-tailing reaction (using a cut tip, as the mixture is extremely viscous) and pipette to mix. 3. Incubate at 17° overnight. 4. Add 170/zl of TE [ 10 mM Tris-HC1 (pH 7.6), 1 mM EDTA], place the tubes in a magnetic rack, and remove the supernatant. Wash the Dynabeads again with 200/zl of TE (twice). 5. Resuspend the Dynabeads in 40 #1 of distilled water for PCR. Transfer to 0.5-ml tubes.

Second-RoundExponentialPolymerase Chain Reaction. The procedure is carried out with template DNA bound to Dynabeads. Ten picomoles of second-round nested region-specific primer 2 and Upper Oligo primers are required per sample. For each new primer set, the number of cycles necessary during the exponential step will need to be determined empirically and almost always falls between 16 and 22 cycles. It is important that the number of cycles falls in the exponential range, especially when quantifying bands. 1. Prepare a PCR cocktail mix in a 1.5-ml tube, using the volumes given below after multiplying by the number of samples. 10x Reaction buffer [as provided with enzyme; 10x is 10.0/zl 100 mM Tris-HCl (pH 9.0 at 25°), 500 mM KCI, and 1% (v/v) Triton X-100] MgC12 (25 mM) 8.0/zl Distilled water 20.0 #1

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Mixture containing 2.5 mM each dNTP (Amersham 10.0/zl Pharmacia Biotech) Oligonucleotide primer 2 (10.0 pmol/#l) 1.0/~1 Upper Oligo linker primer (10.0 pmol/#l) 1.0 #1 2. Place 50.0 #1 of the PCR cocktail mix into the 0.5-ml tubes with the Dynabeads from the ligation reaction; mix well. 3. In a separate 1.5-ml tube, dilute Taq DNA polymerase (5 units/#l; Promega) to 0.2 units/#l in a total volume of 10/zl per sample, using distilled water. Add 10 #1 (2 units) of diluted Taq DNA polymerase to each 0.5-ml tube; tap gently to mix. 4. Denature at 94 ° for 5 min, and then cycle 22 times at 94 ° for 1 min, 68 ° for 2 min, and 74 ° for 1 min plus a 1-sec extension per cycle. For complete extension of all products, incubate at 74 ° for 5 min.

5' End Labeling of Primer 3for Third-Round Polymerase Chain Reaction. Five picomoles of oligonucleotides is required per sample for third-round PCR. Primer 3 is phosphorylated with T4 polynucleotide kinase, using kits (GIBCO-BRL) with forward reaction buffer. Once labeled the primer is stored at - 2 0 ° until ready for use in the third-round PCR. 1. To label the 5' terminus of the third-round primer 3, incubate in a 1.5-ml tube: Oligonucleotide primer 3 (10 pmol//zl), per sample (plus one 0.5 #1 extra) 5 x Forward reaction buffer [as provided with enzyme; 5 x is 6 #1 350 mM Tris-HC1 (pH 7.6), 50 mM MgCI~, 500 mM KC1, 5 mM 2-mercaptoethanol] [F-32p]ATP (10 #Ci/#l; Amersham Pharmacia Biotech) 1 #1 T4 polynucleotide kinase (10 units//zl) 1 #1 Bring the volume to 30 #1 with distilled water. 2. Incubate at 37 ° for 30 min. 3. Add distilled water to bring the total volume to 5 #1 per sample (plus one extra). 4. Purify labeled primer from unincorporated nucleotide by centrifuging through a Bio-Spin-6 spin column (Bio-Rad, Hemel Hempstead, UK).

Third-Round Labeling Polymerase Chain Reaction. This is carried out immediately after the second-round exponential PCR, using the labeled nested primer 3. 1. Prepare a PCR cocktail mix in a 1.5-ml tube, using the volumes given below after multiplying by the number of samples.

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10x Reaction buffer [as provided with enzyme; 10x is 1.0 #l 100 mM Tris-HC1 (pH 9.0 at 25°), 500 mM KC1, and 1% (v/v) Triton X-100] MgC12 (25 mM) 1.0 #1 Distilled water 1.8 #1 Mixture containing 2.5 mM each dNTP (Amersham 1.0 #1 Pharmacia Biotech) Labeled oligonucleotide primer 3 (1.0 pmol//zl) 5.0 #1 Taq DNA polymerase (5 units//zl; Promega) 0.2 #1 Tap gently to mix. 2. Place 10/zl of the PCR cocktail mix into the 0.5-ml tubes from the secondround exponential PCR. 3. Cycle five times at 94 ° for 1 min, 68 ° for 2 min, and 74 ° for 1 min plus a 1-sec extension per cycle. For complete extension of all products incubate at 74 ° for 5 min. 4. Centrifuge the 0.5-ml tubes for 10 sec at 11,600g and transfer 100 #1 of supernatant (not Dynabeads) to fresh 1.5-ml tubes. Add 100/zl of distilled water to 0.5-ml tubes to wash the Dynabeads, respin, and transfer 100 #1 of supernatant (not Dynabeads) to the 1,5-ml tubes. 5. Add 20 #1 of 3 M sodium acetate (pH 5.2) and precipitate the DNA with 3 volumes of 95% (v/v) ethanol (-20°). Vortex, and then place the samples in a dry ice bath for 10 min and centrifuge for t5 min at 13,000g at 4 °. 6. Wash the DNA pellet with 200/xl of 70% (v/v) ethanol (room temperature) by gently inverting the tube, centrifuging for 5 min at 11,600g at room temperature, and removing the supernatant. Repeat the wash and dry the DNA pellet under vacuum. 7. Resuspend the DNA in 5 /xl of sequencing gel-loading buffer [96% (v/v) formamide (deionized), 20 mM EDTA, 0.03% (w/v) xylene cyanol, 0.03% (w/v) bromphenol blue]. It is important that the samples be completely and uniformly resuspended. To achieve this, vortex each sample vigorously for at least 10 sec and centrifuge at 11,600g for 10 sec at 4 ° to collect the liquid. Store at - 2 0 °.

Gel Electrophoresis 1. Separate the radiolabeled amplified fragments in a 50 cm x 21 cm x 0.4 mm, 6% (w/v) denaturing polyacrylamide sequencing gel [National Diagnostics, Hull, UK; 5.7% (w/v) acrylamide, 0.3% (w/v) bisacrylamide, 8.3 M urea, 0.1 M Trisborate (pH 8.3), 2 mM EDTA]. 2. Pre-electrophorese the gel for 30 min prior to loading. The gels are run in l x TBE buffer [89 mM Tris-borate, 2 mM EDTA (pH 8.3)] at 1700-2000 V for 2 hr or until the bromphenol blue dye reaches the bottom of the gel.

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3. Denature the samples at 95 ° for 2-3 rain prior to loading, cool on ice, and load samples into individual wells with an elongated, flat tip (Anachem, Luton, UK). 4. After the run, cover the gel with Saran Wrap and place the gel onto Whatman (Maidstone, UK) 3MM paper, supported by a layer of Whatman DE 81 paper, which binds the shorter fragments that pass through the 3MM paper. Dry the gel on a gel dryer (Bio-Rad) under vacuum at 80 ° for at least 1 hr. 5. Expose the gel to X-ray film (X-Omat LS; Eastman Kodak, Rochester, NY) or phosphoimaging screen for the appropriate time. If an intensifying screen is to be used, the film is exposed at - 7 0 ° . Exposure to X-ray film is usually for about 18 hr without the aid of an intensifying screen, and exposure to phosphoimager screens is usually 2 hr to visualize the bands using these conditions. An example of the method is shown in Fig. 2. Cisplatin adducts are clearly seen in the human c-jun gene when naked genomic DNA is treated with 5 #M drug. A 10-fold higher concentration of drug is required to treat cells in order to detect lesions sufficiently above background. However, the pattern of cisplatin binding is preserved in this sequence in intact cells.

Antibody Purification of DNA A problem with existing genomic sequencing procedures such as sslig-PCR or TD-PCR is that at low doses it is difficult to distinguish between bands representing lesions and background. Antibodies to specific lesions bind only fragments of DNA that contain one or more of these lesions. Purification of these damaged fragments provides an enriched template DNA for PCR. Therefore, an antibody purification procedure (Fig. 3) produces a high signal-to-noise ratio that allows bands representing positions of adducts generated at low doses to be determined by these techniques. Intrastrand cisplatin adduct-specific monoclonal antibodies (ICR4, clone CP9/19) were developed in hooded rats as described. 18 These antibodies can also bind the chemically identical intrastrand adducts produced when carboplatin binds DNA. Antibodies to 6-4 photoproducts, thymidine dimers, and antibodies to the 8oxoguanine adduct to study oxidative DNA damage are available commercially (Kamiya Biomedical, Seattle, WA). Antibodies to melphalan adducts are also available. 19 To visualize bands at low doses, the number of cycles necessary during the second-round exponential PCR step may need to be increased relative to unpurified 18 M. J. Tilby, C. Johnson, R. J. Knox, J. Cordell, J. J. Roberts, and C. J. Dean, CancerRes. 51, 123 (1991). 19 M. J. Tilby, J. M. Styles, and C. J. Dean, Cancer Res. 47, 1452 (1987).

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Cells I

Cisplatin (,uM)

Naked DNA II

0

50 100

[ 1 7]

I

0

5

'Full-Length'

FIG. 2. Detection of cisplatin adducts on the nontranscribed strand of a coding region of the human

c-jun gene by the PCR procedure. The bands on the autoradiograph indicated by arrows represent fragments of DNA of varying length, which correspond to the positions of cisplatin intrastrand adducts along the sequence. The intensity of each adduct-specific band directly relates to the frequency of adducts at that particular position. Full-length represents the position of the downstream restriction enzyme cut. K562 cells were treated with the indicated concentrations ofcisplatin for 18 hr, and isolated DNA was treated for 1 hr. The sequences of the oligonucleotide primers used (JT 1, JT2, and JT3) were as previously described. 17

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Damaged GenomicDNA Denature DNA

Bind Adduct Specific Antibody /J~

2, Bind IgG /J~

l

Bind Protein A SepharoseBeads Q

? Spin Beads Down and Wash Supematant Containing Unbound DNA Away LinearPCR using BiotinylatedPrimer 1.B (See Figure 1) FIG. 3. Antibodypurificationof adductedDNA. An intrastrandcisplatinadductin the startingDNA is indicated by a small triangle.

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DNA. However, the high signal-to-noise (S/N) ratio obtained indicates that this can be achieved without a significant increase in background. Although not purified by antibody, the chemically cleaved products produced when DNA is treated for Maxam and Gilbert sequencing can be amplified by using the same number of cycles without affecting the ability to read the sequence. Preparation ofDNA. Both naked treated DNA and cells treated with drug and controls are purified by this procedure. It is important to completely digest the DNA with restriction enzymes to ensure maximum sensitivity. Large uncut fragments may be more difficult to isolate by immunoprecipitation. Restriction enzymes that cut as close to the biotinylated primer binding site as possible should be selected to minimize isolation of fragments of DNA that possess an adduct 3' of the primer-binding site. We have obtained the best results with primer and restriction enzyme combinations that produce a cut immediately upstream of the first biotinylated primer-binding site. If possible, this could be carried out as a one-step double-digest reaction when cutting at the site downstream of the primer-binding site. Alternatively, it may be necessary to perform two separate reactions. Binding of Antibodies to DNA. Both rabbit anti-mouse IgG and rabbit anti-rat IgG can be used to bind the cisplatin-specific antibodies. Antibodies are stored in small aliquots at - 8 0 ° and TN-milk is kept in aliquots at - 2 0 °. 1. Place 3/zg of damaged DNA sample in a 0.5-ml tube. Bring the volume to 10/zl with distilled water. 2. Denature damaged DNA at 100 ° for 3 min, cool on ice for 3 min, and centrifuge at 10,500g for 5 sec. 3. Add 10 #1 of cisplatin monoclonal antibody culture supernatant (10 #g/ml) to the 0.5-ml tube. Incubate for 1 hr at 37 °. 4. Dilute IgG (1.8-2.5 mg/ml; Sigma, Poole, UK) 10-fold with TN-milk [10 mM Tris-HCl (pH 7.5), 140 mM NaC1, 5% (w/v) nonfat dry milk powder, and yeast tRNA (0.1 g/liter; GIBCO-BRL)]. 5. Add 10/zl of diluted IgG to the 0.5-ml tube. Incubate at 37 ° for 1 hr.

Treatment of Protein A-Sepharose. Before incubation with the antibody-DNA mix, the protein A-Sepharose beads must be treated to reduce nonspecific binding. 1. Swell and wash the protein A-Sepharose beads (Sigma) according to the manufacturer instructions and store at 4 ° . 2. Transfer the protein A-Sepharose beads (250 mg/ml) to a 1.5-ml tube. Use 5/~1 per sample plus an extra 10/zl of beads and resuspend in 5/zl of TN-milk per sample plus 10/zl extra of TN-milk (i.e., for 10 samples resuspend 60 ~1 of beads in 110/zl of TN-milk).

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3. Incubate at room temperature for 1 hr with occasional agitation. 4. Centrifuge at 2600g for 30 sec at 4 °, remove the TN-milk supernatant, and resuspend in 5 #1 of fresh TN-milk per sample plus 20 #1 extra of TN-milk.

Capture of Antibody Complex on Beads 1. Spin down and transfer antibody-DNA mixture to flesh 0.5-ml tubes. 2. Add 10 #1 of pretreated beads mixture to each 0.5-ml tube (with a cut tip, as bead mixture is fairly viscous). Incubate at 37 ° for 1 hr with occasional agitation. 3. Centrifuge the 0.5-ml tubes at 2600g for 30 sec and remove 200 #1 of supernatant carefully, leaving behind the white bead pellet. 4. Resuspend the bead mix in 200 #1 of TN [ 10 mM Tris-HC1 (pH 7.5), 140 mM NaCI] to wash the beads, centrifuge at 2600g for 30 sec at 4 °, and remove 200/~1 of supernatant carefully, again leaving behind the white bead pellet. 5. Repeat step 4, first with 200 #1 of TN and again with 200/~1 of TE [10 mM Tris-HC1 (pH 7.6), 1 mM EDTA] and finally with 200 #1 water. 6. Dry the beads-antibody-DNA mixture under vacuum. 7. Resuspend in I 0/~1 of distilled water for use in PCR and store at 4 ° until ready for TD-PCR. The first-round linear PCR is then carried out with the Sepharose beads and antibodies bound to template DNA. Figure 42o shows the results obtained when antibody-purified DNA is processed by the PCR procedure. Adducts are visualized at 5 #M, approximately 10-fold lower than possible with existing procedures (e.g., see Fig. 2). This increased sensitivity demonstrates that there is minimal nonspecific binding of antibody. The evident dose-dependent increase in adduct formation indicates that this technique can be used for quantitative measurement of lesions. Conclusions This modified/shortened protocol PCR protocol based on single-strand ligation PCR (sslig-PCR) 3-6 and terminal transferase-dependent PCR (TD-PCR) v using Taq polymerase works effectively in combination with antibody-purified template DNA to produce a highly sensitive method for detecting covalent lesions at the nucleotide level in intact cells. The sensitivity is such that lesions can be measured at pharmacologically relevant doses and studies of the sequence specificity and repair of individual lesions can be performed. The antibody-PCR combined procedure is extremely sensitive and highly reproducible.

2oD. Rozekand G. P. Pfeifer,Mol. Cell.Biol. 13, 5490 (1993).

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CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

Cisplatin (~uM) 0

[ 17]

5 10 20 40 80 'Full-Length'

FIG. 4. Detection of cisplatin intrastrand adducts by the PCR procedure, using immunoprecipitated DNA as template. The nontranscribed strand of the promoter region of tile human c-jun gene is shown. K562 cells were treated with the indicated concentrations of the cisplatin for 18 hr. Full-length represents the position of the downstream restriction enzyme cut. The sequences of the oligonucleotide primers used (JD 1, JD2, and JD3) were as previously described. 2°

Acknowledgment This work was funded as part of a Program Grant from the Cancer Research Campaign (SP2000/ 0402) and a Cancer Research Campaign studentship to C. J. McGurk.

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INTERACTIONS

377

[18] Targeting DNA through Covalent Interactions of Reversible Binding Drugs B y DAVID E. GRAVES

Introduction The interactions of small molecules with nucleic acids have provoked considerable interest in the field of drug design over the past three decades. This interest is motivated by the unique functional role of DNA in living organisms and the structural features of DNA afforded by the planar stacking of aromatic base pairs perpendicular to the long axis of the double helix. J'2 Perturbations to the sequential readout of the D N A bases and/or the ability of this information to be identified and read by proteins and enzymes result in marked changes in the normal functioning of basic cellular processes such as replication, transcription, and DNA repair processes, making DNA an interesting and unique target for potential chemotherapeutic agents. 3-5 Studies as early as 1947 by Michaelis 6 noted the utility of DNA-binding agents as effective chemotherapeutic agents and later the studies of Kirk, the Kerstens, and Goldberg examining the structural and functional features of the interactions of actinomycin D with nucleic acids predicted the true potential of D N A as a powerful target for disrupting cellular metabolism and terminating replication and/or transcription. 7-9 One of the earliest classes of DNA-binding compounds to be associated with targeting DNA was the class of acridines. In the 1940s, simple acridine-based compounds such as 9-aminoacridine (structure shown in Fig. 1) and proflavin were used as antibacterial agents l°'ll and were subsequently used as probes to examine mutagenesis. 12,13 In 1956, correlations between biological activities and DNA binding were implicated in studies conducted by Peacocke and Skerrett, who I X. Qu and J. B. Chaires, Methods Enzymol. 321,353 (2000). 2 j. B. Chaires, Curr. Opin. Struct. Biol. 8, 314 (1998). 3 H. M. Berman and E R. Young, Annu. Rev. Biophys. Bioeng. 10, 87 (1981). 4 W. D. Wilson and R. L. Jones, Adv. Pharmacol. Chemother 18, 177 (1981). 5 S. Neidle and Z. Abraham, CRC Crit. Rev. Biochem. 17, 73 (1984). 6 L. Michaelis, Cold Spring Harb. Syrup. Quant. Biol. 12, 131 (1947). 7 j. M. Kirk, Biochim. Biophys. Acta 42, 167 (1960). 8 H. M. Rauen, H. Kersten, and W. Kersten, Z. Physiol. Chem. 321, 139 (1960). 9 E. Reich and I. H. Goldberg, Prog. Nucleic Acid Res. Mol. Biol. 3, 182 (1964). l0 C. N. Hinshelwood."The Kinetics of Bacterial Change." Clarendon Press, Oxford, UK, 1946. II A. C. R. Dean and C. N. Hinshelwood. "Growth, Function, and Regulation in Bacterial Cells" Clarendon Press, Oxford, UK, 1966. 12S. Brenner, L. Barnett, E H. C. Crick, and A. Orgel, J. Mol. Biol. 3, 121 (1961). 13K H. C. Crick, L. Barnett, S. Brenner, and R. J. Wattg-Tobin,Nature (London) 192, 1227 (1961).

METHODS IN ENZYMOLOGY,VOL. 340

Copyright ~3 2001 by Academic Press All rights of reproduction in any form reserved. 0076-6879/00 $35.00

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CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

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Sara. L-N-MeVal~

2N~NH2

O

Sar

L-Pro L - P r o

\L-N-MeVal

D-Val D-Val

T.rJ

\Thr

I

J

OC

CO

OCH3

.O

"]~

0

CH3

FlG. 1. Chemical structures of selected DNA-binding agents used to study nucleic acid structure and function. Note the similar heterocyclic chromophores of ethidium (left), 9-aminoacridine (center), and actinomycin D (right).

demonstrated that the antibacterial activity of proflavin was directly related to the interaction of acridine with DNA.14 The intercalative mode of binding of acridines to DNA, originally proposed by Lerman in 1961, is now well characterized and a common binding motif for many anticancer agents. 15 The cytotoxic properties observed for DNA-binding agents have led to the extensive use of these compounds as chemotherapeutic agents in the treatment of numerous diseases including cancer16; hence, the mechanism through which small molecules such as chemotherapeutic agents, mutagens, and carcinogens interact with nucleic acids is of central biological significance. Biophysical studies correlate the physicochemical properties associated with formation of these complexes with various biological effects that are induced by these DNA-binding agents, such as inhibition of DNA replication and transcription, topoisomerase II inhibition, mutagenesis, and carcinogenesis. 17 Significant progress has been made toward unraveling the structural, thermodynamic, and kinetic properties associated with the interactions of ligand-DNA complexes. These studies provide pivotal insight into the design and development of more effective second- and third-generation chemotherapeutic agents that are being used in the successful treatment of many types of diseases; however, despite extensive research efforts, many molecular mechanisms responsible for eliciting the biological actions remain unknown. A key problem associated with examining the interaction of a drug with its DNA-binding site centers on the reversible nature of the ligand-DNA interaction; 14 A. R. Peacocke and N. J. H. Skerrett, Trans. Faraday Soc. 52, 261 (1956). 15 L. S. Lerman, J. Mol. Biol. 3, 18 (1961). 16 M. J. Waring, Annu. Rev. Biochem. 50, 159 (1981). 17 A. H.-J. Wang, in "Nucleic Acids and Molecular Biology" (F. Eckstein and D. M, Lilley, eds.), Vol. 1, pp. 32-54. Springer-Verlag, New York, 1987.

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hence limiting the microscopic characterization of the complex. 18,19 One strategy used to circumvent this problem is photoaffinity labeling. 2° The usefulness of photoreactive agents as tools to probe structural and/or functional properties of the active sites of proteins has been well documented for more than three decades; however, relatively few applications of photoaffinity labeling have been directed toward the study of ligand-DNA interactions. 21 In the 1970s, Yielding and coworkers realized the usefulness of this method and successfully developed photoreactive analogs of ethidium and acridines as tools for examining ligand-DNA interactions. 22'23 Their studies demonstrated the utility of this method in characterizing ligand-DNA interactions and were extended toward the examination of mitochondrial mutagenesis, bacterial frameshift mutagenesis, and eukaryote DNA repair processes. 24-27 Studies by Graves and Yielding, using a variety of thermodynamic and kinetic measurements, determined that placement of the azido moiety on the phenanthridine ring (ethidium monoazide) did not significantly alter the intercalative DNAbinding properties of the ethidium photoreactive analogs from those of the parent ethidium.28'29 The photoreactive ethidium monoazide was shown to exhibit DNAbinding affinity, binding site size, and base-sequence specificity (in the absence of light) equivalent to those of the parent ethidium bromide; however, on photolysis by visible light, the azide moiety was converted to a highly reactive nitrene resulting in formation of a covalent bond in situ. 3° Over the past two decades, this laboratory and others have exploited this facility for covalent attachment to DNA of ethidium and other photoaffinity analogs of DNA-binding agents in an effort to more closely examine DNA-binding specificities at markedly lower ligand concentrations than had been possible with studies involving the reversibly binding parent molecules. Through covalent attachment of the ligand, much greater sensitivity in characterizing the sequence-selective binding of ligands to DNA is achieved. 31,32 We have extended these studies to include photoaffinity analogs of actinomycin 18 D. E. Graves, in "Advances in DNA Sequence Specific Agents" (L. Hurley and J. Chaires, eds.), Vol. 2, pp. 169-186. Jai Press, Greenwich, Connecticut, 1996. 19j. R. Knowles, Acc. Chem. Res. 5, 155 (1972). 20 E. Neilson and O. Buchardt, Photochem, Photobiol. 35, 317 (1982). 21 W. Lwowski, Ann. N.Y. Acad. Sci. 346, 491 (1980). 22 D. E. Graves, L. W. Yielding, C. L. Watkins, and K. L. Yielding, Biochim. Biophys. Acta 479, 98 (1977). 23 S. C. Hixon, W. E. White, Jr., and K. L. Yielding, Biochem. Biophys. Res. 66, 31 (1975). 24 W. E. White, Jr., and K. L. Yielding, Methods Enzymol. 46, 644 (1977). 25 M. Fukunaga and K. L. Yielding, Mutat. Res. 80, 91 (1981). 26 S. C. Hixon, W, E. White, Jr., and K. L. Yielding, J. Mol. Biol. 92, 319 (1975). 27 L. W. Yielding, W. E. White, Jr., and K. L. Yielding, Mutat. Res. 34, 351 (1976). 28 L. W. Yielding, D. E. Graves, and B. R. Brown, Biochem. Biophys. Res. Commun. 87~ 424 (1979). 29 O. E. Graves, C. L. Watkins, and L. W. Yielding, Biochemistry 20, 1887 (1981). 30 N. Turro, Ann. N.E Acad. Sci. 346, 1 (1980),

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D (7-azidoactinomycin D) and 4'-(9-acridinylamino)methanesulfon-m-anisidide (m-AMSA)(3-azido-m-AMSA) to probe ligand-DNA interactions. 33-35 DNA I n t e r c a l a t o r s Intercalating agents constitute an important class of drugs that have been used for decades to probe DNA structure and function and in the treatment of various diseases including cancer. 36 The intercalative mode of binding involves the insertion of the planar polycyclic aromatic chromophore of the ligand between adjacent base pairs of the DNA duplex. Besides the chromophore involvement in binding, additional chemical substituents residing on the ligand are highly influential in determining the chemical properties such as charge, hydrophobicity, and aqueous solubility that in turn direct the thermodynamic binding mechanism(s), the geometry of the ligand-DNA complex, and the sequence selectivities observed for many of these agents. 37-39 There are numerous intercalators classified according to the structure of the chromophore, as shown in Fig. 1. Classic intercalators such as ethidium bromide and proflavin were among the first compounds to be studied with regard to their abilities to interact with DNA. Pioneering studies of Lerman4° revealed that addition of acridine, proflavin, or acridine orange to DNA resulted in marked changes in the viscosity and sedimentation coefficients. These observations led Lerman to predict that the planar acridine chromophore inserted between adjacent base pairs of the DNA helix, resulting in perturbations to the helical structure of the DNA. In 1965, the first biophysical characterization of the ethidium-DNA complex was presented by Waring, 41 who described the intercalative interaction of ethidium with DNA by using a variety of spectrophotometric methods. In 1967, these studies were extended by LePecq and Paoletti to include the characterization of the ethidium-DNA complex by fluorescence spectroscopy.42 Over the next several decades, numerous investigators have probed the interactions of small 3t j. M. Hardwick, R. S. von Sprecken, K. L. Yielding, and L. W. Yielding, J. Biol. Chem. 259, 11090 (1984). 32 R. L. Rill, G. A. Marsch, and D. E. Graves, J. Biomol. Struct. Dynam. 7, 591 (1989), 33 R. M. Wadkins and D. E. Graves, Biochemistry 30, 4277 (1991). 34 G. W. Rewcastle, W. A. Denny, W. R. Wilson, and B. C. Baugley, Anti-Cancer Drug Des. 1,215 (1986). 35 T.-L. Shieh, P. Hoyos, E. Kolodziej, J. G. Stowell, W. M. Baird, and S. R. Byrn, J. Med. Chem. 33, 1225 (1990). 36 M. J. Waring, Annu. Rev. Biochem. 50, 159 (1981). 37 j. B. Chaires, Anti-Cancer Drug Des. 11, 569 (1996). 38 R. M. Wadkins and D. E. Graves, Biochemiso 3' 30, 4277 (1991). 39 j. M. Crenshaw, D. E. Graves, and W. A. Denny, Biochemistry 34, 13682 (1995). 4o L. S. Lerman, J. Mol. Biol. 3~ 18 (1961). 41 M. J. Waring, J. Mol. Biol. 13, 269 (1965). 42 j. B. LePecq and C. Paoletti, J. Mol. BioL 27, 87 (1967).

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38 1

molecules with nucleic acids by a variety of spectroscopic and crystallographic methods and have provided pivotal information as to the mechanisms of action of numerous biologically active compounds used in the treatment of a variety of diseases, including cancer. From these studies key insights have been gained regarding sequence-selective interactions of small molecules and proteins with DNA, thermodynamic and kinetic properties associated with the interactions of these compounds with DNA, and the abilities of these agents to alter DNA structures and/or enzymatic activities. The vast majority of these studies have dealt with reversibly binding drugs; hence, precluding a detailed characterization of the structural features and perturbations induced by formation of the ligand-DNA complex. Hence, in an effort to circumvent many of the problems associated with characterization of the ligand-DNA complex formed by reversible binding compounds, novel approaches such as photoaffinity labeling were applied. The remainder of this chapter describes the use of photoreactive analogs of several DNA-binding drugs that interact with DNA in a reversible manner. P h o t o r e a c t i v e A n a l o g s of E t h i d i u m Ethidium bromide and 9-aminoacridine are paradigms of DNA-intercalating agents. Biological activities of these compound have been known and exploited for more than half a century because of their propensities for inhibiting nucleic acid synthesis in a variety of parasitic organisms. 43-47 The trypanocidal activity of ethidium was linked to its inhibition of DNA polymerase as early as 1963 and postulated to arise from its interaction with nucleic acids. 48'49 Acridine, first isolated from coal tar more than a century ago, has been used extensively in the development of biologically active analogs, such as 9-aminoacridine and proflavin, that came into broad clinical use as antibacterial agents during World War II. 5°'51 Antimalarial qualities of quinacrine were recognized in the 1930s and the drug continues to be used in the treatment of this disease. 52 Both ethidium and the acridine analogs have been extensively used as DNA-intercalating agents to probe

43 B. A. Newton, Annu. Rev. Microbiol. 19, 209 (1965). 44 D. Kerridge, J. Gen. Microbiol. 42, 71 (1966). 45 j. E Henderson, Prog. Exp. Tumor Res. 6, 84 (1965). 46 R. Tomchick and 3H. G. Mandel, J. Gen. Microbiol. 36, 225 (1964). 47 B. A. Newton, in "Metabolic Inhibitors" (R. M. Hochster and J. H. Quastel, eds.). Academic Press, New York, 1963. 48 M. J. Waring, Biochim. Biophys. Acta 87, 358 (1964). 49 W. n. Elliott, Biochem. J. 86, 562 (1963). 50 A. C. R. Dean, in "Acridines" (R. M. Acheson, ed.), pp. 789-814. John Wiley & Sons, New York, 1975. 51 W. Schulemann, Proc. R. Soc. Med. 25, 897 (1931). 52R. M. Pinder, in "Medicinal Chemistry" (A. Burger, ed.), pp. 508-621. Wiley-lnterscience, New York, 1970.

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and characterize DNA structure and function; however, investigation using these agents is limited because of the reversible nature of the ligand-DNA complexes. Photoaffinity labeling, a technique that involves the design and synthesis of analogs with the ability to covalently attach at their binding sites when photoactirated, provides a means of overcoming the limitation of reversible binding. Several photoreactive analogs of ethidium have been synthesized by replacing either or both of its amino groups at the 3- or 8-positions of the phenanthidine ring with an azido substituent (structures shown in Fig. 2A and B ) . 22'53 A

3~:H:anal o og H 2 N ~ (~

C2H5

NH2

N 3 ~

NH2

~H~_..~ N 8-~C a2zid~analog

etparent hidium C2H5 "~.~ 3,8diazidanal o og FIG. 2. (A) Conversion of the parent ethidium (left) to photoreactive analogs (right). Chemical synthesis involves conversion of one or both of the amino groups to the diazonium salt followed by substitution to the azido moiety. The three photoreactive analogs generated are 3-azido8-aminophenanthridinium chloride (top right), 8-azido-3-aminophenanthridinium chloride (middle right), and 3,8-diazidophenanthridinium chloride (bottom right). For synthesis details, see Graves et al. 10°. (B) Conversion of the parent acridine (left) to photoreactive analogs (Hght) including 9-azidoacridine (top right), 3-azido-6-aminoacridine (middle right), and 3-azido-6-amino-10methylacridine (bottom right). For synthesis details, see Graves et al. i00

[18]

COVALENT CROSS-LINKING OF L I G A N D - - D N A INTERACTIONS

383

B ]3

H 9-azd io

I

H2N~N3 H]

H

3-azido-6-amino

acridine

H2N~N [

CH3

3

3-azido-6-amino-

10-methyl

FIG. 2. (continued)

The value of ethidium as a photoaffinity analog is reflected by its ability to mimic the noncovalent DNA interaction of the parent ethidium bromide in forming reversible complexes (in the absence of light) identical to that of the parent compound. Biophysical data comparing the properties associated with these interactions are provided in Table I. Comparative studies of the interactions of the parent ethidium and the 8-azido analog (ethidium monoazide) with DNA reveal that the interactions of ethidium monoazide closely mimic those of the parent compound. 29 In contrast, the structural conformer, 3-azido-8-amino-5-ethyl-6phenyl-phenanthridinium chloride, was reported by LePecq and co-workers to bind DNA with an orientation that was inverted compared with that of the parent ethidium, with the phenyl and ethyl moieties residing in the helical major groove, in contrast to the minor groove interaction of the parent ethidium and ethidium monoazide. 54 53 W. J. Firth, C. L. Watkins, D. E. Graves, and L. W. Yielding, J. Heterocyclic Chem. 20, 759 ( 1981 ). 54 R Laugaa, A. Delbarre, J. B. Le Pecq, and B. P. Roques, Eur J. Biochem. 134, 163 (1983).

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C H E M I C A L AND M O L E C U L A R B I O L O G I C A L A P P R O A C H E S

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TABLE 1 BIOPHYSICAL PROPERTIES OF PHOTOREACTIVE ANALOGS OF ETHIDIUM BROMIDE

Characteristic

Parent ethidium

Ethidium monoazide (8-azido-3-amino)

Ethidium diazide

Visible properties •~max )~max + DNA

Efree ~°bound Fluorescence properties )-max em ~-max em + DNA

Relative intensity Relative intensity+DNA Pbotolysis rate

480 nm 518 nm 5680M 1 cm-I 2500 M - 1 c m - I

458 nm 495 nm 5 2 2 0 M - t cm 1 3560M tcm-l

432 nm 445 nm 5850M-1 cm-i 3880M -lcm t

6 0 0 nm 590 n m 1 21

590 nm 580 nm 1 21 K = 5.7 x 1 0 - 6 s e c - I (at80J I M-2sec-l

500 nm 507 nm 1 0.1 K = 6.1 x 10 5sec I (at80J-I M-2sec I

Yielding and co-workers revealed that the interactions of the ethidium monoazide with DNA resulted in relatively large increases in the fluorescence emission signal on complex formation, suggesting that the monoazide intercalates in a manner similar to that of the parent compoundfl 9 Stopped-flow studies revealed that the parent ethidium and photoreactive ethidium azide exhibit identical apparent rate constants for the dissociation of these drugs from D N A Y Similarly, the intrinsic binding constants and site exclusion sizes for the two compounds are identical, demonstrating the usefulness of the 8-azido-3-aminoethidium analog to characterize ethidium-DNA interactions as described in Table I. The utility of ethidium monoazide to probe biological functions such as mutagenesis and DNA repair processes has been well documented by Yielding and co-workers. Their studies demonstrate that covalent attachment of the ethidium analog to DNA resulted in induction of petite mutants for yeast mitochondria, frameshift mutagenesis in Salmonella, and repairable DNA lesions in human lymphocytes 56-6° Control experiments with the parent ethidium bromide and/or ethidium monoazide under nonphotolyzed conditions did not elicit these biological effects, indicative of a requirement for covalent attachment for their induction.

55 E Garland, D. E. Graves, L. W. Yielding, and H. C. Cheung, Biochemistry 19, 3221 (1980). 56 S. C. Hixon, W. E. White, Jr., and K. L. Yielding, J. Mol. Biol. 92, 319 (1975). 57 L. W. Yielding, W. E. White, Jr., and K. L. Yielding, Murat. Res. 34, 351 (1976). 58 C. E. Cantrell, K. M. Pruitt, and K. L. Yielding, Mol. Pharmacol. 15, 322 (1979). 59 M. Morita and K. L. Yielding, Mutat. Res. 54, 27 (1978). 60 M. Fukanaga and K. L. Yielding, Biochem. Biophys. Res. Commun. 84, 501 (1978).

[ 18]

COVALENT CROSS-LINKING OF L I G A N D - - D N A INTERACTIONS

385

In 1985, Kulkarni and Yielding demonstrated that DNA lesions produced by photolysis of ethidium in intact human lympocytes were alkali labile. 61 Alkali treatment of the ethidium-DNA adduct resulted in fragmentation of the DNA, presumably at the site of covalent modification. This discovery was coupled with emerging advances in DNA sequencing methodology and allowed the exact location of DNA lesions resulting from covalent attachment of the drug (alkylation) to be mapped. Application of this method was further refined by Rill and co-workers, using piperidine treatment of the ethidium-modified DNA adducts to generate DNA strand breaks at the site of covalent modification. 32'62Covalently attached ethidium was initially probed by this method; however, the method was rapidly extended to include photoreative analogs of 9-aminoacridine, quinacrine, actinomycin D, and m-AMSA. Because the ligand is irreversibly bonded to the DNA, the total ligand concentration could be decreased by 100- to 1000-fold compared with the reversible-binding parent compound. Hence, footprinting studies used to discern DNA sequence preferences were markedly more sensitive, allowing observations of novel high-affinity sites. Footprinting studies with the parent ethidium reported this compound to bind DNA in a random manner with no explicit preferences for DNA base sequence. However, studies by Rill and co-workers, employing covalently attached ethidium at low ligand-DNA binding densities, indicated a nonrandom distribution of binding sites, with the order of preference for covalent attachment being G > C, T > A. Analyses of neighboring base effects showed that guanine adducts were most reactive when flanked on the 5' side by another guanine and on the 3' side by either guanine or thymine. Cytosine adducts were most reactive when flanked on the 5' side by another cytosine and on the 3' side by guanine. Interestingly, the reactivities of ethidium with the d(CpG) and d(GpG) sequences were predicted from optical and nuclear magnetic resonance (NMR) studies of Reinhardt and Krugh. 63-65 Overall analyses of all possible intercalation sites revealed a binding order of d(CpG) ~ d(GpG) > d(GpC) ~ d(TpG) d(GpT) > d(GpA) ~ d(ApG) "-- d(TpA) > d(ApT) > d(ApA). 32'62 Ethidium Monoazide as Probe for Topoisomerase H-Mediated DNA Strand Cleavage Many compounds used in the treatment of cancer have the enzyme topoisomerase II as their molecular target. 66 This enzyme is responsible for controlling

61 M. S. Kulkarni and K. L. Yielding, Chem. Biol. Interact. 56, 89 (1985). 62 G. A. Marsch, D. E. Graves, and R. L. Rill, Nucleic Acids Res. 23, 1252 (1995). 63 C. G. Reinhardt and T. R. Krugh, Biochemistry 17, 4845 (1978). 64 R. V. Kastrup, M. A. Young, and T. R. Krugh, Biochemistry 17, 4855 (1978). 65 C. G. Reinhardt and T. R. Krugh, Bioehemistry 16, 2890 (1977). 66 S. J. Froelich-Ammon and N. Osheroff, J. Biol. Chem. 270, 21429 (1995).

386

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

[1 8]

the topology of the DNA in cells through enzyme-mediated double-strand breakage concomitant with strand passage of an intact DNA duplex through the break and followed by religation of the double-strand DNA break. Through this catalytic cycle, the enzyme controls over- and underwinding of the DNA duplex and can alleviate torsional stress that accumulates ahead of the replication forks. 67'68 The mechanism(s) through which topoisomerase II-targeting agents such as m-AMSA, daunomycin, Adriamycin (doxorubicin), mitoxantrone, actinomycin D, and ellipticine influence the activity of topoisomerase II remain unknown despite more than two decades of intense investigation. 69 Interestingly, these compounds are all classified as reversible DNA-binding agents, with their presumed mode of DNA binding via intercalation. However, not all DNA-binding agents are capable of altering the catalytic activity of topoisomerase II. While m-AMSA is a potent topoisomerase II inhibitor and its presence results in the enhanced stimulation of topoisomerase II-mediated single- and double-strand breaks, the structural conformer o-AMSA is ineffective in eliciting such a response. 7° In this case, both m-AMSA and o-AMSA bind to DNA through intercalation. In addition, o-AMSA h~s been shown to have a binding affinity that is four times greater than that of m-AMSA. 38 A second example of a DNA-intercalating drug that does not influence the catalytic activity of topoisomerase II is the classic intercalator, ethidium bromide. Early studies of Ross and co-workers demonstrated ethidium to be ineffective in eliciting topoisomerase II-mediated DNA cleavage at drug concentrations up to 100/zM. Indeed, the stimulation of topoisomerase II-mediated DNA cleavage by m-AMSA could be reversed with increasing concentrations of ethidium. 71 Hence, intercalation does not appear to be an absolute requirement for the modulation of topoisomerase II activity. In an effort to probe the influence of intercalative DNA binding on modulating topoisomerase II activity, photoaffinity labeling with ethidium monoazide was applied. Ethidium monoazide was chosen for these experiments because of its similarities in DNA binding to that of the parent ethidium bromide. 2~'22 Ethidium monoazide forms a complex with DNA (in the absence of light) that is identical to that of the parent ethidium. However, on photolysis with visible light, the azido moiety is activated to the reactive nitrene and covalent attachment of the ethidium in s i t u is achieved. These data (shown in Fig. 3) demonstrated that covalent attachment of ethidium to DNA resulted in stimulation of the topoisomerase II-mediated cleavage reaction, markedly enhancing both double- and single-strand breaks in the DNA at low covalently attached drug densities. A comparable level of 67j. C. Wang,Annu. Rev. Biochem. 65, 635 (1996). 68N. R. Cozzarelliand J. C. Wang,"DNATopologyand Its BiologicalEffects."Cold Spring Harbor LaboratoryPress, Cold Spring Harbor,New York, 1990. 69A. Y. Chen and L. F. Liu, Annu. Rev. Pharmacol. Toxicol. 34, 191 (1994). 70E. M. Nelson, K. M. Tewey,and L. E Liu, Proc. Natl. Acad. Sci. U.S.A. 81, 1361 (1984). 71W. E. ROSS,W.C. Rowe, B. Glisson, J. Yalowich,and L. R. Liu, CancerRes. 44, 5857 (1984).

[ 1 8]

387

COVALENT CROSS-LINKING OF L I G A N D - D N A INTERACTIONS

35 Single-Strand DNA Breaks

30 t~

~* 25 o < z

a

20

"0

=

10

0

o

~-

5 0 0

20

40

60

80

Covalently Attached Drugs Per Plasmid FIG. 3. Influence of covalently attached ethidium on the enhancement of topoisomerase If-mediated single-strand (©) and double-strand (ff]) DNA breaks. Control experiments using nonphotolyzed ethidium monoazide ( 0 ) and parent ethidium bromide (11) show no enhancement. The cleavage reaction was carried out as described in Marx et al. 74

single- and double-strand cleavage is observed at bound drug concentrations that are 150 times lower than required for equivalent cleavage stimulation by the known topoisomerase II poisons, m-AMSA and etoposide. 72'73 The capacity for covalent attachment of the drug to the DNA provides novel insight into the mechanism(s) through which DNA-binding drugs may influence topoisomerase II activity and provides details of the structural and functional properties of the ternary complex. The study revealed that covalent attachment of ethidium to the DNA resulted in the conversion of an intercalative binding ligand that was ineffective in modulating topoisomerase II activity to one that was highly active in enhancing topoisomerase II-mediated DNA single- and double-strand breaks. Covalent attachment of the ligand to the DNA prior to addition of the enzyme allowed control of the assembly of the ternary complex. In addition, insight into the recognition of DNA structural and/or stability properties by the enzyme was explored through studies comparing the binding affinities of the enzyme to native and ethidium-modified DNAs. These experiments revealed that the enzyme recognizes and binds to modified 72 M. J. Robinson and N. Osheroff, Biochemistry 29, 2511 (1990). 73 N. Osheroff, Biochemistry 28, 6157 (1989). 74 G. Marx, H. Zhou, D. E. Graves, and N. Osheroff, Biochemistry 36, 15844 (1997).

388

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES NHSO2CH3

NHSO2CH 3

NHSO2CH 3

NH

NH

NH

H

H

H

m-AMSA

o-AMSA

[1 8 ]

3-azido-m-AMSA

FIG. 4. Structures of the acridine-based topoisomerase II poison, m-AMSA (left); the inactive conformer, o-AMSA (center); and the photoreactive analog, 3-azido-m-AMSA (right).

DNA (1 drug covalently bonded per 60 base pairs) with a binding affinity 5-fold greater than that of native DNA, indicative of the importance of DNA structural perturbations in ternary complex formation.74 P h o t o r e a c t i v e A n a l o g s of m-AMSA (Amsacrine) In the early 1970s, Cain and co-workers at the Cancer Research Laboratory in Auckland reported the synthesis of an anilinoacridine analog [4'-(9-acridinylamino)methanesulfon-m-anisidide] along with its enhanced activity against a variety of experimental tumors and leukemias (structure shown in Fig. 4). 75-77 Subsequent studies by Kohn and co-workers revealed m-AMSA to induce the formation of protein-associated single- and double-strand breaks in nuclear DNA through a ternary complex formed between the m-AMSA, DNA, and topoisomerase II. 78-8° The exact nature of the spatial orientation and role of m-AMSA within the ternary complex remained unknown until photoaffinity labeling using the photoreactive analog 4'-(3-azido-9-acridinylamino)methanesulfon-m-anisidide (3-azidom-AMSA) was applied by Freudenreich and Kreuzer. sl Prior biophysical studies 75 B. E Cain, G. J. Atwell, and W. A. Denny, J. Med. Chem. 18, 1110 (1975). 76 S. W. Hall, J. Friedman, and S. S. Legha, Cancer Res. 43, 3422 (1983). 77 W. Marsoni and R. Wittes, Cancer Treat. Rep. 68, 77 (1984). 78 y. Pommier, M. R. Mattem, R. E. Schwartz, L. A. Zwelling, and K. W. Kohn, Biochemistry 23, 2927 (1984). 79 j. Minford, Y. Pommier, J. Filipski, K. W. Kohn, D. Kerrigan, M. R. Mattem, S. Michaels, R. Schwartz, and L. A. Zwelling, Biochemistry 25, 9 (1986). so y. Pommier, J. Covey, D. Kerrigan, W. Mattes, J. Markovits, and K. W. Kohn, Biochem. Pharmacol. 36, 3477 (1987). st C. H. Freudenreich and K. N. Kreuzer, Proc. Natl. Acad. Sci. U.S.A. 91, 11007 (1994).

[18]

COVALENTCROSS-LINKINGOF LIGAND-DNA INTERACTIONS

389

by Byrn and co-workers demonstrated that DNA-binding properties of 3-azido-mAMSA were comparable to those of the parent m-AMSA. 82 In 1994, Freudenreich and Kreuzer extended the utility of photoaffinity labeling using 3-azido-m-AMSA to provide the first biophysical characterization of a ternary complex formed between the drug, DNA, and topoisomerase II. Prior to photolysis, the 3-azido-m-AMSA demonstrated stimulation of topoisomerase II-mediated DNA strand breaks comparable that of the parent m-AMSA. However, on photolysis, the photoreactive ligand covalently bonded to the substrate DNA. Mapping of the DNA-binding site of the covalently attached ligand revealed the drug to be covalently bonded to the DNA bases immediately adjacent to the two phosphodiester bonds that were cleaved by the enzyme. From these studies, Freudenreich and Kreuzer demonstrated that topoisomerase II created and/or stabilized preferential DNA-binding sites for the inhibitor precisely at the two sites of enzyme-mediated DNA cleavage. 81 P h o t o r e a c t i v e A n a l o g s of A c t i n o m y c i n D Actinomycin D (structure shown in Figs. 1 and 5) has long been a paradigm for sequence-selective binding to DNA. Early studies using native DNAs of heterogeneous base sequence by Gellert et al., Wells and Larson, and Mtiller and Crothers investigated the hydrodynamic, kinetic, and thermodynamic properties of actinomycin D-DNA interactions and demonstrated actinomycin D to bind strongly to double-strand DNAs (typically Kint of 1-5 x 106 M -1) with binding strength directly correlated with GC content of the DNAs. 83-85 Comparisons of the reversible DNA-binding properties of the parent actinomycin D and 7-azidoactinomycin D are provided in Table II. The structure of actinomycin D complexed with DNA was first determined by Sobell and co-workers, who proposed an intercalative binding model involving insertion of the phenoxazone chromophore between the d(GpC) step and placement of the cyclic pentapeptide side chains within the minor groove of the DNA. This orientation allows the drug to form hydrogen bond contacts between the carbonyl oxygen of a threonine residue with the 2-amino group of guanine; hence, dictating the d(GpC) sequence preference. 86'87 Further evidence of the preferred binding to the d(GpC) step was subsequently presented, using a variety of biophysical methods and DNase I footprinting studies. 82T.-L. Shieh, P. Hoyos,E. Kolodziej,J. G. Stowell, W. M. Baird, and S. R. Byrn,J. Med. 1225 (1990). 83M. Gellert,C. E. Smith, D. Neville,and G. Felsenfeld,J. Mol. Biol. 11, 445 (1965). 84W. MUllerand D. M. Crothers,J. Mol. Biol. 35, 251 (1968). 85R. D. Wells and J. E. Larson,J. Mol. Biol. 49, 319 (1970). 86S. C. Jain and H. M. Sobell,J. Mol. Biol. 68, 1 (1972). 87H. M. Sobell,Sci. Am. 231, 82 (1974).

Chem.

33,

390

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

pentapeptideside

[1 8]

chain~

phenoxazone chromophore

pentapeptsididchai ee n Front View

Side View

FIG. 5. Three-dimensional structure of the sequence-selective DNA-binding agent, actinomycin D. These structures reveal the spatial relationships between the planar phenoxazone chromophore flanked above and below b y the cyclic pentapeptide side chains. The structure was obtained from the Nucleic A c i d Database file (ddh048) ( T a k u s a g a w a

et al? 9) a n d visualized with Insight lI (MSI).

In 1992, Takusagawa and co-workers reported the structure of actinomycin D complexed with the self-complementary deoxyoctanucleotide d(GAAGCTTC)2 by X-ray crystallographic methods. This study revealed that on binding actinomycin D, the DNA becomes severely distorted (i.e., kinked) and, interestingly, displayed an asymmetry of the pentapeptide side chains of actinomycin D residing in the minor groove. Several contacts between the DNA and the ligand were observed, including hydrophobic interactions between the surface of the minor groove and the proline, sarcosine, and methylvaline residues of the cyclic pentapeptides as shown in Fig. 6. 88,89 Early studies by White and Phillips 9° and Fox and Waring 91 revealed complex association and dissociation kinetics needed to describe the actinomycin T A B L E 11 BIOPHYSICAL PROPERTIES OF PARENT ACTINOMYCIN D AND ITS PHOTOREACTIVE ANALOG, 7-AZIDOACTINOMYCIN D

Characteristic

Parent actinomycin D

7-Azidoactinomycin D

Visible properties

)'-max )~max + DNA ~ee 8bound D N A - b i n d i n g properties Kinl Binding site size Sequence specificity Photolysis rate

4 0 0 nm 4 5 0 nm 2 2 , 5 0 0 M l cm I 12,000 M - I cm - l

462 nm 472 nm 29,800 M - 1 c m - I 13,560 M ] c m 1

6 x 105M I(bp) 6 (bp) GC

5 × 105 M -1 (bp) 6 (bp) GC K = 6.2 x 10 3 s e c I (at 80 J I M 2 sec 1

[ 18]

COVALENTCROSS-LINKINGOF LIGAND-DNA INTERACTIONS

391

duplex DNA sequence (5'-CTATTGCA TAC-3')

phenoxazone ring

pentapeptide side chains

FIG.6. Energy-minimizedstructure showing the complexformed between actinomycinD and the 5'-CTATTGCATAC-3'/5'-GTATGCAATAG-3' duplex. The viewis fromthe minorgrooveof the DNA, illustrating the intercalativegeometryof the phenoxazonering at the GpC step and the pentapeptide side chains residing tightly within the minor grooveof the DNA.The structure was generated by Discover and Insight II (MSI). D - D N A interaction. Using homogeneous DNA systems (i.e., synthetic oligo- and polynucleotides), the association kinetics of the ligand-DNA interactions were characterized by several slow, unimolecular processes with qualitatively little or no sequence or length dependence in the binding kinetics. By contrast, utilizing heterogeneous DNAs (i.e., calf thymus), the association kinetics were shown to be highly complex. In 1986, Fox and Waring described a binding model to explain the time-dependent DNase I footprinting patterns, coining the term "shuffling hypothesis." In this model, initial interactions of actinomycin D with the DNA lattice are nonsequence specific. The ligands are presumed to "shuffle" along the lattice sampling binding sites until preferred sites are found. On interaction with the preferred sites, tight binding with slow dissociation is observed. 91 In collaboration with Waring and Bailly, we provided a direct test of the shuffling hypothesis by using the photoreactive analog of actinomycin D, 7-azidoactinomycin D. 92 The 7-azidoactinomycin D analog provides a unique probe for examining the time-dependent binding of actinomycin D to DNA. Similarly, photolysis of the azido moiety to the reactive nitrene converts this analog into a DNAalkylating agent that retains the intercalative DNA-binding characteristics of the s8 S. Kamitori and E Takusagawa,J. Mol. Biol. 225, 445 (1992). 89F. Takusagawa, L. Wen, W. Chu, Q. Li, R. G. Carlson, K. T. Takusagawa, and R. E Weaver, Biochemistry 35, 13240 (1996). 9oR. J. White and D. R. Phillips, Biochemistry 28, 6259 (1989). 91 K. R. Fox and M. J. Waring,Biochemistry 25, 4349 (1986). 92C. Bailly, D. E. Graves, G. Ridge, and M. J. Waring,Biochemistry 33, 8736 (1994).

392

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

[18]

parent actinomycin D molecule. In this study, the photoreactive actinomycin D analog was allowed to interact with a 32p-labeled restriction fragment for specified lengths of time ranging from 20 sec up to 45 min. After incubation, the drug-DNA complex was photolyzed, rendering the drug irreversibly bonded to the DNA. The modified DNA was then treated with hot piperidine and analyzed on a DNA sequencing gel. Results were quantitated by densitometric analysis and revealed time-dependent movements from selected sites on the DNA lattice; with classically low-affinity sites decreasing in reactivity (i.e., actinomycin D binding) and high-affinity sites increasing in reactivity as shown in Fig. 7. These studies provide conclusive support for the shuffling hypothesis and demonstrate that the locations of actinomycin D binding to the DNA lattice change significantly with time. 92'93 Of interest was the observation that even at the shortest times, the DNA sequencing patterns revealed actinomycin to be absent from DNA sequences having high AT content, indicating that the initial event in the sequence recognition process requires recognition of a global structural feature such as minor groove geometry. The minor groove of the DNA duplex within localized sites having high AT content is narrow in comparison with other regions of DNA with high GC or mixed sequences. The inability of actinomycin D to bind the AT-rich sites even at early equilibration times indicates an initial recognition of a sequence-dependent structural feature necessary for ligand interaction. 92 By the same analogy, DNA sequences with high GC or mixed content seemed to accommodate actinomycin D fairly readily, so that early binding of the ligand was observed at many such sequences (i.e., 5'-GpC-3' and 5'-GGG-3'), as shown in Fig. 3. Photolysis of the 7-azidoactinomycin D-DNA complex results in photoactivation of the azide to the reactive nitrene and adduct formation. The ability to covalently bond the actinomycin D analog gave us a highly sensitive probe to examine sequence selectivity of actinomycin D. Covalent attachment of the drug allows sequence specificity studies to be performed at much lower drug concentrations than in comparable studies using reversible binding compounds; thus, much greater specificity (i.e., high-affinity sites) can be observed. As shown by Kulkarni and Yielding 61 and H61bne and co-workers, 94'95 DNA photoadducts of ethidium monoazide and 3-azidoproflavin linked to DNA are alkali labile, thus circumventing the reliance on DNA cleavage agents such as DNase I or chemical cleavage, both of which are hampered by the need for single-hit kinetics and nonrandom DNA cleavage. Because the ligand-DNA adduct is selectively cut at the site of covalent attachment by treating the adduct with piperidine as for a sequencing reaction, the resolution of the sequence specificity is markedly enhanced. 93G. S. Ridge, C. Bailly, D. E. Graves,and M. J. Waring,Nucleic: Acids Res. 22, 5241 (1994). 94p. Cieplak,S. N. Rao, C H616ne,T. Montenay-Garestier,and P. A. Kollman,J. Biomol. Struct. 5, 361 (1987). 95C. H61~neand N. T. Thuong, Genome 31,413 (1989).

Dyn.

[181

COVALENT CROSS-LINKING OF L I G A N D - - D N A INTERACTIONS

393

40 o o o

5'-A-T-C-C-G-C-T-C-A-C-T-A-G-G-C-G-A-G-T-G-5'

35

x,.E. 30 position 47

~ 25 c-

-a 20 C

m 15 "0

[]

5'-A-T-C-C-G-C-T-C-A-C-

(1)

__-N-- 10

E o

Z

5

position 43

0

i

i

I

i

10

20

30

40

50

Incubation Time Prior to Photolysis (min) FIG. 7. Influence of DNA base sequence on binding of 7-azidoactinomycin D within the DNA sequence 5t-GTGAGCGGAT-3 ' as a function of equilibration time. The photoreactive analog of actinomycin D was added to the DNA and equilibrated for specific times prior to photolysis. At times less than 1 min, actinomycin D is shown to interact with DNA in a relatively nonspecific manner. However, as the equilibration time is increased, preferred binding sites show an increase in binding density, whereas less preferred sites show decreased binding densities. Experiments were performed as described in Bailly et al. 92 and Ridge et al. 93

Using this method, Rill and co-workers examined the sequence-specific interactions of 7-azidoactinomycin D and compared these results with those obtained by DNase I footprinting of the parent compound. 97 Results from these studies are consistent with a predominant preference for intercalation of the 7-azidoactinomycin D phenoxazone ring into dGpdC steps. The intercalation geometry is such that both the flanking G and C residues readily react to yield a piperidine-labile adduct. In addition, these studies revealed that the DNA bases that flank the intercalation site strongly influence the binding specificity of actinomycin D. 97 Using this method, 7-azidoactinomycin D was observed to exhibit a strong binding preference for a novel nontraditional sequence, the d[T(G)nT] motif, where n equals 2 to 4. Careful examination of this motif revaled actinomycin D to bind with the strongest affinity to the d(TGGGT) sequence. 9s,99

96 j. Goodisman, R. Rehfuss, B. Ward, and J. C. Dabrowiak, Biochemistry 31, 1046 (1992). 97 R. L. Rill, G. A. Marsch, and D. E. Graves, J. Biomol. Struct. Dyn. 7, 591 (1989). 98 S. A. Bailey, D. E. Graves, R. Rill, and G. Marsch, Biochemistry 32, 5881 (1993). 99 S. A. Bailey, D. E. Graves, R. Rill, and G. Marsch, Biochemistry 33, 11493 (1994).

394

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

[18]

Methodologies Using Photoaffinity Analogs as DNA-Binding Agents Photoaffinity labeling is a valuable yet underutilized tool for studying nucleic acid structure and function. This method is easily adaptable to both in v i t r o and in v i v o conditions and can provide key insight into the mechanism(s) of action of many biologically active DNA-binding agents. Conditions for the preparation and purification of the photoreactive analogs of DNA-binding agents are as varied as the agents themselves. The synthetic procedures and chemical properties of all the photoaffinity analogs described in this chapter are published and referenced accordingly.100 105 Utilization of the azido analogs as DNA-binding probes requires that the work be carried out under photographic "darkroom" conditions. In all synthesis, storage, and subsequent experiments, the extent of exposure to light must be rigorously restricted. Exposure to room light for even a few minutes will result in degradation of the photoreactive agent. Red safety lights have been used with no observed degradation of the photoreactive analog. Covalent attachment of the photoaffinity analog to DNA is accomplished by the following general procedure. Photolysis conditions may be adapted to fit the necessary conditions dictated by the nature of the DNA sample being modified. In general, DNA samples in 10 mM buffer (pH 7.0) containing 1 mM disodium EDTA, and 0.01 to 0.1 M sodium chloride, are equilibrated with known concentrations of the photoreactive ligand. The temperature of the ligand-DNA solution may vary, dependent on the length and topology of the target DNA (i.e., native DNAs with higher melting temperatures may be modified at room temperature whereas deoxyoligonucleotides may require lower temperatures to ensure the duplex structure of the DNA target). Care must be taken to prevent premature exposure to light; hence, the ligand is prepared and the concentration is determined under photographic safelight conditions. After an appropriate equilibration time, the ligand-DNA complex is photolyzed for a brief period via exposure to visible light. The method of choice in this laboratory has been to use two Haake Buchler (Paramus, NJ) light boxes, each containing two General Electric daylight No. F15T8-D fluorescent light bulbs delivering a total of ~80 J. m -2 sec-l. A photolysis kinetic profile in buffer is advisable to gauge the amount of time and energy required for photolysis of the ligand. For the ethidium monoazide adducts, the photolysis is carried out at 5 ° to minimize heating of the sample by the close proximity of the light boxes. The nominal time and efficiency of photolytic attachment vary with the particular drug analog. For example, covalent attachment of the ethidium monoazide to calf thymus DNA is around 10oD. E. Graves,L. W. Yielding, C. L. Watkins,and K. L. Yielding, Biochim. Biophys. Acta 479, 98 (1977). Jo~D. E. Gravesand R. M. Wadkins,J. Biol. Chem. 264, 7262 (1989). Jo2W. A. Denny, B. E Cain, G. J. Atwell, C. Hansch, A. Panthananickal, and A. Leo,J. Med. Chem. 25, 276 (1982). Io3W. J. Firth, S. G. Rock, B. R. Brown, and L. W. Yielding,Mutat. Res. 81, 295 (1981). io4 K. L. Yieldingand L. W. Yielding,Ann. N. 1( Acad. Sci. 346, 368 (1980). Io5W. E. White, Jr., and K. L. Yielding,Methods Enzymol. 44, 644 (1977).

[18]

COVALENTCROSS-LINKINGOF LIGAND--DNAINTERACTIONS

395

30% with extended photolysis times. In contrast, the photolytic efficiency of 7azidoactinomycin D is >95% under identical conditions. Prior to photolysis, the ligand-DNA complex is reversible; hence, during photolysis, ligand is undergoing association/dissociation and can react with water to yield the reversible binding hydroxylamine photoproduct. After photolysis, all ligand that has not bonded to the DNA must be removed. In the case of charged ligands such as ethidium monoazide or m-AMSA, this is readily accomplished by small spin columns filled with Chelex resin (Bio-Rad, Hercules, CA). 106'107 For 7-azidoactinomycin D, which is neutral, size-exclusion chromatography is used. The ligand-DNA adducts made in this laboratory have been relatively stable when maintained at 5 ° and kept in the dark. Ethidium adducts formed with supercoited plasmid DNA have maintained the supercoiled topology for up to 6 months with no appreciable degradation of the supercoiled topology when compared with nonmodified supercoiled plasmid stored at 5 ° and wrapped in aluminum foil. However, exposure to room light (prior to and during gel electrophoresis) will result in substantial single-strand cleavage, comparable to that observed for the parent ethidium-supercoiled DNA complex. Conclusions The use of photoaffinity labeling to study ligand-DNA interactions provides a powerful tool for the overall assessment of ligand targeting, complex structure, and biological and/or molecular mechanism(s) resulting from these interactions. Many of the reversible binding ligands that interact with DNA do so with high affinities, have chromophores that absorb in the visible range, and hence are amenable as potential candidates or parent compounds for the development of photoreactive analogs. The utility of the photoaffinity analog depends on the following criteria: (1) the ligand should be photoreactive at visible wavelengths such that cellular and/or DNA damage is not incurred; (2) the lifetime of the reactive product (nitrene) should be sufficiently short to assure quenching by solvent interactions and exclusion of any secondary perturbations; (3) the reaction time of the reactive nitrene should be sufficiently fast to ensure that the covalent adduct formed is representative of the bound ligand in its reversible state; and (4) the resulting ligand-DNA adduct must be stable such that isolation and characterization of the adduct is possible. This chapter has dealt primarily with ethidium, acridine, and actinomycin derivatives, providing key studies utilizing the photoreactive analogs to examine a number of biophysical and structural problems; however, the potential for applying this methology to other DNA-binding agents offers exciting avenues for future research. Io6E L. Gilbert,D. E. Graves,M. Britt, and J. B. Chaires,Biochemistry 30, 10931 (1991). 107p. L. Gilbert,D. E. Graves,and J. B. Chaires,Biochemistry 30, 10925(1991).

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[19]

[19] Chemical Cross-Linking of Drugs to DNA By RYAN A. LUCK and PAUL B. HOPKINS Introduction It is well known that a number of antitumor agents and carcinogens garner their activities through "chemically cross-linking" DNA. These are compounds that become covalently linked to the DNA duplex. "Chemical cross-linking," however, is an imprecise term in that there are a number of categories of DNA lesions that can be considered the result of these modifications. Four of these categories are considered here. The first of these is interstrand cross-links, which are characterized by covalent linkages formed between the two different strands that form the DNA duplex. Second, there are intrastrand cross-links, covalent linkages formed between two different sites on one strand of DNA. Third, there are monoadducts, which are characterized by the covalent bonding of the compound to one site on one strand of the DNA duplex. Finally, there are a small number of agents that are able to covalently bind DNA via a second endogenous bridging compound. All four categories of chemical cross-links are important in understanding the activity of antitumor agents and carcinogens. As a result, it is important to understand the methods used in their analysis. Initially, this chapter discusses methods for analyzing DNA interstrand crosslinks and includes methods for their detection, location, and determination of their covalent structure. After this, we discuss methods used to analyze intrastrand cross-links and monoadducts. The fourth category of cross-link, wherein agents are covalently linked through endogenous bridging compounds, is not as prevalent as the other categories. As a result, there are fewer published methods for analysis; however, we have developed methodologies in our laboratory for analyzing the formaldehyde-mediated coupling of doxorubicin (Adriamycin) to DNA. This new methodology is explained in the final part of the chapter.

D e t e c t i o n of I n t e r s t r a n d C r o s s - L i n k s

Ethidium Bromide Fluorescence Assay The first challenge for analysis of interstrand cross-links is detection. A number of methods have been developed for detecting the presence of interstrand crosslinks. One of the earliest techniques was the ethidium bromide fluorescence assay. The ethidium bromide fluorescence assay takes advantage of the increase in fluorescence that results when ethidium bromide binds to double-stranded DNA. This assay was initially developed by Morgan and Paetkau in 1972.1 In broad terms,

METHODS IN ENZYMOLOGY, VOL. 340

Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. 0076-6879/00 $35.00

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a DNA sample is exposed to an interstrand cross-linking agent. The sample is then mixed with an ethidium bromide solution and the fluorescence of the resuiting mixture is measured. This is followed by heat denaturation, a fairly short cooling period, and another measurement of the fluorescence. If there is no interstrand cross-linking, the fluorescence of the sample after the cooling period will be greatly diminished. The short cooling period does not allow enough time for the single-stranded DNA sequences to find their complements and reform the duplex. However, if an interstrand cross-link is present, then the strands were never completely separated, resulting in the relatively quick reforming of the duplex. Lusthof and co-workers examined the interstrand cross-linking activity of 2,5bis(1-aziridinyl)-3,5-bis(carboethoxyamino)-1,4-benzoquinone (AZQ, I) and several related compounds, using the ethidium bromide fluorescence assay. 2 The specific procedure is outlined below.

(I) 2,5-bis(1-aziridinyl)-3,5-bis(carboethoxyamino)- 1,4-benzoquinone (AZQ) A solution of the reduced form of AZQ (I) (40/~1, 0.05 mM) is combined with 360 #1 of calf thymus DNA solution (6.7 x 10-1 mg/ml) and allowed to react at 37 ° for 1 hr. Excess cross-linking agent is removed with Sephadex G-50 and ethanol precipitation. The DNA precipitate is dissolved in 100 #1 of water overnight at 4 °. Thirty microliters of this solution is combined with 2 ml of aqueous ethidium bromide solution and the fluorescence is measured. The samples are then heat denatured at 95 ° for 3 min, quickly followed by cooling at 20 ° for 7 min. Immediately after this, the fluorescence of the sample is measured. The ratio of the fluorescence after and before denaturation [(FJFu)s] is compared with the same ratio for a blank sample [(FJFu)u]. Using these ratios, an interstrand cross-link percentage is calculated, using Eq. (1): Cross-link percentage = L

l~(~/~)~b)b

J

100%

(1)

Lusthof and co-workers are careful to note that the cross-link percentage is not directly correlated to the percentage of base pairs covalently linked. Instead, it is

I A. R. Morgan and V. Paetkau, Can. J. Biochem. 50, 210 (1972). 2 K. J. Lusthof, N. J. De Mol, L. H. Janssen, W. Verboom, and D. N. Reinhoudt, Chem. Biol. Interact. 70, 249 (1989).

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CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES I

I

r

[

m

m

/

native duplex DNA

[1 9]

~

Interstrand Cross-link

m

DNA w/ cross-linking agent

FIG. 1. Schematicof DPAGEassay used for detection of interstrand cross-linking. correlated to the percentage of DNA that is in duplex form after the heat denaturation and quick cooling process. Denaturing Polyacrylamide Gel Electrophoresis

Gel electrophoresis is a widely used technique for manipulation of a diversity of biomolecules, including DNA, RNA, and proteins. With the increased availability of relatively short, custom-made oligonucleotides, denaturing polyacrylamide gel electrophoresis (DPAGE) has become a commonly performed assay for manipulating DNA and assessing its interactions with a variety of other compounds. DPAGE is a special type of gel electrophoresis in which the matrix of the gel has been loaded with some type of chemical denaturant. (This denaturant, often urea, is usually a component of the sample loading buffer as well.) The denaturant hinders the formation of the normal DNA duplex by keeping the single-stranded oligonucleotides from base pairing with their complement. This technique is used in a number of ways, including sequencing and purification. DPAGE is also a useful method for the detection of interstrand cross-links. When a double-stranded oligonucleotide is denatured, the single strands move with approximately twice the mobility (compared with the original duplex) through the gel matrix. If an interstrand cross-link has been formed, however, the strands cannot separate. As a result, they retain their retarded mobility and form a discrete band that does not progress nearly as far through the gel (Fig. 1). This slower mobility band can be detected and quantified either through UV shadowing of the gel or by prior radiolabeling of the oligonucleotide. Huang et al. investigated the interstrand cross-linking activity of formaldehyde in 1992. 3 Utilizing a library of self-complementary 17-mer oligonucleotides, they determined which sequences yielded interstrand cross-links, using the DPAGE assay. Solutions of radiolabeled DNA duplexes that have been incubated with 3 H. Huang, M. S. Solomon,and R B. Hopkins,J. Am. Chem. Soc. 114, 9240 (1992).

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formaldehyde are ethanol precipitated. The precipitate is dissolved in 10 ttl of water followed by the addition of 10 #1 of sample loading buffer [90% (v/v) formamide, 10 mM Tris [pH 7.5], 0.1% (w/v) xylene cyanol, and 0.1 mM EDTA]. This mixture is heated at 90 ° for 2 min, iced for 2-3 min, and then loaded onto the gel. DPAGE is conducted on a Hoefer (San Francisco, CA) thermojacketed Poker Face gel stand, using a 25% (w/v) gel (19 : 1 acrylamide : bisacrylamide, 8 M urea, 0.35 mm thick, 33 x 41 cm, 20-tooth comb). The gel is run at 65-70 W at approximately 55 ° until the xylene cyanol dye had run 14-16 cm. The gel is then transferred onto Whatman (Clifton, N J) 3M filter paper, covered with Saran Wrap, and dried with a Bio-Rad (Hercules, CA) model 583 gel drier. Autoradiography is then used to visualize the single-stranded and cross-linked duplexes. Our laboratory has found that DPAGE is an extremely robust method for detection of interstrand cross-linked DNA. This method has been used for detecting interstrand cross-linking by a variety of compounds, including cisplatin, mitomycin C, nitrogen mustard, formaldehyde, and others. Variations in size and thickness of the gel, urea concentrations, acrylamide : bisacrylamide ratio, temperature, loading buffer composition, and power usually can affect only the aesthetic appearance of the resulting data, not the conclusions. With regard to the loading buffer, it should be noted that as long as the sample is heat denatured, the buffer does not necessarily need to contain a chemical denaturant. Furthermore, cross-linking reactions have been directly loaded onto the gel with good results. If the sample is ethanol precipitated, however, it is important first to dissolve the sample in water prior to adding a urea-based loading buffer. A high concentration of urea often makes it difficult for the DNA precipitate to dissolve, resulting in a DNA sample that never migrates into the gel matrix. D e t e r m i n i n g S e q u e n c e L o c a t i o n of I n t e r s t r a n d C r o s s - L i n k s

Use of Fenton Chemistry The DPAGE assay described in the previous section provides a method for determining which DNA base sequences yield interstrand cross-linking. Using this method, it is possible to began making inferences about which specific DNA bases are involved in formation of an interstrand cross-link. Further investigation is needed, however, in order to make definitive conclusions about the bases involved. One method for determining the sequence location of interstrand cross-links combines the DPAGE methodology with Fenton chemistry. In 1985, Tullius and Dombroski demonstrated that a mixture of iron(II), hydrogen peroxide, EDTA, and ascorbate resulted in cleavage of the DNA backbone. 4 In this mixture, iron(II) reduces hydrogen peroxide to give a hydroxyl radical. The 4 T. D. Tulliusand B. A. Dombroski,Science230, 679 (1985).

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CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

5'

* 3'

Fe(EDTA)2H202 ascorbate

[ 19]

* 3' *3 ' - -

*3'--

+ cross-linked fragments

*3'--

Radiolabeled products FIG. 2. Hydroxyl radical cleavage of interstrand cross-linked DNA.

hydroxyl radical abstracts a hydrogen atom from the deoxyribose sugar that composes the DNA backbone, resulting in nearly sequence-random cleavage. Ascorbate is present in order to reduce the resulting iron(III) back to iron(II), thereby continuing the process. This procedure, known as hydroxyl radical footprinting, has been used to examine interactions between DNA and proteins as well as global structural changes associated with unusual DNA sequences. Hydroxyl radical footprinting has been used in our laboratory to pinpoint the sequence location of interstrand cross-links. 5 In a DNA duplex that has only one interstrand cross-link, single-hit cleavage generated by Fenton chemistry results in a mixture of products. If the 3' end of one strand is radiolabeled, there are only two categories of products observed when the mixture is analyzed by DPAGE (Fig. 2). The first category is composed of the radiolabeled products that still contain the interstrand cross-link. The second category consists of the short fragments that correspond to cleavage on the 3' side of the cross-link on the radiolabeled strand. When the mixture is run on the gel, the two categories have different mobilities; the fragments that still contain the cross-link are much larger and move much more slowly through the gel. The location of the interstrand cross-link can then be determined by simply analyzing the length of the short fragments generated. This technique also works if the 5' terminus of one of the single strands is selectively radiolabeled. Radiolabeling of either both 5' termini or both 3' termini can also work, as long as the sequence is self-complementary.

[2+ l H3N, ,NH3 Pt cl" "cI

(II) - cis-diamminedichloroplatinum(II) (cisplatin) This technique has been used in our laboratory to determine the exact sequence location of the interstrand cross-link formed by cisplatin (II). 6 Radiolabeled DNA 5 M. E Weidner, J. T. Millard, and E B. Hopkins, J. Am. Chem. Soc. 111, 9270 (1989). 6 E B. Hopkins, J. T. Millard, J. Woo, M. E Weidner, J. J. Kirchner, S. T. Sigurdsson, and S. Rancher, Tetrahedron 47, 2475 (1991).

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that has been incubated with cisplatin is initially analyzed by DPAGE. The interstrand cross-linked product, which demonstrates approximately half the mobility of the single strands, is visualized by autoradiography and the band is removed from the gel matrix with a razor blade. The gel slice is crushed with a glass rod and incubated at 37 ° in aqueous solution (0.5 M NH4OOCCH3, 1 mM EDTA) for 16 hr. The eluate is extracted with a glass pipette and loaded onto a Waters (Milford, MA) Sep-Pak C~8 cartridge in order to remove salts and gel particulate. On this cartridge, the sample is sequentially washed with 10 ml of aqueous NHaOOCCH3 (10 mM) and 10 ml of water. The sample is then recovered by washing the cartridge with 4 ml of 25% CH3CN-water. This mixture is evaporated in vacuo and the sample is redissolved in water. The 20-#1 reaction mixture for the hydroxyl radical cleavage is then formulated using DNA (100,000 cpm by Geiger counter), (NH4)2Fe(SO4)2 (50/zM), EDTA (100/zM), sodium ascorbate (1 mM), H202 (10 mM), and Tris (pH 7.5, 5 mM). After 1 min at room temperature, the reactions are stopped by addition of 2 #1 of aqueous thiourea (50 mM). The mixture is then lyophilized, dissolved in 5 #1 of sample loading buffer, heated to 90 ° for 3 min, cooled to 0 °, and analyzed by DPAGE. DPAGE in this instance is a 25% (w/v) acrylamide gel cross-linked with 5% (w/v) bisacrylamide, 48% (w/v) urea, 0.35 mm thick, 41 x 37 cm. It is run at 2000 V on a Hoefer thermojacketed Poker Face gel stand at 64-68 °. It has been the experience in our laboratory that the components of the hydroxyl radical cleavage reaction need to the made immediately prior to use, particularly the Fe(II) and sodium ascorbate solutions. Once again, however, the specific composition of the gel used for the DPAGE is not critical. The relatively high temperature of the gel portion of the assay is used in order to make the separation between the various short fragments and the cross-link as well defined as possible. This high temperature is not necessary, however, for the experiment to work. Piperidine Cleavage at Interso'and Cross-Linked Deoxyguanosine Residues

Heating DNA with aqueous piperidine leads to cleavage at deoxyguanosine (dG) residues that are alkylated at N7. There are two steps to this process: (1) depurination and (2) cleavage at the resulting abasic site. This reactivity can be used to demonstrate the presence of interstrand cross-links at N7. Rink et al. and Millard et al. used this assay to determine the sequence preference of the interstrand cross-link formed by mechlorethamine (III).7'gA single-stranded oligonucleotide containing the sequence 5'-d(GJG2G3C) is radiolabeled and annealed to its complement. The resulting duplex is treated with mechlorethamine and the interstrand cross-linked material is isolated by DPAGE and "crush and soak" methodology

c,Ha

cI~N~cI (III) - mechlorethamine

402

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[ 19]

described previously. This DNA is then heated in 50 #l of 1 M piperidine in a sealed microcentrifuge tube for 1 hr. The mixture is dried in vacuo, lyophilized twice from 25 #1 of water, dissolved in sample loading buffer, and analyzed by DPAGE. DPAGE analysis shows that cleavage on the radiolabeled strand occurs primarily at G 2. This result, combined with experiments involving the complement strand and other duplexes, helped to determine that the sequence selectivity of the mechlorethamine interstrand cross-link targeted 5'-d (GNC). C o m p o s i t i o n of I n t e r s t r a n d C r o s s - L i n k s

Acid Hydrolysis of Cross-Linked DNA Generally, the next step in the analysis of interstrand cross-links is to determine the exact chemical composition and structure of the covalent linkage. This includes not only information about the cross-link itself, but also its points of attachment to the bases. The quickest way to gather information about the structure of the cross-link is to cleave the oligonucleotides into their single-base components for analysis. This section addresses various methods for this process; it does not detail, however, the various methods for analyzing the resulting components. The methods for identifying these products are similar to those for identifying natural products, including nuclear magnetic spectrometry (NMR), mass spectrometry, authentic sample comparisons, and others. These methods are too broad to be fully explained here and are addressed elsewhere. The initial methodology for breaking DNA into its single base components was acid hydrolysis. In 1961, Brookes and Lawley used acid hydrolysis to identify the products of a variety of alkylating agents and DNA. 9 From these experiments came the first evidence that mechlorethamine (III) was a DNA cross-linking agent, although it was unclear to the authors whether this was an intrastrand or interstrand cross-link. Unfortunately, the details of the acid hydrolysis procedure were not published. Two different procedures for acid hydrolysis of cross-linked DNA were published, however, in regard to investigations of the cis-syn-thymine photodimer (IV). 1°,11 The cis-syn-thymine photodimer is formed at adjacent thymines when DNA is exposed to ultraviolet light. This cross-link is an intrastrand cross-link and thus does not involve any exogenous alkylating agents, but the procedures should be applicable to interstrand cross-links as well (provided they are acid stable). 7 S. M. Rink, M. S. Solomon, M. J. Taylor, S. B. Rajur, L. W. McLaughlin, and E B. Hopkins, J. Am. Chem. Soe. 115, 2551 (1995). 8 j. T. Millard, S. Raucher, and E B. Hopkins, J. Am. Chem. Soc. 112, 2459 (1990). 9 E Brookes and E D. Lawley, Biochem. J. 80, 496 (1961). l0 A. J. Varghese and S. Y. Wang, Nature (London) 213, 909 (1967). 11 D. Weinblum, Biochem. Biophys. Res. Commun. 27, 384 (1967).

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CHEMICALCROSS-LINKINGOF DRUGS TO DNA

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Varghese and Wang exposed approximately 5 g of native calf thymus DNA to ultraviolet conditions. Afterward, the DNA sample is evaporated to dryness, and the material is dissolved in 200 ml of trifluoroacetic acid (TFA). l° This sample is transferred to 20 Pyrex glass tubes, which are then sealed and heated at 170 ° for 90 min. To remove insoluble material, the sample is then filtered. The tubes and the insoluble material are washed twice with 0.1 N HCI. The filtrate is evaporated to dryness. Varghese and Wang than dissolve the residue in TFA again and analyze the components by thin-layer chromatography and NMR.

O~CH3CH3J? HN")x""~-~ ''" %NH

(IV) - cis-syn thymine photodimer Weinblum uses a different method for hydrolyzing DNA containing the cissyn-thymine photodimer, li Approximately 1.5 g of salmon sperm DNA is exposed to the cross-linking conditions. The dried, cross-linked DNA is then dissolved in 3 ml of 70% (w/v) perchloric acid. This is heated to I00 ° for 45 min. Water 150 ml is added and the solution is neutralized by 10 N KOH. The solution is left overnight; the KC104 precipitate is then filtered off. The filtrate is adjusted to pH 10, using ammonium hydroxide, and the hydrolysis products are separated by Dowex chromatography. Enzymatic Digestion of lnterstrand Cross-Links

There are relatively few examples of the utilization of acid hydrolysis to isolate cross-linked components of DNA. This is because the harsh conditions of the procedure can potentially alter the structure of the cross-links, or even reverse their formation. In 1977, Dubelman and Shapiro addressed this limitation by developing methodology for digesting cross-linked DNA enzymatically.12,13 They accomplished this by performing a 98-hr digest of cross-linked DNA, using a combination of deoxyribonuclease I and snake venom phosphodiesterase. A variety of procedures for enzymatic digestion of DNA have since been developed and are outlined here (Table I; structures V and VI). As can easily be seen, a number of enzyme recipes have been used for digestion of cross-linked oligonucleotides. It should be noted that in the case of the nitrous acid cross-link, only the minimum enzymes are used--alkaline phosphatase and 12S. Dubelmanand R. Shapiro,NucleicAcids Res, 4, 1815 (1977). ~3R. Shapiro, S. Dubelman,A. M. Feinberg, P. E Crain, and J. A. McCloskey,J. Am. Chem. Soc. 99, 302 (1977).

TABLE I SAMPLING OF ENZYMATICDIGESTION METHODS FOR CROSS-LINKEDDNA Cross-linking agent CH20

cis-DEP (V)

Mitomycin C (VI)

AZQ (I)

BCNU (VII)

Nitrous acid

Enzymatic digestion procedure initial sample of cross-linked calf thymus DNA (ca. 60 mg) is brought up to 50 ml with digestion buffer (5 mM Tris-HC1, I mM NaCI, 1 mM MgCI2, pH 8.8). Deoxyribonuclease I (2 mg, 4000 units) is added at - 3 0 ° for 16 hr. Venom phosphodiesterase (1.5 rag, 30 units) and spleen phosphodiesterase are added at - 3 0 ° for another 6 hr. Alkaline phosphatase (150/zl, 250 units) is added at - 3 0 ° for 6 hr. Reaction mixtures are analyzed by HPLC Calf thymus DNA ( 100 #g) that has been incubated with cis-DEP (V) is ethanol precipitated and dried. It is then resuspended in digestion buffer (20 mM sodium acetate, 50 mM NaCI, 1 mM ZnSO4, 10 mM MgCI2, pH 4.6). Bovine pancreas deoxyribonuclease I (200 Kunitz units) is added at - 3 0 ° for 4 hr. SI nuclease (1000 units) is added at - 3 0 ° for 16 hr. One-tenth volume of solution of Tris (1.0 M, pH 9) and 5 units of alkaline phosphatase are added at - 3 0 ° for another 4 hr. Reaction mixtures are analyzed by HPLC Calf thymus DNA that has been cross-linked is brought to 3 A260 units/ml in digestion buffer (0.005 M Tris-HC1, 0.001 M MgC12, pH 7.0). Digestion is accomplished at 37 ° with this sequence of enzymes: DNase 1 (16 units/A260 unit) at 0 and 1 hr; snake venom phosphodiesterase (1.25 units/A260 unit, pH increased to 5.5) at 2 and 5 hr; alkaline phosphatase (0.5 unit/A260 unit) at 7 hr, incubation going until 24 hr. Reaction mixtures are analyzed by HPLC Cross-linked DNA (1.0 A260 unit) is dissolved in 100/zl of digestion buffer (50 mM Tris, 10 mM MgCI2, pH 7.5) and digested by DNase I (50 units), DNase 11 (21 units), snake venom phosphodiesterase (5 units), and calf intestinal alkaline phosphatase (15 units) at 25 "~for 18 hr. Reaction mixtures are analyzed by HPLC Cross-linked DNA sequence [5t-d(GC4G4C)]2 (0.4 A260 unit) is digested in a 30-#1 total volume of digestion buffer (50 mM Tris, 10 mM MgC12, pH 8.5) by a combination of DNase I (10 units), calf intestinal phosphatase (10 units), DNase II (4 units), phophodiesterase I (0.5 unit), and Sl nuclease (180 units). Reactions are allowed to go for 3 hr at 3T ~ and are analyzed by HPLC Cross-linked DNA sequence [5'-d(CG)612 (0.35 A260 unit) is digested in a 75-/zl total volume of digestion buffer (50 mM Tris, 10 mM MgCI2, pH 8.9) by a combination of calf intestinal phosphatase (2/~1, 20 units) and phophodiesterase I (6/~1, 4 units). Reactions are allowed to go 8 hr at 37 ° and are analyzed by HPLC

Ref.

h,,

~t

e

Y

a y. E Chaw, L. E. Crane, E Lange, and R. Shapiro, Biochemistry 19, 5525 (1980). t~A. Eastman, Biochemistry 24, 5027 (1985). c A. Eastman, Biochemistry 22, 3927 (1983). d M. Tomasz, D. Chowdary, R. Lipman, S. Shimotakahara, D. Veiro, V. Walker, and G. L. Verdine, Proc. Natl. Acad. Sci. U.S.A. 83, 6702 (1986). e S. C. Alley, K. A. Brameld, and E B. Hopkins, J. Am. Chem. Soc. 116, 2734 (1994). fP. L. Fischhaber, A. S. Gall, J. A. Duncan, and E B. Hopkins, CancerRes. $9, 4363 (1999). g J. J. Kirchner, S. T. Sigurdsson, and P. B. Hopkins, J. Am. Chem. Soc. 114, 4021 (1992).

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CHEMICALCROSS-LINKINGOF DRUGS TO DNA

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phosphodiesterase. These are the two primary enzymes needed. As the amount of DNA or the structural perturbations introduced by the cross-link increase, other digestive enzymes are called on to help. It should also be mentioned that with small amounts of DNA, digestion can be accomplished at room temperature in less than 1 hr. [

I

H2N, ,NH2 Pt CI" "CI (V) - cis-dichloro(ethylenediamine)platinum(II)

(cis-DEP) O

H2N~O~NH2 o (VI) - mitomycin C Generally, it is the practice in our laboratory to analyze the results of the enzymatic digestion ofinterstrand cross-linked DNA by using reversed-phase highperformance liquid chromatography (HPLC). Analytical HPLC runs are generally performed on an Alltech (Deerfield, IL) 5-#m, Cls, 250 × 4.6 mm Econosphere column. A typical solvent gradient is run at 1 ml/min. A good starting place for gradient development is as follows: solvent A, 10 mM ammonium acetate; solvent B, 100% acetonitrile; isocratic 92% solvent A for 7 min, a 13-min linear gradient to 70% solvent A, a 10-min linear gradient to 60% solvent A, and a 10-min linear gradient to initial conditions. Conditions are then adjusted to increase separation of the nucleosides and cross-linked material. On occasion, the peak percentage of acetonitrile over the gradient must be increased in order to elute the cross-linked material from the column. In this manner, the cross-linked bases can be isolated and identified by other analytical techniques.

Controlled Depurination of Cross-Links Involving N7 Previously, methodology for piperidine-mediated cleavage of DNA at sites of N7 alkylation has been described. Similarly, N7-alkylated sites are vulnerable to controlled depurination by exposure to heat. This characteristic is advantageous when it is thought that an interstrand cross-link is made between N7 of two purines. Tong and Ludlum have used this method to isolate the interstrand cross-link from DNA that had been treated with N,N'-bis(2-chloroethyl)-N-nitrosourea (BCNU,

406

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

[1 9]

VII). 14 The procedure they used for controlled depurination is fairly straightforward. Approximately 16 mg of calf thymus DNA that had been exposed to BCNU is ethanol precipitated, washed with ethanol, and dried. The precipitate is then dissolved in 2 ml of sodium cacodylate buffer (25 mM, pH 7.0). This is followed by heating at 100° in a water bath for 15 rain and then cooling in an ice bath. The depurinated oligonucleotides are precipitated by the addition of 0.2 ml of 1 N HCi; this precipitate is then removed by centrifugation. The supernatant is evaporated in v a c u o and redissolved in 200 #l of water. The pH is adjusted to pH 6.0 with about 50 #l of 1 N NH4OH so that the sample is suitable for HPLC. Using this methodology, Tong and Ludlum were able to isolate 1,2-(diguan-7-yl) ethane (VIII), the major interstrand cross-link formed by BCNU. N"0 H I

I

CIJ~N'~N~cI 0 (VII)

- N,

N'-bis(2-chloroethyl)-N-nitrosourea O

O

(VIII) - 1,2-(diguan-7-yl) ethane It should be mentioned that enzymatic digestion can be used to isolate nearly every cross-link that can be isolated by controlled depurination. There are potential scenarios, however, in which controlled depurination would be the preferred method. One possible situation would be if the cross-linking were performed under conditions that prohibited the activity of the enzymes. Another possible situation would be if the researcher wanted only N7-alkylated purines isolated, because of yield or sensitivity issues. If controlled depurination is not necessary, however, enzymatic digestion is the preferred method in our laboratory. G l o b a l S t r u c t u r e of I n t e r s t r a n d C r o s s - L i n k s The final challenge for complete analysis of interstrand cross-links is the determination of the global structure of the entire oligonucleotide. Although there are numerous techniques for obtaining limited information about the global structure of oligonucleotides, the two most powerful methods are X-ray crystallography

J4W. E Tongand D. B. Ludlum,Cancer

Res.

41, 380 (1981).

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CHEMICALCROSS-LINKINGOF DRUGS TO DNA

407

and nuclear magnetic resonance spectrometry. The details involved in performing either of these types of analysis are too broad to be covered here and are explained elsewhere in this volume.

Analysis of Intrastrand Cross-Links and Monoadducts Most of the techniques described for the analysis of interstrand cross-links can also be used, with little or no modification, for the analysis of intrastrand cross-links or monoadducts. With respect to detection of cross-linking, DPAGE is an excellent technique for observing intrastrand cross-links or monoadducts. (For obvious reasons, the ethidium bromide fluorescence assay is not useful for this goal.) Intrastrand cross-links and monoadducts generally do not hold the duplex together under the denaturing conditions. The presence of a lesion, however, retards the mobility of single-stranded DNA through the gel matrix. As a result, both monoadducts and intrastrand cross-links can be observed by DPAGE, although the separation is not as dramatic as it is with interstrand cross-links. The most proficient method for determining the sequence location involved in interstrand cross-links, the use of Fenton chemistry, is not nearly as applicable with regard to intrastrand cross-links and monoadducts. This technique can be used only if it is possible to detect which hydroxyl radical-generated fragments have the intrastrand cross-link or monoadduct present. Because the change in mobility is less dramatic, this feat is more difficult than it is with an interstrand cross-link. Determination of the location of monoadducts is further complicated by the greater probability that there is more than one lesion. Analysis of results of piperidinemediated cleavage is potentially a useful technique, but only in those cases that involve purine alkylation at N7. After detection of intrastrand cross-links or monoadducts, the most common route in our laboratory is to break the DNA into its components and determine which bases are involved. As it was with interstrand cross-links, the methods of acid hydrolysis, enzymatic digestion, and controlled depurination are all applicable for this process. Also as it was with interstrand cross-links, however, enzymatic digestion is by far the most common method. The same enzyme recipes and analytical techniques used for determining the composition of intersti'and cross-links are the starting point for determining the composition of intrastrand cross-links and monoadducts.

Formaldehyde-Mediated

C o u p l i n g of DNA to A d r i a m y c i n

BackgroundInformation Until now, this chapter has not addressed the fourth category of "chemical cross-links" that was outlined in the introduction. These are the cross-links that result from the coupling of an agent to DNA via a third compound. As an example

408

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

O

OH

O OH30

OH O HO NH 2

IX

OH

O

OH O

O

O DNA + CH20

O

o

[1 9]

H3c'~O

N-~LL- N -H

3

- - ~

.~--o

~"~CH2~''~"

X

FIG. 3. Formaldehyde-mediated coupling of Adriamycin to DNA.

of this category, we outline the methodology we developed for investigating the formaldehyde-mediated coupling of doxorubicin (Adriamycin, IX; Fig. 3) to DNA. The formaldehyde-mediated coupling of Adriamycin (and the structurally similar daunorubicin) to DNA was discovered by Wang and co-workers using X-ray crystallography. 15'16 In this structure, Adriamycin is linked via its amino group to N2 of a deoxyguanosine residue through a methylene group (X, Fig. 3). The methylene bridge presumably results from successive nucleophilic attacks on formaldehyde by the amino groups of Adriamycin and dG. This structure became of interest to our laboratory when Koch and co-workers related it to the putative interstrand cross-links formed by Adriamycin, as seen by Phillips and co-workers. When DNA was exposed to reductively activated Adriamycin, the Phillips laboratory used DPAGE methodology and observed a band that was consistent with the presence of an interstrand cross-link. 17.=8The Koch laboratory used electrospray mass spectrometry to demonstrate that lesion X was formed under the same reductive conditions.19,2° Koch and co-workers suggested that the formaldehyde resulted either from H202-promoted oxidation of buffer components or from BaeyerVilliger oxidation of Adriamycin itself. (The H202 is generated by reduction of O2 by the reducing agents in the reaction mixture.) Koch and co-workers proposed, but did not definitively prove, that the formation of lesion X was responsible for observation of the putative interstrand cross-links. (For a more complete history of the contributions of various laboratories in this research area, please see Luce et al. 2j ) Our laboratory was able to provide supporting evidence for Koch's suggestion by 15 A. H. Wang, Y. G. Gao, Y. C. Liaw, and Y. K. Li, Biochemistry 30, 3812 (1991). 16 j. Wang, M. Chao, and A. H. Wang, Adducts of DNA and anthracycline antibiotics. In "Anthracycline Antibiotics," Vol. 574, pp. 168-182. American Chemical Society, Washington, D.C., ] 99 l. 17 C. Cullinane, A. van Rosmalen, and D. R. Phillips, Biochemistry 33, 4632 (1994). 18 S. M. Cutts and D. R. Phillips, Nucleic Acids Res. 23, 2450 (1995). 19 D. J. Taatjes, G. Gaudiano, K. Resing, and T. H. Koch, J. Med. Chem. 39, 4135 (1996). 20 D. J. Taatjes, G. Gaudiano, K. Resing, and T. H. Koch, J. Med. Chem. 40, 1276 (1997). 21 R. A. Luce, S. T. Sigurdsson, and P. B. Hopkins, Biochemistry 38, 8682 (1999).

[19]

CHEMICALCROSS-LINKINGOF DRUGSTO DNA

409

confirming that either H202 or CH20 mixed with Adriamycin and DNA generated the same double-stranded band when analyzed by DPAGE. The problem with DPAGE, however, is that it is not a consistent detection method for lesion X. Adriamycin is covalently attached to only one of the two strands of the DNA duplex. Noncovalent interactions between Adriamycin and the nonalkylated strand, as well as noncovalent interactions between the two DNA strands themselves, are essential for keeping the duplex together under the denaturing conditions of DPAGE. This makes the results of the DPAGE experiment difficult to predict for various oligonucleotides, depending on their G-C base pair content and length. If the length of the DNA was too short or if there were not enough G-C base pairs, no double-stranded band would be seen, even if it contained the [5'-d(GC)]2 sequence that was essential for its formation. Another problem is that small variations in experimental conditions lead to drastically different results. The temperature, time length, percentage of urea, and composition of loading buffer all affect the outcome of the DPAGE experiments when detecting the lesion. One could imagine that these different variables might somehow be combined and computed to ascertain how "denaturing" a particular DPAGE experiment was. An experiment must be denaturing enough to pull apart the two strands of a DNA duplex when Adriamycin is simply noncovalently bound. Experimental conditions cannot be so denaturing, however, that the lesion is not strong enough to hold the duplex together. This level of denaturing would obviously vary according to the oligonucletide sequence. Given this level of complexity, we decided to attempt to develop another assay for detection of lesion X that would be quantitative and flexible for a variety of oligonucleotides.

Synthesis of N-Methyl Adriamycin The simplest way to detect the presence of lesion X would be to isolate the formaldehyde-mediated conjugate of Adriamycin with deoxyguanosine by employing the enzymatic digestion methods previously described in this chapter. Attempts to isolate this conjugate, however, returned only unmodified nucleosides and Adriamycin. It is presumed that when lesion X is free from duplex DNA, the aminal in X is too vulnerable to hydrolysis. (For similar reasons, there is no detectable monoadduct when DNA containing lesion X is analyzed by DPAGE.) Therefore, the strategy that we chose was to capture the Adriamycin iminium ion intermediate that was generated when this aminal decomposed. Prior to implementing this strategy, however, it was necessary to synthesize a sample ofN-methyladriamyxin (XI). To synthesize N-methyl Adriamycin, a reaction mixture composed of Adriamycin (100 #M), NaCNBH3 (67 #M), and CH20 (2.5 raM) is assembled in a solution of 2 : 1 CH3CN : H20. After 1.5 hr at room temperature in the dark, the reaction mixture is analyzed by analytical HPLC [Alltech, 5 #m, Cls, 250 x 4.6 mm Econosphere column, 1 ml/min; solvent A, 50 mM ammonium acetate (pH 4.0);

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CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[ 19]

solvent B, acetonitrile; isocratic 50% solvent A]. HPLC gives three broad peaks, the first of which is determined to be Adriamycin by comparison of retention times and coinjection. The second and third peaks are determined by electrospray mass spectrometry to be N-methyl Adriamycin (XI) (MH + = 558.5), and N,N-dimethyl Adriamycin (MH + = 572.5), respectively.

CH30

0

OH

0

OH 0

0

H3C'~ HO NH I CH3 (XI) N-methyl adriamycin

Gas Chromatography-Mass Spectrometry Analysis The procedure for generation of samples for gas chromatography-mass spectrometry analysis (GC-MS) is based on that developed by Blakeney and co-workers for separating and analyzing mixtures of amino sugars. 22 The overall strategy for capturing the iminium ion intermediate is outlined here (Fig. 4, structures X I I XV). 2t Reactions of DNA with Adriamycin activated by a variety of different agents are ethanol precipitated after completion of the reaction. After the supernatant is removed and the DNA pellet dried down, 5/zl of glucosamine solution (10 mM) is added as an internal standard. Immediately, 50/zl of aqueous sodium borohydride (5 M) is added, the microcentrifuge tube used as the reaction vessel is pierced with a needle in order to prevent pressure from building up, and the reduction is allowed to progress at 65 ° for 1.5 hr. After the reduction, 50/zl of glacial acetic acid is added to consume the excess sodium borohydride. To acetylate the amino sugars, 100/zl of N-methylimidazole and 1 ml of acetic anhydride are added. After 30 min at room temperature, the acetic anhydride is quenched by addition of 2 ml of H20 and the acetylated amino sugars are extracted with 1 ml of dichloromethane. The dichloromethane is then dried down in vacuo. The samples are kept dry until immediately prior to GC-MS analysis, at which point such sample is dissolved in 100/zl of acetonitrile. GC-MS analysis is performed with a Hewlett-Packard (Palo Alto, CA) 5890 gas chromatograph and a Micromass Trio2000 quadrupole mass spectrometer. GC-MS is conducted with a DB-5 column, a source temperature of 200 °, and an 22 A. B. Blakeney, E J. Harris, R. J. Henry, and B. A. Stone, Carbohydr. Res. 113, 291 (1983).

[ 19]

CHEMICALCROSS-LINKINGOF DRUGS TO DNA 0

OH

O CH30 O

DNA

1OH

0

OH O

H+j OH2 O

k.)"

O

loss of chromophore

S

=

5 M NaBH4

CH30 O

OH O

411

.... t.;NZUN

acetic anhydride,

HH~NHH--Rl-methylimidazole H--t--OH

.o--i--H

OH3 XII: R = H XIII: R = Me

CH2OAc

H'Ji~N~RH+H-Ac H'-I--OAc

AoO--I--H

OH3 XIV: R = H XV: R = Me

HN+

IX + CH20 FIG. 4. Strategy for detection of lesion X.

oven temperature gradient as follows: 140 ° for 1 min, increase at 25°/min to 215 °, increase at 6°/min to 240 °, increase at 70°/min to 280 °, hold at 280 ° for 3 min. Autosampling is performed with a Hewlett-Packard 7674a autosampler, and data analysis is performed with Windows-based Micromass MassLynx software. Along with the samples generated by exposure of DNA to Adriamycin and various activating agents, known concentrations of Adriamycin and N-methyl Adriamycin isolated by HPLC are subjected to the reduction and acetylation procedure. These are the first samples analyzed by GC-MS; they are monitored with total ion current. This is done in order to determine the approximate retention times of the Adriamycin derivatives and glucosamine standard. Once these retention times have been determined, single-ion monitoring (SIM) is used to more precisely measure the most intense signals for XIV (98.1), XV (112.1), and the glucosamine standard (84.1). Quantification of [XV] in the DNA reaction mixtures is then performed in all GC-MS experiments by using the least-squares regression line determined for the N-methyl Adriarnycin standards versus its GC-MS response. In this manner, we have been able to find a correlation between the amount of XV observed by GC-MS and the amount of double-stranded band seen by the DPAGE

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CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[20]

methodology. More importantly, however, we have been able to detect significant amounts of XV in DNA duplexes that contained the sequence [5'-d(GC)]2 but contained no other G-C base pairs in the flanking sequence. These are sequences that generally do not show the double-stranded DNA band when analyzed by DPAGE (data not shown). The GC-MS assay described herein opens avenues for further investigation of the sequence specificity of lesion X formation, as well as its possible generation in biological systems.

Caveats about Denaturing Polyacrylamide Gel Electrophoresis Methodology The experience of our laboratory with Adriamycin underscores a serious and potentially general failure of DPAGE analysis with regard to interstrand crosslinking. The literature regarding Adriamycin, as well as a number of its derivatives, presumes that the thermal lability of this putative interstrand cross-link is the obstacle to isolation of the cross-link. It turned out that in this system, noncovalent interactions were evidently strong enough to leave the DNA duplex intact under the conditions of DPAGE. It seems unlikely that the Adriamycin family of DNAbinding agents is the only family in which this is the case.

[20] High-Resolution Complexes

Using

Footprinting Chemical

and

By K E I T H R. Fox and M I C H A E L

Studies

of Drug-DNA

Enzymatic

Probes

J. WARING

Introduction Footprinting is a method that has been developed for determining the sequencespecific binding of small molecules, oligonucleotides, or proteins to DNA. The technique, which was originally designed for analyzing protein-DNA interactions,1 is based on the ability of ligands to protect DNA from enzymatic or chemical cleavage at their binding sites. The footprinting probe cuts or modifies free DNA, while regions to which a sequence-selective agent are bound are protected from attack. The DNA fragment is labeled at the 3' or 5' end of one strand, usually with 32p, but 33p or fluorescent labels can also be used, and the regions of protection are detected by autoradiography or scanning after separating the products of digestion on denaturing polyacrylamide gels. Several enzymatic and chemical agents have been developed as footprinting probes, and each has its own characteristic advantages and disadvantages. I D, J. G a l a s and A. Schmitz, Nucleic Acids Res. 5, 3157 (1978).

METHODS IN ENZYMOLOGY,VOL.340

Copyright@ 2001 by AcademicPress All rights of reproductionin any form reserved. 0076-6879/00$35.[10

412

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[20]

methodology. More importantly, however, we have been able to detect significant amounts of XV in DNA duplexes that contained the sequence [5'-d(GC)]2 but contained no other G-C base pairs in the flanking sequence. These are sequences that generally do not show the double-stranded DNA band when analyzed by DPAGE (data not shown). The GC-MS assay described herein opens avenues for further investigation of the sequence specificity of lesion X formation, as well as its possible generation in biological systems.

Caveats about Denaturing Polyacrylamide Gel Electrophoresis Methodology The experience of our laboratory with Adriamycin underscores a serious and potentially general failure of DPAGE analysis with regard to interstrand crosslinking. The literature regarding Adriamycin, as well as a number of its derivatives, presumes that the thermal lability of this putative interstrand cross-link is the obstacle to isolation of the cross-link. It turned out that in this system, noncovalent interactions were evidently strong enough to leave the DNA duplex intact under the conditions of DPAGE. It seems unlikely that the Adriamycin family of DNAbinding agents is the only family in which this is the case.

[20] High-Resolution Complexes

Using

Footprinting Chemical

and

By K E I T H R. Fox and M I C H A E L

Studies

of Drug-DNA

Enzymatic

Probes

J. WARING

Introduction Footprinting is a method that has been developed for determining the sequencespecific binding of small molecules, oligonucleotides, or proteins to DNA. The technique, which was originally designed for analyzing protein-DNA interactions,1 is based on the ability of ligands to protect DNA from enzymatic or chemical cleavage at their binding sites. The footprinting probe cuts or modifies free DNA, while regions to which a sequence-selective agent are bound are protected from attack. The DNA fragment is labeled at the 3' or 5' end of one strand, usually with 32p, but 33p or fluorescent labels can also be used, and the regions of protection are detected by autoradiography or scanning after separating the products of digestion on denaturing polyacrylamide gels. Several enzymatic and chemical agents have been developed as footprinting probes, and each has its own characteristic advantages and disadvantages. I D, J. G a l a s and A. Schmitz, Nucleic Acids Res. 5, 3157 (1978).

METHODS IN ENZYMOLOGY,VOL.340

Copyright@ 2001 by AcademicPress All rights of reproductionin any form reserved. 0076-6879/00$35.[10

[20]

FOOTPRINTINGSTUDIES

413

Enzymes, such as DNase I, are often easy to use but typically overestimate the size of the footprint (on account of their size) and generate uneven cleavage patterns in the absence of added ligand. Chemical agents, such as dimethyl sulfate, osmium tetroxide, or diethyl pyrocarbonate (DEPC), are of limited usefulness because they react only with specific DNA bases. Other chemical probes such as methidiumpropyl-EDTA-Fe(II) [MPE-Fe(II)] act by intercalation and so perturb the DNA structure. Arguably the best cleavage agent for accurately mapping small molecule binding sites on DNA is the hydroxyl radical. P r i n c i p l e s of F o o t p r i n t i n g The DNA fragment of interest is first labeled at a 3 t or 5 f end, to allow detection of the cleavage products. After formation of the ligand-DNA complex, using appropriate buffers and equilibration conditions, the complex is subjected to cleavage. The concentration of the cleavage agent and the time of incubation are adjusted so that on average there is only one cleavage event per DNA strand (i.e., "single-hit" kinetics). To achieve this low level of cutting, conditions are chosen so that at least 50% (preferably >90%) of the DNA remains uncut. C h o i c e of DNA F r a g m e n t Footprinting fragments are usually obtained by restriction enzyme digestion of appropriate plasmids. These restriction fragments are usually between 50 and 200 base pairs in length. Longer fragments can be used, but it is more difficult to limit the digestion to conform with single-hit kinetics and the longer products are not properly resolved. Different laboratories have adopted various natural fragments as standard footprinting substrates, although both our laboratories have extensively used the 160-base pair t y r T DNA fragment 2'3 (Fig. 1). Other laboratories have used various fragments cut out of pBR322 or pBS. In many ways it would be convenient if a few fragments became recognized standards, as this would facilitate direct comparison of the affinities and specificities of ligands examined in different laboratories. It should also be noted that a limitation of the footprinting technique is that the (unknown) preferred binding site(s) for a ligand must be present within the DNA substrate. This may not be a problem for small ligands, which recognize two or three base pairs, but it can become a limitation for compounds with greater sequence selectivity unless that selectivity is known or predicted, in which case "designer" sequences or plasmid inserts can be employed. Because there are 10 possible dinucleotide steps, 64 different trinucleotides, 136 tetranucleotides, and 256 pentanucleotides, it is clear that as the specificity of the ligand increases there 2 H. R. Drew and A.A. Travers,Cell 37, 491 (1984). 3 K. R. Fox and M. J. Waring,Nucleic' Acids Res. 12, 9271 (1984).

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CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[[20]

TyrTDNA 5'-AATTCCGGTTACCTTTAATCCGTTACGGATGAAAATTACGCAACCAGTTCATTTT 3'-AAGGCCAATGGAAATTAGGCAATGCCTACTTTTAATGCGTTGGTCAAGTAAAA TCTCAACGTAACACTTTACAGCGGCGCGTCATTTGATATGATGCGCCCCGCTTCCCGA AGAGTTGCATTGTGAAATGTCGCCGCGCAGTAAACTATACTACGCGGGGCGAAGGGCT TAAGGGAGCAGGCCAGTAAAAAGCATTACCCCGTGGTGGGGGTTCCC ATTCCCTCGTCCGGTCATTTTTCGTAATGGGGCACCACCCCCAAGGGCT-5'

Multisite DNAinsert 5'-GGATCCATATGCGGCAATACACATGGCCGATTTCCAACTGCACTAGTCGTAGCGCGA 3'-CCTAGGTATACGCCGTTATGTGTACCGGCTAAAGGTTGACGTGATCAGCATCGCGCT TCAAGGTTAAGCTCCCGTTCTATCCTGGTATAGCAATTAGGGCGTGAAGAGTTATGTA AGTTCCAATTCGAGGGCAAGATAGGACCATATCGTTAATCCCGCACTTCTCAATACAT AAGTACGTCCGGTGGGGTCTGTTTTGTCATCTCAGCCTCGAATGCGGATCCTTCATGCAGGCCACCCCAGACAAAACAGTAGAGTCGGAGCTTACGCCTAGG-

3 ' 5 '

FIG. 1. Sequence of tyrT and multisite DNA fragments. The lower strand of TyrT DNA is usually labeled by filling in at the 3' end of the EcoRI site with [~-32p]dATP while the upper strand is labeled at the 3' end of the Aval site with [ot-32p]dCTP. Two clones of the multisite fragment are used in which the sequence is inserted into the BamHI site of pUC 18 in opposite orientations. HindlII-Sacl restriction fragments are then labeled at the Y end of the HindlIl site, visualizing the upper strand of MSI and the lower strand of MS2.

is a reduced chance of finding the correct site within any given restriction fragment. Even if the preferred binding site is present, a proper analysis of the specificity should examine the binding to related sequences, which may also not be present. In addition, although many ligands specifically recognize one or two bases, their binding affinity may be influenced by the nature of the surrounding bases. For these reasons one of us has prepared a multisite DNA footprinting substrate that contains all 136 tetranucleotide sequences. 4 It is freely available on request as a footprinting standard. The sequences of tyrT and the multisite fragment are presented in Fig. 1. This fragment is used to illustrate the various footprinting reactions described below. Plasmids harboring suitable DNA fragments are often prepared by cesium chloride/ethidium bromide density gradient centrifugation. However, this is expensive and time consuming and the ethidium must be removed at a later stage. Because the final stage in the preparation of a radiolabeled restriction fragment usually involves purification from a polyacrylamide gel the initial plasmid preparation need not be of the highest quality, but needs only to be of sufficient purity 4 M. Lavesa and K. R. Fox, Anal. Biochem. 293, in press (2001).

[20]

FOOTPRINTING STUDIES

415

to enable restriction enzyme digestion and polymerase activity. We routinely use Qiagen (Valencia, CA) or Promega (Madison, WI) Wizard minipreparations for extraction of pUC plasmids from 5-ml cultures and find that this generates between 100 and 200 #1 of radiolabeled DNA, which is suitable for use in footprinting reactions. DNA fragments can also be prepared by polymerase chain reaction (PCR) amplification. These products can be cut with suitable restriction enzymes and labeled at the 3' end. Alternatively, if the 5' end of one primer is blocked, then the PCR products can be directly labeled at the 5' end of the other strand. Another approach is to use a radiolabeled primer directly in the PCR mixtures. R a d i o l a b e l i n g DNA DNA fragments can be labeled at either the 3' or 5' ends. 3' Labeling is generally preferable for DNase I and hydroxyl radical footprinting as the products of digestion comigrate with Maxam-Gilbert sequence markers (see below), whereas 5' end labeling is most convenient for digestion with micrococcal nuclease and DNase II. Chemical cleavage agents, such as DEPC and OsO4, can be used with either. 5' End labeling is achieved with polynucleotide kinase and [y-32p]ATP after removal of the native Y-phosphate group with alkaline phosphatase (not necessary with synthetic oligonucleotides). 3' End labeling is achieved by filling in 3' ends left by digestion with restriction enzymes, using enzymes such as DNA polymerase (Klenow fragment) or reverse transcriptase. Enzymes DNase 1

DNase I was the cleavage agent used in the original footprinting experiments. J This enzyme is a double strand-specific endonuclease that produces single-strand nicks, cutting the O3'-P bond. 5 The enzyme requires divalent cations and has optimal activity in the presence of magnesium and calcium. 6 Manganese has been shown to stimulate the action of DNase I without altering the cleavage pattern and it is often added to buffers containing magnesium. 2 Nonetheless, the enzyme is also active in the presence of calcium as the sole divalent cation. 7 It cuts all phosphodiester bonds, and does not display any simple sequence selectivity. However, DNase I cleavage patterns are often uneven, reflecting sequence-dependent variations in local DNA structure. 2 The enzyme binds by inserting an exposed loop into

5 M. Laskowski. in "The Enzymes" (P. D. Boyer, ed.), Vol. 4, p. 289. Academic Press, London, 1971. 6 p. A. Price, J. Biol. Chem. 250, 1981 (1975). 7 K. R. Fox, Biochem. Biophys. Res. Commun. 155, 779 (1988).

416

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[9,0]

the DNA minor groove, bending the DNA toward the major groove. 8,9 As a result An-T~ tracts are poor substrates for the enzyme because they possess a narrow minor groove. GC-rich regions are also poor substrates, as they have a more rigid structure that impedes the bending that is thought to be an essential feature of the enzymatic reaction.l° The DNA-binding surface of DNase I covers about 10 base pairs, explaining why the enzyme leads to overestimations of the size of the drugbinding site. Closest phosphates positioned across the DNA minor groove are not attached to a single base pair, but are staggered by two or three bases. As a result, DNase I footprints on opposing strands are staggered by two or three bases in the 3' direction relative to each other. DNase I cuts the O3'-P bond, generating products having a phosphate at the 5' terminus and hydroxyl at the 3' end. In contrast, Maxam-Gilbert cleavage generates fragments that terminate with phosphate groups at both 3' and 5' ends.lJ For 3' end-labeled DNA substrates Maxam-Gilbert markers comigrate with the products of DNase I cleavage, because both types of labeled fragments possess 5'-phosphates. However, for 5' end-labeled DNA the radiolabeled products terminate with a 3'-hydroxyl group for DNase I but a T-phosphate for Maxam-Gilbert markers. As a result Maxam-Gilbert marker lanes, with the extra phosphate, migrate faster than corresponding DNase I cleavage products. This difference is often overlooked when footprinting with 5' endqabeled fragments, but is significant for short fragments, where the difference in mobility may be as great as two bands. In contrast, enzymes that cut the O5'-P bond, such as micrococcal nuclease or DNase II, generate products that comigrate with Maxam-Gilbert markers when labeled at the 5' (but not the 3') end. Protocol. DNase I (purchased from Sigma, St. Louis, MO) is dissolved in 0.15 M NaC1 containing 1 mM MgC12, and stored (at a concentration of about 7200 Kunitz units/ml) at - 2 0 °. This stock solution is stable to frequent freezing and thawing and is diluted to working concentrations immediately before use. Mix 2 #1 of radiolabeled DNA (dissolved in 10 mM Tris-HC1, pH 7.5, containing 0.1 mM EDTA at a concentration of > 10 cps as determined on a hand-held Geiger counter) with 2 #1 of ligand (dissolved in a suitable buffer such as 10 mM Tris-HCl, pH 7.5, containing 10 mM NaC1). It is usual to examine a range of ligand concentrations both above and below the minimum concentration required to generate a footprint. The ligand concentration is usually taken as that in the mixture before adding the enzyme. Strictly speaking, for ligands that are in rapid exchange with DNA, the concentration should refer to that after adding the enzyme, whereas for those in slow exchange (e.g., triplexes) the concentration should 8 A. Lahm and D. Suck, J. Mol. Biol. 221,645 (1991). 9 S. A. Weston, A. Lahm, and D. Suck, J. Mol. Biol. 226, 1237 (1992). ") M. E. Hogan, M. W. Robertson, and R. H. Austin, Proc. Natl. Acad. Sci. U.S.A. 86, 9273 (1989). It A. M. Maxam and W. Gilbert, Methods Enzymol. 65, 499 (1980).

[20]

FOOTPRINTINGSTUDIES

417

refer to that before adding the enzyme. It all depends on how much reequilibration of the ligand-DNA complex may be expected to occur during the time after the small dilution caused by adding enzyme. The mixture is then left to equilibrate for an appropriate length of time (usually less than 30 rain for small molecules, although it may need to be much longer for triplexes). Digestion is initiated by adding 2 #1 of DNase I (dissolved in 2 mM MgCI2, 2 mM MnC12, 20 mM NaC1). The enzyme concentration is usually determined empirically, and will depend on the reaction conditions (pH, ionic strength, temperature, etc.) as well as the DNA concentration. However, at 20 ° with 10 mM NaC1 and pH 7.5 a suitable enzyme concentration is about 0.03 Kunitz units/ml. The reaction is terminated after 1 rain by adding 3.5 ~1 of 80% (v/v) formamide containing 10 mM EDTA, 2 mM NaOH, and 0.1% (w/v) bromphenol blue. EDTA is the essential component in this stop solution, as it chelates the divalent metal ions that are necessary for DNase I activity. After digestion samples are denatured by heating to a temperature near boiling, followed by rapid cooling on ice. Samples can be loaded directly from the near boiling conditions, although heating for excessive periods can lead to depurination. The products of digestion are then resolved on a denaturing polyacrylamide gel (typically 6-12%, depending on the length of the DNA fragment). R e a c t i o n Conditions. DNase I is active over a wide range of experimental conditions and can tolerate temperatures between 4 and 70 °, ionic conditions up to 1 M, and pH values between 4.5 and 9.0. DNase I is also active in the presence of various organic solvents, which are often necessary for dissolution of insoluble ligands. A 2% (v/v) concentration of methanol or dimethyl sulfoxide (DMSO) can be included in the reaction mixture without significantly affecting enzyme activity. DMSO concentrations as high as 50% (v/v) can be tolerated, although under these conditions the cleavage pattern becomes more even as a result of changes in DNA structure. Some ligands with low sequence selectivity do not produce DNase I footprints because they are in rapid exchange with DNA. In some instances footprints can be induced by lowering the temperature, thereby decreasing the dissociation rate constant(s). 12 Although enzyme concentrations under different conditions are usually empirically determined, approximate values for relative enzyme concentrations are shown in Table I. Similarly, magnesium can be replaced with calcium, although about 10-fold more enzyme is required in the absence of magnesium. By contrast, the enzyme is inhibited by millimolar concentrations of cobalt or zinc. Some workers include a known concentration of unlabeled carrier (calf thymus) DNA in the reaction mixture. This enables accurate determination of the total DNA concentration, because the concentration of radiolabeled DNA is usually vanishingly low, and therefore produces consistent extents of cleavage with different preparations of radiolabeled DNA. However,

12K. R. Fox and M. J. Waring,Nucleic Acids Res. 15, 492 (1987).

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TABLE 1 RELATIVE CONCENTRATIONSOF DNase I THAT ARE REQUIREDTO PRODUCE EQUIVALENTAMOUNTSOF CLEAVAGEUNDERVARIOUSREACTIONCONDITIONSa Temperature (°C) 4 20 37 50 65

Relative concentration

Ionic strength

Relative concentration

10 1 0.5 1 2

0.01 0.1 1.0

1 5 20

pH

Relative concentration

5.0 6.0 7.0 8.0

50 3 1 1

a The values are relative to the concentration required to produce single-hit kinetics in pH 7.5, 10 mM NaCI at 20 °.

this can complicate the quantitative analysis of footprinting, and can usually be cmitted. Limitations and Advantages. DNase I is the most commonly used footprinting agent because it is inexpensive and simple to use. Digestion products can be loaded directly onto polyacrylamide gels without further purification, precipitation, or extraction. The major problem with using DNase I as a footprinting probe is that it produces an uneven ladder of cleavage products, so that the boundaries of the footprint may be difficult to define. In addition, because the enzyme is much larger than most DNA-binding agents it overestimates the ligand-binding site size. A further complication is that enhanced cleavage is often observed around the ligand-binding site, especially in regions such as An. Tn, which are normally poor substrates for the enzyme.13 This is also evident in triplex footprints, where cleavage of the bond at the 3' end of the target purine strand is usually (but not always) enhanced. 14'15 These enhancements have often been interpreted as arising from ligand-induced changes in the local structure of the DNA helix that render it more susceptible to enzyme cleavage. However, it should alsobe noted that apparent intensification of cleavage can also arise from changes in the ratio of free DNA to enzyme. ~6 Because DNase I cuts from the minor groove it might be expected that it would detect only the binding of ligands that interact with the minor groove. However, it is one of the most successful footprinting probes for detecting triplex formation in the DNA major groove. 15'17 This may arise from oligonucleotideinduced changes in either DNA structure or flexibility. Surprisingly, DNase I can

13 C. M. L. Low, H. R. Drew, and M. J. Waring, Nucleic Acids Res. 12, 4865 (1984). 14 T. J. Stonehouse and K. R. Fox, Biochim. Biophys. Acta 1218, 322 (1994). 15 M. D. Keppler and K. R. Fox, Nucleic Acids Res. 25, 4464 (1997). 16 B. Ward, R. J. Rehfuss, J. D. Goodisman, and J. C. Dabrowiak, Nucleic Acids Res. 16, 1359 (1988). 17 K. R. Fox, E. Flashman, and D. Gowers, Biochemistry 39, 6714 (2000).

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FOOTPRINTING STUDIES

419

be used to determine accurately the 3' end of triplex-binding sites on the purinerich target strand, and the enzyme has been successfully employed for comparing the binding sites of closely related oligonucleotides. ~7 Example. Figure 2 shows representative DNase I footprints for echinomycin (a bifunctional intercalator that is known to bind to the dinucleotide step CpG ~3'~8,t9) and Hoechst 33258 [a minor groove-binding ligand that is selective for (A/T)4 tracts2°'21]. The fragment used in this study is the multisite fragment (Fig. 1), which contains all possible tetranucleotide sequences. This fragment has been cloned into the BamHI site of pUC18 in both orientations; labeling the 3' end of the HindlII site visualizes the top strand of MS 1, while for MS2 the bottom strand is visualized. The use of these two clones together enables clear data to be obtained for both ends of this relatively long footprinting substrate. DNase I cleavage of the drug-free controls reveals an uneven cleavage ladder in which some bonds are cut more efficiently than others. The unevenness is less pronounced with this fragment than with tyrT DNA because it was designed to avoid the presence of oligopurine - oligopyrimidine tracts. Inspection of the cleavage patterns reveals that, at these concentrations, echinomycin induces footprints around each of the CpG steps. At several locations this is accompanied by enhanced cleavage, presumably reflecting drug-induced changes in the local DNA structure caused by bis-intercalation of this ligand. Hoechst 33258 produces footprints at the (A/T)4 tracts, although as previously noted TTAA, TATA, and TTAT do not present good binding sites for this ligand. 21 It should also be noted that Hoechst 33258 does not produce any regions of enhanced DNase I cleavage, presumably because it does not distort the DNA on binding. Differential cleavage plots derived from these data are presented in Fig. 6 (see below). Other Enzymes

DNase H DNase II is also a double strand-specific nuclease that produces single-strand breaks in DNA. 22 It cuts the O5'-P bond in a reaction that does not require the presence of divalent metal ions. As a result, cleavage with this enzyme cannot be terminated by addition of EDTA. A further disadvantage of DNase II is that it requires conditions of low pH. Much less is known about the structural preferences 18 M. W. Van Dyke and P. B. Dervan, Science 225, 1122 (1984). 19 G. Ughetto, A. H. J. Wang, G. J. Quigley, G. van der Marel, J. H. van Boom, and A. Rich, Nucleic Acids Res. 13, 2305 (1985). 20 M. A. Carrondo, M. Coil, J. Aymami, A. H.-J. Wang, G. A. van der Marel, J. H. Van Boom, and A. Rich, Biochemistry 28, 7849 (1989). 2t A. Abu-Daya, P. M. Brown, and K. R. Fox, Nucleic Acids Res. 23, 3385 (1995). 22 G. Bernardi, in "The Enzymes" (P. D. Boyer, ed.), Vol. 4, p. 271. Academic Press, London, 1971.

420

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

MS1 Echy

GC~

[20]

MS2 Hoechst

Echy

Hoechst

CCGG ACGT ACGC

ACGG

TCGCGC ACGA

TCGG

TO

CCGC

FIG. 2. DNase I digestion patterns of fragments MS l and MS2 in the presence of echinomycin or Hoechst 33258. The DNA fragments were each labeled at the 3' end of the HindIIl site, so that the sequence runs 3'---~5' from bottom to top of the gel. Fragment MS I corresponds to the top strand of the sequence shown in Fig. 1, while MS2 corresponds to the bottom strand. Ligand concentrations (micromolar) are shown at the top of each gel lane. The bars indicate the location of the potential ligand-binding sites: CpG for echinomycin and (A/T)4 for Hoechst 33258. Tracks labeled GA are Maxam-Gilbert sequence markers specific for purine bases.

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FOOTPRINTING STUDIES

421

of this enzyme, although it produces a less even pattern of cutting than DNase I. 23 Regions of good cleavage do not correlate across the two strands and it cuts best in oligopurine tracts that contain both A and G. This enzyme has been used only rarely as a footprinting probe. 13 Protocol. DNase II (purchased from Sigma) is dissolved in 20 mM ammonium acetate, pH 5.4, containing 1 mM EDTA and stored (at a concentration of about 1000 units/ml) at 20 °. Mix 2 #1 of radiolabeled DNA (dissolved in 10 mM TrisHC1, pH 7.5, containing 0.1 mM EDTA) with 2 #1 of ligand (dissolved in DNase II digestion buffer: 20 mM ammonium acetate, pH 5.4, containing 1 mM EDTA). Digestion is initiated by adding 2 #1 of DNase II diluted in 20 mM ammonium acetate, pH 5.4, containing 1 mM EDTA. The enzyme concentration is usually determined empirically but will typically be about 1 unit/ml. The reaction is stopped after 3 min by adding 4 #1 of formamide containing 10 mM EDTA and 0.1% (w/v) bromphenol blue. Because DNase II does not require a divalent metal ion, this is not sufficient to inactivate the enzyme and further cleavage is prevented either by placing the reaction on dry ice or by immediately heating to 100 °. Micrococcal Nuclease

Micrococcal nuclease, which is widely used for digesting chromatin to prepare nucleosome core particles, cleaves the O5'-P bond. 24 Cleavage is almost exclusively located at pA and pT bondsY '26 It is thought to bind short regions of single-stranded DNA in a cleft on the enzyme, thereby explaining its preference for cleaving at AT residues. 27 Because of its AT sequence selectivity it is not widely used as a footprinting probe, but it can be used for detecting ligand-induced changes in DNA structure and dynamics. 2~ Protocol. Micrococcal nuclease (purchased from Sigma) is dissolved in 50 mM Tris-HCl, pH 7.6, containing 2 mM CaCI2 and stored (at a concentration of about 2000 units/ml) at - 2 0 °. Mix 2 #1 of radiolabeled DNA (dissolved in 10 mM Tris-HCl, pH 7.5, containing 0.1 mM EDTA) with 2 #1 of ligand (dissolved in a suitable buffer such as 10 mM Tris-HC1, pH 7.5, containing 10 mM NaC1). Digestion is initiated by adding 2 #1 of micrococcal nuclease diluted in 50 mM Tris-HC1, pH 7.6, containing 2 mM CaC12. The enzyme concentration is usually determined empirically and will typically be about l unit/ml. Because micrococcal nuclease possesses exonuclease as well as endonuclease activity it is especially important not to overdigest 23 H. R. Drew, J. Mol. Biol. 176, 535 (1984). 24 A. Deiters, J. C. Galluci, and R. R. Holmes, J. Am. Chem. Soc. 104, 5457 (1982). 25 W. Horz and W. Altenburger, Nucleic Acids Res. 9, 2643 ( 1981 ). 26 C. Dingwall, G. E Lomonossoff, and R. A. Laskey, Nucleic Acids Res. 9, 2659 ( 1981 ). 27 F. A. Cotton, E. E. Hazen, and M. J. Legg, Proc. Natl. Acad. Sci. U.S.A. 76, 2551 (1979). 28 K. R. Fox and M. J. Waring, Biochim. Biophys. Acta 909, 145 (1987).

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CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[9~0]

the samples. The reaction can be stopped after 3 min by adding 4 ~tl of formamide containing 10 mM EDTA and 0.1% (w/v) bromphenol blue. Chemical Cleavage Agents

Hydroxyl Radicals The hydroxyl radical, a small, highly reactive and freely diffusible species, generates an even ladder of cleavage products in the absence of added ligand and can be used to assess accurately the size and location of a DNA-binding site as well as to glean some information on local DNA structural variations. 29,3° The hydroxyl radical probe is generated by the Fenton reaction: Fe 2+ + H202 --+ Fe 3+ + OH- + .OH The Fe z+ is usually complexed with EDTA, generating a negatively charged species, thereby preventing direct interaction of Fe 2+ with the DNA. Although the exact mechanism of hydroxyl radical cleavage is still not known with certainty, it is generally accepted that the free radicals attack the DNA backbone at the C4' or C 1' positions, leading to subsequent base removal and strand scission. 3k32 Most of the cleavage products contain 5'- and 3'-phosphate ends. However, a small amount of 3'-phosphoglycolate is formed as well; this product migrates slightly faster than the T-phosphate and is therefore apparent as a weaker additional band when examining short fragments that are labeled at the 5' end. As a consequence hydroxyl radical cleavage patterns are often cleaner for 3' endlabeled DNA fragments. Because hydroxyl radicals are thought to attack DNA from positions that are accessible from the minor groove, this cleavage agent is most useful for detecting the interaction of proteins and ligands that bind in the minor groove. It is a great advantage that hydroxyl radicals produce even DNA cleavage ladders, with little sequence dependence. Single- and double-stranded nucleic acids are cut with similar efficiency.33'34 Several studies have employed hydroxyl radical footprinting to examine the interaction of AT-selective minor groove-binding ligands with both natural 35 and synthetic 21'36 DNA fragments. These substances produce footprints about three base pairs long, which are staggered by about two bases in the 3' direction across

29 T, D. Tullius, B. A. Dombroski, M. E. A. Churchill, and L. Kam, Methods Enzymol. 155, 537 (1987). 3o M. A. Price and T. D. Tullius, Methods Enzymol. 212, 194 (1992). 3t W. K. Pogozelski, J. McNeese, and q\ D. Tullius, J. Am. Chem. Soc. 117, 6428 (1995). 32 W. K. Pogozelski, T. J. McNeese, and T. D. Tullius, J. Biomol. Struct. Dyn. 8, 167 (1991). 33 M. J. Jezewska, W, Bujalowski, and T. M. Lohman, Biochemistry 28, 6161 (1989) [see correction in Biochemistry 29, 5220 (1990)]. 34 D. W. Celander and I". R. Cech, Biochemistry 29, 1355 (1990). 35 j. Portugal and M. J. Waring, FEBS Lett. 225, 195 (1987). 36 A. Abu-Daya and K. R. Fox, Nucleic Acids Res. 25, 4962 (1997).

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423

the two DNA strands, typical of agents that cut from the DNA minor groove. It has been suggested that at some sites cleavage of one strand is affected more than its complement, possibly providing information about the strand against which the ligand molecules are most closely stacked. 35 The interaction of mithramycin with GC-rich sequences has also been probed by hydroxyl radical footprintingS and has been used to provide information about variations in the location of the antibiotic when bound to different arrangements of GC residues. 38 In some instances hydroxyl radical footprinting was able to detect multiple ligand-binding sites in regions where DNase I produced a single footprint. Protocol. Mix 3/zl of radiolabeled DNA (dissolved in 10 mM Tris-HCl, pH 7.5, containing 0.1 mM EDTA at a concentration of > 10 cps as determined on a hand-held Geiger counter) with 10 #1 of ligand (dissolved in a suitable buffer such as 10 mM Tris-HC1, pH 7.5, containing 10 mM NaC1). The mixture is left to equilibrate for an appropriate time. Digestion is initiated by adding 8 #1 of freshly prepared "hydroxyl radical mix," which is prepared as follows. Stock solutions of the following are prepared in the best quality water available: 100 mM ferrous ammonium sulfate (stored frozen at - 2 0 °), 0.5 M EDTA (pH 8.0), 100 mM ascorbic acid, 30%(v/v) hydrogen peroxide (Sigma). From these the following dilutions are made: Solution Solution Solution Solution

A: Add 2 #1 of 100 mM ferrous ammonium sulfate to 1 ml of water. B: Add 4 #1 of 0.5 mM EDTA pH 8.0, to 1 ml of water. C: Add 100 ttl of 100 mM ascorbic acid to 1 ml of water. D: Add 10 #1 of 30% (v/v) hydrogen peroxide to 1 ml of water.

Prepare the hydroxyl radical mix by mixing solutions A, B, C, and D in the ratio 1 : 1 : 2 : 2 and use immediately. Hydroxyl radical digestion is conducted for 10 min and stopped by adding 2 #1 of 3 M sodium acetate and 65 #1 of ethanol. The samples are left for 10 min on dry ice, before spinning at 13,000 rpm for 10 min in an Eppendorf centrifuge. The supernant is removed and the pellet is washed with 100 #1 of 70% (v/v) ethanol before drying. The radiolabeled DNA pellet is then redissolved in 8 #1 of gel loading buffer [formamide containing l0 mM EDTA, 1 mM NaOH, and 0.1% (w/v) bromphenol blue]. Samples are boiled for 3 min, before loading onto a gel as described for DNase I. Limitations. Hydroxyl radical footprinting offers a powerful means of identifying ligand- and protein-binding sites on DNA and for assessing variations in local helical structure. However, it has some limitations that need to be taken into consideration. First, hydroxyl radicals can only be used to assess the interaction with agents that bind in the DNA minor groove. Compounds that do not occlude the minor groove do not produce hydroxyl radical footprints. As a result, hydroxyl radicals cannot be used for detecting the formation of DNA triple helices. Second, 37B. M. G. Cons and K. R. Fox, NucteicAcids Res. 17, 5447 (1989). 38M. L. Carpenter,J. N. Marks, and K. R. Fox, Eur J. Biochem, 215, 561 (1993).

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C H E M I C AAND L MOLECULARBIOLOGICALAPPROACHES

[20]

some sequence-selective ligands including actinomycin, echinomycin, and nogalamycin do not generally produce hydroxyl radical footprints. 39'4° The reasons for this are not clear, because these agents produce footprints with other footprinting probes. A further limitation in using hydroxyl radicals is that the reaction is sensitive to the presence of free radical scavengers. Agents such as glycerol (used as a stabilizer in many protein preparations) and dimethyl sulfoxide or methanol (used to dissolve insoluble drugs) inhibit the cleavage reaction and therefore in their presence higher concentrations of EDTA-Fe (iI) are required. E x a m p l e . Representative hydroxyl radical footprints for the interaction of Hoechst 33258 with the multisite DNA fragment are shown in Fig. 3. It can be seen that hydroxyl radicals produce an even cleavage in the drug-free controls, Although hydroxyl radicals produce a more precisely defined assessment of the ligand-binding site than does DNase I, the footprints themselves are less clear and appear only as attenuations in the local cleavage pattern. As a result the footprints are best identified from densitometric analysis of the data; examples are shown in Fig. 4. Inspection of these plots reveals that the ligand protects four bands from cleavage around each (A/T)4 site, and that two bonds at the center of each footprint are more strongly affected. The footprints are staggered toward the 3' end of the binding site, as expected for a cleavage probe that attacks from the minor groove. Rather surprisingly, this footprinting tool is able to detect binding to some sites (such as TTAA and TTAT) that were not evident with DNase I (Fig. 2). Methidiumpropyl-EDTA-Fe( ll )

The chemical cleavage agent methidiumpropyl-EDTA-Fe(II) was devised by Dervan and colleagues. 4j'42 It consists of an intercalating moiety (methidium, originally known as dimidium) tethered to a moiety of EDTA. In the presence of Fe 2+ and under reducing conditions, free radicals are generated that cleave the DNA. MPE produces a fairly even ladder of cleavage products although, because its action is based on intercalation of its phenanthridinium chromophore, An" Tn tracts, which are poor intercalation sites, show attenuated cleavage. Although M P E Fe(II) is widely used as a footprinting probe for studying the binding of proteins and small molecules to DNA there have been surprisingly few studies in which it has been used for examining triplex formation, 17,43,44 where it produces less clear footprints than does DNase I. It is likely that this footprinting tool does not reveal clear triplex sites because it binds to triplex as well as duplex DNA and perturbs the triplex-duplex equilibrium, as does the parent molecule ethidium. 39M. E. A. Churchill, J. J. Hayes, and T. D. Tullius, Biochemistry 29, 6043 (1990). 4oK. R. Fox, Anticancer Drug Des. 3, 157 (1988). 41 M. W. Van Dyke and R B. Dervan, Nucleic Acids Res. 10, 5555 (1983), 4z M. W. Van Dyke, R. R Hertzberg, and R B. Dervan, Proc. Natl. Acad. Sci. U.S.A. 79, 5470 (1982). 43 1~A. Beal and E B. Dervan, J. Am. Chem. Soc. 114, 4976 (1992). 44C. Marchand, C. Bailly, C. H. Nguyen, E. Bisagni, T. Garestier, and C. H61~ne,Biochemistry 35, 5022 (1996).

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FOOTPRINTING STUDIES MS1

TTAA TATA A ATTA

425

MS2

TTTA, ATAA TAATT TATA

TTAA TTAT TAAA

TTTT

AAAT

TATT

ATAT

FIG. 3. Hydroxyl radical digestion of the multisite fragment in the presence of Hoechst 33258. The fragments were labeled at the 3' end of the HindlIl site, revealing the top strand of Fig. 1 for MS1 and the bottom strand for MS2. Ligand concentrations (micromolar) are shown at the top of each gel lane. The bars indicate the location of the (A/T)4 sites. Tracks labeled GA are Maxam-Gilbert sequence markers specific for purine bases.

Protocol. Radiolabeled D N A (2 #1) is m i x e d with ligand (4 #1) diluted in an appropriate buffer and left to equilibrate. Five microliters of a solution containing 1 # M M P E (Sigma) and 1 /zM ferrous a m m o n i u m sulfate is then added and the mixture is left to equilibrate for a further 5 min. C l e a v a g e is initiated by adding 3 #1 o f 10 m M dithiothreitol ( D T T ) and stopped after 60 min by ethanol precipitation, in the presence o f 0.3 M sodium acetate.

426

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[20]

MS1 TTTT

TATT

ATTAAATAT 2,~ ¢v ~..1~• ~ ,'

vv"~j V

MS2

Hoechst

TATA

i

TTAT

TAAA I . . . . .

AATT ,A. 1,1~,£~i.

T_TAAT~ ATAT • IIJ[ , A. A )

.A~AA~,A~AI~AA~I~lt~/~, ,~,v,'w~,~" ~ V

JControl FIG. 4. Densitometric analysis of the hydroxyl radical cleavage plots shown in Fig. 3 in the presence of 3 # M Hoechst 33258. Note that sequences are written 3'---*5' so that the left-hand side corresponds to the bottom of the gel.

Potassium Permanganate and Osmium Tetroxide The reagents potassium permanganate and osmium tetraoxide oxidize thymines 45,46 (and to a lesser extent cytosines) by reaction across the C5-C6 double bond. Double-stranded DNA is resistant to this oxidation because the stacked bases do not permit the out-of-plane attack by the oxidizing agent. The reagents are therefore much more reactive toward single-stranded DNA, or regions in which the C5-C6 bond of thymine is exposed. Osmium tetroxide has been used to detect echinomycin-induced changes in DNA structureY whereas permanganate oxidizes certain thymines in the presence of bleomycin. 4s The exact molecular basis for these changes in reactivity is not known, but it is thought to involve ligandinduced helix perturbations that may well include extension and unwinding. The permanganate reaction is performed by mixing 3 #1 of DNA with 3 #1 of ligand in an appropriate buffer (10 mM Tris-HCl, pH 7.5), followed by addition of 3/zl of 50 mM KMnO4. The reaction is stopped after 3 min by adding 2 #1 of 2-mercaptoethanol, and the DNA is precipitated by adding 10 #1 of 3 M sodium acetate, 75 #1 of water, and 300 #1 of ethanol. After washing and drying the pellet, 45 E. Palecek, Methods Enzymol. 212, 139 (1992). 46 C. M. Rubin and C. W. Schmid, Nucleic Acids Res. 8, 4613 (1980). 47 M. J. McLean, E Seela, and M. J. Waring, Proc. Natl. Acad. Sci. U.S.A. 86, 9687 (1989). 48 K. R. Fox and G. W. Grigg, Nucleic Acids Res. 16, 2063 (1988).

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FOOTPRINTINGSTUDIES

427

strand cleavage is achieved by adding 100/zl of 10% (v/v) piperidine and boiling for 30 min. Diethyl Pyrocarbonate

Diethyl pyrocarbonate (DEPC) reacts predominantly with N7 of purines (located in the DNA major groove of B-DNA) so that subsequent treatment with piperidine leads to strand scission. Native duplex DNA is relatively unreactive to this reagent, so it provides a means of examining DNA structures in which the N7 atom may be more exposed than usual. It has therefore been used to detect the local unwinding adjacent to drug intercalation sites. 49 Echinomycin induces DEPC-hyperreactive sites at adenines 3' to the drug-binding site (i.e, at CGA), 5° whereas bleomycin enhances DEPC reactivity at adenines in the sequence GYA. 45 The reaction is performed by mixing 3 #1 of DNA with 3 #1 of ligand in an appropriate buffer (10 mM Tris-HC1, pH 7.5). The reaction is started by adding 5/zl of DEPC. Because this reagent is immiscible with water it is necessary to agitate the tube at regular intervals. The reaction is stopped after 20 min by precipitating the DNA with 15/zl of 0.5 M sodium acetate and 60 #1 of ethanol. After washing and drying the pellet, strand cleavage is achieved by adding 100/zl of 10% (v/v) piperidine and boiling for 30 min. Example. The effects of echinomycin on DEPC and permanganate modification of the multisite fragment are shown in Fig. 5. In the drug-free controls DEPC produces weak cleavage at purine residues. In the presence of echinomycin a few bands are rendered more susceptible to reaction with DEPC. These all correspond to adenines and the strongest are located on the 3' side of echinomycin-binding sites at TCGA and ACGA. Other adenines that are not close to echinomycinbinding sites are also affected, although to a lesser extent. These may correspond to long-range changes in DNA structure that are transmitted into regions that are several nucleotides removed from the ligand-binding site. Alternatively, they may indicate the location of secondary ligand-binding sites. It is possible that the interaction of echinomycin with these secondary sites is too transient to produce a footprint, but DEPC may be able to detect the unwound DNA structure associated with these short-lived species. It should also be noted that echinomycin does not affect the reaction of DEPC with CCGA and GCGA, suggesting that the ligand has less effect on the structure of sites that are flanked by G and C residues. Figure 5 also shows that echinomycin affects the modification of several thymines by permanganate. Although one of the strongest sites of enhancements occurs at TCGT, most of the other enhancements are not close to CG sites. In

49C. Jeppesenand E E. Nielsen,FEBS Lett. 231, 172 (1988). 5oj. Portugal, K. R. Fox, M. J. McLean,J. L. Richenberg,and M. J. Waring, Nucleic Acids Res. 16, 3655 (1988).

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CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[9,0]

MS2

MS1 DEPC KMBO4

DEPC

KMBO,

<

CCGA TCCAA CA2OA

TCGI

ATCAA CCGT

CA~

ACCA | AOAA |

TAATT TATA

GCGT ATGI

ACOA I ACGT

~AIC GTATT

TCGA CA!A

~cIc

]cIc

FIG. 5. DEPC and permanganate modification of fragments MSI and MS2 in the presence of echinomycin. The fragments were labeled at the 3' end of the HindlIl site, revealing the top strand of Fig. 1 for MS 1 and the bottom strand for MS2. Ligand concentrations (micromolar) are shown at the top of each gel lane. Tracks labeled GA are Maxam-Gilbert sequence markers specific for purine bases. Sites of increased modification are indicated alongside the lane with the reactive base underlined. The asterisks show echinomycin-binding sites at which adjacent A's and T's are not affected.

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FOOTPRINTINGSTUDIES

429

addition, no enhancements are seen at CCG _T, GCGT, and A C G ! . It appears that permanganate is able to detect a peculiar feature of the interaction of echinomycin with a subset of its binding sites. A n a l y s i s of F o o t p r i n t i n g D a t a Differential Cleavage Plots Footprinting data are often presented in the form of a differential cleavage plot. 3'j3 In this analysis the fractional cleavage of a bond in the presence of the ligand (J]) is compared with the fractional cleavage of the same bond in the control (f~). The differential cleavage Ji/f~, or ln(f/f~), is then plotted against the DNA sequence. Differential cleavage values less than one correspond to regions that are protected from cleavage whereas values greater than one correspond to regions of ligand-induced enhanced cleavage. Although band intensities should properly refer to the integrated intensity of each band in the cleavage pattern, it is often easier to examine the height, rather than the area, of bands in densitometer plots. Although this procedure is not valid for determining the absolute cleavage at each position, it can be justified when comparing cleavage patterns in a single experiment. In addition, it should be noted that many commercial packages (e.g., ImageQuant) do not produce true Gaussian fits to densitometric data, but merely use rectangles derived from the positions of minium intensity. Figure 6 shows differential cleavage plots derived from the footprints in Fig. 2, for the interaction of Hoechst 33258 and echinomycin with portions of the multisite DNA fragments. Looking at the data for Hoechst (solid circles, Fig. 6) it can be seen that the footprints at isolated sites cover five or six base pairs and are staggered in the 3' direction relative to the expected binding sequences. A clear example of this effect is seen for the footprint at TTTT on fragment MS1, which covers the sequence 5'-TTGTCA. Quantitative Analysis A proper analysis of quantitative footprinting data is presented by Trauger and Dervan (see [22] in this volumeS1). However, a simple and effective procedure examines the intensity of a chosen band within the footprint or the entire footprint over a range of ligand concentrations. These values are then normalized with respect to other bands in the DNA fragment that are not affected by the ligand, so as to correct for differences in gel loading and digestion. Footprinting plots 52 are then constructed, describing band intensity as a function of ligand concentration. These are fitted with the simple binding equation 1/17o = Cso/(L + C50), where 1 and 5J j. W. Traugerand E B. Dervan,Methods Enzymol. 340, [22] 2001 (this volume). 52j. C. Dabrowiak,J. Goodisman,and B. Ward, Methods Mol. Biol. 90, 23 (1997).

430

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[20l

30

o.0~

-

GCATGAAATGTATTGAGAAGTGCGGGATTAACGA

3o1 MS2

0.03 3'-ATACGCCGTTATGTGTACCGGCTAAAGGTTGACGTGATCAGCATCGCGCTAGTTCCAATTCGAGGGCAA

FIG. 6. Differential cleavage plots, showing the interaction of 30 #M echinomycin and 3 /IM Hoechst 33258 with portions of fragments MS 1 and MS2. The open circles correspond to echinomycin, whereas the closed circles show the data for Hoechst 33258. The abscissa shows the intensity of each band in the drug-treated lane divided by the intensity of the same band in the control, plotted on a logarithmic scale. Note that sequences are written Y-->5' so that the left-hand side corresponds to the bottom of the gel.

I0 are the band intensities in the presence and absence of the ligand respectively, L is the ligand concentration, and C50 is the ligand concentration that reduces the intensity of bands in the footprint by 50%. The use of this equation to analyze footprinting data assumes that the DNA concentration is low (lower than the dissociation constant of the ligand). Under these conditions, for which the absolute DNA concentration need not be known, C50 is equal to the ligand dissociation constant. Acknowledgments Work in the authors' laboratories is funded by grants from the Cancer Research Campaign (M.J.W. and K.R.E), the Association for International Cancer Research (K.R.E), and the European Union (M.J.W.).

[21]

DRUG-RNA FOOTPRINTING

[21] Drug-RNA

431

Footprinting

By MARK P. MCPIKE, JERRY GOODISMAN, and JAMES C. DABROWIAK Introduction In the footprinting experiment, the binding sites of a drug or ligand on a DNA or RNA fragment are determined by cleaving the fragment, which has previously been allowed to come to equilibrium with the ligand, and measuring the amounts of oligomers of various lengths produced in the cutting reaction. If the bound ligand inhibits cleavage at one or more positions on the fragment, certain oligomers will be underrepresented in the cleavage products; this constitutes the "footprint" of the bound ligand on the fragment. The most convenient way of setting up the footprinting experiment is to radiolabel the original fragment at one end and detect labeled oligomers only, so that from the length of a detected oligomer the position at which cleavage took place can be determined. To avoid ambiguities when using single end-labeled substrates, the probability of more than one cleavage event occurring on any single full-length fragment should be made small (the "single-hit" regime); this means that no more than ~20% of the full-length substrates should be cleaved. With RNA, it is common to detect the site of breakage or modification on the polymer by extending a labeled DNA primer by using a DNA polymerase. Although primer extension is useful for qualitative work, stops and pauses in the extension and uncertainty concerning the precise termination site of the extended DNA make the approach unsuited for quantitative studies in which binding constants from footprinting data are being sought. The footprinting experiment involves (1) establishing equilibrium between the labeled RNA and the ligand, (2) cleaving the RNA, (3) separating the products of cleavage in a sequencing gel, (4) measuring intensities of bands corresponding to different-length fragments, and (5) comparing the intensities obtained in the control experiment, in which no ligand is present, with those obtained in experiments with ligand. In quantitative footprinting, one studies how the intensities of spots (proportional to amounts of fragments) corresponding to fragments of various lengths vary with the total amount of ligand present. For equilibrium-binding ligands that inhibit cleavage, the decrease in the amount of a fragment produced by cleavage at a given site depends on the binding constant of the ligand for the site. The site-specific binding constant can easily be determined from a plot of amount of cleavage versus concentration of free ligand, but usually only the total ligand concentration is known in the footprinting experiment. However, from "footprinting plots," showing the amount of cleavage at a site as a function of total ligand concentration, it is possible to obtain relative binding constants of sites. In addition, we have shown that absolute values of binding constants can

METHODS IN ENZYMOLOGY, VOL. 340

Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. 0076-6879/00 $35.00

432

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

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be derived if enough footprinting plots are obtained? -3 This requires knowing the concentrations of the binding site and ligand (but not the cleavage agent). We have also shown that binding constants can be derived from footprinting data without knowing the association and dissociation rates of the drug for its site or the cleavage rate of the cutting agent. 4 It is required only that the system be at equilibrium during the time course of the digest. Footprinting plots need not be simple exponential-like decay curves representing inhibition of cleavage, i-6 Sometimes an increase in cleavage with ligand concentration is observed, Such "enhancements" can be due to changes in polymer structure as ligand loading occurs, or to a mass action effect associated with redistribution of the cleavage agent on the polymer. The latter effect is important when the concentration of cleavage agent is small compared with the concentration of cleavage sites, so that, when a drug prevents the cleavage agent from approaching a site, the cleavage agent is displaced to other sites, increasing cleavage there. The mass action enhancement should be unimportant when there is enough cleavage agent to saturate the polymer, and, unlike structural changes, should affect cleavage at all sites. By studying all the footprinting plots together, it is possible to determine which mechanism is operating, and also to obtain absolute values of ligand-binding constants at many different sites, By quantitatively analyzing footprinting plots valuable information can be obtained about the structure of RNA in the presence of the drug. For example, if two groups of complementary contiguous sites separated in sequence from each other give identical footprinting plots, it may be inferred that the two sites are part of a double-stranded region and that drug is binding to the duplex structure. This would occur if the drug were binding to the stem portion of a stem-loop or other duplex regions in RNA. One may also observe identical footprinting plots for nucleotides that are well separated from each other in sequence and are not complementary. This may mean that folding brings distant parts of the structure into proximity with one another to form a binding "pocket" for the drug. The folded structure could exist in the unligated RNA or it could be formed when sites on the polymer load with drug, that is, the binding of the drug could induce a conformational change in RNA. If it is suspected that enhancements are the result of a change in RNA structure and not simply mass action effects associated with I j. C. Dabrowiak, J. Goodisman, and B. Ward, in "Drug-DNA Interaction Protocols" (K. R. Fox, ed.), p. 23. HumanaPress, Totowa, New Jersey, 1997. 2 j. C. Dabrowiak,A. A. Stankus, and J. Goodisman, in "Nucleic Acid Targeted Drug Design" (C. L. Probst and T. J. Perun, eds.), p. 93. Marcel Dekker, New York, 1992. 3 j. Goodisman, R. Rehfuss, B. Ward, and J. C. Dabrowiak,Biochemistry 31, 1046 (1992). 4 j. Goodisman and J. C. Dabrowiak,J. Biomol. Struct. Dyn. 2, 967 (1985). 5 j. Portugal and M. J. Waring,Biochirnie 69, 825 (1987). 6 j. Portugal, K. R. Fox, M. J. McLean, J. L. Richenberg, and M. J. Waring,Nucleic Acids Res. 16, 3655 (1988).

[21]

DRUG-RNA FOOTPRINTING

433

the cutting agent, enhancement occurring in the same drug concentration range as a binding event gives evidence that the two events (binding and the structural change) are correlated. Study of the footprinting plots may also reveal cooperative binding of the ligand, wherein the amount of binding to a site depends not only on the free ligand concentration, but on binding events at other sites. One possible mechanism for cooperative binding is that a ligand molecule bound at one site attracts a second ligand molecule. Another is that ligand bound at a site prevents ligand from binding nearby (this would be anticooperativity). These effects may occur by ligand-induced distortion of the local structure of the RNA, which makes nearby binding sites more or less attractive for another ligand. The packaged viral information of the human immunodeficiency virus exists as a dimer having two genomic-length RNAs linked together through Watson-Crick hydrogen bonds. 7'8 The dimer region, called the packaging region or ~pelement, spans about 360 nucleotides (nt) near the 5' end of the genome. A proposed structure of the packaging region of human immunodeficiency virus type 1 (HIV-1) (LAI) is shown in Fig. 1. The dimeric form of RNA in the virion of the virus is bound by ~2000 copies of a small zinc fnger protein called the nucleocapsid protein (NCp7). Because the packaging region is critically important for insertion of the correct RNA into the budding virus particle and is involved in a number of other steps in the life cycle of the virus, it is a potential new target for drugs directed against acquired immunodeficiency syndrome (AIDS). Whereas drug binding to the trans-activation response (TAR)9-13 and Rev-responsive element (RRE) sites TM15 ofHIV RNA has been studied, the possibility of using the packaging region, in its monomeric and dimeric forms and when complexed to protein, as a target for drug action has not been explored. In this chapter we describe quantitative footprinting methods to study the binding of the aminoglycoside drug paromomycin (Fig. 2) to a 176-nt segment of RNA from the packaging region of HIV- 1 (LAI). The drug-binding sites were identified 7 j. Coffin, in "RNA Tumor Viruses" (R. Weiss, N. Trich, H. Varmus, and J. Coffin, eds.). Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York, 1985. s j. L. Clever, D. A. Eckstein, and T. G. Parslow, J. Virol. 73, 101 (1999). 9 V. Peytou, R. Condom, N. Patino, R. Guedj, A.-M. Aubertin, N. Gelus, C. Bailly, R. Terreux, and D. Cabrol-Bass, J. Med. Chem. 42, 4042 (1999). l0 L. Dassonneville, E Hamy, E Colson, C. Houssier, and C. Bailly, Nucleic Acids Res. 25, 4487 (1997). I I H.-Y. Mei, M. Cui, A. Heldsinger, S. M. Lemrow, J. A. Loo, K. A. Sannes-Lowery, L. Sharmeen, and A. W. Czarnik, Biochemistry 37, 14204 (1998). 12 H. An, B. D. Haly, and P. D. Cook, Bioorg. Med. Chem. Lett. 8, 2345 (1998). 13 E Hamy, V. Brondani, A. Florsheimer, W. Stark, M. J. J. Blommers, and T. Kilmkait, Biochemistry 37, 5086 (1998). 14 M. L. Zapp, S. Stern, and M. R. Green, Cell 74, 969 (1993). 15 M. Hendrix, E. S. Priestley, G. E Joyce, and C.-H. Wong, J. Am. Chem. Soc. 119, 3641 (1997).

434

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with RNase I, which cuts RNA with low specificity, and to a more limited extent using RNase T 1, which cleaves at unpaired guanosine residues.16 Although drugbinding constants were not calculated from the footprinting data, we developed important experimental protocols and scanning procedures, analyzed representative footprinting plots, and identified regions of binding and enhancement on the 176-met. S y n t h e s i s of RNA The strategy employed for synthesizing RNA involves obtaining a plasmid containing the DNA sequence of interest and amplifying a specific region by polymerase chain reaction (PCR). One of the PCR primers is constructed so that amplified DNA contains the T7 polymerase promoter contiguous to coding sequences in a target DNA sequence. The amplified DNA fragment is used as a template for synthesis of RNA, using T7 polymerase in the presence of the four ribonucleoside triphosphates.17,r8 This approach, similar to that used by Gluick and Draper, 19 allows for the synthesis of RNA having a minimum of undesirable sequences in the transcript. The RNA used in this study is obtained from the plasmid pBluescript lI K S ( + / - ) (Stratagene, La Jolla, CA) containing an insert from the ~ element of HIV-1 (LAI). This plasmid is amplified by transfection into Escherichia coli Is and DNA is recovered with a Wizard Plus Minipreps DNA isolation kit (Promega, Madison, WI). A 176-nt ~p fragment containing the hairpins SL1 through SL4, within a 206-nt HIV-1 (LAI) insert in the plasmid, is targeted. This fragment is amplified by PCR with primers having the following sequences: 3' primer, 16 G. D'Alessio and J. E Riordan, "Ribonucleases: Structures and Functions." Academic Press, San Diego, California, 1997. ~7j. R. Wyatt, M. Chastain, and J. D. Puglisi, BioTechniques I t , 764 (1991). 18 j. Sambrook, E. E Fritsch, and T. Maniatis, "Molecular Cloning: A Laboratory Manual." Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York, 1989. 19 T. C. Gluick and D. E. Draper, J. Mol. Biol. 241,246 (1994).

436

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

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5'-TAATACGACTCACTATAGGAAACCAGAGGAGCTCTCTC; 5' primer, 5'-GGTTCCCATCGAATCTAATTCTCC. The 3' primer, a 38-mer, contains the T7 promoter (boldface type) followed by a 21-mer from HIV- 1 (LAI). The denaturing, annealing, and elongation temperature profile is 98, 55, and 75 ~', respectively. Analysis of the amplified fragment by agarose gel electrophoresis with incremental 25-bp markers shows the presence of the desired 192-mer template DNA. After phenol extraction, the DNA fragment is isolated with a Microcon 50 minicentrifuge filter (Millipore, Bedford, MA). Using RiboMAX (Promega) and the 192-mer template DNA containing the T7 promoter, a 176-mer ~ RNA was synthesized. Purification by polyacrylamide gel electrophoresis (PAGE), followed by electroelution and desalting procedures, results in milligram quantities of 176-mer RNA. The molar extinction coefficient of the 176-mer has been determined to be 1,220,000 M -1 cm -1 by complete hydrolysis to mononucleotides, using a procedure outlined earlier. 2° Although the sequence of the DNA template is that of the coding sequences in HIV-1 (LAI) having the appended T7 promoter, subsequent analysis of the RNA transcript in the footprinting experiments reveals that the 176-mer RNA is missing four nucleotides at positions ~304-307. At these positions the DNA may form a small hairpin loop that is not transcribed by the RNA polymerase, giving rise to a slightly shorter than expected transcript. However, for the sake of clarity, the transcript is referred to as the "176-mer" and the correct HIV-1 (LAI) coding sequences are shown in Figs. 1 and 8. The 176-met RNA is prepared for labeling with 32p by removing the 5'triphosphate on the G at position 211 in the sequence. This triphosphate, which is introduced in synthesis of RNA, is removed with calf intestinal phosphatase (CIP; Boehinger Mannheim, Indianapolis, IN), in 60/zl of I x CIP buffer (10× supplied with CIP enzyme) containing ~5 taM RNA and 1 unit of CIP. 18 The reaction is incubated at 50 ° for 1 hr followed by removal of the enzyme by phenol extraction and purification of the sample by ethanol precipitation. RNA is labeled with 20 units of T4 polynucleotide kinase (Promega) in kinase buffer, and 50/zCi of [y-32p]ATP (Amersham, Arlington Heights, IL). The labeling solution is incubated at 3T ~for 1 hr, after which time the reaction is stopped by ethanol precipitation. Labeled RNA is purified by denaturing PAGE in 1 x TBE running buffer (90 mM Tris-borate, 2 mM EDTA, pH 8). After visualizing RNA by autoradiography, the band containing the sample is excised and RNA is recovered by "crushing and soaking" the gel slab in 500 t~1 of nuclease-free water at 4 ° overnight. A 5-#1 sample of 32p-labeled RNA recovered by this method yields ~60,000 cpm on scintillation counting. Because of contamination with RNase, maintaining the full-length labeled RNA for any extended period of time can be extremely difficult.

20M. E Shubsda,M. P. McPike,J. Goodisman,and J. C. Dabrowiak,Biochemistry38, 10147(1999).

[21 ]

DRUG-RNA FOOTPRINTING

437

To minimize RNA degradation it is necessary to use nuclease-free products, that is, buffers, tips, water, and so on. Footprinting Experiments Preliminary experiments are necessary to determine the amount of each enzyme and the cutting time that will result in "single hit" kinetic conditions, in which less than "~20% of the full-length RNA is cut. In these experiments, the amount of enzyme is fixed and the cutting time is varied. Labeled and unlabeled RNA are mixed in the footprinting buffer, 11.1 mM Tris-HC1 (pH 7), in a total volume of 9/_tl and 1 /zl of the enzyme is added. The final concentrations of unlabeled RNA (in strands) and labeled RNA are 1.1 and ~0.05 # M (~, 100,000 cpm). After addition of the enzyme, the reaction medium is briefly mixed by pipette action and the reaction is allowed to proceed for 10 min at room temperature (',~20°). The cutting reaction is stopped by addition of 5 ~1 of loading buffer containing 80% (v/v) formamide, 10 mM EDTA, and bromphenol blue (1 mg/ml). After mixing it by pipette action, an 11-#1 sample of the resulting solution is loaded immediately onto a denaturing polyacrylamide gel. Electrophoresis is carried out at 500 V for 10-12 hr at a running temperature of ~20 ° to separate fragments of different lengths. To ascertain the amount of cleavage as a function of cutting time, the intensity corresponding to full-length RNA or "parent band" is measured. This involves a short-exposure autoradiograph (X-Omat film; Eastman Kodak, Rochester, NY), because we require an image of the 176-met RNA having an optical density (OD) less than 1.1. The image of the parent band is scanned, using the procedures outlined below, and the spot intensity, corresponding to the amount of full-length RNA, is plotted as a function of time. From this plot it is possible to determine the cutting time required for ~20% cleavage of the full-length RNA or "single-hit kinetics." Using 1 unit of RNase T1 or 17.5 units of RNase I with cutting times of 10 and 1.5 rain, respectively, we have carried out footprinting experiments with the drug paromomycin in a total volume of 10/zl in the Tris-HC1 footprinting buffer at room temperature ('-~20°). The general protocol is to mix radiolabeled RNA, 100,000 cpm, with unlabeled RNA, and to add increasing amounts of drug to successive samples in a total volume of 9 #1. After mixing with a pipette, each solution is allowed to stand for 30 min to reach equilibrium. The appropriate amount of enzyme in a volume of 1 #1 is then added and the sample is briefly mixed. The final concentrations of labeled and unlabeled RNA in the final reaction volume of 10/zl are "~0.05 and 1.1 #M, respectively. The final drug concentrations are given in the captions to Figs. 3 and 4. Depending on the enzyme used, cutting is allowed to proceed for 10 or 1.5 min, after which the cleavage reactions are stopped by addition of 5 #1 of formamide loading buffer. Before samples are loaded onto the

438

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES CC lane 1

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FIG. 3. Footprinting autoradiogram of cleavage of the 176-mer, using RNase 1. Lanes 1 and 2, RNA alone; lane 3, hydroxide ion cleavage ladder; lane 4, RNase TI ladder (G); lane 5, RNase A ladder (C, U). The lane number and concentration of paromomycin (micromolar), respectively, are 6, 0; 7, 2.5; 8, 5; 9, 7.5 (not scanned); 10, 10;, 1 I, 12.5; 12, 15; 13, 17.5; 14, 20; 15, 25 (not scanned); 16, 30; 17, 35; 18, 40; 19, 45; 20, 50. Refer to Fig. g for sequence and the structural designations.

[21 ]

DRtJG-RNA FOOTPRINTING

439

gel, they are heated at 95 ° for 5 min and then immediately placed on ice. Eleven microliters of each sample is placed in a lane of a denaturing polyacrylamide gel and the oligomers are separated by electrophoresis at 800 V for 8 hr at room temperature. The autoradiograph is prepared at - 2 0 °, care being taken to ensure that the optical density in the bands corresponding to cut fragments does not exceed OD I. 1. Figures 3 and 4 show footprinting autoradiograms involving RNase I and RNase T1, respectively. Q u a n t i t a t i o n o f Gel D a t a The autoradiograms are digitized with a Hewlett-Packard (Palo Alto, CA) Scan Jet 5p scanner and Sigma Scan software (landel Scientific, San Rafael, CA), yielding an intensity profile for each lane of the scanned image. The intensity profile is a plot of average intensity as a function of position in a lane, the average being over the width of the measuring line running down the lane (I 1 pixels, about one-third the total width of the band). For band intensities measured on a footprinting autoradiogram to give reliable species concentrations, the maximum optical density of the image must not exceed OD ~ 1,1, the limit for linear response of the film. If this condition is met, band intensities will be proportional to the concentrations of oligomers in the gel that produced the image on the film. Typical intensity profiles for cleavage using RNase I at 0, 10, and 15 ~M paromomycin are shown in Fig. 5. The amounts of oligomers are measured in one of two ways. The first is to fit the intensity profile to a sum of Gaussians, using PeakFit software (Jandel Scientific). Each Gaussian is defined by three parameters: location, width, and area. The area of each Gaussian is considered to be proportional to the amount of the corresponding oligomer. A problem is that it is impossible to ensure that the width of a particular Gaussian (corresponding to an oligomer of a particular length) is the same in all lanes of the gel. The variation of the width leads to appreciable variation in the intensity from lane to lane, producing relatively "noisy" footprinting plots. A better procedure, which obviates the need to carry out a fitting process, is to take the maximum intensity of a given band, minus the intensity of the same band in the control lane, as proportional to the amount of oligomer produced in the cutting reaction. This considerably reduces the scatter in the data, but leads to problems when bands are not completely resolved, so that the maxima are not explicit. Such strongly overlapped peaks occur in the top parts of gels, corresponding to large oligomers. Footprinting plots for selected sites using RNase I as a cleavage agent are shown in Figs. 6 and 7. Because intensities from different lanes (different drug concentrations) for each site are required for a footprinting plot, it is necessary to follow each peak from lane to lane. This cannot be done by using peak locations (distances down a lane) alone, because the location of a particular peak may change from lane to lane

440

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

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[21 ]

DRUG-RNA FOOTPRINTING

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as a result of thermal gradients in the gel. Only by comparing patterns in adjacent lanes, either from the intensity profiles or from the image of the autoradiogram, can the peaks be followed from lane to lane. Because a footprinting plot is a plot of spot intensity versus drug concentration, it is necessary to ensure that intensities in different lanes are comparable. A possible problem here is loading error: all spot intensities in a lane are proportional to the amount of sample loaded into the lane, and this may vary. Although the loading error has proved to be less than 10%, attempts have been made to correct for it as follows. The amount of sample loaded into a lane is proportional to the total intensity in all spots, including the parent band. However, it is difficult to measure the total intensity because, on an autoradiogram for which the optical density of the parent band does not exceed 1.1, the fragment spots are too faint to measure accurately; also, the spots for the shortest fragments are not measurable because they have run off the gel. Instead of the total intensity, either the parent band intensity or the sum of cut fragment intensities is used. The argument for using the former is that the fraction cut is small, and roughly the same in all lanes. The area of the parent band in each lane is measured by fitting it to three Gaussians plus a constant background. A correction factor is then calculated for each lane by dividing the average parent band area by the area of the parent band in the lane, and all spot intensities in a lane are multiplied by this correction factor. Use of the sum of all spot intensities corresponding to cleaved fragments instead of the total intensity is justified because intensities for some fragments should increase with drug concentration and others decrease, and intensities for most fragments should not change much in any case. Using this method, a correction factor is calculated for each lane by dividing the average sum of cleaved fragment intensities by the value of this sum for the lane, and all spot intensities in the lane are multiplied by this correction factor. Neither of the two methods can be used when peak maxima are used instead of areas to estimate amounts of fragments. In this case, uncorrected intensities are used to construct footprinting plots. Unlike DNA, RNA is easily fragmented and, as the RNA ages and is repeatedly handled, cleavages in the end-labeled full-length polymer take place. This can be easily seen in the control lanes shown in the autoradiograms in Figs. 3 and 4. If quantitative analysis is to be performed, the intensities of the sites in the control lane should first be subtracted from the corresponding spot intensities in the other lanes. For all the cases studied the amount of degradation of RNA prior to addition of the cleavage agent is less than ~2%.

FIG. 4. Footprinting autoradiogram of cleavage of the 176-mer, using RNase T I (G). Lanes 1 and 2, RNA alone; lane 3, hydroxide ion cleavage ladder; lane 4, RNase A (C, U) cleavage ladder. The lane number and concentration of paromomycin (micromolar), respectively, are 5, 0; 6, 5; 7, 10; 8, 15; 9, 20; 10, 25; 11, 30; 12, 35; 13, 40; 14, 45; 15, 50 (not scanned); 16, 55; 17, 60. Refer to Fig. 8 for sequence and structural designations.

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We are interested in obtaining the locations of drug-binding sites on the 176-mer, and in identifying any drug-induced changes in RNA structure. Measuring absolute values of binding constants would require a complete analysis of all footprinting plots covering the entire length of the 176-mer. This part of the study is in progress.

Results and Discussion

Structure of 176-mer The proposed structure of the packaging region of HIV-1 (LAI) 8 is shown in Fig. 1. This study used a subsection of the region, a 176-mer, containing SL1, SL2, SL3, and SL4. As mentioned, sequences in the packaging region of HIV- 1 are involved in a monomer--dimer equilibrium. To determine the form of RNA present in our experiments, solutions containing various amounts of unlabeled 176-mer and a small amount of labeled 176-mer RNA were analyzed by native PAGE. These experiments showed that, under the conditions used in the footprinting experiments (low total RNA and salt concentration), the 176-mer is in the monomeric form (data not shown). The presence of the monomer was confirmed by cutting the 176-mer with RNase T1, which cuts at unpaired guanosine residues. Cleavage was observed at positions 257, 259, and 261 in the palindromic loop of SL1

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analyzing RNA secondary structure2j was applied to the 176-mer. This produced the lowest energy RNA structure, shown in Fig. 8. A more detailed structural analysis of the 176-mer in the absence of drug is under way.

Paromomycin Paromomycin is a member of the aminoglycoside class of antibiotics. 22 The drug contains a central 2-deoxystreptamine ring to which are appended monoand disaccharide units (Fig. 2). The five cationic protonated amine groups of paromomycin give the drug solubility in water and provide it with a high electrostatic attraction for RNA. Relatively free rotation about the glycosidic bonds allows the drug to alter its shape and this, along with numerous possibilities for 21 D. H. Mathews, J. Sabina, M. Zuker, and D. H. Turner, J. Mol. Biol. 288, 911 (1991). 22 R. Schroeder, C. Waldsich, and H. Wank, EMBO J. 19, 1 (2000).

[21 ]

D R U G - R N A FOOTPRINT1NG

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TABLE I EFFECT OF PAROMOMYCINON FOOTPRINTING PLOTSa RNase TI

RNase T 1 Site number G223 G224 G226 A235 G237 C238 A239 G241 A242 C243 U244 C245 C248 U249 G251 G254 A255 G257 G259 G261 G266 C267 A268 A269 C274 G275 C284 G285 A286 C287 G289 G290 G292 G294 G298

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SD SD

SD SD

I I I

1 I 1

N, W

RD, W RD, W RD, W RD, W RD, W N N, W M M M, W

N, W

SD M M M

SD M M M

M M M M M

10% gel

6% gel

N

N

1 I

1

N,W

N

SD SD

SD SD

SD, W

SD, W

SD

SD

SD

SD

SD SD SD SD

SD SD SD SD N

M M M

I 1

M M M M 1

M

SD SD SD

N SD SD SD

M, W

a N, Neutral (no change in cleavage with increased drug concentration); I, increasing (increased cleavage with increased drug); RD, rapid decrease (cleavage decreases rapidly with increased drug); SD, slow decrease (cleavage decreases with increased drug, but slowly); M, maximum (cleavage increases initially with increased drug, then goes through a maximum and decreases); W, weak (low intensity). l, These three sites are representative of the region from position 361 through position 369, all of which are G, except positions 362 and 368. It is impossible to resolve individual sites in this region.

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CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

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hydrogen bonding (donation and acceptance), makes it difficult to generalize a paromomycin-RNA binding model. However, because of its large size the drug is not likely to bind in the major groove of a normal A-form helix such as would occur in stem regions of the 176-mer (Fig. 8). Puglisi and co-workers 23 used nuclear magnetic resonance (NMR) to uncover details of the binding of paromomycin to the A-site in 16S ribosomal RNA. They found that the drug, which spans about five nucleotides of RNA, adopts an "L-shape" and binds to an asymmetric internal loop caused by noncanonical base pairs in the A-site.

Footprinting Analysis of Paromomycin Binding to 176-mer Inspection of the footprinting plots for RNase I and RNase T1 cleavage of the 176-mer reveals that the former, because of its low base/sequence specificity,16 yields more information about the location of paromomycin-binding sites. By varying the electrophoresis time and the composition of the gel it is possible to resolve most of the sites (single nucleotide resolution), but those farthest from the label, especially in the experiments involving RNase I, remain unresolved. The analysis of the footprinting plots for RNase I and RNase T1 is summarized in Table I. In general, information is included only for sites that have significant intensity and for which cleavage changes noticeably with drug concentration. A footprinting gel for RNase I is given in Fig. 3. The kinetics of cutting in the case of RNase I appear to be zero order in enzyme. In this case the rate of cleavage of phosphodiester linkages of RNA is independent of enzyme concentration, so that apparent increases in the rate on addition of drug must be structural in origin. The zero-order kinetic regimen was used in DNase I footprinting experiments involving the anticancer drug actinomycin D. 24 A footprinting gel involving RNase TI is shown in Fig. 4. The kinetic regimen used in the footprinting experiments involving RNase T1 seems to be first order in enzyme. In this case, the rate of cleavage of a phosphodiester linkage, (rate)i, is given by ki [RNase T 1 ] i , where ki iS the cleavage rate constant at the site and [RNase T 1]i is the concentration of enzyme at site i. This rate law implies that drug binding at certain sites will shift enzyme to other sites, increasing (rate)i through a mass action effect and giving rise to enhancements in the footprinting data. Similar mass action effects were earlier observed in DNase I footprinting experiments involving actinomycin D and a 139-mer DNA restriction fragment. 3'25 Some footprinting plots, all for RNase I, are illustrated in Figs. 6 and 7. As can be seen from the "noise" in the plots, the error in any one optical density measurement is about 0.02 OD unit. The optical densities plotted in Figs. 6 and 7 have been corrected by subtracting the optical density in the control lane from that 23 M. 1. Recht, D. Fourmy, S. C. Blanchard, K. D. Dahlquist, and J, D. Puglisi, J. Mol. Biol. 262, 421 ( ! 996)~ 24 K. D. Bishop, P. N. Borer, Y. Q. Huang, and M. J. Lane, Nucleic Acids Res. 19, 871 ( 1991 ). 25 B. Ward, R. Rehfuss, J. Goodisman, and J. C. Dabrowiak, Nucleic Acids Res. 16, 1359 (1988).

[21]

DRUG-RNA FOOTPRINTING 260

~ A

DIS

447

SD ~G u {ZI'.AG

.%

c G

UG'¢ A ~ t ~

~u"G c

SLI

280

SL2

C.o_~

' ~'c~ .c~.G^ ~GGCG.c-300

G GCu'~ A G

24o.G _'Zc

AAA A A

.uu u G

,,AGA',.,, U A

SL3

AOG.cGAAU "~ ~G ~U A G" "G-/ U" G AAGA',;~CG IjO'GG :560 A U .~.gag '~C GG AG

uc~G C "A~ G G A

A.~U A

220,,G.~U AG

A

,

G"~ A~.G

340

320

~.~-.G

SL4 ~ . c G AGA

cA.u G A A C "G A "G A G 380 G G.~C A A

176-mer H I V - ~ - R N A FIG. 8. Energy-minimized structure of the 176-met from the packaging region ~ of HIV- 1 (LAI). The main stem includes positions 213-238 and 361-388; stem-loop 1 (SLI) includes the dimer initiation site (DIS), having the palindromic sequence 5'-GCGCGC-3'; stem-loop 2 (SL2) includes the 5' splice donor site (SD); stem-loop 3 (SL3); stem-loop 4 (SL4) includes the start codon 5'-AUG-3' for the gag gene (circled).

in the drug lane. The control lane shows fragmentation of the 176-mer prior to the addition of enzyme. Because the amount of degradation of the 176-met is minor, there is some uncertainty in locating the position in the control lane corresponding to the position of a given spot in a drug lane. This could lead to an incorrect value of the optical density to be subtracted, and is responsible for some of the optical densities in Figs. 6 and 7 becoming negative by several hundredths of a unit. No correction for loading error, which can be as large as 10% in a given lane, has been made. Because of this, and because of the "noise," it is best to be wary of any interpretation based on a single point in a footprinting plot. The plots for sites 235, 237, and 266-269 show the simple decrease in intensity with drug concentration associated with a drug-binding site. The last four are associated with adjacent cleavage sites, indicating the same binding event. Because optical density drops to zero when drug concentration exceeds 15 #M, the

448

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

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drug-binding constant is large for this site. For other footprinting plots, such as those associated with sites 300-309, optical density is still decreasing after 30/~M, indicating a much smaller drug-binding constant. The plots for sites 348 and 350 (Table I) show steady increases, or enhancements, which, because of the kinetic regimen of the cutting reaction, are most likely structural. Almost all the other plots of Fig. 6 and 7 show a rapid rise to a maximum, followed by a decrease. The rapid rise must be a structural effect; if it were due to mass action, all sites would show a similar rise, and they certainly do not. Note that the rise occurs over the same concentration range as the drug binding at the strongest binding sites, which suggests an association with those drug binding events. Both RNase I and RNase T 1 reveal paromomycin binding via decreased cleavage with added drug. The decreases occur at lower drug concentrations with RNase I than with RNase T1. Because competitive binding between drug and enzyme is being dealt with, the difference may be due to RNase T1 binding more strongly than RNase I to the RNA, so that higher amounts of drug are required to saturate the site. Without more detailed analysis it is not possible to determine whether one or more than one drug molecule is binding in any region. Considering the RNase I results (Table I and Figs. 6 and 7), it can be seen that the highest-affinity binding sites occur near positions 235,237,266--269, and 274. The first two are at a bulge in the main stem of the 176-met, and the others are near a bulge in the stem of SL1. The footprinting plots for many sites in the experiments with RNase I pass through a maximum, indicated by "M" in Table I. This occurs for regions 238-249 (which includes the linker region between the main stem and SLI as well as part of SL 1), 298-318 (the linker region between SL2 and SL3 and part of the stem of SL3), 325-336 (part of SL3 and SL4 as well as the linker region between them), and 351-361 (stem of SL4 plus the linker between SL4 and the main stem). The maxima appear at ~ 5 - 1 0 #M, the concentration range over which the primary binding sites are accepting drug. Thus, binding to the strongest sites appears to cause a change in the structure of the 176-mer. The decrease in cutting at M sites at higher drug concentration, after the maximum in the footprinting plot, shows that they are secondary sites, binding ligand at higher drug concentration. It is also possible that a decrease in cutting at high drug concentration is not due to drug binding but that a second structural change is occurring. This could be caused by a binding event in another part of the 176-mer. Interestingly, non-contiguous nucleotides sometimes appear to be involved in the same binding process. For example, compare the RNase I footprinting plots (Fig. 6) for nucleotides 235 (A) and 237 (G) with those for sites 266-269 (GCAA). Sites 266-269 flank the bulge in SL1, whereas nucleotides 235 and 237 are in a bulge in the main stem. It is possible that these are separate drug-binding events. However, because loading is taking place in the same concentration range for all of the sites (including nucleotide 274), they may be forming a single site that accepts one drug molecule. Site-directed mutagenesis of the 176-mer to selectively

[21]

DRUG-RNA FOOTPRINTING

449

modify sites and complete quantitative analysis of the footprinting plots should help determine which situation is correct. The footprinting plots for sites 298-318 strongly resemble those for sites 325336 (see Table I and Fig. 7). These nucleotides are not contiguous. All the footprinting plots show maxima at about 10 #M, indicating structural enhancements connected to the primary binding events. It is not unreasonable that all the sites are reacting to the same structural change, but it is unusual that the binding (decrease in cleavage after the maximum) is also the same at all sites. Again, it may be that the sites form a pocket that accepts one drug molecule, or it may be that more than one drug site, having by coincidence the same properties, is involved. The RNase T1 footprinting plots show drug binding at sites 223,224, and 226 (a bulge in the main stem of the 176-mer), 251 and 254 (in the stem of SL 1), 289294 (in the stem and loop region of SL2), 328-342 (near bulges in the stems of SL3 and SL4, plus the linker region), and the region 361-369 (near a bulge in the main stem). The footprinting plots for these sites show a decrease in intensity with increasing drug concentration, sometimes after an initial increase. As is evident from the RNase TI experiment shown in Fig. 4 and the data collected in Table I, enhancements are observed in the palindromic loop of SL1, sites 257, 259, and 261. The enhancements could be due to altered structure, or simply a mass action effect, because the kinetics of cutting are believed to be first order for this enzyme. Summary and Conclusions RNase I and RNase T1 can be used to obtain high-quality footprinting information for paromomycin binding to a 176-mer RNA from the packaging region of HIV-1 (LAI). Controls and scanning procedures are necessary for quantitation of autoradiographic data, so that footprinting plots showing cutting behavior as a function of drug concentration can be used to identify binding sites and regions of altered structure on the 176-mer. From the RNase I footprinting results the primary paromomycin binding sites on the 176-mer are on the main stem and on the stem of SL1, but noncontiguous sequences may be involved in the same binding event. Strong enhancements in cleavage with added drug are also observed, indicating drug-induced structural changes. Drug binding may cause linker regions between stem-loops of the 176-mer to change structure, possibly providing a site or sites for additional drug binding. Because drug binding changes the structure of the packaging region, which may alter its function, paromomycin analogs with enhanced specificity for HIV ~RNA have potential as a new class of agent for treating AIDS. Acknowledgments We thank P. N. Borer for the energy-minimized structure of the 176-mer and for helpful discussions pertaining to the research. This work was in part supported by a grant, GM32691, to P. N. Borer.

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[22] Footprinting Methods for Analysis of Pyrrole-lmidazole Polyamide/DNA Complexes B y JOHN W. TRAUGER a n d PETER B. DERVAN

Introduction The design of synthetic ligands that read the information stored in the DNA double helix has been a long-standing goal at the interface of chemistry and biology. Cell-permeable small molecules that target predetermined DNA sequences enable a potential approach for the regulation of gene expression. The development of "pairing rules" for minor groove-binding polyamides offers a code to control sequence specificity.l'2 Crescent-shaped polyamides containing three aromatic amino acids, N-methylimidazole (Im), N-methylpyrrole (Py), and N-methyl3-hydroxypyrrole (Hp), bind double-helical DNA with affinities and specificities comparable to those of natural DNA-binding proteins. 3-6 Sequence specificity is controlled by side-by-side pairs of rings that afford unique set(s) of sequencespecific hydrogen bonds with the minor groove edges of the intact Watson-Crick base pairs. 7'8 The pair Im/Py specifies a G- C base pair while Py/Im targets C. G. The Py/Py pair binds both A. T and T. A. Hp/Py and Py/Hp pairs discriminate T. A from A. T.l-8 Although the generality of these pairing rules has been demonstrated by synthesizing a large number of polyamides that bind many different sequences, we find that there are limitations with regard to some sequence contexts, which are probably due to the sequence-dependent microstructure of the DNA helix. Remarkably, eight-ring polyamides have been shown to permeate cells and, in a few encouraging examples, inhibit the expression of target genes in cell culture experiments .9,10

I p. B. Dervan and R. W. Biirli, Curr. Opin. Chem. Biol. 3, 688 (t999). 2 D. E. Wernmer, Annu. Rev. Biophys. Biomol. Struct. 29, 439 (2000). 3 j. W. Trauger, E. E. Baird, and E B. Dervan, Nature (London) 382, 559 (1996). 4 S. White, E. E. Baird, and P. B. Dervan, Chem. Biol. 4, 569 (t997). 5 j. M. Turner, S. E. Swaltey, E, E. Baird, and E B. Dervan, J. Am. Chem. Soc. 120, 6219 (1998). 6 S. White, J. W. Szewczyk, J. M. Turner, E. E. Baird, and E B. Dervan, Nature (London) 391, 468

(1998). 7 C. L. Kielkopf, E. E. Baird, P. B. Dervan, and D. C. Rees, Nat. Struct. Biol. 5, 104 (1998). 8 C. L. Kielkopf, S. White, J. W. Szewczyk~ J. M. Turner, E. E. Baird, P. B. Dervan, and D. C. Rees, Science 288, 111 (1998). 9 j. M. Gottesfeld, L. Neely, J. W. Trauger, E. E. Baird, and P. B. Dervan, Nature (London) 387, 202

(1997). 10 L. A. Dickinson, IL J. Gulizia, J. W. Tranger, E. E. Baird, D. E. Mosier, J. M, Gottesfeid, and P. B. Dervan, Proc. Natl. Aead. Sci. U.S.A. 95, 12890 (1998).

METHODS1NENZYMOLOGY,VOL.340

Copyright© 2001by AcademicPress ~1 rightsof reproduction~nany.formreserved. 0076-6879/00$35.00

[22]

FOOTPRINTINGTO ANALYZEPy-Im/DNA

451

A pivotal step in the discovery--evaluation process of new polyamide motifs is the characterization of the affinity and specificity of next-generation molecules following the design-sythesis phase (i.e., structure-function relationships). In the design phase, it is often the case that the sequence preference, as well as the energetics of any new molecule binding at each potential site, are not perfectly understood and it is crucial to scan "libraries" of many potential DNA-binding sites in order to identify true high-affinity binding sites. Characterization of equilibriumassociation constants must guide the choice of sequence context for DNA : ligand complexes selected for subsequent structure elucidation by nuclear magnetic resonance (NMR) or X-ray methods. A DNA restriction fragment, typically a few hundred base pairs in length, provides a library of contiguous small molecule-binding sites on a string addressed by a 32p end label. The high-resolution separation of nucleic fragments by gel electrophoresis underpinning "footprinting" methods has revolutionized the field of small molecule-DNA discovery. This chapter describes three complementary footprinting methods and the protocols used for analysis of polyamide : DNA complexes: (1) MPE-Fe(II) footprinting,1 l- 14 (2) affinity cleavage,15-J7 and (3) quantitative DNase I footprint titration 18-21 (Fig. 1). Footprinting with methidiumpropylEDTA-Fe(II) [MPE-Fe(II)] is used to identify high-affinity polyamide-binding sites to near nucleotide resolution (Fig. 1A). Affinity cleavage is used to determine the orientation of the bound polyamide in the minor groove of DNA (Fig. 1A). Quantitative DNase I footprinting is used to determine equilibrium association constants (K~) for polyamide-DNA complexes at previously identified match and mismatch sites (Fig. 1B).

Description of Methods Footprinting was first described for analysis of protein-DNA complexes using the enzyme DNase I. 22 DNase I has been used to footprint small molecules as

l! M. W. Van Dyke, R. P. Hertzberg, and P. B. Dervan, Proc. Natl. Acad. Sci. U.S.A. 79, 5470 (1982). J2 M. W. Van Dyke and P. B. Dervan, Cold Spring Harbor Syrup. Quant. Biol. 47, 347 (1982). 13 M. W. Van Dyke and E B. Dervan, Nucleic Acids Res. 11, 5555 (1983). 14 M. W. Van Dyke and E B. Dervan, Science 225, 1122 (1984). 15 j. S. Taylor, E G. Schultz, and E B. Dervan, Tetrahedron 40, 457 (1984). 16 p. G. Sehultz and E B. Dervan, J. Biomol. Struct. Dyn. 1, 1133 (1984). 17 p. B. Dervan, Science 232, 464 (1986). 18 M. Brenowitz, D. E Senear, M. A. Shea, and G. K. Ackers, Methods Enzymol. 130, 132 (1986). 19 M. Brenowitz, D. E Senear, M. A. Shea, and G. K. Ackers, Proc. Natl. Acad. Sci. U.S.A. 83, 8462 (1986). 20 D. F. Senear, M. Brenowitz, M. A. Shea, and G. K. Ackers, Biochemistry 25, 7344 (1986)~ 21 M. Brenowitz, D. E Senear, E. Jamison, and D. Dalma-Weisha~sz, in "Footprinting of Nucleic Acid-Protein Complexes" (A. Revzin, ed.), p. 1. Academic Press, San Diego, California, 1993. 22 D. Galas and A. Schmitz, Nucleic Acids Res. 5, 3157 (1978).

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CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

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(A)

5'

3'

5'

3'

(B) C

Increasing [Ligancl] B==

1 0.8 Eo.6 a:~ 0 . 4

Site I

Ref[

0.2-

o lO-ll

10.1o

.... 1~0.9 '

l!O.a ',

1~0.7

Increasing [Ligand]

FIG. 1. (A) Schematic of (left) MPE-Fe(II) footprinting and (right) affinity-cleaving techniques. Cleavage products obtained on a denaturing gel with DNA end labeled either on the 5' or 31 strand are shown. (B) LeJ?: Cleavage pattern generated by quantitative DNase I footprint titration, on a 31 endlabeled DNA fragment in the presence of increasing ligand concentrations. Right: Langmuir binding titration isotherm obtained from DNase I data.

well as proteins. 13,23 However, DNase I does not cleave all sequences equally and footprints can appear significantly larger than the actual base pairs occupied by the D N A - b i n d i n g ligand. 13 Both natural product D N A - b i n d i n g ligands, as well as designed polyamides, typically cover only five or six base pairs and accurate assignment of the b i n d i n g site sequence becomes important. The chemical footprinting reagent, M P E - F e ( I I ) , based on the nonspecific intercalator m e t h i d i u m with E D T A - F e attached, cleaves D N A 24,25 with m i n i m a l sequence specificity in 23 K. R. Fox and M. J. Waring, Nucleic Acids Res. 12, 9271 (1984).

10-s

[22]

FOOTPRINTINGTO ANALYZEPy-Im/DNA

453

the presence of a reducing agent and 02, and allows a higher resolution footprint. By directly attaching the EDTA-Fe moiety to the DNA-binding ligand, the orientation of a ligand in its complex with DNA can be determined in a technique called affinity cleavage. 15-17 In addition, affinity cleavage experiments are useful for determining the groove location of a ligand, because distinct 5'- or T-shifted cleavage patterns are observed for cleavage in the major versus the minor groove. Minor groove-binding ligands produce 3'-shifted cleavage patterns j7 (Fig. 1A). Quantitative DNase I footprinting procedures were developed by Ackers and co-workers, and are the foundation of the protocol presented here. 18-21 For quantitative footprinting experiments, equilibrium mixtures of 32p end-labeled DNA and a range of polyamide concentrations are subjected to partial digestion with DNase I. The resulting cleavage products are separated by gel electrophoresis and visualized by autoradiography. DNA sites bound by a polyamide are protected from cleavage, resulting in a gap in the ladder of bands on the gel. The tractional protection of a binding site as a function of polyamide concentration can be determined by densitometric analysis of the gel, and these data can be fitted to a theoretical binding isotherm to determine the equilibrium association constant (Ka) (Fig. 1B). The data analysis for this assay assumes that the polyamide concentration is much higher than the DNA concentration, allowing the approximation [L]free = [L]tot, where [L]fre°is the concentration of polyamide free in solution and [L]tot is the total polyamide concentration. The total equilibration volume specified in this protocol is 400 #1 and the DNA concentration is ~5 pM. As a consequence, apparent association constants greater than 4 2 x 10 ~° M ~ should be regarded as lower limits. In the protocol described here, polyamide and radiolabeled DNA are equilibrated in the absence of any unlabeled carrier DNA (such as calf thymus DNA) to avoid underestimation of association constants due to polyamide binding to sites on the carrier DNA. Materials

DNase I [fast protein liquid chromatography (FPLC) pure], calf thymus DNA (sonicated, deproteinized), and NICK columns are purchased from Pharmacia (Piscataway, NJ) Restriction enzymes, polynucleotide kinase, Taq DNA polymerase, Klenow DNA polymerase, and glycogen are obtained from Boehringer Mannheim (Indianapolis, IN). Sequenase (version 2.0) is purchased from United States Biochemicals (Cleveland, OH). [ot-32p]Thymidine 5'-triphosphate (>3000 Ci/mmol) and [ot-32p]deoxyadenosine 5'-triphosphate (>6000 Ci/mmol) are purchased from Du Pont-NEN (Boston, MA). [F-32p]adenosine triphosphate (endlabeling grade) is from ICN (Costa Mesa, CA). Denaturing polyacrylamide gel 24R. E Hertzbergand P. B. Dervan,J. Am. Chem. Soc. 104, 313 (I 982). 25R. R Hertzbergand R B. Dervan,Biochemistry 23, 3934 (1984).

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CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

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mix, Tris-borate-EDTA (TBE) packets, and gel plates, spacers, and stands are obtained from GIBCO-BRL (Gaithersburg, MD). Presiliconized 1.5-ml tubes are obtained from Sorenson Biosciences, Inc.

P r e p a r a t i o n of 32p E n d - L a b e l e d DNA End-labeled DNA is prepared from plasmid DNA containing the binding site(s) of interest. The desired DNA fragment is generally prepared by restriction digest and 3' fill-in, using a DNA polymerase and [a-32p]dNTPs (Procedures 1 and 2, below), but can also be prepared by polymerase chain reaction (PCR), using a 5/ 32p-labeled primer (Procedure 3, below). When labeling by 3' fill-in, it is best to cut the end to be labeled with an enzyme whose 51 overhang contains only two different nucleotides because the overhang can then be completely filled in with two labeled dNTPs. The DNA-binding sites should ideally be located 40-100 base pairs from the labeled end of the DNA fragment, although sites of up to ~200 base pairs from the label can usually be resolved. The size of the DNA fragment can range from 100 base pairs to more than 500 base pairs.

Procedure l Procedure 1 is the most convenient labeling procedure, and can be used if (l) both enzymes cut efficiently in the same buffer, and (2) the enzyme that cuts at the unlabeled end leaves either a blunt end or a 3' overhang. Enzyme combinations that work well are EcoRI*/PvuII (this example) and AflII*]FspI (the asterisk denotes the labeled end). 1. Combine plasmid DNA (4 #g) and water in a total volume of 55/zl. Add 7 #1 of H buffer (Boehringer Mannheim), 4/~1 of EcoRI (20 units/#l), and 4 #1 of PvuII (20 units/#l). Incubate for 2-4 hr at 37 °. Add 8 ttl of water, 5 #1 of H buffer, 10 #1 of [~-32P]dATR 10/zl of [a-32p]dTTP, and 10 #1 of Klenow DNA polymerase (2 units/#l). Allow the reaction to proceed for 25 rain at room temperature. Add 5 ttl of dATP/dTTP mix (each at 5 mM) and allow the reaction to proceed for 5 rain. 2. Add 30 #1 of 5x Ficoll loading buffer, mix, and load on a nondenaturing 7% (w/v) polyacrylamide gel (two lanes, each ~75 #1). Run the gel and isolate the DNA as described below.

Plvcedure 2 In this example (EcoRI*/HindIII), the second cutter, HindIII, produces a 5' overhang and requires a buffer different from that required by EcoRI. For these reasons, the second cut is done after the first cut and fill-in.

[221

FOOTPRINTINGTO ANALYZEPy-Im/DNA

455

1. Combine plasmid DNA (4 #g) and water in a final volume of 46/zl. Add 8/zl of H buffer (Boehringer Mannheim), 10 #1 of [a-32p]dATP, 10/xl of [c~-32p]dTTP, 4 #1 of EcoRI (20 units/ml), and 2 #1 of Sequenase (8-25 units/ml). Incubate at 37 ° for 4 hr. Add 5/zl of dATP/dTTP mix (each at 5 mM) and 0.5 #1 of Sequenase, and incubate at 37 ° for 20 min. Pass through a NICK column equilibrated with water to remove unincorporated nucleotides. To the 400/zl of eluate add 25/zl of 4 M NaC1, 1/xl of glycogen (20 mg/ml), and 860 #1 of absolute ethanol. Centrifuge at 14,000 rpm for 30 min in a cold room and decant the ethanol solution. Wash the pellet with 70% (v/v) ethanol (100 #1), decant, and dry briefly in a Speed-Vac (Savant Instruments, Hicksville, NY). 2. Dissolve the DNA pellet in 42/~1 of water, 5 #1 of buffer B (BoehringerMannheim), and add 3 #1 of HindlII. Incubate at 37 ° for 2-4 hr. Add 15 #1 of 5x Ficoll loading buffer and load on a 7% (w/v) nondenaturing gel. Run the gel and isolate the DNA as described below. Procedure 3 When suitable restriction sites are not present, a 5' 32p end-labeled DNA fragment can be prepared by PCR. 1. To 60 pmol of primer A (the primer to be labeled) add 80 #1 of water, 10 #1 of 10× kinase buffer, 4 #1 of [y-32p]ATP (~250 #Ci), and 6 #1 of polynucleotide kinase. Incubate at 37 ° for 30 min. Add 5 #1 of 0.5 M EDTA and extract with 25 : 24 : 1 (v/v/v) phenol-chloroform-isoamyl alcohol (four times, 100 #1 each). Purify the DNA with a NICK column equilibrated with water to remove unincorporated ATP. To the 400/zl from the NICK column add 24 #1 of 4 M NaC1, 1 #1 of glycogen (20 mg/ml), and 860 #1 of absolute ethanol, mix, and centrifuge (14,000 rpm) for 30 min in a cold room. Decant, wash with 70% (v/v) ethanol (100 #1), decant, and dry briefly in a Speed-Vac. 2. Dissolve 60 pmol of primer B in 60 #1 of water. To the pellet from step 1, add 50/zl of the primer B solution, 33 #1 of water, 10 #1 of PCR buffer (Boehringer Mannheim), 3.7 #1 of plasmid DNA (0.003/zg/ml), 2 #1 of dNTP mix (each at 10mM), and 1 #1 of 100x bovine serum albumin (New England BioLabs, Beverly, MA). Transfer the solution to a PCR tube. 3. Heat the tube at 70 ° in a thermocycler for 5 min (hot start PCR). Carefully add 4 units (0.8 #1) of Taq DNA polymerase, mix, and cover the solution with mineral oil. 4. Thermocycle as follows: 30 cycles of (1) 94 ° for 1 min, (2) 54 ° for 1 min, (3) 72 ° for 1.5 min, and then 72 ° for 10 min. 5. Carefully transfer the aqueous phase to a new tube. For each 100 #1 of solution, add 30/A of 5 x Ficoll loading buffer. Load onto a 7% (w/v) nondenaturing gel and purify as described below.

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Gel Purification of End-Labeled DNA 1. Prepare 7% (w/v) nondenaturing gel mix as follows: to a 125-ml Erlenmeyer flask add 8.1 g of acrylamide (caution: acrylamide is toxic!), 290 mg of bisacrylamide, and 24 ml of 5x TBE, and dilute to 120 ml with water and stir until homogeneous. Initiate polymerization by adding 700 ttl of 10% (w]v) ammonium persulfate (APS) and 42 #1 of N,N,N',N'-tetramethyl ethylene diamine (TEMED), and pour the gel. Prepare 1 liter of I x TBE gel running buffer. Run the gel at ~200 V for ~ 2 hr. 2. Carefully take the gel down and cover it with Saran Wrap. Expose the gel to X-Omat film (Kodak, Rochester, NY) for 30-60 sec. Place the developed film under the gel and cut out the desired band, using a razor blade. Transfer the gel slice to a 1.5-ml tube. 3. Crush the gel slice with a pipette tip and add 700 #1 of elution buffer (20 mM Tris-HC1, 250 mM NaCI, pH 8). Place the tube in a shielded container and soak overnight (8-16 hr) in a 37 ° shaker. Filter the DNA, using a plastic centrifugal filter (Quiksep; Isolab, Singapore) into a 15-ml Falcon 2059 tube, and transfer the eluate to a 1.5-ml tube. Add 1.5 volumes of 2-propanol, mix by inversion, and precipitate by spinning in a microcentrifuge at 14,000 rpm for 30 rain in a cold room. Decant, and then add 100 #1 of 75% (v/v) ethanol, spin briefly (30-90 sec), decant, and dry the DNA pellet briefly in a Speed-Vac. 4. Dissolve the DNA in 100 #1 of water and pass it through a NICK column equilibrated with water. Divide the 400-#1 eluate into aliquots, which are then counted in a scintillation counter and stored at - 8 0 ° until use. P r e p a r a t i o n of P o l y a m i d e S e r i a l D i l u t i o n s Solid-phase methods for the synthesis of polyamide have been described, z6 Polyamide stock solutions are conveniently prepared from dry 25-nmol aliquots. Concentrations are determined by UV absorption, using an experimentally determined extinction coefficient. Extinction coefficients can be estimated on the basis of the number of aromatic rings, using the relation 8690 M- 1 cm- 1/aromatic ring for the absorption maximum between 290 and 315 nm (e.g., for an eightring polyamide, ~ ~ 69,500 M ~ cm -1 at the maximum between 290 and 315 nM). Polyamides having at least one positive charge per eight rings are generally freely soluble up to a concentration of at least 500 ttM. The presence of multiple f%alanines can reduce solubility. It is good practice, especially for polyamides expected to have relatively low solubility, to spin the initial stock solution in 26 E. E. Baird and P. B. Dervan, J. Am. Chem. Soc. 118, 6141 (1996).

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FOOTPRINTINGTO ANALYZEPy-lm/DNA

457

a microcentrifuge for several minutes at high speed (14,000 rpm) to pellet any insoluble material and transfer the supernatant to a new tube before measuring the concentration. For quantitative footprinting experiments, the goal is to prepare binding reactions over a range of polyamide concentrations that produce from 0 to 100% fractional saturation of the DNA-binding site(s) of interest. This generally requires at a minimum a three order of magnitude concentration range. In a typical titration, 10-13 data reactions, 1 control reaction (with no polyamide), and l undigested DNA control should be prepared. Prepare polyamide stock solutions as follows: Dissolve 25 nmol of polyamide in water to give an approximately 10 #M solution. Determine the concentration of this solution by UV absorption, and then prepare 10× polyamide stock solutions with nearly equal spacing on a log scale (e.g., 1, 2, 5, and 10 nM). Quantitative DNase I Footprinting Procedure 1. Prepare a 1.14× DNA solution: Combine 2.06 ml of 5x TKMC, pH 7.0 (50 mM Tris-HC1, 50 mM KC1, 50 mM MgC12, 25 mM CaC12, pH 7.0}, 7.0 ml of water, and 32p-labeled DNA (400,000 cpm). The amount of DNA used is calculated to give a final loading of --~15,000 cpm per lane. 2. Prepare equilibrium binding mixtures: To a 1.5-ml presiliconized Eppendoff tube add 40 #1 of 10x polyamide solution (or water for the control lanes) and 350/zl of 1.14× DNA solution. Allow the mixtures to equilibrate at room temperature for 4-24 hr. Polyamides with association constants over 109 M -l should be allowed to equilibrate for at least 12 hr. 3. Prepare DNase I stop buffer: Combine 40 #1 of glycogen (20 mg/ml), 40/zl of 1 mM bp calf thymus DNA, 107 ~1 of water, 788 #1 of 4 M NaC1, and 425/zl of 0.5 M EDTA, pH 8.0. The EDTA in this buffer quenches DNase I activity by chelating the essential Mg 2+ and Ca 2+ ions. The other ingredients provide for good ethanol precipitation and subsequent resuspension. 4. Prepare a DNase I stock solution: Combine 975/zl of water and 20 #1 of 50 mM dithiothreitol (DTT), and chill the solution on ice. Add 5/zl of DNase I (Pharmacia FPLCPure, 7500 units/ml) and mix the resulting 38 units/ml-solution by inverting the tube several times. Prepare the final stock solution by adding 15-70 #1 of DNase I (38 units/ml) to a prechilled mixture of 20/zl of 50 mM DTT and 930-965 #1 of water (the total final volume should be 1000/zl). Keep the DNase I solution on ice throughout the experiment (prepare this solution freshly from the 7500-units/ml stock and use within l hr). The exact amount of DNase I to use depends on the batch of DNase I and the restriction fragment being used. The goal is to achieve ~50% digestion of the restriction fragment. Although the exact amount of DNase I to use can be determined by running a titration, it is

458

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usually possible to estimate on the basis of the length of the restriction fragment. For a 220-bp fragment, dilute ~ 5 0 / z l of the 38-units/ml stock solution to 1 ml. Shorter restriction fragments require more DNase I, longer fragments less (e.g., for a 440-bp fragment, use ~25 #1 of the 38-units/ml solution). 5. Digest the DNA: To each tube (except the intact DNA control) add 10 #1 of the final DNase I solution and mix by vortexing. Allow the reaction to proceed for 7 rain at room temperature, and then add 50/~1 of "stop buffer" and mix by vortexing. Add 975 #1 of room temperature absolute ethanol and mix by inversion. Carry out this step while using a stopwatch: First, open the tops of all tubes, then start the stopwatch. Every 15 sec, add DNase I to a tube, close the top, mix by vortexing, and place the tube in a microcentrifuge. When all the tubes are in the microcentrifuge, spin them briefly (~2 sec), remove from the centrifuge, and open all the tops. When 7 rain has elapsed, reset the stopwatch. Add stop buffer every 15 sec and vortex. Finally, add ethanol and mix. 6. Precipitate the DNA: Spin the tubes in a microcentrifuge in a cold room at 14,000 rpm for 25-30 min (start the samples spinning right away, and do not spin for longer than 30 min: spinning longer may make resuspension of the DNA more difficult). After precipitation the pellets should be visible. Carefully decant the supernatant, and then add 300 #1 of 70% (v/v) ethanol. Vortex the tube briefly to thoroughly wash the pellet. Spin the tubes briefly (10-20 sec) in a microcentrifuge, and then carefully decant the supernatant. 7. Resuspend the DNA: To each tube add 15 /zl of water and vortex (5-10 sec) to resuspend the DNA (dissolving the samples in water and reconcentrating them increases the chances that the DNA will resuspend properly and not hang in the wells). Freeze the samples by placing them in liquid nitrogen, or in a - 8 0 ° freezer. The samples may be stored overnight at this point at - 8 0 °. Concentrate the solution to dryness in a Speed-Vac (to reduce the chances of hanging lanes, do not overdry; it is best to take the samples out as soon as they are dry). Next, add 7 #1 of 80% (v/v) formamide-1 x TBE loading buffer (prepared by mixing 8 ml of formamide with 2 ml of 5 x TBE). The loading buffer should be stored at 4 ° and discarded after 1-2 months (use of old loading buffer can result in smearing of bands on the gel). Thoroughly vortex each tube (~30 sec) to resuspend the DNA. 8. Denature and load the gel: Denature the DNA by heating for 10 min at 85-90 °, and then immediately place the samples on ice. Load 5/~1 per lane on a prerun 8% (w/v) [ 19 : 1 (w/w) acrylamide-bisacrylamide] denaturing polyacrylamide gel (prerun the gel for 1 5 4 0 min until the temperature of the front gel plate reaches 50-55°). Load one or more chemical sequencing reaction lanes? 7,2s Run the gel at 27A. M. Maxamand W. S. Gilbert,Methods Enzymol. 65, 499 (1980). 28B. L. Iversonand P. B. Dervan,Nucleic' Acids Res. 15, 7823 (1987).

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FOOTPRINTINGTO ANALYZEPy-Im/DNA

459

'-~2000 V (running bromphenol blue to the bottom of the gel is good for resolving DNA sites 40-80 base pairs from the labeled end). To help ensure that the gel sticks to only one glass plate when the plates are separated, one of the gel plates may be treated with a silanizing reagent (e.g., SigmaCote; Sigma, St. Louis, MO) occasionally (once every 10 runs or so). 9. Transfer the gel to drying paper, cover with plastic wrap, and dry on a gel dryer for 60 min at 80 °. Expose the gel to a storage phosphor screen (Molecular Dynamics, Sunnyvale, CA) for 8-16 hr.

Data Analysis Image the gel with a phosphorImager such as the Molecular Dynamics 400S PhosphorImager. Intensity data are obtained from the gel image by quantitation using ImageQuant software (Molecular Dynamics). Background-corrected (i.e., quantitate the intact DNA control lane and subtract this background value) volume integration of rectangles encompassing the footprint sites and a reference site at which DNase I reactivity is invariant across the titration provides values for the site intensities (/site) and the reference intensities (Iref), respectively. The apparent fractional occupancy (0app) of the sites is then calculated using Eq. (1): 0app = 1 --

(lsite/lref)/(lOsite/l°ref)

(1)

w h e r e I0site a n d / O r e f are the site and reference intensities, respectively, from a control lane to which no polyamide was added. The ([L], 0app) data points are then fitted to a general Hill equation [(Eq. (2)] by minimizing the difference between 0ap p and 0fit: 0fit = 0mi n + (0ma x --

Omin)(Kna[L]n/1 + K"a[L]n)

(2)

where [L] is the total polyamide concentration, Ka is the equilibrium association constant, and 0mi n and 0max are the experimentally determined site saturation values when the site is unoccupied or saturated, respectively. The data are fitted by a nonlinear least-squares fitting procedure (such as KaleidaGraph software; Synergy, Reading, PA) with Ka, 0min, and 0max as the adjustable parameters, and with n fixed at either 1 or 2. A good fit to a Langmuir isotherm [Eq. (2), n = 1] is consistent with formation of a 1 : 1 polyamide-DNA complex, whereas a good fit to a cooperative isotherm (Eq. (2), n = 2] is consistent with cooperative dimeric binding. 29 The data are normalized by Eq. (3): 0norm = (0app -- 0min)/(0max -- 0min)

(3)

29 C. R. Cantor and E R. Schimmel, "Biophysical Chemistry, Part III: The Behavior of Biological Macromolecules," p. 863. W. H. Freeman, New York, 1980.

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Reported association constants are the average values obtained from three independent footprinting experiments.

M e t h i d i u m p r o p y l - E D T A - F e ( I I ) F o o t p r i n t i n g P r o t o c o l a n d Affinity Cleavage Protocols The protocol reported here avoids the use of carrier DNA and gives apparent affinities that are the same as those obtained using the DNase I footprinting assay described above. The final solution conditions are 20 mM HEPES, 200 mM NaCI glycogen (50 tig/ml), 5 mM DTT, pH 7.3, and either (1) 0.5 tiM MPE-Fe(II) for footprinting experiments or (2) 1 tiM Fe(II) for affinity cleavage experiments. 1. Prepare a 1.43× DNA solution: To a 15-ml Falcon 2059 tube add 1286 til of 5× cleavage buffer (100 mM HEPES, 300 mM NaCI, pH 7.3), 225 #1 of 4 M NaC1, 3 ml of water, and labeled DNA (250,000 cpm). 2. Prepare 10× polyamide (or EDTA-polyamide) stock solutions. 3. Set up equilibrations: To a 1.5-ml Eppendorf tube add 280 #1 of 1.43 × DNA solution, 40 #1 of 10x polyamide, and 40 #1 of glycogen (0.5 mg/ml). Allow the solution to equilibrate for 1-18 hr. 4. Prepare a precipitation buffer by combining 35 #1 of glycogen (20 mg/ml), 35 #1 of calf thymus DNA (1 mM bp), and 180 #1 of water. 5a. For MPE footprinting experiments: Prepare 5 #M MPE-Fe(II) by combining equal volumes of 10 tiM MPE and freshly prepared 10 #M Fe(NH4)2(SO4)2. To the polyamide-DNA solution add 40 til of 5 tiM MPE-Fe(II); allow to equilibrate for 10 min. Add 10 #1 of 200 mM DTT (use a freshly thawed DTT aliquot). Allow cleavage to proceed for 30 min. Add 1 ml of ethanol, mix the tubes by inversion, and spin briefly. 5b. For affinity cleavage experiments: To the EDTA-polyamide/DNA solution add 20 til of freshly prepared Fe(NH4)2(SO4)2 and equilibrate for 10-30 min. Add 40 til of DTT (use a freshly thawed aliquot) and allow cleavage to proceed for 30 min. Add 1 ml of ethanol, mix the tubes by inversion, and spin briefly. 6. Precipitate the DNA and run the gel: Add 10 #1 of precipitation buffer to each tube and mix by inversion. Spin the tubes at 14,000 rpm in a cold room for 30 min. Decant, wash with 75% (v/v) ethanol (350 #1), and decant. Resuspend the DNA in 16 til of water, freeze, and Speed-Vac dry. Resuspend in 7 til of 80% (v/v) formamide-1 x TBE loading buffer, heat denature (10 min at 85-90 °, then place on ice), and load 5 til per lane on a denaturing 8% (w/v) gel and subject to electrophoresis. 7. Transfer the gel to paper, cover with plastic wrap, and dry on a gel dryer (45-60 min) at 80 °. Expose the gel to a storage phosphor screen for 8-16 hr and image with a Phosphorlmager.

[22]

FOOTPRINTINGTO ANALYZEPy-Im/DNA o, HN

461

I N

~/, .

,

o o

I

~

. N2N'JL'.~"N~:

I

1 (R=H) 1-E (R=RE)

".

".

O

O

.

.~ O I

.

RE = I ~ . - ' N ' - . P ' N ~, '

~,. N'~

I

N O

o

c

,%0 °

H H H+ "~j...~N....~-.,~N~.~/N-R '~' ~ ~ "

FIG. 2. Structures of polyamide ImPy-fl-lmPy-y-lmPy-/3-ImPy-/3-Dp (1) and EDTApolyamide ImPy-/3-ImPy-y-lmPy-/%ImPy-/3-Dp (l-E) (/3,/3-alanine; y, y-aminobutyric acid; Dp, dimethylaminopropyl).

Data Analysis The extent of cleavage protection [for MPE-Fe(II) footprinting] and cleavage intensities (for affinity cleavage experiments) are determined by quantitation of the gel, using ImageQuant software. For MPE-Fe(II) footprinting experiments, 0app values are calculated for each band, using Eq. (1). For affinity cleavage experiments, quantitation of cleavage products provides the cleavage efficiency directly. C h a r a c t e r i z a t i o n of I m P y - ~ - I m P y - y - l m P y - ~ - I m P y - ~ - D p We reported previously that the eight-ring polyamide ImPy-j6-1mPy-y-lmPy]%ImPy-fl-Dp (1) (Fig. 2) binds to 5'-(A,T)GC(A,T)GC(A,T)-3' target sequences within the human immunodeficiency virus type I (HIV- i) promoter region that are adjacent to binding sites for the cellular transcription factors TBP and Ets- l, blocks HIV-I transcription in vitro, and inhibits HIV-I replication in cell culture, l° As an example application of the quantitative DNase I footprinting protocol described here, we present results of experiments that (1) assessed the specificity of I for a match site versus several mismatch sites, and (2) compared equilibrium association constants determined with TKMC buffer (10 mM Tris-HCl, 10 mM KCI, I0 mM MgCI2, 5 mM CaCl2, pH 7.0), which provides optimal DNase I activity, with those obtained with solution conditions approximating those expected to prevail within a living cell. Complexes of 1 with binding sites on the 282-bp EcoRI/PvuII restriction fragment from pJT2B2 were characterized by quantitative DNase I footprinting, MPEFe(II) footprinting, and affinity cleavage experiments. Plasmid pJT2B2 was prepared by hybridizing the complementary oligonucleotides 5'-CCGGCTTAAGTTC GTGGGCCATGCTGCATTCGTGGGCCATGGTGGATTCGTGGGCCATGTTA CATTCG-3' and 5'-TCGACGAATGTAACATGGCCCACGAATCCACCATGGC

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CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

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5I

I

A A C A G C ]-T

I TC G A C

] T C

T A A C A

C A T

C A

I

3' FIG. 3. Storage phosphor autoradiogram of a denaturing 8% (w/v) polyacrylamide gel used to separate the products generated by DNase 1 digestion in a quantitative footprinting experiment with polyamide L Lane 1, A-specific sequencing lane (see Dervan17); lane 2, DNase I digestion products obtained in the absence of polymide; lanes 3-12, DNase I digestion products obtained in the presence of 0.002, 0.005, 0.01, 0.02, 0.05, 0.1, 0.2, 0.5, 1, and 2 nM polyamide 1, respectively; lane 13, intact DNA. Polyamide-binding sites and the reference site (shaded bracket) are indicated along the right side of the autoradiogram. All reactions contained 15 kcpm of 32p end-labeled restriction fragment, 10 mM Tris-HCl, 10 mM KC1, 10 mM MgCI2 and 5 mM CaCI2 (pH 7.0, 24°).

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TABLE 1 EQUILIBRIUMASSOCIATIONCONSTANTSOF POLYAMIDE 1 FOR ITS MATCH SITE 5'-TGCTGCA-3 'a Buffer t'

Temperature(°C)

Ka (M -I)

TKMC

24 37

1.5 x 10 t° (0.3) 8.4 × 109 (2.3)

lntracellular

24 37

1.9 × 10 I° (0.6) 1.1 x 101°(0.2)

a Values reported are the mean values from at least three DNase I footprint titration experiments. The standard deviation for each value is indicated in parentheses. ~' TKMC buffer: 10 mM Tris-HCl, 10 mM KC1, 10 mM MgCI2, and 5 mM CaCI2, pH 7.0 at 24 °. Intracellular buffer: 10 mM HEPES-HCI, 140mMKCI, 10 mM NaCI, 1 mMMgCI2, I mM spermine, pH 7.2.

5'-AGCTGTT-3' and the "reverse orientation" site 26 5'-TCGTCGA-3'. The (0norm, [L]) data points for 1 binding to these sites were well fit by Langmuir binding isotherms [Eq. (2), n = 1], consistent with formation of the expected 1 : 1 "hairpin" polyamide-DNA complexes. Polyamide 1 is >50-fold specific for its match site 5'-TGCTGCA-3' relative to the double-base pair mismatch sites 5'-TC_C_CAC_CA-3' and 5'-TGTAACA-3'. Additional footprinting experiments indicate that increasing the equilibration temperature from 24 to 37 °, and changing the solution conditions from standard polyamide assay conditions (i.e., TKMC buffer: 10 mM Tris-HC1, 10 mM KC1, 10 mM MgCI2, 5 mM CaC12, pH 7.0 at 24 ° to conditions modeling those encountered within a typical mammalian cell 3° (140 mM KCI, 10 mM NaC1, 1 mM MgC12, I mM spermine, pH 7.2) has little effect on the affinity of polyamide 1 for its match site 5'-TGCTGCA-3' (Table I). We note that DNase I activity is nearly 100-fold lower in the model intracellular buffer compared with TKMC. MPE-Fe(II) footprinting and affinity cleavage experiments confirm that 1 binds to its match site 5'-TGCTGCA-3' (Fig. 6). The observed affinity cleavage pattern, which consists of roughly equal cleavage patterns at both ends of the binding site, is expected because the ligand can bind its pseudosymmetric match site in two orientations. Cleavage patterns are shifted toward the 3' end of each strand at the binding site, consistent with binding of the ligand in the minor groove of DNA. 30 R. J. Jones, K. Y. Lin, J. F. Milligan, S. Wadwani, and M. D. Matteucci, J. Org. Chem. 58, 2983 (1993).

[22]

FOOTPRINTINGTO ANALYZEP y - I m / D N A

A

B

q u_ t,- t~.

1 2 3 4 5

465

t-

Z

I nM1

6 7 8 9

5' - T G G C C C A C G A A T ( ~ C A ( ~ . A T G G C C C 3' - ACCGGGTGCTTACGTCGTACCGGG-

1 nM1-E

,11,

,111,

5 ' - TGGCCCACGAATGCAGCATGGCCC3' -ACCGGGTGCTTACGTCGTACCGGG-

'II'

3' 5'

'II'

3 ' 5 '

5' ~ ~

T C G

C ~

~ ~,

~

~

m

S

T

"]

A

,-I

n

T ....

C A

. . . . . . .

C

.,1 C

A

..... ....

3j

FIG. 6. (A) Storage phosphor autoradiogram of a denaturing 8% (w/v) polyacrylamide gel used to separate the products generated by an MPE-Fe(II) footprinting experiment with polyamide 1 and by an affinity cleavage experiment with polyamide | - E . Lanes 1, 6, and 8, A sequencing lanes; lanes 2 and 4, MPE-Fe(II) cleavage products obtained in the absence of 1; lane 3, MPE-Fe (II) cleavage products obtained in the presence of 1 nM 1; lane 5, intact DNA [no MPE-Fe(II)]; lane 7, cleavage products obtained in the presence of 1 nM l - E ; lane 9, intact DNA (no I-E). The reactions contained 15 kcpm of restriction fragment, 20 mM HEPES, 200 mM NaCI, glycogen (50 #g/ml), and 5 mM DTT at pH 7.0, and 0.5/zM MPE-Fe(II) [for MPE-Fe(II) footprinting, lanes 2 4 ] or 1 # M Fe(II) (for affinity cleavage, lanes 7 and 9). (B) Results of MPE-Fe(II) footprinting (top) and affinity cleavage experiments (bottom) with polyamides I and I-E, respectively. Horizontal and vertical bar heights are proportional to the amount of cleavage protection and cleavage, respectively, at the indicated base.

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Acknowledgments We are grateful to the National Institutes of Health (General Medical) and the National Foundation for Cancer Research for research support, and to the National Science Foundation and the Ralph M. Parsons Foundation for predoctoral fellowships to J.W.T.

[23] High-Resolution Transcription Assay for Probing Drug-DNA Interactions at Individual Drug Sites By DON R. PHILLIPS,SUZANNEM. CUTTS, CARLEENM. CULLINANE,and DONALD M. CROTHERS Introduction Approximately half the anticancer agents in routine current clinical use are known to interact with DNA by one or more of the following mechanisms: (1) intercalation [e.g., doxorubicin (Adriamycin), mitoxantrone], (2) groove binding (e.g., distamycin), (3) formation of covalent adducts and/or cross-links (e.g., cisplatin, melphalan, mitomycin C), and (4) incorporation of modified bases (e.g., 5fluorouracil, 6-thioguanine). Although the exact mechanisms of action of these agents remain unresolved in some cases, the apparent critical role of DNA has prompted the initiation of a wide range of studies of these drug-DNA interactions. A detailed understanding of the chemical/biochemical aspects of such interactions with DNA (particularly categories 1-3 above) would be expected to provide the necessary insight to design new generations of more active derivatives. New therapeutic approaches have indeed become apparent as a result of improved understanding of the molecular detail of drug-DNA interactions. Some of the fundamental properties of drug-DNA complexes include (1) the DNA sequence involved in the interaction, (2) the kinetics of the interaction, and (3) the affinity of the interaction. There have been two distinct phases in the development of experimental approaches to determine these parameters. Early procedures relied on a variety of physicochemical methods such as detergent sequestration, equilibrium dialysis, and spectrophotometric/spectrofluorimetic binding studies. Although these procedures all yield useful and valuable information concerning the overall drug-DNA interaction, a general limitation is that they usually yield only average binding parameters resulting from the multiple equilibria occurring at a multitude of binding sites on heterogeneous DNA and do not provide details of the drug-DNA interaction at individual sites on the DNA. To overcome this limitation several new approaches were subsequently developed that relied on the use of identical sequences of DNA (usually from plasmids), rather

METHODS IN ENZYMOLOGY,VOL. 340

Copyright(c) 2(101by AcademicPress All rightsof reproductionin any form reserved. 0076-6879/00$35.00

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Acknowledgments We are grateful to the National Institutes of Health (General Medical) and the National Foundation for Cancer Research for research support, and to the National Science Foundation and the Ralph M. Parsons Foundation for predoctoral fellowships to J.W.T.

[23] High-Resolution Transcription Assay for Probing Drug-DNA Interactions at Individual Drug Sites By DON R. PHILLIPS,SUZANNEM. CUTTS, CARLEENM. CULLINANE,and DONALD M. CROTHERS Introduction Approximately half the anticancer agents in routine current clinical use are known to interact with DNA by one or more of the following mechanisms: (1) intercalation [e.g., doxorubicin (Adriamycin), mitoxantrone], (2) groove binding (e.g., distamycin), (3) formation of covalent adducts and/or cross-links (e.g., cisplatin, melphalan, mitomycin C), and (4) incorporation of modified bases (e.g., 5fluorouracil, 6-thioguanine). Although the exact mechanisms of action of these agents remain unresolved in some cases, the apparent critical role of DNA has prompted the initiation of a wide range of studies of these drug-DNA interactions. A detailed understanding of the chemical/biochemical aspects of such interactions with DNA (particularly categories 1-3 above) would be expected to provide the necessary insight to design new generations of more active derivatives. New therapeutic approaches have indeed become apparent as a result of improved understanding of the molecular detail of drug-DNA interactions. Some of the fundamental properties of drug-DNA complexes include (1) the DNA sequence involved in the interaction, (2) the kinetics of the interaction, and (3) the affinity of the interaction. There have been two distinct phases in the development of experimental approaches to determine these parameters. Early procedures relied on a variety of physicochemical methods such as detergent sequestration, equilibrium dialysis, and spectrophotometric/spectrofluorimetic binding studies. Although these procedures all yield useful and valuable information concerning the overall drug-DNA interaction, a general limitation is that they usually yield only average binding parameters resulting from the multiple equilibria occurring at a multitude of binding sites on heterogeneous DNA and do not provide details of the drug-DNA interaction at individual sites on the DNA. To overcome this limitation several new approaches were subsequently developed that relied on the use of identical sequences of DNA (usually from plasmids), rather

METHODS IN ENZYMOLOGY,VOL. 340

Copyright(c) 2(101by AcademicPress All rightsof reproductionin any form reserved. 0076-6879/00$35.00

[23]

In Vitro TRANSCRIPTIONASSAY

467

than the heterogeneous sequences previously used with bacterial or mammalian DNA. These approaches include DNA footprinting (utilizing DNase I or hydroxyl radicals) to yield preferred drug-binding regions, and inhibition of the processivity of DNA-dependent enzymes such as DNA polymerase, DNA exonucleases, and RNA polymerase. The RNA polymerase inhibition assay results in the accumulation of truncated transcripts arising from blockage of the movement of RNA polymerase along the DNA at specific drug-binding sites. An overview of the procedure is shown in Fig. 1. The length of the truncated RNAs therefore reveals the location of these drug-binding sites. In vitro procedures that are based on blocking the progression of RNA polymerase along double-stranded DNA (dsDNA) are now referred to as "in vitro transcription assays" (and are analogous to DNA polymerase inhibition assays), whereas the term "bidirectional transcription footprinting" denotes the use of two counterdirected promoters, such that truncated transcripts arising from transcriptional blockages to both sides of a drug site define the physical "footprint" of the drug-occupied region. In addition to defining the sequence specificity of drug-binding sites on DNA, the transcription assay can also be exploited to yield the relative drug occupancy at each site as well as the complex lifetime at each individual site. The transcription assay is therefore a powerful technique that is able to yield several molecular parameters simultaneously to define how and where a drug interacts with DNA, and this information can be determined at individual drug sites on the DNA. In this chapter we outline the experimental approach to the in vitro transcriptional analysis of drug-DNA interactions. The approach requires a synchronized population of initiated transcription complexes containing RNA of a common length, which is achieved with promoters with a range of specific characteristics: (1) they should not require additional activating elements such as catabolite activator protein (CAP) and cAMP (these would add an unnecessary degree of complexity to the assay); (2) they should be "strong" promoters (i.e., the RNA polymerase should have a high affinity for the promoter region); (3) "slippage" of the start site of transcription should be minimal (the fidelity of the start site of transcription should be >99%, or capable of being "forced" to that level of fidelity); (4) the sequence of the nascent RNA must be such that a stable initiated complex can be formed in the absence of one or more of the four nucleotide triphosphates (5) the rate of formation of the initiated transcripts should be rapid (for experimental convenience); and (6) the half-life of the initiated transcription complex should be at least several hours, A range of promoters appears to satisfy all these criteria (Table I). Because the UV5 promoter had been extensively characterized it was used for the development of this assay, l and has continued to be the major promoter used in our laboratories. I D. R. Phillips and D. M. Crothers,Biochem&try25, 7355 (1986).

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~[ RNAPOLYMERASE

HEPARIN

I

INITIATIONNUCLEOTIDES

DRUG

I

-

-

I

ELONGATIONNUCLEOTIDES

~ |

FIG. 1. Overviewof the transcriptionassay. The majorsteps are bindingof RNA polymeraseselectivelyto the lacUV5 promoter, formationof a synchronizedinitiatedtranscriptioncomplex(see Fig. 2 for details), reactionof the initiatedtranscriptioncomplex with the drug of interest, and elongationof the transcriptioncomplex to yield drug-inducedblocked transcripts.

I s o l a t i o n of 5 1 2 - B a s e P a i r lacUV5 DNA F r a g m e n t The lacUV5 promoter is obtained from the plasmid pHWO1, which contains a single copy of the LSUV5 double mutant, 203-bp lac promoter cloned into the EcoRI site of pHW1.2 Only the UV5 mutation at - 9 (numbering with respect to the mRNA start site at + 1) is significant for this work because this confers strong "up"

2 H. Wu and D. M. Crothers, Nature (London) 308, 509 (1984).

[23]

In Vitro TRANSCRIPTION ASSAY

469

TABLE I PROMOTERS THAT YIELD SYNCHRONIZEDINITIATEDTRANSCRIPTIONCOMPLEXES Promoter

Initiation dinucleotide

Absent initiation nucleotide

Initiated transcript length

Ref. a

UV5 N25 TetR SP6 T3 T7 ~-PL

GA AU AG AG AG AG AU

CTP CTP GTP GTP CTP UTP UTP

10 29 1I 9 12 13 15

1 2 3 4 4 4 3

"Key to references: (1) R. J. White and D. R. Phillips, Biochemistry 27, 9122 (1988); (2) R. J. White and D. R. Phillips, Biochemistry 28, 6259 (1989); (3) H. Trist and D, R. Phillips, Nucleic" Acids Res. 17, 3673 (1989); (4) R. J. White and D. R. Phillips, Biochemistry 28, 4277 (1989).

p r o m o t e r c h a r a c t e r i s t i c s to the p r o m o t e r a n d i n i t i a t i o n o f t r a n s c r i p t i o n d o e s not r e q u i r e a c t i v a t i o n b y CAP. 3 A g o o d s u m m a r y o f the s e q u e n c e c h a r a c t e r i s t i c s o f the 2 0 3 - b p f r a g m e n t is available. 4 T h e 2 0 3 - b p f r a g m e n t was ligated into the E c o R I site o f p B R 3 2 2 to yield pRW1.5 T h e U V 5 p r o m o t e r c a n b e e x c i s e d as a 4 9 7 - b p +1

I

5 ' - GGAATTGTGAGCGGATAACAA'I-I'TCACACAGG 3 ' - CC-I-I-AACACTCGCCTATTGTTAAAGTGTGTCC

GpA E.

coli RNA polymerase

ATP, GTP, [32P]UTP +1 I v 5 ' -GGAATTGTGAGCGGATAACAATTTCACACAGG 3 ' - CC-FFAACACTCGCCTATTGTTAAAGTGTGTCC

GAAUUGUGAG ~",xx

T e m p ' l a t e DNA s t r a n d

Radiolabeled

nascent RNA

FIG. 2. Synchronized initiated transcription complex. Initiation of the lacUV5 promoter with E. coli RNA polymerase, GpA, ATP, GTP, and [c~-32p]UTPresults in a stable transcription complex containing a nascent RNA mainly 10 nucleotides in length, because CTP is absent (denoted by the arrowhead). Minor amounts of 17-mer and 23-mer transcripts are also formed, presumably resulting from CTP contaminants of other components (or from misincorporation of other nucleotides). The nascent RNA begins at the - 1 position with G of GpA present in the initiation mixture. The first nucleotide of the transcript formed under normal conditions is denoted as + 1. Radiolabel (32p) is incorporated into the nascent RNA at three sites, denoted with asterisks.

470

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

[23]

PvulI/SalI fragment. The 497-bp fragment from pRW1 (containing the UV5 promoter) has also been cloned directionally into pSP64 to yield a much higher copy number plasmid, pCC 1. The plasmid yield from this vector is significantly greater than from pBR322-derived vectors. The UV5 promoter is then excised as a 512-bp PvulI/HindlII fragment. 6 The lacUV5 promoter is also available commercially in the vector pGH6 [American Type Culture Collection (ATCC), Manassas, VA]. Procedure

1. Restriction digest pCC 1 with PvulI and HindlII [or use appropriate restriction enzymes/polymerase chain reaction (PCR) primers to isolate the lacUV5 promoter from other sources]. 2. To separate the resulting two DNA fragments, subject the restriction digest to electrophoresis, using a 1.5% (w/v) minisubmarine agarose gel in Tris-borateEDTA (TBE) buffer (lacking ethidium bromide) for 2 hr at 10 V/cm. If ethidium bromide is included in the agarose gel, then single-strand nicks may be induced in the DNA, and this will result in a high background of truncated transcripts during the elongation phase of the transcription assay. 3. Cut a thin slice lengthwise from the gel and stain with ethidium bromide (0.5 #g/ml). 4. Visualize the location of the 512-bp fragment in the gel slice with a transilluminator. 5. Align the gel slice with the main gel and excise the 512-bp fragment from the main agarose gel. 6. Place the agarose gel slice in a Biotrap chamber (Schleicher & Schuell, Keene, NH) and electroelute the DNA fragment. The Biotrap electroelution procedure has a high efficiency of recovery of DNA from agarose (typically greater than 95%). 7. Purify the DNA further by extracting with an equal volume of phenolchloroform and then subject to ethanol precipitation. 8. Redissolve the DNA in TE buffer to a concentration of approximately 100 #g/ml. F o r m a t i o n of S y n c h r o n i z e d I n i t i a t e d T r a n s c r i p t i o n C o m p l e x e s Formation of synchronized complexes initially requires an interaction between Escherichia coil RNA polymerase and DNA containing the [acUV5 promoter.

The use of heparin displaces bacterial RNA polymerase from nonspecific binding 3 A. E. Silverstone,R. R. Arditti, and B. Magasnik,PJvc. Natl. Acad. Sci. U.S.A. 66, 773 (1970). 4 F. Schaeffer,A. Kolb,and H. Buc,EMBOJ. 1, 99 (1982). 5R. J. White and D. R. Phillips,Biochemistry 27, 9122 (1988). 6C. Cullinaneand D. R. Phillips,Nucleic Acids Res. 21, 1857(1993).

[23]

In Vitro TRANSCRIPTIONASSAY

471

sites on the DNA, including the ends of the linear DNA, which have modest affinity for the polymerase. This procedure ensures that only single-copy transcripts result from the subsequent elongation step because the RNA polymerase will be unable to rebind to the promoter because of competition with heparin. Initiation is accomplished by the addition of a nucleotide mix containing a radiolabeled nucleotide (usually UTP) but lacking CTP. The resulting initiated transcription complex comprises a nascent RNA predominantly 10 nucleotides long, which is paused at the first dGTP of the template strand (see Fig. 2), and is sufficiently stable for most drug-DNA studies, with a half-life of 23 hr at 37 °.7 Transcription from the UV5 promoter does not begin exclusively from the + 1 site. When all four nucleotides are present only 59% of transcripts begin at the + 1 site, with 29, 7, and 9% beginning from the - l , +2, and -t-5 positions, respectively.8 To ensure that transcription begins from one site only, all nucleotides are maintained at <5 #M. Because these levels are too low for incorporation of the first nucleotide into the transcription complex, little initiation occurs. Initiation can therefore be achieved with great selectivity by using a high concentration of the dinucleotide GpA, which creates a nascent RNA starting from the - 1 location (see Fig. 2). If a longer half-life is required then a longer initiating oligonucleotide must be employed. Additional stability of the transcription complex may be gained by ensuring that the nascent RNA is longer than a 10-mer, and this can be achieved by initiating the transcription complex with the trinucleotide dGGA, or by using other promoters with early-transcribed sequences appropriately constituted to yield long nascent RNA from only three nucleotides in the initiation mixture. 9- ~1 It is important to use high-purity, sterile water to prepare all solutions for this procedure because the presence of trace amounts of metal ions, bacteria, or nucleases can completely destroy transcription complexes. Because of the limited lifetime of dithiothreitol (DTT), especially under alkaline conditions, use only fresh aliquots. The use of an RNase inhibitor is optional for short reaction and elongation times, but becomes increasingly necessary for reactions in the 2- to 20-hr time range. The exact MgC12 concentration in the transcription buffer is critical to ensure efficient transcription and minimal natural pausing/2 Different buffer conditions are required for bacteriophage RNA polymerases.13 Fresh 32p nucleotides (less than 2 weeks old) should be used because radiolytic degradation products can inhibit transcription.

7 C. Cullinane and D. R. Phillips, Bioehemistry 29, 5638 (1990). 8 A. J. Carpousis, J. E. Stefano, and J. D. Gralla, J. Mol. Biol. 157, 619 (1982). 9 H. Trist and D. R. Phillips, Nucleic Acids Res. 17, 3673 (1989). 10 R. J. White and D. R. Phillips, Biochemistry 28, 6259 (1989). I l D. C. Straney and D. M. Crothers, Cell 43, 449 (1985). 12 j. E. Stefano and J. Gralla, Biochemistry 18, 1063 (1979). 13 R. J. White and D. R. Phillips, Biochemisny 28, 4277 (1989).

472

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

[23]

Procedure

1. To a microcentrifuge tube add 5 til of lacUV5 DNA fragment (approximately 50 nM final concentration), 3 til of 10x transcription buffer [Tc; 1 x Tc consists of 40 mM Tris-HC1 (pH 8.0), 100 mM KCI, 3 mM MgCI2, 0.1 mM EDTA], 1.5 #1 of bovine serum albumin (BSA, 3 mg/ml), 1.5 til of 200 mM DTT, 1.5 #1 of RNase inhibitor (~30 U/#I), and Milli-Q water to a total volume of 30 #1. 2. Add 0.5 #1 ofE. coli RNA polymerase (1 U/til), mix gently, and incubate for 15 rain at 37 °. The entire mixture up to this point is referred to as the transcription mix (TM). 3. Add 7.5 til of heparin (2 mg/ml) in 1 × Tc and incubate for 5 rain at 37 c . 4. Add 7.5 #1 of initiation mix (IM) and incubate for a further 5 rain at 37 °. The initiation mix comprises 1.2 mM GpA, 30 tiM GTP, 30 tiM ATE and [a-32p]UTP (2 tiCi/til; 2500 Ci/mmol) in 1× Tc. 5. Take a 5-til aliquot of the initiated complex and add to 5 #1 of termination/loading buffer on ice. Termination/loading buffer consists of 9 M urea, 10% (w/v) sucrose, 40 mM EDTA, 0.1% (w/v) xylene cyanol, 0.1% (w/v) bromphenol blue in 2× TBE buffer, pH 7.5. P r e p a r a t i o n of R N A - S e q u e n c i n g R e a c t i o n s Sequencing of the DNA template is achieved by adding chain terminating 3'O-methylnucleotides into RNA transcripts in the elongation phase of transcription. This allows the subsequent sequence assignment of transcripts that are terminated by DNA sequence-specific drug blokages to RNA polymerase. The methoxynucleotides provide a statistical probability of terminating the elongation phase of transcription, and yield C and G sequencing lanes analogous to dideoxy-terminated DNA-sequencing lanes. If sequencing lanes are required for RNA longer than approximately 150 nucleotides, the Y-methoxynucleotide : nucleotide ratio must be reduced to enable the RNA polymerase to be able to transcribe further along the DNA before transcription is terminated by incorporation of the methoxynucleotide into the nascent RNA. Dideoxy-CTP and dideoxy-GTP can be used as an alternative to 3'-methoxynucleotides to generate sequencing lanes. Procedure

1. Place two 5-#1 aliquots of the initiated transcript into separate Eppendorf tubes. 2. Add 2.5 til of 3'-methoxy-CTP/CTP (9 : 1) (referred to as MeC mix) to one and 3'-methoxy-GTP/GTP (9: 1) (MEG) to the other. The sequencing mixes are made up as 3 x mixes and can be made for any of the four ribonucleotides. For example, the MeC mix comprises 6 mM ATE 6 mM GTP, 6 mM UTP, 270 tiM 3'-methoxy-CTP, 30 tiM CTP, and 1.2 M KC1 in 1 x Tc.

[23]

In Vitro TRANSCRIPTIONASSAY

473

3. Incubate at 37° for 5 min, and then add an equal volume of termination/loading buffer to both samples. R e a c t i o n of I n i t i a t e d T r a n s c r i p t s w i t h D r u g The drug of interest is routinely added directly to the initiated transcription complex, and an outline of the overall procedure is shown in diagrammatic form in Fig. 3. However, an alternative method is to prereact the DNA with drug and then remove unreacted drug from the DNA before the above-described initiation steps (Fig. 4). This alternative method is preferred when covalently binding drugs react with other components of the transcription assay (e.g., alkylation of DNA by nitrogen mustards is inhibited by the transcription buffer itselfl4; cisplatin reacts with RNA polymerase) or when drugs interact with the unprotected promoter region, leading to a decrease in the RNA polymerase-promoter complex, t5 For both procedures subsaturating concentrations of drug are normally employed to ensure that most drug-DNA sites are not occupied (i.e., less than one lesion per fragment). This precaution results in a range of different drug sites being subsequently detected in the elongation phase of transcription. If high drug loadings were employed the first drug site would be completely occupied and RNA polymerase would not proceed past that site and therefore be unable to probe additional downstream drug sites. Natural pausing of RNA polymerase is minimized by the use of high levels of all four nucleotides during the elongation process, as well as by high ionic strength (0.4 M KC1). 5 The high level of nucleotides also ensures that additional incorporation of (x-32P label into the growing RNA chain is effectively eliminated, meaning that all transcripts, irrespective of length, contain the same amount of radiolabel. Plvcedure

1. Divide the remaining initiated transcription complex into two equal parts. In the flow chart shown in Fig. 3, this consists of two 15-/zl aliquots. 2. To one tube, add the drug of interest in a small volume (approximately 2 #1 in 1x Tc) and to the other half add the same volume of 1x Tc. 3. Take 5-#1 aliquots for studies of drug-DNA reaction time dependence and allow to react for appropriate times at 37 ° . 4. Add 2.5/zl of elongation mix (EM) to each of the drug- and non-drug-treated initiation mixtures and mix rapidly. Allow elongation to proceed for 5 rain at 37 ° and then add an equal volume of loading/termination buffer and place samples

14p. j. Gray,C. Cullinane, and D. R. Phillips, Biochemistry 30, 8036 (1991). 15S. M. Cutts, P. G. Parsons, R. A. Sturm, and D. R. Phillips, J. Biol. Chem. 271, 5422 (1996).

474

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

U

[23]

TM (30 I.tL)

15 rain f 7 . 5 gL heparin 5 rain ,--7.5 gL IM 5 rain

5 gL

5 gL

5 gL

15 gL

U

tl~.,,~ U~.,~ ~,~ MeG MeC 1xTc

(b

$

~

15 laL

,,- D,,,g"

~L2 MLdrug inl xTe "+Drug"

/ 5 ~L lh

5 ~L 4h

U

U

,"-2.5 , L

W

5'

5'

@

@

2.5 , L

5 ~L 8h

5 ~L lh

U

U

If 5'

2.5 , L

@

2.5 , L 5'

@

5 ~L 4h

5 ~L 8h

U

U

k"2"5 , L 5'

@

2.5 , L 5'

@

FIG. 3. Typical in vitro transcription protocol for drug-DNA reaction time dependence. After preparation of the transcription mix (TM), four types of samples are required. These are the initiation (I), sequencing (C and G), drug-treated (D1-D8), and control (non-drug-treated) samples (C1--C8). The reaction conditions parallel the procedures outlined in text (formation of synchronized initiation complexes and formation and sequencing of drug-induced blocked transcripts). The circled letters represent the end point of transcription for each of these samples, where an equal volume of termination/loading buffer is added and the samples are placed on ice before being denatured and subjected to electrophoresis.

[23]

In Vitro TRANSCRIPTIONASSAY

475

DRUG

I

•

IL RNA POLYMERASE A

A w

INITIATION NUCLEOTIDES A

,.7

i

A

ELONGATION NUCLEOT!DES

~ | . .

.

.

.

1 ....

FIG. 4, Alternative in vitro transcription assay for covalently binding drugs. An alternative to adding drug directly to the initiated transcription complex is to react DNA first with drug, and then to subject it to the initiation and elongation steps of transcription.

on ice. The elongation mix is made up as a 3 x solution and comprises a 6 mM concentration of each of the four ribonucleotides and 1.2 M KC! in 1x Tc. S e p a r a t i o n of B l o c k e d T r a n s c r i p t s The RNA transcripts are separated on high resolution sequencing gels. If the control lanes (lacking drug) show low levels of transcription or significant levels of background (i.e., resulting from pausing of the transcription complex prior to formation of full-length transcripts) the most likely causes are insufficient purity of the promoter-containing DNA fragment, or degradation of the DNA by the presence of single-strand or double-strand nicks, High-quality template DNA can usually be obtained by subjecting the DNA to an additional phenol--chloroform

476

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

[23]

extraction. If quality of DNA is not the problem, it is prudent to make fresh stock solutions of all reagents, as this has invariably proved quicker than trying to identify the individual contaminated component. Procedure

1. Prepare a conventional 12% (w/v) acrylamide denaturing sequencing gel (19 : 1 acrylamide : bisacrylamide, containing 7 M urea) in TBE buffer. 2. Subject the gel to preelectrophoresis for 1 hr to heat up to approximately 60 ° (typically 2000 V, approximately 100 W). 3. Denature all transcription samples (in termination/loading buffer) at 90 ° for 5 min, and then place on ice. 4. Load 4/zl of each sample onto the gel and continue electrophoresis until the bromphenol migrates approximately 75% of the length of the gel (approximately 2 hr). Fix the gel in 10% (v/v) glacial acetic acid-10% (v/v) methanol and then dry it. Q u a n t i t a t i o n of B l o c k e d T r a n s c r i p t s The relative amount of each length of RNA can be determined either by conventional autoradiography or by a phosphorimaging process. Both procedures are outlined below. The phosphorimaging process is preferable because it is faster, more sensitive, has a bigger dynamic range, and is fully computerized. An example of increasing transcriptional blockages induced by Adriamycin is shown in Fig. 5 at increasing times of reaction of the drug with DNA. 7 The relative site occupancy of Adriamycin (quantitated from the 16-hr lane of Fig. 5) is shown in Fig. 6. Autoradiography

Amersham (Arlington Heights, IL) Hyperfilm-g max X-ray film is routinely used for final quantitative work because of the low background absorbance and high contrast of this film. The time of exposure of the gel must be modified to ensure that photographic linearity is maintained. For Kodak (Rochester, NY) XAR X-ray film linearity is restricted to a 0-1 absorbance range. 16 Procedure

1. Place the dried gel in contact with Amersham Hyperfilm-g max or Kodak XAR-5 X-ray film overnight, without intensifying screens and at room temperature. 16j. C. Dabrowiak,A. Skorobogaty,N. Rich,C. E Vary,and J. N. Vournakis,Nucleic Acids Res. 14, 489 (1986).

In Vitro TRANSCRIPTION ASSAY

[23]

I

1

2

Control 4816243140481

477

Ad 2

4

8 162431 4 0 4 8 C

G

1 2

4

Ad+Fe(lll) 81624314048

FIG. 5. Blocked transcripts induced by Adriamycin (doxorubicin). The image shows the dependence of transcriptional blockages on the reaction time (3T ') of 10 #M Adriamycin (Ad) with the lacUV5containing DNA fragment in the absence and presence of 75/zM Fe(IlI), for reaction times of 1-48 hr. Control reactions in the absence of both Adriamycin and Fe(lII) ions were also left for the same reaction times. Lane I is the initiated complex prior to elongation. All other samples were subjected to elongation conditions for 5 min. Lanes C and G are sequencing lanes. The length of some of the major blocked transcripts is shown on the right-hand side of the autoradiogram. [Reprinted with permission from C. Cullinane and D. R. Phillips, Biochemistry 29, 5638 (1990). Copyright :~' 1990 American Chemical Society.[ 2. S c a n the a u t o r a d i o g r a m w i t h a d e n s i t o m e t e r (laser light s o u r c e to m a x i m i z e resolution), c o u p l e d to a n integrator. 3. S u m the total area ( p r o p o r t i o n a l to r a d i o a c t i v i t y ) in e a c h lane a n d e x p r e s s e a c h b a n d as a f r a c t i o n o f the total. T h i s yields the m o l e f r a c t i o n o f b l o c k e d t r a n s c r i p t s in e a c h r e a c t i o n m i x t u r e .

478

[23]

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

I I I I I

o I I m ! I I I ¢

m

I

~

~

I

'

i

'~

I

~--,~

"~ ~

~'.se~e.._ ~

~ " ~

=

I

o

'

.=~= ~.~

I

"~

0

•

~,~" "

=

~~

~ ~.~ ~.~.

~=z~ Z 0 l.Ij ~

~o ~

~ ~'~-. ,~ ~

.,~ ~'~

,

[23]

In Vitro TRANSCRIPTIONASSAY

479

Phosphorimaging The phosphorimaging process currently offers two major advantages over the photographic process: it is at least 250-fold more sensitive for the detection of 32p, and has a linear dynamic range at least 400 times greater than that of film processes. 17

Procedure l. Place the dried gel in contact with the phosphor plate for 1 hr (or up to overnight if necessary). 2, Scan the phosphor plate with a phosphorimaging system. 3. Normalize each transcript with respect to the total intensity in each lane to yield the mole fraction of each transcript in each reaction mixture. Relative O c c u p a n c y a n d D r u g D i s s o c i a t i o n K i n e t i c s A true dissociation rate constant can only be estimated from the first drug site encountered by the initiated transcription complex. All subsequent dissociation rate constants are distorted to some degree by readthrough of RNA polymerase from earlier (upstream) sites, and by the fact that all sites downstream of the first site are underestimated because less RNA polymerase reaches them compared with the first site. For these reasons, estimates of drug occupancy and dissociation kinetics are only good approximations (except for the first site) when the drug occupancy at each site is low. For a more rigorous approach a Monte Carlo simulation can be employed, is but a compartmental analysis yields identical results and is far simpler and quicker. 19 The mole fraction of RNA at each drug site is an indication of the relative occupancy of drug at each site. However, a true correlation between these two parameters exists only at infinite dilution of the drug. In practice this means using the lowest drug level possible to detect blockages, so that under these conditions most drug sites will not be occupied. Therefore, the majority of RNA polymerases in the initiated transcription complex will elongate fully to yield a fulllength transcript. Decay of transcriptional blockages can be quantitated to reveal the time-dependent loss of drug-DNA adducts at individual sites. Figure 7 shows the loss of Adriamycin-induced blockage sites with increasing elongation time, together with the increase in full-length transcript. 2° The first-order kinetic analysis of the decay of two Adriamycin blockage sites (Fig. 7B) shows that Adriamycin adducts are more stable at GpC sites (37-mer) compared to an isolated guanine residue (61-mer). 17 R. E Johnston, S. C. Pickett, and D. L. Barker, Electrophoresis l l , 355 (1990). E8D. R. Phillips, R. J. White, D. Dean, and D. M. Crothers, Biochemistry 29, 4812 (1990). t9 D. R. Phillips, P. J. Moate, and R. C. Boston, Anti-Cancer Drug Des. 9, 209 (1994). 2o A. van Rosmalen, C. Cullinane, S. M. Cutts, and D. R. Phillips, Nucleic Acids Res. 23, 42 (1995).

480

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[23]

A C

G

1

2

3

4

5

6

7

8

9 10

82

§0

37

FIG. 7. Elongation of the transcription complex past Adriamycin-induced blockage sites. (A) Adriamycin (10/zM) was reacted with the 497-bp fragment containing the lacUV5 promoter. Subsequent elongation of the initiated transcription complex was allowed to proceed for 1, 15, or 30 min or for 1, 2, 4, 6, 12, 24, or 48 hr (lanes 1-10). C and G denote sequencing lanes. The numbering represents the

[23]

In Vitro TRANSCRIPTIONASSAY

481

12

B

10 _=

I

10

r

20

i

30

I

I

40

i

50

Time (h) FIG. 7. (continued)

Procedure

1. Form initiated transcription complexes as outlined above. 2. React the initiated complexes with a range of drug concentrations (typically 0.1-100 # M for a preliminary study) for the required time, as outlined above. Once the initiated transcript has been formed it is possible to reduce the temperature to as low as 4 ° to study DNA interactions of drugs with fast dissociation kinetics. 3. Measure blocked transcripts as outlined above. 4. Determine the drug concentration that yields approximately 90% full-length transcripts. 5. Repeat steps 1-3 at this drug concentration and vary elongation times at 37 ° (or the selected lower temperature). When the elongation mix is added to initiated complexes, mix rapidly because this is zero time for subsequent kinetic analysis. 6. Plot ln[RNA] (where [RNA] is the mole fraction of blocked transcript) against elongation time for the first few drug-induced blockages. The slope of these plots, if linear, yields the rate constant for dissociation of drug from each site. 7'2° The non-linear kinetics for the dissociation of the 37-mer (Fig. 7B) can be resolved as two rates of loss at this site with half-lives of 3 hr and 40 hr. length of the RNA transcript, beginning with G of the initiating GpA dinucleotide. (B) The amount of RNA transcript at the 37-mer (i) and 6 l-mer ([]) blockage sites was quantitated by phosphorimager analysis and subjected to first-order kinetic analysis. ]Reprinted with permission from A. van Rosmalen, C. Cullinane, S. M. Cutts, and D. R. Phillips, Nucleic Acids Res. 23, 42 (1995). Copyright © 1995 Oxford University Press.]

482

C H E M I C A AND L MOLECULAR BIOLOGICAL APPROACHES

Bidirectional Transcription

[23]

Footprinting

The in v#ro transcription assay described above has been extremely successful in detecting the 5' end of drug-induced blockage sites on D N A but does not indicate the physical size of the blocking unit. To obtain this information, the blockage can be probed by R N A polymerase from both directions. This assay, in which two counterdirected promoters are employed (see Fig. 8), is referred to as bidirectional transcription footprinting.l° If the kinetics of readthrough past occupied drug sites

A

q [~N25

B

"

,,

.P:

0¢ INITIATE

C

A

DRUG

D

.~

I

I' ....... ELONGATE

E A l l

| A

FIG. 8. Schematic representation of bidirectional transcription footprinting. The DNA fragment containing the counterdirected UV5 and N25 promoters is shown in (A). Deactivationof either one of the promoters yields (B) and addition of E. coli RNA polymerase and initiation nucleotides yields the initiated transcription complex (C). Reaction with drug yields (D) and subsequent elongation results in a range of drug-inducedblocked transcripts (E). The two sets of blocked transcripts are summarized together in (F) to reveal bidirectionaltranscription footprints of drug sites.

[23]

In Vitro TRANSCRIPTIONASSAY

483

are not of interest, then any counterdirected commercial promoter system could be employed (e.g., SP6/T7 or T3/T7, both of which are available from a variety of suppliers, together with the SP6, T3, and T7 RNA polymerases) and are ideally suited to the detection of permanent lesions in any defined DNA. These systems enable insertion of any DNA sequence at the multiple cloning site between the promoters, and are particularly useful because the bacteriophage polymerases are sensitive to the presence of lesions and adducts. 13 An example of the bidirectional footprint of Adriamycin adducts is shown in Fig. 9,7 using the counterdirected bacterial promoters UV5 and N25. Procedure

1. Restriction digest pRW2t° with PvulI and XhoI and isolate the 315-bp fragment containing counterdirected UV5 and N25 promoters. 2. Restriction digest the 315-bp fragment with DraI, to deactivate the N25 promoter. 3. Initiate the UV5 promoter with an initiation mix (6x) comprising 1.2 mM GpA, 30/zM GTP, 30/zM ATP, and [u-32p]UTP (2/zCi/#l) in 1× Tc. 4. React the initiated UV5 fragment with drug and elongate and quantitate blocked transcripts as outlined previously. 5. Digest the 315-bp fragment with BstNI to deactivate the UV5 promoter. (Note: Steps 5-7 should be carried out in parallel with steps 2-4.) 6. Initiate the N25 promoter with an initiation mix (6×) comprising 1.2 mM ApU, 30 #M GTP, 30 #M ATE and [ot-32p]UTP (2 #Ci/#l) in 1x Tc. 7. React the initiated N25 promoter fragment with drug and elongate and quantitate blocked transcripts as outlined above. 8. Correlate the mole fraction of blocked transcripts from both promoters and plot as a histogram to reveal bidirectional transcription footprints. 7"10 The lacUV5 transcription assay has been used in our own laboratory to identify the sequence specificity of a number of reversibly binding drugs (actinomycin D, Adriamycin, daunomycin, echinomycin, mithramycin, mitoxantrone, and nogalomycin)21 as well as drug-induced DNA adducts (Adriamycin, barminomycin, cyanomorpholino-Adriamycin, nitrogen mustards, cisplatin, mitomycin C). 21'22 Transcription assays employing bacteriophage and mammalian RNA polymerases have also been used successfully as probes of sequence-specific drugDNA interactions. The bacterial E. coli RNA polymerase has been used in other laboratories as a probe for the detection of DNA binding by actinomycin D 23 and 21 D. R. Phillips, in "Advances in DNA Sequence Specific Agents" (J. B. Chaires and L. H. Hurley, eds.), p. 101. JAI Press, Greenwich, Connecticut, 1996. 22 L. C. Perrin, C. Cullinane, K. Kimura, and D. R. Phillips, Nucleic Acids Res. 27, 1781 (1999). 23 V. A. Aivasashvilli and R. S. Beabealashvilli, FEBS Lett. 160, 124 (1983).

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

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psoralen adducts. 24,25 However, the bacteriophage RNA polymerases T7 and SP6 have been used widely to detect the sites of interaction of nitrogen mustards, 26,27 cisplatin, 2s-3° nitrosoureas, 3° benzo[a]pyrene, 31 benzodiazepines, 32 and several intercalating agents. 13,21 Although RNA polymerase II-based assays have been developed, the complexity of such systems limits their use to probing the effects of DNA-binding drugs on transcription initiation, pausing, and formation of full-length transcripts. 33-38 24 y. B. Shi, H. Gamper, and J. E. Hearst, Nucleic Acids Res. 15, 6843 (1987). 25 y. B. Shi, H. Gamper, and J. E. Hearst, J. Biol. Chem. 263, 527 (1988). 26 R. O. Pieper, B. W. Futscher, and L. C. Erickson, Carcinogenesis 10, 1307 (1989). 27 R. O. Pieper and L. C. Erickson, Carcinogenesis It, 1739 (1990). z8 M. A. Lemaire, A. Schwartz, A. R. Rahmouni, and M. Leng, Proc. Natl. Acad. Sci. U.S.A. 88, 1982 (1991). 29 M. Decoville, A. Schwartz, D. Locker, and M. Leng, FEBS Lett. 323, 55 (1993). 3o R. O. Pieper, S. L. Noftz, and L. C. Erickson, Mol. Pharmacol. 47, 290 (1995). 31 B. D. Thrall, D. B. Mann, M. J. Smerdon, and D. L. Springer, Carcinogenesis 13, 1529 (1992). 32 M. S. Puvvada, S. A. Forrow, J. A. Hartley, P. Stephenson, I. Gibson, T. Jenkins, and D. E. Thurston, Biochemistry 36, 2478 (1997). 33 y. Corda, M.-E Anin, M. Leng, and D. Job, Biochemistry 31, 1904 (1992). 34 y. Corda, C. Job, M.-E Anin, M. Leng, and D. Job, Biochemistry 32, 8582 (1993). 35 j. Mote, P. Ghanouni, and D. Reines, J. Biol. Chem. 236, 725 (1994). 36 B. A. Donahue, S. Yin, J.-S. Taylor, D. Reines, and P. C. Hanawalt, Proc. Natl. Acad. Sci. U.S.A. 91, 8502 (1994). 37 B. A. Donahue, R. P. Fuchs, D. Reines, and P. C. Hanawalt, J. Biol. Chem. 271, 10588 (1996). 38 C. Cullinane, S. J. Mazur, J. M. Essigmann, D. R. Phillips, and V. A. Bohr, Biochemistry 38, 6204 (1999).

[241 Use of DNA Molecules Substituted with Unnatural Nucleotides to Probe Specific Drug-DNA Interactions B y CHRISTIAN BAILLY a n d MICHAEL J. WARING

Introduction DNase I footprinting has long been used for qualitative, and more recently quantitative, analyses of drug-DNA interactions. DNA restriction fragments of 100-300 base pairs containing diverse arrangements of A - T and G. C base pairs are usually employed for these experiments. Specific target sequences can be inserted into the substrate DNA but up until the 1990s it was not fully realized that the DNA can be completely designed to contain unnatural nucleotides. In 1992, a relatively long DNA fragment was engineered to have one of its bases replaced with

METHODSIN ENZYMOLOGY.VOL.340

Copyright© 2001 by AcademicPress All rightsof reproductionin anyformreserved. 0076-6879/00$35.00

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DNA CONTAININGMODIFIEDBASES

485

psoralen adducts. 24,25 However, the bacteriophage RNA polymerases T7 and SP6 have been used widely to detect the sites of interaction of nitrogen mustards, 26,27 cisplatin, 2s-3° nitrosoureas, 3° benzo[a]pyrene, 31 benzodiazepines, 32 and several intercalating agents. 13,21 Although RNA polymerase II-based assays have been developed, the complexity of such systems limits their use to probing the effects of DNA-binding drugs on transcription initiation, pausing, and formation of full-length transcripts. 33-38 24 y. B. Shi, H. Gamper, and J. E. Hearst, Nucleic Acids Res. 15, 6843 (1987). 25 y. B. Shi, H. Gamper, and J. E. Hearst, J. Biol. Chem. 263, 527 (1988). 26 R. O. Pieper, B. W. Futscher, and L. C. Erickson, Carcinogenesis 10, 1307 (1989). 27 R. O. Pieper and L. C. Erickson, Carcinogenesis It, 1739 (1990). z8 M. A. Lemaire, A. Schwartz, A. R. Rahmouni, and M. Leng, Proc. Natl. Acad. Sci. U.S.A. 88, 1982 (1991). 29 M. Decoville, A. Schwartz, D. Locker, and M. Leng, FEBS Lett. 323, 55 (1993). 3o R. O. Pieper, S. L. Noftz, and L. C. Erickson, Mol. Pharmacol. 47, 290 (1995). 31 B. D. Thrall, D. B. Mann, M. J. Smerdon, and D. L. Springer, Carcinogenesis 13, 1529 (1992). 32 M. S. Puvvada, S. A. Forrow, J. A. Hartley, P. Stephenson, I. Gibson, T. Jenkins, and D. E. Thurston, Biochemistry 36, 2478 (1997). 33 y. Corda, M.-E Anin, M. Leng, and D. Job, Biochemistry 31, 1904 (1992). 34 y. Corda, C. Job, M.-E Anin, M. Leng, and D. Job, Biochemistry 32, 8582 (1993). 35 j. Mote, P. Ghanouni, and D. Reines, J. Biol. Chem. 236, 725 (1994). 36 B. A. Donahue, S. Yin, J.-S. Taylor, D. Reines, and P. C. Hanawalt, Proc. Natl. Acad. Sci. U.S.A. 91, 8502 (1994). 37 B. A. Donahue, R. P. Fuchs, D. Reines, and P. C. Hanawalt, J. Biol. Chem. 271, 10588 (1996). 38 C. Cullinane, S. J. Mazur, J. M. Essigmann, D. R. Phillips, and V. A. Bohr, Biochemistry 38, 6204 (1999).

[241 Use of DNA Molecules Substituted with Unnatural Nucleotides to Probe Specific Drug-DNA Interactions B y CHRISTIAN BAILLY a n d MICHAEL J. WARING

Introduction DNase I footprinting has long been used for qualitative, and more recently quantitative, analyses of drug-DNA interactions. DNA restriction fragments of 100-300 base pairs containing diverse arrangements of A - T and G. C base pairs are usually employed for these experiments. Specific target sequences can be inserted into the substrate DNA but up until the 1990s it was not fully realized that the DNA can be completely designed to contain unnatural nucleotides. In 1992, a relatively long DNA fragment was engineered to have one of its bases replaced with

METHODSIN ENZYMOLOGY.VOL.340

Copyright© 2001 by AcademicPress All rightsof reproductionin anyformreserved. 0076-6879/00$35.00

486

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[24]

a nonconventional base--specifically, with inosine residues substituted for all the guanosines in a 160-base pair DNA fragment made by the polymerase chain reaction (PCR).I Initially the enzymatic procedure was long and laborious, requiring more than 50 PCR tubes to obtain a tiny amount of the modified DNA, just enough for one or two footprinting experiments. However, this landmark experiment opened up a completely new avenue for the study of sequence-specific drug binding to DNA and rapidly led to optimization of procedures for synthesizing DNA containing unnatural bases, so that the methodology is now available to probe the interaction of DNA with a large diversity of small molecules and proteins. The strategy has become a cornerstone of pharmacology in the field of drug-DNA interactions. Background

DNA Helix: A Double Ribbon Coated with Exocyclic Groups The double-helical structure of DNA delimits two channels that provide distinct routes of access to the genetic information encoded in its sequence of bases (Fig. 1). The major groove is not only more accessible than the minor groove but also displays a characteristic pattern of hydrogen bond donors and acceptors that enable the four base pairs to be distinguished. The edges of the A. T and G. C base pairs have quite different hydrogen-bonding potentials and can therefore engage in distinct directional interactions with a ligand having hydrogen-bonding complementarity. Thus, a G. C base pair seen from the major groove has a pattern of acceptor-acceptor-donor that is distinct from that for the reverse C. G base pair. In contrast, A. T and T. A pairs exhibit identical major groove acceptor/donor patterns but can be distinguished by the presence of the exocyclic methyl group of thymine, which introduces an asymmetry into the groove. Viewed from the minor groove, the hydrogen-bonding capabilities of the base pairs afford much less opportunity for discrimination: the only available hydrogen bond donor group is the 2-amino group of guanine, which occupies a large space and must impede the access of ligands, proteins, and drugs to the minor groove surface o f G . C and C. G base pairs. In fact, the C2 exocyclic amino group of guanine, which protrudes out from the stack of base pairs, is the major chemical difference in the minor groove between A. T and G. C base pairs. This substituent plays a significant role in the control and maintenance of DNA architecture. In B-DNA, on average the width of the minor groove is about 6 A and its depth is 8.2 ~. The depth of the major groove is not much different (8.5 ]~) but its width is considerably greater, about 11.6 ~.2 The local width of the minor groove of the DNA helix correlates to a first approximation with its base composition because A-T tracts appear to be associated with a narrowed groove, whereas G-C tracts have a widened minor groove. I C. Marchand, C. Bailly, M. J. McLean, S. E. Moroney, and M. J. Waring, Nucleic AcMs Res. 20, 5601 (1992). 2 S. Neidle, "DNA Structure and Recognition." IRL Press, London, 1994.

[24]

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488

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

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DNA Bases: A, T, G, C, and All the Others The common idea that DNA is composed of only four bases, A, T, G, and C, is an oversimplification because most types of DNA, including human DNA, contain naturally modified forms of the canonical bases. 5-Methylcytosine (5-MeC or M; Fig. 1) was the first modified nucleobase to be discovered more than half a century ago. 3 5-MeC completely replaces C in the DNA from Xanthomonas phage XP-12. 4 Nowadays, it is well established that this unusual but naturally occurring base participates in the control of gene expression in higher organisms. 5 Over the last 50 years many other modified bases have been discovered, particularly in DNA from microorganisms. DNA from the bacteriophage SP-15, which infects Bacillus subtilis, contains a modified pyrimidine [5-(4',5'-dihydroxypentyl)uracil] that replaces 62% of the thymines, and other types of B. subtilis phages have T residues replaced by uracil (U) or 5-bydroxymethyluracil (hm5U). 6'7 Just as T residues are found in tRNA, U residues can be a normal constituent of DNA. It is quite common among diverse types of bacteriophages that one of the normal pyrimidines of DNA is completely or partially replaced by a modified derivative. 6 But the modified base can also be a purine. One of the most interesting is 2-aminoadenine (abbreviated DAP or D for 2,6-diaminopurine; Fig. 1) that replaces adenine in the cyanophage S-2L, which infects the blue-green algae Synechococcus elongatus. 8 Thus, DAP in DNA is compatible with normal DNA function although the DAP. T base pair possesses an extra hydrogen bond compared with A- T because of the additional -NH2 group that hydrogen bonds to the 2-keto of thymine in the minor groove of the double helix (Fig. 1). By contrast, the inosine nucleoside (I) lacks the -NH2 group of guanosine and therefore I. C base pairs can form only two hydrogen bonds instead of the three typical of G. C base pairs. Consequently D and I have become favorite tools with which to examine the critical role of the exocyclic 2-amino group of guanine in DNA recognition by various antibiotics. DNA C o n t a i n i n g I n o s i n e a n d / o r D i a m i n o p u r i n e R e s i d u e s Synthesis The phosphoramidite forms of the I and DAP nucleosides are available by total synthesis and can readily be incorporated into synthetic oligonucleotide 3 R. D. Hotchkiss, J. Biol. Chem. 168, 315 (1948). 4 M. Ehrlich, K. Ehrlich, and J. A. Mayo, Biochim. Biophys. Acta 395, 109 (1975). 5 X. Cheng, Annu. Rev. Biophys. Biomol. Struct. 24, 293 (1995). 6 M. Ehrlich and X.-Y. Zhang, in "Chromatography and Modification of Nucleosides" (C. W. Gehrke and K. C. T. Kuo, eds.). Elsevier, Amsterdam, 1990. 7 M. Ehrlich and K. C. Ehrlich, J. Biol. Chem. 256, 9966 (1981). s I. Y. Khudyakov, M. D. Kimos, N. I. Alexandrushkina, and B. E Vanyushin, Virology 88, 8 ( 1978).

[24]

DNA CONTAININGMODIFIEDBASES

489

sequences. 9 There are numerous instances in which these bases have been used to investigate nucleic acid stability, structure, or interaction with ligands. DAP is frequently introduced into DNA to increase its melting temperature ~° and/or into RNA duplexes It as well as for related applications such as making primers for sequencing and PCR 12'13 or fingerprinting.14 Inosine is also commonly used to probe unusual DNA structures such as parallel-stranded DNA 15 or intrinsically curved sequences, 16 and to evidence minor groove binding to DNA of certain proteins such as high mobility group (HMG) proteins (see below) or the tyrosine kinase c-Abl.17 The potential uses of I and DAP are wide ranging.18 Alternatively, I and DAP can be incorporated into DNA by enzymatic methods via the use of the corresponding triphosphate and polymerases. The triphosphate of 2-amino-adenosine acts as a true analog of ATP in transcription. 19 dITP has long been commercially available. By contrast, dDTP was not manufactured until more recently. It is currently sold by TriLink Biotechnologies (San Diego, CA). Methods have been described to convert 2,6-diaminopurine-2'-deoxyribonucleoside into dDTE The synthesis involves converting the nucleoside to its 5'-monophosphate derivative dDMP, followed by pyrophosphorylation. 2°m Other chemical routes have been reported. 22'23 Incorporation of nucleoside triphosphate analogs by polymerases is a useful method of examining miscoding by different DNA polymerases, dDTP and to a lesser extent dITP are good substrates for a number of polymerases, including heat-stable enzymes such as Taq polymerase. We have established a protocol to incorporate inosine and]or diaminopurine into both strands of a 160-mer fragment containing the Escherichia coli tyrT promoter, 24 which offers a great variety of 9 B. S. Sproat, A. M. Iribarren, R. G. Garcia, and B. Beijer, Nucleic Acids Res. 19, 733 (1991). l0 j. D. Hoheisel and H. Lehrach, FEBS Lett. 274, 103 (1990). I1 G. M. Lamm, B. J. Blencowe, B. S. Sproat, A. M. Iribarren, U. Ryder, and A. I. Lamond, Nucleic AcidsRes. 19, 3193 (1991). 12 T. L. Azhykina, S. I. Veselovskaya, V. A. Myasnikov, V. K. Potapov, and E. D. Sverdlov, Proc. Natl. Acad. Sci. Russia 330, 624 (1993). 13 y. Lebedev, N. Akopyants, T. Azhikina, Y. Shevchenko, V. Potapov, D. Stecenko, D. Berg, and E. Sverdlov, Genet. Anal. 13, 15 (1996). 14 M. 1. Prosnyak, S. 1. Veselovskaya, V. A. Myasnikov, E. J. Efremova, V. K. Potapov, S. A. Limborska, and E. D. Sverdlov, Genomics 21, 490 (1994). 15 S. Mohammadi, R. Klement, A. K. Shchyolkina, J. Liquier, T. M. Jovin, and E. Taillandier, Biochemistry 37, 16529 (1998). 16 N. E. M~llegaard, C. Bailly, M. J. Waring, and E E. Nielsen, Nucleic Acids Res. 25, 3497 (1997). 17 M. H. Dav id-Cordonnier, M. Hamdane, C. B ailly, and J. C. D'Halluin, Biochemistry 37, 6065 (1998). 18 C. Bailly and M. J. Waring, NuHeic Acids Res. 26, 4309 (1998). 19 H. R. Rackwitz and K. H. Scheit, Eulz J. Biochem. 72, 191 (1977). 20 N. N. Kahn, G. E. Wright, L. W. Dudycz, and N. C. Brown, Nucleic Acids Res. 13, 6331 (1985). 21 A. Chollet and E. Kawashima, Nucleic Acids Res. 16, 305 (1988). 22 C. A. Brennan and R. I. Gumport, Nucleic Acids Res. 13, 8665 (1985). 23 L. W. McLaughlin, T. Leong, E Benseler, and N. Piel, Nucleic Acids Res. 16, 5631 (1988).

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binding sites and has been extensively employed over the last 20 years in footprinting experiments with numerous drugs and proteins. 25 The four DNA molecules containing either natural bases or inosine residues in place of guanosines (G ~ I substitution), or 2,6-diaminopurine residues in place of adenines (A ~ DAP substitution), or both I and DAP residues, can be synthesized by PCR amplification, using the parameters described in Table I. The sequence of the tyrT fragment and the two primers (Watson primer 1 and Crick primer 1) used for the PCR are specified in Fig. 2. For the sake of convenience, the two strands of the duplex are named the Watson (Fig. 2, top, coding) and Crick (Fig. 2, bottom, noncoding) strand. The use of primers in which the 5' terminal nucleotide bears a 5'-OH or a 5'-NH2 terminus is convenient as it enables selective labeling of one or the other strand in the PCR product. The most straightforward procedure is to initiate the PCR amplification with primers already radiolabeled, but the use of unlabeled primers has the advantage that quantities of the modified DNA may be stored for subsequent use. DAP-containing DNA is routinely obtained in good yield, but making the inosinesubstituted DNA can be more tricky. We found that commercial stock solutions of dITP are not stable and should be replaced frequently (every month). Moreover, because of the limited thermal stability of DNA containing inosine residues in both strands the yield of the reaction can vary substantially from one experiment to another, mostly depending on the freshness (purity) of the dITP preparation. This DNA is by far the least stable of the four species, and is best used promptly. The protocol can be adapted to engineer DNA molecules containing the modified base(s) on one strand only of the duplex. Molecules asymmetrically substituted with inosine and/or 2,6-diaminopurine are best made with different PCR primers that produce complementary strands of different length. 26 For example, to synthesize a duplex DNA containing DAP residues on the radiolabeled Watson strand only, a PCR is set up with the tyrT template, the Watson primer 1 and Crick primer 2, and dDTP plus dTTP, dCTP, and dGTP (Fig. 3). This PCR produces a 162-base pair duplex with DAP residues on both strands. In parallel, a PCR is set up with the four natural nucleotides, using Watson primer 2 and Crick primer 1. It produces a normal duplex of 155 base pairs. Both PCR products are then "polished" by treatment with reverse transcriptase to eliminate any incomplete strands (which could subsequently become labeled) by converting them to DNA molecules of homogeneous length. The normal and DAP-containing duplexes are mixed, denatured by heating, and slowly cooled to allow reannealing of the complementary single strands. The products comprise the parental PCR species plus hybrid molecules containing strands originating from the two different PCRs, but only one duplex can be radiolabeled in the presence of [o~-32p]dCTPand avian myetoblastosis virus 24 C. Bailly and M. J. Waring, Nucleic Acids Res. 23, 885 (1995). 25 M. J. Waring and C. Bailly, J. Mol. Recognit. 7, 109 (1994). 26 S. Jennewein and M. J. Waring, Nucleic Acids Res. 25, 1502 (1997).

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TABLE I POLYMERASE CHAINREACTIONCONDITIONSUSED TO INCORPORATEINOSINE AND/OR 2,6-DIAMINOPURINERESIDUESINTOtyrT DNA" Amplification cycles (20--30) Denaturation Primer annealing Polymerization

Normal and DAP DNA

Inosine and I + DAP DNA

1 min at 94 ° 2 min at 45 ° 10 min at 72 °

1 min at 84 ° 2 min at 45 ° 10 min at 62 °

a In all cases, PCR mixtures contained l0 ng of the DNA template, a 1 #M concentration of each primer, a 250/~M concentration of each appropriate dNTP (dTTP, dCTP, plus dATP or dDTP, and dGTP or dlTP according to the desired DNA), and 5 units of Taq polymerase in a volume of 50 #1 containing 50 mM KC1, 10 mM TrisHCI (pH 8.3), 0.1% (v/v) Triton X-100, and 1.5 mM MgC12. To prevent unwanted primer-template annealing before the cycles began, the reactions were heated to 60 ° before adding the Taq polymerase. After the last cycle, the extension segment was continued for an additional 10 min at 72 ° followed by a 5-min segment at 55 ° and a 5-min segment at 37 °. The purpose of these final segments was to maximize annealing of full-length product and to minimize annealing of unused primer to full-length product. 24

(AMV) reverse transcriptase. This is the hybrid duplex bearing a free 3'-hydroxyl group with a single-stranded template 5' overhang that contains a natural Crick strand and an artificial Watson strand, as schematized in Fig. 3. Finally, purification is easy because of the difference in length between the hybrid and either of the

5"5"5'3'-

GTTACCT AATT CCGGTTACCT AATT CCGGTTACCT 0 i0 GGCCAATGGA

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40 TAATGCGTTG

CAGTTCATTT 50 GTCAAGTAAA

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AACACTTTAC

AAGAGTTGCA

TTGTGAAATG

TCGCCGCGCA

CATTTGATAT 90 GTAAACTATA

GATGCGCCCC i00 CTACGCGGGG

GCTTCCCGAT ii0 CGAAGGGCTA

AAGGGAGCAG

60

GCCAGTAAAA 130 CGGTCATTTT

AGCGGCGCGT

70

AGCATTACCC CGTGGTGGGG 140 150 TCGTAATGGG GCACCACCCC (Crick primer 1) 3 ' - G G G C A C C A C C C C

FIG. 2. Sequences of the

80

120 TTCCCTCGTC

GTTC-3'

158 CAAGGGCT -5' CAAGGGCT -5'

tyrT (A93) DNA and the PCR primers used to prepare the modified

DNA.

492

CHEMICAL AND MOLECULAR

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TABLE II MELTING TEMPERATURES FOR NATURAL AND ARTIFICIALtyrT DNA FRAGMENTSa Fragment Normal DNA DAP DNA Inosine DNA I -I- DAP DNA

Tm value (°C) 62.7, 65.9, 42.4, 49.5,

70.0, 72.8 77.4, 78.6 43.8 51.2

~' Tm measurements were performed in BPE buffer, pH 7.1 (6 mM Na2HPO4, 2 mM NaH2PO4, 1 mM EDTA), at 260 nm with a heating rate of 0.2°/rain. 3°

parental molecules, allowing separation by electrophoresis on a neutral acrylamide gel. The same procedure can be adapted to produce DNA molecules asymmetrically substituted with inosine or with inosine + diaminopurine residues. 26 Physical and Structural Studies It is well known that G ---> I substitution significantly reduces the thermal stability of DNA because of the loss of a hydrogen bond. By contrast, incorporation of DAP into short DNA oligomers increases the thermal stability of the duplex by 0-2 ° per D. T base pair. 2v The melting temperature (Tin) of a given oligo- or polynucleotide containing Y% of DAP increases by a factor of 0.14 x Y (ATm).14 However, the dependence of the Tm on the diaminopurine content is not linear. 2s,29 Thermal denaturation experiments performed with tyrT DNA containing artificial bases have provided additional information. Unexpectedly, the Tm analysis of the unsubstituted normal DNA revealed three discrete transitions (Table II), probably reflecting the heterogeneity of the sequence, which contains mixed AT/GC stretches as well as several tracts of contiguous purely A. T or purely G. C base pairs. With the fully DAP-substituted DNA we also measured three Tm values, each 3-7 ° higher than those obtained with the natural DNA fragment (Table 1I).3o The Tm elevation resulting from introduction of a 2-amino group on to A residues is much smaller in the deoxy series than in the ribo series.31In the deoxyribo series the stabilizing contribution arising from the formation of a third hydrogen bond in D. T pairs is opposed by a destabilization due to disruption of the spine of hydration in the minor groove of B-form DNA. 32 As one would anticipate, the Tm values 27 S. Gryaznov and R. G. Schultz, Tetrahedmn Lett. 35, 2489 (1994). 28 M. Muraoka, H. T. Miles, and E B. Howard, Biochemistry 19, 2429 (1980). 29 j. Sagi, E. Szakonyi, M. Vorlickova, and J. Kypr, J. Biomol. Struct. Dyn. 13, 1035 (1996). 3o C. Bailly, S. Dongchul, M. J. Waring, and J. B. Chaires, Biochemistry 37, 1033 (1998). 31 E B. Howard and H. T. Miles, Biochemistry 22, 597 (1983). 32 E B. Howard and H. T. Miles, Biochemistry 23, 6723 (1984).

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CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

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measured for the inosine DNA are considerably lower than those of normal DNA. The destabilizing effect of inosine is pronounced and surely accounts for some, at least, of the technical difficulty encountered during PCR synthesis. Only two transitions were measured with both the I-containing DNA and I + DAP-containing DNA (Table II). Substitution with both DAP and inosine produces a molecule that is significantly less stable than the natural molecule, although complementary absorption and circular dichroism measurements revealed that all four DNA species, with modified or unmodified bases, remain in the duplex from with little or no alteration of the global secondary B-type conformation. 3° However, the incorporation of I or D residues into DNA does induce significant local variation of the structure, particularly at runs of A- T base pairs. It affects the electrophoretic mobility in nondenaturing polyacrylamide gels; inosine-substituted DNA migrates more slowly than normal DNA whereas DAP-substituted DNA migrates more rapidly. The A --+ DAP substitution has a marginally greater influence than the G --+ I substitution. These results can be interpreted on the basis that removal of the guanine 2-amino group makes the former G. C base pairs, now I. C, behave like A. T base pairs, adopting a high propeller twist 33 and permitting short homopolymeric (dI). (dC) tracts to confer intrinsic curvature on the DNA fragment, which consequently migrates more slowly. 34 Conversely, substitution of DAP for adenine flattens the 2-amino-A. T pairs, renders them more rigid, and abolishes the intrinsic curvature conferred by short (dA). (dT) tracts. Studies with the substituted tyrT DNA species have validated previous work suggesting that the exocyclic 2-amino group plays a significant role in the sequence conformational microheterogeneity of the DNA helix. 33,34 Photocleavage of substituted DNA by uranyl nitrate at acidic pH is modulated quite differently from natural DNA, consistent with a marked narrowing of the minor groove at sites of G --~ I substitution and widening at sites of A --+ DAP replacement. The latter exerts the dominant effect. Changes in groove width are equally evident in the patterns of susceptibility to DNase I cleavageY

Protein-DNA Recognition PCR products substituted with I and/or DAP residues have proved useful for investigating sequence recognition by proteins. A major groove-binding protein, the E. coli factor for inversion stimulation (FIS), markedly bends the double helix. 36 Band-shift assays of FIS protein binding revealed facilitated interaction 33 S. Diekmann, J. M. Mazzarelli, L. W. McLaughlin, E. von Kitzing, and A. A. Travers, J. Mol. Biol. 225, 729 (1992). 34 H.-S. Koo and D. M. Crothers, Biochemistry 26, 3745 (1987). 3.s C. Bailly, N. E. M¢llegaard, P. E. Nielsen, and M. J. Waring, EMBO J. 14, 2121 (1995). 36 S. E. Finkel and R. C. Johnson, Mol. Microbiol. 6, 3257 (1992).

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DNA CONTAININGMODIFIEDBASES

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with inosine-containing DNA and reduced binding to DAP-containing DNA, attributable to altered bendability. DNase I footprinting experiments confirmed that fewer sites would bind FIS in DAP-containing DNA at a given protein concentration, whereas higher levels of binding occurred with inosine-containing molecules. It was concluded that base substitutions that affect the placement of the purine 2-amino group in the minor groove can affect both the intrinsic curvature and the bendability of DNA. 37 The minor groove-binding protein HMG-D, one of the Drosophila melanogaster counterparts of the abundant chromosomal protein HMG-1, 38 also bends DNA, and again DAP-containing DNA is a poor substrate for the protein whereas HMG-D binds strongly to the inosine-containing DNA. With the doubly substituted DNA, the positive effect of the G --~ I substitution is counterbalanced by the negative effect of the A -+ DAP substitution. 39 In this case the substitution of DAP for A appears not only to reduce the initial affinity of HMG-D for the probe DNA but also the cooperativity of binding. The reduction in affinity could be a simple consequence of partial steric exclusion consequent on the presence of the additional 2-amino groups whereas the loss of cooperativity could be related to a reduction in the flexibility of DNA as well. Drug-DNA Recognition Because most small molecules bind to DNA via the minor groove of the double helix, it has long been suspected that the exocyclic 2-amino group of guanine plays a key role in determining their preferred binding sites. 4° PCR-generated DNA molecules containing inosine and/or diaminopurine furnish a unique means to verify this notion by simply footprinting the ligands on the substituted DNA molecules as substrates. With few exceptions,41 the footprints of all drugs and antibiotics are radically altered by the G --~ I and A --~ DAP substitution. For example, the intercalative transcription inhibitor actinomycin42 normally binds to GpC steps in DNA but interacts predominantly with DpT steps in the doubly substituted I + DAP DNA, with apparently the same affinity as for the sites in natural DNA. By contrast, binding of the bisintercalating antibiotic echinomycin to the artificially created TpD sites is considerably enhanced (up to 1000 times) compared with its canonical sites surrounding CpG steps. 24 The preferential interaction of minor groove binders such as Hoechst 33258, berenil, and distamycin with AT-rich sequences of DNA is also dependent on 37 C. Bailly, M. J. Waring, and A. A. Travers, J. Mol. Biol. 253, 1 (1995). 38 C. R. Wagner, K. Hamana, and S. C. R. Elgin, Mol. Cell. Biol. 12, 1915 (1992). 39 C. Bailly, D. Payet, A. A. Travers, and M. J. Waring, Proc. Natl. Acad. Sci. U.S.A. 93, 13623 (1996). 4o M. J. Waring, Nature (London) 219, 1320 (1968). 41 C. Bailly, M. Brafia, and M. J. Waring, Eul: J. Biochem. 240, 195 (1996). 42 S. Kamitori and E Takusagawa, J. Am. Chem. Soc. 116, 4154 (1994).

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the position of the 2-amino group of guanine. 24 As for the intercalating drugs, relocating it onto adenines within the minor groove suffices to induce a major redistribution of the drug-binding sites. Figure 4 illustrates a typical DNase I footprinting gel obtained with the antibiotic netropsin. On binding to normal DNA well-resolved footprints appear at several sites, particularly between nucleotide positions 60-70 and 80-90, whereas cleavage by DNase I at the intervening sequence between positions 70 and 80 is massively enhanced in the presence of the ligand. Densitometric analysis yielded the differential cleavage plots shown in Fig. 5. The intense footprints centered around positions 65 and 85 correspond to sequences composed essentially of contiguous A. T base pairs as is the case with all other identified binding sites (indicated by black rectangles). Nuclear magnetic resonance (NMR) and crystallographic studies have confirmed that netropsin fits deeply into the narrow minor groove of (A. T)4 tracts. 43 Both the local geometry and the negative electrostatic potential in the minor groove favor the AT specificity of netropsin. 44 The footprints of netropsin are radically altered by the combined G --+ I and A ~ D substitutions, so that the antibiotic is displaced from its canonical sites at positions 60-70 and 80-90 and binds decisively to the intervening IC-rich cluster created in the doubly substituted DNA, formerly the most conspicuous region of enhanced DNase I cleavage (Fig. 4). The radically changed pattern of binding is emphasized in the differential cleavage plots of Fig. 5. The sites of binding (protection) and enhanced cutting are almost perfectly inverted in the doubly substituted polynucleotide. This observation corroborates previous resuits obtained with the singly substituted DNA species 45 indicating that the purine 2-amino group behaves as a negative effector for sequence-specific recognition of DNA by netropsin. The AT-specific antibiotic fails to bind to PCR product DNA in which the minor groove is completely obstructed with a 2-amino group on every base pair, as happens with DAP DNA, but binds nonspecifically all over molecules lacking the exocyclic amino group completely (inosine DNA). These effects are readily explicable in terms of changes in the geometry of the minor groove surface affecting the complementarity of the drug-DNA interface and hence direct recognition. 39 The only instance in which we found that repositioning the 2-amino group of G. C base pairs did not influence the recognition of preferred sequences by a small molecule was with a bisintercalating bisnaphthalimide drug, elinafide (LU79553), which is normally selective for sequences having an alternating purine-pyrimidine motif, particularly those containing GpT (ApC) and TpG (CpA) steps. Substitution with inosine and/or 2,6-diaminopurine has little effect on the distribution of binding sites for elinafide. This observation together with other data, particularly the fact 43 D. S. Goodsell, M. L. Kopka, and R, E. Dickerson, Biochemistry 34, 4983 (1995). 44 C. Bailly and J. B. Chaires, Bioconjug. Chem. 9, 513 (1998). 45 M. J. Waring and C. Bailly, Gene 149, 69 (1994).

Normal DNA

I+DAP DNA

netropsin

netropsin

FIG. 4. DNase I footprinting of netropsin on the tyrT (A93) DNA fragment containing the four natural nucleotides (normal DNA) or inosine and DAP residues in place of guanosine and adenine, respectively (I + DAP DNA). The products of DNase I digestion were identified by reference to the Maxam~3ilbert markers (lanes G + A), taking into account the difference in mobility of the fragments due to the presence or absence of a 3'-phosphate group. Control lanes show the products resulting from limited DNase I digestion in the absence of ligand. The remaining lanes show the products of digestion in the presence of the indicated antibiotic concentrations (micromolar). Numbers at the side of the gels refer to the numbering scheme of the DNA fragment used in Fig. 2.

498

[24]

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES I

I

I

I

0

0 < ,<

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[24]

D N A CONTAINING MODIFIED BASES

499

that the drug affects the reaction of D N A with dimethyl sulfate, suggests that the interaction must occur within the major groove of the double helix. 41

DNA-Containing

Uridine and/or

5-Methylcytosine

Residues

The same PCR-based strategy described above can be adapted to determine the influence of the other exocyclic groups of the nucleobases on DNA structure and l i g a n d - D N A interaction, for example, the effect of the 5-methyl group of pyrimidines on nuclease cleavage and sequence recognition by drugs. The 5-methyl group of thymidine residues occupies a pocket in the major groove of B-DNA, where it affects the structural and dynamic properties of the double helix. In particular, it appears to contribute to the unusual structures adopted by d A . dT tracts and participates to some extent in the stabilization of curved DNA sequences. 33'46 This substituent can be deleted from thymidines by substitution with uridine, and it can be added to cytidine residues by replacement with 5-methylcytidine (M). The combined C --+ M and T ~ U replacements are equivalent to relocating the 5-methyl group from T to M residues (Fig. 1). The appropriate PCR protocol is the same as that utilized for normal DNA (Table I), and both U and M can be efficiently incorporated into DNA by the heat-stable Taq polymerase. The deletion of the 5-methyl group of thymidines greatly affects both DNA structure and p r o t e i n - D N A interaction. Homopolymeric tracts of d A . dT become significantly more susceptible to DNase I cleavage when uridine is substituted for thymidine residues. 47 This observation was rather unexpected because there are no direct contacts between the enzyme and the major groove of DNA, only between DNase I and substituents within the minor groove. 48 Evidently removal of the thymidine methyl group from the major groove at AT tracts must induce perturbations that transmit into the opposite minor groove, where they can be detected by endonuclease probing. The perturbations might include electronic 46 p. j. Hagerman, Biochemistry 29, 1980 (1990). 47 C. Bailly, S. Crow, A. Minnock, and M. J. Waring,J. Mol. Biol. 291, 561 (1999). 48 D. Suck, J. Mol. Recognit. 7, 65 (1994).

FIG.5. Differential cleavage plots comparing the susceptibility of tyrT(A93) DNA to cutting by DNase I in the presence of increasing concentrationsof netropsin (25 nM to 25 tzM). Top: Cleavage of normal DNA containing the four natural nucleotides. Bottom: Cleavage of I + DAP DNA containing inosine and DAP residues in place of guanosine and adenine, respectively. Positive and negative values correspond, respectively, to enhanced or diminished DNase I cutting at each internucleotide bond. The values plotted compare the measured probabilities of cleavage expressed in logarithmic units. The binding sites for netropsin on normal DNA (filled rectangles) correspond essentially to AT-richsequenceswhereasthose on the doubly substitutedI + DAP DNA (hatchedrectangles)coincide predominantly with IC-rich sequences. As in Fig. 4, the tyrT DNA was labeled at the 5' end of the Watson strand.

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CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

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effects transmitted through the Watson-Crick base pairing, a phenomenon that seems to affect the reactivity of CpG steps toward mitomycin C and certain carcinogens in the minor groove.49 There is also evidence that removal of the bulky methyl group from an A. T pair in the major groove results in a change in the width of the opposite minor groove. 5° This could explain why AU tracts become more susceptible to DNase I cleavage than AT tracts, because the opening of the minor groove should facilitate the access of DNase I to the surface of the groove.47 Sequence-specific drug-DNA recognition in the minor or major groove of the double helix is minimally affected by the removal of the thymine methyl groups. Neither the DNA-threading intercalator nogalamycin, nor the aforementioned bisintercalator elinafide, both of which bind preferentially to sequences including GpT or TpG steps, shows any major sign of sensitivity to the removal of the methyl group. 47 All the drugs we have tested so far are unaffected by the T --~ U substitution. This is illustrated in the footprinting gel presented in Fig. 6, where it can be seen that the bisintercalator echinomycin binds equally well to a number of sites within the normal and uridine-substituted tyrT DNA. The antibiotic recognizes the same CpG-containing sequences irrespective of the presence or absence of the thymine methyl group. Other intercalating agents including ditercalinium, actinomycin, and daunomycin have been tested but in no case did the T --+ U substitution affect drug binding, as judged from similar DNase I footprinting experiments. If small molecules are blind to the T ~ U substitution, this is not the case for the HMG-D protein, which interacts 5-10 times more strongly with uridinecontaining tyrT DNA than with its counterpart containing natural nucleotides. This sensitivity to removal of the exocyclic methyl group of thymidine in the major groove, despite binding of the protein in the minor groove, is further evidence that structural changes in the major groove can lead to compensatory changes that translate into the minor groove. Whereas the removal of the methyl group (replacement of T with U) generally has little effect on sequence recognition by a variety of drugs, addition of a methyl group (replacement of C with 5-methyl-C), in contrast, does generate new binding sites for some DNA intercalators such as the acridine-4-carboxamide antitumor agents DACA and SN 16713, and the anthracyclines. Daunomycin, which is one of the most widely used anticancer drugs, intercalates preferentially at 5'-(A/T)GC or 5'-(A/T)CG triplets. 3°'51 Methylation of the cytosine residues in tyrT DNA causes the appearance of at least one new daunomycin-binding site at a G. M-rich region. 52 49A. Das, K. S. Tang, S. Gopalakrishnan, M. J. Waring,and M. Tomasz,Chem. Biol. 6, 461 (1999). 50G. Zhou and P. Ho, Biochemistry 29, 7229 (1990). 51j. B. Chaires, J. E. Herrera, and M. J. Waring,Biochemistry 29, 6145 (1990). 52A. Minnock,S. Crow,C. Bailly, and M. J. Waring,Biochim. Biophys. Acta 1489, 233 (2000).

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D N A CONTAINING MODIFIED BASES

501

echinomycin Uridine DNA

Normal DNA

GA ~,~o~q~ ~, ~ ~o Ct ~ b ~ q ~ b

¢o CtGA

m

w

-50

-40 J

J~o

-30 m

FIG. 6. DNase I footprinting of echinomycin on the tyrT(A93) DNA containing the four natural nucleotides (normal DNA) or uracil residues in place of thymines (uridine DNA). Control lanes (Ct) show the DNase I digestion in the absence of the test drug. Other details as in Fig. 4.

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Conclusions The PCR-based strategy to incorporate modified bases into DNA has been optimized with the tyrTfragment, which is 160 bp long. The protocol should work as well with other DNA fragments of 100-250 base pairs (providing that they do not contain too many long runs of contiguous A- T or G. C pairs, which may reduce considerably the efficiency of incorporation of certain modified bases). The protocol has been successfully used in our laboratory to make analog-containing variants of many DNA fragments in this size range. Moreover, it is possible to effect several types of modifications concomitantly. A totally synthetic tyrT DNA species containing inosine, methylcytosine (I. M) and diaminopurine, uridine (D-U) base pairs has been prepared in good yield by PCR. 53 This DNA contains no natural nucleotides at all, and its quadruple substitution completely switches the patterns of base pair hydrogen bonding and attachment of exocyclic groups. However, the radical alteration of its internal architecture does not prevent it from interacting with proteins. Although composed exclusively of I. M and D. U base pairs this DNA appears to bind to a histone octamer more tightly than does natural DNA and, surprisingly, its rotational orientation on the surface of the protein is not affected.53 DNA is highly adaptable and can evidently tolerate extensive modifications at the nucleotide level as well as at the level of its sugar-phosphate backbone. I, M, D, and U are only four representative examples of modified bases that can be incorporated into DNA via the PCR methodology presented here. But the method is general and many other types of modified bases can be introduced into the DNA sequences. 5-Fluoro-, 5-bromo-, and 5-nitrouracil bases were successfully incorporated into the tyrT fragment by much the same procedure as for preparing the uridinesubstituted DNA. Borane derivatives and many other synthetic nucleoside triphosphates can be employed to engineer DNA molecules. The proposed PCR procedure is widely applicable and can be extremely useful to help comprehend how ligands, small and large, recognize structural elements of the DNA double helix. Acknowledgments The authors thank all the past and present members of the Waring laboratory in Cambridge whose work is discussed here: Eric Sayers, Christophe Marchand, Dean Gentle, Andrew Minnock, Steve Crow, Mao Guo, Stefan Jennewein, and Johanna Virstedt, together with our collaborators: Andrew Travers, Brad Chaires, Niels Erik MOllegaard, and Peter Nielsen. This work was done with the support of research grants (to C.B.) from the Ligue Nationale Franqaise contre le Cancer and (to M.J.W.) from the European Union, the Wellcome Trust, and the Cancer Research Campaign. The authors thank the Sir Halley Stewart Trust for a grant to assist cooperation.

53 M. Buttinelli, A. Minnock, G. Panetta, M. J. Waring, and A. A. Travers, Proc. Natl. Acad. Sci. U.S.A. 95, 8544 (1998).

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503

[25] Drug Interactions with Nucleosomes and Chromatin By Jose PORTUGAL Introduction Considerable attention has been focused on understanding the nature of the interaction between chemotherapeutic drugs and their DNA targets in vitro. However, in the cell, the DNA is a component of chromatin, which consists of DNA complexed with a collection of histones and other nuclear proteins.l-3 Determining whether drugs are able to bind to DNA within the chromatin, and what effects this might have, is important to our understanding of how drugs can interact with DNA in eukaryotic cells. The basic subunit of chromatin is the nucleosome core particle. I It consists of two copies of each histone protein (H2A, H2B, H3, and H4) assembled into an octamer, and about 146 bp of DNA that are wrapped around the proteins in a left-handed superhelix. Individual core nucleosomes are separated by variable lengths of internucleosomal linker DNA. In the presence of histone H 1, the nucleosome (i.e., nucleosome core plus linker DNA and H 1) compacts into higher order helices. 4-6 Although nucleosomes can contain many different DNA sequences, there are grounds for considering that they adopt well-defined locations on DNA sequences. Consequently, nucleosomes are often positioned at preferred locations near regulatory elements, yet the observed position of the nucleosome arrays is not always uniquely defined, nor are they static, but can be involved in the modulation of gene expression. The tight wrapping of DNA about the histone octamer suggests that the bendability of DNA may be the main determinant of its positioning (phasing) on the nucleosome core particle. Hence, DNA sequences that are more difficult to bend are likely to be excluded from the nucleosome core particle. 7 Moreover, in the nucleosome core particle AT runs are preferentially placed with the minor groove facing approximately inward toward the protein octamer, whereas the GC runs tend to occupy positions where the minor groove faces outward; although it is I K. Luger, A. W. Mader, R. K. Richmond, D. E Sargent, and T. J. Richmond, Nature (London) 389, 251 (1997). 2 j. L. Workman and R. E. Kingston, Annu. Rev. Biochem. 67, 545 (1998). 3 R. D. Kornberg and Y. Lorch, Cell 98, 285 (1999). 4 j. T. Finch and A. Klug, Proc. Natl. Acad. Sci. U.S.A. 73, 1897 (1976). 5 D. L. Bates, P. J. Butler, E. C. Pearson, and J. O. Thomas, Eu~: J. Biochem. 119, 469 (1981). 6 j. Ausi6, N. Borochov, D. Seger, and H. Eisenberg, J. Mol. Biol. 177, 373 (1984). 7 H. R. Drew and A. A. Travers, ,l. Mol. Biol. 186, 773 (1985).

METHODS IN ENZYMOLOGY,VOL. 340

Copyright~ 2001 by AcademicPress All rightsof reproductionin any form reserved. (X)76-6879/00$35.00

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unlikely that all the local bending preferences will be satisfied by a single precise path of DNA on the histone octamer. The mechanisms of drug binding to nucleosomes, and any structural changes involved, are well documented, but they are not fully understood, s-l I Comparatively, studies devoted to the analysis of drug binding to large chromatin are rather scarce. ~2-~4 A better understanding of the action of these drugs is important for our comprehension of their mechanism of action in cell nuclei.15

Preparation

of Chromatin

Chromatin can be prepared from chicken erythrocytes by the method of Lutter, 16 with some modifications. 17 It is also possible to use other biological materials, such as rat liver or calf thymus glands, in which case mechanical homogenization of the samples is required. J8 The following protocol is to prepare chromatin from chicken blood. For other biological sources see Kornberg et al., 18 and references therein. Chicken blood can usually be collected from a local slaughterhouse, or via heart puncture from live chickens (when this procedure is performed by a trained person it does not seriously injure the animal). All the steps are carried out at 4 ~ unless otherwise indicated. In any case, the protocols using previously purified nuclei (see step 6 below) are equivalent for all. 1. Mix 50 ml of fresh chicken blood with about 7 ml of 85 mM sodium citrate (pH 7.0) to prevent coagulation. Dilute the blood to 500 ml with 0.34 M sucrose in buffer A [buffer A: 15 mM potassium cacodylate (pH 6.0), 60 mM KC1, 15 mM NaC1, 0.5 mM spermidine, 0.15 m M spermine] supplemented just before use with 0.5 m M EGTA, 15 mM 2-mercaptoethanol, 0.2 m M phenylmethylsulfonyl fluoride (PMSF), and 1 mM benzamidine. 2. Centrifuge the resulting mixture at 500g for 3 min. Discard the supernatant. Resuspend the pellet in 500 ml of the same solution described in step 1. 8 j. B. Chaires, N. Dattagupta, and D. M. Crothers, Biochemisoy 22, 284 (1983). 9 C. M. L. Low, H. R. Drew, and M. J. Waring,Nucleic Acids Res. 14, 6785 (1986). 10j. Portugal and M. J. Waring, Biochimie 69, 825 (1987). I I p. M. Brown and K. R. Fox, Methods Mol. Biol. 90, 81 (1997). 12E. J. Gabbay and W. D. Wilson, Methods" Cell Biol. 18, 351 (1978). 13C. Bourdouxhe, P. Colson, C. Houssier, J. S. Sun, T. Montenay-Garestier, C. H616ne, C. Rivalle, E. Bisagni, M. J. Waring, J. P. H6nichart, and C. Bailly, Biochemistry 31, 12385 (1992). 14A. Rabbani, M. Iskandar, and J. Ausi6, J. Biol. Chem. 274, 18401 (1999). ].5j. S. Mymryk, E. Zaniewski, and T. K. Archer, Proc. Natl. Acad. Sci. U.S.A. 92, 2076 (1995). 16L. C. Lutter, J. Mol. Biol. 124, 391 (1978). 17H. R. Drew and C. R. Calladine,J. MoL Biol. 195, 143 (1987). 18R. D. Kornberg,J. W. LaPointe, and Y. Lorch, Methods EnzymoL 170, 3 (1989).

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3. Repeat the previous step twice. The supernatant should be fairly clear, but white blood cells can form a thin layer on top of the erythrocytes, and can be removed with a pipette. 4. The pellet is resuspended in 500 ml ofthe solution described in step 1,which has been previously supplemented with 0.1% (w/v) Nonidet P-40 (NP-40) and the pH brought to 7.5 with Tris base. This will release the cell nuclei. Centrifuge at 1000g for 5 min and discard the supernatant. Resuspend the pellet in 500 ml of the same solution. 5. Repeat step 4 twice. The final pellet, which contains washed nuclei, should be milky white. The washed nuclei are resuspended in buffer A containing 0.34 M sucrose, 0.2 mM PMSF, and 15 mM 2-mercaptoethanol, and centrifugated at 1000g for 3 min. 6. Resuspend the pellet in 100 ml of the same buffer, previously adjusted to pH 7.5 with Tris base. Any undissolved material must be broken into smaller pieces by gentle pipetting. The DNA concentration of the solution can be determined at 260 nm in 0.1 M sodium hydroxide, and the absorbance adjusted to 50 units/ml of nuclei (about 5 mg/ml of protein plus DNA).I 1 7. Adjust the solution to 1 mM CaCI2 to prepare it for micrococcal nuclease digestion. Incubate at 37 ° for 3 min. Add micrococcal nuclease to a concentration of 40 units/ml and digest the sample for 5 rain. It is worth performing a trial digestion, using an aliquot of the solution, to find the optimum digestion time. 8. Stop the digestion by adding EDTA up to 2 mM. Centrifuge the solution at 3000g for 10 min. Discard the supernatant. 9. Resuspend the pellet in 10 mM Tris-HC1 (pH 8.0) containing 0.2 mM EDTA and 0.2 mM PMSF at approximately half the previous volume. 10. Keep the solution on ice for about 30 rain. Shake it gently to bring the chromatin back into solution. After 30 min, centrifuge at 3000g for 10 min. Discard the pellet and measure the supernatant volume accurately. To prepare large chromatin containing all the histone complement, the following steps must be omitted and the sample should be stored directly as described in step 13. 11. A stock 4 M NaCI solution is added dropwise while stirring until a final concentration of 0.65 M is obtained. This step releases histones H5 and H1 quantitatively. 12. Apply the previous solution to a column of Sepharose 4B (2 × 80 cm) equilibrated in 20 mM sodium cacodylate (pH 6.0), 0.63 M NaC1, 0.2 mM PMSF, 0.2 mM EDTA. The H1/H5-depleted chromatin elutes, in the same buffer, after 4-6 hr. A plot of absorbance at 260 nm versus fraction number should show two peaks: the first, which contains the HI-depleted chromatin, and the second

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containing linker DNA plus H1 and H5 proteins. Fractions corresponding to the first peak are pooled. The absence/presence of H 1 and H5 can be checked with a small gel, containing 18% (w/v) polyacrylamide and 0.1% (w/v) sodium dodecyl sulfate (SDS), stained with Coomassie blue. 13. The Hl/H5-depleted chromatin sample is concentrated to 4 mg/ml, using an Amicon (Danvers, MA) PM10 membrane. The sample can be stored at 4 ° for 2 months, or at - 2 0 ° for 6 months in 50% (v/v) glycerol.

Purification of 175-Base Pair Nucleosomes and Nucleosome Core Particles 1. Take a small portion of the chromatin preparation (step 13, above) and adjust it to 20 mM Tris-HC1 (pH 7.5) and 1 mM CaC12. Digest with micrococcal nuclease (100 units/ml) for about 30 min, removing 20/tl every 2 rain, and adjusting it to 2 mM EDTA. Extract chromatin DNA by proteinase K digestion (0.5 #g/ml) in 0.1% (w/v) SDS, followed by phenol extraction and ethanol precipitation. The digested samples should be checked on an 8% (w]v) polyacrylamide gel containing 7 M urea, stained with ethidium bromide, and the time course of digestion visualized under ultraviolet light. 17 A point of time must be chosen at which a 175-bp band (core particle plus intemucleosomal linker DNA), or a 145-bp band (core particle), is most abundant. 2. Adjust the bulk of the nucleosome preparation to 20 mM Tris-HCl (pH 7.5) and 1 mM CaCI2 and digest it according to the conditions determined in the previous step. Stop the reaction by adding EDTA up to 2 mM. 3. Centrifuge the digested material (3000g for 5 min). Discard the pellet, and apply the supernatant to a Sepharose 6B column (2 × 80 cm) that has been previously equilibrated in 20 mM sodium cacodylate (pH 6.0), 100 mM NaCl, 0.2 mM EDTA, 0.2 mM PMSE 4. The nucleosome core particles elute after about 6 hr. A plot of absorbance versus fraction number should have two peaks: the first contains the nucleosome core particles (or 175-bp nucleosomes when the micrococcal nuclease digestion time was adjusted to obtain them). The fractions containing the nucleosomes are combined into a single volume and concentrated with an Amicon PM10 membrane. Nucleosomes can be stored at 4 ° for 2 months or at - 2 0 ° for up to 6 months in 50% (v/v) glycerol containing 1 mM benzamidine. Histones should be electrophoreticaUy checked for proteolysis. They can be conveniently analyzed on 18% (w/v) polyacrylamide gels containing 0.1% (w/v) SDS, and stained with 0.1% (w/v) Coomassie blue. Alternatively, 175-bp nucleosomes have been obtained in sufficient purity by centrifugation in a 5-20% (w/v) sucrose gradient. 8

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Preparation of Chromatin of UniformLength An oligonucleosome chromatin of uniform length (either containing all the histones, or as H 1/H5-depleted chromatin) can be obtained from a general preparation as follows. 1. Centrifuge the nuclear lysate (obtained as described above) at 12,000g for 15 rain. 2. Fractionate the superuatant, using a 5-20% (w/v) sucrose gradient in 10 mM Tris-HC1 (pH 7.5) containing 20 mM NaC1 and 0.5 mM EDTA. Our protocol uses an SW28 rotor (Beckman, FuUerton, CA) at 82,000g (4 °) for 3 hr. 14 A homogeneous preparation of monomers, dimers, trimers, and teramers may be obtained by sedimenting pooled fractions in a second sucrose gradient.18 3. Collect fractions of 1 ml and determine their absorbance at 260 nm. 4. Take an aliquot of some fractions and extract chromatin DNA by proteinase K digestion (0.5 mg/ml) in 0.1% (w/v) SDS, followed by phenol extraction and ethanol precipitation. 5. Run the DNAs on a 1-1.2% (w/v) agarose gel in 45 mM Tris-borate (pH 8.4), 1 mM EDTA to analyze their lengths. 5 6. Take the fractions of interest (those containing DNA of similar size should contain on average the same number of nucleosomes), combine and dialyze them against the desired buffer, supplemented with 0.2 mM PMSF and 1 mM benzamidine, and store at 4 ° until required (it is better to use immediately) to assay the chromatin-drug interactions. The average number of nucleosomes in a chromatin preparation can be estimated indirectly from the size of the DNA fragments, determined in step 5, considering a nucleosome repeat length to be 210 bp (for chicken erythrocyte chromatin).

Q u a n t i t a t i v e A n a l y s i s of D r u g B i n d i n g to N u c l e o s o m e s and Chromatin This section describes methods to quantify the binding of drugs to DNA wound around the histone octamer, and to compare with the binding to the linker DNA in either nucleosomes or large (polynucleosomal) chromatin samples. Either nucleosome core particles or 175-bp nucleosomes, prepared as described above, can be used in the binding experiments. The objectives of a quantitative analysis may include comparison of the binding constants between free DNA and DNA wound around the protein. We might anticipate from a qualitative point of view that any drug will prefer free DNA regions over protein-bound regions.

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The results reported to date using large chromatin and intercalating drugs provide conflicting conclusions. 12'19'2° Two main reasons for these discrepancies appear to be the use of poorly characterized chromatin preparations, in which the stability of the protein has not been determined, and the different buffers and ionic strengths employed. The first is difficult to overcome because of the labile nature of chromatin. The use of freshly prepared nucleosomes, whose histone integrity is electrophoreticaily checked, is advisable. The ionic strength of the buffer should be selected both to maintain the shape of the nucleoprotein and to allow significant binding of the drug, because the extent of binding is expected to decrease with increasing salt content. It should be emphasized that the three-dimensional folding of chromatin is salt dependent. 4,6 For either large chromatin, or H1/H5-depleted chromatin, circular dichroism and sedimentation analyses can be used to test the integrity of nucleosome organization of chromatin. 21"22 Binding isotherms can be obtained, using either nucleosomes or chromatin, from equilibrium dialysis or fluorescence titration experiments using an adapted version of the protocols described for free DNA (for a more extensive discussion on these methods see [1] in this volume23). In fluorescence studies, the changes in the fluorescence properties of a drug can be analyzed with addition of chromatin or nucleosomes. To obtain meaningful results the changes in fluorescence should be qualitatively almost the same for chromatin and DNA in terms of shifting of the maximum wavelengths, and enhancement or reduction of quantum yield. 12 Equilibrium dialysis is mostly used for drug-chromatin binding studies in solution.S, 12,14 General Protocol f o r Determining Binding Isotherms by Equilibrium Dialysis

1. Fill a Spectra/Por 2 dialysis tube (Spectrum, Los Angeles, CA) with about 1 ml of a solution of chromatin, or nucleosomes, corresponding to about 40/_tg of DNA per milliliter, in 10 mM Tris-HC1 (pH 7.4), containing 0.1 mM EDTA and 75 mM NaC1 (the salt concentration can be altered as required). For this technique, numerous types of apparatus and methods of membrane preparation might be satisfactory. 2. Dialyze the contents of the tube against the drug solution in the same buffer, at 4 ° . Equilibrium is normally reached within 48-72 hr. 3. Measure the concentration of free drug in the dialysate, using absorbance or fluorescence intensity. The amount of drug inside the dialysis tube can also be determined directly by dissociating the complex with 1% (w/v) SDS. 19L. Kleiman and R. C. Huang,J. Mol. Biol. 55, 503 (1971). 20C. T. McMurrayand K. E. van Holde, Proc. Natl. Acad. Sci. U.S.A. 83, 8472 (1986). 21j. Ausi6, R. Sasi, and G. D. Fasman, Biochemistry 25, 1981 (1986). 22R. L. Rill, B. R. Shaw, and K. E. Van Holde, Methods Cell Biol. 18, 69 (1978). 23j. B. Chaires, Methods Enzymol. 340, [1] 2001 (this volume).

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It is advisable to measure the drug concentration on both sides of the membrane to determine the bound and unbound ligand directly, thus avoiding errors arising from drug binding to the dialysis membrane. The procedures to measure free and bound drug concentrations, and the way to analyze them to obtain reliable binding isotherms, are mostly the same as those used for DNA-drug interactions, and they are presented in [1] in this volume. 23 So far, analysis of the binding of different drugs to chromatin has been undertaken with four different kinds of preparation: nucleosome core particles, nucleosomes (HI-depleted 175-bp nucleosomes), large chromatin fragments, and H 1-depleted large chromatin. Several drugs, including minor-groove binders, have been used in quantitative studies of drug-chromatin interactions, but only ethidium bromide and the anthracycline antibiotics have been widely studied, s,~3,19,20,24 For all the anthracyclines studied, the binding to the nucleohistone appears to increase in the following order: large chromatin < HI-depleted chromatin < core particles < 175-bp nucleosomes < DNA.S,14,25 Several anthracyclines have also been ranked with respect to their relative binding to nucleosomes. 25-27 The presence of the core histones usually reduces the affinity of the anthracyclines for DNA, as do H 1 and H5 in internucleosomal DNA. Moreover, with a large-chromatin sample, when the ionic strength increases the nucleosomes come close together and an important fraction of the internucleosomal DNA is hindered. The binding of ethidium bromide to nucleosomes is controversial, 2°'24,2s illustrating the difficulties in comparing experimental results that come from biological samples prepared in different laboratories. Cooperative binding to nucleosomes, with initial binding weaker than binding to DNA, has been described, 29 whereas another study indicated that ethidium, like other intercalators, unfolds nucleosomes, and that the binding was noncooperative and stronger than binding to free DNA. 24 More recently it has been suggested that slow dissociation of the protein may take place after drug binding, and therefore when the binding isotherm is obtained by correcting total binding for that seen with free DNA, the cooperativity becomes evident, 28 but it seems that a critical amount of drug would be required for the dissociation to take place. 28 It should be mentioned here that ethidium bromide has been used to analyze different aspects of nucleosome dynamics because it applies

24 H. M. Wu, N. Dattagupta, M. Hogan, and D. M. Crothers, Biochemistry 19, 626 (1980). 25 H. Fritzsche, U. W~hnert, J. B Chaires, N. Dattagupta, E B. Schlessinger, and D. M. Crothers, Biochemiso'y 26, 1996 (1987). 26 C. Cera and M. Palumbo, Nucleic" Acids Res. 19, 5707 (1991). 27 C. Cera, G. Palu, S. M. Magno, and M. Palumbo, Nucleic" Acids Res. 19, 2309 (1991). 28 C. T. McMurray and K. E. van Holde, Biochemistry 30, 5631 (1991). 29 M. Erard, G. C. Das, G. de Murcia, A. Mazen, J. Pouyet, M. Champagne, and M. Daune, Nucleic' AcidsRes. 6, 3231 (1979).

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a positive constraint through its intercalation. 3° It seems that mononucleosomes on DNA minicircles can tolerate large positive supercoiling. A n a l y s i s of E f f e c t s of D r u g s o n R e c o n s t i t u t e d N u c l e o s o m e s b y D N a s e I or H y d r o x y l R a d i c a l C l e a v a g e

Preparation and Radioactive Labeling of Cloned DNA Molecules Approximately 500 ng of the DNA to be reconstituted into nucleosome core particles should be cut out of the carrying plasmid, using appropriate restriction enzymes, and labeled selectively at one of the recessed ends, as described in [20] in this volume. 31

Reconstitution of Nucleosomes with Radiolabeled DNA Fragments There are several methods by which radioactive DNA fragments can be efficiently reconstituted into nucleosome core particles. We describe one of them: the salt exchange method. 7 Other well-established methods might also be used, such as the histone transfer method or reconstitution using acidic assembly factors. 32 A preparation of H 1-depleted chromatin (described above) can be used directly as a source of histones for reconstitution experiments. The pros and cons of reconstitution by the salt exchange method (also known as the salt gradient method) have been discussed previously. 32 The preparation of purified nucleosome core particles is not usually required for preparing nucleosomes reconstituted with DNA fragments shorter than 200 bp.l In the salt exchange method, radioactive DNA is combined with the nucleosome core histones by the exchange of the proteins onto DNA at high salt concentration. 1. Dissolve about 120,000 cpm of end-labeled DNA in 10 #1 of TE buffer [20 mM Tris-HC1 (pH 7.5), 1 mM EDTA]. Reactions should be carried out in siliconized tubes to avoid the attachment of radiolabeled DNA to the walls. 2. Add 8 #1 of 80 mM Tris-HCl (pH 7.5) containing 4 M NaCI and 1 mM EDTA, followed by 2 #1 of 50 mM PMSK 3. Add 20 #1 of either HI-depleted chromatin or nucleosome core particles (from a 4-mg/ml stock). 4. Incubate at 37 ° for 30 min. In a few minutes, the large excess of nucleosome cores promotes exchange between the labeled DNA and nucleosome core DNA.

30 A. Sivolob and A. Prunell, J. Mol. Biol. 295, 41 (2000). 31 K. R. Fox and M. J. Waring, Methods Enzymol. 340, [20] 2001 (this volume) 32 D. Rhodes and R. A. Laskey, Methods Enzymol. 170, 575 (1989).

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5. To "freeze" the equilibrium, the NaC1 concentration must be slowly reduced to about 100 mM. This is carried out, at room temperature, by slow stepwise addition of 20 mM Tris-HC1 (pH 7.5) containing 0.1% (w/v) Nonidet P-40:6 additions of 10/zl followed by 11 additions of 20/zl, leaving 5 min between each addition. 6. Store the radioactive reconstituted nucleosomes at 4 °. They can be used over 7-10 days without significant degradation. The efficiency of reconstitution can easily be monitored in 0.7% (w/v) agarose gels run at 25 mA in 45 mM Tris-borate (pH 8.3), 1 mM EDTA, and observed by autoradiography. Figure 1B displays a typical example: DNA reconstituted into nucleosomes shows a slower mobility than the free DNA. However, this result does not necessarily mean that the nucleosome core particles have been correctly reconstituted, but only demonstrates a stable DNA-protein interaction. 11 An authentication of reconstituted nucleosome core particles would require DNase I cleavage analysis, v,16 The resulting cleavage pattern should present cuts occurring every i0 nucleotides, as described for native nucleosome core particles.

Footprinting Cleaving Agents DNase I is stored at - 2 0 ° at a concentration of 7000 units/ml in 150 mM NaC1, and diluted to working concentrations in 20 mM NaC1, 2 mM MgCI2, and 2 mM MnC12. Solutions for hydroxyl radical cleavage must be prepared immediately before use. Mix 0.2 mM ferrous ammonium sulfate, 0.2 mM EDTA, 10 mM L-ascorbic acid, and 0.1% (v/v) hydrogen peroxide in a ratio of 1 : 1 : 2 : 2 , respectively.

DNase I Footprinting Digestion of the reconstituted material with DNase I is performed in parallel with the digestion of an aliquot of the same labeled DNA that has not been reconstituted into nucleosomes. 9 1. Take 10/zl of reconstituted nucleosome core particles (this volume can be modified to obtain the approximately 3000 cpm that are required for each digestion). 2. Add 10 #1 of the drug solution in 10 mM Tris-HC1 (pH 7.5) containing 100 mM NaC! to obtain the desired final concentration to be incubated with the core particle. In parallel, prepare a drug-DNA complex as a control lane. Incubate at 37 ° for 20 min. If any organic solvent has been used to dissolve the drug, its final concentration should be lower than 5% in the final volume of incubation. 3. Adjust the solution to 1 mM MgC12 and incubate with DNase I for 2 min at a final enzyme concentration of 2 U/ml for core, or 0.01 U/ml for free DNA (the

A. +

Net.

Cor,

B.

Core

DNA

Core

÷

Netropsln

Cl

1,0

®

Q,5

.0,5 ~10

o. og

.VV

-

100 Bas~ numbe~

FIG. 1. (A) DNase I footprinting ofnetropsin bound to flee tyrT DNA or to nucleosome core particles containing this DNA fragment. Asterisks indicate the positions of some new bands that appear in the nuclease cleavage of core particles in the presence of increasing amounts of netropsin (Net.), 10 and 20/~M, respectively. The new electrophoretic bands might be produced after a change in the DNA phasing about the nucleosome core particle (see text for details). (B) An example of the reconstitution of the histone octamer with tyrT DNA. The reconstituted DNA-core particle migrates more slowly than DNA alone (on a 0.7% agarose gel). The other track shows that the presence of 30/~M netropsin, a concentration higher than that used in the footprinting experiment displayed in (A), produces only a slight detachment of DNA from the core particles (J. J. P6rez, and J. Portugal, unpublished observations, 2001 ). (C) Plot representing the orientation of DNA on the core particle in the presence of four netropsin molecules bound to the DNA, as measured by the angle 0 between the place of contact of DNA with the protein surface and the middle of the minor groove. The plot shows the values of - c o s (0) versus the base pair numbering of the DNA. The positive values represent the DNA minor groove facing outward from the protein surface. The arrows indicate the base pairs that are deemed to be more sensitive to nuclease attack according to footprinting results (A) [J. Portugal and M. J. Waring, Nucleic Acids' Res. 14, 8735 (1986)]. The new electrophoretic bands lie approximately midway between the digestion maxima observed in the absence of the ligand. [(C) Reproduced, with permission, from J. J. P6rez and J. Portugal, Nucleic Acids Res. 18, 3731 (1990).]

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exact amount of enzyme required, as well as the digestion time, should be adjusted for each set of experiments in order to obtain "single-hit" kinetics). 4. Stop the reaction by adding EDTA to 2 mM, and adjust it to 0.1% (w/v) SDS. 5. Digest the sample with proteinase K (0.5 mg/ml) at 37 ° for 15 min. Extract the entire reaction volume twice with phenol-chloroform, and remove the residual phenol with water-saturated ether. 6. Adjust the resulting solution to 0.35 M sodium acetate (pH 5.8) and add 3 volumes of absolute ethanol to precipitate the DNA. After centrifugation, wash the pellet twice with 75% (v/v) ethanol and dry it briefly. 7. Redissolve the dried pellet in 8 #1 of 85% (v/v) formamide containing 10 mM EDTA (pH 8.0) and 0.05% (w/v) bromphenol blue. Before gel loading (see below), the samples must be heated at 90 ° for 2 rain and cooled immediately on ice.

Hydroxyl Radical Footprinting Hydroxyl radical cleavage of free DNA is characterized by an even pattern of cleavage products, while the hydroxyl radical produces a manifest 10-bp modulation in DNA fragments from the nucleosome core particle. 33 The following is based on previously described protocols. 11'33 The nucleosome core particles should contain no glycerol, which acts as a hydroxyl radical quencher. 1. Prepare the DNA~trug and nucleosome-drug complexes as described above for DNase I footprinting (20-#1 final volume). 2. Add 40 #1 of the hydroxyl radical mixture, described under Footprinting Cleaving Agents (above), freshly prepared immediately before use. 3. Allow the reaction to take place for about 5 min (the cleavage time might require some adjustment in each experiment), but shorter reaction times are required for free DNA. 4. Add 100 #1 of phenol-chloroform to stop the reaction; recover the aqueous phase by centrifugation. 5. Extract the sample again with phenol-chloroform. Combine the aqueous phases, and remove the residual phenol with water-saturated ether. 6. Prepare the samples for electrophoresis as described in steps 6 and 7 for DNase I footprinting.

Gel Electrophoresis of Footprinting Samples Load the samples containing digested materials onto a denaturing polyacrylamide gel [0.3 mm × 40 cm, 8% (w/v) polyacrylamide gel containing 8 M urea]. Run the gel at 40 W (about 1500 V) for 1.5-2 hr in 90 mM Tris-borate (pH 8.3), 2 mM EDTA. To obtain a better resolution of the longer DNA fragments the sample 33M. E. Churchill,J. J. Hayes,and T. D. Tullius,Biochemistry29, 6043 (1990).

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can be applied in two loadings, in parallel, with a ~45-min delay between them. After running the gels can be fixed in 10% (v/v) acetic acid, or simply soaked in distilled water, transferred to Whatman (Clifton, NJ) 3MM paper, and dried under vacuum at 80 °, and subjected to autoradiography. Data obtained from the microdensitometric scanning of those autoradiographs can be quantitatively compared to produce differential cleavage plots (e.g., drugcore minus untreated core particle) normally displayed on a logarithmic scale, in which positive values would represent enhanced cleavage at any bond, and negative values blockage.9' l0 Footprinting Detection of Drug Effects on Chromatin: Alteration of Rotational Positioning of DNA on Nucleosome Core Particles The use of DNase I, or hydroxyl radicals, to cleave DNA wound around the core particle in the presence of binding drugs provides us with direct answers to fundamental questions: Can drugs still bind to DNA associated with histones, and if so, what are the recognized DNA sequences? What effect does the bound drug have on the DNA bound to the protein, or in the three-dimensional nucleosome structure? The sequence-specific interaction of several intercalating drugs and minor groove binders has been studied with nucleosome core particles reconstituted with various DNA fragments. 9,34-36 Figure 1 shows an example of the digestion of nucleosome core particles containing the tyrT DNA fragment, incubated with the minor groove-binding drug netropsin. In the presence of the ligand a new DNase I digestion pattern appears, in which new electrophoretic bands are localized midway between the maxima seen in the digestion of the core particles alone. Many such bands (see Fig. 1A and C) are new, because they are not observed in the cleavage of free DNA or in the presence of the ligand. This kind of experimental result is consistent with a model in which the presence of bound drug causes the DNA to rotate by roughly 180 ° on the nucleosome surface. In this way, the DNA regions to which a drug binds are turned toward the protein while those that were facing in are turned out. It seems likely that the induced change in DNA setting takes place because the binding of the drug molecules to their preferred sequences would serve to optimize nonbonded contacts between the ligand and the polynucleotide backbone, and to minimize the potential electrostatic repulsion between the cationic end of many drugs and the charged groups in the histone core surface. Molecular modeling studies have shown that the more stable rotational setting of DNA on a histone octamer can be changed after the binding of four netropsin molecules, thus giving a new structure with an energetic minimum achieved when DNA had turned around 180 ° on the protein surface 37 (see Fig. 1C). 34j. Portugal and M. J. Waring, NucleicAcids Res. 14, 8735 (1986). 35p. M. Brownand K. R. Fox, J. Mol. Biol. 262, 671 (1996). 36j. Portugal and M. J. Waring,NucleicAcids Res. 15, 885 (1987).

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The footprinting experiment displayed as an example in Fig. 1A, has been confirmed by a series of hydroxyl radical and DNase I footprinting analyses using nucleosomes reconstituted with different DNA fragments. 35'36 In hydroxyl radical cleavage experiments some drugs had little effect on the cleavage pattern, II which might be related to the particular location of binding sites in the reconstituted DNA fragment. It is noteworthy that the total DNA concentration is higher in the nucleosome footprinting experiments than in experiments using free DNA because in the nucleosome core preparations there is a large amount of unlabeled DNA present apart from the radiolabeled nucleosomal DNA. 35 In the absence of histones, drug-DNA interactions are dictated by the equilibrium binding constant, whereas for the core particles the amount of bound drug is closely related to the drug : DNA stoichiometric ratio. 35 Although most of the ligands analyzed to date produce changes in the cleavage patterns of the DNA around the nucleosome, this is not always the case. For example, mithramycin appears to have no.effect on the phasing of DNA complexed with nucleosome core particles, 38 whereas with daunomycin some enhanced electrophoretic bands appear at positions different from those of the core panicle cleavage, but they do not conform to any regular pattern. I° Actinomycin D has no effect at low concentrations, but there is a large displacement of DNA from the nucleosome core particle as the drug concentration is raised, l° The occurrence of conformational changes associated with the binding of drugs, without gross detachment of DNA from the core panicle, requires a low occupancy of the potential binding sites in a DNA fragment. 9,35.36 S e d i m e n t a t i o n A n a l y s i s of C h r o m a t i n - D r u g C o m p l e x e s Sedimentation velocity experiments permit the analysis of perturbations in the hydrodynamic parameters of chromatin and nucleosomes caused by drug binding. Moreover, changes in the sedimentation coefficient of samples studied under different salt conditions can indicate certain characteristics of the preferential binding of a drug to, for example, the exposed linker DNA versus the DNA wrapped around the histones core, as well as the effects of DNA-binding drugs on the folding of chromatin fibers, or changes in the shape of the nucleosomes. The analysis of drug-nucleosome binding is complicated by the ability of several drugs to promote nucleosome aggregation and/or disruption of the core particle. The binding of ethidium bromide to core particles can result, for example, in partial DNA unwinding 24 and in a stepwise dissociation of the nucleohistone structure, a° Sedimentation velocity analysis of drugs incubated with either nucleosomes, chromatin, or HI-depleted chromatin can be performed, at the desired salt 37 j. j. Prrez and J. Portugal, Nucleic' Acids Res. 18, 3731 (1990). 38 K. R. Fox and B. M. Cons, Biochemistry 32, 7162 (1993).

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concentrations, in a Beckman XL-A analytical ultracentrifuge, using an aluminum An-55 rotor and double sector aluminum-filled Epon centerpieces. 14 Alternatively, it is possible to use, for example, a Beckman model E analytical ultracentrifuge and an AnF rotor. 8 Runs are usually performed at 20 °, and sedimentation coefficients are routinely obtained from the slope of plots of In X versus time (where the radial position X is the distance from the center of rotation to the half-height of the UV scanner traces)/ The sedimentation coefficients of chromatin and H1/H5-depleted chromatin increase with ionic strength because of chromatin folding, which also depends on the presence of the linker-bound histones H 1 and H5.4'6 Hence, the salt concentration can play a dual role via an effect on chromatin fiber folding or a direct effect on the magnitude of binding. For example, the binding of daunomycin has been widely analyzed using either nucleosomes or chromatin. 8' 14 The anthracyclines usually induce chromatin aggregation in experiments with either nucleosome or oligonucleosome preparations, which is easily detected in sedimentation analysis. This behavior has been tentatively considered to arise from nucleosome core unfolding followed by aggregation, a' J4,20

C i r c u l a r D i c h r o i s m A n a l y s i s of D r u g I n t e r a c t i o n s w i t h Nucleosomes or Chromatin Circular dichroism (CD) has frequently been used to analyse DNA conformation, but also to study several aspects of chromatin structure 22'39 and chromatindrug interactions. 13.26,40 An extensive discussion on the use of CD to study DNAdrug interactions is presented by Eriksson and Nord6n ([4] in this volumeal). Although the basic CD spectrum of DNA and the protein CD spectra are available to help interpret the circular dichroism of chromatin, the complex spectrum of the nucleohistone is not easily unraveled into its component parts. Notwithstanding, nucleosomes and whole chromatin show CD spectra that are easily distinguished from those of DNA alonefl 2

Analysis of Circular Dichroism Data The chromatin and chromatin~:trug solutions to be measured must be optically pure. Chromatin preparations, especially those containing large chromatin, or even oligonucleosomes, can become turbid, possibly owing to charge neutralization by the combined presence of protein and charged drugs.

39 G. D. Fasman, Methods Cell Biol. 18, 327 (1978). 40 L. Vergani, P. Gavazzo, G. Mascetti, and C. Nicolini, Biochemistry 33, 6578 (1994). 41 M. Eriksson and B. Nord~n, Methods Enzymol. 340, [4] 2001 (this volume).

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Qualitative insight into drug effects on nucleosome cores can be gained considering certain well-determined changes in the CD spectra: 1. Nucleosomes and large chromatin show some differences in their signals in the UV region. The chromatin spectrum presents two positive maxima around 272 and 284 nm, whereas for nucleosomes only the maximum at 272 nm is observed, which resembles the maximum of free DNA. 4° 2. At the 284-nm maximum nucleosomes and chromatin show a molecular ellipticity that is lower than for DNA. 3. In nucleosome core particles the cross-over point is shifted from 258 nm (free DNA) to 264 nm. 4. The nucleosome core particle has a small negative signal around 295 nm. Binding of drugs to nucleosomes can produce changes in the CD spectra in both the ultraviolet and visible regions. The following aspects can be used to infer some characteristics of drug binding to chromatin. 5. The positive 272-nm signal may increase when drug is added. Because the intensity of this signal is sensitive to the DNA winding angle, it could be caused by the DNA unwinding produced by an intercalating drug. 6. No other spectral changes are normally seen in a chromatin-drug complex within the UV region. Because the chromatin, or nucleosome, CD spectra below 240 nm are attributable to the histones, 39 a clearly altered spectrum in this region could be due to displacement of DNA from the histone core. 7. Commonly, DNA-binding drugs lack intrinsic optical activity, but they become optically active when they bind to chromatin (as they do when binding to DNA). If an induced CD signal, (Cotton effect) is observed in the visible region of the drug absorption spectrum it reveals drug binding. 8. The induced CD signal can provide information about the orientation of the drug in the complex, especially when the drug is a known intercalator. For example, the CD signal measured in the visible region might first increase (or decrease) at low drug-to-DNA binding ratios, but it can, thereafter, decrease (or increase) at higher binding ratios to become either negligible or a negative (positive) signal.13"27 These changes in the CD spectra may be used to infer the orientation of an intercalated moiety within a binding site. 42 For some ligands the band sign and the position of the induced maxima can be similar for free DNA and chromatin. 13 Hence, it is assumed that histones do not locally perturb the binding mode of those drugs. In other cases, as for acridine orange, there are indications of different mechanisms of interaction with chromatin and DNA. 12 42E. Tuite and B. Nord6n,J. Am. Chem.

Soc.

116, 7548 (1994).

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Developments

Progress in the analysis of the binding of drugs to chromatin has been significant, but it is still somewhat slow compared with achievements in the analysis of D N A - d r u g interactions. Many useful results have been obtained using the techniques described above, as well as others that are not detailed here. s' 13,28 However, the intrinsic complexity of chromatin organization, and the differences in drug binding to DNA occasioned by the presence of histones as well as the conformational polymorphism of chromatin, lead to difficulties that are not easy to overcome. It has been shown that chromatin structures can modulate the extent of DNA damage in precisely positioned nucleosomes and in a transcriptionally active gene. 43-45 Results of this kind underscore the necessity of addressing the effect of drugs on chromatin experimentally within the more dynamic environment of gene expression. This may require considering the role of chromatin-remodeling complexes. The clear correlation found between changes in chromatin and the development of certain cancers 46 indicates that there is an urgent requirement to develop methods to investigate the effects of antitumor drugs on active chromatin, both in vivo and in vitro. The analysis of dynamic aspects of chromatin, as well as the effects of histone acetylation, 2'3 should foster new studies in the field of chromatin~:lrug interactions.

Acknowledgments Financial support from the Direccion General de Ensefianza Superior e Investigacion Cientifica, Spain (PB98-0469 and PB96-0812), and the Generalitat de Catalunya is acknowledged.

43 L. Yu, I. H. Goldberg, and R C. Dedon, J. Biol. Chem. 269, 4144 (1994). 44 B. L. Smith, G. B. Bauer, and L. K Povirk, J. Biol. Chem. 269, 30587 (1994). 45 j. Wll, J. Xu, and R C. Dedon, Biochemistry 38, 15641 (1999). 46 S. Jacobson and L. Pillus, Curt Opin. Genet. Dev. 9, 175 (1999).

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[26] Locating Cobalt-Binding Sites on DNA Using Restriction Endonucleases By ANGELA M. SNOW and RICHARD D. SHEARDY

Introduction The interactions of simple metal complexes with DNA have been widely studied.l The interaction specificities of these molecules range from simple outside binding, such as observed with [C0(NH3)613+, 2-4 to covalent binding as observed with Pt(NH3)2C12, 5-7 to reactions involving DNA oxidative cleavage through Fenton or related chemistries. 8-j° One important feature of such interactions, especially those leading to covalent binding, is the sequence specificity of the reaction. It has been well established that platination of DNA by Pt(NH3)2C12 is highly sequence specific with a preferential reaction at - G G - sites, although reactions involving - G A - and - G C - sites have also been noted. Reaction at an isolated - G G - or - G A - site results in an intrastrand cross-link composed of a pur(N7)-Pt-pur(N7) linkage, whereas the reaction at a - G C - site results in an interstrand cross-link with the same type of linkage. 5-7 The sequence specificity of these and related reactions is due to the accessibility to N7 of purines, located in the major groove of the DNA, and to the nucleophilicity of N7, particularly that of guanine bases. We have been interested in the interaction specificities of simple Co(Ill) complexes with DNA. Our initial interest stemmed from the ease of the B-to-Z transition of alternating pur-pyr DNA oligomers induced by micromolar concentrations of [C0(NH3)613+. 11 The replacement of one of the ammine ligands by the more labile aquo ligand dramatically altered the interaction specificities of the complex

I T. Tullius,in "MetaI-DNAChemistry" (T. Tullius,ed.). AmericanChemical Society,Washington, D.C., 1989. 2 R. V. Gessner, G. J. Quigley,A. H.-J. Wang, G. A. van der Marel, J. H. van Boom, and A. Rich, Biochemistry 24, 237 (1985). 3 W. H. Braunlin,C. E Anderson,and M. T. Record, Jr., Biochemistr3, 26, 7724 (1087). 4 W. H. Braunlinand Q. Xu, Biopolymers 32, 1703 (1992). 5 S. L. Bruhn,J. H. Toney,and S. J. Lippard, Prog. lnorg. Chem. 38, 477 (1990). 6 W.Brabec, J. Reedjil,and M. Leng,Biochemistry 31, 12307 (1992). 7 p. B. Hopkins,J. T. Millard,J. Woo,M. F. Weidner,J. J. Kirchner,S. T. Sigurdsson,and S. Raucher, Tetrahedron 47, 2475 (1992). 8 D. A. Sigman,D. R. Graham, V. D'Aurora, and M. A. Stern,J. Biol. Chem. 254, 12269 (1979). 9 R. P. Herzberg and P. B. Dervan,J. Am. Chem. Soc. 104, 313 (1984). l0 T. D. Tulliusand B. A. Dombroski,Science 230, 679 (1985). II D. M. Calderone,E. J. Mantilla,M. Hicks, D. H. Huchital,W. R. Murphy,Jr., and R. D. Sheardy, Biochemistry 34, 13841 (1995).

METHODS IN ENZYMOLOGY, VOL. 340

Copyright © 2001 by Academic Press All rights of reproduction in any form reserved. 0076-6879/00 $35.00

520

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES COTAR1 :

5'-ATTAAT-C/TTAAG-ATTAAT-3' Y-TAATTA-GAATT/C-TAATTA-5'

COTAR2:

5'-ATTAAT-GIGATCC-ATTAAT-3' 3'-TAATTA-CCTAG/G-TAATTA-5'

COTAR3:

5'-ATTAAT-A/AGCTT-ATTAAT-3' 3'-TAATTA-TTCGA/A-TAATTA-5'

COTAR4:

5'-ATTAAT-TT/CGAA-ATTAAT-3' 3'-TAATTA-AAGCfrT-TAATTA-5'

[26]

FIG. 1. The DNA oligomers used to assess the sequence specificity of their reaction with [Co(NH3)5(OH2)] 3+ via a thermodynamic approach. 12 Note that each has a unique restriction site possessing at least one G base: COTARI, Aflll (C/TTAAG); COTAR2, BamHI (G/GATCC); COTAR3, HindllI (A/AGCTT); COTAR4, BstbI (TT/CGAA). Further, all have an AsnI (AT/TAAT) site.

with DNA. 11 Whereas [Co(NH3)6] 3+ will induce the reversible B-to-Z transition of [d(5meC-G)4]2, [Co(NH3)5(OH2)] 3+ inhibits the transition. In fact, incubation of that oligomer with [Co(NH3)5(OH2)] 3+ leads to permanent modification of the DNA due to incorporation of cobalt into the DNA. Because of the similarities between [Co(NH3)5(OH2)] 3+ and Pt(NH3)2C12, this modification is most likely due to the formation of a covalent linkage from N7 of guanine to the cobalt center with concomitant loss of water. To assess the sequence specificity, if any, of the reaction of [Co(NH3)5(OH2)] 3+ with DNA, a series of four different model DNA oligomers (Fig. 1) was synthesized and treated with the cobalt complex according to established protocols. Each oligomer is self-complementary and possesses a unique restriction endonuclease site containing one or two G bases. A thermodynamic approach was used to assess the sequence specificity of the interaction. To summarize the results of these studies, each oligomer was incubated with [Co(NH3)5(OH2)] 3+ and then exhaustively dialyzed to remove nonspecifically bound cobalt complex. Each oligomer was then reconstituted in phosphate buffer possessing NaC1 but no cobalt complex. Optical melting curves for both the untreated and cobalt treated oligomers were then carfled out, followed by a comparison of thermodynamic parameters obtained via a van't Hoff analysis. 12 Whereas COTAR1 and COTAR4 were apparently unaffected by the cobalt treatment, COTAR2 was destabilized (lower Tin, lower AHvH of helix formation) and COTAR3 was stabilized (higher Tm, higher AHvH of helix formation) relative to their untreated parents. In addition, the ion dissociation term, An, changed only for COTAR2 and COTAR3 on treatment with the cobalt complex. Finally, atomic absorption studies determined the presence of bound cobalt in treated COTAR2 and 12 M. Hicks, G. Wharton IlI, D. H. Huchital, W. R. Murphy, Jr., and R. D. Sheardy, Biopolymers 42, 549 (1997).

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COTAR3 but none in the other treated oligomers. These studies clearly indicate a robust sequence specificity in the interaction of [Co(NH3)5(OH2)] 3+ with the short synthetic DNA oligomers. Thus, [Co(NH3)5(OH2)] 3+ has a high specificity for isolated - G G - sites (COTAR2), a slight preference f o r - G C - sites (COTAR3), and, apparently, no specificity f o r - G A - sites (as found in COTAR1) or - C G sites (as found in COTAR4) under the particular reaction conditions described. (The details of this work can be found in Hicks et al. J2) The next series of experiments undertaken was to assess the effect of bound cobalt on restriction enzyme digestion. Examination of the sequence of COTAR2 reveals a BamHI site (-G/GATCC-). Digestion of cobalt-modified COTAR2 with BamHI indicated nearly 30% inhibition of the enzyme relative to untreated oligomer (our unpublished results, 2000). Hence the presence of the cobalt bound in the recognition site of the enzyme inhibited enzyme activity. BamHI requires N7 of guanine in its recognitionl3; therefore, blocking N7 with cobalt most likely is the source of inhibition due to diminished enzyme binding. A similar strategy was used to assess sequence specificity in the reaction of [Co(NH3)5(OH2)] 3+ with polymeric DNA. A description of the methodology is the focal point of this chapter. To summarize our approach, pBR322 plasmid is first treated with the cobalt complex via established protocols and then linearized by using AseI, which has a single restriction site on the DNA. The resultant linearized DNA is subsequently subjected to cleavage with a second, different restriction endonuclease. The primary criteria for the choice of that particular nuclease are (1) the presence of a G base at or near its recognition site, and (2) the presence of only one such site on the DNA (for simplicity). Agarose electrophoresis of the treated and untreated DNA follows for quantitation and comparison of the cleavage reactions. The utility of the method is its wide applicability. First, any plasmid of known sequence can be used. The first step of the process is reaction of the plasmid with the DNA-modifying reagent via established protocols. The modified plasmid is linearized by an endonuclease with only one unique site. Cleavage of the linearized, modified plasmid by a second endonuclease follows. The choice of endonuclease to linearize the modified plasmid and endonuclease to subsequently cleave the linearized plasmid depends on the suspected sequence specificity of the reagent used to initially modify the DNA. In our situation, preliminary results indicated that the cobalt complex targeted G bases. Hence, our linearization endonuclease had no G bases in its recognition site whereas the cleavage endonuclease used had at least one G base in or near the recognition site. For comparison, we also used a cleavage endonuclease that did not have a G base in its site. 13 M. Newman, T. Strzelecka, L. E Dorner, I. Schildkraut, and A. K. Aggarwal, Science 269, 656 (1995).

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Experimental Design and Methods

Selection of DNA Plasmid For this approach, any plasmid DNA of known sequence can be used. Because the DNA sequence would be known, a map of the restriction sites would also be available. For this study, we chose pBR322. The pertinent restriction map of pBR322 is shown in Fig. 2.

Selection of Restriction Endonucleases For simplicity, we primarily use restriction endonucleases with only one unique site on the DNA. Table I lists the enzymes used along with their respective recognition sequences. The last enzyme listed (AseI) is used to linearize the DNA. The other enzymes are used to assess the sequence specificity of the cobalt modification reaction. Because the cobalt complex apparently prefers N7 of guanine bases, the selection of these enzymes is premised on the presence of at least one G base at or near the restriction site. As a control, we also selected one enzyme without a G base at or near the restriction site (Dral).

Materials pBR322 plasmid and all restriction enzymes are purchased from New England BioLabs (Beverly, MA). Agarose and ethidium bromide are purchased from Bio-Rad Laboratories (Hercules, CA). [Co(NH3)5OHz](C104)3 is a gift from W. R. Murphy (Seton Hall University, South Orange, NJ). All other chemicals EcoRI a ~

Dra I 3941 ~

Hind HI 29 , 1BamHI

PstI . . ~ - r . . . . . . 3607/-

o.

N

3537{ 4,~61basepa:~s ] D~I ..~ 3249 . ~ 3 2

/ 3

0

I

~

~Pvu II 2064 FIG.2. Map of the pBR322 DNAplasmidshowingrelevantrestriction sites. Note that the site used to linearize (AseI)is well removedfrom the other sites.

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LOCATING COBALT-BINDINGSITES ON DNA

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TABLE I RESTRICTIONENDONUCLEASESANDTHEIRRESPECTIVE RECOGNITIONSITES Enzyme BamHI Pvull HindIII EcoRV DraI

Asel

Recognitionsequencea CCCGTCCTGTG/GATCCTCTACGCCGG GGGCAGGACACCTAG/GAGATGCGGCC AGCTTTACCGCAG/CTGCCTCGCGCGT TCGAAATGGCGTC/GACGGAGCGCGCA TATCATCGATA/AGCTTTAATGCGGTAG ATAGTAGCTATTCGA/AATTACGGATC CCTCTTGCGGGAT/ATCGTCCATTCCG GGAGAACGCCCTA/TAGCAGGTAAGGC CCTAGATCCTTTT/AAATTAAAAATGAA GGATCTAGGAAAA/TTTAAT1T~ACTT AAAATGAAGTTTT/AAATCAATCTAAA TTTTACTTCAAAA/TTTAGAATGATTT ATAGCAGAACTTT/AAAAGTGCTCATC TATCGTCTTGAAA/TTTTCACGAGTAG AT/TAAT TA/ATTA

a The recognitionsite is in boldface and the slash (/) indicatesthe site of strandcleavage.Asel was usedto linearize the plasmid. For the other enzymeslisted, which were usedto monitorthe bindingof cobalt,the 10 base pairs flankingthe recognitionsites are also shown. Note that DraI has three unique sites on the plasmid; all others have only one uniquesite.

are reagent grade. Water for all experiments is deionized and glass distilled before use. Binding o f Cobalt to DNA

Cobalt is bound to DNA by incubation of 20 /zg of pBR322 with 1 mM [Co(NH3)5OH2](C104)3 in 10 mM phosphate buffer, pH 7, containing 50 mM Na + for a m i n i m u m of 48 hr at 37 °. Most reactions are incubated for 85 hr before the DNA is precipitated with ethanol to remove any unreacted cobalt complex. AseI Linearization

Linearization of 5 # g of untreated or cobalt-treated pBR322 plasmid is performed by incubation in 50 m M Tris-HCI, pH 7.9, containing 100 m M NaC1, 10 mM MgCI2, and 1 mM dithiothreitol with the addition of 50 units of AseI (1 unit is the amount of enzyme required to cleave 1/zg of ~. DNA in 1 hr at 37°).

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The reaction sample is incubated for 3 hr at 37 °. The completeness of linearization is checked by 1% (w/v) agarose minigel (50 V, 30 min). PvuH Cleavage Reactions Cleavage reactions are carried out in 10 mM Tris-HCI, pH 7.9, containing 50 mM NaC1, 10 mM MgCI2, and 1 mMdithiothreitol on 0.25/xg of AseI-linearized pBR322 plasmid or AseI-linearized cobalt-modified pBR322 plasmid for 30 min at 37 ° . The amount of enzyme used is that necessary to achieve complete cleavage of the control untreated pBR322 plasmid DNA. Reactions are immediately analyzed by agarose gel electrophoresis. BamHI Cleavage Reactions Cleavage reactions are carried out in 10 M Tri s-HC1, pH 7.9, containing 150 mM NaC1, 10 mM MgC12, 1 mM dithiothreitol, and bovine serum albumin (BSA, 100 #g/ml) on 0.25 /zg of AseI-linearized pBR322 plasmid or AseI-linearized cobalt-modified pBR322 plasmid for 30 rain at 37 °. The amount of enzyme used is that necessary to achieve complete cleavage of the control untreated pBR322 plasmid DNA. Reactions are immediately analyzed by agarose gel electrophoresis. Hindlll Cleavage Reactions Cleavage reactions are carried out in 10 mM Tris-HCl, pH 7.9, containing 50 mMNaC1, 10 mMMgCI2, and 1 mM dithiothreitol on 0.25/zg of AseI-linearized pBR322 plasmid orAseI-linearized cobalt-modified pBR322 plasmid for 30 min at 37 ° . The amount of enzyme used is that necessary to achieve complete cleavage of the control untreated pBR322 plasmid DNA. Reactions are immediately analyzed by agarose gel electrophoresis. DraI Cleavage Reactions Cleavage reactions are carried out in 20 mM Tris-acetate, pH 7.9, containing 50 mM potassium acetate, 10 mM magnesium acetate, and 1 mM dithiothreitol on 0.25 #g ofAseI-linearized pBR322 plasmid or AseI-linearized cobalt-modified pBR322 plasmid for 30 min at 37 °. The amount of enzyme used is that necessary to achieve complete cleavage of the control untreated pBR322 plasmid DNA. Reactions are immediately analyzed by agarose gel electrophoresis. EcoRV Cleavage Reactions Cleavage reactions are carried out in 10 M Tris-HC1, pH 7.9, containing 50 mM NaC1, 10 mM MgC12, 1 mM dithiothreitol, and BSA (100 #g/ml) on 0.25 #g of AseI-linearized pBR322 plasmid or AseI-linearized cobalt-modified pBR322

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plasmid for 30 rain at 37 °. The amount of enzyme used is that necessary to achieve complete cleavage of the control untreated pBR322 plasmid DNA. Reactions are immediately analyzed by agarose gel electrophoresis.

Determination of Amount of Enzyme to Use To determine the minimum amount of enzyme to use to ensure near 100% cleavage of the linearized untreated pBR322, the plasmid is prepared in the corresponding buffer system and titrated with increasing amounts of particular enzyme. Reactions are run for 30 min at 37 ° followed immediately by agarose gel electrophoresis. The concentration of enzyme in the lane indicating 100% cleavage of the DNA is then used for all subsequent cleavage reactions.

Agarose Gel Electrophoresis Horizontal gel electrophoresis is carried out on 1% (w/v) agarose gels prepared with 90 mM Tris-HCl (pH 7), 90 mM boric acid, and 2 mM EDTA containing ethidium bromide (1.5 #g/ml). Gels are run at 60 V for 1.5 hr. The DNA is visualized by ultraviolet light. The gels are photographed with Polaroid 667 film.

Gel Quantitation Agarose gel photographs were quantitated using a Hewlett Packard Scan-Jet 4c, SigmaGel, and Microsoft Excel software. Results and Discussion Figure 3 shows a schematic of the typical agarose gel of the various forms of pBR322. As can be seen, the supercoiled plasmid, the nicked or relaxed plasmid, and the linearized plasmid all have unique mobilities through the gel. Although the pBR322 plasmid purchased had a small fraction of form II, it was used without further purification. On linearization, only the band for form III (i.e., linearized form) was observed. Figure 4A shows a typical gel for the determination of the minimum concentration of PvulI needed to obtain near 100% cleavage of the untreated, linearized pBR322 DNA. As can be seen, 0.35 unit of enzyme per microliter is insufficient for cleavage whereas 0.7 unit (or more) of enzyme per microliter is (compare lane 8 with lanes 4 and 6). Hence, for all subsequent cleavage reactions using PvulI, for both untreated and cobalt-treated pBR322, at least 0.7 unit of enzyme per microliter was used. Similar approaches were used for all other enzymes to determine their minimum concentrations required for 100% cleavage of untreated, linearized pBR322.

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CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

O

Open Circul~ Form II

[26]

Wells II III

Linear Form 1II

•

I

Supercoiled Form I

FlG. 3. Schematic of typical agarose gel of the various forms of pBR322 plasmid. From left to right, lane 1 contains form I (supercoiled) with a little form II (nicked), lane 2 contains form III (linear), lane 3 contains all three forms, and lane 4 contains only form II.

Another feature revealed in Fig. 4A is that even 1.4 units of enzyme per microliter is insufficient to completely cleave the cobalt-treated, linearized pBR322 DNA (compare lane 4 with lane 3). Hence, much higher concentrations of enzyme are required to completely cleave the cobalt-treated pBR322 than are required for the untreated pBR322. A comparison of the cleavage of untreated and

A 1

13 2

3

4

5

6

7

8

1

2

3

4

5

6

7

8

FIG. 4. Typical agarose gel comparing digestion of the linearized, cobalt-treated, and untreated pBR322 DNA polymer. For each gel, the band of lowest mobility is the linearized plasmid; the bands of higher mobilities are for the cleavage products. (A) Determination of minimum concentration of PvulI needed for complete digestion of linearized pBR322. Lane 1, cobalt-treated DNA, no enzyme; lane 2, untreated DNA, no enzyme; lane 3, cobalt-treated DNA, enzyme at 1.4 units//zl; lane 4, untreated DNA, enzyme at 1.4 units/#l; lane 5, cobalt-treated DNA, enzyme at 0.7 unit/p.1; lane 6, untreated DNA, enzyme at 0.7 unit/#l; lane 7, cobalt-treated DNA, enzyme at 0.35 unit//zl; lane 8, untreated DNA, enzyme at 0.35 unit//zl. (B) Comparison of digestion oflinearized, cobalt-treated, and untreated DNA by DraI, EcoRV, and Pvull. Lane 1, cobalt-treated DNA, no enzyme; lane 2, cobalt-treated DNA, Dral at 2.0 units/tzl; lane 3, untreated DNA, DraI at 2.0 units/#l; lane 4, cobalt-treated DNA, EcoRV at 2.0 units//zl; lane 5, untreated DNA, EcoRV at 2.0 units//zl; lane 6, cobalt-treated DNA, Pvull at 1.1 units//zl; lane 7, untreated DNA, PvulI at 1.1 units//~l; lane 8, cobalt-treated DNA, no enzyme.

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LOCATING COBALT-BINDING SITES ON D N A

527

<

z ¢'~ 100 Cq

~

80

•~

60

0

~ 40 go

~

20

~

o Pvu I

Eco RV

Barn HI

Hind III

Dra I

Restriction Endonuclease FIG. 5. Comparison of the cleavage of treated and untreated linearized pBR322 at particular sites.

For eachenzyme,the solid columnrepresents the percentcleavageof untreated DNA and the hatched column represents the percent cleavage of the cobalt-treated DNA. Further, for each enzyme, the comparisonsare madewiththe minimumenzymeconcentrationrequiredfor near 100%ofthe untreated, linearized pBR322.

cobalt-treated DNA by three different enzymes (DraI, EcoRV, and PvulI) can be seen in Fig. 4B. The result here clearly indicates that cleavage of cobalt-treated pBR322 by DraI is not inhibited (compare lane 3 with lane 2), whereas cleavage of cobalt-treated pBR322 by the other enzymes is inhibited. The results for all five enzymes are shown in Fig. 5 and summarized in Table II, As can be seen, the cleavage of cobalt-treated pBR322 is dramatically inhibited for all enzymes except DraI. For each enzyme listed, the percent inhibition listed in Table I1 is the difference in percent cleavage of the cobalt-treated pBR322 relative to the untreated DNA, using the minimum enzyme concentration required to achieve near 100% cleavage of the untreated pBR322. Our previously published results noted above indicated a high sequence specificity of the cobalt complex f o r - G G - and, to a lesser extent, - G C - sitesJ 2 Examination of the cleavage sites for the various enzymes used (Table I) reveals that BamHI has one and EcoRV has two potential - G G - sites/strand whereas PvulI has three and HindlII has one potential - G C - site/strand. However, the DraI site is vacant of either-GG- o r - G C - sites. The data presented in Table II clearly indicate that treatment of pBR322 with the cobalt complex leads to inhibition of enzyme activity only for those enzymes that have - G G - or - G C - sites in or immediately adjacent to their recognition sites. Hence, this study supports our previous results

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CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

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TABLE II ENZYME INHIBITIONSTUDIES Enzyme

PotentiaLcobalt-binding sites/stranda

Percent inhibition h

BamHI Pvull Hindlll EcoRV Dral

- G G - ( 1) -GC (3) - G C - ( 1) - G G - (2) None

24.8 29.7 21.6 26.6 1.3

a These are sites within or near the enzyme recognition site that have the same sequences that bind cobalt as demonstrated in previous studies.12 Each number in parentheses indicates the number of potential cobalt-binding sites/strand within or immediately adjacent to the enzyme recognition site. b Percent inhibition is determined as the difference in percent cleavage of the cobalt-treated pBR322 relative to the untreated DNA, using the minimum enzyme concentration required to obtain near 100% cleavage of the untreated DNA.

with regard to the sequence specificity in the reaction of [Co(NH3)5OH2](CIO4)3 with DNA. Conclusions We have presented here a facile and fast method that can be employed to assess the sequence specificity of a ligand's reaction with DNA. Although the reagent used here covalently binds to DNA, this method should also work for reversible interactions as well, as long as the ligand has a high affinity for the DNA. The advantages of the method are its flexibility and wide applicability. Further, it does not require the use of radioisotopes because the electrophoresis bands are visualized via ethidium bromide fluorescence.

Acknowledgments The authors thank the Research Corporation Partners in Science Program and Bristol-MeyersSquibb (Grant #527179) for support of this project. In addition, we thank Peter Glod for assistance on the project and Prof. Wyatt Murphy of Seton Hall University for the kind donation of [Co(NH3)5OH2](CIO4)3.

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[27] Exploiting Anthracycline Scaffold for Designing DNA-Targeting Agents B y W A L D E M A R PRIEBE, IZABELA FOKT, TERESA PRZEWLOKA, JONATHAN B . CHAIRES, JOSI~ PORTUGAL, a n d JOHN O. TRENT

Introduction Mutations in DNA, the fundamental code of life, cause many diseases including cancer. Thus, the practical value of fully analyzing and deciphering the human genome will lie in applying what is learned to medical needs. For instance, because it is ideally preferable to treat the source of a disease and not its symptoms, it follows that it is better to target the actual DNA responsible for a disease (i.e., the pathogenic genes) and not its downstream effects. One possible approach to controlling the effects of pathogenic genes is to control their expression at the transcription level. The selective inhibition of transcription by the specific targeting of protein-binding sites in DNA became a promising field of research 1-5 when it was discovered that small molecules could control gene expression by interfering with DNA-protein and protein-protein interactions. Thus it is reasonable to expect that high-affinity, sequence-selective small molecule DNA-binding ligands will allow for precise regulation of gene expression by interfering with the binding of specific protein transcription factors to DNA. However, only a small number of high-affinity DNA-binding agents are currently available, and their sequence selectivity is limited. Clearly, there is an urgent need to develop new molecules that are small enough to permeate cells, can bind tightly and selectively to extended sequences of DNA, and can consequently regulate transcription of specific genes. The approach we present here is new and complementary to the current research efforts of other laboratories. A number of approaches to targeting DNA with small molecules have been investigated, including ones involving intercalators, minor groove binders, threading agents, and affinity agents. However, these approaches have two limitations. First, most of them produce molecules that target short (2M bp) sequences (with the obvious exception of the approach of Dervan, 6'7 who has combined N-methylpyrrole derivatives and imidazole moieties to target longer sequences. Second, the current I M. Bellorini, V. Moncollin, M. D'lncalci, N. Mongelli, and R. Mantovani, Nucleic Acids Res. 23, 1657 (1995). 2 A. Vaquero and J. Portugal, Eur. J. Biochem. 251,435 (1998). 3 S.-Y. Chiang, J. C. Azizkhan, and T. A. Beerman, Biochemistry 37, 3109 (1998). 4 L. A. Dickinson, R. J. Gulizia, J. W. Trauger, E. E. Baird, D. E. Mosier, J. M. Gottesfeld, and P. B. Dervan, Proc. Natl. Acad. Sci. U.S.A. 95, 12890 (1998). 5 B. Martfn, A. Vaquero, W. Priebe, and J. Portugal, Nucleic Acids" Res. 27, 3402 (1999).

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approaches tend to focus on one class of ligands at a time and use structure-based design principles to a limited extent. As an alternative, we have developed a new approach to the design of highaffinity sequence-selective DNA binders that can incorporate different classes of ligands. We have named it a "modular design approach" to illustrate our strategy. The whole process involves (1) identifying existing and creating new basic structural fragments (blocks) that display an ability to interact with DNA (for the clarity of the concept these fragments can be treated as the molecular equivalents of Lego blocks), (2) assessing them for their binding affinity and sequence specificity, and (3) assembling the resulting blocks into high-affinity DNA-binding agents. Our approach has several advantages. First, molecular Lego blocks can be identified and their DNA-binding characteristics readily assessed by analyzing in detail the DNA-binding properties of known DNA-binding agents. Second, such well-analyzed molecular blocks can be used to create DNA-binding agents via either a rational structure-based design [molecular modeling, based on nuclear magnetic resonance (NMR) and X-ray crystallographic structures, and readily assembled into tight-binding ligands targeting specific sequences] or a combinatorial approach (preparation of libraries of DNA-binding agents by combinatorial chemistry methods). Because by their very nature anthracyclines (1) bind tightly to DNA and (2) are already modular in having two clearly defined domains (one intercalating and one minor groove binding), our work with anthracyclines has consequently demonstrated the effectiveness and usefulness of the modular design approach. Whereas structure-based design has been an important strategy used to design sequence-specific binding agents, relatively little attention was paid to the energetic and dynamic factors that govern specific binding reactions. In our rational modular design approach, in addition to the structures of related complexes, we take into consideration the thermodynamics and kinetics of complex formation. We have carried out, described below, detailed thermodynamic studies of selected anthracycline antibiotic and assessed energetic contributions to the free energy of binding of individual fragments of anthracyclines. Such well-analyzed fragments can potentially be used as building blocks to assemble novel high-affinity sequence-selective DNA-binding agents. In brief, using our modular approach we have designed, synthesized, and evaluated new high-affinity agents that can bind to specific (up to 6-bp) DNA sequences. Our selection of building blocks, and design of new DNA-binding agents, was based on the analysis of the crystallographic structure of the daunorubicin-DNA complex and energetics of anthracycline interactions with DNA.

6 p. B. Dervan and R. W. Burli, Curr. Opin. Chem. Biol. 3, 688 (1999). 7 C. L. Kielkopf, R. E. Bremer, S. White, J. W. Szewczyk, J. M. Turner, E. E. Baird, E B. Dervan, and D. C. Rees, J. Mol. Biol. 295, 557 (2000).

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Anthracyclines and Their DNA-Binding Properties Anthracyclines emerged in the late 1960s and early 1970s as an important group of anticancer agents. 8-12 Daunorubicin (DNR) and doxorubicin (DOX) were the first anthracyclines approved for clinical use and still are the most effective anticancer agents against leukemias, lymphomas, breast carcinoma, and sarcomas. DOX is especially noted for its broad spectrum of activity, even though the toxic side effects of DOX (e.g., chronic cardiotoxicity and acute myelosuppression) and its clinically approved analogs can limit dose level and the number of drug courses. Although it is generally accepted that the mechanism of action of anthracycline antibiotics is multifactorial, DNA is recognized as their primary cellular target. DOX and DNR bind tightly to DNA, and the fact that the anthracyclines are so clinically important has led to numerous studies that have made anthracyclines one of the most analyzed classes of DNA-binding agents. The magnitude of this work is illustrated by the more than 20 high-resolution structures of anthracycline complexes with DNA oligonucleotides reported to date. DNR and DOX in particular have been studied in great detail for their binding sequence specificity by highresolution footprinting analysis ~3and computational chemistry ~4and been shown to prefer binding to sequences of 5'-(A/T)GC or 5'-(A/T)CG. In addition, detailed thermodynamic and kinetic studies of DNR binding to DNA 15-17 have allowed the approximate DNA-binding affinity of new analogs to be determined. All of these available data have allowed us to assess the potential of anthracyclines as a model of our modular design approach to DNA-binding agents. Figure ! illustrates the structures of DNR and DOX and identifies functional domains. The first domain is a flat anthraquinone ring system that intercalates into DNA; the second domain is daunosamine, a positively charged sugar moiety, that acts as minor groove binder and provides additional favorable polyelectrolyte contribution to the binding free energy; the third domain is formed by substituents on ring A, which by forming hydrogen bonds with DNA bases anchor and stabilize the complex. This is shown clearly by the X-ray crystallographic structure by Wang TM (Fig. 2). 8 F. Arcamone, "Doxorubicin Anticancer Antibiotics." Academic Press, New York, 1981. 9 F. Arcamone, Cancer Res. 45, 5995 (1985). 10 j. W. Lown (ed.), "Anthracycline and Anthracenedione-Based Anticancer Agents." Elsevier Science, Amsterdam, 1988. J l W. Priebe and R. Perez-Soler, Pharmacol. Ther. 60, 215 (1993). 12 W. Priebe, "Anthracycline Antibiotics: Novel Analogues, Methods of Delivery, and Mechanisms of Action." American Chemical Society, Washington, D.C., 1995. J3 j. B. Chaires, J. E. Herrera, and M. J. Waring, Biochemistry 29, 6145 (1990). 14 B. Pullman, Anticancer Drug Des. 7, 95 (1991). 15 H. Fritzsche and H. Berg, Gazz. Chim. ltal. 117, 331 (1987). 16 j. B. Chaires, Biophys. Chem. 35, 191 (1990). 17 j. B. Chaires, W. Priebe, D. E. Graves, and T. G. Burke, J. Am. Chem. Soc. 115, 5360 (1993). 18 A. H.-J. Wang, G. Ughetto, G. J. Quigley, and A. Rich, Biochemistry 26, 1152 (1987).

532

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

[27]

Intercalation ~..

~..

~.H

~

Anchor

and N3 of

H

~

Minor groove binding and AGpe

R = OH - Doxorubicin R = H - Daunorubicin FIG. 1. The functional domainsof DNR and DOX importantfor DNA binding. The current approaches to DNA-binding agent design generally follow the classic rules, such as hydrogen bonding, shape complementarity, and electrostatic interactions, and in most cases ignore binding affinity during the design process. Only recently, however, have we shown that the driving force (free energy) for the binding is not solely based on the classic rules. Therefore, we undertook the assessment of energetic contributions made by specific fragments and substituents of anthracycline antibiotics to DNA binding.17' 19 In brief, we set out to determine the energetic contribution of different fragments of anthracyclines by examining the DNA binding of a carefully selected series of anthracyclines (Fig. 3) by fluorescence titration methods over a wide range of NaC1 concentrations. ~7'19 The results are summarized in Table I. As the data show, DOX displayed higher affinity than any of the other tested anthracyclines, and the binding affinity was clearly influence by the sugar moiety. Daunosamine itself contributed 3.6 to 3.7 kcal mo1-1, or close to 40%, to the total free energy of binding of either DNR or DOX, whereas on the other hand, the decreased DNA binding of the fl-anomer of DOX indicated that daunosamine orientation within the minor groove is critical and that any alteration of such orientation decreases binding. Application of polyelectrolyte theory to these data allowed us to dissect the binding free energy into two parts, the "nonelectrostatic" contribution (~xGt) and the polyelectrolyte contribution (AGej). The former reflects the contribution of such stabilizing interactions as hydrogen bonding and van der Waals interaction, J9j. B. Chaires, S. Satyanarayana,D. Suh, 1. Fokt, T. Przewloka,and W. Priebe, Biochemistry 35, 2O47(1996).

[27]

NOVEL HIGH-AFFINITYDNA-BINDING AGENTS

533

FIG.2. Daunorubicinbinding to DNA.Crystalstructureof two daunorubicin moleculesbinding to d(CGTACG)2.18 whereas the latter reflects primarily the entropic contribution of cation release from the DNA resulting from the binding of a positively charged anthracycline. The "electrostatic contribution" (AGel) to DNA binding of all 3'-aminated analogs ranged from - 2 . 2 to - 2 . 6 kcal mol -l, whereas the AGel of WP159 was - 0 . 4 kcal moi -l. The difference in binding energy between DOX and WP159 ( - 1 . 9 kcal mo1-1) was a direct result of deamination and introduction of hydroxyl at C-3'. The same magnitude of binding energy difference is expected for other pairs of T-amino- and 3'-hydroxyanthracyclines. The contribution of other key structural fragments can be quantitatively evaluated with the data in Table I and is summarized in Table II. The "nonelectrostatic" free energy contribution is shown in Table I for the compounds studied. Differences (AAGt) relative to the reference compound, DOX (Table II), may be interpreted as the energetic contribution of the simple functional groups or larger structural fragments. The hydroxyl groups at C-9 and C-14 each contributed about 1 kcal to the binding free energy. The whole

534

~ CH30

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

CH2R1

O

~

C

H

3

CH300 O~

OH i(p

[9,7]

CH3

•

CH30

o

H NH2 Daunorubicin: FII= H Doxorubicin: I:11= OH

~ CH~O

WP608

CH2OH

'0

~ CH~O

9-Deoxydoxorubicin

~,

CHzOH H

O H OH2 ~o Adriamycinone: R 1= C)H Daunomycinone: RI= H

Hydroxyrubicin: I:11= OH

13Anomer of Doxorubicin

FIG. 3. Structures of anthracycline analogs selected for examination of energetics of DNA-binding energetics (boxes mark alteration in the structure when compared with DNR or DOX).

TABLE 1 ENERGETICS OF ANTHRACYCLINEANTIBIOTICBINDINGTO DNAa

Compound

K/106 (M t)

-AG ° (kcal/mol)

--,~ log K/ 31ogNa +

-AGel (kcal/mol)

-~xGt (kcal/mol)

Doxorubicin Daunorubicin WP608 9-Deoxydoxorubicin Hydroxyrubicin (WP159) fl-Anomer of doxorubicin Adriamycinone (no sugar) Daunomycinone (no sugar)

29,6 11.6 8.0 4.2 0.35 0.11 0.052 0.022

9.98 9.44 9.20 8.86 7.4 6.7 6.3 5.8

0.97 1.08 1.28 0.91 0.18 0.91 0.26 --

2.3 2.6 3.1 2.3 0.4 2.2 0.6 --

7.7 6.8 6.1 6.56 7.0 4.5 5.7 5.2

"Binding constants (K) and standard free energy changes (A Gr~)refer to solution conditions of 6 mM Na2HPO4, 2 mM Na2PO4, 1 mM Na2EDTA, pH 7.0, at 20 °. The electrostatic contribution to the standard free energy change was calculated from the relation AGel = RTS ln[NaC1], where S = (~log K/6 log Na. The thermodynamic free energy change was calculated by the difference AGt = AG ° - AGel.

CH2R I

[27]

535

NOVEL HIGH-AFFINITY D N A - B 1 N D I N G AGENTS TABLE II ENERGETIC COST OF STRUCTURALALTERATIONSIN ANTHRACYCLINEANTIBIOTICSa Compound Doxorubicin Hydroxyrubicin Daunorubicin 9-Deoxydoxorubicin WP608 Adriamycinone Daunomycinone fl-Anomer of doxorubicin

Structural alteration Reference compound 3'-NH2 ---> Y-OH 14-OH --+ 14-H 9-OH --+ 9-H 3'-NH2 ~ Y-OH; 4'-OH ~ 4~-NH2; 14-OH ~ 14-H Sugar removed Sugar removed; 14-OH ---> 14-H Sugar in/3 orientation

A AGt (kcal/mol) 0.0 0.7 0.9 1. I 1.6 2.0 2.5 3.2

a The quantity AAGt refers to the difference in the thermodynamic binding free energy relative to the reference compound DOX. In all cases, AAGt is positive, indicating a less favorable binding interaction resulting from the structural alteration.

daunosamine "nonelectrostatic" free energy contribution was approximately 2 kcal. This substantial contribution to the binding free energy emphasizes the role of the sugar moiety as a tiny DNA groove binder. These studies allowed the molecular determinants of DNA binding of the anthracyclines to be specified in considerable detail and complemented the structural studies of anthracycline-DNA complexes that emerged from X-ray crystallographic studies. 18,20 M o d u l a r D e s i g n of DNA-Binding A g e n t s Using A n t h r a c y c l i n e - B a s e d B u i l d i n g Blocks The major advantages of our modular design approach are (1) its general applicability to different classes of DNA binders, (2) simplification of a structurebased design process, and (3) its compatibility with combinatorial chemistry methods, enabling facile creation of focused libraries of high-affinity sequence-specific small-molecule DNA binders. The potential of small-molecule binders has long been appreciated and there have been attempts to create anthracycline-based bisintercalating molecules2~-23 with expectations that DNA binding can be dramatically increased to the value of 20 A. H.-J. Wang, in "Nucleic Acids and Molecular Biology" (F. Eckstein and D. M. J. Lilley, eds.). Springer-Verlag, Berlin, 1987. 21 D. W. Henry and G. L. Tong, U.S. Patent 4,112,217, September 5, 1978. 22 D. R. Phillips, R. T. C. Brownlee, J. A, Reiss, and P. A. Scourides, Invest. New Drugs 10, 79 (1992). 23 A. Skorobogaty, R. T. C. Brownlee, C. J. Chandler, I. Kyratzis, D. R. Phillips, J. A. Reiss, and H. Trist, AnticancerDrug Des. 3, 41 (1988).

536

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

[27]

FIG.4. Structure of daunorubicin with daunomycinone, an intercalating aglycone, and daunosamine, a minorgroove-bindingsugar, represented by Legoblocks. the square of the binding constant of the monointercalator and that the binding site size of bisintercalator as well as its selectivity will be increased relative to the monointercalator. However, from the chemist's point of view, such attempts were not optimized for success because they focused on exploring chemically accessible linking positions (C- 13 and C- 14) on the side chain of the aglycon, the problem being that the side chain participates in specific interactions with DNA and so any modification at those positions would ultimately reduce the binding affinity of each monomer. In exploring the use of anthracycline scaffolds in the design of high-affinity agents, we have utilized our results of how anthracyclines interact with DNA. In our modular approach, a molecule of DNR can be presented as two blocks-an intercalating aglycon (daunomycinone) and a minor groove-binding sugar (daunosamine)--connected via a glycosidic bond (Fig. 4). Simple combination of an aglycon with a sugar substituent gives 3-bp binding modules, for example, DNR, which preferentially binds to 5'-(AFF)GC or 5'-(A/T)CG sequences. Even though the quantity and categories of blocks and modules can be easily increased and utilized by modular design, we limit our discussion to anthracycline-based blocks and selected linkers and demonstrate their application in two different approaches to high-affinity agents: (1) a rational structure-based design approach and (2) a combinatorial approach. These modular design approaches are exemplified by two compounds: (1) WP631 (structure-based design) and (2) WP760 (combinatorial approach). Rational Structure-Based

Design

Further analysis of the DNR molecule dissected into its two blocks clearly indicated that DNR block constituents can be combined in different ways. Indeed, both blocks can be individually combined with other nonanthracycline types of DNA-binding molecules. Our first attempt demonstrated the possibility of using our structure-based design approach to design agents with a binding specificity of at

[27]

NOVELHIGH-AFFINITYDNA-BINDINGAGENTS

537

least 6 bp. We had already established that one DNR molecule is a 3 bp-recognizing module; therefore the proper linking of two DNR molecules should afford a 6 bpbinding agent (Fig. 5). Information critical to our design was obtained by analysis of the crystallographic structure of a DNR-hexanucleotide (5'-CGTACG)2 complex and from our own studies of how monointercalating anthracyclines interact with DNA. In the complex of (5'-CGTACG)2 and DNR, the two DNR molecules are arranged such that they intercalate at the two CG steps and their groove-binding sugar moieties face each other at the center of the complex (Fig. 2). Clearly, designing a proper linker that would facilitate aglycon intercalation and sugar minor groove binding and its attachment point was of the utmost importance (Fig. 5). Further analysis of the DNR-DNA complex indicated that the DNA template places the T-amines at a distance of 6-7 ,~. Therefore, the simplest way to obtain a molecule that would bind tightly to the 6-bp sequence (5'-CGTACG)2 was to design a linker that would cover the required 6- to 7-,~ distance and fit into the groove. One such potential linker we considered was a p-xylyl molecule. Molecular modeling studies could easily verify whether such a linker can fit into the minor groove while holding both DNRs in their original binding sites. Below we present methods used to model our target molecule. Molecular Modeling

Modeling studies successfully reproduced the NMR and X-ray crystallographic structures of DNR : DNA and DOX : DNA. This partially validates the modeling protocols, which is the first crucial step in the rationalization, prediction, and design of DNA-interactive ligands. Simulations were performed in the isothermal isobaric ensemble (300 K, 1 atm) with the AMBER 5.0 program and AMBER95 force field, using periodic boundary conditions and the particle mesh-Ewald algorithm. Molecular dynamics (MD) simulations used the Sander routine (1.5-fsec time step), with SHAKE to freeze all bonds involving hydrogen. Detailed methods for the simulation of DNA intercalators and groove binders can be found in [14] in this volume. TM The optimization of the linker is a two-step process: the generation of starting models and initial screening, and the fully solvated molecular dynamics simulations. The implicit solvation (GB/SA algorithm) can be used to coarsely screen many linker candidates with several different sequence lengths. Some models will be too strained to be stable. The most stable structures are then used for fully solvated molecular dynamics to determine the relative stabilities. It is important to include explicit solvation effects and counterions, as DNA is a polyanionic molecule, and to more accurately represent the hydrophobic and hydrophilic nature of the ligand. Also, the minor groove hydration may be disrupted (which can be reproduced by the fully solvated molecular dynamics protocol), which should be taken into account in the overall stabilization of the 23aj. O. Trent,Methods Enzymol. 340, [14] 2001 (this volume).

538

[27]

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

©

i

©

X..-

;:m

t'.,,i ~Z e-

•~ ._=

,< Z

+

o

'z,

~9 .,.., o e',

.o

*"

o

o

~D

.-'-~,

~9 •

._

--

~-01

Z

O

b __2E 2

[27]

NOVEL HIGH-AFFINITYDNA-BINDING AGENTS

539

DNA : ligand complex. The same protocols can be used to rationalize experimental results and design new modules and linkers. Combinatorial Approach While there is a great appeal of the rational structure-based design and clearly there has been significant progress in molecular modeling methodology, there are experimental conditions that dictate DNA conformation that are not reproduced at present by molecular modeling. Combinatorial approaches can effectively complement the rational approach. Our modular design approach is perfectly suited for both rational design and combinatorial methodology. In our initial work we chose particular building blocks, which included two intercalating blocks (adriamycinone and daunomycinone), two minor groovebinding sugars (daunosamine and 4'-amino-3'-hydroxy-ol-L-lyxo-hexopyranose), and a range of linkers (Fig. 6). Adriamycinone was selected because the 14-hydroxy group plays an important biological function by distinguishing doxorubicin from daunorubicin. In addition, our studies indicated (Tables I and II) that 14-OH contributes approximately I kcal/mol to the free energy of binding to DNA. Our studies (Tables I and II) also indicated that 4'-amino-3'-hydroxy-ol-L-lyxo-hexopyranose, a sugar used in compound WP608, does not reduce DNA-binding affinity, and thus its contribution to the free energy is similar to that of daunosamine. Molecular modeling indicates that substitution at the 4'-amino group should not obstruct DNA binding of the monomer. Therefore, in addition to the 3'-amine, the 4'-amine can also be used as a convenient attachment point for a linker. Our initial efforts resulted in the parallel synthesis of approximately 65 compounds, and their studies continue. Compounds linked at C-4' appeared to have interesting DNA-binding and biological properties, which are still under evaluation. The first compound from the 4'-linked bisanthracycline family was bisintercalator WP652. Its DNA-binding mode was analyzed by IH NMR, and the structure of its complex with d(TGTACA)2 is shown in Fig. 7. 24 Its interaction with DNA differs from that of WP631, which is able to recognize 6-bp sequences (data for WP631 are shown below), whereas WP652 prefers tetranucleotide sequences. The DNA-binding affinity of WP652 is significantly higher than that of DNR. The energetics of WP652 interactions with DNA are now being studied in detail. One of the possible and immediately available tests to assess the biological potential and selectivity of our newly assembled potential DNA-binding agents is the National Cancer Institute (NCI, Rockville, MD) in vitro disease-oriented primary antitumor screen, which consists of 60 human tumor cell lines. The majority of the compounds we have prepared have been tested in this screen, and the most

24H. Robinson,W. Priebe,J. B. Chaires, and A. H.-J. Wang,Biochemisto'36, 8663 (1997).

540

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

~ CH30

O

CHs

~

C

H

=

O

H

[27]

0•/1•-3•

OH

OH OH

CH~O

Daunomycinone

OH OH

Adriamycinone

HO~~0

O==~H

O

HaN OH Daunosamine

4-Amino-L-lyxo-hexopyrans~

H%O

Ct 2

() HO

0

Br

.o~o Linkers

FIG. 6. Building blocks used for preparation of the first library: daunomycinone, adriamycinone, daunosamine, 4'-amino-L-lyxo-hexopyranose, and linkers (only selected linkers are shown).

WP652 FIG. 7. Chemical structure of WP652 (left) and its complex with d(TGTACA)2 DNA oligonucleotide

(right). 24

[27]

NOVEL HIGH-AFFINITY DNA-BINDING AGENTS

541

WP760

FIG. 8. Chemical structure of WP760 (left) and molecular modeling-derived structure of WP760DNA complex (right).

interesting pattern of antitumor activity we noticed was for a close analog of WP652, which for a change was a doxorubicin-based DNA-binding agent (WP760, NSC 688130) (Fig. 8) connected like WP652 at C-4' rather than at C-3', like WP631. The NCI screen revealed that WP760 is selectively cytotoxic against melanoma cell lines. Results presented in Figs. 9 and 10 (Fig. 9 containing dose-response curves and Fig. 10 containing mean graphs) clearly indicate the selective cytotoxicity of WP760. In the mean graphs (Fig. 10), bars extending to the fight represent sensitivity of the cell line to WP760 in excess of the average sensitivity of all tested lines. Because the bar is logarithmic, a bar 2 units to the right indicates that the compound achieved a response parameter [i.e., median lethal concentration (LCs0)] at 1/100th the mean concentration required over all cell lines, and thus the cell line is unusually sensitive to that compound. Thus, WP760 appeared to be 10-fold to more than 100-fold more cytotoxic against melanoma cell lines than against other cell lines in the NCI panel. These results are even more striking because for most cell lines, the LCs0 values could not determined even at the highest tested concentration (those are proceeded by ">"). This fact is well illustrated by

542

CHEMICAL

AND

MOLECULAR

BIOLOGICAL

APPROACHES

[9.7]

(A) CNS Cancer tO0

I

[

I

I -7

I -6

I -5

50

~o

-50

-lOe

I -8

-9

L o g 10 o f S a m p l e SF-268 SNB-75

SF-295 U251

Concentration

__ ~ _ _ . _ 4,--- -

SF-539

-4

(Molar) _ _ _ A- - -

SNB-19

. . . . . [] . . . . .

(B) Melanoma 1oo

,~..

~

I

I

~,, ,%~,

50

"-.

~o +), \ \

.50

-9

-8

-7 Log l0 of Sample

LOXIMVI SK-MEL-28

0 _ _• __

FIG. 9. Dose-response

MALME-3M SK-MEL-5 curves

__~..._ _. _ ~ _ _

-6

(B).

-5

Concentration MI4 UACC-257

for a typical carcinoma,

and the selective activity in melanoma

...,,

CNS

-4

(Molar) .__.~___ _ .. ~ .. _

cancer,

SK-MEL-2 UACC-62

. . . . . [] . . . . .

with no specific activity (A),

Panel/Cell Line Leukemia CCRF-CEM HL-60(TB) K-562 MOLT-4 RPMI-8226 SR Non-Small Cell Lung Cancer A549/ATCC EKVX HOP-62 HOP-92 NCI-H226 NCI-H23 NCI-H322M NCI-H460 NCI-H522 Colon Cancer COLO 205 HCT-116 HCT-15 HT29 KM 12 SW-620 CNS Cancer SF-268 SF-295 SF-539 SNB-19 SNB-75 U25 ) Melanoma LOX IMVI MALME-3M MI4 SK-MEL-2 SK-MEL-28 SK-MEL-5 UACC-257 UACC-62 Ov~'ian Cancer IGROVI OVCAR-3 OVCAR-4 OVCAR-5 OVCAR-8 SK-OV-3 Renal Cancer 786 0 ACHN CAKI-I RXF 393 SNI2C TK-IO UO-31 Prostate Cancer PC-3 DU-145 Breast Cancer MCF7 NCl/ADR-RES MDA-MB-23 I/ATCC HS 578T MDA-MB-435 MDA-N MG_MID Delta Range

LC50

LOglo LC50 >

m

-4.60

> -4.60 > -4.60 > -4.60 -4.84

m

-460 -460 -4.60 -4.60 -4.60 -6.54 > -4.60 > -4.60 -7.12

m

> -4.60 > -4.60 > -460 > -4.60 > -4.60

m

> > > > >

mm l• /

7. m m

> > > > > >

-4 60 -4.60 -4.60 -4.60 -4.60 -4.60

>

-460 -6.47 -5.95 -686 -6.18 -6.79 -4.71 -6,94

I

-4,60 -4.60 -4.60 -4.60 -4,60 -4.60

ll

> > > > > >

7. I m

/ m I

m

m m

> -4.60 > -4.60 > -4.60 > >

II

-460 -4.60 -4.98

> -4.60 > -4.60 > > > >

-4.60 -460 -4,60 -460

>

-4.60 -4.92 220 2.52 I

I

I

+3

+2

+1

0

I

I

-1

-2

FIG. 10. Excerpt from the NCI Developmental Therapeutics Program mean graphs lbr the 60-eel line screen. LogloLC5o values are shown.

544

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

[27]

the dose-response curves, and it further indicates that in practice the difference in cell line sensitivity can be even higher. In summary, identified compound WP760 displays high activity against melanoma tumors and it is hoped that its use in treating melanomas may not be limited by myelosuppression typically caused by anthracyclines. These promising in vitro data warrant further studies of WP760, aimed at assessing its clinical potential on the one hand and its DNA-binding properties and sequence specificity on the other hand. Our studies clearly demonstrated that the process of randomly assembling even a limited number of DNA-binding molecular blocks can lead to novel DNA-binding agents with unique biological activity and, consequently, validate a modular design of high-affinity DNA ligands by the combinatorial approach.

S y n t h e s i s a n d E v a l u a t i o n of B i s i n t e r c a l a t o r W P 6 3 1

Synthesis and Chemical Characterization of WP631 The DNR molecule contains a primary amino group at C-Y, and this amine can be selectively reacted (e.g., alkylated or acylated) in the presence of other functional groups present on DNR. The most direct approach to creating a p-xylyl link was direct alkylation using commercially available a,a-dibromo-p-xylyl (Fig. 11 ). Reaction gave one main product, which could be purified by on a conventional silica gel column and thin-layer chromatography (TLC). The exact experimental procedures are given below. Experimental Procedure. Daunorubicin hydrochloride (282 rag, 0.5 mmol) is dissolved in a mixture of N,N-dimethylformamide (DMF) and CH2C12 (1 : 1, v/v; 6 ml). Na2CO3 (100 rag) and a,cd-dibromo-p-xylene (68.7 mg, 0.26 mmol) are then added. The reaction mixture is stirred at room temperature for 2028 hr. The progress of the reaction is monitored by TLC [chloroform-methanolNH4OH(aqueous), 86 : 13 : 1, v/v/v). After the reaction is complete, the reaction mixture is diluted with CH2C12 (100 ml) and poured into water (100 ml). The organic layer is separated and washed with water until neutral pH is attained, dried over anhydrous Na2SO4, and evaporated under diminished pressure. The crude product is purified by column chromatography (Silicage160, 230-400 mesh; Merck, Rahway, NJ), eluted with CHCI3, and then eluted with CHC13-methanol at ratios of 98 : 2 and 95 : 5 (v/v). The final product is isolated as the free amine (WP630) and precipitated from CH2C12-hexane to give a red solid (178 mg, 0.153 mmol), yield 61.5%. Elemental analysis for (C62H64N2020 x H20) C, H, N: IH NMR (CDC13) 6:13.95 (s, 2H, 6-OH, ll-OH), 8.01 (d, 2H, JI,2 ----7.7 Hz, HI), 7.77 (dd, 2H, J2.3 = 7.7 Hz, H-2), 7.38 (d, 2H, J3,2 = 8.5 Hz, H-3), 7.18 (s, 4H, p-xylene), 5.50 (d, 2H, Jl'.2' ----3.4 Hz, H-I'), 5.28 (bs, 2H, H-7), 4.66 (bs, 2H, 9-OH), 4.07 (s, 6H, OCH3), 4.05 (q, 2H, J5',6' = 6.25 Hz, H-6'), 3.77 (d,

[27]

NOVEL HIGH-AFFINITY DNA-BINDING AGENTS

545

CH30 O O~ CH3

I 1 CH30 OII OO~

+

~

r

CH2 Br

HD

HCI.HI~

IH 2

Daunorubicin

o

OH WP631

FIG. 11. Synthesis of WP631. 2H, J = 12.78 Hz, CH2 from p-xylene), 3.63 (m, 4H, CH2 from p-xylene and H-4'), 3.22 (d, 2H, Jl0a,10e = 18.8 Hz, H-10e), 2.96 (d, 2H, Jt0a,10e = 18.8 Hz, H-10a), 2.98-2.93 (m, 2H, H-3'), 2.41 (s, 6H, 14-CH3), 2.37 (d, 2H, Jga,8e = 14.7 Hz, H-8e), 2.09 (dd, 2H, J8a,8e = 14.9 Hz, Jv,8a = 4.1 Hz, H-8a), 1.76 (td, 2H, J2'a,2'e ~---Jz'a,3' = 13.0 Hz, J2'a,l' = 3.8 Hz, H-2'a), 1.68 (dd, 2H, J3',2'e = 4.7 Hz, Jz'a,Z'e = 13.0 Hz, H-2'e), 1.37 (d, 6H, J5',6' = 6.65 Hz, H-6'). The free amine WP630 is suspended in methanol (2 ml). Then dry 1 N HC1 in methanol is added (to pH 4), followed by an excess of diethyl ether to precipitate the hydrochloride of WP630. The red solid is washed with ether until neutral pH and then dried to give analytically pure WP631 (170 mg, 0.138 mmol), 55.3% yield calculated from daunorubicin, rap: 160-170 ° dec. [ot]Dz5 : 184.7 ° (c 0,05, CHC13-CH3OH, 1 : 1). Analysis (C62H65N202o x 2HC1 x 4H20) C, H, C1, N. 25j. E Brandts and L.-N. Lin, Biochemistry 29, 6927 (1990).

546

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

[27]

DNA-Binding Properties of WP631 Viscosity Studies: Verification that WP631 Binds to DNA by Bisintercalation. In the absence of high-resolution structural data, hydrodynamic studies, especially viscosity, provide the most reliable means of inferring the binding mode of agents that interact with DNA. 26 The theory of Cohen and Eisenberg 27 predicts that, for monointercalation, plots of the cubed root of the relative viscosity [(rl/rl0) 1/3] versus the binding ratio (bound drug per DNA base pair) ought to have a slope of 1.0. The proved monointercalators ethidium and daunorubicin run as controls in the experiment, and both show viscosity changes in excellent agreement with the theory. For bisintercalators, the slope is expected to be twice that observed for monointercalators, an expectation that has been verified for a variety of bisintercalating compounds. The slope observed for WP631 is nearly double that observed for daunorubicin, consistent with a bisintercalative binding mode (Fig. 12). Ultratight Binding of WP631 to DNA. Traditional spectrophotometric methods for measuring affinity fail for ultratight binding expected for WP631. Reliable alternatives for the accurate determination of binding constants are optical melting studies or differential scanning calorimetry. 25'2s 3o UV melting studies were used to determine the binding constant for the interaction of WP631 with herring sperm DNA in BPE buffer (16 mM total Na÷). In the absence of WP631, the Tm of herring sperm DNA was found to be 67.2 °. In the presence of 10 #M WP631, a concentration sufficient to saturate the DNA lattice, the Tm was elevated to 93.5 °. McGhee 29 has shown that the shift in Tm is a function of the ligand-binding constant and site size. Assuming no interaction of ligand with single-stranded DNA, McGhee derived the equation (1/Tin ° - 1/Tin) = (AHm/R) ln(1 + KL) 1/" where Tm° is the melting temperature of the DNA alone, Tm is the melting temperature in the presence of saturating amounts of ligand, AHm is the enthalpy of DNA melting (per base pair), R is the gas constant, K is the ligand-binding constant at Tin, L is the free ligand concentration (approximated at the Tm by the total ligand concentration), and n is the ligand site size. A value of AHm = 7.0 4- 0.3 kcal mol-1 for herring sperm DNA, determined by separate differential scanning calorimetry experiments, was used. From the experimentally determined increase in Tm observed for WP631, a value of K = 8.8 4- 106M -1 (at 93.5 °) was computed, assuming n = 6 bp. Correction of this value to lower temperatures requires knowledge of the binding enthalpy, which was determined by differential scanning calorimetry. 26D. Suh and J. B. Chaires, Bioorg. Med. Chem.3, 723 (1995). 27G. Cohen and H. Eisenberg,Biopolymers8, 45 (1969). 28D. M, Crothers, Biopolymers10, 2147 (1971). 29j. D. McGhee,Biopolymers15, 1345 (1976). 3oG. Hu, X. Shui, E Leng,W.Priebe,J. B. Chaires,and L. D. Williams,Biochemist~36, 5940 (1997).

[27]

NOVEL HIGH-AFFINITY DNA-BINDING AGENTS

547

1.2 pj OJ

j~ °~4

o~ "t"' A

C:,

1.1

.-.......

1.0

0.0

0.1

0.2

Bound Druglb.p FIG. 12. Viscosity studies of WP631-DNA interaction. The cubed root of the relative viscosity

(q/To) is shown as a function of the ratio of bound drug per DNA base pair: solid circles, ethidium bromide; open diamonds, daunorubicin; solid diamonds, WP63 I. Linear least-squares fits to these data yielded the following slopes: ethidium, 0.84 ± 0.02; daunorubicin, 0.89 ± 0.04; WP631, 1.58 ± 0.08.

Experimental for UV Melting Studies. Ultraviolet DNA melting curves are determined with a Cary 3E UV/visible spectrophotometer (Varian, Palo Alto, CA), equipped with a thermoelectric temperature controller. Sonicated herring sperm DNA at a concentration near 20/zM bp in BPE buffer (2 mM NaH2PO4, 6 mM Na2HPO4, 1 mM NazEDTA, pH 7.0) is used for melting studies. Samples are heated at a rate of 1° min -1, while the absorbance at 260 nm is continuously monitored. Primary data are transferred to the graphics program Origin (Microcal, Northampton, MA) for plotting and analysis. Differential Scanning Calorimetry Determination of Binding Enthalpy. Figure 13 shows the results of differential scanning calorimetry experiments using herring sperm DNA in the presence and absence of saturating amounts of WP631. The area under the curves shown in Fig. 13 provides a model-free estimate of the enthalpy of melting of the DNA alone and of the DNA-WP631 complex. By

548

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES ,

1200

[27]

1

13

._UZ2"

W

W

o I

40

,

50

fl0

,

TO

1

I

I

80

90

100

110

Temperature, oC FIG. 13. Results of differential scanning calorimetry studies of the melting of herring sperm DNA alone (A) or in the presence of near-saturating amounts of WP631 (B). Excess heat capacity (cal mo1-1 °C -I) is plotted as a function of temperature.

Hess's law, these data may be used to determine the enthalpy of WP631 binding to DNA. The equilibria to be considered, along with the experimentally determined enthalpy values, are as follows: Duplex ~- 2(single strand), AHI = 7.0 i 0.3 kcal mol -I WP63 l-Duplex ~ 2(single strand) + WP631, AH2 = 11.44- 1.0 kcal mol -I By combining these two reactions, the binding reaction and its enthalpy may be obtained: WP631-duplex ~ duplex + WP631, AH3 = AH2 - AHI

[27]

NOVEL HIGH-AFFINITYDNA-BINDING AGENTS

549

The enthalpy A//3 needs to be corrected for the amount of WP631 bound to the DNA, and the sign changed, to obtain the binding enthalpy, A Hb = -- A H3/(mol of WP631/mol of bp) From five determinations, a value of AHb = - 3 0 . 2 4-2.6 kcal mol -l was obtained for the association of WP631 with DNA. Complete Thermodynamic Profile for Binding of WP631 to DNA. AHb may be used, assuming that it is constant with temperature, to calculate the binding constant at 20 ° by application of the standard van't Hoff equation, yielding a value of 2.7 × 10 lj M -I . In BPE buffer at 20 °, the binding constant for the interaction of daunorubicin with herring sperm DNA is 1.6 × 107 M-J (E Leng and J. B. Chaires, unpublished data, 2000), so the binding constant of WP631 indeed approaches the value expected for an ideal bisintercalator. The magnitude of the WP631 binding constant approaches that observed for the binding of many regulatory proteins for their specific DNA-binding sites. Knowledge of the binding constant and the binding enthalpy allows us to construct the complete thermodynamic profile for the binding of WP631 to DNA. The free energy is obtained from the standard relation A G ° = - R TIn K, yielding a value of -15.3 kcal mol-] at 20 °. The entropy may be evaluated from the equation - T A S = AG - A H , yielding AS = - 5 1 cal mol-l K-T at 20 °. The thermodynamic profile indicates that the large, favorable binding free energy o f - 15.3 kcal mol ~is derived from the large negative enthalpic contribution of - 3 0 . 2 kcal mo1-1. Binding is opposed by an unfavorable entropic contribution of TAS = - 14.9 kcal tool -I at 20 °. Experimental for Differential Scanning Calorimetry, Differential scanning calorimetry (DSC) experiments utilize a Microcal MC2 instrument along with its DA2 software (July 1986 version) for data acquisition and analysis. Sonicated herring sperm DNA at a concentration of 1 mM bp in BPE buffer is used for all experiments. A scan rate of 1° min J is used. Primary data are corrected by subtraction of a buffer-buffer baseline, normalized to the concentration of DNA base pairs, and further baseline corrected using the Cp(0) software option. Baseline-corrected, normalized data are transferred to Origin graphics software for integration and plotting. Samples for DSC of DNA plus WP631 are prepared by weighing appropriate amounts of solid WP631 and dissolving the solid directly into 2 ml of 1 mM DNA solution. Any undissolved drug is removed by low-speed centrifugation. The exact amount of WP631 bound to the DNA is determined spectrophotometrically.

X-Ray Diffraction and Nuclear Magnetic Resonance Studies of WP631 Complex with DNA Hexanucleotide d( CGATCG)2 The most important evidence that WP631 binds as intended was obtained by X-ray diffraction 3° and NMR 24 analysis of the structure of the WP63 l complex with DNA hexanucleotide d(CGATCG)e (see Fig. 14). Both the X-ray crystal and

550

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[27]

FIG. 14. Stereoscopic view of WP631 :DNA hexanucleotide d(CGATCG)2 complex. 24

NMR structures confirm stable binding to d(CGATCG)2 with little distortion of the optimal DNR intercalation sites. The p-xylyl linker is in a different orientation in the two structures; the crystal structure has it parallel to the groove walls and the NMR structure has it perpendicular to the groove walls. This does not affect the linker distance for the two DNR molecules. Fully solvated molecular dynamics simulations show that both orientations are possible and that the p-xylyl position is dynamic in nature. It should be noted that crystallographic and NMR structures both fit data to an empirical force field and, depending on the resolution and nuclear Overhauser effects (NOEs) defining the ligand interactions, respectively, rely to a greater or lesser degree on the particular force field. B i o c h e m i c a l a n d Biological E f f e c t s of W P 6 3 1 WP631 as Potent Inhibitor of Basal and Spl-Activated Transcription in Vitro

The use of transcription assays in vitro to analyze DNA-drug interactions is considered in detail by Phillips et al. ([23] in this volume). 3] Here, we describe a :~l D, R. Phillips, S. M. Cutts, C. M. Cullinane, and D. M. Crothers, Methods Enzymol. 340, [23] 2001 (this volume).

[27]

NOVEL HIGH-AFFINITY DNA-BINDING AGENTS

551

transcription assay system that allows a direct and quantitative disclosure of the effects of bisanthracycline WP631, as well as any other antitumor drug, on in vitro transcription by RNA polymerase I1.5 The templates used are plasmids containing the adenovirus major late promoter linked to a synthetic DNA fragment that lacks cytidine residues on the transcribed strand (a G-less cassette), which generates transcripts without guanosines. Omission of GTP from the reaction leads to elongation arrest at the end of the cassette. In vitro transcriptions are performed in the absence of GTP; in the presence of RNase T1, which acts as a guanylate-specific ribonuclease; and 3'-O-methyl-GTP, which is added to prevent readthrough transcription of the G-less cassette. Using this assay, the unique RNA that accumulates is the RNase Tl-resistant transcript. 32 We sought to determine the influence of WP631 on an Spl-trans-activated promoter in vitro by comparing its effects with those of an Sp i-lacking promoter. The selective targeting of a transcription factor-binding site might provide a way to interfere with transcription-regulatory processes in vivo. Because the Spl site contains (G + C)-rich tracts, including CpG steps, we foresaw WP631 binding to the same regions, although with different binding affinity. In fact, G-less cassettes are suitable for analyzing almost any DNA-binding d r u g . 4'33'34 WP631 was highly efficient at inhibiting transcription initiation when a plasmid containing an Spl-binding site was used under experimental conditions in which Spl acts as a gene activator. 5 The two plasmids used in the analysis of the effects of WP631 on transcription contained a G-less template. 5 This means that we have been able to distinguish unambiguously between effects that are due to the decrease in transcription initiation and those due to elongation. Presented here is a general protocol that, by taking advantage of the characteristics of a G-less cassette, can be used to analyze any DNA-drug complexes. The utility of the protocol is discussed in the context of WP631 and its powerful activity as an inhibitor of Sp 1-activated transcription.5

DNA Vectors pAdML. The pAdML plasmid, 35 also known as pAdML50[180], contains the strong adenovirus major late promoter linked to an ~ 190-bp G-less cassette. pAdSP01. The pAdSP01 plasmid 5 contains an Spl-binding site. It was constructed by inserting the oligonucleotide 5'-GATTCCGGGGCGGGGCGAATC3', which contains a consensus 5'-(GFF)GGGCGG(G/A)(G/A)(C/T)-3' sequence

32 M. Sawadogo and R. G. Roeder, Proc. Natl. Acad. Sci. U.S.A. 82, 4394 (1985). 33 j. Portugal, B. Martfn, A. Vaquero, N. Ferrer, S. Villamarfn, and W. Priebe, Curt. Med. Chem. 8, 1 (2001). 34 C. Cullinane, S. J. Mazur, J. M. Essigmann, D. R. Phillips, and V. A. Bohr, Biochemist~ 38, 6204 (1999). 35 j. Bernu6s, K. A. Simmen, J. D. Lewis, S. 1. Gunderson, M. Polycarpou-Schwarz, V. Moncollin, J. M. Egly, and I. W. Mattaj, EMBOJ. 12, 3573 (1993).

552

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[27]

for Sp I binding, 36 in the unique EcoRI site located immediately upstream of the AdML promoter in pAdML. Cell Extracts and Spl Protein. One of the better characterized cell extracts for studying transcription of eukaryotic genes in vitro is derived from HeLa nuclei. Because HeLa cells can be produced in large quantities and are relatively free of nucleases they are a good choice for producing extracts that work well in transcription assays. 37 HeLa cell extracts contain some Spl protein, which is difficult to remove (e.g., it cannot be properly immunoprecipitated from nuclear extracts). The presence of Spl should therefore be considered whenever activated transcription is analyzed quantitatively. The problem of the presence of endogenous Spl protein in HeLa cells can be avoided by using Drosophila embryo nuclear cell extracts, which are considered to lack Spl protein. 38 Nevertheless, these extracts are difficult to prepare for routine use in transcription studies in vitro. Another alternative is other cell nuclear extracts which have been used in the analysis of the effects of the drug on transcription. 4 Transcription in Presence of WP631. Basal and activated transcription assays are carried out in disposable microcentrifuge tubes. A control tube (containing no added drug) is used together with others containing different amounts of WP631 (or any other drug to be analyzed). It is important to avoid introducing RNase activity and all buffers must be prepared with nuclease-free water. Moreover, potassium salts should be used instead of sodium salts to optimize the RNA polymerase II performance. The experiments should be done at 30 ° . General Protocol for Basal Transcription Assays. The protocol to be used for assaying basal transcription is as follows. 1. Prepare, at 4 °, in a final volume of 20 #1, a reaction mixture containing 30 mM HEPES-KOH (pH 7.9), 7 mM MgC12, 5 # M ZnSO4, 1 mM dithiothreitol (DTT), 0.2 mM EDTA, 2% (w/v) polyethylene glycol 8000 (Sigma, St. Louis, MO), 400 # M each of ATP and CTP, 1 # M UTP, 1 mM 3'-O-methyl-GTP (Amersham Pharmacia Biotech, Piscataway, NJ) 15 U ofRNase TI (Calbiochem, La Jolla, CA), and 10/zCi of [~-32p]UTP (800 Ci/mmol; Amersham, Arlington Heights, IL). 2. Briefly warm the reaction mixture at 30 ° . 3. Add 1 /zl of a 0.2-/zg//zl solution of supercoiled pAdML plasmid. 4. Add the drug (WP631) solutions to the different reaction tubes to obtain the desired final concentrations. 5. Add 50 ktg (1-2 #1) of HeLa nuclear extract (the exact amount will depend on the activity of each extract; it is advisable to check beforehand) and nucleasefree water such that the final volume is 25 #1. 36 M. R. Briggs, J. T. Kadonaga, S. P. Bell, and R. Tjian, Science 234, 47 (1986). 37 j. D. Dignam, P. L. Martin, B. S. Shastry, and R. G. Roeder, Methods Enzymol. 101,582 (1983). 38 A. J. Courey, D. A. Holtzman, S. R Jackson, and R. Tjian, Cell 59, 827 (1989).

[27]

NOVEL HIGH-AFFINITYDNA-BINDINGAGENTS

553

6. Incubate at 30 ° for 60 min. 7. Stop the reaction by adding 100 #l of TESS [1 mM Tris-HC1 (pH 8.0), 2 mM EDTA, 300 mM sodium acetate (pH 7.5), and 0.5% (w/v) sodium dodecyl sulfate (SDS)]. In all the transcription experiments, an internal control for recovery and gel loading [a radioactive (--,300 cpm) unrelated transcript, larger than 250 bp] is also added at this stage. 8. Immediately digest the samples with proteinase K (0.2 mg/ml) at 60 ° for 15 min and extract the entire reaction with 200 #1 of phenol-chloroform. 9. Transfer the upper aqueous phase to a new tube and precipitate it with 100% ethanol. Wash the pellet with 75% (v/v) ethanol. 10. Resuspend the dried pellet carefully in 8-10 #1 of a loading solution consisting of 95% (v/v) formamide, 10 mM EDTA (pH 7.0), 0.02% (w/v) xylene cyanol, and 0.02% (w/v) bromophenol blue. 1 I. Analyze the transcripts by high-voltage electrophoresis in 90 mM Trisborate and 2 mM EDTA (pH 8.3), using 30-cm-long 8% (w/v) polyacrylamide gels containing 7 M urea. 12. After electrophoresis, soak the gel in distilled water, dry under vacuum, and subject it to autoradiography.

Protocolfor Assaying Spl-Activated Transcription. The protocol to be used for assaying Sp 1-activated transcription is practically the same as described above for the basal transcription assays. However, the pAdSP01 plasmid, which contains an Spl-binding site, is used (see above). In step 5, about 20 ng of pure recombinant Spl protein is also added. Pure protein can be prepared in the laboratory36 or purchased from Promega (Madison, WI). Quantitative analyses of transcripts can be carried out with any densitometer. The relative amounts of transcripts observed should be normalized to the total amount of radioactivity loaded, using the internal control for recovery and gel loading. The C50 values (the drug concentrations that reduce electrophoretic band intensity by 50%) can be derived from plots of percentages of transcription versus drug concentration. These C50 values are then used to compare the effects of WP6315 (or any other drug) on basal and activated transcription and to compare the effects of WP631 (or any other drug) with those of other DNA-binding drugs. The inhibition of transcription from either promoter by increasing concentrations of WP631 is apparently due to the effect on transcription initiation by RNA polymerase II. Because there are no CpG or GpC steps in a G-less cassette, there should be no preferred intercalating sites in the transcribed region that would have an effect during the elongation step. WP631 differentially influences promoter-initiated transcription in vitro (Fig. 15). When transcription was analyzed with the AdML promoter, the C50 for WP631 was 0.48 #M. In contrast, when transcription was analyzed using the

554

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

AdML promoter

AdSP01 promoter

WP631

Control 0.44

1,76

2.2

[27]

WP631

2.64 3.52 !aM Control ÷ Spl 0.02

0.07 0,20

0.50 0.90 tdv

,~_Q, I00"~ "6 z~ O

E

2 s ~ ff

o

%% WP631 [~tM]

0 , ,~

,

~, ' ,7

~

xooOo, % % WP631 [laM]

FIG. 15. Inhibitionof RNApolymeraseII transcriptionin vitroby WP631.Left: Effectof increasing concentrationsof WP631 on basal transcriptionfrom the AdML promoter. Right: Effect of WP631 on Sp 1-activatedtranscriptionfrom the AdSP01 promoter. In both panels, the upper part displaysthe results of electrophoretic gel analysisof the RNA transcripts, while the bottom part corresponds to the results of quantitativeanalysisof the relativeamountof transcription.Note that in the experiments shown, an internal standard for recovery and gel loading was used (see text), but is not shown. The results presented clearly agree with those we have described previously5 and emphasize the strong effect of WP631 on activatedtranscriptionin vitro. AdSP01 promoter, the ability of WP631 to inhibit Sp 1-activated transcription was outstanding and the C50 was in the low nanomolar range (only 60 nM). 5 WP631 was also more efficient at inhibiting activated transcription than basal transcription; at a concentration as low as 0.5 #M, it totally inhibited Sp l-activated transcription in vitro. In comparative experiments performed with daunorubicin, the concentrations of this anthracycline that inhibited the transcription of either promoter were similar. 5'33 The strong effect of WP631 on AdSP01 was fairly distinct (Fig. 15) and consequently rather specific.

Using Gel Retardation and DNase I Footprinting Assays to Complement Information Provided by Transcription Assays in Vitro. More direct evidence of the inhibitory effects of WP631 on Sp 1 binding can be obtained by gel retardation (band shift) and DNase I footprinting.5'33 General protocols of the gel retardation technique are widespread, and the characteristics of the techniques have been discussed in detail elsewhere. 39

[27]

NOVEL HIGH-AFFINITY DNA-BINDING AGENTS

555

Gel retardation experiments are carried out with a labeled Sp 1 oligonucleotide and pure Spl protein. Gel retardation (band-shift) assays using WP631 have been performed in 10 mM Tris-HC1 (pH 7.4) containing 50 mM KC1, 1 mM MgCI2, 0.2 mM EDTA, 0.5 mM DTT, bovine serum albumin (30 #g/ml), and 5% (v/v) glycerol. A typical reaction contains about 20 ng of pure Spl protein and about 2 nmol (in base pairs) of the end-labeled double-stranded oligonucleotide 5'GAATTCGGGGCGGGGCGAATTC-3' in the presence of 1 #g of poly[d(I-C)] (Boehringer Mannheim, Indianapolis, IN). Thereafter, the samples are analyzed on nondenaturing 4.5% (w/v) polyacrylamide gels containing 45 mM Tris-borate and 1 mM EDTA (pH 8.3). The gels are run at low voltage (12 V/cm), dried under vacuum, and subjected to autoradiography. The results obtained with WP631 attest that this bisanthracycline might block the binding of Spl to its putative binding site with high efficiency.33 Drug-mediated inhibition of the interaction between Spl and the oligonucleotide shows the destabilizing effect of WP631 on Spl-DNA complexes (it is also possible to use longer DNA fragments containing an Sp 1 site). In the presence of decreasing concentrations of WP631, the formation and stability of the S p l DNA complexes was more apparent, indicating that the DNA-protein complex was sensitive to the bisanthracycline in a concentration-dependent way. DNase I footprinting assays of the binding of WP631 and the Sp I protein to the same sequence can be performed with the pAdSP01 plasmid, which contains a putative Spl-binding site. A 352-bp DNA fragment purified from pAdSp01 can be labeled at the 5' end of the upper strand with [y-32p]ATP and T4 polynucleotide kinase. 5 General aspects of a footprinting assay and details of its use in DNA-drug interactions are considered in [20] in this volume. 4° In a standard experiment, samples containing 3000 cpm of an end-labeled DNA fragment (about 10 pmol in base pairs), along with different concentrations of WP631 and/or of pure Spl protein, are supplemented with a buffer consisting of 10 mM Tris-HC! (pH 7.4), 50 mM KC1, 2 mM MgC12, 2 mM MnC12, 0.5 mM DTT, and 5% (v/v) glycerol to a final volume of 20/zl and then digested with DNase I (Boehringer Mannheim) at a final concentration of 0.01 U/ml for 2 min at 30 °. The reaction mixture is phenol extracted and ethanol precipitated, and the resulting pellet is dissolved in 85% (v/v) formamide, 10 mM EDTA, and 0.02% (w/v) bromphenol blue. Samples are heated at 95 ° for 2 min prior to electrophoresis. Footprints are resolved by high-voltage electrophoresis in 90 mM Tris-borate and 2 mM EDTA (pH 8.3), using 6% (w/v) polyacrylamide gels containing 8 M urea. Our footprinting studies performed in this way have revealed that a ternary complex is formed with some concentrations of Spl and WP631 on the Splbinding site of the AdSP01 promoter. 33 39 j. B. Taylor, A. J. Ackroyd, and S. E. Halford, Methods Mol. Biol. 30, 263 (1994). 4o K. R. Fox and M. J. Waring, Methods Enzymol. 340, [20] 2001 (this volume).

556

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[28] D e s i g n , S y n t h e s i s , and of Polyintercalating

[28]

Characterization Ligands

By VLADIMIR M. GUELEV, M A R K S. CUBBERLEY, MEREDITH M. M U R R , R . SCOTT LOKEY,

and

BRENT L. IVERSON

Introduction Functionalized DNA polyintercalators that bind to DNA in a sequence-selective manner 1-3 represent, at least in principle, an attractive strategy for targeting cellular DNA. As such, polyintercalation could be considered an alternative to approaches such as triple helix-forming oligonucleotides,4,5 minor groove-binding molecules,6 peptide nucleic acids,7 and modified zinc fingers/Indeed, the idea of connecting several intercalating groups via appropriate linkages to create DNAbinding agents with high affinity is certainly not new.9-13 Synthetic trisintercalators have been reported, most notably a polyamine-linked trisacridine compound synthesized by Laug~a et al., 13 that binds to DNA with an affinity n e a r 1014 M -1. Despite noteworthy successes such as this, attempts to extend interactions beyond bisintercalation have often failed to significantly improve binding affinities. In addition, the key issue of sequence specificity has hardly been addressed with polyintercalating systems. Design Considerations There are potential difficulties associated with the design of polyintercalators. As noted above, linking multiple intercalating moieties together does not necessarily result in a corresponding increase in binding constant or even in full 1 j. Chaires, Curt Opin. Strucr Biol. 8, 314 (1998). 2 L. Hurley and J. Chaires (eds.), "Advances in DNA-Sequence Specific Agents," Vol. 2, JAI Press, Greenwich, Connecticut, 1996. 3 E Nielsen, Chem. Eur. J. 3, 505 (1997). 4 D. Gowers and K. Fox, Nucleic Acids Res. 27, 1569 (1999). 5 j. Franqois, J. C. Lacoste, L. Lacroix, and J. L. Mergny, Methods Enzymol. 313, 74 (2000). 6 S. White, J. Szewczyk, J. Turner, E. Baird, and E Dervan, Nature (London) 391, 468 (1998). 7 M. Egholm, O. Buchardt, L. Christensen, C. Behrens, S. Freier, D. Driver, R. Berg, S. Kim, B. Norden, and P. Nielsen, Nature (Londan) 365, 566 (1993). 8 A. Klug, J. Mol. Biol. 293, 215 (1999). 9 L. Wakelin, Med. Res. Rev. 6, 275 (1986). 10 G. Atwell, W. Leupin, S. Twigden, and W. Denny, J. Am. Chem. Soc. 105, 2913 (1983). II j. Hansen, T. Thomsen, and O. Buchardt, J. Chem. Soc. Chem. Commun, p. 1015 (1983). 12 W. Denny, G. Atwell, G. Willmott, and L. Wakelin, Biophys. Chem. 22, 17 (1985). 13 p. Laugfia, J. Markovits, A. Delbarre, J. Le Pecq, and B. Roques, Biochemistry 24, 5567 (1985).

METHODS1NENZYMOLOGY,VOL.340

Copyright~ 2001 by AcademicPress All rightsof reproductionin any formreserved. 0076-6879/(X)$35.00

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557

intercalation. Likely explanations include the entropic cost associated with immobilizing a long flexible molecule on the DNA and/or self-stacking of the adjacent aromatic units in a polyintercalator in the absence of DNA. In addition, there are many examples of naturally occurring and synthetic DNA mono- and bisintercalators 1-3,9 that are well studied, but there are no known examples of trisor tetra-, etc., polyintercalators found in nature on which to base synthetic designs. The pursuit of predictable sequence specificity in a high-affinity polyintercalating system will require a detailed structural understanding, but unfortunately the characterization of DNA-bound structure is complicated in the case of polyintercalators. For example, a presumed polyintercalator can potentially have multiple binding modes; accordingly threading versus nonthreading intercalation (see below) and even groove binding should be considered.14 In addition, if polyintercalation is confirmed, then the linker could be placed in the major or minor grooves and there could be various numbers of base pairs between consecutive intercalated aromatic units. Finally, and perhaps most challenging, polyintercalation involves considerable distortion of the DNA helix, the structural and energetic consequences of which must be taken into account in the design of sequence-specific agents. ~5,~6 On the basis of the above considerations, a viable DNA-readout polyintercalator system should be modular, so that the design of the longer polyintercalator analogs can be based on detailed studies of the shorter modules, that is, dimers. Ideally, the linkers will be held deep in one or both grooves of DNA, facilitating sequence-specific interactions. A viable polyintercalating system also needs to be easily synthesized, including the ability to create numerous derivatives to provide for various sequence specificities. Finally, the system needs to be readily characterized structurally when bound to DNA, presumably via multidimensional nuclear magnetic resonance (NMR). Although initial polyintercalator designs were based on attaching several chromophores as side chains of a linear linker, 9-13 Takenaka et aL constructed a trisintercalator with chromophores attached in a head-to-tail fashion.~7 On the basis of the ability of the central intercalator to "thread through" DNA and access both DNA grooves simultaneously, a threading binding mode was proposed for this molecule, in which the linker alternates between the major and minor grooves of the DNA helix.IV' 18 Aside from purely aesthetic appeal, such a binding mode may have several advantages pertaining to biological activity, such as simultaneous blockade of both DNA grooves, improved sequence specificity, and slower dissociation rates from cellular DNA.

14 D. Suh and J. Chaires, Bioorg. Med. Chem. 3, 723 (1995). 15 E Gago, Methods 14, 277 (1998). J6 j. Chaires, Biopolymers 44, 201 (1997). 17 S. Takenaka, S. Nishira, K. Tahara, H. Kondo, and M. Takagi, Supramol. Chem. 2, 41 (1993). J8 C. Lamberson, Ph.D. Thesis, pp. 23-26. University of Illinois, Chicago, Illinois, 1991.

558

Ht~3N

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

N"A'~-"

,~.~N

2H

4 H

~

O

[9,8]

H

O

n=0-8

NDI amino acid FiG. 1. General structure of the polyintercalators, synthesized by standard SPPS techniques. When specifying polyintercalator composition in text, the 1,4,5,8-naphthalenetetracarboxylic diimide amino acid is referred to as NDI.

1,4,5,8-Naphthatenetetracarboxylic diimide (NDI) derivatives have been shown to monointercalate DNA in a threading manner. 19 They are chemically robust and are easily derivatized into amino acids in which the amino and carboxyl termini are on opposite sides of the molecule (Fig. 1). These features make the NDI unit particularly well suited to be a building block for the construction, via solid-phase peptide synthesis (SPPS) methodology, of modular polyintercalating molecules with a putative threading mode of DNA binding. With the above considerations in mind, we devised a series of polyintercalators in which a number of intercalating NDI units are strung together in a head-to-tail fashion (Fig. 1). 20 The compounds are synthesized by standard 9-fluorenylmethoxycarbonyl (Fmoc)-based solid-phase methods, 21,22 which readily allows for tremendous variation in the linker composition as well as the facile incorporation of even large numbers of intercalating aromatic units. Initially, we reported the first DNA tetraintercalator based on chains of NDI units linked via Gly-Gly-Gly-Lys peptides, 2° and more recently we have expanded the number of intercalating cbromophores to eight units, 23 which appear to fully 19 E Tanious, S. Yen, and W. D. Wilson, Biochemisto' 30, 1813 (1991). 2o R. S. Lokey, Y. Kwok, V. Guelev, C. Pursell, L. Hurley, and B. Iverson, J. Am. Chem. Soc. 119, 7202 (1997). 1 E. Atherton and R. C. Sheppard, "Solid Phase Peptide Synthesis--A Practical Approach." IRL Press, Oxford, (1989). 22 NovaBiochem, "NovaBiochem Catalog & Peptide Synthesis Handbook." NovaBiochem, La Jolla, California, 1999. 23 M. Murr, M. A. Thesis. University of Texas, Austin, Texas, 1999.

[28]

POLYINTERCALATING L|GANDS H2N~

NBOC

TsO HaN O ~ "O" "O

559

Statistical Mixture: RI= R2 = CH2CH2CO2Bz RI= CH2CH2CO2Bz R2 = CH2CH2NBOC RI= R2 = CH2CH2NBOC

OBz

DIEA, i PrOH 94%

O" "N" "O

1 H2, Pd/C 36%

1.) TFA/CH2CI 2

/\

2.) FMOC-AA-OPfp, .OBT, O" "N" "O

/

O

NH

2,6-1utidine, NMP 90 %

O~ "N" "O

/

/

NBOC

NFMOC

2

2a

R~[~NFMOC

3

SCHEME I. Synthetic scheme for the generation of the NDI-amino acid 3, used in the solid-phase synthesis. Although amino acid 2a would be a more convenient building block in the synthesis, we have been unable to isolate this derivative in high yield (see text).

intercalate on DNA binding according to viscometry and UV measurements. We have also used combinatorial methods to isolate a linker that causes a dramatic change in DNA sequence specificity.24 Below we describe a general protocol for the synthesis of NDI-based polyintercalators of up to eight chromophores, as well as the synthesis of mixtures of bisintercalators for library studies. S y n t h e s i s of 1 , 4 , 5 , 8 - N a p h t h a l e n e t e t r a c a r b o x y l i c Diimide C h r o m o p h o r e - A m i n o Acid (3) Scheme 1 (1-3) describes the synthesis of the diimide amino acid 3. Ideally, it would be preferable to eliminate any in-solution couplings, by starting with a protected amino acid such as 2a. Unfortunately, this derivative was found to be highly unstable, having a propensity to spontaneously lose the Fmoc-protecting 24 g. Guelev, M. Harting, R. S. Lokey, and B. Iverson, Chem. Biol. 7, l (2000).

560

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

[28[

group. The mechanism of this decomposition reaction has not yet been investigated. Therefore, the solid-phase synthesis starts with a diimide-amino acid adduct 3, prepared in solution as follows.

N-( 2-te rt-Buto xycarbonylaminoethyl )-N'-( 2-carbo xyethyl )1,4,5,8-naphthalenetetracarboxylic Diimide (2) 1,4,5,8-Naphthalenetetracarboxylic dianhydride (1) (8.96 g, 33.4 retool) is combined with mono-tert-butoxycarbonylaminoethylamine (5.34 g, 33.4 mmol) and fl-alanine benzyl ester p-toluene sulfonate salt (11.72 g, 33.4 mmol) in 2propanol (450 ml). N,N-Diisopropylethylamine (6.4 ml, 37 mmol) is added to the suspension and the mixture is heated at reflux for 19 hr under an inert atmosphere. The mixture is allowed to cool to room temperature and the suspended solid is collected by filtration, washed thoroughly with diethyl ether, and dried under high vacuum. The statistical mixture of diimide products (17.9 g, 94%) is used directly in the next step without further purification. The above mixture of diimide products is suspended in absolute ethanol (250 ml) and purged with argon. Pd/C (9 g, 10%) is added and the mixture is charged with H2. After 48 hr, the mixture is concentrated to 50 ml (bath temperature 30 °, 20 torr) and diluted with 15% (v/v) triethylamine (TEA)-CHzCI2 (250 ml). The suspension is filtered through Celite, which is rinsed with 10% (v/v) TEA-CHzCI2 until the filtrate is colorless. The filtrate is concentrated to dryness and the resulting solid is loaded onto a large column of silica equilibrated with 10% (v/v) TEA-CH2CI2. Flash chromatography using a gradient of 0 --+ 5% (v/v) methanol in 10% (v/v) TEA-CH2CI2 yields the TEA salt of the title compound as the second yellow band. The product fractions are concentrated to dryness and the resulting solid is dissolved in a minimal amount of 10% (v/v) methanol-CHzCl2. Acetic acid (20 ml) is added followed by the slow addition of hexanes (300 ml). The resulting solid is collected by filtration, triturated several times in methanol to remove residual acetic acid, and dried in vaeuo to yield the title compound 2 as a white powder (5.5 g, 73% based on statistical mixture).

N-2-(N ~-9-Fluorenylmethoxycarbonyl-N~-tert-butoxycarbonyl)lysylaminoethyl N'-( 2-carboxyethyl )- l, 4,5,8-naphthalenetetracarboxylic Diimide (3) The following procedure works for the preparation of most Fmoc-protected amino acids, except for Fmoc-glycine, for which a different workup is necessary because of poor solubility (see below). The t-butyloxycarbonyl (Boc)-protected NDI amino acid 2 (1.5 g, 3.1 mmol) is suspended in CH2C12 (10 ml) and trifluoroacetic acid (TFA, 10 ml) is slowly added. After standing for 15 min, the solvent is evaporated and residual TFA is removed by azeotropic evaporation from heptane (twice). The resulting residue is dried in vacuo for 4 hr. The TFA salt is then

[28]

POLYINTERCALATING LIGANDS

561

combined with N-9-fluorenylmethoxycarbonyl-(N~-tert-butoxycarbonyl)lysine (Fmoc-Lys-OH, 2.2 g, 3.2 mmol) and 1-hydroxybenzotriazole (HOBO (0.48 g, 3.1 mmol) in N-methylpyrolidone (NMP, 10 ml). 2,6-Lutidine (1 ml, 6 mmol) is added and the suspension is stirred until clear (approximately 2 hr). Adduct Workup of Lysine and All Other Amino Acids The reaction mixture is diluted with ethyl acetate ( 1000 ml) and quickly washed with 0.1 N HC1 (3 x 100 ml). The organic solution is reduced in volume to "-~500ml and 100 ml ofhexane is added. The resulting solid is collected by filtration and dried in vacuo over KOH to yield the title compound 3 as a yellow powder (2.35 g, 90%). Glycine Adduct Workup The reaction mixture is poured into 200 ml of citrate buffer (0.2 M, pH 4) with stirring and filtered. The filtered material is washed with H20 and dried in vacuo. The dry solid is ground to a fine powder, triturated with diethyl ether and with ethyl acetate several times, and dried in vacuo. Solid-Phase Synthesis Coupling/Deprotection Cycle Reagents Needed Fmoc-amino acid-NovaSyn TGA resin N-methylpyrrolidinone (NMP) 2-propanol dimethyl sulfoxide (DMSO) piperidine (20%, v/v) in NMP N-methylmorpholine (NMM) (Benzotriazol-l-yloxy)tris(pyrrolidino)phosphonium hexafluorophosphate (PyBOP) Fmoc-amino acid-NDI adduct 3 Fmoc-protected amino acids Equipment Needed Solid-phase peptide synthesis (SPPS) reaction vessel with coarse frit filter (freshly silanized) Rotary shaker The Fmoc-amino acid-NovaSyn TGA resin (1 equivalent) is added to the solidphase reactor followed by enough NMP to cover the resin. The resin is allowed to swell for 15-20 min.

562

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

[28]

Deprotection and Wash. The NMP is drained from the resin and piperidineNMP (10 ml/g; 20%, v/v) is added. The mixture is shaken for 15 min before the solvent mixture is drained from the resin and a second portion of piperidine-NMP is added. After 15 min, the solvent mixture is removed and the resin is washed with NMP (three times), 2-propanol (three times), and NMP (three times) in preparation for acylation. Coupling. The Fmoc-amino acid-NDI adduct 3 (3 equivalents) and PyBOP (3 equivalents) are completely dissolved in NMP (sonicate if necessary) to make a 0.2-0.4 M solution. The solution and NMM (6 equivalents) are added to the resin and the mixture is shaken for 45 min. The solvent is drained and the resin is washed with NMP, after which the coupling is repeated. The resin is washed twice with DMSO (to remove all unreacted NDI amino acid) and followed by the usual NMP-2-propanol-NMP wash sequence. Capping. The unreacted amino termini are capped with 5% (v/v) acetic anhydride-5% (v/v) 2,6-1utidine in NMP (30 min). The wash/deprotection/wash/acylation cycle is continued until the desired sequence is achieved. Double couplings and additional DMSO washes are necessary only for the NDI amino acid. Cleavage from Resin and Side Chain Deprotection Reagents and Equipment Needed Glacial acetic acid Dichloromethane (CH2C12) Methanol Diethyl ether Trifluoroacetic acid (TFA) : water mixture (95 : 5, v/v) If Trp, Met, Tyr, Cys, and Phe are used in the synthesis, also needed are phenol, anisole, and 1,2-ethanedithiol (EDT) as cation scavengers.

Equipment Needed Conical flask (25 ml) Plastic centrifuge tube (50 ml) Centrifuge

Procedure. When the desired sequence is achieved, the terminal Fmoc group is removed as described above. The resin is then washed with NMP (three times), acetic acid and CH2C12 (three times), and methanol (three times) before the resin is dried over KOH under high vacuum for several hours. The TFA cleavage mixture (20 mug) is added and the solution is mixed gently. After 1 hr the TFA

[9.8]

POLYINTERCALATING LIGANDS

563

solution is drained into the 25-ml conical flask and the resin is washed with undiluted TFA (two times). The TFA solution is concentrated and added dropwise to cold diethyl ether (50 ml/g) in a 50-ml centrifuge tube. The solution is allowed to precipitate for 12 hr at - 2 0 °. The precipitate is collected by centrifugation at 2000 rpm at 0 ° for 10 min. The ether is removed by decantation and additional ether is added. The suspension is triturated for several minutes and the precipitate collected as before. The trituration is repeated and the collected precipitate is partially dried under reduced pressure to remove any residual ether. Caution: Drying too long may make the compounds difficult to dissolve. The precipitate is dissolved in ~ 5 - 8 ml of water, filtered, and purified as described below. Alternatively, the solid is extracted with diethyl ether and the residual ether is removed from the aqueous layer by rotary evaporation (at room temperature) prior to purification. No~s 1. The Kaiser test for free amino termini does not give reliable results once

the first NDI residue is introduced. 2. Our experience shows that the NDI-amino acid coupling is by far the most difficult. The problem may be due to aggregation of the diimide chromophores. a. TentaGel-type resins (Rapp Polymere, Tiibingen, Germany) work somewhat better than regular polystyrene/divinylbenzene (DVB) resins. b. Double couplings of--~45 min (with PyBOP as the coupling reagent) are required to achieve coupling efficiency >90%. Capping after the first coupling with acetic anhydride ensures lack of nondiimide by-products that can go undetected. c. N-Methylpyrrolidinone (NMP) gives better results as a solvent than N,Ndimethylformamide (DMF) or N,N-dimethylacetamide (DMA). An additional DMSO wash after the couplings of the NDI amino acid removes any residual excess reagent. 3. On the basis of analysis of library mixtures, we have not detected any other problematic coupling steps or specific "difficult sequences. ''22 However, coupling efficiency and the resulting product purity deteriorate rapidly with increasing number of NDI units. Whereas product purity is "~90% for the dimer analogs (Fig. 2), it was only "-~10% for the octamer. 4. N-Fmoc-protected amino acids (or oligopeptides), when not commercially available, can be readily prepared from the unprotected material, using 9-fluorenylmethyloxycarbonyl-N-hydroxysuccinimide(Fmoc-OSu), according to the procedure of Lapatsanis et al. 25 25L. Lapatsanis,G. Milias, K. Froussios,and M. Kolovos,Synthesis, p. 671 (1983).

564

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES a

[28]

1,2 3,4

.

.

.

.

.

.

250

.

.

.

.

,

.

,

.

.

300 350 Wavelength (nm)

.

.

,

400 3

5

10 15 Time (min)

20

25

FIG.2. HPLC analysisof the crude productfrom the synthesisof the bisintercalatorLys-NDI-GlyGly-Gly-Lys-NDI-Gly.Gradient:85% water [0.1% (v/v)TFA] ~ 60% CH3CN[0.1% (v/v)TFA]over 60 min at 0.75 ml/min.Usingthe UV spectra (a) the four majorproducts seen in the chromatogram(b) can be differentiatedas containingone NDI (peaks 1 and 2) and two NDI (peaks 3 and 4) units. Mass spectral analysisidentifiedpeak 3 as the desired product. Peaks 1 and 2 are due to monomericNDI by-products and peak 4 (mass 14 units greater than the major product) is thought to be an N-oxide, formed by oxidationof a lysine side chain.

5. If desired, the number of coupling steps can be reduced by using short peptide segments, for example, Fmoc-N-Gly-Gly-Gly-OH (see Note 1).

P u r i f i c a t i o n b y P r e p a r a t i v e C18 H i g h - P e r f o r m a n c e Liquid Chromatography

Reagents Needed High-performance liquid chromatography (HPLC)-grade acetonitrile Distilled water Trifluoroacetic acid (TFA)

Solvent Systems Solvent A: 0.1% TFA-CH3CN (v/v) Solvent B: 0.1% TFA-water (v/v)

Procedure Typically, a gradient of 100% solvent B -+ 0% solvent B over 240 min at 0.75 ml/min yields good results. Holding the gradient (e.g., at 85% solvent B for dimeric derivatives) is sometimes necessary to resolve especially impure mixtures.

[28]

POLYINTERCALATING LIGANDS NH3

HaN

+NH 3

N~ 0

565

N~

X Y Zd

"~-E "0

H 0

N---if.O (~ ~'~--~' ~

H 0

x,z-{

Skip

.

o

y =

H 0

0

H 0

H 0

FIG. 3. General structure of a 360-member library of bisintercalators synthesized to study the effect of amino acid linker on DNA sequence specificity} 4

Library Synthesis Figure 3 shows the composition of a 360-member library synthesized as described below} 4 Equipment. Synthesis is performed on a parallel synthesizer that consists of 30 open sintered glass filter tubes attached to a Teflon block, equipped on the bottom with a vacuum outlet/argon inlet. The reactor is mounted on a rotary shaker. Note: Coupling conditions are the same as described above for the single compounds.

Procedure The NDI-amino acid coupling is performed in one vessel; prior to deprotection the resin is divided by suspending it in NMP in a graduated cylinder and, while stirring, pipetting out equal amounts into the separate reaction vessels. Deprotection is performed in parallel, with a different amino acid added to each vessel. The resin is then recombined and redivided, and deprotection/coupling is repeated the desired number of times. To increase the number of compounds in a mixture, we have added two amino acids per coupling reaction. To minimize differences in

566

CHEMICAL AND MOLECULARBIOLOGICALAPPROACHES

[28]

reactivity, we group the amino acids, separating primary from secondary amino acids. Half an equivalent of amino acid is used to allow for shorter peptide linkage ("skip" residue). The mixtures are cleaved with approximately 0.5 ml of TFA-H20 (95 : 5, v/v) per 50 mg of resin (adding more TFA would prevent precipitation). The solution is collected in 15-ml centrifuge tubes and the resin is washed with a minimal amount of TFA (~0.5 ml). The product is precipitated with 10 ml of diethyl ether and centrifuged, and the ether is decanted. This step is repeated two more times, the product is then extracted between water and diethyl ether, and the aqueous layer is loaded on a Waters (Milford, MA) Sep-Pak cartridge, washed with several volumes of 0.1% (v/v) aqueous TFA, eluted with a 70 : 30 mixture of water-acetonitrile [0.1% (v/v) TFA each], and lyophilized. Note: If a given mixture does not precipitate on treatment with ether, the procedure described above for the single compounds should be used. Alternatively, the product is extracted with water, neutralized with triethylamine and lyophilized, and then loaded on a Sep-Pak column and purified as described above. Characterization Because of the acetic anhydride capping of the resin after the first NDI coupling, all impurities in the crude product contain at least one NDI moiety that can be used as a diagnostic chromophore to facilitate characterization.

High-Performance Liquid Chromatography Efficient separation of the crude product or of library mixtures of derivatives is achieved on C 18columns with a gradient of 100% water [TFA, 0.1% (v/v)] --+ 100% CH3CN [TFA, 0.1% (v/v)] over 180 min at 0.75 ml/min. Generally, the compounds with higher numbers of NDI units have longer retention times. Depending on the compounds analyzed, the conditions can be optimized, for example, in the case of mixtures of bisintercalators, a gradient of 85% water [TFA, 0.1% (v/v)] 60% CH3CN [TFA, 0.1% (v/v)] over 60 min at 0.75 ml/min is sufficient and can resolve two bis-NDIs with diastereomeric linkers. Monitoring is by UV (see below) at ~. = 384 nm against a baseline at )~ = 450 nm, or, if available, by liquid chromatography-electrospray ionization (LC-ESI; see below).

Ultraviolet-Visible Spectroscopy The naphthalene diimide chromophore exhibits a characteristic UV spectrum. Generally, because of intramolecular stacking of the chromophores in the compounds with more than one intercalating unit, a hypochromic effect leads to a distinct UV profile for those compounds as compared with the NDI monomer (Fig. 2a). This often allows the unambiguous assignment of the desired multimeric

[28]

POLYINTERCALATING LIGANDS

567

TABLE I EXTINCTIONCOEFFICIENTSAT 386 nm FOR NDI OLIGOMERS Compound

Oligomer

na

~F b

Gly-NDI-Lys Gly-NDI-Lys-GIy-Gly-GIy-NDI-Lys GIy-(NDI-Lys-Gly-GIy-GIy)2-NDI-Lys GIy-(NDI-Lys-GIy-GIy-GIy)3-NDI-Lys GIy-(NDI-Lys-GIy-Gly-Gly)7-NDI-Lys

Monomer Dimer Trimer Tetramer Octamer

0 I 2 3 4

20,000 27,400 39,200 51,300 104,500

~'SDSc 23,400 44,000 74,600 96,000 190,000

a Refer to Fig. 1. b Measured in 10 mM Tris buffer, 1 mM EDTA, 50 mM NaCI, pH 7.4. "Measured in 2% SDS.

product from the commonly present monomer impurities by simple HPLC/UV (Fig. 2). For crude concentration estimates, the extinction coefficients shown in Table I can be used for any derivative with a given number of diimide chromophores. However, the observed hypochromic effect is somewhat sensitive to linker variations; therefore, independent measurement of the extinction coefficient for each derivative is important for accurate DNA binding studies such as quantitative footprinting.

Electrospray Ionization-Mass Spectrometry Compounds with basic amino acid side chains are conveniently analyzed by ESI-mass spectrometry (MS) in the positive ionization mode. All possible ionization states are usually observed, with the relative intensities of the different species somewhat dependent on the conditions. If the basicities of the components of a mixture are similar, as is the case in a library mixture of compounds with one lysine residue per NDI unit, the relative amounts of compounds are roughly proportional to the sums of all ionized species observed. Figure 4 and Table II show the ESI-MS data for a mixture of bisintercalators, 24 synthesized as described above (without double coupling for the second NDI chromophore).

Future Directions We have demonstrated that a modular DNA polyintercalator system, amenable to a great number of structural variations, is accessible via facile solid-phase synthetic procedures. 2°,23,24 Therefore, a number of the technical challenges associated with producing polyintercalating agents with antigene activity have been successfully addressed. The next step in the design of the sequence-specific polyintercalators requires structural studies of bisintercalating modules bound to their

568

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[9,8]

1356.6

1oo-~

°%° !I

90 85 60

Ob 75

7"

890~8 i~08

65 60

1285,7 b13m99"6

28,rnI Im114006

679.2

o~ 45

did

>_ 40

(D tY

25 20~ 3646 422.5

',',

' .....

400

,,,,,J

66 567.6 d I I ~]

....

i ........

600

i.,,,i,

8OO

1000

12O0

1400

..... , ..... ;',',,,,

m/z FIG. 4. ESl-mass spectrum of a mixture of bisintercalators synthesized as described in text. The data are tabulated in Table I1. All 11 nonisobaric compounds were identified as the monoprotonated (m) and/or diprotonated (d) species. Most impurities were identified as monomeric NDI derivatives, resulting from incomplete second NDI coupling (i).

preferred DNA sequences. In this respect, there have been several encouraging developments in bisintercalator design.

Binding Affinity Chaires and co-workers have designed an ultratight binding bisdaunorubicin, WP631,26-28 based on the crystal structure of two daunorubicin monomers complexed to DNA, demonstrating the degree to which structural data can be used to fine-tune the energetics of bisintercalation. Interestingly, the daunorubicin monomers, as well as the dimer with optimal linker, span not two, but four DNA base pairs between the two intercalating aromatic units. 27 We have carried out initial NMR studies of a bis-NDI module, and the resulting structure indicates four base pairs might be the optimal inter-intercalator distance for our NDI systems as well, at least for certain DNA sequences.29 Another 26 j. Chaires, E Leng, T. Przewloka, 1. Fokt, Y. Ling, R. Perez-Soler, and W. Priebe, J. Med. Chem. 40, 261 (1997). 27 G. H u , X. Shui, E Leng, W. Priebe, J. Chaires, and L. Williams, Biochemistry 36, 5940 (1997). 28 E Leng, W. Priebe, and J. Chaires, Biochemistry 37, 1743 (1998). 29 g. Guelev, S. Sorey, J. Lee, J. Ward, D. Hoffman, and B. Iverson, in preparation (2001).

[28]

POLYINTERCALATING

LIGANDS

569

Z ¢g ,.A

E b

E z

© [...

e..,

,-r, [... ©

.7 © [-.

< =~<

-7-

e',: ~ 0~ o

o

,-; ~ ~

¢5 ,....; ,.-.: ¢,4

~<~ Pz ,.a < Z

¢~

~" .'¢2 ._E

<

,< [... ¢~

.~

<

V-

(.) eo

.--,

570

CHEMICAL AND MOLECULAR BIOLOGICAL APPROACHES

[9,8]

interesting observation, which may be crucial for the design of tetraintercalators with high binding affinities, is that NDI-based bisintercalators, like other bisintercalators such as echinomycin, exhibit a general propensity to bind cooperatively in pairs. 2°'24

Sequence Specificity We originally reported a general preference for (G + C)-rich sequences for members of the NDI polyintercalator series, as evidenced by slower UV dissociation profiles in SDS from poly(dG-dC) compared with poly(dA-dT), and by DNase I footprinting of specifically designed oligonucleotides. 2° However, in a library screen we showed that the DNA-binding specificity of the bisintercalators (Fig. 3) could vary quite dramatically with variations in the amino acid linkers connecting the NDI units, and is not simply limited to (G + C)-rich versus (A + T)-rich DNA. 24 Consistent with this conclusion, we found preliminary DNase ! footprinting data for the octaintercalating compound difficult to interpret. 23 It now appears that a more detailed analysis of the sequence specificity of the higher analogs should await detailed structural data for the bisintercalating modules. Moreover, the data on sequence specificity raises the exciting possibility that the design of future NDI polyintercalators could be a priori targeted to relatively long specific DNA sequences.

Threading versus Nonthreading Intercalation Although initial UV dissociation measurements of our polyintercalators from poly(dA-dT) duplexes were consistent with a nonthreading binding mode for those compounds] ° these results are in contradiction with NMR data for an NDI bisintercalator, 29 suggesting, perhaps, that the DNA-binding mode depends on DNA sequence. Moreover, results from other laboratories strongly support the feasibility of threading polyintercalation with simultaneous blockage of both DNA grooves. An NMR structure of the bisnaphthalimide LU-79553 bound to DNA 3° revealed that the inter-intercalator linker is placed in the major groove--the first (and long awaited) structurally verified example of such a binding mode. On the basis of limited spectroscopic evidence, a bis-[Ru(phen)2dppz] 2+ intercalator was also thought to bind in a threading manner. 31 These exciting results, coupled with the synthetic advances described here, anticipate the development, in the not-sodistant future, of modular DNA polyintercalators in which new major groovebinding motifs are combined with existing minor groove recognition elements to achieve coverage of extended sequences via both DNA grooves.

30 j. Gallego and B. Reid, Biochemistry 38, 15104 (1999). 31 B. Onfelt, P. Lincoln, and B. Nord6n, J. Am. Chem. Soc. 121, 10846 (1999).

[29]

TARGETING TELOMERES AND TELOMERASE

573

[29] Targeting Telomeres and Telomerase B y DAEKYU SUN a n d LAURENCE H. HURLEY

Introduction Telomeres and Telomerase The ends of eukaryotic chromosomes have unique nucleoprotein structures, termed telomeres, which consist of telomeric DNA and the proteins that bind specifically to these sequences.l-4 Telomeric DNA, found in all vertebrates including humans, consists of tandem repeats of the hexanucleotide sequence (TTAGGG)n.l-4 The size of telomeric DNA varies among different species, tissues, and cell types, and ranges from a few kilobases (kb) to more than 100 kb. 4 Telomeres are essential for maintaining genomic stability by providing a protective cap for the ends of chromosomes. 3'4 Without this telomeric cap, chromosomes would undergo degradation, recombination, or fusion, resulting in the formation of dicentric and multicentric chromosomes. 5-8 The other essential function of telomeres in some cells is to prevent the loss of terminal bases at each end of chromosomal DNA following the completion of linear chromosomal DNA replication. The "end replication" problem, which Watson 9 and Olovnikov l°' I I first independently described, is the inability of the DNA replication machinery to copy the final few base pairs of the lagging strand. Some cell types, such as germ line or cancer cells, could overcome this end replication problem by using mechanisms that replenish telomeric DNA at the ends of chromosomal DNA. In the absence of these mechanisms, telomeric DNA in most human somatic cells becomes progressively shortened. Because the cumulative loss of telomeric DNA after many cell divisions results in a limited replicative capacity, this process is proposed to function as a biological clock, eventually limiting the proliferative life span of somatic cells and leading to cellular senescence. 5' i2-14

I V. A. Zakian, Annu. Rev. Genet. 23, 579 (1989). 2 E. H. Blackburn, Science 249, 489 (1990). 3 E. H. Blackburn, Nature (London) 350, 569 (1991). 4 E. H. Blackburn and W. C. Greider (eds.), "Telomeres." Cold Spring Harbor Press, Cold Spring Harbor, New York, 1995. 5 C. B. Harley, A. B. Futcher, and C. W. Greider, Nature (London) 345, 458 (1990). 6 C. M. Counter, A. A. Avilion, C. E. LeFeuvre, N. G. Stewart, C. W. Greider, C. B. Harley, and S, Bacchetti, EMBO J. 11, 1921 (1992). 7 C. B. Harley and B. Villeponteau, Curt: Opin. Genet. Dev. 5, 249 (1995). 8 B. van Steensel, A. Smogorzewska, and T. de Lange, Cell 92, 401 (1998). 9 j. D. Watson, Nat. New Biol. 239, 197 (1972). 10 A. M. Olovnikov, Dokl. Akad. Nauk (S.S.S.R.) 201, 1496 (1971).

METHODSIN ENZYMOLOGY,VOL.340

Copyright4) 2001 by AcademicPress All rightsof reproductionin any formreserved. I)076-6879/00$35.00

574

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Various immortalized cells, including tumor cells and stem cells, are known to obtain an unlimited replicative capacity through the activation of telomerase. 6 Telomerase was first discovered on the basis of distinctive telomere extension activity in the ciliate Tetrahymena,and, subsequently, in human HeLa cell lines. 15-18 It is a ribonucleoenzyme that consists of RNA and reverse transcriptase components and is able to compensate for the sequence losses resulting from incomplete terminal replication by using a short RNA motif as a template for the synthesis of telomeric D N A J 5'17 Thus, telomerase is a unique polymerase in that it carries its own template as a part of the enzyme. The RNA component of telomerase was first cloned and sequenced from various ciliates and subsequently from two mammalian sources, human and mouse. 17 20 Both mammalian telomerase RNAs are approximately 500 nucleotides in length. 19,20 The protein component of telomerase was first identified in Euplotes aediculatus and has subsequently been identified in Tetrahymena,Sacchalvmyces, Schizosaccharomyces, and mammals. 21-24 These proteins harbor several sequence motifs characteristic of catalytic regions of reverse transcriptase. 2~-26 Disruption of these motifs has been reported to abolish the enzymatic activity of telomerase. 22'26 The current model for the mechanism of telomeric DNA synthesis by telomerase was first proposed by Blackburn. 2 As depicted in Fig. 1, the 3' end of a single-stranded telomeric repeat base pairs with the template domain of telomerase RNA (recognition step), which allows the reverse transcriptase extension of 11 A. M. Olovnikov, J. Theor Biol. 41, 181 (1973). 12 N. D. Hastie, M. Dempster, M. G. Dunlop, A. M. Thompson, D. K. Green, and R. C. Allshire, Nature (London) 346, 866 (1990). 13 j. Lindsey, N. J. McGill, L. A. Lindsey, D. K. Green, and H. J. Cook, Mutat. Res. 256, 45 (1991). 14 M. Z. Levy, R. C. Allsopp, A. B. Futcher, C. W. Greider, and C. B. Harley, J. Mol. Biol. 225, 951 (1992). 15 C. W. Grieder and E. H. Blackburn, Cell 51,887 (1987). 16 G. B. Morin, Cell 59, 529 (1989). 17 C. W. Greider and E. H. Blackburn, Nature (London) 337, 331 (1989). Ix D. Shippen-Lentz and E. H. Blackburn, Mol. Cell. Biol. 9, 2761 (1989). 19 j. Feng, W. D. Funk, S. S. Wang, S. L. Weinrich, A. A. Avilion, C. A. Chiu, R. R. Adams, E. Chang, R. C. Allsopp, J. Yu, S. Le, M. D. West, C. B. Harley, W. H. Andrew, C. W. Greider, and B. Vileponteau, Science 269, 1236 (1995). 20 M, A. Blasco, W. D. Funk, B. Villeponteau, and C. W. Greider, Science 269, 1267 (1995). 21 T. M. Nakamura, G. B. Morin, K, B. Chapman, S. L. Weinrich, W. H. Andrews, J. Lingner, C. B. Harley, and T. R. Cech, Science 277, 955 (1997). 22 C. M. Counter, M. Meyerson, E. N. Eaton, and R. A. Weinberg, Proc. Natl. Acad. Sci. U.S.A. 94, 9202 (1997). 23 j. Lingner, T. R. Hughes, A. Shevchenko, M. Mann, V. Lundblad, and T. R. Cech, Science 276, 561 (1997). 24 L. Harrington, W. Zhou, T. McPhail, R. Oulton, D. S. K. Yeung, V. Mar, M. B. Bass, and M. O. Robinson, Genes Dev. l 1, 3109 (1997). 25 V. Lundblad, Proc. Natl. Acad. Sci. U.S.A. 95, 8415 (1998). 26 j. Nakayama, M. Saito, H. Nakamura, A. Matsuura, and F. Ishikawa, Cell 88, 875 (1997).

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Annealing 5"-TrAGGGTTAGGGTTAGGG-3" + Telomeric Repeats

RNA C o m p o n e n ~ hTR

Protein Component(s)

I~ Translocation

•c , c ~ a ' r ~

Elongation (6 bp)

~ _ _

FIG. I. Proposedmodel for the mechanismof telomericDNA synthesis by telomerase. telomeric repeats by the protein component of telomerase using the telomerase RNA as a template. After the addition of the first four telomeric nucleotides (TrAG) to the 3' end of the single-stranded telomeric repeat (initiation step), the 3' side of the substrate dissociates from its RNA template (dissociation step), and is translocated to the 3' portion of the RNA template (translocation step). Thus, the extended telomeric repeats are subjected to another round of elongation by telomerase in which six nucleotides (GGTTAG) are added to the 3' end of telomere DNA (synthesis). Through this cycle of controlled reactions, telomerase is able to continue to elongate the same telomeric repeats up to hundreds of nucleotides without dissociation. 27 The processive nature of the telomerase reaction is one of its most striking features, and in a cell-free assay, this implies a translocation reaction to reposition the primer and template strands for successive rounds of elongation without separation of the primer from the telomerase.15' 16 There is sufficient evidence to believe that telomerase activity is directly associated with cellular proliferation (see Holt and Shay 28 for review). In model tissue culture systems, telomerase repression was often observed when either tumor or stem cells were induced to move into terminal differentiation, Go quiescence, or drug-induced growth arrest. 28 Results from a telomerase-knockout (KO) mouse 27E. H. Blackburn,Annu. Rev. Biochem. 61, 113 (1992). 28S. E. Holt and J. W. Shay,J. Cell. Physiol. 180, 10 (1999).

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experiment also suggested that progressive telomere loss due to the absence of active telomerase reduces the proliferation and self-renewal potential of regenerative cells and tissues in mice. 29'3° The same studies concluded that telomerase deficiency ultimately produces a reduction in proliferation and an increase in apoptosis in germ cells of male mice. 3° Telomerase is found at high levels in the majority of cancer ceils, but is absent or significantly lower in benign tumors and resting cells. 31-33 A survey regarding the status of telomerase in human cancers has revealed that nearly 85% of the malignant human tumors tested showed high levels of telomerase activity, whereas less than 0.5% of normal cells were telomerase positive. 33 With the cloning of the telomeric catalytic subunit hTERT (human telomerase reverse transcriptase), a critically important question was answered concerning the role of telomerase in cellular senescence, telomere maintenance, and tumorigenesis. Normal human fibroblast and epithelial cells undergo a limited number of cell divisions and ultimately enter a nondividing state, called replicative senescence, as a result of the progressive loss of telomeric DNA. 5,13,14,34 When hTERT is stably transfected into normal cells, the resulting fibroblast and epithelial cell clones, which now express telomerase, are capable of extending their life span by maintaining or even elongating their telomeres. 35-37 The generation of these telomerase-immortalized normal human cells provides the first direct evidence of the essential role of telomerase in preventing telomere shortening and cellular senescence. Other studies suggest that telomerase alone does not cause cancer, despite its critical function in maintaining telomeres and facilitating continued cell growth. 38-4° However, in one report, the ectopic expression of hTERT in combination with two oncogenes [the simian virus 40 (SV40) large-T oncoprotein and an oncogenic allele of H - r a s ] results in direct tumorigenic conversion of 29M. A. Blasco, H. W. Lee, M. E Hande, E. Samper, P. M. Lansdorp, R. A. DePinho, and C. W. Greider, Cell 91, 25 (1997). 3oH. W. Lee, M. A. Blasco, G. J. Gottlieb,J. W. Homer, and C. W. Greider,Nature (London) 392, 569 (1998). 31T. de Lange,Proc. Natl. Acad. Sci. U.S.A. 91, 2882 (1994). 32N. W. Kim, M. A. Piatyszek, K. R. Prowse, C. B. Harley, M. D. West, P. L. C. Ho, G. M. Coviello, W. E. Wright, S. L. Weinrich,and J. W. Shay, Science 266, 2011 (1994). 33j. W. Shay and S. Bacchetti,Eur J. Cancer 33, 787 (1997). 34j. Campisi,Eur. J. Cancer 33, 703 (1997). 35A. G. Bodnar,M. Ouellette,M. Frolkis,S. E. Holt, C. Chiu,G. B. Morin,C. B. Harley,J. W. Shay, S. Lichtsteiner,and W. E. Wright, Science 279, 349 (1998). 36H. Vaziriand S. Benchimol,Curr. Biol. 8, 279 (1998). 37C. M. Counter, W. C. Hahn, W. Wei, S. D. Caddle, R. L. Beijersbergen,P. M. Lansdorp, J. M. Sedivy, and R. A. Weinberg,Proc. Natl. Acad. Sci. U.S.A. 95, 14723 (1998). 38j. Wang,L. Y. Xie, S. Allan,D. Beach, and G. J. Hannon,Genes Dev. 12, 1769 (1998). 39M. M. Ouellette, D. L. Aisner,I. Savre-Train,W. E. Wright, and J. W. Shay, Biochem. Biophys. Res. Commun. 254, 795 (1999). 40j. Zhu, H. Wang,M. Bishop, and E. H. Blackburn,Proc. Natl. Acad. Sci. U.S.A. 96, 3723 (1999).

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normal human epithelial and fibroblast cells. 41 A subsequent study by the same group demonstrated that human tumor cells without telomerase undergo apoptosis after a limited number of cell divisions. 42 These results provide the first direct evidence that telomerase is not a mere by-product of human cancer, but one of the components required during multistep tumorigenesis. Taken together, these results suggest that cancer cell telomeres and telomerase, or more broadly, telomere maintenance mechanisms, could be a selective target for anticancer agents. It is known that telomerase activity in hematopoietic stem cells in bone marrow is upregulated only when the growth is stimulated by cytokines, whereas resting quiescent cells are essentially telomerase negative.43 Thus, it was proposed that telomerase-competent stern cells may regulate telomerase activity to maintain levels sufficient to slow but not prevent telomere shortening. 36,44 Because most cancer cells possess a high proliferative index compared with normal stem or germ line cells, antitelomerase therapy is expected to have minimal side effects, such as bone marrow toxicity, and this would be much safer for patients than conventional chemotherapy. 29 Telomerase-negative tumors have been identifiedY and alternative mechanisms of maintaining telomere length in several human tumor cell lines have been reported. 45'46 Thus, antitelomerase therapy cannot be regarded as a foolproof way of treating human cancers, but it will deprive tumor cells of a growth advantage over surrounding normal cells, which could result in a significant reduction of malignant progression of tumor cells. In addition, the disruption of telomere maintenance could be achieved through targeting components other than telomerase that are involved in the telomere maintenance mechanism (see below). The simultaneous downregulation of telomerase and the disruption of telomere maintenance mechanisms by various means might be expected to produce even greater detrimental effects on cancer cell viability. S t r a t e g i e s to T a r g e t T e l o m e r a s e Not surprisingly, there have been a variety of reports on different strategies for inhibiting telomerase activity in human cells, because telomerase is considered the primary mechanism for telomere maintenance in highly proliferative tumor 4~ W. C. Hahn, C. M. Counter, A. S. Lundberg, R. L. Beijersbergen, M. W. Brooks, and R. A. Weinberg, Nature (London) 400, 464 (1999). 42 W. C. Hahn, S. A. Stewart, M. W. Brooks, S. G. York, E. Eaton, A. Kurachi, R. L. Beijersbergen, J. H. Knoll, M. Meyerson, and R. A. Weinberg, Nat. Med. 5, 1164 (1999). 43 C.-E Chui, W. Dragowska, N. W. Kim, H. Vaziri, J. Yui, T. E. Thomas, C. B. Harley, and R M. Lansdrop, Stem Cells 14, 239 (1996). 44 C. Pan, B.-H. Xue, T. M. Ellis, D. J. Peace, and M. O. Diaz, Exp. CellRes. 321, 346 (1997). 45 T. M. Bryan, A. Englezou, J. Gupta, S. Bacchetti, and R. R. Reddel, EMBO J. 14, 4240 (1995). 46 T. M. Bryan, A. Englezou, L. Dalla-Pozza, M. A. Dunham, and R. R. Reddel, Nat. Med. 3, 1271 (1997).

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cells. 47-56 Here we describe a spectrum of strategies for developing novel agents that interfere with various aspects of human telomerase function. Reviews on development of telomerase inhibitors have been published. 57'58 Targeting H u m a n Telomerase Reverse Transcriptase

hTERT, the catalytic subunit of telomerase, has been implicated as an important contributor to the immortalization process, and the hTERT protein contains well-conserved reverse transcriptase motifs. 59 Therefore, several known reverse transcriptase inhibitors (RTIs) belonging to the family of nucleotide analogs have been tested as potential inhibitors, using an in vitro telomerase assay, and have been further evaluated to test their effects on telomere maintenance in cell culture models in vitro. 48"49 In earlier studies, the effects of azidothymidine triphosphate (AZTTP), arabinofuranosylguanine triphosphate (Ara-GTP), dideoxyinosine triphosphate (ddlTP), ddTTP, and ddGTP on Tetrahymena telomerase were studied. 48'49 All these nucleoside triphosphate analogs inhibited telomerase activity in vitro, with only small variation in potency among the analogs. Significantly, the same study revealed that the analogs AZT, 3'-deoxy-2',3'-didehydrothymidine (d4T), and A r a - G in the nucleoside form also caused a consistent and rapid telomere shortening in vegetatively growing Tetrahymena. In a more recent study, using both a metastatic murine melanoma cell line and a human breast cancer cell line, it was determined whether treatment of cancer cells with A Z T has any effect on telomere length. 5° A fluorescence in situ hybridization (FISH) analysis was used to measure telomere length, and the results showed a significant concentrationdependent reduction of telomeric signal intensity in interphase and metaphase 47 E. Raymond, D. Sun, S.-E Chen, B. Windle, and D. D. Von Hoff, Curr. Opin. Biotechnol. 7, 583 (1996). 48 C. Strahl and E. H. Blackburn,Nucleic Acids Res. 22, 893 (1994). 49 C. Strahl and E. H. Blackburn,Mol. Cell. Biol. 16, 53 (1996). 50 A. S. Multani, C. Furlong, and S. Pathak, Int. J. Oncol. 13, 923 (1998). 51 y. Kondo, S. Koga, T. Komata, and S. Kondo, Oncogene 19, 2205 (2000). 52 D. M. Kushner, J. M. Paranjape, B. Bandyopadhyay,H. Cramer, D. W. Leaman, A. W. Kennedy, R. H. Silverman, and J. K. Cowell, Gynecol. Oncol. 76, 183 (2000). 53 B. Pandit and N. P. Bhattacharyya,Biochem. Biophys. Res. Commun. 251, 620 (1998). 54 A. E. Pitts and D. R. Corey, Proc. Natl. Acad. Sci. U.S.A. 95, 11549 (1998). 55 A. I. Glukhov, O. V. Zimnik, S. A. Gordeev, and S. E. Severin, Biochem. Biophys. Res. Commun. 248, 368 (1998). 56 S. E. Hamilton, C. G. Simmons, I. S. Kathiriya, and D. R. Corey, Chem. Biol. 6, 343 (1999). 57 J.-L. Mergny, P. MaiUiet,E Lavelle, J.-F. Riou, A. Laoui, and Claude H61~ne,Anti-Cancer Drug Des. 14, 327 (1999). 58 S. Neidle and L. R. Kelland, Anti-Cancer Drug Des. 14, 341 (1999). 59 M. Meyerson, C. M. Counter, E. N. Eaton, L. W. Ellisen, E Steiner, S. D. Caddie, L. Ziangra, R. L. Beijersbergen,M. J. Davidoff, Q. Liu, S. Bacehetti, D. A. Haber, and R. A. Weinberg,Cell 911,785 (1997).

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spreads of AZT-treated cells, whereas no reduction in signal intensity was observed in untreated controls. These findings provide validation for the approach to designing telomerase inhibitors based on nucleotide analogs so as to achieve the disruption of telomere maintenance in cancer cells. As an alternative to the nucleotide approach, potent telomerase inhibitors have been identified through screening of chemical libraries in a telomerase assay. In one study, this has resulted in identification of a set of isothiazolone-containing telomerase inhibitors that have submicromolar potency in a telomerase assay. 6° Because isothiazolones are known to react with the reduced thiols of cysteine and dithiothreitol (DTT), these compounds are proposed to interfere with telomerase activity by modifying a cysteine residue(s) in or near the reverse transcriptase active site. 6° This result validates the use of a biochemical screening assay system that targets the reverse transcriptase activity of telomerase for identifying inhibitors. A second approach to targeting hTERT protein is to use antisense oligonucleotides that are designed to hybridize with complementary sequences of hTERT m R N A and consequently deplete hTERT protein levels inside cells. The clinical success of the first antisense drug provides impetus for the further development of these strategies for selective disruption of telomerase expression. 61,62 Because only the RNA component has been targeted in the area of telomerase-directed antisense strategies, in future studies it will be worthwhile to test the effect of an antisense drug designed to target hTERT. Targeting H u m a n T e l o m e r a s e R N A C o m p o n e n t

The RNA component of telomerase is divided into several unique domains that are critical for normal enzymatic function and assembly of the telomerase enzyme. 62-67 An alignment domain facilitates substrate binding to the template domain for polymerization. Other nontemplate domains of the RNA are probably involved with binding to the telomerase protein components and further involved in modulating enzymatic activity. The critical role of the RNA component of telomerase in the enzymatic function has inspired a number of investigators to design telomerase inhibitors based on its sequence and structure. Because the o0L. A. Bare, L. Trinh, S. Wu, and J. J. Devlin, Drug Dev. Res. 43, 109 (1998). ol A Webb, D. Cunningham, E Cotter, E A. Clarke, E di Stefano, P. Ross, M. Corbo, and Z. Dziewanowska,Lancet 349, 1137 0997). 62j. Nemunaitis, G. Eckhardt, A. Don-,J. Pribble, R. Smith, J. Bruce, N. Ogonskie, and D. D. Von Hoff, "ASCO Meeting Abstracts," p. 246a. American Society of Clinical Oncology,Alexandria, Virginia, 1997. 63A. Bhattacharyyaand E. Blackburn, Proc. Natl. Acad. Sci. U.SA. 94, 2823 (1997). 64D. Gilley, M. S. Lee, and E. H. Blackburn, GenesDev. 9, 2214 (1995). 65D. Giily and E. H. Blackburn, Mol. Cell. Biol. 16, 66 (1996). 66 D. Gilley and E. H. Blackburn, Proc. Natl. Acad. Sci. U.S~4.96, 6621 (1999). 67y. Tzfati, T. B. Fulton, J. Roy, and E. H. Blackburn, Science 288, 863 (2000).

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RNA component of telomerase is an ideal target for oligonucleotides that are able to recognize and hybridize to accessible complementary RNA sequences, an antisense strategy has been explored. 52-56 The initial studies have mostly been directed toward targeting the template domain of telomerase RNA with antisense oligonucleotides, and efficient inhibition of telomerase activity has been achieved. In efforts to improve the stability and binding specificity of antisense oligonucleotides, the backbone structure of the administered oligonucleotides has been modified. For example, peptide nucleic acids (PNAs), which are backbone-modified oligomers, 68'69 have shown a clear advantage over phosphorothioate oligonucleotides in both binding efficiency and selectivity. 56,7° More recently, it has been reported that 2'-O-alkylRNA oligomers are even more selective than PNAs in binding to the complementary RNA sequences. 54 Furthermore, results from the same study showed that 2'-O-methyl-RNA-based oligomers complexed with cationic lipids can efficiently inhibit telomerase on transfection of a human prostate tumor cell line. 54 In one study it has been shown that an RNA pseudoknot structure located in the nontemplate domain in ciliate telomerase plays a critical role in the stable assembly of a catalytically active enzyme. 66 Strong evidence has also been obtained for the functional importance of RNA secondary structures outside the template region of the RNA component of telomerase. 67 The disruption of this conserved RNA structure adjacent to the template domain of yeast telomerase caused abnormal synthesis of telomeric DNA, eventually leading to defects in cellular growth, presumably due to the failure of the normal mechanism for telomere maintenance. 67 Thus future studies that employ oligonucleotide targeting of nontemplate domains as well as the template domain of telomerase RNA may prove to be fruitful. 66'67 The pseudoknot may well afford a useful target. R N A / D N A H y b r i d Target

In addition to an accessible single-stranded human telomerase RNA component (hTR) RNA template and secondary structures such as the pseudoknot, the heteroduplex D N A . RNA d(TTAGGG), r(CCCUAA) formed when the singlestranded telomeric DNA binds to the RNA template is another possible telomerasespecific target. This heteroduplex is similar to the Okazaki fragments formed as a consequence of RNA priming for DNA polymerases for which a nuclear magnetic resonance (NMR) structure has been solved. 71 68L. Betts, J. A. Josey, J. M. Veal, and S. R. Jordan, Science 270, 1838 (1995). 69N. J. Peffer, J. C. Hanvey,J. E. Bisi, S. A. Thomson,C. E Hassman, S. A. Noble, and L. E. Babiss, Proc. Natl. Acad. Sci. U.S.A. 90, 10648 (1993). 70S. E. Hamilton, A. E. Pitts, R. R. Katipally, X. Jia, J. P. Rutter, B. A. Davies, J. W. Shay, W. E. Wright, and D. R. Corey,Biochemistry 36, 11873 (1997). 71 O. Y. Fedoroff,M. Salazar, and B. R. Reid, J. Mol. Biol. 233, 509 (1993).

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Targeting Assembly of Active Telomerase Molecular chaperones are a diverse group of proteins that prevent or reverse deleterious aggregation of protein-folding intermediates. 72,73 Their function is not only to facilitate the de novo folding of newly synthesized proteins, but also to allow renaturation of partially denatured proteins. The 90-kDa heat shock protein (Hsp90) and p23 were identified in rabbit reticulocyte lysates (RRLs) as molecular chaperones in the assembly of active telomerase from hTERT and hTR in vitro and in vivo. TM The 90-kDa heat shock proteins are also involved in the folding of a wide range of proteins. 75-77 The benzoquinone ansamycin antibiotics, such as geldanamycin and herbimycin A, are known to bind to the ATP-binding site in the N-terminal domain of Hsp90 and to disrupt the ATPase activity that is essential for its chaperone activity in vivo. 78 Significantly, downregulation of many proteins was observed when functions of Hsp90 were blocked with these antibiotics, resulting in significant antiproliferative effects on tumor cells. 73 The blockage of ATPase activity of Hsp90 with geldanamycin also resulted in downregulation of telomerase activity due to failure in assembly of the active telomerase complex. 74 Because telomerase expression is known to be closely associated with cellular proliferation, downregulation by preventing chaperone-promoted assembly of active telomerase is an alternative strategy for achieving interference with telomere maintenance in human tumors.

Assays for Telomerase Activity Because telomerase is an attractive target for anticancer agents as well as an important diagnostic marker of human c a n c e r , 31-33'79's0 it is important to measure accurately the levels of enzyme activity present in tumor tissue samples and cultured human cells. Currently, two telomerase assay methods are used: the conventional 72 H. Wiech, J. Buchner, R. Zimmerman, and U. Jakob, Nature (London) 358, 169 (1992). 73 j. Buchner, Trends Biochem. Sci. 24, 136 (1999). 74 S. E. Holt, D. L. Aisner, J. Baur, V. M. Tesrner, M. Dy, M. Ouellette, J. B. Trager, G. B. Morin, D. O. Toft, J. W. Shay, W. E. Wright, and M. A. White, Genes Dev. 13, 817 (1999). 75 S. D. Hartson, E. A. Ottinger, W. Huang, G. Barany, P. Bum, and R. L. Matts, J. Biol. Chem. 273, 8475 (1998). 76 S. M. Roe, C. Prodromou, R. O'Brien, J. E. Ladbury, P. W. Piper, and L. H. Pearl, J. Med. Chem. 42, 260 (1999). 77 R. J. Schmacher, R. Hurst, W. P. Sullivan, N. J. McMahon, D. O. Toft, and R. L. Matts, J. Biol. Chem. 269, 9493 (1994). 78 C. E. Stebbins, A. A. Russo, C. Schneider, N. Rosen, E U. Hartl, and N. P. Pavletich, Cell 90, 239 (1997). 79 E. Hiyama, L. Gollahon, T. Kataoka, K. Kuroi, T. Yokoyama, A. Gazdar, K. Hiyama, M. A. Piatyszek, and J. W. Shay, J. Natl. Cancer Inst. 88, 116 (1996). 80 C. B. Harley and N. W. Kim, in "Principles and Practice of Oncology," Vol. 11, p. 1, LippincottRaven, Philadelphia, 1997.

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primer extension assay 16 and the telomeric repeat amplification protocol (TRAP) assay based on the polymerase chain reaction (PCR). 32 In the conventional primer extension assay, the telomerase enzyme extends the primer by adding a number of telomeric repeats. The telomerase reaction is routinely subjected to phenolchloroform extraction followed by ethanol precipitation to separate unincorporated [ct-32p]dGTP and cellular proteins from the product of the primer extension reaction. The extension products are then analyzed by denaturing polyacrylamide gel electrophoresis followed by autoradiography. However, both a high level of background from residual 32p and low specific activity of human telomerase make it difficult to obtain quantitative results.~6 The more recently developed TRAP assay has greatly improved the speed and sensitivity of telomerase activity measurements. The method is reported to be able to detect telomerase levels in as few as --~10 human cells. 32 In this assay, the telomerase substrate is extended by the telomerase enzyme, and then serves as a template for the ensuing polymerase chain reaction. The PCR step greatly enhances the sensitivity of detection of telomerase levels, even in small amounts of clinical samples. The PCR-amplified products of the extended telomerase substrate result in a six-nucleotide DNA ladder, which is easily visualized on a native polyacrylamide gel. The availability of the TRAP assay dramatically increased the number of laboratories evaluating telomerase activity in clinical samples. 33'79'8° Furthermore, the method has been improved for the high-throughput enzyme-linked immunosorbent assay (ELISA) format, allowing the rapid analysis of a large number of samples and the screening of telomerase inhibitors from natural and chemical libraries. 6°'81-83 However, the TRAP assay has a major drawback for screening purposes, because any compound that interferes with Taq DNA polymerase activity will produce a false-positive result. In addition, the reaction products of both processive and nonprocessive telomerase activity cannot be distinguished after PCR amplification of extension products, thereby making the mechanistic study of telomerase difficult, even though it has been somewhat improved by the further modification of methodology. 81-83 The primer extension telomerase assay has been improved by a biomagnetic separation method to overcome the inherent difficulties of the conventional assay. 84 In this method, 5'-biotinylated (TTAGGG)3 is used as a primer for the telomerase reaction. The biotinylated primer is then immobilized onto streptavidin-coated Dynabeads so as to remove 32p-labeled high molecular weight DNA molecules and unincorporated [~-32p]dGTE Using this method, it is possible to reproducibly 81N. W. Kim and F. Wu, Nucleic Acids Res. 25, 2595 (1997). 82C. Poremba, W. Bocker, H. Willenbring, K. L. Schafer, F. Otterbach, H. Burger, R. Diallo, and B. Dockhorn-Dworniczak,Int. J. Oncol. 12, 641 (1998). 83j. n. Ohyashiki,K. Ohyashiki,T. Sano, and K. Toyama,Jpn. J. Cancer Res. 87, 329 (1996). 84D. Sun, L. H. Hurley,and D. D. Von Hoff,BioTechniques 25, 1046 (1998).

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detect telomerase activity without amplification of the signals from cell extracts equivalent to as few as 200-500 cells. This assay is useful in studies requiring more quantitative measures of telomerase activity and in detecting potential telomerase inhibitors. Last, it is useful in probing biochemical aspects of the telomerase reaction. 85,86

Methodologies Used for Telomerase Assays Preparation of Cell Extracts. Cultured human cells are washed once in phosphate-buffered saline (PBS), resuspended in ice-cold washing buffer (10 mM HEPES-KOH, 1.5 mM MgC12, 10 mM KC1, and 1 mM dithiothreitol, pH 7.5), and pelleted by centrifugation at 10,000g for 1 min at 4 °. The washed cells are resuspended in ice-cold lysis buffer containing 10 mM Tris-HCl (pH 7.5), 1 mM MgC12, 1 mM EDTA, 0.1 mM phenylmethylsulfonyl fluoride (PMSF), 5 mM 2-mercaptoethanol (2-ME), 1 mM dithiothreitol, 0.5% (w/v) 3-[(3-cholamidoproyl)dimethylammonio]- 1-propane sulfonate, and RNase inhibitor (50 units/ml), along with 10% (v/v) glycerol (106 cells/20/zl of buffer) and incubated for 30 min on ice. 32 The cell lysates are centrifuged for 30 min in a microcentrifuge at 13,000 rpm at 4 °, and the resulting supematant is adjusted to 20% (v/v) glycerol and stored in a - 8 0 ° freezer. Conventional Telomerase Assay. Telomerase activity is assayed by measuring the amount of extension of telomeric primer (TFAGGG)3 by addition of (TTAGGG)b. The reaction mixture (40/tl), which contains 2 mM dATP, 2 mM dTI'P, 1 # M (TTAGGG)3, and 1.56 IzM [a-32p]dGTP (800 Ci/mmol) in 50 mM Tris-HCl (pH 8.5), 50 mM potassium chloride, 1 mM MgC12, 5 mM 2-mercaptoethanol, and 1 mM spermidine, and 4 #1 of cell extract (150 lzg of protein), is incubated at 30 ° for 1 hr. 16 The stop solution containing either RNase A or proteinase K is added sequentially to the reaction tube to terminate the reaction by destroying the RNA and protein components. After chloroform-phenol extraction, the reaction products are precipitated with ethanol and separated on a denaturing polyacrylamide sequencing gel. The DNA ladders can be visualized by autoradiography or by phosphorimage analysis. Telomeric Repeat Amplification Protocol Assay. The reaction mixture (50/zl) containing cell extracts (0.1-5/zg) is incubated for 30 rain at 25 ° in a final volume of 50/zl containing 20 mM Tris-HC1 (pH 8.3), 1.5 mM MgC12, 63 mM KC1, 0.005% (v/v) Tween 20, 1 mM EGTA, a 50/~M concentration of each deoxynucleoside triphosphate, 0.1 /zM TS primer sequence (5'-AATCCGTCGAGCAGAGTT-3~), 2 U of Taq DNA polymerase, 0.1 # M [c¢-32p]dCTP (800 Ci/mmol), and bovine 85D. Sun, B. Thompson,M. Salazar,S. M. Kerwin,T. Jenkins, S. Neidle, and L. H. Hurley,J. Med. Chem. 40, 2113 (1997). 86D. Sun,C. C. Lopez-Guajardo,J. Quada,L. H. Hurley,and D. D. VonHoff,Biochemistry38, 4037 (1999).

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serum albumin (0.1 mg/ml). After further incubation at 90 ° for 5 min to inactivate telomerase, 0.1/zg of CX primer sequence [5'-(CCCTTA)3CCCTAA-Y] is added, and the resulting mixture is subjected to the polymerase chain reaction for 24 cycles of 30 sec at 94 °, 30 sec at 50 °, and 60 sec at 72 °, and subsequently to extension (72 ° for 10 min). The amplification products are separated by electrophoresis in a nondenaturing 10% (w/v) polyacrylamide gel. The 6-bp DNA ladder can be visualized by autoradiography or by phosphorimage analysis. Telomerase activity is quantitated by densitometric analysis (ImageQuant; Molecular Dynamics, Sunnyvale, CA) of the autoradiogram. Conventional Telomerase Assay Using 5'-Biotinylated Primer. The telomerase reaction mixture (20/zl) containing 50 mM Tris-acetate (pH 8.5), 50 mM KC1, 1 mM MgC12, 5 mM 2-ME, 1 mM spermidine, 1 #M telomere primer [5'-biotinylated-(TTAGGG)3], 2.4/zM [a-32p]dGTP (800 Ci/mmol), 1 mM dATE and 1 mM dTTP with 4/zl of cell lysate is incubated at 30 ° for 1 hr. Reactions are terminated by adding 20 #1 of prewashed streptavidin-coated Dynabead suspension (Dynabeads M-280; Dynal, Great Neck, NY) and then reaction products are immobilized on Dynabeads by continuous agitation of the Dynabead suspension for 1 hr at room temperature. The immobilized reaction products are separated from the suspensions with a magnet (Dynal MPC), and the beads are washed at least five times with washing buffer (1 M NaC1 and 10 mM Tris-HCl, pH 7.5). The reaction products are dissociated from the Dynabeads by incubating the Dynabead-reaction product complex in 200/zl of guanidine hydrochloride solution (final concentration, 5.0 M) at 90 ° for 30 min. The reaction products are separated from the Dynabeads by placing the tubes in a magnetic rack (Dynal MPC). After ethanol precipitation, telomerase reaction products are analyzed by denaturing 8% (w/v) polyacrylamide gel electrophoresis. The DNA ladders can be visualized by autoradiography or by phosphorimage analysis. Telomerase activity is quantified by densitometric analysis (ImageQuant; Molecular Dynamics) of the autoradiogram. S t r a t e g i e s to T a r g e t T e l o m e r e s The unique structure and dynamics of t-loops found at the ends of telomeres provide a number of opportunities to selectively disrupt processes associated with telomere maintenance mechanisms. The t-loop consists of a duplex 6-bp repeat (dATTGGG)n with a G-rich single-stranded end that invades and closes into the duplex region to form a secondary DNA structure of as yet undetermined complexity, although a D-loop has been reasonably postulated to form. 87 In addition to the nucleic acid component, a number of proteins, including TRF1, TRF2, and probably telomerase, are associated with the t-loop structure. TRF1 is associated 87 j. D. Griffith, L. Comeau, S. Rosenfield, R. M. Stansel, A. Bianchi, H. Moss, and T. de Lange, Cell 97, 503 (1999).

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585

t-loop

TelomereD-loop ~ binding proteins (e.g., TRF2)

O

DuplexDNA binding proteins

FIG.2. The t-loopand associatedproteinand secondaryDNA structures.87 with the duplex telomeric repeat region, and its binding is determined by posttranscriptional modification by tankyrase and poly(ADP-ribose) glycohydrolase (PARG). s8 TRF2 appears to be associated with the junction site between the invading single-stranded telomeric sequence and duplex DNA, although it is not restricted to this site (see Fig. 2). Because the single-stranded telomeric DNA sequence is sequestered in the junction with TRF2 and duplex DNA, it is unavailable for telomerase extension. Presumably, through as yet undefined events involving dissociation of TRF1 and TRF2 from the t-loop, the single-stranded telomeric end can become accessible to telomerase for reverse transcriptase activity, which leads to telomeric extension. Thus telomere extension by telomerase involves both structural and dynamic changes in the t-loop that go beyond the telomerase cycle of elongation and translocation. t-Loop Targets

The t-loop and postulated D-loop duplex-single-strand invasion junction are ideal targets for DNA-interactive agents. This is because the t-loop has a duplex telomeric repeat, providing multiple targets of the same sequence, and also specific secondary DNA structures associated with the invasion complex. This presents three unique opportunities to attain unusual specificity with DNAinteractive agents. TelomericDuplexDNA. Telomeric duplex DNA has a 6-bp repeating sequence (5'-TTAGGG)n, and the binding of TRF1 dimers may result in the curvature necessary to form the t-loop. The c-Myb-like domain of TRF1 bound to duplex DNA 89 shows major groove binding of the protein, leaving the minor groove accessible for drug modification. Thus drugs that interact in the minor groove of DNA and have 88M. K. Jacobsonand E. L, Jacobson,Trends Biochem. Sci. 24, 415 (1999). 89p. KOnig,L. Fairall, and D. Rhodes,Nucleic Acids Res. 26, 1731(1998).

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a sequence specificity that targets them toward telomeric sequences and distorts DNA structure may well disrupt the t-loop structure. Telomeric S i n g l e - S t r a n d e d D N A . At the very ends of telomeres are 150- to 200-base single-stranded DNA regions of the 3' (G-rich) strand. Because of the unique properties of G-rich sequences of DNA, they can form an unusual variety of secondary structures known as G-quadruplexes (Han and Hurley, 9° and references therein). During the cell cycle, most likely at the end of the S phase, it seems reasonable that this single-stranded end must become accessible to telomerase or to other factors prior to resequestration within the t-loop junction at the so-called D-loop site by TRF2 (see Fig. 2). Within this window the presence of low molecular weight "driver molecules" could facilitate the formation of secondary DNA structures such as G-quadruplexes (Fig. 3), making these ends unavailable for either telomerase or sequestration into the D-loop structure. A number of groups have been seeking to identify small organic molecules that bind to G-quadruplex structures with the aim of inhibiting telomerase and thus disrupting telomeres.91-93 Although the biological effects of G-quadruplexes might not be limited to telomerase and telomeres, 9° the intended targets of G-quadruplexinteractive agents at this stage are still telomerase and telomeres, primarily because of the lack of data concerning other cellular roles. Our laboratory, along with several others, has adopted a structure-based design and synthesis approach to the development of compounds that interact with G-quadruplexes, which has been successful. To date, several groups of compounds have been identified (Fig. 4), and their interaction with G-quadruplexes has been extensively studied. Anthraquinone analogs were originally developed as DNA-interactive agents cytotoxic to a range of tumor cell lines. 94 The 2,6-diamidoanthraquinones were shown to act as selective triplex-interactive compounds with reduced affinity for duplex DNA. 95 Molecular modeling studies predicted that they might bind to G-quadruplex structures by a threading intercalation model. 96 We first showed by 1H N M R that one of the analogs of the 2,6-diamidoanthraquinone BSU-1051 (Fig. 4) binds to and stabilizes G-quadruplex structures. We also demonstrated by 90H. Han and L. H. Hurley, Trends Pharm. Sci. 21, 136 (2000). 91 S. Sharma, E. Raymond,H. Soda, D. Sun, S. G. Hilsenbeck, A. Sharma, E. Izbicka,B. Windle, and D. D. Von Hoff, Ann. Oncol. 8, 1063 (1997). 92E J. Perry and L. R. Kelland, Exp. Opin. Ther. Patents 8, 1567 (1998). 93L. H. Hurley, R. T. Wheelhouse, D. Sun, S. M. Kerwin, M. Salazar, O. Yu. Fedoroff, E X. Han, H. Han, E. Izbicka, and D. D. Von Hoff, Pharmacol. Ther. 85, 141 (2000). 94M. Agbandje, T. C. Jenkins, R. McKenna, A. P. Reszka, and S. Neidle, J. Med. Chem. 35, 1418 (1992). 95I. Haq, J. E. Ladbury,B. Z. Chowdhry,and T. C. Jenkins, J. Am. Chem. Soc. 118, 10693 (1996). 96E A. Tanious, T. C. Jenkins, S. Neidle, and W. D. Wilson, Biochemistry 31, 11632 (1992).

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TARGETING TELOMERES AND TELOMERASE R

H

587

H

A

H--N

\

/

H

B

H

,

//

,.; 9:

H

H

/

R

C

A k

......'""T'" o,o,°I, o,oo 1 °oo°OO,°ooOO°i ° ....

.............! .....

7

1)

......"'"'T'""

.........."i

FIG. 3. G-tetrad and G-quadruplexes. (A) Four guanine residues forming a planar G-tetrad structure through Hoogsteen hydrogen bonding. (B) A parallel-stranded G-quadruplex model. (C) An intermolecular antiparallel G-quadruplex model. (D) An intramolecular foldover G-quadruplex model. Each parallelogram in (B), (C), and (D) represents a G-tetrad.

588

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0

0 II

"CH H "k"~CH2~ H

H N

CH20H H3C. IN+

)) 0 BSU-1051

CH3

=CH3 ~

H3C-

~/~

H3

-CH 3

" - . . ~ ' - ¢ ~ - " cH~ )CH3

TMPyP2

TMPyP4

PIPER

H2N~NH2

i-(, DODC

EtBr

FIG. 4. G-quadruplex-interactive compounds. BSU-1051: 2,6-diamidoanthraquinone; TMPyP4: tetra(N-methyl-4-pyridyl)porpbine; TMPyP2: tetra-(N-methyl-2-pyridyl)porphine; PIPER: N,N'-bis[2-(1-piperidino)ethyl]-3,4,9,10-perylenetetracarboxylic diimide; DODC: 3,3:-diethyloxyadicarbo cyanine; EtBr: ethidium bromide.

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using the direct telomerase assay that this compound inhibits telomerase through its interaction with the intramolecular G-quadruplex formed from the telomeric primer. 85 A number of anthraquinone analogs have been identified as interacting with G-quadruplexes so as to inhibit telomerase. 92 Some of them are among the most potent small-molecule inhibitors of telomerase reported to d a t e . 92'97 Porphyrins have long been of interest in photodynamic cancer therapy, largely because of their ability to accumulate to a greater extent in tumor tissues rather than in normal tissues. 98 The planar arrangement of the aromatic rings in porphyrin analogs led us to propose that such compounds might be able to bind to G-quadruplexes through interactive stacking with the G-tetrads. 99 By using spectroscopy, circular dichroism, and NMR, two groups have independently found that the porphyrin analog TMPyP4 (Fig. 4) binds to and stabilizes both parallel and antiparallel G-quadruplex structures. 99,1°° Work in our laboratory has further shown that TMPyP4 inhibits telomerase through G-quadruplex interaction, whereas its positional isomer TMPyP2 does not (Fig. 4). 99'101 A photocleavage assay revealed that the difference in G-quadruplex interaction between these two isomers results from their different binding sites in G-quadruplexes J °l Two different G-quadruplex-binding models have been proposed for porphyrins. One model, based on photocleavage data, predicts that porphyrin molecules stack externally to G-tetrads located at the ends of G-quadruplex. 101 The other proposes, on the basis of molecular modeling and stoichiometry measurements, that porphyrin molecules intercalate between two G-tetrads. 1°2 Although the spectroscopy data and molecular modeling studies support both models, photocleavage data provide the best experimental evidence for the external stacking model, l°J Comparative studies using tumor cell lines revealed that TMPyP4 has a higher slowing effect on cell growth than TMPyP2, which is in accord with the telomerase inhibition data.l°3,1°4 TMPyP4 also induces anaphase chromosomal bridges in sea urchins, whereas TMPyP2 does not. 1°4 This result indicates that G-quadruplexinteractive compounds might target the telomeres directly inside cells.

97 p. j. Perry, M. A. Read, R. T. Davies, S. M. Gowan, A. P. Reszka, A. A. Wood, L. R. Kelland, and S. Neidle, J. Med. Chem. 42, 2679 (1999). 98 j. j. Schuitmaker, E Baas, H. L. van Leengoed, R. W. van der Meulen, W. M. Star, and N. van Zandwijk, J. Photochem. Photobiol. B 34, 3 (1996). 99 R. T. Wheelhouse, D. Sun, H. Han, E X. Han, and L. H. Hurley, J. Am. Chem. Soc. 120, 3261 (1998). 10o N. V. Anantha, M. Azam, and R. D. Sheardy, Biochemistry 37, 2709 (1998). 101 E X. Han, R. T. Wheelhouse, and L. H. Hurley, J. Am. Chem. Soc. 121, 3561 (1999). 1o2 I. Haq, J. O. Trent, B. A. Chowdhry, and T. C. Jenkins, J. Am. Chem. Soc. 121, 1768 (1999). 103 E. Izbicka, R. T. Wheelhouse, E. Raymond, K. K. Davidson, R. A. Lawrence, D. Sun, B. E. Windle, L. H. Hurley, and D. D. Von Hoff, CancerRes. 59, 639 (1999). 1o4 E. Izbicka, D. Nishioka, V. Marcell, E. Raymond, K. K. Davidson, R. A. Lawrence, R. T. Wheelhouse, L. H. Hurley, R. S. Wu, and D. D. Von Hoff, Anti-Cancer Drug Des. 14, 355 (1999).

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Perylene as a potential G-quadruplex-interactive agent was first identified by using the structure-based design program DOCK. 1°5 On the basis of molecular modeling data, Fedoroff and co-workers designed and synthesized the N, N'-bis[2-(1-piperidino)ethyl]-3,4,9,10-perylenetetracarboxylic diimide (PIPER), which contains two positive charges at the ends (Fig. 4). This compound turned out to be a specific G-quadruplex-interactive agent that has little interaction with single- or double-stranded DNA. The NMR structure of a PIPER-G-quadruplex complex (Fig. 5, see color insert) the first definitive structure of a ligand~3quadruplex complex, showed that the binding mode of PIPER to G-quadruplexes is similar to that proposed by us for the porphyrins, that is, stacking external to G-tetrads.~°6 As with other G-quadruplex-interactive compounds, PIPER showed good telomerase and DNA polymerase inhibitory activities) °6 However, one of the most striking properties of this compound is its driver-like role in the assembly of G-quadruplex structures. PIPER not only converts the dimerization reaction of dimeric G-quadruplex formation from second order to first order, but it also increases the initial rate of formation about 100-fold at 10/zM concentration. 1°7 A similar effect has been observed in the facilitation of G-quadruplex formation by short G-rich oligonucleotides 1°8 and by the/~ subunit of the Oxytricha telomerebinding protein, which is reportedly a G-quadruplex chaperone) °9 This finding suggests that in addition to passive binding and stabilization, this type of compound may be able to play an inductive role in the formation of G-quadruplex structures within cells. Last, we have shown that PIPER, by binding to G-quadruplexes, inhibits their unwinding by Sgs 1, a G-quadruplex-specific helicase from yeast.lm° D-Loop Structure. It has been proposed that the invasion of the single-stranded telomeric end into the duplex telomeric region may involve a D-loop. It is also conceivable that an alternative secondary structure such as a G-quadruplex may be involved. Successful drug targeting of this DNA structure will be dependent on its detailed identification.

Posttranslational Modification of Telomere-Binding Proteins An attractive feature of the t-loop model is that it provides an explanation for how telomeres escape detection by DNA damage checkpoint proteins. However,

105H. Briem and I. D. Kuntz,J. Med. Chem. 39, 3401 (1996). 106O. Y. Fedoroff,M. Salazar, H. Han, V. V. Chemeris,S. M. Kerwin, and L. H. Hurley,Biochemistry 37, 12367(1998). 107H. Han, C. L. Cliff, and L. H. Hurley,Biochemistry38, 6981 (1999). 1o8y. Marco-Haviv,N. Baran, and H. Manor,J. Mol. Biol. 286, 45 (1999). lo9G. Fang and T. R. Cech,Biochemistry32, 11646(1993). 110H. Han, R. J. Bennett, and L. H. Hurley,Biochemistry39, 9311 (2000).

FIG. 5. NMR-derived structure of the 1:1 d(TAGGGTI'A)4-PIPER complex. Thymine is shown in cyan, adenine in purple, guanine in yellow, and PIPER in green. 1°5

NAD-~ Tankyrase I Nam*-" l

iiiiiii ADPRpolymers~

~ ADPR

LPARG I~-- H20

IL JIrj TRF1I

TRF2I

FIG. 6. A speculative scheme for the involvement of tankyrase and poly(ADP-ribose) glycohydrolase (PARG) in the disassembly and reassembly of a telomere loop structure,ss The black lines represent nontelomere DNA and the yellow and red lines represent C-rich and G-rich strands of telomere DNA, respectively. The G-rich overhang is stabilized by the telomere-specific proteins TRF1 and TRF2 to form a loop structure. Tankyrase-catalyzed addition of negatively charged ADP-ribose (ADPR) polymers to TRFI results in dissociation of TRF1 from telomere DNA and disassembly of the loop. In turn, PARGcatalyzed degradation of TRFl-associated polymers allows reassembly of the loop. Nam, Nicotinamide.

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it also seems logical that this structure must be disassembled and reassembled to allow telomere maintenance mechanisms to function. Figure 6 (see color insert) shows a scheme proposed by the Jacobsons 88 for the involvement of tankyrase and PARG in the disassembly and reassembly of a telomere loop structure. Tankyrase and Poly(ADP-ribose) Glycohydrolase. The discovery of a component of telomeres containing poly(ADP-ribose) polymerase (PARP) activity has provided evidence of the direct involvement of ADP-ribose polymer cycles in telomere maintenance mechanismsJ 11 This PARP, termed tankyrase, is a 142-kDa protein with a catalytic domain homologous to PARP-1, but otherwise it has a different domain structure. Most notably, tankyrase contains 24 tandem repeats of the ankyrin motif, a 33-amino acid sequence motif that often links membrane proteins to the cell cytoskeleton. In contrast to the better known PARP- 1 and PARP-2, tankyrase does not appear to require DNA for activity, and it interacts with and catalyzes polymer modification of the telomere-specific protein TRF1 in vitro. The domain structure for tankyrase and TRF1 and their suggested interactions are shown in Fig. 6. The structure of ADP-ribose polymers endows them with a high density of negative charges, making them effective polyanions that can compete with DNA for DNA-binding proteins. The tankyrase-catalyzed addition of negatively charged polymers to TRF1 is postulated to result in disassociation of TRF1 from telomere DNA and disassembly of the loop. In turn, PARG-catalyzed degradation of TRF 1-associated polymers would result in the removal of the competing polyanion and thus facilitate reassembly of the loop. The model shown in Fig. 6 is not meant to imply that ADP-fibose polymer cycles necessarily result in complete dissociation of the t-loop structure, only that they are capable of increasing access to telomere DNA. Telomere maintenance mechanisms that may involve tankyrase and PARG include telomere replication, telomere repair, telomere length stabilization by telomerase, stable integration of telomerase into the telomere, and telomere recombination.

Disruption of Telomere Maintenance Mechanisms by Targeting Tankyrase and Poly(ADP-ribose) Glycohydrolase. The unlimited proliferation potential of cancer cells is closely linked with their ability to maintain stable telomere structures. Thus, the disruption of telomere stability by targeting tankyrase suggests a new approach for cancer treatment. Most inhibitors developed to inhibit PARP-1 act at or near the nicotinamide region of the NAD-binding site of the enzyme, which is likely to have a similar structure in all PARPs. This is supported by the observations that 3-aminobenzamide inhibits the activity of each of the four known PARPs. H2-114 The discovery of multiple PARPs now prompts the design of 111 S. Smith, I. Giriat, A. Schmitt, and T. de Lange, Science 282, 1484 (1998). 112 G. de Murcia and J. Menissier de Murcia, Trends Biochem. Sci. 19, 172 (1994). 113 j. C. Am6, V. Rolli, V. Schreiber, C. Niedergang, E Apiou, P. Decker, S. Muller, T. Hoger, J. Menissier de Murcia, and G. de Murcia, J. Biol. Chem. 274, 17860 (1999).

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inhibitors specific for different PARPs for both elucidation of function and therapeutic targeting of ADPR polymer cycles. Finally, PARG may be an important therapeutic target. The dynamic nature of polymer cycles indicates that PARPs and PARG function coordinately; thus inhibition (or activation) of PARG is expected to affect functions modulated by polymer cycles. The low homology of PARG to other known proteins and its structurally unique substrate suggest that highly selective PARG inhibitors could be developed. Furthermore, because each of the known PARPs undergoes automodification, modulation of PARG might be an effective way to modulate these enzymes, too.

Assay Methodsfor G-Quadruplex-Interactive Compounds There are a number of assays to identify G-quadruplex compounds, including the standard telomerase assay, 115 a polymerase stop assay, 116 a helicase unwinding assay, 110 and a novel dialysis assay that simultaneously provides comparative binding affinity to a variety of nucleic acid structure types.liT Acknowledgments This research was supported by a National Cooperative Drug Discovery Grant from the National Cancer Institute (CA67760). Particular thanks are due to Dr. Mike Jacobson and Dr. Elaine Jacobson for insight into tankyrase and PARG as drug targets. We are grateful to other members of the NCDDG group for valuable discussions, and to Dr. David M. Bishop for preparing, proofreading, and editing the manuscript and figures.

114 W. A. Kickhoefer, A. C. Siva, N. L. Kedersha, E. M. Inman, C. Ruland, M. Streuli, and L. H. Rome, J. Cell Biol. 146, 917 (1999). 115 D. Sun, L. H. Hurley, and D. D. Von Hoff, BioTechniques 25, 1046 (1998). 116 H. Han, M. Salazar, and L. H. Hurley, Nucleic Acids Res. 27, 537 (1999). 117 j. Ren and J. B. Chaires, Biochemistry 38, 16067 (1999).

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[30] Rapid, High-Throughput Engineering of Sequence-Specific Zinc Finger DNA-Binding Proteins By MARKISALANand YEN CHOO Introduction Zinc fingers engineered to recognize predetermined DNA sequences have been used to control gene expression and also to target catalytic domains, such as restriction enzymes, methylases, and integrases, to specific regions of DNA (reviewed by Choo and Isalanl). The use of these proteins as transcription factors alone has potential applications in functional genomics, gene target validation, engineering of animals or plants with improved phenotypes, and human therapy. With the widespread utility of these domains now evident, and particularly for use as bespoke tools in the postgenomic era, it has become increasingly important to devise a high-throughput, rapid, and reliable engineering procedure capable of producing high-quality zinc fingers that bind to any given DNA sequence. Although we, and others, have been studying the interactions of zinc fingers with DNA in order to classify the protein-DNA contacts into a "recognition code" that could be used for zinc finger design, this code does not allow the engineering of zinc fingers that bind to any given DNA sequence. Quite separately, we believe that the quickest and certainly the most reliable way of engineering sequence-specific zinc finger proteins is by using selection methods such as phage display, and not by use of rational design. This is because selection in the presence of competitor DNA necessarily yields the tightest and most specific DNA binders from a large collection of zinc finger molecules. A correctly constructed library contains all the zinc finger candidates that could be designed using the recognition code, alongside many other candidates that bind DNA using more obscure recognition modes that are not understood at present. In addition, if the phage selection method is well conceived, it can yield zinc finger proteins even quicker than rational design, as a few rounds of phage selection can be performed more quickly than the cloning and subsequent validation of rationally designed zinc finger candidates. We have used a new, rapid, and widely applicable zinc finger-engineering strategy that produces three-finger domains that bind a wide variety of DNA sequences. 2 Here we describe this engineering strategy, placing particular emphasis on methodological aspects, in order to illustrate the increase in the speed and throughput of zinc finger engineering. I y. C h o o a n d M. D. Isalan, Curr. Opin. Struct. Biol. 10, 411 (2000). 2 M. D. Isalan, Y. C h o o , a n d A. Klug, Nat. Biotechnol., in press (2001).

METHODSIN ENZYMOLOGY,VOL.340

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Zinc Finger Engineering Strategies Based on Phage Display Phage display methods have been widely used to select novel zinc finger DNAbinding domains. 3'4 Until more recently, however, the reported methods have generally been restricted to randomizing DNA-contacting residues in one zinc finger, within the context of a three-finger scaffold (reviewed in Choo and Isalan 1). Different engineering strategies have been developed in order to produce entire zinc finger DNA-binding domains, for instance, domains containing three zinc fingers. These strategies have either resulted in proteins that recognize a small subset of DNA sequences, or else have been too complex for high-throughput applications. In one strategy, zinc fingers selected in parallel are spliced together in a second step to produce multifinger domains that bind longer DNA sequences. 5,6 Although this technique is relatively fast and straightforward, it is not able to produce zinc fingers that bind to a wide variety of DNA sequences. To date, the proteins engineered by this method bind only to DNA sequences of the form (GNN)~, i.e., sequences in which guanine occurs at every third base position. In a second strategy of engineering zinc finger DNA-binding domains, selections of single zinc fingers are carried out in series, with the addition of every finger representing a "walk" across the DNA target sequence. 7 This protocol is capable of producing zinc finger domains that bind to more diverse DNA sequences, although the iterative cloning of libraries makes it fairly cumbersome and generally unsuitable for high-throughput applications. B i p a r t i t e - C o m p l e m e n t a r y M e t h o d of Zinc F i n g e r E n g i n e e r i n g

Strategic Considerations in Designing Protocol In devising an improved, rapid, and convenient high-throughput zinc finger engineering protocol, we were conscious of three major considerations. First, in order to recognize a DNA sequence most effectively, zinc fingers use an overlapping (synergistic) mode of DNA binding. 8 This mode is particularly important in engineering zinc fingers that bind to DNA sequences that are not guanine rich. 9 To take advantage of this synergistic effect, an effective zinc finger engineering strategy must simultaneously select the appropriate amino acids at the 3 E. J. Rebar, H. A. Greisman, and C. O. Pabo, Methods Enzymol. 267, 129 (1996). 4 M. D. Isalan and Y. Choo, "Phage Display of Zinc Fingers and Other Nucleic Acid Binding Motifs." Oxford University Press, Oxford, 2000. 5 y. Choo, I. Sanchez-Garcia, and A. Klug, Nature (London) 372, 642 (1994). 6 D. J. Segal, B. Dreier, R. R. Beerli, and C. E Barbas, Proc. Natl. Acad. Sci. U.S.A. 96, 2758 (1999). 7 H. A. Greisrnan and C. O. Pabo, Science 275, 657 (1997). 8 M. Isalan, Y. Choo, and A. Klug, Proc. Natl. Acad. Sci. U.S.A. 94, 5617 (1997). 9 M. lsalan, A. Klug, and Y. Choo, Biochemistry 37, 12026 (1998).

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cooperating positions in any two adjacent zinc fingers.9 Examples of these potential cooperating contacts are shown as arrows that contact the base pair 5X-5'X, in Fig. 1. Second, the engineering protocol must not involve serial library constructions, which are cumbersome and time consuming, but rather should rely mainly on large premade libraries. Once these libraries are constructed, it would be relatively easy to select multiple zinc fingers in parallel, making the procedure suitable for high-throughput protein engineering. Third, the output of the protocol, that is, the product zinc finger protein, must be in a form that allows rapid validation of the activity of the engineered proteins. We have found that the quickest way of achieving this is by carrying out an enzymelinked immunosorbent assay (ELISA) on DNA-binding zinc fingers displayed on phage. We have been using this technique in conjunction with band shift and cell culture analysis of subcloned zinc finger proteins and so far have not observed any inconsistencies between the three methods.

Overview of Bipartite-Complementary Library Method This method utilizes two premade sublibraries (Lib 12 and Lib23) based on the three-finger DNA-binding domain of Zif268. Each library contains randomizations in the u-helical DNA-contacting residues of a pair of adjacent fingers in order to capture cooperative interactions (Fig. 1). Hence Lib 12 contains randomizations in all the base-contacting positions of F1 and certain base-contacting positions of F2, while Lib23 contains randomizations in the remaining base-contacting positions of F2 and all the base-contacting positions of F3 (Fig. 1). Selections from these libraries can be carried out in parallel, to yield two DNA-binding domains, which each recognize new 5-bp sequences. The selected portions of these domains are complementary, in that they may be recombined to produce a novel peptide that recognizes a composite 9- or 10-bp sequence. An overview of the steps required to engineer zinc fingers, using the bipartite-complementary method, is shown in Table 1. This protein engineering strategy has been experimentally validated by the selection of a variety of zinc finger peptides against DNA sequences found in the promoter of human immunodeficiency virus type 1 (HIV-1). 2

Practical Considerations in Library Design The bipartite-complementary library system is made practically feasible by several important features. First, selective randomization of the zinc finger et helix (Figs. 1 and 2) allows simultaneous randomization of adjacent zinc fingers, while constraining the library size to within the practical cloning limits of phage display experiments. This is possible because no sequence space is wasted on amino acids that are unlikely to confer sequence specificity. A large body of zinc finger

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ENZYMOLOGY AND BIOLOGICAL APPROACHES

[30[

w

io

3"

F1

F2

F3

1123456 -1123456 -1123456 ASAALIA ASAALST RSDERKR D DD TE DRDD-H HH K H HH N KN N N KN ,(-Lib12 Q NS Q Q NS R QT R R QT T RV T T RV S V S -1123456

RSDELTR

Lib23

-112345 6 -1123456 RSDHLS A AKAARIA E DNDD KE •f

K N Q R T V

HRHH TK NSKN N Q NS Q R QT R T RV T S V

FIG. 1. Overview of library composition in a bipartite strategy. The final recombined zinc finger products are represented schematically (above), with residues derived from randomized positions shown as circles (numbered relative to their helical positions). The dashed line (indicated by *R) indicates the

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SEQUENCE-SPECIFIC ZINC FINGERS

597

TABLE I ENGINEERINGSEQUENCE-SPECIFICDNA-BINDINGZINCFINGERSBY BIPARTITE-COMPLEMENTARYSYSTEMa Day 1-5

Procedure Phage selection

6

ELISA

7

Recombination

8-9 10-14

Cloning into phage vector Product evaluation

Notes Parallel selection of complementary fingers from Libl2 and Lib23 3-5 rounds 1 round per day Test binding of individual clones obtained from Libl2 and Lib23 Select clones with best affinity/specificity for recombination Recover complementary genes from Libl 2 and Lib23 by PCR Digest with DdeI, religate, recover recombinant cassettes by selective PCR Ligate recombined cassettes into phage vector and transform host bacteria Test binding of individual clones to full-length target DNA by ELISA Carry out further rounds of selection if required DNA sequencing of selected clones

a The approximate time scale for carrying out these procedures is indicated in days.

D N A - b i n d i n g data is n o w available, f r o m w h i c h it is apparent that o n l y a subset o f the 20 a m i n o acids is m o s t l y responsible for D N A recognition 9-n (Fig. 1). A n o t h e r important feature o f the system allows the r e c o m b i n a t i o n in v i t r o o f the c o m p l e m e n t a r y portions o f the two libraries, without the n e e d for purification steps o f any kind (Fig. 4). To do this w e have taken advantage o f selective p o l y m e r a s e chain reaction (PCR), so as to amplify only the products o f r e c o m b i nation. P C R with e n z y m e s lacking 5'---> Y - e x o n u c l e a s e activity cannot p r o c e e d i f primers contain one or m o r e 3' m i s m a t c h e s against their template-binding sites. 10 y. Choo and A. Klug, Proc. Natl. Acad. Sci. U.S.A. 91, 11163 (1994). 11 y. Choo and A. Klug, Curr. Opin. Struct. Biol. 7, 117 (1997).

recombination breakpoint in the two libraries. Recombination is carded out by digestion and religation of both Libl2 and Lib23. The composition of each helical position is indicated in the column of amino acids. Each library contains an invariant wild-type Zif268 region that acts as an anchor to the system, binding a fixed, G-rich, DNA sequence. 12 After selection, the two complementary variable regions of Libl2 and Lib 23 are recombined to make a full-length protein that potentially binds 10 bp of DNA: bases 2x-n'x. of a DdeI restriction enzyme site (CTGAG) that encodes Leu-Ser in positions 4 and 5 of F2 ~

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ENZYMOLOGY AND BIOLOGICAL APPROACHES

A

~

c~-helix -1 1 2 3 4

i

I

Minicassettel

B

~

x

lrl

[30]

linker

4 5 6 I

Minicassette2

x

x

I

Minicassette3

Minicasseffe 1 5'-GCGGAAGAGAGGCCCTACGCATGCCCTGTCGAGTCCTGCGATCGC CGCCTTCTCTCCGGGATGCGTACGGGACAGCTCAGGACGCTAGCGGCGA-5

MinicasseRe 2 F1 - 1 +1 +2 +3 CGCTTTTCTxxxTCGxxxxx AAAGAxxxAGCxxxxxGGAAT

-TOP -BOTTOM Code for:

TOPCGCTTTTCTcrsTCGgmcca CGCTTTTCTcrsTCGgmcav CGCTTTTCTcrsTCGgmcgh CGCTTTTCTcrsTCGmrsca CGCTTTTCTcrsTCGmrsav CGCTTTTCTcrsTCGmrsgh CGCTTTTCTrmcTCGgmcca CGCTTTTCTrmcTCGgmcav CGCTTTTCTrmcTCGgmcgh CGCTTTTCTrmcTCGmrsca CGCTTTTCTrmcTCGmrsav CGCTTTTCTrmcTCGmrsgh

BOTTOMTAAGGtggkcCGAsygAGAAA TAAGGbtgkcCGAsygAGAAA TAAGGdcgkcCGAsygAGAAA TAAGGtgsykCGAsygAGAAA TAAGGbtsykCGAsygAGAAA TAAGGdcsykCGAsygAGAAA TAAGGtggkcCGAgkyAGAA TAAGGbtgkcCGAgkyAGAAA TAAGGdcgkcCGAgkyAGAAA TAAGGtgsykCGAgkyAGAAA TAAGGbtsykCGAgkyAGAAA TAAGGdcsykCGAgkyAGAAA

-i R,Q,H R,Q,H R,Q,H R,Q,H R,Q,H R,Q,H N,D,A,T N,D,A,T N,D,A,T N,D,A,T N,D,A,T N,D,A,T

+2 D,A D,A D,A RQHKSN RQHKSN RQHKSN D,A D,A D,A RQHKSN RQHKSN RQHKSN

+3 H N,S,T V,A,D H N,S,T V,A,D H N,S,T V,A,D H N,S,T V,A,D

Minicassette 3 F1 +4 +5 +6 5'- CCTTAxxxxxCATATCCGCATCCACA xXXXxGTATAGGCGTAGGTGTGGCC TOPCCTTayccrgCATATCCGCATCCACA CCTTaycghaCATATCCGCATCCACA CCTTaycamsCATATCCGCATCCACA

-TOP -BOTTOM

Code for:

BOTTOMCCGGTGTGGATGCGGATATGcyggr CCGGTGTGGATGCGGATATGtdcgr CCGGTGTGGATGCGGATATGsktgr

+5 I,T I,T I,T

+6 R,Q V,A,E K,N,T

Key to Randomised Nucleotides m=A/C r=A/G w=A/T

s=G/C y=T/C k=T/G

b=C/G/T

v=A/C/G

n=A/C/G/T

h=A/C/T d=A/G/T

FIG. 2. Construction of a gene cassette encoding a zinc finger phage display library with "smart" randomizations. (A) The scheme used to generate selective randomization throughout the u helix of a zinc finger. A set of complementary oligonucleotides is used to construct a series of"minicassettes" that

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SEQUENCE-SPECIFIC ZINC FINGERS

599

The two complementary libraries are therefore designed with unique sequences at their 5' and 3' termini, and the corresponding primers are used to amplify any recombinants of the two libraries. Finally, it should be noted that the selection procedure is amenable to a microtiter plate format, so that selections and most subsequent manipulations can be carried out by liquid-handling robots. This allows substantial automation of the zinc finger engineering procedure. The ability to carry out selective PCR implies that the protocol can even be adapted to selecting complementary library portions in the same tube or well. For example, both universal libraries can be coscreened in a single well, thereby increasing the efficiency of high-throughput applications. The output of such combined selections can be monitored either by selective PCR, or by ELISA of samples of isolated clones, as described below. Library Construction Gene inserts for phage libraries may be constructed by end-to-end ligation of selectively randomized dsDNA "minicassettes," made individually by annealing complementary template oligonucleotides (Fig. 2). The resulting genes are amplified by PCR and encode zinc fingers in a suitable reading frame for cloning and expressing as fusions to the phage minor coat protein, pIII. Our work uses the DNA-binding domain of the transcription factor Zif268 as a scaffold, because it contains three Cysz-His2 zinc fingers whose mode of binding is well understood.12,13 To selectively randomize the ot helix of a zinc finger, the coding region is synthesized using DNA minicassettes, such that helical positions - 1 through 4 are encoded by one cassette (minicassette 2; Fig. 2), whereas positions 4 through 6 are encoded by another cassette (minicassette 3; Fig. 2). These double-stranded cassettes are synthesized with complementary overhangs that anneal through the codon for the fourth or-helical residue, which is invariant. Each cassette actually comprises a library of oligonucleotides synthesized with appropriate codon randomizations so as to encode a given subset of amino acids. Figure 2A shows that a "smart library" of zinc finger genes can be created with three minicassettes: the first cassette is a single sequence and encodes the invariant/3-sheet region, 12N. R Pavletich and C. O. Pabo, Science 252, 809 (1991). 13M. Elrod-Erickson, M. A. Rould, L. Nekludova,and C. O. Pabo, Structure 4, 1171 (1996).

can be annealed and ligated togetherto constructthe randomizedportionof the gene. Note that several additional minicassettes, encoding other fingers, need to be constructed to achieve the randomization scheme outlined in Fig. 1. After ligation of all the minicassettes, the full-lengthconstructis recovered by PCR, using primers that containSfiI/Notl restriction enzymesites for cloninginto a phage vector.(B) Examples of the oligonucleotidesused to achieve the selective randomizationof a single zinc finger.

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whereas the second and third cassettes contain randomizations of the ct helix. Figure 2B shows that each of the library minicassettes comprises numerous oligonucleotides created through a limited number of solid-phase syntheses: minicassette 2 requires oligonucleotides from 12 pairs of syntheses, whereas minicassette 3 requires oligonucleotides from 3 pairs of syntheses. Each oligonucleotide synthesis is designed to introduce limited variability into each cassette; the library complexity is increased by the use of oligonucleotides from multiple syntheses and by the combination of the two minicassettes. Library Construction Protocol. Single-stranded template oligonucleotides are phosphorylated in a kinase reaction prior to assembly (100 pmol of each oligonucleotide in 10/zl of I x T4 kinase buffer, containing 1 mM dATP and 10 U of T4 polynucleotide kinase, 37 °, 1 hr). Complementary single-stranded template oligonucleotides are annealed pairwise to form double-stranded minicassettes (Fig. 2): 100 pmol of each oligonucleotide (or, for smart randomization, 100 pmol of each strand mixture) is mixed in 1 x T4 ligase or kinase buffer, to a final DNA concentration of 10 pmol//zl. Annealing is achieved by heating to 94 ° and then cooling slowly (~ 1 hr) to room temperature. The resulting double-stranded DNA (dsDNA) minicassettes are combined and ligated by adding an equal volume of l x T4 ligase buffer and 8 / t l (3200 U) o f T 4 ligase per 100/zl (16°, 20 hr). Full-length genes are amplified by PCR from the ligation mixture with primers that introduce NotI and Sill restriction sites for cloning into phage vector Fd-TETSN.l° Thorough digestion with these endonucleases is essential for high-efficiency ligation into similarly prepared phage vector (200 U of enzyme per 40/zg of DNA, with 8 hr of incubation in appropriate temperatures and buffers, adding enzymes in stages at 2-hr intervals). Typically, 1/zg of pure phage vector is ligated with a 5-fold excess of gene cassette insert (1 x T4 ligase buffer, 3/zl of T4 ligase, 30-~1 total volume, 16°, 20 hr). Ligation reactions are prepared for electroporation by washing twice with an equal volume of chloroform and precipitating by adding a 1/10 volume of sodium acetate (pH 5.5) and 3 volumes of ethanol. 14 DNA pellets are washed with 70% (v/v) ethanol and resuspended in sterile water to a final concentration of 200 ng//zl. The phage library is cloned by electroporation of recombinant vector into a suitable strain of Escherichia coli, such as TG1. Typically, 0.5/zg of recombinant phage vector can be used with 100/zl of electrocompetent cells, 15 yielding up to --d06 library transformants (2-ram path cuvette, 2.5 kV, 25/zF, 200 f2). After pulsing, cells are immediately resuspended in 1 ml of SOC medium (see Appendix) and incubated without shaking (37 °, 1 hr). Fd-TET-SN confers tetracycline resistance, allowing positive selection of bacterial transformants by plating on 2x YT-agar plates, containing tetracycline (15/zg/ml) (37 °, 16 hr). 14j. Sambrook,E. F. Fritsch,and T. Maniatis, in "MolecularCloning: A LaboratoryManual."Cold Spring HarborLaboratoryPress,Cold SpringHarbor,New York, 1989. 15W. J. Dower,J. E Miller,and C. W. Ragsdale,NucleicAcids Res. 16, 6127 0988).

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Selection from Libraries Selection Protocol. Phage are prepared for selections by scraping library transformant colonies into 2× YT liquid medium, containing tetracycline (15 #g/ml) and 50/xM zinc chloride, and incubating in an orbital shaker (30 °, 220 rpm, 16 hr). The culture supernatant containing phage particles is collected by centrifugation (3700g, 15 min). For each selection, the supernatant is diluted 1 : 10 in 1 ml of phosphate-buffered saline (PBS) containing 1% (w/v) Marvel, 1% (v/v) Tween 20, and sonicated salmon sperm DNA (20 #g/ml). Although these conditions generally work well, using specific competitor DNAs and varying binding conditions can have a significant impact on the outcome of individual selections. The phage mixtures are added to streptavidin-coated tubes or wells (Roche, Rahway, NJ) that have been precoated with biotinylated target DNA (made by annealing two complementary oligonucleotides, one of which is biotinylated). The selection procedure described here is for use with streptavidin-coated tubes and a total reaction volume of 1 ml (however, by scaling down to a 200-/zl volume the process can easily be adapted to a 96-well microtiter plate format). Typically, 1 pmol of target DNA is coated on each tube, in 50 #1 of PBS-Zn (--~20°, 15 min). The addition of 1 ml of 4% (w/v) Marvel blocking agent helps to reduce nonspecific binding. After blocking (-,~20°, 1 hr) tubes are emptied, refilled with 1 ml of phage binding mixture, and left to equilibrate (~20 °, 1 hr). Washing steps are then carried out to remove all unbound phage [20 washes with 1 ml of PBS-Zn containing 2% (w/v) Marvel, 1% (v/v) Tween 20, followed by one wash with PBS-Zn alone]. Retained phage are eluted in 100 #1 of 0.1 M triethylamine, removed to a separate container, and immediately neutralized with an equal volume of 1 M Tris-HCl, pH 7.4. Eluted phage can be stored at - 2 0 °. Fifty microliters of the eluted phage are used to infect 0.3 ml of a logarithmic phase culture ofE. coli TG1. The bacteria are derived from colonies grown freshly on M9 minimal agar as this ensures expression of the F' pilus, which is required for phage infection. Bacteria are infected by the addition of phage and incubating without shaking (37 °, 1 hr). Bacteria are then transferred to 2-5 ml of 2× YT containing tetracycline (15 #g/ml) and 50 #M zinc chloride and grown, as before, to prepare phage supematant (30 °, 220 rpm, 16 hr). Subsequent rounds of selection are carried out as described above, although the amount of competitor DNA may be increased in later rounds to increase the stringency of selection. Three to five rounds of selection are usually sufficient to enrich target-binding clones; however, the progress of individual selections may be monitored by plating out infections to estimate phage yield after each round of selection (Fig. 3; see Related Procedures in the Appendix). It should be noted, however, that phage yield does not by itself give an indication of the quality of the selected zinc fingers because fast-growing nonspecific binders can dominate selections. After selections, pools of complementary phage can be recombined directly without preliminary analysis; however, we recommend that individual clones first

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3500 3000 -

=zl. 2500

~

Libl211

2000

Lib23 [ ]

1500 1000 500 0 RI

R2

R3

R4

R5

Round FIG. 3. Phage titers of selections from Libl2 and Lib23, using two DNA target sites, 5'-GCG GCT CTC-31 and 5'-GGA GCG GCG-3r (selected sequences shown in boldface). After each round of selection (R1-5), neutralized phage eluate was used to infect an excess of logarithmic phase E. coli TG1 cells. Serial dilutions of these infections were plated out on TYE-TETplates. Colonies that grew after incubation (15 hr, 37°) were counted to estimate phage yield. Phage yields are shown plotted vertically as number of colony-formingunits per microliter of selection eluate. undergo testing for binding to their respective 5-bp target sites, using an ELISA (see below).

Recombination of Complementary Libraries After three to five rounds of selection, PCR is used to amplify zinc finger genes from Libl2 and Lib23, and the selected portions are recombined by digestion and religation at a unique DdeI restriction site. The DNA template for the initial PCR can be derived from the entire pool of selected phage, or preferably from one or several individual clones previously validated for DNA binding. Individual clones are chosen on the basis of sequence-specific DNA binding, verified by phage ELISA (see below). Recombination begins with a PCR step to amplify all the zinc finger genes present, using primers that are external to the SfiI/NotI gene cassette. One microliter of phage supernatant(s) provides sufficient DNA template on lysing the virions (94 °, 9 min). The PCR itself comprises 30 cycles, using Taq polymerase. After PCR, 50/zl of the reaction is phenol/chloroform extracted, diluted in 2 volumes of DdeI buffer (New England BioLabs, Beverlay, MA), and digested with 60 U of DdeI per 150 #l. The enzyme is heat inactivated (65 °, 20 min) and the reactions are precipitated with a 1/10 volume of sodium acetate (pH 5.5) and 3 volumes of ethanol. Each DNA pellet is resuspended in 10 ttl of T4 DNA ligase buffer (New England BioLabs) and 400 U o f T 4 ligase is added per reaction (20 °, 15 hr). Recombinants, comprising the selected portions of Libl2 and Lib23, are amplified selectively by PCR from 1/zl of the ligation mixture (Fig. 4). For optimal selective amplification, only 20 cycles of PCR are carried out with

[30]

SEQUENCE-SPECIFIC ZINC FINGERS ,,~

W

Ub12

i

I

"

\

603

Ub23

I

W

I

Dde I

Dde I

x.

M

I PCR, Dde I cut, ligation W

I

W

M

W

I

I

M ,~

I W

\

M

SelectivePeR (M/M primers)

Recombinedproduct FIG.4. SelectivePCR to recovercorrectlyrecombinedportionsof the two complementarylibraries. PCR primers that bind the 3t and 5~ends of zinc finger genes are designedto bind either the wildtype Zif268 codingsequence (W) or a mutantDNA sequencecontainingalternativecodons (M). The mutations affect priming events in the PCR but do not alter the protein sequence when translated. The two selected portions of the bipartite libraries(shown in black and gray) containmutant primer recognitionsites.Therefore,two mutant(M/M)primersare requiredto recoverthe correctrecombinants from the mixedpool of ligationproducts. a nonproofreading DNA polymerase such as Taq. It is worth noting that enzymes such as Pfu, which contain 5'--->3' proofreading activity, erode the DNA ends, thus eliminating selective priming. PCR products are cloned back into phage vector as described above (see Library Construction, above). The resulting products of the recombination step, which often comprise pools of clones (defined by the various permutations in which portions from the two libraries can recombine), may be reselected by phage display so that the optimal zinc finger phage is isolated. Alternatively, a number of different clones may be screened for binding to the target DNA sequence.

Coselection of Two Complementary Libraries In devising the present strategy we initially envisaged the simultaneous selection of complementary zinc finger domains from a mixture of L i b l 2 and Lib23

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in a single tube. This should be possible as different DNA sequences are used to select zinc fingers from the two libraries, meaning that these would not compete with each other for the DNA used in their selection. We further imagined that the mixture of selected zinc fingers could then be recombined directly to yield product zinc fingers (without previously carrying out ELISAs to verify the presence of binders from each library). Although this approach is feasible in theory, and indeed in practice we have used it to engineer a few DNA-binding domains, we have found that physically separating the selections from the two libraries increases the probability that the engineering will be successful. It often emerges that after coselection of the two libraries, the large majority of selected phage are from just one of the two libraries (this can be checked either by phage sequencing or by selective PCR) and therefore recombinant DNA-binding domains cannot be produced. The reason for this is that certain phage clones have a distinct growth advantage that contributes to the selection process (alongside DNA binding), allowing a few clones to dominate the selection. Thus, separating the selections from the two libraries increases the chance of obtaining phage from both libraries.

Phage Enzyme-Linked Irnmunosorbent Assay Determination of D N A - B i n d i n g S p e c i f i c i t y a n d Affinity Phage ELISA may be used to assess the DNA-binding affinity and specificity of a zinc finger phage clone at any stage of the engineering procedure, but particularly prior to recombination of clones from the two libraries, and at the end point, at which time it is important to validate DNA recognition by the product. By using closely related variants of the DNA binding site, or a variety of binding-site concentrations, it is possible to obtain either a profile of the sequence discrimination and/or the apparent Kd values of interaction.

Enzyme-Linked Immunosorbent Assay Protocol Single bacterial colonies, infected with phage clones derived from library selections, are picked from agar plates. Colonies are transferred to wells in sterile, round-bottom, 96-well plates containing 150/zl of 2 x YT, tetracycline (15/zg/ml), and 50/zM zinc chloride. Plates are incubated with orbital mixing (30 °, 225 rpm, 16 hr). Phage supematant is prepared from the bacterial cultures by centrifuging the 96-well plates (3700g, 15 min), in a suitable swinging-bucket centrifuge. A phage binding mixture is prepared by diluting supernatant 1 : 10 in 1 ml of PBSZn containing 2% (w/v) Marvel, 1% (v/v) Tween 20, and sonicated salmon sperm DNA (20/zg/ml). DNA target sites for ELISA can be made by synthesizing and annealing two complementary oligonucleotides, one of which is biotinylated. These targets are

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SEQUENCE-SPECIFIC ZINC FINGERS

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bound to streptavidin-coated wells in 96-well microtiter plates (Roche). Typically, 0-5 pmol of DNA site is bound in 50/zl of PBS-Zn ('--20°, 15 min). Wells are then blocked (20 °, I hr), using 150 #1 of PBS-Zn containing 4% (w/v) Marvel. Blocking solutions are discarded and 50 #1 of the phage binding mixture is applied to each well. Binding mixtures are left to equilibrate (~20 °, 1 hr) and are then discarded. Wells are washed seven times with 200/~1 of PBS-Zn containing 1% (v/v) Tween 20. A further three washes are carried out with 200 #1 of PBS-Zn alone. Fifty microliters of PBS containing 2% (w/v) Marvel and a 1 : 5000 dilution of horseradish peroxidase (HRP)-conjugated anti-M 13 IgG antibody (Pharmacia Biotech, Piscataway, N J) is added to each well. After incubating (~20 °, 1 hr), antibody binding reactions are discarded and the wells are washed three times with 200 #1 of PBSZn containing 0.05% (v/v) Tween 20. A further three washes are carried out with 200/zl of PBS-Zn alone. ELISAs are developed with 100 #1 of HRP substrate, such as the 3',3',5',5'-tetramethylbenzidine (TMB)-based, ELISA developer solution described below. Colorimetric reactions are stopped after ~5 min by adding 100 #1 of 1 M H2SO4. ELISA signals are quantitated with a spectrophotometer fitted with a microtiter plate reader. Signals should be compared with negative control wells that either lack target DNA or have been coated with unrelated DNA sites.

Scanning Mutagenesis Binding Assays In this phage ELISA variation, each position in the ~10-bp binding site is sequentially mutated in a series of oligonucleotide targets in order to help assess the DNA sequence specificity of the engineered zinc finger protein. The single transition mutations used in the control DNA sites are more stringent controls of DNA-binding specificity than their counterpart transversion mutations. This is primarily because of the size and charge similarities of purines and pyrimidines (the purines A and G are both larger and partially negatively charged, whereas the pyrimidines C and T are smaller and partially positively charged). Transition mutations therefore represent a good test of the ability of zinc fingers to differentiate between closely related DNA sites. Typically, binding is carried out at a DNA concentration of 5-10 riM, which is close to the expected apparent Kd of zinc fingers. In the example shown in Fig. 5, a clone engineered from the bipartite-complementary library against the target site GGA-GCT-CTC is tested against a series of transition mutant sites. Although the construct binds well to its own target site, it appears that discrimination is not flawless throughout the binding site. In particular, the assay reveals that a C-+T transition (Fig. 5; column 8) is tolerated by the protein. Otherwise, however, the overall binding specificity is shown to be fairly good. This type of assay provides an indication of the DNA-binding specificity for a given protein but obviously does not allow a comprehensive study. To achieve

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Selection target GGA F3

GCT F2

CTC F1

I Phage ELISA Well

1

2

3

4

5

6

7

8

9

10

G G A G C T C T C

a

G A G C T C T C

G a A G C T C T C

G G g G C T C T C

G G A a C T C T C

G G A G t T C T C

G G A G C e C T C

G G A G C T t T C

G G A G C T C e C

G G A G C T C T t

ii@@@@@@ii@@

I Plot ELISA Well 0.6

~

1

2

3

4

5

6

7

8

9

10

T

G

G

A

G

C

T

C

T

C

E

liiliilll

0.3

0.0

T

a

a

g

a

t

c

t

c

t

FIG. 5. Scanning mutagenesis ELISA binding assay. A zinc finger phage clone was tested for binding both to its full target site (GGA-GCT-CTC) and to sites containing single transition mutations. Each well of tfie ELISA microliter plate is coated with a DNA sequence that contains a different transition mutation (shown as a lower-case nucleotide). DNA binding is revealed by the ELISA absorbance reading (A45o-A650),which is plotted vertically in the graph. All binding reactions were carried out at a DNA concentration of 8 nM.

[301

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this we have developed microarrays of double-stranded DNAs printed on glass slides. 16 These microarrays can be used to perform phage ELISA in a procedure similar to that described above, but the high density of DNA spotting allows parallel sampling of many thousands of DNA sequences. A u t o m a t i o n of Zinc F i n g e r E n g i n e e r i n g P r o c e s s Most of the steps of the engineering process--bacterial growth, phage selection, colony picking, phage ELISA, PCR, and cloning--can be automated by using commercially available instruments. At Gendaq (London, UK) we use 96or 384-well microtiter plates to carry out phage selections, ELISA reactions, and PCR preparation, on a liquid-handling robotic platform. A robotic arm shuttles the microtiter plates between a pipetting station, a plate hotel, a plate washer, a spectrophotometer, and a PCR block. A colony-picking robot is used to inoculate microcultures of bacteria in microtiter plates in order to provide monoclonal phage for ELISA. A robot that interfaces with the spectrophotometer is capable of returning to the liquid culture archive in order to "cherry-pick" particular clones that are suitable for recombination, or that should be archived. A bar-coding system is used to keep track of the various plates used for phage selections, phage ELISAs, or for archiving interesting clones. Concluding Remarks Previous strategies for engineering zinc fingers by phage display were labor intensive and/or limited in their utility.l If zinc fingers were selected in parallel and subsequently assembled into a multifinger domain, the DNA specificity of the domain was limited to binding only a guanine-rich binding site. 6 If, on the other hand, the selection of zinc fingers was carried out serially, the protein engineering procedure became laborious. 7 The "bipartite library" protocol described here solves the problem of engineering sequence-specific zinc fingers to bind diverse DNA sequences, and at the same time increases the speed and throughput of the engineering process. Appendix: Solutions and Related Procedures

Solutions Enzymes and buffers: T4 DNA kinase, T4 DNA ligase, Taq DNA polymerase, and SfiI, Nod, and DdeI endonucleases are all used under conditions specified by manufacturers (e.g., New England BioLabs, Promega) 16M. Bulyk,G. Church, and Y. Choo,in preparation(2001).

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[30]

Escherichia coli TG117: supE hsdA thiA(lac-proAB) F' [traD36 proAB + laqI q laqZAM 15] 2 x YT: Bacto-tryptone, 16.0 g/liter; Bacto-yeast extract, 10.0 g/liter; NaCI, 5.0 g/liter. For growth of zinc finger phage add tetracycline (15/zg/ml) and 50/zM zinc chloride 2x YT-agar: Bacto-tryptone, 16.0 g/liter; Bacto-yeast extract, 10.0 g/liter; NaC1, 5.0 g/liter; agar, 15.0 g/liter SOC medium: Bacto-tryptone, 20.0 g/liter; Bacto-yeast extract, 5.0 g/liter; NaC1, 10 raM; KCI, 2.5 mM; MgC12, 10 raM; MgSO4, 10 mM; glucose, 20 mM Minimal agar medium: Na2HPO4, 6.0 g/liter; KH2PO4, 3.0 g/liter; NH4C1, 1.0 g/liter; NaC1, 0.5 g/liter. After autoclaving, add 1 M MgSO4, 1.0 ml/liter; glucose, 2.0 g/liter; 1 M CaCI2, 0.1 ml/liter; 1 M thiamine hydrochloride 1.0 ml/liter; agar, 15.0 g/liter PBS-Zn: PBS (10x stock solution: NaC1, 80 g/liter; KC1, 2 g/liter; Na2HPO4. 7H20, 11.5 g/liter; KHzPO4, 2 g/liter); make up to 1 x and add 50 #M zinc chloride immediately before use TMB-based ELISA developer solution: 0.1 M sodium acetate pH 5.5; 3',3', 5',5'-tetramethylbenzidine (TMB; Sigma, St. Louis, MO ), 0.5 mg/ml; dimethyl sulfoxide (DMSO), 1% (v/v); H202, 0.05% (v/v) Related Procedures Cesium Chloride Gradient Purification of Replicative Form Phage Vector. If Fd-TET-SN is required to clone large phage display libraries, vector DNA is prepared from a l-liter bacterial culture by using a large-scale plasmid preparation kit [e.g., Qiagen (Valencia, CA) Maxiprep], followed by additional purification on a cesium chloride gradient. 14Only double-stranded phage DNA purified on a cesium chloride gradient is suitable for library construction; the library size that may be cloned is severely reduced by lower quality vector. Ten to 100 #g of phage DNA is dissolved in 4 ml of Tris-EDTA (TE) buffer containing 4 g of cesium chloride and 0.1 ml of ethidium bromide solution (10 mg/ml). Samples are well mixed to dissolve and dispensed into 5-ml ultracentrifuge tubes (Beckman, Fullerton, CA). Tubes are centrifuged with a Beckman Vti 65.2 rotor (20°, 370,000 g, 20 hr). Two bands of DNA are visible under UV light. The lower (supercoiled plasmid) band is collected with a syringe and washed four times with water-saturated butanol to remove all traces of ethidium. The resulting solution is diluted 3-fold in TE buffer and then ethanol precipitated to recover pure phage vector. Preparation of Electrocompetent Cells. Various precautions must be taken to achieve the highest level of electrocompetence.15 Escherichia coli colonies are initially selected from freshly inoculated minimal medium plates, which promote 17T. J. Gibson,CambridgeUniversity,Cambridge,UK, 1984.

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SEQUENCE-SPECIFIC ZINC FINGERS

609

the growth of bacteria containing the F' plasmid. Cells are then transferred to the rich 2x YT medium to encourage rapid growth and division. Also, all inoculations are carried out in prewarmed medium to prevent the interruption of cell growth. At the cell-harvesting stage, all centrifugations are carried out with precooled (4°), sterile centrifuge pots and buffers to minimize cell damage. To further maintain cell viability, centrifugations are performed at the minimum speed and for the minimum time required to pellet cells, so that resuspensions may be carried out gently. Escherichia coli TG1 cells are streaked out onto minimal medium plates and incubated (37 °, ~15-24 hr). A single colony is transferred into 5 ml of 2x YT medium and incubated until turbid (37 °, ~3 hr). The 5-ml culture is then added to 250 ml of 2x YT medium and incubated until the culture reaches an optical density of 0.6 at 600 nm (37 °, -,~3-4 hr), equivalent to approximately 108 cells/ml. At this point the culture is transferred to 4 × 50 ml sterile tubes and harvested by centrifugation (4 °, 3500 g, 5 min). The cell pellets are resuspended in 4 x 50 ml of cold, sterile Milli-Q water, and then centrifuged as before. The pellets are resuspended in 2 x 50 ml of cold, sterile Milli-Q water, and centrifuged again. These pellets are resuspended in 20 ml of 10% (v/v) glycerol in cold, sterile Milli-Q water and centrifuged once more. The final pellets are resuspended in 0.7 ml of 10% (v/v) glycerol in cold, sterile Milli-Q water. At this stage, the cell suspension is subdivided into 50- to 70-#1 aliquots, and either used directly for electroporation or frozen on dry ice and stored at -80". Cells prepared by this method yield approximately 109 transformants per microgram of pUC-19 supercoiled plasmid DNA, as measured by a standard electroporation assay using 15 pg of DNA. For best results, transformations are carried out by electroporation of freshly prepared (not previously frozen) electrocompetent cells. Estimating Phage Yield. Phage titer from selection eluates and culture supernatants can be estimated by using 1 #1 of phage sample to infect 1 ml of a logarithmic phase culture of E. coli (37 °, 1 hr, no shaking). Infections are serially diluted 10-fold with individual dilutions being spread on 2× YT-agar plates containing tetracycline (15/~g/ml). After incubation (37 °, 16 hr), individual colonies are counted to give an indication of the colony-forming units (phage titer) in the original sample (see Fig. 3). Acknowledgments Y. C. and M. 1. were staff scientist anti graduate student, respectively, at the Medical Research Council Laboratory of Molecular Biology (Hills Road, Cambridge CB2 2QH, UK). Both authors currently work at Gendaq, Ltd. (1-3 Burtonhole Lane, London NW7 lAD, UK).

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[31]

[31] DNA Relaxation and Cleavage Assays to Study Topoisomerase I Inhibitors By C H R I S T I A N

BAILLY

Introduction The history of the topoisomerase I enzyme is intimately interwoven with that of the plant alkaloid camptothecin (CPT).I This antitumor drug contributed importantly to the elucidation of the DNA-cleaving properties of the enzyme and much of its biochemistry. 2-4 Topoisomerase I is a monomeric t r a n s - e s t e r a s e . The enzyme catalyzes a reversible DNA cleavage reaction by transferring a phosphodiester bond to the active site tyrosine residue. 5'6 This reaction, which does not require an energy cofactor, produces a covalent intermediate in which the enzyme is covalently attached to the 3' terminus of a DNA single-strand break (Fig. 1). This transient enzyme-DNA covalent complex (often called the cleavage complex) is the key element of the reaction. The identification of specific inhibitors is essentially based on their capacity to stabilize this covalent complex. In the absence of inhibitor, the covalent complex is highly labile and freely dissociates to regenerate the active enzyme and the religated DNA molecule with intact double strands. This catalytic cycle is essential for relieving DNA torsional stress during all manipulations of DNA in cells, including replication and transcription. 7 But in the presence of a poison, such as camptothecin, the religation step of the topoisomerase I catalytic reaction can be inhibited, thus producing single-stranded DNA breakages that are subsequently transformed into double-strand breaks lethal for the growing cells. Camptothecin converts topoisomerase I into a cell poison by blocking the religation step, thereby enhancing the formation of persistent DNA breaks responsible for cell death. This is also the case for various synthetic analogs endowed with potent antitumor activities such as the clinically used drugs topotecan and SN38 (the active metabotite of irinotecan) as well as derivatives 9-aminocamptothecin, 9-nitrocamptothecin, DX-8951 f, GG-211, CKD602, and the homocamptothecin derivative BN 80915. But in addition to the camptothecins, an important 1 M. E. Wall and M. C. Wani, CancerRes. 55, 753 (1995). 2 M. Gupta, A. Fujimori, and Y. Pommier, Biochim. Biophys. Acta 1262, I (1995). 3 y. Pommier, E Pourquier, Y. Fan, and D. Strumberg, Biochim. Biophys. Acta 1400, 83 (1998). 4 j. j. Champoux, Plvg. Nucleic Acid Res. Mol. Biol. 60, 111 (1998). 5 M. R. Redinbo, L. Stewart, P. Kuhn, J. J. Champoux, and W. G. J. Hol, Science 279, 1504 (1998). 6 L. Stewart, M. R. Redinbo, X. Qiu, W. G. Hol, and J. J. Champoux, Science 279, 1534 (1998). 7 j. C. Wang, Annu. Rev. Biochem. 65, 635 (1996).

METHODS IN ENZYMOLOOY.VOL 34(1

Copyright~¢;~200J by AcademicPress All righlsof reproductionit] any tbml reserved. 0076-6879/00$35.00

[3 1]

TOPOISOMERASE I INHIBITORS

dissociation

Topo I

binding and recognition

DNA cleavage

61 1

) covalent

re~a~n

complex

FIG. 1. Schematicrepresentation of the topoisomerasecatalyticcycle. The topoisomerizationreaction involves binding of topoisomerase I to DNA, followedby DNA cutting and then religation. group of topoisomerase I inhibitors with disparate structures and origins has been identified. 8- l0 Benzophenanthridine alkaloids, certain bis- and terbenzimidazole derivatives, and several glycosylated indolocarbazole derivatives hold great promise as topoisomerase I-targeted anticancer agents. ~ Several experimental approaches have been employed to identify these inhibitors and to characterize their mechanism of action. In most cases, in vitro assays using purified topoisomerase I and a radioactively labeled DNA substrate have been employed to evidence the poisoning effects of these drugs. This chapter focuses on two frequently used experimental approaches to characterize topoisomerase I inhibitors. These procedures are illustrated mainly with examples from our own work on topoisomerase I-targeted drugs. Further detailed discussion of the effect of topoisomerase I inhibitors at the cellular level is beyond the scope of this chapter. The experiments presented here can be performed with commercial topoisomerase I from calf thymus (Life Technologies, Rockville, MD) or the human enzyme (TopoGen, Columbus, OH). Methods to overexpress and purify topoisomerase I from different organisms, including bacteria, yeast, human placenta, and baculoviruses have been described. 12

8 y. Pommier, Biochimie 80, 255 (1998). 9 y. Pommier, E Pourquier, Y. Urasaki, J. Wu, and G. S. Laco, Drug Resistance Updates 2, 307 (1999). 10C. Bailly, Curt Med. Chem. 7, 39 (2000). 11B. H. Long and B. N. Balasubramanian, Exp. Opin. Ther. Patents 10, 635 (2000). 12M.-A. Bjornstiand N. Osheroff(eds.), DNAtopoisomeraseprotocols.I. DNAtopologyand enzymes. In "Methods in Molecular Biology,"Vol. 94, Humana Press, Totowa,New Jersey, 1999.

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ENZYMOLOGYAND BIOLOGICALAPPROACHES

(-) ethidium

(+) ethidium

z o0-C0 al-~ I-'1F'IF"lF-I

topoisomer~¢

[3 1 ]

z

<._~}f~..j~

Sc'~

09

~yf°'J~

FIG.2. Effect of ethidium bromide on the electrophoretic separation of different DNA species. Native supercoiledpKMp27 DNA (0.5/zg) (lane DNA) was incubated with 4 units of topoisomerase 1 in the absence (lane Topoi)or presence of 10 #M camptothecin(lane CPT) or ascididemin (lane ASC). In both cases, reactions were stopped with sodium dodecyl sulfate and treatment with proteinase K. DNAsamples were separatedby electrophoresison an agarosegel (a) withoutethidium or (b) containing ethidium bromide at 1 /zg/ml. The supercoiled DNA (Sc), nicked DNA (Nck), and relaxed DNA (Rel) species are depicted. DNA Relaxation Assays Topoisomerase I can remove supercoils from plasmid DNA by making transient nicks in the plasmid and allowing the passage of a single-stranded DNA molecule through the nick. Plasmid DNA represents a useful substrate to study the activity of topoisomerase I as well as the inhibitory effect of drugs. Depending on the system used to lyse bacteria (alkaline or detergent lysis, boiling method) and to purify the plasmid [CsC1 centrifugation, selective precipitation, Qiagen (Valencia, CA) columns], the preparation usually contains 90-99% of negatively supercoiled circular DNA molecules (form I) and 1-10% nicked circular DNA molecules (form II). In spite of their identical sequence and molecular weight, these two DNA forms can be easily separated by electrophoresis on agarose gels because of their different shapes. Supercoiled DNA has a compact structure and thus migrates much more rapidly than the nicked DNA with an extended structure. On treatment with eukaryotic topoisomerase I, the supercoiled DNA is transformed into a population of relaxed DNA topoisomers (Fig. 2a). Each band in the ladder represents a single topoisomer differing from its neighbors by a linking difference of ± l ; the more supercoiled the species, the faster the migration.13 To visualize the different forms of the DNA, the gel is soaked in a solution containing a drug that becomes brightly fluorescent once bound to the DNA. The intercalating drug ethidium bromide is usually employed but, alternatively, the Is R. Bowater, K Aboul-Ela, and D. M. Lilley. Methods Enzymol. 212, 105 (1992).

[31]

TOPOISOMERASE I INHIBITORS

613

DNA in agarose gels can be stained with other fluorescent probes such as SYBR gold (Molecular Probes Engene, OR). Although ethidium is toxic and carcinogenic (therefore it must be handled with care), it provides satisfactory staining of all DNA forms and can easily be extracted from the DNA if needed. For this reason, this chemical remains the most widely used DNA-staining reagent. If an identical experiment is performed with an agarose gel prestained with ethidium bromide, then the result is completely different. Indeed, in the case of an agarose gel containing ethidium (usually at a concentration of 0.5-1 /zg/ml), the DNA species moving toward the anode during the electrophoresis will become progressively saturated with ethidium molecules. The intercalator inserts itself between the stacked base pairs and this effect causes an untwisting of the double-helical structure. In contrast, the electrophoretic mobility of the nicked DNA form II is practically unaffected because intercalation has no effect on the writhe of a nicked DNA ring (Fig. 2b). It should be noted that supercoiled and nicked DNAs do not stain with ethidium bromide to the same extent. Owing to its topological constraints, the closed circular DNA form accommodates much less ethidium than the open circular form. Fluorescence quantitation measurements have shown that for plasmid pBR322, the closed circular form bound about 1.8 times less ethidium than an equivalent amount of open circular form. 14 This observation explains why decreases in fluorescence of the open circular DNA band are not strictly matched by increases in the closed circular DNA form.

Applications 1. Ascididemin is a planar pentacyclic alkaloid isolated from the Mediterranean ascidian Cystodytes dellechiajei. 15 Intercalation of this drug between the base pairs of DNA affects the relaxation of plasmid DNA by topoisomerase I (Fig. 2). Without ethidium added to the electrophoresis gel, the effect of ascididemin is at first sight comparable to that of camptothecin (Fig. 2a). But when the experiments is repeated with ethidium in the agarose gel, then it can clearly be shown that the marine alkaloid is not a potent poison for topoisomerase I. In contrast to camptothecin, the band corresponding to nicked DNA is only weakly increased in the presence of ascididemin. This drug poorly stabilizes the formation of covalent DNA-topoisomerase I complexes. 16 Nonspecific effects due to DNA intercalation and specific effects resulting from the poisoning of topoisomerase I can thus be differentiated by this relaxation assay. 14 E Boege, T. Straub, A. Kehr, C. Boesenberg, K. Christiansen, A. Andersen, F. Jakob, and J. KShrle, J. Biol. Chem. 271, 2262 (1996). 15 I. Bonnard, N. Bontemps, S. Lahmy, B. Banaigs, G. Combaut, C. Francisco, E Colson, C. Houssier, M. J. Waring, and C. Bailly, Anti-Cancer Drug Des. 10, 333 (1995). 16 L. Dassonneville, N. Wattez, B. Baldeyrou, C. Mahieu, A. Lansiaux, B. Banaigs, I. Bonnard, and C. Bailly, Biochem. Pharmacol. 60, 527 (2000).

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ENZYMOLOGY AND BIOLOGICALAPPROACHES

[31 ]

2. Glycosylated indolocarbazoles related to the antibiotics rebeccamycin and BE-13793C have been identified as a potent topoisomerase I inhibitors.17-2° Despite their different chemical structures, camptothecins and indolocarbazoles share common steric and electronic features and recognize similar structural elements of the topoisomerase I - D N A covalent adductfl I The effects of four indolocarbazole drugs (Fig. 3A) on the relaxation of DNA by topoisomerase I are shown in Fig. 3B. Negatively supercoiled plasmid pKMp27 was incubated with the purified enzyme in the presence of increasing concentrations of the test drugs. The DNA samples were treated with sodium dodecyl sulfate (SDS) and proteinase K to remove any covalently bound protein and resolved in a 1% (w/v) agarose gel containing ethidium bromide to differentiate the specific (poisoning) and nonspecific effects. As shown in Fig. 3B, the relaxed DNA migrates faster than the supercoiled plasmid because of ethidium-induced DNA unwinding effects. A marked increase in the intensity of the band corresponding to nicked DNA molecules can be detected with camptothecin used as a positive control, and with three of the indolocarbazole drugs. Compounds 1, 2, and 3 (Fig. 3) efficiently stabilize topoisomerase I - D N A complexes. In contrast, the bisglycosyl derivative 4 appeared incapable of stimulating DNA cleavage by topoisomerase I. 22 In addition to the changes in intensity of the nicked DNA band, a strong shift in the mobility of supercoiled plasmid (form I) was observed with increasing concentrations of 1, 2, and 4, but not with 3. The 2, 10-dihydroxy analog 3 exhibits DNA-binding properties distinct from those of the other three indolocarbazoles. 23 The effects of 1, 2, and 4 on the electrophoretic mobility of supercoiled DNA can be attributed to a decrease in plasmid DNA linking number due to intercalation. The tumor-active drug NB-506 (1) and its analog 2 both behave as specific topoisomerase I inhibitors trapping the cleavable complexes and as nonspecific inhibitors of a DNA-processing enzyme acting via DNA bindingfl 4 On the other hand, the 2, 10-isomer 3 behaves exclusively as a specific topoisomerase I poison, as is the case with camptothecin. 23 17T. Yoshinari, M. Matsumoto, H. Arakawa, H. Okada, K. Noguchi, H. Suda, A. Okura, and S. Nishimura, Cancer Res. 55, 1310(1995). 18T. Yoshinari, A. Yamada,D. Uemura, K. Nomura, H. Arakawa, K. Kojiri, E. Yoshida, H. Suda, and A. Okura, Cancer Res. 53, 490 (1993). 19C. Bailly,J. E Riou, R Colson,C. Houssier, E. Rodrigues-Pereira,and M. Prudhomme,Biochemisto, 36, 3917 (1997). 2oC. Bailly, X. Qu, D. E. Graves, M. Prudhomme, and J. B. Chaires, Chem. Biol. 6, 277 (1999). 2J C. Bailly, C. Carrasco, E Hamy, H. Vezin, M. Prudhomme,A. Saleem, and E. Rubin, Biochemisto' 38, 8605 (1999). 22X. Qu, J. B. Chaires, M. Ohkubo,T. Yoshinari, S. Nishimura, and C. Bailly,Anti-CancerDrug Des. 14, 433 (2000). 23C. Bailly, L. Dassonneville, E Colson, C. Houssier, K. Fukasawa, S. Nishimura, and T. Yoshinari, Cancer Res. 59, 2853 (1999). 24C. Bailly, X. Qu, J. B. Chaires, P. Colson, C. Houssier, M. Ohkubo, S. Nishimura, and T. Yoshinari, J. Med. Chem. 42, 2927 (1999).

.~H NHCHO

A

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N"

)

0

*

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)

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1

B

2

3

4

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FIG. 3. (A) Structures of four glycosylated indolocarbazole drugs and (B) their effects on the relaxation of plasmid DNA by topoisomerase I. Native supercoiled pKMp27 DNA (0.5 /zg) (lane DNA) was incubated with 6 units of topoisomerase in the absence (lane Topoi) or presence of drug at the indicated concentration (micromolar). Reactions were stopped with sodium dodecyl sulfate and treatment with proteinase K. Camptothecin (lane CPT) was used at 20/~M. DNA samples were separated by electrophoresis on an agarose gel containing ethidium bromide at 1 #g/ml. The gel was photographed under UV light. Nck, Nicked; Rel, relaxed; Sc, supercoiled.

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ENZYMOLOGYAND BIOLOGICALAPPROACHES

[3 1]

DNA Cleavage Assays Although supercoiled DNA is the major substrate for topoisomerase I, the enzyme can also bind to and cleave linear DNA molecules. The minimum DNA duplex region required for topoisomerase I-mediated reaction in vitro is a sequence containing nine nucleotides on the scissile strand and five nucleotides on the noncleaved strand. 25 Cleavage assays can be performed with either synthetic oligonucleotide duplexes containing a few topoisomerase I cleavage sites, usually one or two, or DNA restriction fragments cut out from genes, DNA viruses, or plasmids. Use o f Synthetic Oligonucleotides

Various oligonucleotide sequences can be used to study topoisomerase I-mediated DNA cleavage. For example, duplexes 1, 2, and 3 each contain a unique topoisomerase I cleavage site (indicated by an arrow): Duplex 1: Y-GGCGCGGAGACTTSGGAGAAATTTGGCGCGG-Y Y-CCGCGCCTCTGAA CCTCTTTAAACCGCGCCC-5'

Duplex 2: 5'-GATCTAAAAGACTT'I'GGAAAAATTTTTAAAAAAGCTCa *-y Y -CTAGATTI-I-CTGAA CCTT-I-rTAAAAATTTTIq-CGAG- 5~

Duplex 3: 5'-CAAAGTCAGGTTGAT$GAGCATA-I-rTTACTCa*-Y Y-GTTTCAGTCCAACTA CTCGTATAAAATGAG-5'

The 30-base pair DNA duplex 1 was designed to contain a high-efficiency topoisomerase I cleavage site. 25'26The 36-base pair duplex 2 derives from the Tetrahymena rDNA hexadecameric sequence, 27 whereas the 30-base pair duplex 3 is found in simian virus 40 (SV40) DNA. 28'29 Duplex 1 has been employed to characterize 25j. Q. Svejstrup, K. Christiansen, A. H. Andersen, K. Lund, and O. Westergaard,J. Biol. Chem. 265, 12529 (1990). 26K. Christiansen, J. Q. Svejstrup, A. H. Andersen, and O. Westergaard, J. Biol. Chem. 268, 9690 (1993). 27 B. J. Bonven, E. Gocke, and O. Westergaard, Cell 41,541 (1985). 28C. Jaxel, K. W. Kohn, and Y. Pommier,Nucleic Acids Res. 16, 11157 (1988). 29C. Jaxel, G. Capranico,D. Kerrigan,K. W. Kohn,and Y.Pommier,J. Biol. Chem. 266, 20418 (1991).

[31]

TOPOISOMERASE I INHIBITORS

617

the effect of CPT derivatives. 30'31 Duplex 2 has also been used to compare the inhibitory potency of camptothecin derivatives32 as well as a variety of non-CPT topoisomerase I poisons such as aclacinomycin A . 33 On treatment of the Y-labeled oligonucleotide terminated by [u-32p]cordycepin, marked a*) with topoisomerase I in the presence of CPT or SN38, the active metabolite of the anticancer drug irinotecan generates the underlined 23-mer oligonucleotide, which can easily be separated from the full-length oligonucleotide on a denaturing polyacrylamide gel. In addition, this one-site oligonucleotide also provides a convenient system to measure the reversibility of covalent complex formation. The covalent topoisomerase I-DNA complex is highly reversible on drug removal or if exposed to a high salt concentration. By adding NaC1 (usually 0.5 M, at 25 °) directly to the drug-containing reaction mixture (prepared in a standard low salt reaction buffer such as 20 mM Tris-HC1, pH 7.2) after formation of the cleavage complex, one can thus estimate the stability of the covalent complex and compare the stabilizing action of drugs. 34 Distinguishing b e t w e e n Cleavage and Religation Reactions Short synthetic oligonucleotides also provide useful substrates to uncouple the cleavage and ligation reactions of topoisomerase I. The model proposed by Hecht and collaborators is diagrammed in Fig. 4. 35,36 A partial DNA duplex containing a high-efficiency topoisomerase I cleavage site is reacted with the enzyme, which catalyzes a nucleophilic attack via its active site tyrosine OH group on the phosphodiester bond at the TpA step (Fig. 4A). The resulting 3'-O-phosphorotyrosine bond ensures the covalent attachment of the enzyme to the oligonuclotide and the 5'-OHAGAGA oligonucleotide is released. Sequential religation cannot occur because of the instability of the duplex involving the truncated strand downstream from the site of cleavage. The topoisomerase I-DNA covalent intermediate is thus trapped. For this reason, the initial partial duplex is designated as a "suicide" substrate. The covalent 5t-GGCGCGGAGACTT-topoisomerase I binary complex thus formed can be purified. 37 The covalently bound enzyme remains catalytically competent, 30 X. Wang, G. F. Short, W. D. Kingsbury, R. K. Johnson, and S. M. Hecht, Chem. Res. Toxicol. 11, 1352 (1998). 31 X. Wang, L. K. Wang, W. D. Kingsbury, R. K. Johnson, and S. M. Hecht, Biochemistry 37, 9399 (1998). 32 M. Valenti, W. Nieves-Neira, G. Kohlhagen, K. W. Kohn, M. E. Wall, M. C. Wani, and Y. Pomrnier, Mol. Pharmacol. 52, 82 (1997). 33 j. L. Nitiss, E Pourquier, and Y. Pommier, Cancer Res. 57, 4564 (1997). 34 A. Tanizawa, K. W. Kohn, G. Kollhagen, E Leteurtre, and Y. Pommier, Biochemistry 34, 7200 (1995). 35 K. A. Henningfeld and S. M. Hecht, Biochemistry 34, 6120 (1995). 36 X. Wang, K. A. Henningfeld, and S. M. Hecht, Biochemistry 37, 2691 (1998). 37 K. A. Henningfeld, T. Arslan, and S. M. Hecht, J. Am. Chem. Soc. 118, 1170l (1996).

618

ENZYMOLOGY AND BIOLOGICAL APPROACHES

[31 ]

high efficiency Topoi site

A 5' - G G C G C G G A G A C T T A G A G A 3, - C C G C G C C T C T G A A T C T C T T T A A A C C G C G C C C

partial DNA duplex (suicide substrate)

3' - C C G C G C C T C T G A A T C T C T T T A A A C C G C G C C C

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B

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branched substrate

F-T-r.T.!..-T-T.Z-~3--~-T-r-r.. .I-~-T-- "4, nicked substrate T~-~-T'F-=~'~-',r,T" -3 ' gapped substrate

[3 1]

TOPOISOMERASE I INHIBITORS

6 19

as it can engage in ligation (Fig. 4B) with an oligonucleotide acceptor containing a free Y-OH group to afford the full duplex product. The religation can be performed with complementary acceptors 26'38'39 but also with nonhomologous acceptors of varying lengths and sequences 35"39 or structurally altered substrates such as an acceptor oligonucleotide having a Y-terminal 3'-deoxyadenosine analog. 4° A variety of such suicide substrates and modified acceptors have been utilized. 3°,3j ,36,41 43 A few are representated in Fig. 4C. Modifications include reactions that form DNA insertions, deletions, and mismatches. 35 The design of specific oligonucleotide sequences containing various types of natural or nonnatural bases represents a powerful experimental approach to characterize the role of single nucleotides within the recognition domain in substrate recognition and cleavage by topoisomerase I. In addition, this model system is convenient for drug studies. For example, the assay has been used to compare the effects of the topoisomerase I inhibitors CPT, topotecan, nitidine, and coralyne. Interestingly, the ligation proved much more efficient with the two phenanthridine alkaloids than with the two camptothecins when the 5'-terminal residue on the acceptor oligonucleotide was an adenine residue required to form a C- A mismatch. 36 Similar model systems using branched, nicked, and gapped DNA substrates have been successfully used to compare the topoisomerase 1-poisoning activities of numerous CPT analogs. 3°'31

Use of Restriction Fragments Oligonucleotides can be easily manipulated and their sequences can be chosen at will, but they generally offer a limited number of topoisomerase I cutting sites. In most cases, within a 30-mer oligonucleotide no more than three or four cleavage sites can be mapped at nucleotide resolution. To analyze a large number of sites so as to determine possible base preferences around topoisomerase I cleavage sites it 38 j. Q. Svejstrup, K. Christiansen, I. I. Gromova, A. H. Andersen, and O. Westergaard, ,I. Mol. Biol. 222, 669 (1991). 39 S. Shuman, J. Biol. Chem. 267, 8620 (1992). 4o T. Arslan, A. T. Abraham, and S. M. Hecht, Nucleosides Nucleotides 17, 515 (1998). 41 T. Arslan, A. T. Abraham, and S. M. Hecht, J. Biol. Chem. 273, 12383 (1998). 42 p. Pourquier, L.-M. Ueng, G. Kohlhagen, A. Mazumder, M. Gupta, K. W. Kohn, and Y. Pommier, J. Biol. Chem. 272, 7792 (1997). 43 p. Pourquier, A. A. Pilon, G. Kohlhagen, A. Mazumder, A. Sharma, and Y. Pommier, J. Biol. Chem. 272, 26441 (1997).

FIG. 4. (A) Oligonucleotide substrates used to uncouple the cleavage and religation reactions of topoisomerase I. The enzyme induces cleavage of the suicide substrate at the 5'-TpA step (arrow) and then religates the cleaved strand to the acceptor sequence to generate the full duplex. (B) Topoisomerase l-mediated ligation reaction. (C) Schematic representation of the different acceptor oligonucleotides used to generate structurally modified substrates.

620

ENZYMOLOGY AND BIOLOGICAL APPROACHES

[31

]

is appropriate to use DNA fragments of 50-300 base pairs in length. Restriction fragments obtained from plasmid DNA offer a range of DNA substrates suitable for sequencing studies. Almost any restriction fragment, whatever its origin, can be used but as with oligonucleotides, it is recommended to label the DNA substrate at its 3' end. In the covalent complex, topoisomerase I is linked to the 3' terminus of the DNA break that it generates. Therefore, if a 5' end-labeled DNA were used, the labeled strand would be covalently attached to the enzyme and this could perturb significantly the electrophoretic mobility of the DNA in the gel unless the protein molecules were totally digested by a protease prior to electrophoresis. DNA fragments from SV40 DNA, phagemids and plasmids, human oncogenes, and diverse cDNA molecules have been tlsed. 28'29'34'44'45 For example, we found it particularly useful to employ the 117-bp and 265-bp fragments obtained from the plasmid pBS (Stratagene, La golla, CA codigested with the enzymes PvuII and EcoRI. 19 Once a series of at least 30 cleavage sites has been mapped, the analysis of the base preferences around topoisomerase I cleavage sites in the presence and absence of a given drug can be performed by aligning the topoisomerase I-induced cleavage sites relative to the broken phosphodiester b o n d . 46"47 Applications

1. Homocamptothecins containing a seven-membered /%hydroxylactone, in place of the conventional six-membered ~-hydroxylactone ring found in camptothecins, represent a new series of potent inhibitors of topoisomerase I. 48,49 The lead compound in the homocamptothecin series is a 9,10-bisfluoro derivative referenced BN80915. This drug has revealed exceptional profiles of antitumor activity both in vitro and in vivo and has been advanced to clinical trials. 5° The capacity of BN80915 to inhibit DNA topoisomerase I was compared with that of SN38, the

44 S. E. Porter and J. J. Champoux, Nucleic Acids Res. 21, 8521 (1989). 45 E Leteurtre, A. Fujimori, A. Tanizawa, A. Chabra, A. Mazumder, G. Kohlhagen, H. Nakano, and Y. Pommier, J. Biol. Chem. 269, 28702 (1994). 46 y. Pommier, K. W. Kohn, G. Capranico, and C. Jaxel, in "Molecular Biology of DNA Topoisomerases and Its Application to Chemotherapy" (T. Andoh, H. Ikeda, and M. Oguro, eds.), p. 215. CRC Press, London, 1993. 47 G. Capranico and M. Binaschi, Biochim. Biophys. Acta 1400, 185 (1998). 48 O. Lavergne, L. Lesueur-Ginot, E Pla Rodas, G. Kasprzyk, J. Pommier, D. Demarquay, G. Prevost, G. Ulibarri, A. Rolland, A.-M. Schiano-Liberatore, J. Harnett, D. Pons, J. Camara, and D. C. H. Bigg, J. Med. Chem. 41, 5410 (1999). 49 L. Lesueur-Ginot, D. Demarquay, R. Kiss, P. G. Kasprzyk, L. Dassonneville, C. Bailly, J. Camara, O. Lavergne, and D. C. H. Bigg, Caneer Res. 59, 2939 (1999). 50 E Philippart, L. Harper, C. Chaboteaux, C. Decaestecker, Y. Bronckart, L. Gordover, L. LesueurGinot, H. Malonne, O. Lavergne, D. C. H. Bigg, P. M. da Costa, and R. Kiss, Clin. Cancel" Res. 6, 1557 (2000).

[31]

TOPOISOMERASE I 1NHIBITORS

621

o

°

' - V

-

-

F

A

SN38(~M)

_

BN80915

indolocarbazoles

(~M) -

1

3

~..,,=lll ~ml I -

OO00Q

n

-110 -100

SOI~

+IEi.=++

`=~

i +o

-8o

~

ii ~ l i ~ ' -

i -70

70-ii m

i

i -.60

i i l i i ~

++

l i + i i + +

....

-

.....

+

II

7--~" .

Ill + + ~

~

t0-~

~

i

,

"

i

. t.50 .

++

~llliiliil+,-

... ,.+

.

+

+

+

>+

i

+'+' i

40- i ~

-40

411

+o-+

. = -30

t

FIG. 5. (A) Cleavage of a 116-bp DNA restriction fragment from plasmid pTayD by human topoisomerase I in the presence of graded concentrations of the homocamptothecin derivative BN80915 and the conventional camptothecin SN38. The 3' end-labeled fragment (lane DNA) was incubated in the absence (lane Topoi) or presence of the test drug at the indicated concentration (micromolar). Topoisomerase I cleavage reactions were analyzed on a 8% denaturing polyacrylamide gel. Numbers at the right-hand side of the gel show the nucleotide positions, determined with reference to the guanine tracks labeled G. The positions of I 0 major cleavage sites are indicated. (B) Cleavage of a 131-bp DNA restriction fragment from plasmid pTayB by human topoisomerase I in the presence of CPT or the two glycosylated indolocarbazole derivatives 1 and 3 (structures indicated in Fig. 3A).

active metabolite of the clinically used antitumor drug irinotecan. For this purpose, a 116-bp EcoRI-HindlII restriction fragment from plasmid pTayD, uniquely 3' end labeled at the EcoRI site, was used as a substrate for the topoisomerase I cleavage reaction. The cleavage products were analyzed on a sequencing polyacrylamide gel as shown in Fig. 5A. Both BN80915 and SN38 promote DNA cleavage by topoisomerase I, but the effect is more pronounced with the homocamptothecin

622

ENZYMOLOGYAND BIOLOGICALAPPROACHES

[3 1]

derivative compared with the conventional camptothecin. Moreover, the patterns of cleavage sites are different. For example, sites 2, 3, 4, 7, 8, and 10 are cleaved more efficiently with BN80915 than with SN38. The converse effect occurs at sites 1 and 5 whereas the cleavage intensities at sites 6 and 9 are comparable for the two drugs. The results confirm that the replacement of the camptothecin lactone E-ring with a homologous seven-membered lactone ring changes the sequencespecificity of drug-induced DNA cleavage by topoisomerase I. In particular, several sites encompassing the sequence 5'-AAC'I'G were found to be specifically cleaved in the presence of homocamptothecin (hCPT) but not with camptothecin. 51 Such cleavage studies with restriction fragments have provided essential information about the role of the o~/fl-hydroxylactone ring of CPT/hCPT in the poisoning of topoisomerase I. 2. Indolocarbazoles derived from the antibiotic rebeccamycin are promising antitumor agents targeting topoisomerase I. l°'j~ Unlike the camptothecins, these drugs bind to DNA even in the absence of topoisomerase I. The planar indolocarbazole chromophore of the tumor active drug NB-506 (compound 1 in Fig. 3A) intercalates between two consecutive base pairs in the DNA double helix, thus placing the appended glucose residue into one of the helical grooves, most likely the minor groove. 19 We used a series of restriction fragments to compare the patterns of topoisomerase I-mediated DNA cleavage obtained in the presence of 1 or its regioisomeric analog 3, which contains two hydroxyl groups at positions 2 and 10 instead of at positions 1 and 11. A typical sequencing gel obtained with a 131-bp EcoRI-HindIII restriction fragment from plasmid pTayB, uniquely 3' end labeled at the EcoRI site, is presented in Fig. 5B. The two drug isomers are equally potent at maintaining the integrity of the topoisomerase I-DNA covalent complexes but stimulate cleavage at different sites on DNA. Compound 1 stabilizes topoisomerase I preferentially at sites having a pyrimidine (T or C) and a G on the 5' and 3' sides of the cleaved bond, respectively, whereas compound 2 induces topoisomerase I-mediated cleavage principally at TG sites only. The two drugs preferentially induce cleavage by the enzyme at different nucleotide sequences. Therefore, we concluded that the positioning of the two hydroxyl groups on the drug chromophore exerts a significant effect on the sequence specificity of topoisomerase I-induced DNA cleavage. 23 Cleavage assays can also be performed with a linear plasmid DNA 3' end labeled, such as the EcoRI-linearized pBR322 DNA. In this case, the cleavage products of the DNA fragment are denatured with alkali (0.45 M NaOH) prior to electrophoresis on agarose gels containing a detergent [0.1% (w/v) SDS]. This procedure does not permit identification of the cleavage site at nucleotide resolution, 51C. Bailly, A. Lansiaux, L. Dassonneville,D. Demarquay,H. Coulomb,M. Huchet, O. Lavergne, and D. C. H. Bigg,Biochemistry 38, 15556 (1999).

[31 ]

TOPOISOMERASE I INHIBITORS

623

but it is relatively quick and the radioactive substrate is easy to prepare. This is a convenient assay for comparing the inhibitory potencies of series of drugs. 52 Conclusion The relaxation and cleavage assays presented here are the most frequently used in vitro methods to study the effects of topoisomerase I inhibitors. However, other

technical approaches can be used. For example, a filter binding assay can also be employed to evidence the stabilizing action of drugs on DNA-topoisomerase I covalent complex formation. 53 Neither topoisomerase I alone nor DNA is retained on nitrocellulose filters whereas the large covalent complexes bind selectively to nitrocellulose in the presence of SDS. The addition of a topoisomerase I poison such as CPT, or certain flavones, 14 increases significantly the amount of material retained on the filter, thus reflecting the drug-induced formation of additional covalent complexes. This assay may be used as a rapid screening procedure for screening libraries but it is not sensitive and requires relatively large amounts of enzyme. In this chapter, the two most widely used in vitro approaches to characterize topoisomerase I inhibitors have been briefly outlined. Not only do these assays help to identify novel topoisomerase I poisons; they can provide important structural data. They can easily be performed by any laboratory that is experienced in methods of molecular biology. We hope that this description will encourage researchers to apply these approaches to discover new anticancer or antiparasitic agents directed against topoisomerase I. Acknowledgment Support from the Association pour la Recherche sur le Cancer is acknowledged.

52 j. F. Riou, P. FossE, C. H. Nguyen, A. K. Larsen, M. C. Bissery, L. Grondard, J. M. Saucier, E. Bisagni, and E Lavelle, Cancer Res. 53, 5987 (1993). 53 S. M. Hecht, D. E. Berry, L. MacKenzie, R. W. Busby, and C. A. Nasuti, J. Nat. Prod. 55, 401 (1992).

624

ENZYMOLOGY AND BIOLOGICAL APPROACHES

[32]

[32] In Vitro Human Immunodeficiency Virus Type 1 Integrase Assays B y CHRISTOPHE M A R C H A N D , NOURI NEAMATI, a n d YVES POMMIER

Introduction Integration of the human immunodeficiency virus type 1 (HIV- 1) genome into host cell DNA is an essential step for viral replication. Soon after cell penetration, the single-stranded viral RNA is reverse transcribed into a double-stranded proviral DNA. The next major event is integration, whereby the viral DNA is inserted into the host chromosome to create the proviral state. This event is catalyzed by a viral enzyme called integrase, encoded in the po! gene and generated after proteolysis of the Gag-Pol fusion protein precursor by the HIV-1 protease (Pig. 1). Integrase is responsible for the removal in the cytoplasm of a GT dinucleotide immediately 3' from a conserved CA dinucleotide at the Y-end of both extremities of the viral genome [U3 and U5 long terminal repeats (LTRs)]. After this first step, called Yprocessing, integrase remains bound to the LTRs and this preintegration complex migrates to the nucleus, where the second step of the integration reaction (3'-end joining) occurs. The T-end joining reaction consists of the direct nucleophilic attack of the Y-recessed viral ends on the host chromosome. Both termini of the viral DNA, kept in close proximity, integrate with a 5-bp stagger toward the 51-ends of the target chromosomal DNA. Completion of the integration process requires removal of the two unpaired nucleotides at the 5'-ends of the viral DNA and gap filling, probably accomplished by cellular enzymes {for review see Refs. 1-3). HIV-1 integrase is a 32-kDa protein of 288 amino acids. It can be divided into three functional domains {Pig. 2). The central domain represents the enzyme catalytic core and is required for all enzymatic activities. The N- and C-terminal domains are only required for full enzymatic activity and oligomerization. The catalytic core of HIV-I integrase {residues 50-212) contains three acidic amino acids (Asp-64, Asp-116, and Glu-152), which are highly conserved among integrases from all retroviruses, retrotransposons, and some bacterial transposases. These conserved residues are absolutely required for all integrase activities and are referred to as the D,D(35)E motif. In HIV-l-infected patients, at present, only reverse transcriptase and protease inhibitors are used as a combination in highly active antiretroviral therapy I y. Pommier, A. A. Pilon, K. Bajaj, A. Mazumder, and N. Neamati, Antivira[ Chem. Chemother 8, 463 (1997). 2 p. O. Brown, in "Retroviruses" (J. M. Coffin, S. H. Hughes, and H. E. Varmus, eds.), pp. 161-203. Cold Spring Harbor Press, Cold Spring Harbor, New York, 1998. 3 A. M. Skalka, "Advances in Virus Research." Academic Press, San Diego, California, 1999.

METHODSIN ENZYMOLOGY.VOL.340

In Vitro H I V - 1 INTEGRASE ASSAYS

[32]

U3

R U5

s,I_~.nc'z"z'ccc~

625

U3 R Gag

Pol

Env

U5

3'

ATCGTCA 1

5'

3'-Processing I

C 3'-end joining

1

Hosl

5'-processing gap filling

FIG. 1. Schematic representation of the integration reaction steps. HIV-1 integrase catalyzes 3rprocessing in the cytoplasm and 3t-end joining in the nucleus. The conserved CA dinucleotide is outlined and the 3t-processed GT dinucleotide is underlined. The mechanisms tbr gap filling and 5'-end joining are unknown.

626

[32]

ENZYMOLOGY AND BIOLOGICAL APPROACHES

PR

RT

IN

s

s

s ~

s~ p s

~

s s

s~ I~

/" ts ~4~J I I

,

H 1 2 H 1 6 C 4 0 C43

zinc binding

DD(35)E

'

~................................................................... I

D64

D116

K...K...K 211

catalytic core

E152

.... R...R..R

.... K...K...K 244

DNA binding

FIG. 2. The three domains of HIV-1 integrase. The central core domain contains three conserved acidic residues [D,D(35)E] common to all retroviral integrases.

(HAART). Because HIV-1 integrase is absolutely required for viral replication and because it has no cellular equivalent, this enzyme is an excellent target for a chemotherapeutic approach. 4'5 In vitro integration assays are used to elucidate the enzyme mechanisms and to discover, design, and develop integrase inhibitors. Other assays using subcellular fractions from infected cells have been reported (preintegration complex assays, PICs). 6 They are not described in the present chapter. Human Immunodeficiency Virus Type 1 Integrase Preparation The HIV- 1 integrase protein commonly used is a soluble double mutant bearing the mutations F185K and C280S. This mutant is as catalytically active as the wildtype protein. Its higher solubility provides better purification yields. 7 Transformation

The HIV-1 IN/F185K/C280S gene has been cloned into a pET-15b ampicillinresistant vector (Novagen, Milwaukee, WI). Twenty microliters ofEscherichia coli 4 y. Pommier and N. Neamati, Adv. Virus Res. 52, 427 (1999). 5 N. Neamati, C. Marchand, and Y. Pommier, Adv. Pharmacol. 49, 147 (2000). 6 M. S. T. Hansen, G. J. I. Smith, T. Kafri, V. Molteni, J. S. Siegel, and F. D. Bushman, Nat. BiotechnoL 17, 578 (1999).

[32]

In Vitro HIV-1 INTEGRASEASSAYS

627

BL21 (DE3) competent cells (Novagen) are placed into a 1.5 ml prechilled tube (Eppendorf Scientific, Westbury, NY) and kept on ice. Five nanograms of the pET15b/HIV- 1 IN/F 185K/C280S plasmid is then added to the cells and the suspension is incubated for 5 min on ice. The competent bacteria are heat shocked by placing the tube in a water bath at 42 ° for 30 sec and then back on ice for 2 min. Bacteria are diluted by adding 250 #1 of SOC medium (Novagen) and placed for 1 hr at 37 ° in a rotary shaker. The entire tube volume is spread on an agar plate containing ampicillin (100 #g/ml; K. D Medical, Columbia, MD) and grown overnight at 37 °. At least three colonies are picked up and amplified for 12 hr in separate tubes containing 6 ml of Luria-Bertani (LB) broth medium (Digene, Beltsville, MD) and ampicillin (100 #g/ml; Boehringer Mannheim, Indianapolis, IN). A QIAprep Spin minipreparation (Qiagen, Valencia, CA) is performed with 5 ml of culture and the remaining 1 ml is diluted to 40% (v/v) glycerol (GIBCO-BRL/Life Technologies, Rockville, MD), properly labeled and stored at - 8 0 °. The plasmid resulting from the minipreparation is checked on a 1% (w/v) agarose gel and its concentration is determined by UV absorption. The presence of the integrase gene insert can be checked by sequencing of the plasmid. Amplification

BL21 (DE3) competent cells containing the pET- 15b/HIV- 1 IN/F185K/C280S vector are spread on an agar plate containing ampicillin (100/zg/ml) and grown overnight. The colonies are then harvested, expanded in 1 liter of fresh LB broth medium containing ampicillin (100 #g/ml), and incubated in a rotary shaker at 37 °. When the cells reach an optical density of 0.5 at 600 nm, overexpression is induced by adding fresh isopropyl-fi-D-thiogalactopyranoside (IPTG; GIBCO-BRL/Life Technologies) to a final concentration of 0.4 mM. Cells are then incubated for an additional 3 hr. Purification

Cells are harvested by centrifugation (10 min, 3000 rpm) and the pellet is resuspended in ice-cold lysis buffer containing 20 mM HEPES, pH 7.5 (Boehringer Mannheim), 1 M NaC1 (Digene), 5 mM imidazole (Sigma-Aldrich, Milwaukee, WI), 2 mM 2-mercaptoethanol (Sigma-Aldrich), and fresh lysozyme (0.2 mg/ml; Boehringer Mannheim) (15 ml of lysis buffer per 500 ml of culture). The lysate is incubated on ice for 30 min with occasional stirring, sonicated on ice for approximately 1 min and centrifuged (20 min, 30,000g, 4°). The supernatant is then applied to a Sepharose column (diameter 1 cm, volume 15 ml; Bio-Rad, Hercules, CA) containing 1 ml of chelating Sepharose Fast Flow (Amersham Pharmacia Biotech, Piscataway, N J) saturated with a solution of 0.4 M NiSO4 (Sigma-Aldrich). Excess NiSO4 has been previously removed by 1 column 7T. M. Jenkins, A. Engelman,R. Ghirlando,and R. Craigie,J. Biol. Chem.271, 7712 (1996).

628

ENZYMOLOGYAND BIOLOGICALAPPROACHES

[32]

volume of elution buffer containing 25 mM HEPES (pH 7.5), 0.5 M NaC1, 2 mM 2-mercaptoethanol, and 5 mMimidazole. The supernatant is washed with 3 column volumes of the same elution buffer but with increasing concentration of imidazole (20, 60, and 250 mM). HIV-1 integrase protein is eluted and fractionated with 5 ml of elution buffer at 750 mM imidazole. Protein fractions are checked for integrase on a 12-20% gradient (w/v) Tricine-SDS-protein gel (Novex, San Diego, CA) revealed by Gel Code Blue (Pierce, Rockford, IL). HIV-1 integrase-containing fractions are pooled and dialyzed twice against 1 liter of buffer containing 50 mM HEPES (pH 7.5), 1 M NaC1, 4 mM EDTA (Quality Biological, Gaithersburg, MD), 2 mM dithiotbreitol (DTT; Boehringer Mannheim), and 20% (v/v) glycerol. Integrase concentration is checked by Micro Protein assay (Bio-Rad) and the protein is diluted 2-fold with a 60% (v/v) glycerol solution. The HIV-I integrase is stored "as is" at - 2 0 ° in 25 mM HEPES (pH 7.5), 0.5 M NaC1, 2 mM EDTA, 1 mM DTT, and 40% (v/v) glycerol. Assays

5'-End Labeling of DNA Oligonucleotides In all the following assays (except choice of nucleophile), double-stranded oligonucleotides are 5'-end labeled by T4 polynucleotide kinase (GIBCO-BRL/ Life Technologies). The single-stranded oligonucleotide (10 pmol) is incubated at 37 ° for 30 min in 50 #1 of kinase buffer containing 10/zCi of [y-32p]ATP (Amersham Pharmacia Biotech) and 10 units of kinase. The labeling solution is then applied to the top of a Sephadex G-25 Quick Spin column (Boehringer Mannheim) and the filtrate is annealed with 20 pmol of the complementary strand for 5 min at 95 ° and for 30 min at 37 °.

3'-End Labeling of DNA Oligonucleotides In one of the following assays (choice of nucleophile), the 21-mer singlestranded oligonucleotide A (Table I) is T-end labeled as follows. Ten picomoles of oligonucleotide A is incubated at 37 ° for 60 min in 50 #1 of terminal transferase buffer containing 10 #Ci of ~-cordycepin TP (NEN Life Science Products, Boston, MA) and 15 units of terminal transferase (GIBCO-BRL/Life Technologies). The labeling reaction results in the addition of a single 32p-labeled nucleotide. The labeling solution is then applied to the top of a Sephadex G-25 Quick Spin column and the filtrate is annealed with 20 pmol of oligonucleotide G (complementary strand) for 5 min at 95 ° and for 30 min at 37 °.

Standard Reaction Conditions Unless otherwise indicated, the integrase assays are performed in "integration buffer" containing 25 mM morpholinepropanesulfonic acid (MOPS, pH 7.2),

In Vitro HIV- 1 INTEGRASEASSAYS

[32]

629

TABLE I OLIGONUCLEOTIDESUSED IN ASSAYS

Oligonucleotide

Size

Sequence

A B C D E F G Au

21-mer 21-mer 19-mer !5-mer 34-mer 30-mer 22-mer 21-mer

5'-GTG TGG AAA ATC TCT AGC AGT-3' 5'-ACT GCT AGA GAT TTT CCA CAC-3' 5'-GTG TGG AAA ATC TCT AGC A-3' 5'-GAA AGC GAC CGC GCC-3' 5'-GTG TGG AAA ATC TCT AGC AGG GGC TAT GGC GTC C-3' 5'-GGA CGC CAT AGC CCC GGC GCG GTC GCT TTC-3' 5'-ACT GCT AGA GAT TTT CCA CAC T-3' 5'-GTG TGG AAA ATC TCT AGC UGT-3 ~

7.5 mM MnC12, 14.3 mM 2-mercaptoethanol, and bovine serum albumin (BSA; final concentration, 0.1 mg/ml).

Integration Assay Measuring 3'-Processing and Strand Transfer In the integration assay (Fig. 3A), a 21-mer double-stranded DNA oligonucleotide (oligonucleotide *A, 5'-end labeled and annealed to oligonucleotide B), corresponding to the last 21 bases of the U5 viral LTR (Table I), is used to follow both T-processing and T-end joining (strand transfer) reactions. In the 3'processing reaction, integrase liberates a GT dinucleotide at the 3'-end of the labeled strand, resulting in the generation of a 19-mer labeled product. The strand transfer (3'-end joining) reaction consists of the insertion of a T-processed oligonucleotide into another DNA target. This strand transfer leads to higher and lower molecular weight species migrating slower and faster, respectively, than the original 21-mer substrate (Fig. 3A). The higher molecular weight species (strand transfer products, STP) are generally used to evaluate strand transfer (integration). For inhibitor testing, the compound to be tested is incubated in integration buffer for 30 min at 37 ° with 400 nM HIV- 1 integrase and 5 nM 5'-labeled doublestranded DNA template (*A/B) in a total volume of 10 #1. The reaction is stopped by adding the same volume of electrophoresis denaturing dye containing 99% (v/v) formamide (Sigma-Aldrich, Milwaukee, WI), 1% (w/v) sodium dodecyl sulfate (SDS), bromphenol blue (0.2 mg/ml; Sigma-Aldrich), and xylene cyanol blue (0.2 mg/ml; Sigma-Aldrich). A preincubation for 15 min at 37 ° with either integrase or DNA can be adopted in order to provide advantage to the drug-protein or drug-DNA interaction, respectively. Samples are then heated for 5 min at 95 ° and loaded on 20% (w/v) acrylamide : bisacrylamide (19 : 1) denaturing gel (Accugel; National Diagnostics, Atlanta, GA) containing 7 M urea (GIBCO-BRL/Life Technologies) in TBE (GIBCO-BRL/Life Technologies).

630

ENZYMOLOGY AND BIOLOGICAL APPROACHES

[321

B

A

IN

A~

21 3AGT 21

B

3'-P ~

+

IN

C* 8

t9 2A 21

GT 19

19

S.T, ,'I

3A

21

3A

21

STP

21 3AGT

~*i~L

19

+

E D* F

- +

D

C

|5~110~ 15

IN

IN

~AGT*A T

22

BQi '~ qb|

3O

I/Glycerol or

~ Disintegration[ ~ ~ 4

~|

Water,

I~IA-3'OH

Glycerol-GT*A G TWA

o,~,l

IG) lCl

5'*OH-GT*A

FIG. 3. Catalytic activities in vitro of HIV-l integrase. The asterisk marks the location of the 32p radiolabel. Oligonucleotides are denoted by capital letters with reference to Table I. In the autoradiograms to the right of each panel, the left lane represents DNA alone and the right lane represents DNA plus integrase (1N). T-E Y-Processing; S.T., strand transfer (3' end joining); STP, strand transfer products. (A) Dual assay for 3'-processing and strand transfer, using a blunt-ended 21-mer; (B) strand transfer assay using a "precleaved" 19-mer; (C) disintegration assay using a Y-shaped oligonucleotidic structure; (D) assay to determine the choice of nucleophile, using a Y-labeled double-stranded 22-met oligonucleotide.

Strand Transfer Assay In the strand t r a n s f e r a s s a y (Fig. 3B), a " p r e c l e a v e d " 1 9 - m e r Y - e n d l a b e l e d o l i g o n u c l e o t i d e C is a n n e a l e d to its 2 1 - m e r c o m p l e m e n t a r y s t r a n d B. T h i s substrate, c o n t a i n i n g a r e c e s s e d Y - e n d , is u s e d to d e t e r m i n e w h e t h e r the 3 ' - e n d j o i n i n g

[32]

In Vitro HIV- 1 INTEGRASEASSAYS

63 t

reaction is truly inhibited by a compound or whether the decrease in strand transfer products observed in the integration assay is primarily due to an inhibition of the T-processing step. The conditions of this assay are the same as those used for the integration assay measuring T-processing and strand transfer (see above).

Disintegration Reaction The integration reaction (Fig. 3C), catalyzed in vitro by integrase, is in equilibrium with the reverse reaction called disintegration. Disintegration is the only reaction that the core enzyme (without the N- and C-terminal domains) can catalyze. Therefore the disintegration reaction is commonly used to test whether inhibitors act on the integrase core domain. Disintegration can be monitored by using an oligonucleotide that mimicks an integrated product (Fig. 3C). In this reaction, 10 pmol of a 15-mer 5'-end labeled single-stranded oligonucleotide *D is annealed with 20 pmol of oligonucleotides E, B, and F to form a branched Yshaped double-stranded structure (see 5'-End Labeling of DNA Oligonucleotides, above). During catalysis, HIV-I integrase removes the branch and liberates a 30-met 5'-end labeled double-stranded oligonucleotide. Conditions used in this assay are identical to those used for the integration assay.

Choice of Nucleophile in 3~-Processing Reaction The choice of nucleophile in the T-processing reaction (phosphodiester cleavage) can be analyzed with a 22-mer double-stranded DNA oligonucleotide (oligonucleotide A* T-end labeled and annealed to oligonucleotide G) (see 3'-End Labeling of DNA Oligonucleotides, above). Water, glycerol, or T-OH DNA can be used by integrase as nucleophiles. Products resulting from hydrolysis (L), glycerolysis (G), and nucleotide circularization (C) can be monitored by gel electrophoresis (Fig. 3D). If all these products are inhibited to the same extent by a compound, this compound can be described as exerting a global inhibition on the 3'-processing reaction. The reaction conditions are similar to those used in the integration assay.

DNA-Binding Assays Two binding assays are used to monitor integrase-DNA interactions. The first is a photocross-linking assay in which 254 nm ultraviolet light is used to generate free radicals responsible for cross-linking HIV- 1 integrase to the DNA (Fig. 4A). In this assay, the compound to be tested is incubated in integration buffer containing no BSA for 15 min at 37 ° with 1.5 # M HIV- 1 integrase and 5 nM 5'-labeled doublestranded DNA template (A*/B) in a total volume of 10 #1. BSA is removed from the integration buffer to prevent formation of nonspecific photocross-linking products between BSA and the target DNA template. Samples are then irradiated for 15 min under a 254 nm portable UV lamp (UVP, Upland, CA) at room temperature and at

632

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a distance of 1 cm from the light source. One volume of 2x SDS-Tricine loading buffer (Novex) is added to the samples, which are then heated 5 min at 95 ° and resolved in a 12-20% gradient (w/v) Tricine-SDS-polyacrylamide precast gel (Novex). The second method depends on covalent chemical trapping, via reduction with sodium borohydride, of a Schiff base formed between an appropriately positioned e-amino group of a lysine present on the enzyme, and an aldehydic abasic site, enzymatically introduced into the DNA (Fig. 4B). 8 In this Schiff base assay, an abasic site is generated in the DNA template (A*/B) by using a 21-mer single-stranded oligonucleotide that contains a uracil base incorporated in the oligonucleotide (oligonucleotide Au). The Y-end labeled Au oligonucleotide is annealed to its 21mer complementary strand (oligonucleotide B), and treated in the kinase buffer with 1 unit of uracil DNA glycosylase (GIBCO-BRL/Life Technologies) for 60 min at 37 °. The labeling solution is then applied to the top of a Sephadex G-25 Quick Spin column. In this assay, the compound to be tested is incubated in integration buffer containing no BSA for 15 min at 37 ° with 1.5 # M HIV- 1 integrase and 5 nM Y-labeled abasic site-containing double-stranded DNA template (*Au/B) in a total volume of 10/zl. The covalent trapping is initiated by reduction of the Schiff base, using I/zl of fresh 1 M sodium borohydride. After 5 rain at room temperature, samples are treated with I volume of 2x SDS-Tricine loading buffer, heated for 5 min at 95 °, and loaded on 12-20% gradient (w/v) Tricine-SDS-polyacrylamide gels. Conclusion Success obtained with highly active antiretroviral therapy (HAART) has changed the prognosis of acquired immunodeficiency syndrome (AIDS). However, these treatments are limited by side effects and viral resistance. Discovering inhibitors for new targets such as HIV-1 integrase will help to decrease viral load and emergence of drug resistance. The in vitro HIV-1 integrase assays described here can be modified for high-throughput screening 9 and are routinely used for the discovery of HIV-1 integrase inhibitors for the treatment of AIDS.

s A. Mazumder, N. Neamati, A. Pilon, S. Sunder, and Y. Pommier,J. Biol. Chem. 271, 27330 (1996). 9 D. J. Hazuda, J. C. Hastings, A. L. Wolfe, and E. A. Emini, Nucleic Acids Res. 22, 1121 (1994).

FIG.4. DNA-binding activities of HIV-1 integrase. The asterisk marks the location of the 32p radiolabel. Oligonucleotidesare denotedby capital letters with referenceto TableI. In the autoradiogram to the right of each panel, the left lane represents DNA alone and the right lane represents DNA plus integrase. DPC, DNA-proteincross-links.(A) Photocross-linkingassayfor DNAbinding; (B) chemical cross-linking assay for DNA binding.

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[33] Use of X e n o p u s Egg Extracts to Study Effects of DNA-Binding Drugs on Chromatin Assembly, Nuclear Assembly, and DNA Replication B y ASMITA KUMAR, HONGZHI XU, a n d GREGORY H. LENO

Introduction

Xenopus egg extract has been extremely valuable for the analysis of numerous integral processes in cell biology including chromatin assembly] ,2 nuclear assembly and nuclear protein import, 3'4 DNA replication, 5 and cell cycle control. 6 One crucial feature of the Xenopus egg, and egg extract, that makes it so useful for studying these processes is the stockpile of components that are required to support the rapid cell division cycles following fertilization. The female germ cell or oocyte is arrested in prophase of meiosis I within the ovary of the adult frog. During this stage of meiotic arrest oocyte growth or oogenesis occurs, resulting in the production of a stockpile of macromolecules and organelles that are required to support the rapid cell cycles in the early embryo. Fully grown oocytes are then induced to complete meiosis I and enter a second stage of arrest in metaphase of meiosis II. The mature oocyte then passes down the oviduct and is released by the frog as an "unfertilized egg." Extracts are prepared from these unfertilized eggs. On fertilization, the egg is released from metaphase arrest and enters interphase, with the first mitotic cell cycle lasting approximately 90 min and the next 11 cycles lasting only 30 min each. The stockpile of components present within the egg not only supports these remarkably rapid embryonic cell cycles in vivo, but also supports the decondensation and remodeling of sperm chromatin, the assembly of remodeled chromatin into functional nuclei, and the replication of nuclear DNA in egg extracts, mimicking the events within the intact egg. Thus, this cell-free system has remarkable potential for determining the effects of DNA-binding drugs on replication-related processes, with the goal of identifying mechanism(s) leading to cytotoxicity and the possible rational design of new anticancer drugs. In this chapter, we describe how to establish and maintain a Xenopus frog colony, how to obtain unfertilized eggs and prepare egg extracts, and how to isolate sperm chromatin as a source of template DNA. We then provide a general t A. R Wolffe and C. Schild, Methods Cell Biol. 36, 541 (1991). 2 G. H. Leno, Methods Cell Biol. 53, 497 (1998). 3 D. D. Newmeyer and K. L. Wilson, Methods Cell Biol. 36, 607 ( 1991 ). 4 M. J. Lohka, Methods" Cell Biol. 53, 367 (1998). 5 G. H. Leno and R. A. Laskey, Methods Cell Biol. 36, 561 (1991 ). 6 A. W. Murray, Methods CellBiol. 36, 581 (1991).

METHODS IN ENZYMOLOGY~VOL.34(1

Copyright~) 2001 by AcademicPress All rightsof reproductionin any formreserved. (X176-6879/00$35.0(I

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approach for the analysis of drug effects on the remodeling of sperm chromatin, nuclear assembly, and DNA replication, and provide detailed methodology for analysis of each of these processes. Finally, we provide examples of drug effects on nuclear assembly and DNA replication, using this cell-free system. Establishing and Maintaining Frog Colony

Xenopus laevis females may be purchased from one of several supply companies (e.g., Nasco, Fort Atkinson, WI; Xenopus I, Ann Arbor, MI). Frogs may be purchased immature or as sexually mature females. However, given that it may be several weeks before high-quality eggs can be obtained from newly purchased sexually mature frogs, obtaining immature females and housing them until maturity may be the most cost-effective approach. This is especially true if mature females are present within an established colony and new frogs are obtained to replace older animals or to increase colony size. Sperm cells may be obtained immediately after purchase of sexually mature male frogs. Frogs are maintained in covered polyethylene tanks, 66 cm long ×45 cm wide x 30 cm high. The animals are housed at or below a density of one frog per 4 liters of water. Tank water is supplied as well water filtered through activated charcoal and water quality (volatile organic chemicals) is analyzed quarterly. Charcoal filters are changed as needed but no less than twice per year. Ambient room temperature is maintained at approximately 20 °, with the optimal temperature range between 17 ° and 22 °. Illumination is maintained according to a 12-hr light and 12-hr dark cycle. Frogs are fed daily, Monday-Friday. Approximately 1.5 g of Purina Trout Chow is provided for each female frog whereas the smaller, male flogs are provided approximately 1 g of chow each. One hour after adding food pellets to the tanks, any remaining food is removed with a fine mesh fish net. Tank water is exchanged on Monday, Wednesday, and Friday of each week after feeding. Water is removed from each tank with a pump and the tank walls are scrubbed with a brush to remove any debris. The tank is then filled with fresh charcoal-filtered water. The flogs remain within the tank during this procedure and exhibit no adverse effects. Every 2 weeks the tanks are sanitized by thoroughly scrubbing all surfaces with 2 M NaC1. All debris is removed and the tanks are rinsed with charcoal-filtered water. Water pumps and tubing are sanitized in a like manner. Each frog is removed and placed into a freshly cleaned tank prior to the sanitation process. Obtaining Unfertilized Eggs from Sexually Mature Frogs Mature oocytes are arrested at metaphase of meiosis II within the ovary, awaiting ovulation and fertilization. Unfertilized eggs, from which extracts are prepared, are obtained in the following way. Each frog is primed for ovulation by injection

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of 100 IU of pregnant mare serum gonadotropin (PMSG) 4 days before egg collection. This results in follicle cell stimulation and oocyte growth. Twelve to 15 hr before egg collection, each primed frog is injected with 500 IU of human chorionic gonadotropin (HCG) to induce ovulation. Hormone is prepared in deionized water and administered by subcutaneous injection into the dorsal lymph sac. The frogs are then placed in individual tanks containing high-salt Barth's solution [110 mM NaCI, 15 mM Tris-HC1 (pH 7.4), 2 mM KCI, 2 mM NaHCO3, 1 mM MgSO4, 0.5 mM Na2HPO4],7 which prevents spontaneous activation of the eggs. After egg laying slows or stops, the frogs are returned to filtered water and allowed to recover for a minimum of 3 months before they are reinjected with hormone.

Preparing Extracts from Unfertilized Eggs Eggs from each frog are transferred in high-salt Barth's solution to a l-liter glass beaker for processing, keeping the eggs from each frog separate. The external jelly coat of the egg is removed by incubation in 2% (w/v) cysteine hydrochloride, pH 7.8-7.9, for approximately 10 min at room temperature. The beaker should be rocked during incubation to facilitate removal of the jelly coat, which results in the eggs "settling" together, with no visible space between adjacent eggs. Dejellied eggs are then rinsed with Barth's solution [88 mM NaC1, 15 mM Tris-HC1 (pH 7.6), 2 mM KC1, 1 mM MgC12, 0.5 mM CaC12](7) until debris is no longer visible in the rinse solution. Extract quality will be improved if discolored or misshapen eggs are removed before further processing. After eggs are dejellied, they are artificially "activated." Activation is the entry of the cell into interphase and results in contraction of the pigment in the animal hemisphere. This is achieved by incubation of the eggs with the calcium ionophore A23187 for approximately 5 min (0.2-0.5 #g/ml in Barth's solution). After activation, eggs are rinsed extensively in ice-cold extraction buffer [50 mM HEPES-KOH (pH 7.4), 50 mM KC1, 5 mM MgC12, 2 mM 2-mercaptoethanol] (7). After the final rinse in extraction buffer, eggs are gently poured into ice-cold centrifuge tubes. Transferring in a larger volume of buffer will avoid flattening of the eggs against the glass beaker, which can cause them to rupture. After the eggs have settled to the bottom of each tube, much of the excess buffer can be removed with a pipette. The preparation and activities of egg "low-speed" supernatant (LSS) and "high-speed" supematant (HSS) are illustrated in Fig. 1. The eggs are packed by FIG. 1. The preparation of Xenopus egg extracts and their activities. Stage 1 decondensation (adapted from Lu et al?8). Chromatin remodeling: The core histones H2A, H2B, H3, and H4, the spermspecific basic proteins X and Y, the embryonic linker histone B4 (arrowheads), and the high mobility group protein 1 (HMG 1, arrow), are indicated in these two-dimensional Triton-acid-urea (TAU)-SDSPolyacrylamide gels stained with Coomassie blue (from Philpott and Leno 13). Replication time course (from Leng and Lenol°).

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centrifugation at 134g for 1 min at 4 ° in an SW50. 1Ti rotor (Beckman, Fullerton, CA). Buffer should be removed from the surface to prevent excessive dilution of the extract. To separate the various egg components, packed eggs should be crushed by centrifugation at 21,000g for 10 min at 4 ° in an SW50.1Ti rotor (Beckman). Egg components separate into three distinct fractions: a lipid cap, a golden cytoplasm layer, and a plug of yolk platelets and pigment. The cytoplasmic layer is collected by breaking through the lipid cap with a cooled Pasteur pipette. Cytochalasin B, aprotinin, pepstatin A, and leupeptin are all added to the cytoplasmic "extract" to a final concentration of 10 #g/ml. The extract obtained after crushing of the eggs is contaminated with yolk, pigment, and lipid material. This material is "cleared" from the extract by centrifugation at 84,000g for 15 min at 4 c'. The cleared extract, or LSS, may be used to study stage I decondensation and remodeling of sperm chromatin, nuclear assembly and stage II decondensation, and DNA replication (Fig. 1). If LSS is the extract of choice, it may be used fresh or frozen as beads in liquid nitrogen for later use. To prevent damage, frozen extracts should be supplemented with glycerol to 1% (v/v). Separation of the soluble extract components from membrane vesicles is achieved by high-speed centrifugation of the LSS. s Approximately 1.5 ml of LSS is placed in a precooled ultraclear (Beckman) centrifuge tube and overlaid with chilled liquid paraffin. The extract is centrifuged at 114,000g for 1 hr at 4 ° in an SW50.1Ti rotor (Beckman). Three major fractions are produced: a transparent soluble fraction or HSS; a membrane vesicle fraction; and a golden, hard ribosomal pellet (Fig. 1). If fractionation is incomplete after 1 hr, the extract should be centrifuged for an additional 30 rain. If the initial volume of LSS is > 1.5 ml, further centrifugation will nearly always be required. The HSS and membrane fractions are collected by side puncture of the tube with a syringe. As with the LSS, the HSS can be used fresh or frozen for later use after addition of glycerol to 7% (v/v). The HSS is able to decondense and remodel sperm chromatin (Fig. 1) but is unable to assemble this chromatin into a functional nucleus and replicate DNA. Under optimal conditions, nuclear assembly and DNA replication can be reconstituted by combining the HSS with the membrane fraction. It is important to note that not all frogs injected with hormone will produce eggs. Furthermore, even if eggs are produced, their quality or quantity may make extract preparation difficult, if not impossible. We find that in a typical group of animals, one-half to three-quarters of the frogs lay eggs that can be used for extract preparation. It is also important to note that the quality of extract is reduced when eggs are allowed to remain in high-salt Barth's solution for long periods of time. Thus, once a sufficient number of high-quality eggs have been produced,

7 j. j. Blowand R. A. Laskey,Cell 47, 577 (1986). M. A. Sheehan,A. D. Mills, A. M. Sleeman,R. A. Laskey,andJ. J. Blow,J. CellBiol. 106, l (1988).

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EXTRACTS

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they should be processed into extract immediately. All these factors should be considered when determining the number of frogs needed for an injection series. The number of eggs needed to make an extract depends on the chosen method of preparation. If large-scale preparation procedures are employed, as described here, a minimum of "-~1000 eggs is required. This can be a disadvantage, especially if smaller frogs, which often produce fewer eggs than larger animals, are used. Smallscale preparations, using benchtop centrifugation procedures, 9 require fewer eggs and may be the method of choice under these conditions. However, preparation of extract in bulk offers many advantages including the thorough characterization of a single extract and its subsequent use in many experimental settings. "Pooling" eggs from different frogs to increase quantity is not recommended even if the eggs appear to be of similar quality. One bad batch of eggs can adversely affect the activity of the whole pooled extract. P r e p a r a t i o n of Xenoptrs S p e r m Nuclei To prepare Xenopus sperm nuclei, 1° six male X. laevis frogs are injected with 50 IU of HCG one week prior to the removal of the testes. Frogs are killed by injection of a lethal dose (120 mg/frog) of tricaine methane sulfonate (MS-222) into the dorsal lymph sac. Administration of MS-222 is considered an acceptable method of euthanasia for amphibian species by the American Veterinary Medical Association. 11 Testes are then removed from the frogs by dissection and placed in 25 ml of modified Barth's saline [MBS; 20 mM HEPES-KOH (pH 7.5), 110 mM NaC1, 1 mM KC1, 0.75 mM CaC12, 0.82 mM MgSO4, 2.4 mM NaHCO3] in a petri dish on ice. Testes are homogenized by chopping them with a razor followed by repeated passage through a 10-ml syringe without a needle. This material is then transferred to two 50-ml conical tubes and the petri dish is rinsed with an additional 20 ml of MBS. The supernatant is then split into two aliquots and each is layered over 10 ml of 70% (v/v) Percoll, prepared in modified Barth's saline, in a 50-ml conical tube. Mature sperm cells are then sedimented by centrifugation at 1510g for 30 min at 4 °. A single centrifugation step is normally not adequate for the recovery of all mature sperm from the testicular homogenate. Therefore, this material should be removed, gently shaken to loosen clumps of debris, layered on fresh Percoll, and recentrifuged as described above. The sperm pellets are then resuspended in buffer NIM [10 mM HEPES (pH 7.5), 2.5 mM MgC12, and 0.2 M sucrose], combined, and resedimented by centrifugation at 1730g for 15 min at 4 °. The pellet is then resuspended in 10 ml of buffer NIM containing lysophosphatidylcholine (LPC, 1.0 mg/ml) with 100 mM phenylmethylsulfonyl fluoride, 9 j. Newport, Cell 48, 205 (1987). ~o F. Leng and G. H. Leno, J. Cell. Biochem. 64, 476 (1997). It E. J. Andrews, J. Ant. Vet. Med. Assoc. 202, 229 (1993).

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and kept on ice for 15 min. During this period the sample should be mixed by inverting the tube every minute. LPC disrupts the plasma and nuclear membranes, allowing free movement of macromolecules into the cell. This membrane "permeabilization" is stopped by adding 5 ml of ice-cold buffer NIM containing 3% (w/v) bovine serum albumin (BSA). Permeabilized sperm nuclei are then pelleted by centrifugation at 430g for 15 min at 4 °, washed once in buffer NIM containing 0.4% (w/v) BSA, and finally resuspended in buffer XN-50% (v/v) glycerol [buffer XN: 50 mM HEPES-KOH (pH 7.0), 250 mM sucrose, 75 mM NaC1, 0.5 mM spermidine, 0.15 mM spermine]. The DNA concentration can be determined by counting nuclei with a hemocytometer and assuming a DNA mass of 3.15 pg per haploid genome. 12 On average, >100 #g of DNA is obtained from the testes of a single frog. The nuclei are stored at - 8 0 ° in buffer XN-50% (v/v) glycerol. S t r a t e g i e s for A n a l y s i s of D r u g E f f e c t s o n C h r o m a t i n R e m o d e l i n g , N u c l e a r A s s e m b l y , a n d DNA R e p l i c a t i o n in Xenopus E g g E x t r a c t There are three general strategies for analysis of drug effects on the decondensation and remodeling of sperm chromatin, the assembly of remodeled chromatin into functional nuclei, and DNA replication (Fig. 2). In the first approach (Fig. 2A), drug, Xenopus sperm nuclei (XSN), egg extract (LSS or HSS), and a reaction mixture (Rx Mix) are combined and incubated at 23 °. The type of extract (LSS vs. HSS), and the time of incubation, will depend on the process to be analyzed (see Fig. 1). In the second approach (Fig. 2B), drug and XSN are preincubated in buffer for 15-30 rain to promote drug-DNA interaction in the absence of extract. The XSN are then sedimented by centrifugation at 770g for 10 min at 4 ° and the buffer containing the drug is removed. Egg extract (LSS or HSS), with or without drug, and Rx Mix are then added and the total mixture is incubated as described above. In the third approach (Fig. 2C), drug is preincubated with extract (LSS or HSS) for 15-30 min to facilitate interaction of drug with extract components in the absence of DNA. The XSN and Rx Mix are then added and the sample is incubated as described above. Chromatin Remodeling In general, the remodeling of sperm chromatin that accompanies stage I decondensation involves the replacement of sperm-specific basic proteins (e.g., X and Y) with histones H2A and H2B from the egg, resulting in the formation of nucleosome cores. It is now clear that in Xenopuseggs, this remodeling is mediated by specific factors that participate in both assembly and disassembly processes. 13 In addition 12 I. B. Dawid, J. Mol. Biol. 12, 581 (1965).

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to the core histones, the embryonic linker histone B4, and high mobility group protein I (HMG1), are assembled on sperm chromatin by the extract. T M These changes in chromatin composition and structure occur within the first 3 to 10 rain of incubation and can be analyzed by one- or two-dimensional polyacrylamide gel electrophoresis and micrococcal nuclease digestion (Fig. 3).

Analysis of Chromatin Remodeling by Egg Extract Fresh or freshly thawed Xenopus egg extract (LSS or HSS) is supplemented with (1) a reaction mixture (Rx Mix) containing an energy-regenerating system [creatine phosphokinase (150 lzg/ml), 60 mM creatine phosphate],7 cycloheximide to a final concentration of 100/zg/ml, ATP to a final concentration of 2 mM, and deoxynucleoside triphosphates (dNTPs) to readjust pool sizes to 50 # M 7' 15 after J3 A. Philpott and G. H. Leno, Cell69, 759 (1992). 14S. Dimitrov, G. Almouzni,M. Dasso, and A. E Wolffe,Dev.Biol. 160, 214 (1993). 15C. J. Hutchison, R. Cox, R. S. Drepaul, M. Gomperts, and C. C. Ford, EMBOJ. 6, 2003 (1987).

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dilution of the extract16; (2) drug or an equal volume of diluent alone; and (3) XSN at 77 ng of DNA per microliter of extract. The extent of extract dilution is always kept constant at 30% of the initial volume for all incubations. Samples are incubated at 23 ° for 3 to 10 min. Electrophoretic Analysis of Chromatin Proteins To isolate chromatin-associated proteins after incubation of sperm nuclei in Xenopus egg extract, the mixture should be diluted with buffer A [15 mM TrisHC1 (pH 7.4), 60 mM KC1, 15 mM NaC1, 1 mM 2-mercaptoethanol, 0.5 mM spermidine, 0.15 mM spermine] and the nuclei sedimented by centrifugation at 770g for 10 min at 4 °. The nuclei are then resuspended in buffer A, centrifuged under the same conditions, and finally resuspended in a small volume of buffer H [20 mM H E P E S - K O H (pH 7.4), 50 mM KC1]. Basic proteins are extracted from the sperm chromatin by addition of HC1 to a final concentration of 0.5 M. The sample is immediately centrifuged at 16,000g for 10 rain at 4 ° and the resultant supematant is frozen in liquid nitrogen and lyophilized. For one-dimensional analysis, lyophilized proteins are dissolved in water, combined with 2× sodium dodecyl sulfate (SDS) sample buffer and separated by SDS-polyacrylamide gel electrophoresis (SDS-PAGE) on a 17% (w/v) gel, using the Laemmli buffer system, 17 and visualized with Coomassie blue stain or by silver stain (Fig. 3). Is For two-dimensional analysis by Triton-acid-urea (TAU) (first dimension, left to right)/SDS-PAGE (second dimension, top to bottom), lyophilized proteins are dissolved in 8 M urea, 5% (v/v) acetic acid, 4% (v/v) 2-mercaptoethanol, and 0.01% (w/v) pyronin Y, loaded onto a 1.5-mm tube gel containing 15% (w/v) acrylamide, 8 M urea, and 6 mM Surfact-Amps Triton X-100 (Pierce, Rockford, IL), and separated by running for 16 hr at 70 V. Tube gels are removed and equilibrated for 20 to 30 min with buffer [80 mM Tris-HC1 (pH 6.8), 50 mM dithiothreitol (DTT), 2% (w/v) SDS, 0.01% (w/v)bromphenol blue] and loaded onto 17% (w/v) SDS-polyacrylamide slab gels run using the Laemmli buffer system. 17 After electrophoresis, slab gels are fixed and stained with silver stain or with 0. 1% (w/v) Coomassie Blue R-250 dissolved in 50% (v/v) J6L. S. Cox and G. H. Leno, J. Cell Sci. 97, 177 (1990). 17U. K. Laemmli,Nature (London) 227, 680 (1970). 18Z. H. Lu, D. B. Sittman, D. T. Brown, R. Munshi, and G. H. Leno, J. Cell Sci. 110, 2745 (1997).

FIG. 3. Analysis of chromatin remodeling in Xenopus egg extracts. SDS-PAGE(adapted from Lu et al?8). TAU-SDS two-dimensional PAGE: The core histones H2A (2A), H2B (2B), H3 (3), and H4 (4), and the sperm-specific basic proteins X and Y, are indicated (from Philpott and Lenol3). Micrococcalnuclease digestion of chromatin (adapted from Lenoe).

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ENZYMOLOGY AND BIOLOGICAL APPROACHES

[33]

methanol-7% (v/v) acetic acid. Gels stained with Coomassie blue are destained in 5% (v/v) methanol-7% (v/v) acetic acid 13 (Fig. 3). M i c r o c o c c a l N u c l e a s e D i g e s t i o n of C h r o m a t i n The enzyme micrococcal nuclease has been used to investigate the subunit structure of sperm chromatin before and after decondensation in egg extracts. 13,J9 Specifically, sperm nuclei are used without extract incubation (XSN) or after incubation in egg extract (LSS or HSS) for 3 to 10 min at a DNA concentration of 77 ng/#l extract. Samples are diluted with 10 volumes of buffer A and overlaid onto a 90% (v/v) Percoll cushion in buffer A. Sperm nuclei are sedimented onto the Percoll cushion by centrifugation at 770g for 10 min at 4 °. Chromatin (5 ~g of DNA) is resuspended in 200 #1 of MN digestion buffer [ 15 mM Tris-HCl (pH 8.5), 60 mM KC1, 15 mM NaC1, 0.34 M sucrose, 15 mM 2-mercaptoethanol, 0.5 mM spermidine, 0.15 mM spermine, 2 mM CaC12] and incubated with 0.05 U of micrococcal nuclease (Sigma, St. Louis, MO) at 23 ° for 3 min. Two hundred microliters of MN termination buffer [20 mM Tris-HCl (pH 8.0), 20 mM EDTA, 0.5% (w/v) SDS] is added along with proteinase K (0.5 mg/ml; Sigma) followed by incubation for 16 hr at 37 °. DNA is then purified by phenol-chloroform extraction followed by ethanol precipitation. Dried samples are dissolved in TE buffer [10 mM Tris-HCl (pH 7.4), 1 mM EGTA] containing RNase A (1 Izg/#l) and incubated at 37 ° for 1 hr. DNA fragments are resolved on a 1.8% (w/v) Metaphor agarose gel (FMC, Rockland, ME) with TBE buffer in an 8-V/cm electric field. The gel is stained with ethidium bromide (Fig. 3). The remodeling of XSN by egg extract results in the formation of nucleosomes (in Fig. 3, compare XSN with XSN + LSS). Nucleosome repeat length is determined by estimating the size of DNA fragments on the basis of electrophoretic mobility. DNA molecular weight markers at 100-base pair (bp) intervals are used to plot the standard electrophoretic mobility versus DNA fragment size interpolation curve.

Selecting Appropriate System Both low-speed and high-speed supernatants (LSS and HSS) promote stage I decondensation and remodeling of sperm chromatin.13'20 However, chromatin remodeling in LSS (Fig. 3) is more efficient than in HSS (Fig. 1). The basis for this difference is not clear, but could involve the loss of specific remodeling factors from the LSS after high-speed centrifugation or, alternatively, a reduction in remodeling factor efficiency in the HSS. In spite of this disadvantage, the HSS may be the extract of choice when drug-membrane interactions need to be avoided or if long exposures of DNA to drug are necessary without the potential complications

19 S. Dimitrov, M. C. Dasso, and A. E Wolffe, J. Cell Biol. 126, 591 (1994). 20 A. Philpott, G. H. Leno, and R. A. Laskey, Cell 65, 569 (1991).

[33]

USE OF Xenopus EGG EXTRACTS

645

associated with the formation of higher order chromatin structure, nuclear assembly, and DNA replication Nuclear Assembly A functional nucleus is assembled from remodeled sperm chromatin after approximately 30 min in egg LSS. This involves the formation of higher order chromatin structure, the assembly of an intact nuclear membrane, the import of nuclear protein, the formation of a nuclear lamina, and chromatin swelling (stage II decondensation). Three features of nuclear structure and function, namely, the assembly of the nuclear membrane and the nuclear lamina, and the import of nuclear protein (Fig. 4), are routinely used to assess nuclear assembly in egg extract.

Assembly of Nuclear Membrane Determining the presence of a nuclear membrane surrounding decondensed sperm chromatin can be achieved with the lipid dye, Nile red (Sigma). Nile red fluoresces primarily within a hydrophobic environment, thereby reducing background fluorescence in the assay. Nuclear membranes are visualized by placing an aliquot of unfixed sample (e.g., 2 #1) on a slide and mixing it with an aliquot of Nile red (0.15/zg/ml) and Hoechst 33258 (5 #g/ml) to label total DNA. A continuous, peripheral Nile red fluorescence (Figs. 1 and 4) indicates complete nuclear membrane formation around sperm chromatin (Fig. 1) in the extract. 16 The anthracycline antibiotic daunomycin has been shown to disrupt nuclear membrane formation in the extract in a concentration-dependent manner. ~0At high drug concentrations (>20 #M) membrane vesicles do not associate with sperm chromatin (Fig. 5), whereas at a lower drug concentration (5/~M) membrane assembly is complete within most nuclei (see Fig. 3 in Leng and Lenol°).

Assembly of Nuclear Lamina The formation of a complete nuclear lamina by egg extract is a late event in the process of nuclear assembly. 21'22 After formation of a transport-competent nuclear membrane (see below), lamin LjII, the most abundant lamin species in egg extract, accumulates within the nucleus, leading to the formation of a complete lamina structure. 23 If the majority of lamin Lln is removed from egg extract by immunodepletion, the resultant nuclei are unable to initiate replication, 24-26

21 C. J. Hutchison, R. Cox, and C. C. Ford, Development 103, 553 (1988). 22 C. J. Hutchison and I. Kill, J. Cell Sci. 93, 605 (1989). 23 C. J. Hutchison, J. M. Bridger, L. S. Cox, and I. R. KillJ. Cell Sci. 107, 3259 (1994). 24 j. W. Newport, K. L. Wilson, and W. G. Dunphy, J. Cell Biol. 111, 2247 (1990). 25 j. Meier, K. H. S. Campbell, C. C. Ford, R. Stick, and C. J. Hutchison, J. Cell Sci. 98, 271 ( 1991). 26 H. Jenkins, T. Holman, C. Lyon, B. Lane. R. Stick, and C. Hutchison, J. Cell Sei. 106, 275 (1993).

646

[33]

ENZYMOLOGY AND BIOLOGICAL APPROACHES

Nut:lear Assembly XSN LSS Rx Mix +/- Drug

Jna Ni'°arLam' 30'

Structural Analysis: Nuclear Membrane &

Nuclear Membrane

Functional Analysis: Nuclear Protein Import

Nucle~arProtsth Imoort Transport probe added T7 RNA Polymarase+/- NLS

Nuclear Lamina

Unfixed sample on slide

~0'-1

h

Samples diluted 70-fold with buffer A Overleyed on 30% Sucrose in buffer A

Lipid dye: Ni~eRed (0.15pg/ml) DNA stain: Hoechst 33258 (5~g/rnl)

~

Nuclear Membrane

- - Scin~lletion ~al

Diluted sample

/I

300 Sucrose

Polylysine.coated coverslip Centdfuge 770xg 5' 230C

Nuclei pelleted on coverslips Rinsed with D-PSS Fixed in 4% Paratarmaldehyde/D-PBS10' 230C Rinsed with D-PBS Membranes solubtlizedwith Methanol.20oC t0'

/

Rinsed with D-PBS Blocked twice with TTBS-SSA

Probe with primely antibody Mouse anti-lamin LIII monoolonalantibody Lo46F7(t :250) 1h 23OC

4

Rinsed with TTBS-BSA Sheep anti-mouse Texas Red-conjugated antibody (1:25) r l h 23oc Rinsed with TTBS-BSA DNA stainedwith 5p,g/mlHoechst33258 Nuclear Lamina

Probe with primely antibody Rabbit anti-T7 RNA polyrnaraseantiserum(1:10,000) ~ 1h23~ Rinsed with TTBS-BSA Donkey anti-rabbitfluorescein-conjugatedantibody (1:25) ~ lh23~C Rinsed with "I-FBS-BSA DNA stained with 5pg/ml Hoechst 33258 Nuclear Transport TT-NLS T7+NLS

FIG. 4. Analysis of nuclear assembly in Xenopus egg extracts. Nuclear lamina (from Lu et al. 18).

[33]

USE OF Xenopus EGG EXTRACTS

647

Nuclear Membrane

Daunomycin (50 I.tM) FIG. 5. A high concentration of daunomycin inhibits assembly of the nuclear membrane in Xenopus egg extract. Xenopus sperm nuclei and daunomycin (50 #M) were incubated simultaneously (Fig. 2A) in egg extract for 90 min. Unfixed nuclei were stained with Nile Red and viewed by confocal microscopy. These nuclei lack the continuous peripheral fluorescence surrounding control nuclei (Fig. 4, Nuclear Membrane) indicating that assembly of the nuclear membrane is incomplete. The fluorescence associated with the sperm chromatin is due to autofluorescence of daunomycin that is visible at high drug concentrations (from Leng and Lenol°).

demonstrating that initiation of DNA replication in egg extract requires the formation of a complete nuclear lamina. To assay nuclear lamina assembly around sperm chromatin, each sample is diluted 70-fold with buffer A, then layered on 30% (w/v) sucrose in buffer A, and the nuclei are sedimented onto a polylysine-coated coverslip by centrifugation at 770g for 5 min at 23 °. The coverslip containing sedimented nuclei is then rinsed in Dulbecco's phosphate-buffered saline (D-PBS) and the nuclei are fixed by incubation in 4% (w/v) paraformaldehyde-D-PBS for 10 min at 23 °. The coverslip is then rinsed in D-PBS and treated with methanol at - 2 0 ° for 10 min. The use of both paraformaldehyde and methanol may not be necessary or even beneficial in all cases. The coverslip is then rinsed in D-PBS, blocked twice with TTBS-BSA [0.1% (v/v) Tween 20, 25 mM Tris-HC1, (pH 8.0), 137 mM NaC1, 2.7 mM KC1, 1.5% (w/v) BSA], and probed for 1 hr or longer at 23 ° with the primary antibody (e.g., mouse anti-lamin Llu monoclonal antibody, L046F72v diluted with T T B S BSA (e.g., 1 : 250). The coverslip is then rinsed with TTBS-BSA and incubated for 1 hr at 23 ° with a fluorochrome-conjugated secondary antibody (e.g., sheep anti-mouse Texas Red antibody) diluted with TTBS-BSA (e.g., 1 : 25). The coverslip is then rinsed with TTBS-BSA and stained with Hoechst 33258 to label DNA. An example of nuclear lamina staining is shown in Fig. 4.18

27 D. Lourim, A. Kempf, and G. Krohne, J. Cell Sci. 109, 1775 (1996).

648

ENZYMOLOGY AND BIOLOGICALAPPROACHES

[33]

Nuclear Protein Import

Nuclear transport is essential for DNA replication in Xenopus egg extract. 28 To determine whether extract-assembled nuclei are capable of nuclear protein import, one can use a transport probe such as bacteriophage T7 RNA polymerase. Natural T7 RNA polymerase is too large to diffuse through the nuclear pore complex, and as such, is excluded from an intact nucleus. However, T7 RNA polymerase, constructed with the nuclear localization signal (NLS) from the simian virus 40 (SV40) large-T antigen, 29 is actively transported through the nuclear pore complex and accumulates within an intact nucleus. 3° T7 RNA polymerase, with the NLS (T7 + NLS) or without it ( T 7 - N L S ) , is expressed in Escherichia coli BL2131 and purified according to the procedure described by Zawadzki and Gross. 32 Nuclear protein import is assayed in the following way. Sperm chromatin, at 1-3 ng of DNA per microliter of LSS, is incubated for 30 min to allow for nuclear assembly. T7 RNA polymerase is then added to the extract and incubated an additional 30 min to 1 hr, giving a total time in extract of 60 to 90 min. Samples are processed as described for the nuclear lamina, and as illustrated in Fig. 4. After fixation, the coverslip is rinsed in D-PBS, blocked twice with TTBS-BSA, probed for 1 hr or longer at 23 ° with the primary antibody (e.g., rabbit anti-T7 RNA polymerase antiserum), and diluted (e.g., 1 : 10,000), with TTBS-BSA. The coverslip is then rinsed with T T B S - B S A and incubated for 1 hr at 23 ° with a fluorochrome-conjugated secondary antibody (e.g., donkey anti-rabbit fluorescein-conjugated antibody), and diluted (e.g., 1 : 25) with TTBS-BSA. The coverslip is then rinsed with T T B S - B S A and stained with Hoechst 33258 to label DNA. An example o f T 7 + NLS accumulation within the nucleus, and of T 7 - N L S exclusion from the nucleus, is shown in Fig. 4. No fluorescence is observed surrounding the nucleus incubated with T 7 - N L S because the nuclei are sedimented through 30% (w/v) sucrose onto a coverslip and away from excluded protein. DNA Replication Low-speed supematants from Xenopus eggs can assemble sperm chromatin into functional nuclei that are able to initiate and complete semiconservative DNA replication under complete cell cycle control] We routinely use three assays for DNA replication as outlined in Fig. 6. 28L. S. Cox, J. CellSci. 101, 43 (1993). 29B. M. Benton, W. Eng, J. J. Dunn, F. W. Studier, R. Sternglanz, and P. A. Fisher, Mol. Cell. Biol. 10, 353 (1990). 30j. j. Dunn, B. Krippl, K. Bernstein, H. Westphal, and E W. Studier, Gene 68, 259 (1988), 31j. Grodbergand J. J. Dunn, J. Bacteriol. 170, 1245 (1988). 32g. Zawadzki and H. J. Gross, Nucleic Acids Res. 19, 1948 (1991).

DNA Replication XSN LSS P,x Mix

+/- Drug

Single Round of Mass of DNA Synthesized

DNA Replication w/In Individual Nuclei ~

Semiconssrvatlve DNA Replication

Terminate reaclion w/ SDS/EDTA buffer

Samples diluted 70-toid wl buffer A Ovedayed on 30% sucrose in buffer A

Terminate reaction wt SDS/EDTA buffer

D~ke~~n'~,e

Degrade protein w l protelnase K (0.5mg/ml) Jr ~ lh

Degrade woteln w/ pmteinase K (0,5mghnl) 37~Cth

~

~

Extract DNA w/ phen ol:chloroform:isoamyl alcohol (25:24:1)

phenei->phenol-chlomform->chlomform

Centrifuge 770xg 5' 23oC

Spot DNA on G F- C flit ers

Nuclei pelleted on coverslips

N p=

TE buffer to 0.Sml 2.5 m1111%+CsCI/TE buffer Refractive Index 1,41

;

Membranes solubittzed w/

1 Precipitate DNA on Washed filters w/ 10% TCA-2% sodium pyrophosphate

Wash filters 3x wl 5% TCA -> 2x w/95% ETOH

0f

1~ eachMap Dry washed filters & count all filters m liquid scin~llant

|

;

Rinsed w/buffer A

~

] -

ndedayaredw/~,

stock CsCI/'rE solution

I~ Texas Red streptevidtn + Hoechst 33258 or Ruorescein sfreptavidin + pmpidium iodide + RNasa A

I I

,00;0, Llqu~@c~a~a~

Time(rain) 0

0

Fraction

FIG. 6. Analysis of DNA replication in Xenopus egg extracts. Density substitution (adapted from Lu et al.36).

650

ENZYMOLOGYAND BIOLOG|CALAPPROACHES

[33]

Incorporation and Detection of Radiolabeled Deoxyribonucleotides The rate and extent of DNA replication in egg extract can be determined by the incorporation of radiolabeled deoxyribonucleotides. The radiolabeled deoxynucleoside triphosphate (dNTP) [ot-32p]dATR is routinely used at 100 #Ci/ml (800 Ci/mmol) to determine the mass of DNA synthesized from a known mass of DNA added to the extract. This requires knowledge of the endogenous dNTP pools in the extract. Using isotope dilution analysis, the dATP 7 and dCTP 15 pools were shown to be approximately 50 #M. All four dNTPs, that is, dATE dCTP, dTTP, and dGTP, are added to the reaction mixture in replication assays to a final concentration of 50 #M to readjust pool sizes after dilution. This not only ensures sufficient dNTP precursors for replication but also allows direct comparison of radiolabeled precursor incorporation between reactions. Sperm nuclei replicate most efficiently when added between 2 and 10 ng of DNA per microliter of extract. 7 Thus, there appears to be an upper and lower threshold of template concentration tolerated by egg extracts. After incubation of the radiolabeled percursor in extract, the reactions are stopped by adding termination buffer [0.5% (w/v) SDS, 20 mM EDTA, 20 mM Tris-HC1 (pH 8.0)]. Proteinase K (0.5 mg/ml) is added and the sample is incubated for 1 hr at 37 °. The DNA is then extracted with phenol, phenol-chloroform, and chloroform and applied to glass fiber filters (GF-C; Whatman, Clifton, N J). A set of two filters is labeled for each sample. Sixty microliters of sample is spotted onto one filter ("washed") and 15/~1 is applied to the other filter ("unwashed"), representing a 4-fold difference in sample volume. The filters are then allowed to dry. Each filter containing the larger volume of sample is then treated with 10% (w/v) trichloroacetic acid (TCA)-2% (w/v) sodium pyrophosphate for 30 min to precipitate the DNA. These filters are then washed three times with 5% (w/v) TCA and then twice in 95% (v/v) ethanol. The washed filters are dried with a heat lamp and then counted, by liquid scintillation, along with the unwashed filters. An average from duplicate filters for each sample is determined. The percentage of [32p]dATP incorporated multiplied by 0.654 yields the mass of DNA synthesized as nanograms per microliter of extract (Fig. 6).33 Titration experiments have shown that daunomycin effectively inhibits replication in the extract, with 50% inhibition occurring at a total drug concentration of 2.7 # M (see Fig. 2 in Leng and Lenol°).

Incorporation and Detection of Deoxynucleotide Analogs Two deoxynucleotide analogs of thymidine, 5-bromodeoxyuridine triphosphate (BrdUTP) and biotinylated deoxyuridine triphosphate (biotin-dUTP), can be

33A. D. Mills, J. J. Blow,J. G. White, W. B. Amos,D. Wilcock,and R. A. Laskey,J. CellSci. 94, 471 (1989).

[33]

USE OF X e n o p u s EGG EXTRACTS

651

used for analysis of DNA replication in the extract. BrdUTP is nearly entirely substituted for thymidine in nascent DNA 34 and biotin-dUTP incorporation has been shown to increase linearly with DNA content in X e n o p u s sperm nuclei replicating in egg extract. 35 Thus, the incorporation of these analogs accurately represents the extent of DNA replication in this system. Two specific features of these analogs, namely density and specificity of detection, can be exploited for the analysis of DNA replication. The dense precursor BrdUTP is combined with [~-32p]dATP, used as a tracer, in density substitution experiments. This procedure allows semiconservative DNA replication to be distinguished from repair synthesis or incomplete strand synthesis and demonstrates whether replication is occurring under cell cycle control, that is, one round per S p h a s e ] DNA (3 ng/#l) is incubated in LSS containing [c¢-32p]dATP at 100 #Ci/ml (800 Ci/mmol) and 0.25 mM BrdUTP for 3 hr. After incubation, the reaction is stopped with termination buffer (see above), proteinase K (0.5 mg/ml) is added, and the sample is incubated for 1 hr at 37 °. The DNA is then extracted with phenol-chloroform-isoamyl alcohol (25 : 24 : 1, v/v/v), brought to a volume of 0.5 ml with TE buffer, mixed with 2.5 ml of 111% CsC1-TE (refractive index, 1.41), and underlaid with 3 ml of stock CsC1-TE solution. Substituted DNA is then separated from unsubstituted DNA by centrifugation to equilibrium, using a 70.1 Ti rotor (Beckman) run at 84,000g for 60 hr at 20 c'. The resultant gradient is fractionated (0.23 ml/min, 200/zl/fraction) and the refractive index of every fifth fraction is determined. Fractions are spotted onto GF/C (Whatman) filters containing 50 # g of herring sperm DNA as carrier and the filters are washed with TCA and counted by liquid scintillation. A typical density substitution profile is shown in Fig. 6. 36 A single peak of radioactivity is detected at a density of 1.75 g/ml (heavy/light DNA, HL), indicating a single round of semiconservative DNA replication and ruling out extensive DNA repair or partial strand synthesis, which would resolve at densities between hemisubstituted (HL) and unsubstituted (light/light, LL) DNA. 37'38 The presence of completely substituted (heavy/heavy, HH) DNA would indicate rereplication within a single incubation, that is, a single cell cycle. Care must be taken when centrifuging high concentrations of cesium chloride as excessive rotor speed, excessive CsCI concentration, overfilled tubes, or low temperature can all cause crystallization and rotor failure. Nascent DNA can be visualized in replicating nuclei through the use of another TTP analog, biotinylated dUTP. Biotin-dUTP incorporated into nascent DNA can be visualized by staining with fluorescent streptavidin. Using flow cytometry 34 R. M. Harland and R. A. Laskey, Cell 21, 761 (1980). 35j. j. Blow and J. V. Watson, EMBO J. 6, 1997 (1987). 36Z. H. Lu, D. B. Sittman, R Romanowski, and G. H. Leno, Mol. Biol. Cell 9, 1163 (1998). 37 H. M. Mahbubani, J. R J. Chong, S. Chevalier, R Thommes, and J. J. Blow, J. Cell Biol. 136, 125 (1997). 38T. Krude, M. Jackman, J. Pines, and R. A. Laskey, Cell 88, 109 (1997).

652

ENZYMOLOGY AND BIOLOGICAL APPROACHES DNA

[33]

Biotin

Daunomyein

(60 rain.) FIG. 7. Daunomycin disrupts the coordinate initiation of DNA replication in Xenopus egg extract. Xenopus sperm nuclei and daunomycin (5 #M) were incubated simultaneously (Fig. 2A) for 60 rain in egg extract containing biotin-dUTP. Nuclei were sedimented onto coverslips, fixed, and stained with Hoechst 33258 (DNA) and Texas Red streptavidin (biotin) to label nascent DNA.

to detect the fluorescent streptavidin-biotin-dUTP complex, Blow and Watson 35 demonstrated that biotin-dUTP incorporation is proportional to DNA synthesis. The biotin-streptavidin system reveals DNA replication within individual nuclei, an approach that has proved extremely useful for analysis of DNA replication in Xenopus extracts. From 20 to 40 # M 5-biotin-16-dUTP is added to the sample, in the Rx Mix, and incubated at 23 ° for 30-120 min. The sample is diluted 70-fold with buffer A, then layered on 30% (w/v) sucrose in buffer A, and the nuclei are sedimented onto a polylysine-coated coverslip by centrifugation at 770g for 5 min at 23 °. The coverslip containing sedimented nuclei is then rinsed in Dulbecco's phosphatebuffered saline (D-PBS) and the nuclei are treated with methanol at - 2 0 ° for 10 min. Nuclei are rinsed with buffer A and stained with fluorescent streptavidin and a total DNA stain such as Hoechst 33258 or propidium iodide. To eliminate any possible nonspecific signal from labeled RNA, RNase A (50 k~g/ml) may be included in the stain mixture when propidium iodide is used. Biotin-labeled nuclei, stained with Texas Red-streptavidin, are shown in Fig. 6. Initiation events occur synchronously or nearly synchronously at the beginning of S phase within individual sperm nuclei replicating in egg extract. 35 Daunomycin has been shown to disrupt the coordination of initiation events, resulting in asynchronous origin firing in egg extract (Fig. 7).1°

Concluding Remarks The interaction of specific drugs with DNA within the cell can affect essential cellular processes, resulting in cytotoxicity. In many cases, the nature of the

[33]

USE OF Xenopus EGG EXTRACTS

653

interaction is known; however, the mechanisms by which drugs induce cytotoxicity are not. Xenopus egg extract provides an excellent cell-free system for the identification of these mechanisms that may provide the basis for the design of novel anticancer drugs. Acknowledgments We thank Drs. Paul Fisher and Miguel Berrios for the gift of bacteriophage T7 RNA polymerase and antiserum, and Dr. Georg Krohne for the gift of monoclonal antibody L046F7. We also appreciate help and advice from Zhi Hong Lu and Jennifer Johns. Our work is supported by the National Science Foundation.

A u t h o r Index

A

Amirikyan, B. R., 170 Amos, W. B., 650 An, H., 433 Anantha, N. V., 138, 141(91), 589 Andersen, A. H., 612, 616, 619, 619(26), 623(14) Anderson, C. E, 9, 20, 147, 195,519 Anderson, J. R., 242, 247(35; 36) Andrew, W. H., 574 Andrews, D. T., 291 Andrews, E. J., 639 Andrews, W. H., 574 Anin, M.-F., 485 Apiou, F., 591 Arakawa, H., 614 Arakawa, S., 154 Arcamone, E, 21,531 Archer, T. K., 504 Arditti, R. R., 469(3), 470 Armitage, B., 338 Armstrong, R. W., 77 Arnott, S., 319 Arrow, A., 314, 315(142), 316(142) Arslan, T., 617,619 Arthanari, H., 138 Asensio, J.-L., 118, 131(33), 256, 260(17), 269, 278,278(17; 69), 279, 280(69; 81), 313, 314, 315(144), 317 Asensio Alvarez, J. L., 347, 351(35), 353(35) Asseline, U., 343 Atherton, E., 558 Atwell, G. J., 388, 394, 556, 557(10; 12) Aubertin, A.-M., 433 Auer, G., 291 Ausi6, J., 503,504, 507(14), 508, 508(6), 509(14), 516(6; 14) Austin, R. H., 18, 106, 416 Avilion, A. A., 573,574, 574(6) Aymami, J., 212,419 Azam, M., 138, 141(91), 589 Azhikina, T., 489 Azhykina, T. L., 489 Azizkhan, J. C., 529

Abildgaard, E, 257 Aboul-Ela, E, 54, 61(10), 612 Abraham, A. T., 619 Abraham, Z., 377 Abu-Daya, A., 224, 225,419, 422, 422(21) Ackers, G. K., 451,453(18-20) Ackroyd, A. J., 554(39), 555 Adams, R. R., 574 Adhya, S., 249 Afshar, M., 22 Agbandje, M., 133, 134(73), 136(73), 586 Aggarwal, A. K., 521 Agrawal, S., 291 Ahsen, U. V., 22(17), 23 Aich, E, 24, 25(41) Aida, M. J., 170 Aisner, D. L., 576, 581 Aivasashvilli, V. A., 483 ,~kerman, B., 73 Aki, T., 249 Akopyants, N., 489 Alaiya, A. A., 291 Alessi, K., 111, 114(18), 115(18) Alexandrushkina, N. I., 486 Allan, S., 576 Allawi, H. T., 170, 171(35), 172(35), 174(35), 175(35), 176, 176(35), 178(35), 329 Allen, E H., 301 Allersma, M. W., 240 Alley, S. C., 404 Allfrey, V. G., 336, 337 Allison, D. P., 235(3), 236, 238,249 Allshire, R. C., 573(12), 574 Allsopp, R. C., 573(13), 574, 576(14) Almouzni, G., 641 Almqvist, N., 250 Altenburger, W., 421 Altona, C., 257 Amaratunga, M., 167, 168(11), 169(11) Am6, J. C., 591 655

656

AUTHOR INDEX

B Baas, P., 589 Babcock, M. S., 306 Babiss, L. E., 335,580 Bacchetti, S., 573,574(6), 576, 577,578, 581(33), 582(33) Baer, L. M., 281 Baguley, B. C., 24(43), 25 Bailey, S. A., 393 Bailly, C., 10(34), 11, 24, 25(35), 26, 29(51), 99, 107(12), 108(12), 347,391,392, 392(92), 393(92; 93), 424, 433,485,486, 489, 489(24), 490, 493,494, 495,495(24), 496, 496(24; 39), 499, 499(41), 500, 500(30), 504, 509(13), 516(13), 517(13), 518(13), 610, 611,612, 614, 620, 620(19), 622, 622(10; 19; 23) Baird, E. E., 291,450, 456, 464(26), 529, 529(7), 530, 556 Baird, W. M., 380, 389 Bajaj, K., 624 Bajic, M., 26, 29(51) Baker, L. J., 230, 233(29) Balagurumoorthy, P., 143 Balasubramanian, B. N., 611,622(11) Baldeyrou, B., 612 Baldridge, K. K., 300, 302(99), 311(99), 318(99), 324(99) Bambirra, S., 29 Bamdad, C., 240, 241(26) Banaigs, B., 612 Bandyopadhyay, B., 578,580(52) Bankman, I. N., 242 Bannwarth, W., 212 Bansal, M., 143 Banville, D. L., 11, 14(40) Baran, N., 590 Barany, G., 581 Barbas, C. F., 594 Barber, J., 212 Bare, L. A., 579, 582(60) Barfield, M., 281 Barker, D. L., 479 Barlow, S., 337 Barner, J., 254 Barnett, L., 377 Barton, J. K., 289 Bass, M. B., 574 Basu, S., 138

Bates, A. D., 51(2), 52 Bates, D. L., 503,507(5) Bates, P. J., 29, 118, 131(31), 291,313,314, 315(142), 316(142) Batey, R. T., 281 Baty, D., 88 Batz, H. G., 338 Baudoin, O., 344, 356(18) Bauer, C. J., 258,270(25) Bauer, G. B., 518 Baugley, B. C., 380 Baur, J., 581 Bax, A., 254 Bayly, C. I., 293,296(31), 302, 304(31), 305(31), 307(31), 308(31), 311(114; 115), 312(31), 318(114; 115), 319(31), 324(114; 115) Beabealashvilli, R. S., 483 Beach, D., 576 Beal, P. A., 132, 424 Beams, J. W., 150 Bear, D. G., 248 Beau, J.-M., 281 Becker, M. M., 100 Beebe, T. P., 240, 241(25) Beekman, J. M., 131 Beerli, R. R., 594 Behrens, C., 329, 333(5), 556 Beijer, B., 489 Beijersbergen, R. L., 576, 577,578 Bell, S. P., 552, 553(36) Bellard, S., 301 Bellorini, M., 529 Belotserkovskii, B. P., 332, 334 Benatar, L., 237 Benchimol, S., 576, 577(36) Benight, A. S., 165, 166, 167, 167(8), 168, 168(8; 11-13), 169(11-13), 170, 170(8; 15), 171,171(1; 8; 15; 26), 172(1; 8; 15; 26; 31), 173(8), 175(8), 176(8), 178(8) Bennett, R. J., 590, 592(110) Benseler, F., 489 Bentin, T., 332, 334, 336(27) Benton, B. M., 648 Beoge, F., 612, 623(14) Berding, C., 329 Berg, D., 489 Berg, H., 531 Berg, R. H., 90, 329, 330(2), 331(1), 332, 333(5), 338(1), 556

AUTHOR INDEX Berger, I., 283, 299 Bergman, J., 24, 25(41) Bergqvist, S., 313 Berkowitz, M. L., 294 Berman, H. M., 111,283,300, 307(104; 105), 311(105), 324(104), 377 Bemardi, G., 419 Bemardou, J., 212 Bernasconi, C. E, 228(27), 229 Berno, A., 166 Bernstein, K., 648 Bernu6s, J., 551 Berova, N., 69 Berrman, T. A., 529 Berry, D. E., 623 Berry, R. S., 153 Betts, L., 580 Beveridge, D. L., 292, 294, 295,296, 296(52; 53), 297(52; 53; 7l), 300, 307(105), 311(105) Bezailla, M., 247 Bhat, T. N., 300, 307(104), 324(104) Bhattacharyya, A., 579 Bhattacharyya, N. P., 578,580(53) Bianchi, A., 584, 585(87) Bianchi, N., 29 Bibby, M. C., 215 Bichenkova, E. V., 22(14), 23 Bigam, C. G., 257 Bigey, P., 338 Bigg, D. C. H., 620, 622 Binaschi, M., 620 Birchall, A. J., 263 Birch-Hirschfeld, E., 29 Bisagni, E., 99, 110, 131,131(7), 343, 345,346, 347, 347(23; 25), 349, 350, 351(25), 354, 354(19), 355,356, 356(19; 45), 424, 504, 509(13), 516(13), 517(13), 518(13), 623 Bischoff, G., 29 Bischoff, R., 29 Bishop, K. D., 165, 171(1), 172(1), 446 Bishop, M., 576 Bisi, J. E., 335,580 Bissery, M. C., 623 Bjornsti, M.-A., 611 Blaber, M., 130 Blackburn, E. H., 137,573, 574, 574(2), 575, 575(15), 576, 578,579 Bladen, S. V., 27 Blake, R. D., 170

657

Blakeney, A. B., 410 Blanchard, S. C., 22(1 l; 13; 33), 23, 24, 446 Blasco, M. A., 574, 576, 577(29) Blencowe, B. J., 489 Bl6cker, H., 170, 174(24) Blommers, M. J. J., 22(25), 23(25; 32), 24, 433 Bloomfield, V. A., 3, 6(7), 55,168,208,282 Blow, J. J., 637(7), 638,641 (7), 648(7), 650, 650(7), 651, 651(7), 652(35) Blume, S., 131 Boatz, J. A., 300, 302(99), 311 (99), 318(99), 324(99) Bocker, W., 582 Bodnar, A. G., 576 Boelens, R., 259 Boesenberg, C., 612, 623(14) Boffa, L. C., 336, 337 Bogdan, F. M., 22 Bogusz, S., 294 Bohley, C., 29 Bohr, V. A., 485,551 Boles, T. C., 51(3), 52 Bolton, P. H., 138, 144, 280, 299, 324 Bonaventura, F. L., 216 Bond, J. P., 195 Bond, P. J., 319 Bonnard, I., 612 Bontemps, N., 612 Bonven, B. J., 616 Bonvin, A. M. J. J., 259 Borer, P. N., 119, 262, 446 Borgias, B. A., 253 Borochov, N., 503,508(6), 516(6) Bosshard, H. R., 114, 115(25) Bostock-Smith, C. E., 216, 297 Boston, R. C., 479 Botstein, D., 291 Bottomley, L. A., 234, 235(3), 236, 242, 243(34), 247, 247(34-36) Bourdouxhe, C., 504, 509(13), 516(13), 517(13), 518(13) Bourne, P. E., 300, 307(104), 324(104) Bowater, R., 54, 61(10), 612 Boyd, F. L., 99, 109 Boyd, J., 263 Boykin, D. W., 4, 22, 22(21; 22), 23(21; 22), 24, 24(5), 25(5; 35), 26, 29, 29(51), 50(58) Brabec, V., 519 Bradbury, E. M., 248 Brahmachari, S. K., 143

658

AUTHORINDEX

Brameld, K. A., 404 Brafia, M., 495,499(41) Branddn, L. J., 337 Brandts, J. E, 115, 121(26), 128,545 Brandts, J. M., 128 Braunlin, W. H., 519 Breaker, R. R., 291 Bremer, R. E., 529(7), 530 Brennan, C. A., 489 Brenner, S., 377 Brenowitz, M., 451,453(18-21) Breslauer, K. J., 18, 24, 24(42), 25, 25(40), 106, 114, 115(23), 117, 127(23), 128, 142(29), 143, 143(29), 146(29), 168, 170, 171, 172(38), 174(24), 175(23; 38), 177(17), 178(38), 346, 352, 352(32) Breslin, D. T., 242,247(36) Bresloff, J. L., 106 Breusegem, S. Y., 212,219, 225(19) Brice, M. D., 301 Bridget, J. M., 645 Briem, H., 590 Briggs, M. R., 552, 553(36) Britt, M., 11, 12(35), 395 Bronckart, Y., 620 Brondani, V., 22(25), 23(25; 32), 24, 433 Brookes, P., 163,402 Brooks, B. R., 292,293,294 Brooks, C. L., 294 Brooks, M. W., 577 Broude, N. E., 337 Brown, B. R., 394 Brown, D. G., 271,274(74), 354 Brown, D. T., 643,646(18), 647(18) Brown, N. C., 489 Brown, P. M., 224, 225,419, 422(21), 504, 510(11), 511(11), 513(11), 514, 515(11) Brown, P. O., 291,624 Brown, T., 256, 259, 260(17), 261,262(37), 264(40), 265(38; 40), 269, 269(40), 278, 278(17; 69), 279, 280(69; 81), 284, 285(11), 295,313,314, 315(144), 317, 347, 351(35), 353(35) Browne, K. A., 247 Brownlee, R. T. C., 535 Broyde, S., 164 Bruccoleri, R. E., 293 Bruce, J., 579 Bruhn, S. L., 519 Bruice, T. C., 247

BriJnger, A. T., 259 Bryan, T. M., 577 Buc, H., 27,469(4), 470 Buchardt, O., 90, 329, 330, 330(2), 331,331(1), 332, 332(14), 333(5; 1 l), 335,336, 336(29), 338(1; 14), 379, 556, 557(11) Buchner, J., 581 Buckin, V. A., 150, 156, 157 Buckle, M., 27 Bujalowski, W., 422 Bukanov, N. O., 336, 337, 337(33) Bukhman, Y. V., 22, 23(9) Bulichov, N. V., 150 Bulyk, M., 607 Burger, H., 582 Burgin, A. B., 34 Burke, T. G., 53l, 532(17) Burli, R. W., 291,450, 529(6), 530 Burn, R, 581 Busby, R. W., 623 Bushman, E D., 626 Bustamante, C., 240, 249, 250 Butler, P. J., 503, 507(5) Buttinelli, M., 502 Byrn, S. R., 380, 389

C Cabrol-Bass, D., 433 Caddle, S. D., 576, 578 Cai, L., 99, 283 Cain, B. F., 388,394 Calderone, D. M., 519 Caldwell, J. W., 293,296(31), 304(31 ), 305(31), 307(31), 308(31), 312(31), 319(31) Calladine, C. R., 504, 506(17) Callo, E J., 170, 171(26), 172(26) Camara, J., 620 Campbel, I. D., 263 Campbell, K., 292 Campbell, K. H. S., 645 Campbell, V. W., 312 Campisi, J., 576 Cantor, C. R., 119, 130, 459 Cantrell, C. E., 384 Capaldi, D. C., 313 Capobianco, M., 21 Capp, M. W., 168 Capranico, G., 616, 620, 620(29)

AUTHOR INDEX Carlson, J. W., 239 Carlson, R. G., 23(34), 24, 390(89), 391 Carpaneto, E. M., 336, 337 Carpenter, M. L., 423 Carpousis, A. J., 471 Carr, R., 278,280(81), 314, 315(144), 317 Carrasco, C., 614 Carrascosa, J. L., 249 Carrondo, M. A., 212,419 Carter, C. W., Jr., 282 Cartright, B. A., 301 Cary, R. B., 248 Casasfinet, J., 27 Cascio, D., 83 Case, D. A., 254, 296, 300, 307(103) Cassidy, S. A., 132, 313,347,349, 351(35), 353, 353(35) Cathers, B. E., 299 Caufield, C., 300, 303(98), 304(98), 307(98), 311 (98), 319(98), 320(98) Cech, T. R., 117, 138,422,574, 590 Celander, D. W., 422 Cera, C., 509, 516(26), 517(27) Chaboteaux, C., 620 Chabra, A., 620 Chaiken, I. M., 48, 49(77) Chaires, J. B., 3, 4, 5, 8, 8(13), 9(25), 11, 11(16), 12(16; 35), 18, 19, 19(45), 37, 50, 65, 99, 100, 100(18; 19), 101,101(10; 11), 102, 106, 107(12), 108(10-12), 109, 111, 114(20), 115(20), 117(20), 123(20), 124(20), 126, 126(22), 135, 135(20; 39), 157,203,204, 204(14), 205,207,207(14), 216, 223(15), 234, 245,246, 291,298, 308, 309, 319, 348,377,380, 395,493,496, 500, 500(30), 504, 506(8), 508, 508(8), 509, 509(8; 23), 516(8), 518(8), 529, 531,532, 532(17), 539,540(24), 546, 549(24; 30), 550(24), 556, 557,557(1; 2), 568,592,614 Chalikian, T. V., 168, 177(17) Champagne, M., 509 Champoux, J. J., 610, 620 Chan, P., 291,336 Chan, S. S., 18, 106 Chan, V., 169 Chandler, C. J., 535 Chandler, S. E, 354 Chang, A. H., 22(18), 23 Chang, E., 574

659

Chang, G., 300, 303(98), 304(98), 307(98), 311(98), 319(98), 320(98) Changeux, J.-P., 16 Chanteloup, L., 281 Chao, M., 408 Chapman, A., 131 Chapman, K. B., 574 Chartier, M., 88 Chastain, M., 33(70), 34, 435 Chattopadhyaya, R., 167, 168(11), 169(11) Chaudhary, D., 312 Chaw, Y. E, 404 Cheatham, T. E., 292, 294, 294(47), 295,296, 296(57-59), 297(57; 59; 61) Chemeris, V. V., 138, 323,590 Chen, A. Y., 386 Chen, D. J., 248 Chen, L., 99, 283 Chen, Q., 23(30), 24 Chen, S.-E, 578 Cheng, J.-W., 226, 253, 277(7) Cheng, X., 486 Cheng, Y.-K., 294(49), 295,298,298(49) Chemy, D. I., 248,249 Cherny, D. Y., 332 Cheung, C. L., 238 Cheung, H. C., 384 Chevalier, S., 651 Chiang, S.-Y., 529 Chiu, C. A., 574, 576 Choi, S.-D., 347 Chollet, A., 489 Chong, J. E J., 651 Choo, Y., 593,594, 595(2; 9), 597,607, 607(1) Chou, S.-H., 226 Chow, C., 22 Chowdary, D., 404 Chowdhry, B. Z., 4, 100(20), 101,109, 111, 114(20), 115(20), 117(20), 123(20), 124(20), 126(22), 128, 130(47), 134, 135(20), 139, 141(94), 142(94), 216, 223(15), 266, 271(59a), 276(59a), 299, 320(97), 324(97), 347,348(37), 586, 589 Chrisey, L. A., 240, 241(28) Christensen, L., 329, 330, 333(5; 11), 334, 334(8), 338(8), 556 Christiansen, K., 612, 616, 619,619(26), 623(14) Chu, T. M., 151

660

AUTHOR INDEX

Chu, W., 390(89), 391 Chui, C.-P., 577 Church, G., 607 Churchill, M. E. A., 422,424, 513 Cieplak, P., 293,295,296(31; 58), 302, 304(31), 305(31), 307(31), 308(31), 311(114; 115), 312(31), 318(114; 115), 319(31), 324(114; 115), 392 Clairac, R. L. d., 291 Clairac, R. P. d., 291 Clark, G. R., 222, 230, 233(29) Clarke, P. A., 579 Clegg, R. M., 4, 5(21), 212, 213(11), 216(10), 217,219, 225(19), 228(28), 229 Clegg, W., 301 Cleland, A. N., 251 Clement, R. M., 156 Cleveland, J. P., 237, 239, 240 Clever, J. L., 433,443(8) Cliff, C. L., 590 Clore, G. M., 262 Coffin, J., 433 Cohen, G., 546 Cole, M. D., 312 Coil, M., 212, 419 Collier, D. A., 131,133,340, 343, 351(15) Colonna, E P., 21 Colson, P., 24, 433,504, 509(13), 516(13), 517(13), 518(13), 612, 614, 620(19), 622(19; 23) Combaut, G., 612 Comeau, L., 584, 585(87) Condom, R., 433 Conn, G. L., 22, 23(10) Connelly, P. R., 128 Conney, A. H., 163 Cons, B. M. G., 423,515 Conte, M. R., 258,270(25), 271, 274(76) Conway, B. E., 151 Cook, H. J., 573(13), 574, 576(13) Cook, P. D., 433 Coombs, D., 30, 48(67) Cooper, M. A., 29 Corbo, M., 579 Corda, Y., 485 Cordell, J., 371 Corey, D. R., 334, 336(26), 578,580, 580(54; 56)

Cornell, W. D., 293,296(31), 302, 304(31), 305(31), 307(31), 308(31), 311(114; 115), 312(31), 318(114; 115), 319(31), 324(114; 115) Comilescu, G., 254 Correia, J. J., 8, 9(25), 37,245 Cosman, M., 163, 164 Cotter, E, 579 Cotton, E A., 421 Coull, J. M., 330, 333(11), 338 Coulomb, H., 622 Counter, C. M., 573,574, 574(6), 576, 577, 578 Courey, A. J., 552 Coury, J. E., 242, 243(34), 247,247(34-36) Covey, J., 388 Coviello, G. M., 137,576, 581(32), 582(32) Cowell, J. K., 578,580(52) Cox, L. S., 643,645,645(16), 648 Cox, R., 645 Cozzarelli, N. R., 51(3), 52, 386 Craigie, R., 626(7), 627 Crain, P. F., 403 Cramer, H., 578, 580(52) Crane, L. E., 404 Crane-Robinson, C., 168 Crenshaw, J. M., 380 Crescenzi, V., 157 Crick, E H. C., 377 Crooke, S. T., 25,291 Crothers, D. M., 3, 6, 6(7), 8(23), 11(23), 16, 18(43), 51, 54, 55, 62(13), 64(14), 65, 65(14), 99, 100, 102, 106, 194, 262, 389, 466, 467,468,471,479, 494, 504, 506(8), 508(8), 509, 509(8), 515(24), 516(8), 518(8), 546, 550 Crouch, R. J., 30 Crow, S., 499, 500 Crowley, M. E, 295,296, 296(59), 297(59) Cubberley, M. S., 556 Cubero, E., 298 Cui, M., 22(23), 23(23), 24, 25(23), 433 Cui, Q., 292(26), 293 Cullinane, C., 408,466, 470, 471,473,476(7), 477,478,479, 481,483,483(7), 484, 485, 550, 551 Cundliffe, E., 109 Cunningham, D., 579 Curto, E. B., 264 Cutts, S. M., 408,466, 473,479, 481,550

AUTHOR INDEX Czamik, A. W., 22(23; 24), 23(23; 24), 24, 25(23), 433

D Dabrowiak, J. C., 10(33), 11,100(19), 101,418, 429, 431,432, 436, 446, 446(3), 473 da Costa, P. M., 620 Daggett, V., 293 Dahl, O., 334, 335(28), 339 Dahlquist, K. D., 22(13), 23, 23(33), 24, 446 Dale, R., 314, 315(142), 316(142) D'Alessio, G., 435 Dalke, A., 306 Dalla-Pozza, L., 577 Dalma-Weishausz, D., 451,453(21) Danielson, U. H., 26, 49(52) Danyluk, S. S., 260 Daragan, V. A., 262 Darden, T. A., 294, 294(47), 295 Das, A., 500 Das, G. C., 509 Dasso, M. C., 641,644 Dassonneville, L., 24, 433,612, 614, 620, 622,622(23) Dattagupta, N., 16, 18(43), 65, 102,504, 506(8), 508(8), 509, 509(8), 515(24), 516(8), 518(8) Daune, M. P., 156, 509 D'Aurora, V., 519 Dauter, Z., 139, 324 David-Cordonnier, M. H., 489 Davidoff, M. J., 578 Davidson, K. K., 138,589 Davies, B. A., 580 Davies, D. B., 260 Davies, D. R., 130, 301 Davies, J., 22(17), 23 Davies, R. T., 589 Davin, D., 312 Davis, D. R., 29, 255 Davis, J., 130 Davis, P. W., 118 Davis, T. M., 22, 42, 43(76) Dawid, I. B., 640 D'Costa, N. P., 242 Dean, A. C. R., 377,381 Dean, C. J., 371 Dean, D. A., 336, 479

661

Debbs, R., 203 Debizemont, T., 355 Decaestecker, C., 620 Decker, P., 591 Decout, J.-L., 130, 340 Decoville, M., 485 Dedon, P. C., 4, 518 Degrooth, B. G., 239 Deiters, A., 421 de Jong, R., 258,260(27) Delage, S., 350 Delaglio, F., 254 Delaine, E., 240, 248 de Lange, T., 573,576, 581(31), 584, 585(87), 591 Delbarre, A., 383,556, 557(I 3) Delben, F., 157 Delcourt, S. G., 170 De los Santos, C., 164 delos Santos, C., 315 Demaret, J., 297 Demarquay, D., 620, 622 Demeny, T., 300, 307(105), 311(105) Demidov, V. V., 332, 333,334, 336, 337, 337(33), 338,339(20) Demin, V. V., 249 De Mol, N. J., 397 Dempster, M., 573(12), 574 de Murcia, G., 509, 591 Denissenko, M. F., 359 Denny, W. A., 380, 388,394, 556, 557(10; 12) DePinho, R. A., 576, 577(29) Deprew, D. E., 54 DeRose, J. A., 240 Dervan, P. B., 100, 130, 132, 278,291,340, 356(2), 419, 424, 429, 450, 451,452(13; 15-17; 24; 25), 453,456, 458, 462(17), 464(26), 519, 529, 529(6; 7), 530, 556 Descounts, P., 241 Desnoyers, J. E., 151 Devlin, J. J., 579, 582(60) de Vroom, E., 258,260(27) D'Halluin, J. C., 489 Dhesi, J., 313 Dhingra, M. M., 311 Diallo, R., 582 Diaz, M. O., 577 diCera, E., 3, 6(5) Dickerson, R. E., 83, 212, 226, 306, 496 Dickinson, L. A., 450, 529

662

AUTHOR INDEX

Diekmann, S., 212, 213( 11), 494, 499(33) Dignam, J. D., 552 Dimitrov, S., 641,644 D'Incalci, M., 529 Ding, D., 4, 26, 29, 29(51), 50(58) Dingley, A. J., 281 Dingwall, C., 421 di Stefano, F., 579 Dobrowiak, J. C., 393 Dobrynin, V. N., 131 Dockhorn-Dwomiczak, B., 582 Doktycz, M. J., 170, 171(26), 172(26), 249 Dombroski, B. A., 399, 422,519 Dominey, R. N., 291 Dominy, B. N., 294 Donahue, B. A., 485 Dongchul, S., 493,500(30) Donner, P. O., 312 Domer, L. E, 521 Dorr, A., 579 Dosanjh, H. S., 29, 118, 131(31; 33), 141,299 Doubleday, A., 301 Doudna, J. A., 22(26), 23(26), 24 Douglas, K. T., 22(14; 15), 23,212,214, 215, 222(12), 254 Douthwaite, S., 24 Dower, W. J., 600, 608(15) Dragan, A. I., 168 Dragowska, W., 577 Draper, D. E., 22, 23(9; 10), 156, 435 Draper, J. E., 24 Dreier, B., 594 Dremaldi, K. A., 252 Drenth, J., 282,284(2) Drepaul, R. S., 641 Drew, H. R., 413,415(2), 418,419(13), 421, 421(13), 429(13), 503,504, 506(17), 510(7), 511(7; 9), 514(9), 515(9) Dritschilo, A., 248 Driver, D. A., 329, 333(5), 556 Drobny, G. P., 258 Drohat, A. C., 284, 285(13) Droz, E., 241 Drukier, A. K., 337 Drushlyak, K. N., 131 Duan, Y., 296 Dubelman, S., 403 Dudycz, L. W., 489 Dueholm, K., 330, 333(11) Duff, R. J., 22(16), 23

Dumortier, L., 4, 5(21), 212, 216(10), 217 Dunham, M. A., 577 Dunlop, M. G., 573(12), 574 Dunn, J. J., 648 Dunphy, W. G., 645 Dupuis, M., 300, 302(99), 311 (99), 318(99), 324(99) Durand, M., 346 Durchschlag, H., 155 Durland, R. H., 130 Durselen, R., 239 Duval-Valentin, G., 99, 110, 131(7), 355 Duvic, M., 130 Dy, M., 581 Dynan, W. S., 248 Dyson, H. D., 257 Dziewanowska, Z., 579 Dziowgo, C., 157

E Eastman, A., 404 Eastwood, J. W., 294 Eaton, E. N., 574, 577,578 Ebbinghaus, S., 291 Eblings, V., 237 Ebrahimi, S. E. S., 22(15), 23,214, 215,222(12) Ecker, D. J., 25 Eckhardt, G., 579 Eckstein, D. A., 433,443(8) Edsall, J. T., 3 Edwards, K. J., 271 Efremova, E. J., 489 Egan, W. M., 257 Egholm, M., 90, 329, 330, 330(2), 331, 331(1), 332, 332(14), 333(5; 11), 335, 336, 336(29), 338,338(1; 14), 556 Egly, J. M., 551 Ehrlich, K., 486 Ehrlich, M., 486 Eisen, M. B., 291 Eisenberg, H., 503,508(6), 516(6), 546 Elber, R., 306 Elbert, S. T., 300, 302(99), 311(99), 318(99), 324(99) Eldrup, A. B., 339 Elgin, S. C. R., 495 Elliott, W. H., 381 Ellis, T. M., 577

AUTHORINDEX Ellisen, L. W., 578 Elrod-Erickson, M., 599 Elvingson, C., 73, 88 Emini, E. A., 633 Eng, W., 648 Engelke, J., 262 Engelman, A., 626(7), 627 Englezou, A., 577 Erard, M., 509 Erickson, J. W., 27 Erickson, L. C., 485 Eriksson, M., 68, 90, 92,516 Erkkila, K. E., 289 Escud6, C., 340, 345, 349, 350, 354, 354(19), 356(19; 45) Essigmann, J. M., 485, 551 Essmann, U., 294

F Faaland, C. A., 354 Fairall, L., 585 Faldesz, B. D., 165,171(1), 172(1) Fan, J. B., 166 Fan, P., 219 Fan, Y., 610 Fang, G., 590 Fang, Y., 242 Faruqi, A. F., 313, 336 Fasman, G. D., 12, 508,516, 517(39) Faucon, B., 99, 110, 131(7) Faunt, L. M., 8 Fawthrop, S. A., 266 Fede, A., 212 Fedoroff, O. Y., 138, 323,580, 586, 590 Feig, M., 296 Feigon, J., 110, 139, 142(14), 144, 253,278, 278(5), 281,314, 315(143), 317 Feinberg, A. M., 403 Feigner, P. L., 203,337 Felsenfeld, G., 130, 389 Feng, J., 574 Feng, Z., 300, 307(104), 324(104) Ferguson, D. M., 293,296(31), 304(31), 305(31), 307(31), 308(31), 312(31), 319(31) Feriotto, G., 29 Fernandez, C., 281 Fernandez-Saiz, M., 22, 24(5), 25(5)

663

Ferrara, J. D., 144 Ferre-D'Amare, A. R., 22(26), 23(26), 24 Ferrell, T. L., 235(3), 236, 238 Ferrer, N., 551,554(33), 555(33) Fiala, R., 23(31), 24 Filippov, S. A., 131 Filipski, J., 388 Filoche, M., 297 Finch, J. T., 216, 503,508(4), 516(4) Finkel, S. E., 494 Firth, W. J., 382(53), 383,394 Fisher, C., 27 Fisher, J., 266 Fisher, P. A., 648 Fisher, R. J., 27 Fivash, M., 27 Flanagan, J. M., 280 Flashman, E., 418,419(17), 424(17) Florsheimer, A., 22(25), 23(25; 32), 24, 433 Fokt, I., 19,291,308,529,532, 568 Footer, M., 338 Ford, C. C., 641,645 Ford, K. G., 324 Forrow, S. A., 485 Forsyth, G., 345 Foss6, P., 623 Fourcade, A., 248 Fourmy, D., 22(11-13), 23, 23(29; 33), 24, 446 Fox, K. R., 11, 12(35), 110, 117(12), 125, 130(12), 131,131(12), 132, 132(12; 67), 215,224, 225,271,274(78; 79), 291,298, 339, 347,349, 350, 35 I(35), 353,353(35), 354, 391,391(91), 412, 413,414, 415,417, 418,419, 419(17), 421,422, 422(21), 423, 424, 424(17), 426, 427,429(3), 432,452, 504, 510, 510(11), 511(11), 513(11), 514, 515,515(11), 555,556 Fox, T., 293,294(47), 295,296(31; 59), 297(59), 304(31), 305(31), 307(31), 308(31), 312(31), 319(31) Francisco, C., 612 Francois, J. C., 343,556 Frank, R., 170, 174(24) Frankel, A. D., 291 Frank-Kamenetskii, M. D., 110, 117(9), 130, 131,131(9), 170, 176, 332, 333,334, 336, 337,337(33), 338,339(20), 341 Franzen, B., 291 Frau, S., 212 Frazier, J., 301

664

AUTHOR INDEX

Frederick, C. A., 86, 164, 212 Frederick, N. A., 238 Freeman, R., 255 Freier, S. M., 118, 329, 333,333(5), 336, 556 Freire, E., 114, 115(23), 127(23) Frenkiel, T. A., 266, 273,275(64) Fresco, J. R., 170 Freudenreich, C. H., 388 Friedman, J., 388 Friedman, R. A., 9, 10(29; 30) Fritsch, E. E, 363,435,600, 608(14) Fritz, M., 240 Fritzsche, H., 509, 531 Fritzsche, W., 238 Froelich-Ammon, S. J., 385 Frolkis, M., 576 Froussios, K., 563 Frydenlund, H., 338 Fuchs, R. E, 485 Fujimori, A., 610, 620 Fukanaga, M., 384 Fukunaga, M., 379 Fukusawa, K., 614, 622(23) Fulton, T. B., 579 Funk, W. D., 574 Furlong, C., 578 Futcher, A. B., 573,573(13), 574, 576(5; 14) Futscher, B. W., 485

G Gabbay, E. J., 504, 508(12), 517(12) Gaffney, B. L., 117, 142(29), 143, 143(29), 146(29), 256 Gago, E, 557 Galan, A. A., 22(24), 23(24), 24 Galas, D. J., 412, 451 Gale, E. E, 109 Gall, A. A., 240, 241,241(24) Gallego, J., 570 Gallie, D. R., 250 Gallo, E J., 165, 166, 167(8), 168(8), 170(8), 171(1; 8), 172(1; 8), 173(8), 175(8), 176(8), 178(8) Gallo, M. A., 354 Galluci, J. C., 421 Gambari, R., 29 Gamper, H., 485 Gandhi, V., 131

Gao, Q., 287 Gao, X., 203,315 Gao, Y.-G., 283,408 Garbay-Jaureguiberry, C., 287 Garbesi, A., 21 Garcia, R. G., 489 Garcia, R. J., 249 Garcia de la Torre, J., 263 Garestier, T., 131,340, 341,344, 345,346, 347, 347(23; 25), 349,350, 351(25), 354, 354(19), 356, 356(18; 19; 45), 424 Garland, E, 384 Gaub, H. E., 251 Gaudiano, G., 408 Gaudin, E, 281 Gavazzo, P., 516 Gazdar, A., 581 Geacintov, N. E., 24(42), 25, 163, 164, 352 Gee, J. E., 131 Geierstanger, B. H., 291 Geiger, A., 329 Gelasco, A., 164 Gelbin, A., 111,300, 307(105), 311(105) Gellert, M., 389 Gelus, N., 24, 25(35), 433 Gessner, R. V., 519 Ghandour, G., 166 Ghanouni, E, 485 Ghirlando, R., 626(7), 627 Giancotti, V., 157 Gibbs, S. J., 263,269(53) Giberson, R., 239 Gibson, I., 485 Gibson, Q. H., 214 Gibson, T. J., 608 Gieselmann, R., 240 Giesen, U., 329 Giessner-Prette, C., 254 Gilbert, D. E., 110, 142(14) Gilbert, E L., 395 Gilbert, W., 361,416, 458 Gill, S. J., 3, 6(3) Gilley, D., 579 Gillies, G. T., 150, 153(8) Gilliland, G. L., 284, 285(13), 300, 307(104), 324(104) Gimenez-Arnau, E., 297 Giovannangeli, C., 131,340, 343,354 Giriat, I., 591 Gisself~ilt, K., 73

AUTHOR INDEX Glazer, P. M., 291,313,336 Glisson, B., 386 Gluick, T. C., 24, 435 Glukhov, A. I., 578, 580(55) Gmeiner, W. H., 23(28), 24 Gochin, M., 253 Gocke, E., 616 Godfrey, B. J., 239 Gold, L., 291 Goldberg, I. H., 377,518 Goldstein, B., 30, 48(67) Goldstein, R. E, 168, 170, 170(15), 171, 171(15; 26), 172(15; 26) Goljer, I., 280 Gollahon, L., 581 Gomperts, M., 641 Gonzalez, C., 259 Goodisman, J., 10(33), 11,393, 418, 429, 431, 432, 436, 446, 446(3) Goodman, S. R., 336 Goodsell, D. S., 83,496 Gopalakrishnan, S., 500 Gordeev, S. A., 578,580(55) Gordon, M. S., 300, 302(99), 311 (99), 318(99), 324(99) Gordover, L., 620 Gorenstein, D. G., 253,264 Gorenstein, J., 337 Gotfredgen, C. H., 314, 315(143), 317 Gotoh, O., 170 Gottesfeld, J. M., 450, 529 Gottlieb, G. J., 576 Gould, I. R., 293,296(31), 304(31), 305(31), 307(31), 308(31), 312(31), 319(31) Gowan, S. M., 141,299, 589 Gowers, D., 418, 419(17), 424(17), 556 Graham, D. R., 519 Gralla, J. D., 291, 471 Graslund, A., 24, 25(41) Graves, D. E., 377, 379, 379(32), 380, 382(22; 53; 100), 383,383(29), 384, 384(29), 385, 385(32; 33), 386(22; 38), 387,388(74), 391,392, 392(92), 393,393(92; 93), 394, 395,531,532(17), 614 Graves, D. J., 169 Gray, E. J., 222 Gray, J. C., 29 Gray, P. J., 473 Green, D. K., 573(12; 13), 574, 576(13) Green, M. R., 22(20; 22), 23(20; 22), 24, 433

665

Greenberg, W. A., 26, 29(48; 49), 36(48) Greider, C. W., 137, 573,573(13), 574, 574(6), 575(15), 576, 576(5; 14), 577(29) Greig, M. J., 333 Greisman, H. A., 594, 607(7) Grek, S., 130 Greve, J., 239 Griffey, R. H., 25,333 Griffieth, M. C., 336 Griffith, J. D., 584, 585(87) Griffith, M. C., 333 Grigg, G. W., 426 Grigoriev, M., 331 Grimaldi, K. A., 358,358(5; 6), 359, 364(3-6), 375(3~5) Grodberg, J., 648 Gromann, U., 29 Gromova, I. I., 619 Grondard, L., 623 Gronenborn, A. M., 262 Gross, H. J., 648 Grunewald, U., 239 Gryaznov, S., 493 Grzesiek, S. J., 281 Grzeskowiak, K., 212, 226 Grzeskowial, K., 83 Gu, M., 238 Guarcello, V., 314, 315(142), 316(142) Guedj, R., 433 Gueguen, G., 313 Guelev, V. M., 556, 558,559, 565(24), 567(20; 24), 568,570(20; 29) Gueron, M., 263,297 Guida, W. C., 300, 303(98), 304(98), 307(98), 311(98), 319(98), 320(98) Guieysse, A. L., 110, 331,341 Gukovsky, V. P., 150 Gulizia, R. J., 450, 529 Gumport, R. I., 489 Gunderson, S. I., 551 Gunnell, S., 130 Guntherodt, H. J., 238,240 Guo, Q., 138, 143 Gupta, G., 311 Gupta, J., 577 Gupta, M., 610, 619 Gurley, G., 237 Gurskii, G. V., 194 Gutell, R. R., 22, 23(10) Guthold, M., 250

666

AUTHOR INDEX

H Haaima, G., 334 Haber, D. A., 578 Habhoub, N., 130, 340 Haefke, H., 238 Hagerman, P. J., 499 Hagmar, E, 88 Hahn, W. C., 576, 577 Hakkinen, A., 262 Halford, S. E., 554(39), 555 Halim, N. S., 22(24), 23(24), 24 Hall, S. W., 388 Haly, B. D., 433 Hamalained, M., 26, 49(52) Hamana, K., 495 Hamasaki, K., 25 Hamdane, M., 489 Hamilton, S. E., 578, 580, 580(56) Hampel, K. J., 131,345 Hamy, E, 22(25), 23(25; 32), 24, 25(35), 433,614 Han, E X., 138, 141(87), 586, 589 Han, H., 138, 141(87), 278,323,586, 589, 590, 592, 592(110) Han, W. H., 237 Han, Z., 301 Hanawalt, P. C., 485 Hande, M. E, 576, 577(29) Hanlon, S., 167, 168(11), 169( 11) Hannon, G. J., 576 Hansch, C., 394 Hansen, H. E, 334 Hansen, J., 556, 557(11) Hansen, M. S. T., 626 Hansma, H. G., 240, 247,248, 251 Hansma, P. K., 237,238,239, 240, 250, 251 Hanvey, J. C., 131,335,340, 580 Haq, 1,4, 100(20), 101,109, 111,114(19; 20), 115(19; 20), 117(20), 123(20), 124(20), 126(22), 134, 135(20), 139, 141(94), 142(94), 216, 223(15), 266, 271(59a), 276(59a), 299, 320(97), 324(97), 347, 348(37), 586, 589 Harada, K., 291 H~ird, T., 76, 79, 81(13), 219 Hardenbol, E, 303 Hardin, C. C., 117

Hardwick, J. M., 379(31), 380 Harel-Bellan, A., 331 Harland, R. M., 651 Harley, C. B., 137,573,573(13), 574, 574(6), 576, 576(5; 14), 577,581,581(32), 582(32) Harnett, J., 620 Harper, L., 620 Harrington, L., 574 Harrington, R. E., 240, 241 (24) Harris, P. J., 410 Harrison, J. R., 299 Hartl, F. U., 581 Hartley, J. A., 252, 358,358(5; 6), 359, 364(3-6), 375(3-6), 485 Hanson, S. D., 581 Haruki, M., 30 Harvey, S. C., 294, 311 Hassman, C. E, 335,580 Hastie, N. D., 573(12), 574 Hastings, J. C., 633 Hawkins, C. A., 291 Hawley, R. C., 294 Hawn, D. D., 24 l Hayakawa, K., 203 Hayes, J. J., 424, 513 Haynie, D. T., 130 Hazen, E. E., 421 Hazuda, D. J., 633 He, S. M., 106 He, X., 30, 48(67) Hearst, J. E., 150, 485 Hecht, S. M., 22(16), 23,617, 619, 619(30; 31; 35; 36), 623 Hegner, M., 240, 241(27) Heim, G., 248 Heldsinger, A., 22(24), 23(24), 24, 433 H61bne, C., 99, 110, 130, 131,131(7), 137,331, 340, 341,343,344, 345,346, 347,347(23; 25), 349, 350, 351(15; 25), 354, 354(19), 355, 356, 356(18; 19; 45), 392, 424, 504, 509(13), 516(13), 517(13), 518(13), 578 Henderson, E., 117, 238, 239 Henderson, J. E, 167, 168(11), 169(11 ), 381 Hendricksen, T., 294, 300, 303(98), 304(98), 307(98), 311(98), 319(98), 320(98) Hendrix, M., 26, 29(48; 49; 50), 31(50), 36(48), 38(50), 433 H6nichart, J. E, 504, 509(13), 516(13), 517(13), 518(13)

AUTHOR INDEX Henningfeld, K. A., 617,619(35; 36) Henry, D. W., 535 Henry, R. J., 410 Hermann, T., 22, 22(19), 23(19), 24 Hernandez, B., 292(27), 293 Herne, T. M., 29 Herrera, J. E., 11, 12(35), 18, 106, 500, 531 Hertzberg, R. P., 424, 452(24; 25), 453, 519 Heus, H. A., 258,260(27) Hicks, M., 519,520, 521(12), 527(12), 528(12) Higgs, H., 301 Hi|bers, C. W., 257,258,260(27) Hill, H. D. W., 255 Hilsenbeck, S. G., 586 Hingerty, B. E., 164 Hinshelwood, C. N., 377 Hiramoto, R. A., 312 Hirose, Y., 138 Hirshberg, M., 293 Hixon, S. C., 379, 384 Hiyama, E., 581 Hiyama, K., 581 Ho, E, 500 Ho, R L. C., 137,576, 581(32), 582(32) Ho, E S., 100 Hochleitner, K., 359 Hockney, R. W., 294 Hoff, E. V., 156, 157(26) Hoffman, D., 568,570(29) Hoffman, E. E, 337 Hoffman, S., 29 Hofstadler, S. A., 25 Hogan, M. E., 16, 18, 18(43), 65, 106, 130, 131, 416, 509, 515(24) Hoger, T., 591 Hoh, J. H., 242 Hohiesel, J. D., 489 Hoiland, H., 155 Hol, W. G. J., 610 Holbrook, J. A., 168 Holman, T., 645 Holmes, C. E., 22(16), 23 Holmes, R. R., 421 Holt, S. E., 575,576, 581 Holtzman, D. A., 552 Hommel, U., 263

667

Honda, K.-J., 170, 175(36), 176(36), 178(36) Honig, B., 300, 307 Hopkins, P. B., 396, 398,400, 401(7; 8), 402, 404, 408, 519 Homer, J. W., 576 Horz, W., 421 Hotchkiss, R. D., 486 Houssier, C., 24, 433,504, 509(13), 516(13), 517(13), 518(13), 612, 614, 620(19), 622(19; 23) Howard, E B., 301,493 Howell, D. K., 167, 168(12), 169(12) Howland, R., 237 Hoyos, P., 380, 389 Hoyt, P. R., 249 Hsie, L., 166 Hsieh, S. H., 300, 307(105), 311(105) Hu, G., 546, 549(30), 568 Hu, G. G., 284, 285(12) Hu, J., 238 Hu, S., 271 Huang, C. Y., 4 Huang, H., 398 Huang, L., 203,337 Huang, P., 131 Huang, R. C., 508,509(19) Huang, W., 581 Huang, Y. Q., 446 Huang, Z., 238 Huber, P. W., 22(23), 23(23), 24, 25(23) Huchet, M., 622 Huchital, D. H., 519, 520, 521(12), 527(12), 528(12) Huckriede, B. D., 257 Hudson, B. P., 289 Hughes, T. R., 574 Hui, C., 338 Huizenga, D. E., 291 Hummelink, T., 301 Hummerlink-Peters, B. G., 301 Humphrey, W., 306 Hunenberger, P. H., 294 Hunter, W. N., 284, 285(11) Hurley, L. H., 99, 109, 138, 141(87), 323,556, 557(2), 558,567(20), 570(20), 573,582, 583,586, 589, 589(85), 590, 592, 592(110) Hurst, R., 581 Hutchison, C. J., 641,645 Hyrup, B., 329

668

AUTHOR INDEX

lbanez, V., 164 Ikeda, N., 138 Ikuta, S., 167, 168(11), 169(11) Inada, M., 281 lng, N. H., 131 lngacio Tinoco, J., 3, 6(7) Inman, E. M., 592 Ippel, J. H., 257,258, 260(27) Iribarren, A. M., 313,489 lsalan, M. D., 593,594, 595(2; 9), 607(1) Ishihara, T., 334, 336(26) Ishikawa, E, 574 lskandar, M., 504, 507(14), 509(14), 516(14) Itsuki, H., 154 lverson, B. L., 458, 556, 558,567(20), 568, 570(20; 29) Izbicka, E., 138,586, 589 lzvolsky, K. I., 336, 337,337(33)

,J Jackman, M., 65 l Jackson, S. E, 552 Jacobs, B. L., 240, 241(24) Jacobson, E. L., 585,591(88) Jacobson, M. K., 585,591(88) Jacobson, S., 518 Jagadeesh, J., 284, 285(13) Jain, S. C., 389 Jakob, F., 612,623(14) Jakob, U., 581 James, R., 312, 355 James, T. L., 253,257, 258, 259, 259(26), 262, 264(26), 265(26), 270, 270(26), 271, 277(26) Jamison, E., 451,453(21) Janin, Y., 345,354(19), 356(19) Janmey, E, 240 Janota, V., 181, 185 Janssen, L. H., 397 Jares-Erijman, E. A., 159 Jaxel, C., 616, 620, 620(28; 29) Jayaram, B., 294, 295,296(52), 297(52) Jayaraman, K., 131 Jelesarov, 1., 114, 115(25), 168 Jen-Jacobson, L., 223 Jenkins, H., 645

Jenkins, T. C., 4, 29, 99, 100(20), 101,109, 110, 111,113(21), 114(20), 115(20), 117(13; 20), 118, 118(21), 123(20), 124(20; 21), 126, 126(22), 130(13), 131,131(13; 31; 33; 40), 132(13; 67), 133, 133(13), 134, 134(73), 135, 135(20; 39), 136(73-75), 137(13; 16), 139, 141,141(94), 142(13; 94), 157,216, 223(15), 261, 262(37), 264(40), 265(38; 40), 266, 269(40), 271,271 (59a), 273,274(74-76; 79), 275(64), 276(59a; 75), 295,298, 299, 313,320(97), 324(97), 347, 348(37), 350, 485,583,586, 589,589(85) Jenkins, T. M., 626(7), 627 Jennewein, S., 490, 493(26) Jensen, J. J., 300, 302(99), 311 (99), 318(99), 324(99) Jensen, K. K., 329 Jeppesen, C., 427 Jezewska, M. J., 422 Jia, X., 580 Jiang, L., 23(31 ), 24 Jin, R., 117, 142(29), 143, 143(29), 146(29) Jing, T. W., 237 Job, D., 485 Job, E, 4 Johnson, C., 371 Johnson C. S., Jr., 263,269(53) Johnson D. K., 249 Johnson M. L., 8 Johnson R. C., 494 Johnson R. K., 617,619(30; 31) Johnson W. C., 100 Johnston, R. E, 479 Jones, D. E., 291, 312 Jones, R. A., 117, 142(29), 143, 143(29), 146(29), 256 Jones, R. J., 464 Jones, R. L., 11, 14(40), 377 Jonsson, M., 73 Jordan, S. R., 580 Joselevich, E., 238 Josey, J. A., 580 Jovin, T. M., 159, 239,248,489 Jovin, T. E, 228(28), 229 Joy, D. C., 238 Joyce, G. E, 26, 29(50), 31 (50), 38(50), 433 Jucker, F. M., 281 Jujawinski, E., 281 Jung, M., 248

AUTHOR INDEX

K Kadonaga, J. T., 552, 553(36) Kafri, T., 626 Kahlon, J. B., 291 Kahn, N. N., 489 Kainosho, M., 262,281 Kakefuda, T., 54 Kallansrud, G., 119 Kallenbach, K. R., 138 Kallenbach, N. R., 143 Kam, L., 422 Kamitori, S., 390(88), 391,495 Kandimalla, E. R., 291 Kang, C. H., 283 Kankia, B. 1., 149, 156, 159, 165 Kankiya, B. I., 150, 157 Kaptein, R., 259 Karplus, M., 292(26), 293 Kasas, S., 250 Kashin, I., 165 Kashlev, M., 250 Kasprzyk, G., 620 Kasprzyk, P. G., 620 Kastrup, R. V., 385 Katahira, M., 23(27), 24 Kataoka, T., 581 Kathiriya, I. S., 578,580(56) Katipally, R. R., 580 Kawano, T, L., 138 Kawashima, E., 489 Kay, L. E., 262, 281(44) Keams, D. R., 219 Kedersha, N. L., 592 Kehr, A., 612, 623(14) Keiderling, T. A., 181 Kelland, L. R., 141,299, 578,586, 589, 589(92) Kelland, S. M., 299 Keller, D. J., 240 Keller, W., 54 Kelly, K., 312 Kempf, A., 647 Keniry, M. A., 299 Kennard, O., 301 Kennedy, A. W., 578,580(52) Kennedy, M. A., 258, 280 Keppler, M. D., 131,132(67), 298, 354, 418 Kemen, P., 240, 241(27) Kerper, P. S., 249

669

Kerridge, D., 381 Kerrigan, D., 388 Kersten, H., 377 Kersten, W., 377 Kerwin, S. M., 138,299, 323,583,586, 589(85), 590 Kerwood, D. J., 253 Kessler, D. J., 18, 130, 131 Kessler, O. J., 106 Kharakoz, D. R, 154 Khil, P. P., 249 Khudyakov, I. Y., 486 Kickhoefer, V. A., 592 Kielkopf, C. L., 289,450, 529(7), 530 Kiely, J. S., 333 Kill, I., 645 Kilmkait, T., 433 Kim, H. G., 291,312, 355 Kim, J.-H., 332 Kim, K., 248 Kim, K.-H., 332 Kim, M.-S., 347 Kim, N. W., 137,576, 577,581,581(32), 582, 582(32) Kim, S.-G., 265,270(59) Kim, S. K., 83, 90, 329,333(5), 346, 347, 556 Kimura, K., 483 Kingsbury, W. D., 617,619(30; 31) Kingston, R. E., 503,518(2) Kirchner, J. J., 400, 404, 519 Kirk, L M., 377 Kirnos, M. D., 486 Kirolos, M. A., 24, 25(40), 346 Kiselyov, A. S., 132, 345 Kiss, R., 620 Kleider, W., 329 Kleiman, L., 508,509(19) Klement, R., 159, 489 Klimkait, T., 22(25), 23(25; 32), 24, 25(35) Klinov, D. V., 249 Klotz, I. M., 3, 6(4), 100, 106(14) Klug, A., 216, 503,508(4), 516(4), 556, 593, 594, 595(2; 9), 597 Knoll, J. H., 577 Knops, J., 298,304(90) Knowles, J. R., 379 Knox, R. J., 371 Koberle, B., 252 Koch, T. H., 338,408

670

AUTHOR INDEX

Koga, S., 578 Kohn, K. W., 291,388,616, 617,619, 620, 620(28; 29) K6hrle, J., 612, 623(14) Koizumi, M., 291 Kojima, C., 262 Kojiri, K., 614 Kolb, A., 469(4), 470 Kollhagen, G., 617, 619,620 Kollman, E A., 11, 14(38), 292, 293, 294(47), 295,296, 296(31; 57-59), 297(57; 59; 61), 298,302, 304(31), 305(31), 307(31), 308(31), 311(115), 312(31), 318(115), 319(31), 324(115), 392 Kolodziej, E., 380, 389 Kolovos, M., 563 Komata, T., 578 Komura, J., 359, 364(7), 375(7) Kondo, H., 557 Kondo, S., 578 Kondo, Y., 578 Kondoh, A., 27 K6nig, E, 585 Koo, H.-S., 332,494 Kool, E. T., 250 Kopka, M. L., 496 Kornberg, R. D., 503,504, 506(18), 518(3) Kosaak, J. C. T., 203 Koseki, S., 300, 302(99), 311 (99), 318(99), 324(99) Koshlap, K. M., 253,278(5) Koudriakova, T. B., 359 Kowlski, D., 363 Kozloski, S. J., 104 Kozlowski, S. A., 103 Krawczyk, S. H., 313 Kreuzer, K. N., 388 Krippl, B., 648 Krohne, G., 647 Kron, S., 338 Krude, T., 651 Krugh, T. R., 18, 106, 160, 385 Kubinec, M. G., 266 Kubista, M., 69, 88 Kubo, Y., 262 Kuhn, H., 332, 333,337, 339(20) Kuhn, P., 610 Kuklenyik, Z., 315 Kukreti, S., 345, 354, 354(19), 356(19)

Kulkarni, M. S., 385,392(61) Kumar, A., 4, 22, 22(21; 22), 23(21; 22), 24, 24(5), 25(5), 26, 29, 29(51), 50(58), 259, 262, 270, 634 Kumar, P., 23(27), 24 Kumar, S., 29, 118, 131(31), 144, 324 Kung, H. C., 280 Kung, P. E, 256 Kuntz, I. D., 23(30), 24, 590 Kupke, D. W., 149, 150, 151, 153(8), 154, 155(18), 156, 157(18), 158(4), 159, 162(18), 163 Kurachi, A., 577 Kurakin, A., 336, 339(38) Kuroi, K., 581 Kurpiewski, M. R., 223 Kurucsev, T., 69, 77 Kushner, D. M., 578, 580(52) Kwok, Y., 558, 567(20), 570(20) Kypr, J., 493 Kyratzis, I., 535

L Laaksonen, A., 297 Labhardt, A., 212 Laco, G. S., 611 Lacoste, J. C., 343,556 Lacroix, L., 343,556 Ladbury, J. E., 4, 100(20), 101, 111, 114, 115(24), 126(22), 134, 216, 223(15), 347, 348(37), 581,586 Laemmli, U. K., 643 Lagutina, I. V., 249 Lahm, A., 416 Lahmy, S., 612 Lamberson, C., 557 Lamm, G. M., 297,489 Lamond, A. L., 313,489 Lancelot, G., 281 Landsorp, P. M., 576, 577(29) Lane, A. N., 118, 126, 131(33; 40), 252,253, 256, 258,259,260(17), 261,262(37), 263, 264(40), 265(38; 40), 266, 269, 269(40), 270(25), 271,273,274(74-76), 275(64), 276(75), 277(7; 34), 278,278(17; 69), 279, 280(69; 81), 295,313,314, 315(144), 317, 347,351(35), 353(35) Lane, B., 645

AUTHORINDEX Lane, M. J., 165, 166, 167(8), 168(8), 170(8), 171(1; 8), 172(1; 8), 173(8), 175(8), 176(8), 178(8), 446 Laney, D. E., 240, 248 Lange, P., 404 Langley, D. R., 296, 297(72; 75) Langmore, J. P., 138 Langowski, J., 248 Lansdorp, P. M., 576, 577 Lansiaux, A., 612, 622 Laoui, A., 137,578 Lapatsanis, L., 563 Laplante, S. R., 262 LaPointe, J. W., 504, 506(18) Latimer, E W., 249 Larsen, A. K., 623 Larsen, H. J., 332,336, 337, 339(37; 38) Larsen, T. A., 83 Larson, J. E., 202, 389 Larsson, A., 29 Laskey, R. A., 421,510, 634, 637(7; 20), 638, 641(7), 644, 648(7), 650, 650(7), 651, 651 (7) Laskowski, M., 415 Latimer, L. J. P., 131,345 Latt, S. A., 194 Laug'~a, P., 383,556, 557(13) Laughlan, G., 324 Laughton, C. A., 29, 118, 131(31), 271,274(79), 280, 296, 297,298,298(62), 306, 313, 314(62), 317 Lavelle, F., 137,578,623 Lavergne, O., 620, 622 Lavery, R., 300, 308(102), 317, 319(102) Lavesa, M., 414 Lawley, P. D., 402 Lawrence, D. A., 359 Lawrence, R. A., 138,589 Le, H., 250 Le, S., 574 Leadon, S. A., 359 Leaman, D. W., 578, 580(52) Leavitt, A. J., 240, 241(25) Lebedev, A. V., 150 Lebedev, Y., 249, 489 Le Doan, T., 130, 340, 343 Lee, C.-H., 54 Lee, G. U., 240, 241(28) Lee, H., 294 Lee, H. W., 576, 577(29)

671

Lee, J., 568,570(29) Lee, J. K., 291 Lee, J. S., 131,345 Lee, M. S., 579 LeFeuvre, C. E., 573,574(6) Legault, P., 253(9), 254, 281 Legg, M. J., 421 Legha, S. S., 388 Le Grimellec, C., 250 Leharne, S. A., 128, 130(47) Lehn, J. M., 344, 356(18) Lehrach, H., 489 Lemaire, M. A., 485 Lemrow, S. M., 433 Leng, E, 5, 291,546, 549(30), 568,639, 645(10), 647(10), 650(10), 652(10) Leng, M., 485,519 Leno, G. H., 634, 637(13; 20), 639,641,643, 643(2; 13), 644, 644(13), 645(10; 16), 646(18), 647(10; 18), 650(10), 651,652(10) Leo, A., 394 Leonard, G. A., 284, 285(1 l) Leong, T., 489 LePecq, J. B., 380 Le Pecq, J. B., 383,556, 557(13) Lepple, F. K., 156, 157(26) Lerman, L. S., 51,378,380 Leroy, J.-L., 263 Leseur-Ginot, L., 620 Lesnik, E. A., 333 Lesser, D. R., 223 Lesuer-Ginot, L., 620 Leteurtre, E, 617,620 Leupin, W., 212, 222, 266, 556, 557(10) Levenson, C., 132 Levitt, M., 293 Levy, G. C., 262 Levy, M. Z., 573(13), 574, 576(14) Lewis, J. D., 551 Lhomme, J., 130, 340 Li, K., 22, 22(21), 23(21), 24, 24(5), 25(5) Li, M., 238 Li, Q., 390(89), 391 Li, Y. K., 408 Liang, K. W., 337 Liang, X., 337 Liaw, Y. C., 408 Lichtsteiner, S., 576 Lieber, C. M., 238 Liepinsh, E., 266, 267

672

AUTHOR INDEX

Lifson, S., 193, 196(3) Ligget, D., 203 Lillehei, P. T., 234 Lilley, D. M., 54, 61(10), 139, 324, 612 Limborska, S. A., 489 Lin, K. Y., 464 Lin, L.-N., 115, 121(26), 128,545 Lincoln, P., 73,347,570 Lindau, S., 29 Linder, S., 291 Lindsay, S. M., 237, 240, 241(24) Lindsey, J., 573(13), 574, 576(13) Lindsey, L. A., 573(13), 574, 576(13) Ling, Y. H., 291,568 Lingner, J., 574 Lipanov, A. A., 212 Lipari, G., 261 Lipman, R., 404 Lippard, S. J., 86, 164, 519 Lipscomb, L. A., 234, 286(16), 287,289 Lipton, M., 300, 303(98), 304(98), 307(98), 311(98), 319(98), 320(98) Liquier, J., 489 Liskamp, R., 300, 303(98), 304(98), 307(98), 311(98), 319(98), 320(98) Liu, K., 301 Liu, L. E, 52, 386 Liu, L. R., 386 Liu, Q., 578 Liu, X., 24, 25(40) Liu, Y., 203 Live, D., 315 Loakes, D., 354 Locker, D., 485 Lockshin, C., 283 Lohka, M. J., 634 Lohman, T. M., 9, 147,422 Lobse, J., 334, 335(28), 338 Lokey, R. S., 556, 558,567(20), 570(20) Lomonossoff, G. P., 421 London, R. E., 255 Long, B. H., 611,622(11) Loo, J. A., 22(24), 23(24), 24, 433 Loontiens, E G., 4, 5(21), 212, 213(11), 216(I0), 217,219, 225(19), 228(28), 229 Lopez-Guajardo, C. C., 583,589(85) Lorch, Y., 503,504, 506(18), 518(3) Louis, J. M., 262 Louissaint, M., 336, 337 Lourenco, A., 235(3), 236

Lourim, D., 647 Loutfy, R. O., 345 Low, C. M. L., 418,419(13), 421(13), 429(13), 504, 511(9), 514(9), 515(9) Lown, J. W., 531 Lu, M., 138, 143 Lu, X. J., 306 Lu, Z. H., 643,646(18), 647(18), 651 Luce, R. A., 396, 408 Luck, G., 24(43), 25 Ludlum, D. B., 406 Luedtke, N. W., 25 Luger, K., 503 Luisi, B., 139, 301,324 Lund, K., 616 Lundberg, A. S., 577 Lundblad, V., 574 Luneva, N. P., 163 Luque, E J., 292(27), 293,296, 298 Luque, F. Q., 301 Lusthof, K. J., 397 Lutter, L. C., 504, 511(16) Luty, B. A., 294 Lwowski, W., 379, 386(21) Lyamichev, V. I., 130, 131 Lyng, R., 79, 81, 81(13), 92 Lyon, C., 645 Lyubartsev, A. P., 297 Lyubchenko, Y. L., 170, 240, 241,241(24), 249

M Ma, Y., 359 Macaya, R. F., 139, 144, 278 MacCross, M., 260 Machinek, R., 159 Mack, D. P., 22(24), 23(24), 24 Macke, T., 300, 307(103) MacKenzie, L., 623 MacKerell, A. D., 293,297 Mader, A. W., 503 Magasnik, B., 469(3), 470 Magno, S. M., 509, 517(27) Mahbubani, H. M., 651 Maher, L. H., I 18 Mahieu, C., 612 Mailliet, P., 137, 578 Makarov, V. L., 138 Malonne, H., 620

AUTHOR INDEX Malvy, C., 248 Mamoon, N. M., 209 Manalili, S., 25 Mandel, H. G., 381 Maniatis, T., 363,435,600, 608(14) Mann, D. B., 485 Mann, M., 574 Manne, S., 239 Manning, D. D., 26, 29(49) Manning, G. S., 9, 10(29; 30) Manor, H., 590 Mantilla, E. J., 519 Mantovani, R., 529 Mar, V., 574 Marathias, V. M., 144, 299, 324 Marcell, V., 589 Marchand, C., 344, 347,350, 356, 356(18), 424, 486, 624, 626 Marco-Haviv, Y., 590 Margeat, E., 250 Margulis, L. A., 164 Mark, A. E., 292(28), 293 Markgren, P.-O., 26, 49(52) Markley, J. L., 257 Markovits, J., 388, 556, 557(13) Marks, D. S., 22(14), 23 Marks, J. N., 423 Marky, L. A., 111, 114(18), 115(18), 138, 143, 149, 150, 151, 155(18), 156, 157, 157(18), 158(4), 159, 162(18), 163, 165, 170, 174(24), 175(23), 348 Marsch, G. A., 379(32), 380, 385,385(32), 393 Marsh, T., 239 Marsoni, W., 388 Martin, A., 262 Martfn, B., 291,529, 551,551(5), 554(5; 33), 555(33) Martin, M. T., 131,345,347(23) Martin, P. L., 552 Martin, R. E, 230, 233(29) Martin, R. G., 262 Marx, G., 387,388(74) Marzilli, L. G., 11, 14(40), 315 Mascetti, G., 516 Masse, J. E., 281 Masters, J. R., 252 Mathews, D. H., 444 Matousek, J., 181,184(43), 186(43), 187(43) Matsudaira, P., 338 Matsugami, A., 23(27), 24

673

Matsumoto, M., 614 Matsunaga, N., 300, 302(99), 311(99), 318(99), 324(99) Matsuoka, Y., 79 Matsuura, A., 574 Mattaj, I. W., 551 Mattern, M. R., 388 Mattes, W., 388 Matteucci, M. D., 313,464 Matts, R. L., 581 Maurizot, J. C., 346 Maxam, A. M., 416, 458 Maxam, M., 361 Maxwell, A., 51(2), 52 Mayfield, C. A., 291,312, 355 Mayo, J. A., 486 Mayo, K. H., 262 Mazen, A., 509 Mazumder, A., 619, 620, 624, 633 Mazur, S. J., 4, 29, 50(58), 485, 55l Mazzarelli, J. M., 494, 499(33) McAdam, S. R., 358,358(6), 359,364(3; 4; 6), 375(3; 4; 6) McCammon, J. A., 294 McCampbell, C. R., 170 McCloskey, J. A., 403 McConnaughie, A. W., 22, 24(5), 25(5) McCurdy, S., 144 McCutchan, R. E, 22, 23(9) McDark, C. J., 252 McFail-Isom, L., 156, 226, 242, 243(34), 247, 247(34-36), 284, 285(12) McGhee, J. D., 6, 7(24), 15(24), 54, 64(15), 65(15), 77, 99, 193, 194, 196(1), 199(l), 201(1), 202(1), 204(9), 209(9), 245, 246(41), 247(41), 546 McGill, N. J., 573(13), 574, 576(13) McGurk, C. J., 358 McHugh, P. J., 252,358 McKenna, R., 133,134(73), 136(73), 586 McKenzie, S. E., 169 McKie, J. H., 214, 222(12) McLaughlin, L. W., 212, 213(11), 313,401(7), 402, 489, 494, 499(33) McLean, M. J., 426, 427, 432,486 McMahon, N. J., 581 McMurray, C. T., 508,509, 509(20), 515(20), 516(20), 518(28) McNeese, J., 422 McPhail, T., 574

674

AUTHOR INDEX

McPhee, J. D., 200, 203 McPherson, A., 282 McPike, M. P., 43 l, 436 McRee, D. E., 282 Meadows, R. E, 264 Mei, H.-Y., 22(23; 24), 23(23; 24), 24, 25(23), 433 Meier, J., 645 Meister, W.-V., 29 Mel'nikov, S. M., 203,206(15) Menissier de Murcia, J., 591 Mergny, J.-L., 99, 110, 131(7), 137,343,350, 351(15), 355,356(45), 556, 578 Merz, K. M., 293,296(31), 304(31), 305(31), 307(31), 308(31), 312(31), 319(31) Meunier, B., 212, 338 Meyerson, M., 574, 577, 578 Michaelis, L., 377 Michaels, S., 388 Michel, J., 313 Michl, J., 69(4), 70 Miles, H. T., 301,319(108; 109), 493 Milias, G., 563 Millard, J. T., 400, 401(8), 402,519 Miller, D. M., 131,291,298, 304(90), 312, 314, 315(142), 316(142), 355 Miller, J. E, 600, 608(15) Miller, J. L., 294(47), 295 Millero, F. J., 151, 154(11), 155(11; 16), 156, 156(11), 157(26) Milligan, J. F., 34, 464 Mills, A. D., 638,650 Minford, J., 388 Minnock, A., 499, 500, 502 Mirkin, S. M., 110, 117(9), 130, 131,131(9), 341 Mischiati, C., 29 Misra, V. K., 156 Missailidis, S., 297 Mittmann, M., 166 Mizan, S., 132, 345 Mizusawa, H., 54 Moate, E J., 479 Modrich, E, 249 Moelling, K., 312 Mohamadi, E, 300, 303(98), 304(98), 307(98), 311(98), 319(98), 320(98) Mohamed, A. J., 337 Mohammadi, S., 489 Mohanty, D., 143 Mc,llegaard, N. E., 332, 335,489, 494

Moiler, A., 103, 104 Mollman, E A., 302, 311(114), 318(114), 324(114) Molteni, V., 626 Moncollin, V., 529, 551 Mongelli, N., 529 Monod, J., 16 Montenay-Garestier, T., 99, 110, 131(7), 343, 351(15), 392, 504, 509(13), 516(13), 517(13), 518(13) Montgomery, J. A., 300, 302(99), 311(99), 318(99), 324(99) Moody, M. H., 324 Moody, E C., 324 Moore, M. H., 324 Mooren, M. M. W., 257 Moraru-Allen, A. A., 347,351(35), 353(35) Moreau, S., 313 Moreland, D. W., 22(24), 23(24), 24 Morgan, A. R., 396(1), 397 Morin, G. B., 574, 575(16), 576, 581,582(16), 583(16) Morita, M., 384 Moroney, S. E., 486 Morozov, V., 156 Morris, E L., 336 Morse, D. E., 238 Mortensen, K., 88 Morton, T. A., 30, 48, 48(69), 49, 49(69; 77) Moseley, H. N. B., 264 Moser, H. E., 130, 340, 356(2) Mosier, D. E., 450, 529 Moss, H., 584,585(87) Mote, J., 485 Motherwell, W. D. S., 301 Moyzis, R., 99, 283 Mrksich, M., 291 Mucenski, M. L., 249 Mueller, P. R., 358 Muller, S., 591 Miiller, W., 54, 64(14), 65(14), 100, 389 Mullis, K. B., 165 Multani, A. S., 578 Munshi, R., 643,646(18), 647(18) Muraoka, M., 493 Murchie, A. I. H., 139, 324 Murphy, M., 131 Murphy, W. R., Jr., 519, 520, 521(12), 527(12), 528(12) Murr, M. M., 556, 558,567(23), 570(23)

675

AUTHOR INDEX Murray, A. W., 634 Musso, M., 341,349 Myasnikov, V. A., 489 Myers, T. G., 291 Mymryk, J. S., 504 Myszka, D. G., 27, 29, 30, 37, 38(75), 39(75), 41(75), 48, 48(68; 69), 49, 49(69; 77), 50

N Nagahara, L. A., 240 Naghibi, H., 126, 135(41) Nair, T. M., 29 Nakamura, H., 247,574 Nakamura, T. M., 574 Nakanishi, K., 69 Nakano, H., 620 Nakano, S.-I., 170, 175(36), 176(36), 178(36) Nakayama, J., 574 Nasuti, C. A., 623 Neamati, N., 624, 626, 633 Neely, L., 450 Negroni, M., 27 Neidle, S., 4, 29, 50(58), 99, 118, 131, 131(31), 132(67), 133, 134(73), 136(73-75), 141, 222, 261,262(37), 264(40), 265(38; 40), 269(40), 271,274(74; 79), 295,298,299, 313,324, 350, 377,486, 578, 583,586, 589, 589(85) Neilsen, E E., 331 Neilson, E., 379 Nekludova, L., 599 Nelson, E. M., 386 Nelson, H. C. M., 216 Nelson, L. D., 341 Nemunaitis, J., 579 Neretina, T., 249 Nesetril, J., 165, 17l(1), 172(1), 181, 184(43), 185,186(43), 187(43) Nettikadan, S., 249 Neuner, E, 313 Neville, D., 389 Newman, M., 521 Newmeyer, D. D., 634 Newport, J. W,, 639, 645 Newton, B. A., 381 Nguyen, C. H., 99, 110, 131,131(7), 337,345, 346,347,347(23; 25), 349, 350, 351 (25), 354, 354(19), 356, 356(19; 45), 424, 623

Nguyen, K. A., 300, 302(99), 311(99), 318(99), 324(99) Ni, E, 264 Nicholls, A., 300, 307 Nicolini, C., 516 Niedergang, C., 591 Nielsen, P. E., 90, 92,248, 329,330, 330(2), 331, 331(1), 332,332(14), 333, 333(5; 11), 334, 334(8), 335,335(28), 336, 336(27; 29), 337,338,338(1; 8; 14), 339, 339(20; 37; 38), 427,489,494, 556, 557(3) Nieves-Neira, W., 617 Nikitin, A. Y., 359 Nikonowicz, E. P., 281 Nilsson, L., 296 Nilsson, E, 29 Nishikawa, S., 23(27), 24 Nishimura, S., 614, 622(23) Nishioka, D., 589 Nishira, S., 557 Nitiss, J. L., 617 Noble, S. A., 335,580 Nod6n, B., 69 Noftz, S. L., 485 Noguchi, K., 614 Nomura, K., 614 Nord6n, B., 24, 25(41), 68, 69(3), 70, 73, 76, 79, 81,81(13), 83, 86, 88, 90, 92, 94, 329, 331, 332, 333(5), 346, 347,516, 517,556, 570 Nordheim, A., 103, 104 Norman, D. G., 324 Norwood, T. J., 258 Nygren, P.-A., 29

O O'Brien, R., 581 O'Connor, T. R., 359 Oden, E I., 240, 241(24) O'Ferrall, C. E., 240, 241(28) Ogata, C., 287 Ogonskie, N., 579 Ohkubo, M., 614 Ohnesorge, E, 239 Ohyahiki, J. H., 582 Ohyashiki, K., 582 Ojemann, J., 18, 106 Okura, A., 614 Olafson, B. D., 293

676

AUTHOR INDEX

Oldfield, E,, 257 Oldstone, M. B. A., 22 Olivas, W. M., 118 Olmstead, M. C., 20 Olovnikov, A. M., 573,573(11), 574 Olson, W. K., 24(42), 25,300, 306, 307(105), 311(105), 352 O'Malley, B. W., 131 Onfelt, B,, 570 Ono, A., 262, 281 Orgel, A., 377 Omstein, R. L., 170 Orozco, M., 280, 292(27), 293,296, 298, 298(62), 301,314(62), 317 Orum, H., 329, 338 Osborne, M. R,, 163 Osheroff, N., 385, 387,388(74), 611 O'Toole, N., 22(14), 23 Otterbach, E, 582 Otting, G., 266, 267, 268 Ottinger, E. A., 581 Otto, P., 170, 171(34), 172(34) Ouellette, M. M., 576, 581 Ouhashi, K., 23(27), 24 Oulton, R., 574 Owczarzy, R., 165, 166, 167(8), 168, 168(8), 170(8; 15), 171(1; 8; 15), 172(1; 8; 15), 173(8), 175(8), 176(8), 178(8)

P Pabo, C. O., 594, 597(12), 599, 607(7) Pace, B., 336 Pace, N, R,, 34 Pack, G, R., 297 Packman, L, C., 29 Padmanabhan, K. P., 144 Paetkau, V., 396(1), 397 Palecek, E., 426 Paloczi, G, T., 251 Palu, G., 509, 517(27) Palumbo, M., 509, 516(26), 517(27) Pan, C., 577 Pancoska, E, 165, 171(1L 172(1), 181,185 Pandit, B., 578,580(53) Paner, T. M., 165, 166, 167(8), 168(8), 170, 170(8), 171(1; 8; 26), 172(1; 8; 26), 173(8), 175(8), 176(8), 178(8) Panetta, G., 502

Pang, D., 248 Panthananickal, A., 394 Paoletti, C., 380 Papas, T., 27 Paquet, E, 281 Paranjape, J. M., 578,580(52) Pardi, A., 253(9), 254, 281 Park, Y. W., 346 Parkinson, J. A., 212,214, 222(12), 254 Parslow, T. G., 433,443(8) Parsons, P. G., 473 Passner, J. M., 18, 106 Patel, D. J., 23(31), 24, 103, 139, 164, 280, 301, 315,341 Pathak, S., 578 Patino, N., 433 Pavletich, N. P., 581,597(12), 599 Payet, D., 495,496(39) Payton, N., 345 Peace, D. J., 577 Peacocke, A. R., 14,378 Pearl, L. H., 324, 581 Pearson, E. C., 503,507(5) Pedersen, H. G., 294 Pedersen, L. G., 294 Pednault, E. P., 306 Peek, M. E., 234, 282, 286(16), 287, 289 Peffer, N. J., 335,580 Perera, L., 294 Pdrez, J. J., 512,515 Perez-Soler, R., 291, 531,568 Perlman, M. E., 255 Perouault, L., 130 Perrin, D., 356 Perrin, L. C., 483 Perrouault, L., 99, 110, 131(7), 340, 343, 354 Perry, P. J., 110, 131,132(67), 137(16), 141, 298,299, 586, 589, 589(92) Persson, B., 29 Perun, T. J., 109 Pervushin, K., 281 Peterson, R. D., 281 Peterson, S. R., 248 Pettitt, B. M., 130, 294(49-51), 295,296, 298, 298(49; 50) Peytou, V., 433 Pfannschmidt, C., 248 Pfeifer, G. P., 358, 359, 364, 365(17), 372(17), 375,376(20) Pham, T~ Q., 144, 324

AUTHOR INDEX Philippart, E, 620 Phillips, D. R., 390(90), 391,408,466, 467,469, 469(5), 470, 471,473,473(5), 476(7), 477, 478,479, 481,482(10), 483,483(7; 10; 13), 484, 485,535,550, 551 Phillips, K., 139, 301,324 Philpott, A., 637(13; 20), 641,643(13), 644, 644(13) Piatyszek, M. A., 137,576, 581,581(32), 582(32) Pickett, S. C., 479 Piel, N., 489 Pieper, R. O., 485 Pietrasanta, L. I., 251 Pilch, D. S., 24, 24(42), 25, 25(40), 131,132, 143,345,346, 347(23; 25), 351(25), 352, 352(32) Pillus, L., 518 Pilon, A. A., 619, 624, 633 Pinder, R. M., 381 Pineda, A. R., 30, 48(67) Pines, J., 651 Piotto, M., 269 Piper, P. W., 581 Pippo, L., 29 Pitts, A. E., 580 Pitts, E. E., 578,580(54) Pjura, E E., 212 Pla Rodas, E, 620 Plaskon, R. R., 170, 234, 286(16), 289 Plaxco, K. W., 251 Plotnikov, V. V., 128 Plukett, W., 131 Plum, G. E., 24, 25(40), 110, 117(10), 118(10), 128, 131(10), 143, 168, 177(17), 208 Pogozelski, W. K., 422 Poland, D., 167, 168(10), 169(10), 193 Polucci, E, 131,350 Polycarpou-Schwarz, M., 551 Pommier, Y., 291,388,610, 611,616, 617, 619, 620, 620(28; 29), 624, 626, 633 Pons, D., 620 Poremba, C., 582 Porter, S. E., 620 Portugal, J., 291,422, 427,432, 503,504, 512, 514, 514(10), 515, 515(10), 529, 551, 551(5), 554(5; 33), 555(33) Post, C. B., 264 Potaman, V. N., 110, 117(15), 118(15), 349 Potapov, V. K., 489

677

Potekhin, S. A., 127, 128 Pourquier, E, 610, 611,617,619 Pouyet, J., 509 Povirk, L. E, 518 Prabhakaran, M, 311 Praseuth, D., 110, 130, 331,340, 341 Prescott, C. D., 22 Prescott, D. M., 138 Presnell, S. R., 234, 247, 286(16), 289 Prestegard, J. H., 280 Preuss, W., 239 Prevost, G., 620 Pribble, J., 579 Price, M. A., 422 Price, E A., 415 Priebe, W., 5, 19, 291,308,529, 531,532, 532(17), 539, 540(24), 546, 549(24; 30), 550(24), 551, 551(5), 554(5; 33), 555(33), 568 Priestley, E. S., 433 Priestley, W. S., 26, 29(48; 50), 31(50), 36(48), 38(50) Pritchard, L. L., 331 Privalov, P. L., 127, 168 Prodromou, C., 581 Prokhorov, V. V., 249 Propst, C. L., 109 Prosnyak, M. I., 489 Prosser, J. K., 117 Prowse, K. R., 137,576, 581(32), 582(32) Prudhomme, M., 614, 620(19), 622(19) Pruitt, K. M., 384 Prunell, A., 510 Przewloka, T., 291,529, 532,568 Puglisi, J. D., 22(11-13), 23, 23(29; 30), 24, 33(70), 34, 281,435,446 Pulleyblank, D. E., 54 Pullman, B., 254, 531 Pulyaeva, H., 337 Pursell, C., 558, 567(20), 570(20) Putman, C. A., 239 Puvvada, M. S., 485 Pyle, A. M., 34

Q Qin, P. Z., 34 Qiu, X., 610 Qu, X., 4, 8(13), 19, 308,377. 614

678

AUTHORINDEX

Quada, J., 583,589(85) Quadrifoglio, E, 157 Quigley, G. J., 283,309, 311(126), 320(126), 419, 519, 531,533(18), 535(18) Quintana, J. R., 212,226

R Rabbani, A., 504, 507(14), 509(14), 516(14) Rackwitz, H. R., 489 Radhakrishnan, I., 280, 315,341 Radmacher, M., 240 Raghavan, S., 99 Raghunathan, G., 301,319(108; 109) Raghuraman, M. K., 117 Ragsdale, C. W., 600, 608(15) Ragunathan, K. G., 22, 24(5), 25(5) Rahmouni, A. R., 485 Rajewsky, M. E, 359 Rajur, S. B., 401(7), 402 Rama Krishna, N., 264 Ramasamy, K., 336 Rando, R. R., 25 Rao, S. N., 11, 14(38), 392 Ratliff, R., 99, 283 Ratmeyer, L., 22(22), 23(22), 24 Raucher, S., 400, 401(8), 402,519 Rauen, H. M., 377 Ravishanker, G., 295,296(53), 297(53) Ray, R., 131 Raymond, E., 138,578,586, 589 Rayner, D. M., 345 Razzano, G., 21 Read, M. A., 13l, 132(67), 141,298,299, 589 Rebar, E. J., 594 Recht, M. I., 22(11; 13), 23, 23(33), 24, 446 Record, M. T., Jr., 9, 20, 147, 168, 195,519 Reddel, R. R., 577 Reddoch, J. E, 291,314, 315(142), 316(142) Redinbo, M. R., 610 Reedjil, J., 519 Rees, D. C., 289, 450, 529(7), 530 Reese, C. B., 313 Reeves, R. H., 242 Regenfuss, P., 4, 5(21), 212,216(10), 217 Rehfuss, R., 393,418,432, 446, 446(3) Reich, E., 377 Reich, N. O., 249

Reid, B. R., 226, 253,258,265,270(59), 277(7), 570, 580 Reid, C. M., 168 Reimann, P., 238 Reines, D., 485 Reinhardt, C. G., 385 Reinhold, W. C., 291 Reinhoudt, D. N., 397 Reiss, J. A., 535 Ren, J., 4, 99, 101(10; 11), 107(12), 108(10-12), 111, 114(20), 115(20), 117(20), 123(20), 124(20), 126, 135(20; 39), 157,298, 348, 592 Rentzeperis, D., 111, 114(18), 115(18), 156, 157, 159, 163,348 Resing, K., 408 Reszka, A. E, 131, 132(67), 133, 134(73), 136(73), 141,298,299, 586, 589 Revenko, |., 248 Rewcastle, G. W., 380 Reynolds, M. A., 259 Reynolds, P. E., 109 Rhee, S., 301 Rhoads, R. E., 166 Rhodes, D., 510, 585 Riccelli, P. V., 165, 171(1), 172(1) Rice, S. A., 153 Rich, A., 99, 103, 104, 130, 212, 283,309, 311(126), 320(126), 419, 519,531, 533(18), 535(18) Rich, N., 473 Rich, R. L., 27 Richards, N. G., 300, 303(98), 304(98), 307(98), 311(98), 319(98), 320(98) Richenberg, J. L., 427, 432 Richmond, M. H., 109 Richmond, R. K., 503 Richmond, T. J., 503 Richter, M., 251 Ridge, G. S., 391,392, 392(92), 393(92; 93) Rief, M., 251 Riggs, A. D., 358, 359, 364(7), 375(7) Rigl, C. T., 22, 24(5), 25(5) Rigl, T., 22(21), 23(21), 24 Rill, R. L., 379(32), 380, 385,385(32), 393,508, 516(22) Rink, S. M., 401(7), 402 Riordan, J. E, 435 Riou, J.-E, 137,578,614, 620(19), 622(19), 623 Rippe, K., 250

679

AUTHOR INDEX Risen, L. M., 333,336 Rivalle, C., 343,504, 509(13), 516(13), 517(13), 518(13) Rivetti, C., 240 Rizzo, V., 21 Roberts, J. J., 371 Robertson, M. W., 416 Robertson, S. A., 291 Robinson, H., 539, 540(24), 549(24), 550(24) Robinson, M. J., 387 Robinson, M. O., 574 Robles, J., 313 Rock, S. G., 394 Rodger, A., 69(3), 70, 81,346 Rodgers, J. R., 30l Rodrigues-Pereira, E., 614, 620(19), 622(19) Rodu, B., 312,355 Roe, J. A., 139 Roe, S. M., 581 Roeder, R. G., 551,552 Roesel, J., 312 Rogers, J., 22(18), 23 Rogers, M. J., 22, 23(9) Rolland, A., 620 Rolli, V., 591 Romanowski, E, 651 Rome, L. H., 592 Roques, B. E, 287,383,556, 557(13) Rosamond, J., 212 Rosamund, J., 254 Roselt, P. D., 313 Rosen, N., 581 Rosenberg, J. M., 296 Rosenbohm, C., 26, 29(49) Rosendahl, G., 24 Rosenfield, S., 584, 585(87) Rosenzweig, A. C., 86, 164 Ross, D. T., 291 Ross, J., 153 Ross, P., 579 Ross, W. E., 386 Roug6e, M., 99, 110, 131,131(7), 343,346, 347(25), 351(15; 25) Rould, M. A., 599 Rouzina, I., 168 Rowe, T. C., 386 Roy, J., 579 Royer, C. A., 250 Rozek, D., 375,376(20) Rubin, C. M., 426

Rubin, E., 614 Ruland, C., 592 Russell, E E., 238 Russo, A. A., 581 Riiterjans, H., 262 Rutigliano, C., 29 Rutter, J. P., 580 Ryberg, H., 88 Ryberg, M., 88 Rychlik, W., 166 Ryder, U., 313,489

S Sabina, J., 444 Sacchi, N., 21 Sadat-Ebrahimi, S. E., 22(14), 23 Sadler, J. E., 144 Saecker, R. M., 168 Saenger, W., 150, 156(1) Sagi, J., 493 Sagui, C., 294 Saito, M., 574 Salazar, M., 138,226, 299, 323,580, 583,586, 589(85), 590, 592 Saleem, A., 614 Sambrook, J., 363,435,600, 608(14) Samper, E., 576, 577(29) Sanaterre, J. E, 203 Sanchez-Garcia, I., 594 Sannes-Lowery, K. A., 22(24), 23(24), 24, 25, 433 Sano, T., 582 Sansom, C. E., 271,274(78) SantaLucia, J., 170, 171(35), 172(35), 174(35), 175(35), 176, 176(35), 178(35), 329 SantaLucia, J., Jr., 168, 170(14), 174(14), 175(14), 176(14), 177(14), 178(14) Sapolsky, R. J., 166 Sargent, D. E, 503 Sarma, R. H., 311 Sarvazyan, A. E, 150 Sasi, R., 508 Sasisekharan, V., 143, 301,319(108; 109) Sasmor, D. J., 25 Sasmor, H., 25 Satyanarayana, S., 100(19), 101,532 Saucier, J. M., 623 Saudek. V., 269

680

AUTHOR INDEX

Sausville, E. A., 291 Savre-Train, I., 576 Sawadogo, M., 551 Scaria, P. V., 106, 143, 343,345(14) Scatchard, G., 54, 62(12), 194, 246 Schaeffer, F., 469(4), 470 Schafer, K. L., 582 Schaffer, T. E., 239, 251 Schaper, A., 248 Scheek, R. M., 270, 277(70) Scheit, K. H., 489 Schellman, J. A., 194 Scheraga, H. A., 167, 168(10), 169(10), 193 Scherf, U., 291 Schiano-Liberatore, A.-M., 620 Schild, C., 634 Schildkraut, I., 521 Schimmel, P. R., 459 Schipper, P. E., 79 Schlessinger, F. B., 509 Schmacher, R. J., 581 Schmid, C. W., 426 Schmidt, C. E, 240 Schmidt, M. W., 300, 302(99), 311(99), 318(99), 324(99) Schmitt, A., 591 Schmitt, L., 250 Schmitz, A., 412, 451 Schmitz, U., 257,258, 259, 259(26), 264(26), 265(26), 270, 270(26), 277(26) Schneider, B., 300, 307(105), 311(105) Schneider, C., 581 Schneider, H. J., 22, 24(5), 25(5) Schreiber, V., 591 Schroeder, R., 22(17; 18), 23,444 Schuitmaker, J. J., 589 Schulemann, W., 381 Schulten, K., 306 Schultz, P. G., 451,452(15; 16) Schultz, R. G., 493 Schultze, P., 139, 144, 278 Schuster, G. B., 242, 247(36), 338 Schwartz, A., 485 Schwartz, R. E., 388 Schwarz, U., 238 Scott, E. V., l 1, 14(40) Scourides, P. A., 535 Scratchard, G., 3, 6(1) Scudiero, D. A., 291

Searle, M. S., 216, 253,254(4), 268, 276(67), 277(4; 67), 297 Sedivy, J. M., 576 Seela, E, 426 Segal, D. J., 594 Seger, D., 503,508(6), 516(6) Sehlstedt, U., 24, 25(41), 83 Sekiguchi, J., 138 Selsing, E., 319 Semenza, G., 240, 241(27) Sen, S., 296 Senear, D. F., 451,453(18-21) Seneviratne, P., 170, 171(35), 172(35), 174(35), 175(35), 176(35), 178(35), 329 Senter, J. P., 153 Sergeyev, V. G., 203, 206(15) Sethson, I., 257 Severin, S. E., 578,580(55) Seymour, C. K., 242 Shafer, R. H., 23(30), 24, 106, 132, 138, 143, 271,291,299, 343,345(14) Shahin, M. A., 9, 10(30) Shanck, B. S., 151 Shao, Z., 337 Shapiro, H., 119 Shapiro, R., 403,404 Sharma, A., 586, 619 Sharma, S., 586 Sharmeen, L., 22(24), 23(24), 24, 433 Sharon, R., 293 Sharp, K., 300 Sharp, S. L., 238 Sharpies, D., 212, 254 Shastry, B. S., 552 Shaw, B. R., 508,516(22) Shay, J. W., 137,575,576, 580, 581, 581(32; 33), 582(32; 33) Shchyolkina, A. K., 489 Shea, M. A., 451,453(18-20) Shea, R. G., 144 Sheardy, R. D., 138, 141(91), 519, 520, 521(12), 527(12), 528(12), 589 Sheehan, M. A., 638 Sheng, S., 337 Sheppard, R. C., 558 Sherer, E., 296 Shevchenko, A., 574 Shevchenko, Y., 489 Shi, Y. B., 485 Shida, T., 138

AUTHORINDEX Shieh, T.-L., 380, 389 Shields, G. C., 280, 296, 298, 298(62), 314(62), 317 Shimizu, M., 131,340 Shimotakahara, S., 404 Shindyalov, I. N., 300, 307(104), 324(104) Shippen-Lentz, D., 574 Shirahama, K., 203 Shirahata, A., 349 Shire, S. J., 143 Shlyaktenko, L. S., 240, 241,241(24), 249 Shokolenko, I., 336 Short, G. F., 617,619(30) Shubsda, M. E, 436 Shui, X., 156, 226, 284, 285(12), 287,546, 549(30), 568 Shultze, P., 314, 315(143), 317 Shuman, S., 619 Shure, M., 54 Siebenlist, U., 312 Siegel, J. S., 626 Sigal, L. H., 349 Sigman, D. A., 519 Sigurdsson, S. T., 400, 404, 408,519 Silverman, R. H., 578, 580(52) Silverstone, A. E., 469(3), 470 Simmen, K. A., 551 Simmerling, C., 306 Simmons, C. G., 578,580(56) Simpson, I. J., 4, 29, 50(58) Sinden, R. R., 349 Sines, C. C., 226 Singh, S. B., 298 Sirr, A., 281 Sittman, D. B., 643,646(18), 647(18), 651 Siva, A. C., 592 Sivolob, A., 510 Sj6berg, B., 88 Skalka, A. M., 624 Skerrett, J. N. H., 14 Skerrett, N. J. H., 378 Sklenar, H., 300, 308(102), 317,319(102) Sklenar, V., 269 Skorobogaty, A., 473,535 Sleeman, A. M., 638 Slickers, E, 313 Sligar, S. G., 239 Smerdon, M. J., 485 Smimov, I., 291 Smith, B. L., 238,250, 251,518

681

Smith C. E., 389 Smith C. I. E., 337 Smith E W., 139, 144, 253,278(5) Smith G. J. I., 626 Smith L. H., 291,359 Smith P. E., 294(49-51), 295,298(49; 50) Smith P. J. C., 319 Smith R., 579 Smith S., 591 Smo orzewska, A., 573 Snow, A. M., 519 Snyder, R. C., 131 Sobell, H. M., 389 Sober, H. A., 194 Soda, H., 586 Soliva, R., 296 Solomon, M. S., 398, 401(7), 402 Song, Y., 209 S6nnichsen, S. H., 338 Sorey, S., 568,570(29) Soto, A. M., 157 Souhami, R. L., 358,364(3), 375(3) Soyfer, V. N., 110, 117(15), 118(15) Spackova, N., 299 Spellmeyer, D. C., 293,296(31), 304(31), 305(31), 307(31), 308(31), 312(31), 319(31) Spencer, W. J., 166 Spink, C. H., 193,203,204, 204(14), 205,207, 207(14) Spisz, T. S., 242 Sponer, J., 299 Sprankle, K. G., 333 Springer, D. L., 485 Sproat, B. S., 313,489 Sprous, D., 294, 295 Squire, C. J., 222,230, 233(29) Srinivasan, A. R., 24(42), 25,300, 307(105), 311 (105), 352 Srinivasan, J., 296 Stahl, M., 301 Stankus, A. A., 10(33), 11,432 Stansel, R. M., 584, 585(87) Star, W. M., 589 Starink, J. P., 239 Stark, W., 22(25), 23(25; 32), 24, 433 States, D. J., 293 Stebbins, C. E., 581 Stec, W., 259 Stecenko, D., 489

682

AUTHOR INDEX

Stefano, J. E., 471 Steiner, R, 578 Stenglanz, R., 648 Stenhag, K., 29 Stephenson, R, 485 Stem, M. A., 519 Stem, S., 22(20), 23(20), 24, 433 Stevens, M. E, 271,274(78), 297 Stevens, R. M. D., 238 Stewart, L., 610 Stewart, N. G., 573, 574(6) Stewart, R. J., 240 Stewart, S. A., 577 Stick, R., 645 Still, W. C., 294, 300, 303(98), 304(98), 307(98), 311 (98), 319(98), 320(98) Stillinger, E H., 151 Stivers, J. T., 284, 285(13) Stone, B. A., 410 Stone, M. E, 18, 106 Stonehouse, T. J., 418 Stowell, J. G., 380, 389 Strahan, G. D., 299 Strahl, C., 578 Straney, D. C., 471 Straub, T., 612,623(14) Straume, M., 8, 114, 115(23), 127(23) Strauss, U. E, 77 Strekowski, L., 132, 345,349, 353,354 Streuli, M., 592 Strumberg, D., 610 Strzelecka, T., 521 Stucky, G. D., 238 Studier, E W., 648 Sturm, J., 156 Sturm, R. A., 473 Sturtevant, J. M., 126, 127, 135(41) Styles, J. M., 371 Su, S., 300, 302(99), 311 (99), 318(99), 324(99) Suck, D., 416, 499 Suda, H., 614 Sugawara, K., 247 Sugimoto, N., 170, 175(36), 176(36), 178(36) Suh, D., 11, 18, 234, 532, 546, 557 Sullivan, W. E, 581 Sulston, I., 313 Sun, D., 138, 141(87), 299, 573,578,582, 583, 586, 589, 589(85), 592

Sun, J. S., 110, 131,340, 341,344, 345,346, 347(23; 25), 349, 350, 351(25), 354, 354(19), 356, 356(18; 19; 45), 504, 509(13), 516(13), 517(13), 518(13) Sun, Y. C., 106 Sunder, S., 633 Suri, A. K., 23(31), 24 Svejstrup, J. Q., 616, 619, 619(26) Sverdlov, E. D., 249, 489 Svinarchuk, F., 248 Swalley, S. E., 450 Swaminathan, S., 144, 293,324 Sweet, R. M., 282 Sykes, B. D., 257 Szabo, A. G., 261,345 Szakonyi, E., 493 Szewczyk, J. W., 450, 529(7), 530, 556 Szoka, E C., 203 Szostak, J. W., 291 Szyperski, T., 281

T Taatjes, D. J., 408 Tabore|li, M., 241 Tagashira, Y., 170 Tahara, K., 557 Taillandier, E., 489 Taira, K., 23(27), 24 Takac, L,, 238 Takagi, M., 557 Takahara, P. M., 86, 164 Takahashi, M., 88 Takashima, K., 203 Takasugi, M., 355 Takenaka, S., 557 Takeyasu, K., 249 Takisawa, N., 203 Takusagawa, E, 23(34), 24, 390(88; 89), 391,495 Takusagawa, K. T., 23(34), 24, 390(89), 391 Tamayo, J., 249 Tamura, A., 126, 135(41) Tan, R. K. Z., 294 Tanabe, L., 291 Tanaka, S., 247 Tanford, C., 150 Tang, C. L., 240 Tang, D., 54

683

AUTHORINDEX Tang, K. S., 500 Tanious, E A., 4, 29, 50(58), 132, 133, 136(74), 345,558,586 Tanious, T. C., 4 Tanizawa, A., 617,620 Tanner, J. E., 263,269(54) Tarlov, M. J., 29 Tate, S., 262 Taylor, J. B., 554(39), 555 Taylor, J.-S., 451,452(15), 485 Taylor, M. J., 401(7), 402 Teat, S. J., 301 Tempczyk, A., 294 Teng, M.-K., 212 Terasawa, S., 154 Terreux, R., 433 Tesmer, V. M., 581 Teulade-Fichou, M.-E, 344, 356(18) Tewey, K. M., 386 Thayaparan, J., 298, 304(90) Thomale, J., 359 Thomas, J. O., 503,507(5) Thomas, S. D., 291,312, 314, 315(142), 316(142) Thomas, T. J., 349, 354, 577 Thommes, E, 651 Thompson, A. M., 573(12), 574 Thompson, B., 299, 583,589(85) Thompson, J. B., 251 Thomsen, T., 556, 557(11) Thomson, N. H., 237,250 Thomson, S. A., 335,580 Thrall, B. D., 485 Thulsprup, E. W., 69(4), 70 Thundat, T., 235(3), 236, 238,240, 249 Thuong, N. T., 130, 131,281,340, 343,346, 354, 392 Thurston, D. E., 485 Tilby, M. J., 358,371 Tilby, N. J., 252 Tinoco, I. T., Jr., 55 Tirado, M. M., 263 Tironi, I. G., 294 Tjerneld, E, 79, 86 Tjian, R., 552, 553(36) Toft, D. O., 581 Tokumasu, F., 249 Tolman, J. R., 280 Tomassetti, M., 29 Tomasz, M., 404, 500

Tomchick, R., 381 Tondelli, L., 21 Toney, J. H., 519 Tong, G. L., 535 Tong, W. E, 406 Tor, Y., 25 Torda, T. E., 270, 277(70) Tordova, M., 284, 285(13) Tomaletti, S, 364, 365(17), 372(17) Toyama, K., 582 Trager, J. B., 581 Tran, H., 156 Tran-Patterson, R., 312 Trant, I. O., 537 Trauger, J. W., 429,450, 529 Travers, A. A., 413,415(2), 494, 495,496(39), 499(33), 502, 503,510(7), 511(7) Trent, J. O., 19, 131,132(67), 139, 141, i41(94), 142(94), 290, 291,296, 298,299, 303(65), 304(90), 308,313, 320(65; 97), 324(97), 529,589 Trinh, L., 579,582(60) Trist, H., 469, 471,535 Tsai, M. J., 131 Tu, Y., 364, 365(17), 372(17) Tuite, E., 347,517 Tulinski, A., 144 Tullius, T. D., 399, 422,424,513,519 Tunis, M.-J. B., 150 Turner, D. H., 444 Turner, J. M., 450, 529(7), 530, 556 Turro, N., 379 Twigden, S., 556, 557(10) Tzfati, Y., 579

U Uedaira, H., 150 Uemura, D., 614 Ueng, L.-M., 619 Uesugi, S., 23(27), 24 Ughetto, G., 283,309, 311(t26), 320(126), 419, 531,533(18), 535(18) Uhlen, M., 29 Uhlenbeck, O. C., 34 Ulibarri, G., 620 Ulyanov, N. B., 259, 270 Urasaki, Y., 611 Usman, N., 212

684

AUTHOR INDEX

V Valenti, M., 617 Vallberg, H., 24, 25(41) Valle, M., 249 Vallone, P. M., 165, 166, 167(8), 168, 168(8), 170(8; 15), 171(8; 15), 172(8; 15), 173(8), 175(8), 176(8), 178(8) Valpuesta, J. M., 249 van Boom, J. H., 22(16), 23,212,258,260(27), 283,419, 519 van der Marel, G. A., 22(16), 23,212, 258, 260(27), 283,419, 519 van der Meulen, R. W., 589 VanDerveer, D., 226 vander Vliet, P. C., 291 Vanderwerf, K. O., 239 Van Dyke, M. W., 303,341 VanDyke, M. W., 349 Van Dyke, M. W., 419, 424, 451,452(13) van Gunsteren, W. E, 270, 277(70), 292(28), 293,294 vanHolde, K. E., 100 van Holde, K. E., 508,509, 509(20), 515(20), 516(20; 22), 518(28) Vanhulst, N. E, 239 Vankatachalam, S., 359 Van Landschoot, A., 228(28), 229 van Leengoed, H. L., 589 van Rosmalen, A., 408,479, 481 van Steensel, B., 573 van Wijk, J., 257 Vanyushin, B. E, 486 Vaquero, A., 291,529, 551,551(5), 554(5; 33), 555(33) Varani, G., 22 Varghese, A. J., 402 Vary, C. P., 473 Vasquez, K. M., 291 Vaziri, H., 576, 577,577(36) Veal, J. M., 580 Veiro, D., 404 Verboom, W., 397 Vercauteren, J., 313 Verdine, G. L., 404 Vergani, L., 516 Veselkov, A. G., 334 Veselovskaya, S. I., 489 Vesenka, J., 239, 240 Vezin, H., 614

Viale, R. D., 230 Viani, M. B., 251 Vicens, Q., 22, 24(4) Vickers, T. A., 336 Vigevani, A., 21 Vigneron, J.-P., 344, 356(18) Vigneswaran, N., 291,298,304(90), 312, 355 Vileponteau, B., 574 Villamarfn, S., 551,554(33), 555(33) Villeponteau, B., 573,574 Vinayak, R., 22(22), 23(22), 24 Vinograd, J., 54 Volenshtein, M. V., 194 V61ker, J., 168, 177(17) Volkovitsky, P., 337 Vologodskii, A. V., 170 von Ahsen, U., 22(18), 23 von Hippel, P. H., 6, 7(24), 15(24), 54, 64(15), 65(15), 77, 194, 204(9), 209(9), 245, 246(41), 247(41), 250 Von Hoff, D. D., 138,578,579, 582, 583,586, 589, 589(85), 592 von Kitzing, E., 494, 499(33) von Sprecken, R. S., 379(31), 380 Vorlickova, M., 493 Vosberg, H.-P., 54 Vournakis, J. N., 473

W Wadkins, R. M., 159, 380, 385(33), 386(38), 394 Wadwani, S., 464 Wagar, E. A., 165 Wagner, C. R., 495 Wagner, P., 240, 241(27) W~ihnert, U., 212, 271,509 Wakasa, M., 30 Wakelin, L., 556, 557(9; 12) Waldsich, C., 444 Walker, G. T., 18, 106 Walker, V., 404 Wall, M. E., 610, 617 Walter, F., 22, 24(4) Waiters, A., 4, 5(20) Waiters, D. A., 237,239 Waiters, P., 301 Waltham, M., 291

AUTHOR INDEX Wang, A. H.-J., 212, 283,378, 408,419, 519, 531,532, 533(18), 535(18), 539, 540(24), 549(24), 550(24) Wang, C., 117, 142(29), 143(29), 146(29), 256 Wang, G., 336 Wang, H., 576 Wang, J. C., 52, 54, 248, 303,386, 408,576, 610 Wang, K. Y., 144, 280, 324 Wang, L. K., 26, 29(51), 617,619(31) Wang, P. U., 106 Wang, S., 22(23), 23(23), 24, 25(23) Wang, S. D., 106 Wang, S. S., 574 Wang, S. Y., 402 Wang, X., 617,619(30; 31; 36) Wang, Y., 139, 167, 168(11), 169(11), 301 Wang, Z., 238 Wani, A. A., 359 Wani, M. C., 610, 617 Wank, H., 444 Ward, B., 119, 356, 393,418, 429, 432, 446, 446(3) Ward, G. K., 156, 157(26) Ward, J., 568,570(29) Waring, M. J., 10(34), 11, 12(35), 109,346, 347, 347(25), 351(25), 378, 380, 381,391, 391(91), 392, 392(92), 393(92; 93), 412, 413,417,418,419(13), 421,421(13), 422, 426, 427,429(3; 13), 432, 452, 485,486, 489,489(24), 490, 493,493(26), 494, 495, 495(24), 496, 496(24; 39), 499, 499(41), 500, 500(30), 502, 504, 509(13), 510, 511(9), 512, 514, 514(9; 10), 515(9; 10), 516(13), 517(13), 518(13), 531,555,612 Warmack, R. J., 235(3), 236, 238,249 Warshaw, M. M., 119 Wartell, R. M., 167, 168(11), 169(11), 170, 172(31), 202 Waterman, M. S., 181 Watkins, C. L., 379, 382(22; 53; 100), 383, 383(29), 384(29), 386(22), 394 Watson, D. G., 301 Watson, D. K., 27 Watson, J. D., 573 Watson, J. V., 651,652(35) Watson, T., 117 Wattez, N., 612 Watts-Tobin, R. J., 377 Weaver, R. E, 23(34), 24, 390(89), 391 Webb, A., 579

685

Weber, G., 3, 6(6) Webster, C. I., 29 Weerasinghe, S., 294(49-51), 295,298(49; 50) Wei, S. L., 106 Wei, W., 576 Weidner, M. E, 400, 519 Weimer, J. J., 241 Weinberg, R. A., 574, 576, 577,578 Weinblum, D., 402, 403(11) Weinrich, S. L., 137, 574, 576, 581(32), 582(32) Weinstein, J. N., 291 Weissig, H., 300, 307(104), 324(104) Weisz, K., 271 Wellman, S. E., 193,209, 210 Wells, R. D., 131,202, 340, 389 Wells, T. N., 241 Wemmer, D. E., 266, 291,450 Wen, L., 390(89), 391 Wendman, M. A., 237 Wenzler, L. A., 240, 241(25) West, M. D., 137, 574, 576, 581(32), 582(32) Westbrook, J., 111,283, 300, 307(104; 105), 311(105), 324(104) Westergaard, O., 616, 619, 619(26) Westhof, E., 22, 22(19), 23(19), 24, 24(4), 150 Westhof, W., 22 Weston, S. A., 416 Westphal, H., 648 Wetmur, J. G., 165,166(5) Wharton, G. III, 520, 521(12), 527(12), 528(12) Wheelhouse, R. T., 138, 141(87), 586, 589 White, J. G., 650 White, J. H., 51(3), 52, 230, 233(29) White, M. A., 581 White, R. J., 390(90), 391,469, 469(5), 470, 471,473(5), 479, 482(10), 483(10; 13) White, S., 291,450, 529(7), 530, 556 White, W. E., Jr., 379, 384, 394 Wiech, H., 581 Wijmenga, S. S., 257,258, 260(27) Wilcock, D., 650 Wiles, N. C., 18, 106 Wilke, 1. G., 312 Wilkosz, P., 296 Willenbring, H., 582 Williams, A. R., 150 Williams, D. H., 29 Williams, H. E. L., 268,276(67), 277(67), 297

686

AUTHORINDEX

Williams, L. D., 156, 226, 234, 242, 243(34), 247, 247(34-36), 282, 284, 285(12), 286(16), 287, 289, 296, 546, 549(30), 568 Williams, R. M., 27 Williamsand, J. M., 240, 241(25) Williamson, J. R., 117, 138, 147,281 Williston, S., 115, 121(26) Willmott, G., 556, 557(12) Wilson, J. H., 291 Wilson, K. L., 634, 645 Wilson, W. D., 4, l 1, 14(39; 40), 22, 22(21; 22), 23(21; 22), 24, 24(5), 25(5; 35), 26, 29, 29(51), 42, 43(76), 50(58), 132, 133, 136(74), 345,349, 354, 377,504, 508(12), 517(12), 558,586 Wilson, W. R., 380 Wilton, A. N., 22(14; 15), 23 Wiltshire, T., 242 Windle, B. E., 138,578,586, 589 Windus, T. L., 300, 302(99), 311(99), 318(99), 324(99) Wiorkiewicz-Kuczera, J., 293 Wiseman, T., 115, 121(26) Wishart, D. S., 257 Wittes, R., 388 Wittung, E, 92, 329, 331,332 Wofsy, C., 30, 48(67) Wohlrab, E, 131,340 Wold, B., 358 Wolfe, A. L., 633 Wolffe, A. E, 634, 641,644 Wong, A. H., 309, 311(126), 320(126) Wong, C.-H., 26, 29(48-50), 31(50), 36(48), 38(50), 433 Wong, L., 297 Wong, S. S., 238 Woo, J., 400, 519 Woo, R. J., 234, 286(16), 289 Wood, A. A., 141,299, 589 Wood, R. D., 252 Woody, R. W., 69 Woolley, A. T., 238 Workman, J. L., 503, 518(2) Wright, G. E., 489 Wright, W. E., 137,576, 580, 581,581(32), 582(32) Wu, C. B., 106 Wu, E, 582 Wu, H. M., 509, 515(24) Wu, H.-W., 65,468

Wu, J., 518, 611 Wu, R. S., 589 Wu, S., 579,582(60) Wurtz, N. R., 291 Wiithrich, K., 266, 267,281 Wyatt, J. R., 33(70), 34, 118,435 Wyman, J., 3, 6(3), 16

X Xiao, G., 22(21), 23(21), 24, 284, 285(13) Xiaoxin, X., 336 Xie, J. X., 106 Xie, L. Y., 576 Ximen, H. Y., 238 Xu, H., 634 Xu, J., 518 Xu, Q., 519 Xu, Y., 203,238 Xu, Z., 24(42), 25,352 Xue, B.-H., 577

Y Yalowich, J., 386 Yamada, A., 614 Yamamoto, R., 23(27), 24 Yamasaki, E. E, 359 Yanagi, K., 226 Yang, D., 262, 281 (44) Yang, G., 250 Yang, J.-C., 266 Yang, L., 294(51), 295 Yao, J., 257 Yao, S., 132, 315,345 Yao, X., 238 Yavnilovich, M. V., 334, 338 Yen, S., 558 Yeung, D. S. K., 574 Yielding, K. L., 379, 379(31), 380, 382(22; 100), 384, 385,386(22), 392(61), 394 Yielding, L. W., 379, 379(31), 380, 382(22; 53; 100), 383,383(29), 384, 384(29), 385,386(22), 394 Yin, S., 485 Yip, R. W., 345 Yokoyama, T., 58 ! Yoneyama, M., 170, 175(36), 176(36), 178(36)

AUTHOR INDEX Yoo, S., 248 York, D. M., 294 York, S. G., 577 Yoshida, E., 614 Yoshikawa, K., 203,206(15) Yoshinari, T., 614, 622(23) Yoshizawa, S., 22(12), 23, 23(29), 24 Young, M. A., 295,296, 296(52; 53), 297(52; 53; 71), 385 Young, P. R., 377 Yu, J., 574 Yu, L., 518 Yui, J., 577

Z Zacharias, W., 298,304(90) Zahler, A. M., 138 Zakian, V. A., 573 Zanatta, N., 262 Zandwijk, N., 589 Zaniewski, E., 504 Zapp, M. L., 22(20; 22), 23(20; 22), 24, 433 Zardecki, C., 283 Zasedatelev, A. S., 194 Zaugg, F., 240, 241(27) Zawadzki, V., 648

Zechel, A., 4, 5(21), 212,216(10), 217 Zegrocka, O., 354 Zelphati, O., 337 Zeman, S. M., 51 Zendegui, J. G , 131 Zhang, J., 306 Zhang, L., 238 Zhang, X., 283 Zhang, X.-Y., 486 Zheng, X.-Y., 238 Zhou, E X., 234, 286(16), 289 Zhou, G., 500 Zhou, H., 387,388(74) Zhou, K., 22(26), 23(26), 24 Zhou, W., 574 Zhu, J., 238,576 Zhu, L., 258 Zhu, N., 203 Zhu, X., 250 Ziaugra, L., 578 Zichi, D. A., 294(48), 295 Zieba, K., 151 Zimmer, C., 24(43), 25,202, 212,271 Zimmer, D. P., 262 Zimmerman, R., 581 Zimnik, O. V., 578,580(55) Zon, G., 11, 14(40), 132, 257,345 Zuker, M., 444 Zwelling, L. A., 388

687

Subject Index

A Acridine DNA intercalation, 379-380 photoreactive azides, 383 4'-(9-Acridinylamino)methanesulfon-manisidide, topoisomerase II interactions azido analog stimulation of DNA strand breaks, 388-389 inhibition, 388 Actinomycin D DNA binding properties, 389-391 kinetics of DNA interactions, 390-391 photoreactive analog, s e e 7-Azidoactinomycin D Actinomycin D DNA binding modes and volume changes, 159-160 nucleosome-drug interactions, 515 Adjacency matrix, s e e DNA duplex stability Adriamycin chemotherapy, 531 energetics of DNA binding, 532-533, 535 formaldehyde-mediated coupling to DNA assay gas chromatography-mass spectrometry, 410--412 N-methyl adriamycin standard synthesis, 409-410 denaturing polyacrylamide gel electrophoresis, 408-409, 412 overview, 407-408 structure of lesion, 408 functional domains, 531 modular drug design using anthracycline-based building blocks, s e e a l s o WP631 combinatorial approach, 539, 541,544 overview, 530, 535-536 rational structure-based design, 536-537, 539

site specificity for DNA, 531 transcription assays of DNA binding, 479, 483 9-Aminoacridine, historical perspective of use, 377-378,381 Antigene, s e e DNA triplex Ascididemin, topoisomerase I inhibition, 613 Association constant equation, 112 noncalorimetric determination for nucleic acid-ligand interactions, 125-126 Atomic force microscopy, s e e Scanning force microscopy 7-Azidoactinomycin D alkali lability of DNA adducts, 392 biophysical properties, 389-390 criteria for ideal photoaffinity analog, 395 DNA binding properties, 389-391 DNA photoreactivity, 391-392 photolysis conditions, 394-395 shuffling hypothesis of binding, 391-392 specificity of binding, 392-393

B Berenil, nuclear magnetic resonance analysis of minor groove binding, 272, 274 BIAcore, s e e Surface plasmon resonance BN80915, topoisomerase I inhibition, 620-622

C Camptothecin, topoisomerase I inhibition, 610, 622 CD, s e e Circular dichroism Cetyltrimethylammonium bromide, McGhee model of DNA melting with ligand, 206 Chromatin~zlrug interactions binding isotherm generation equilibrium dialysis anthracycline antibiotics, 509 dialysis conditions, 508-509 ethidium bromide, 509-510 fluorescence titration, 508

689

690

SUBJECT INDEX

chromatin preparations chicken blood, 504-506 integrity checking, 508 uniform length preparation, 507 circular dichroism analysis, 51 6-517 DNA base distribution in grooves, 503-504 footprinting of reconstituted nucleosomes actinomycin D binding, 515 DNase I footprinting, 511,513-515 gel electrophoresis, 513-514 hydroxyl radical footprinting, 513, 515 netropsin binding, 51 4-515 radiolabeling of DNA, 510 reconstitution with radiolabeled DNA, 510-511 McGhee model of DNA melting with linker histone binding, 209-210 nucleosome purification, 506 prospects for study, 518 sedimentation velocity analysis, 515-516 structure histones, 503 nucleosome, 503 Xenopus sperm chromatin remodeling assays in egg extracts applications, 652-653 DNA replication assays deoxynucleotide incorporation, 650-652 overview, 648 radiolabeled deoxyribonucleotide incorporation, 650 electrophorettic analysis of proteins, 643-644 incubation conditions for drug testing, 641, 643 micrococcal nuclease digestion analysis, 644-645 nuclear assembly assays nuclear lamina assembly, 645,647 nuclear membrane assembly, 645 overview, 645 protein import assay, 648 overview, 640-641 Circular dichroism absorption of polarized light, 69 chromatin~lrug interaction analysis, 516-517 DNA~trug interactions drug dimerization and DNA ineractions, 82 exciton spectra, 97-98

induced signal and structural information, 93, 95-96 modes of binding and spectra interpretation calculations, 82 groove binding, 77, 81 intercalation, 76-77, 79, 81 phosphate binding, 76 signal origin, 69, 75-76 peptide nucleic acid formation, 90, 92 principles, 75-76 RNA-drug interactions, 24-25 triplex DNA~lrug interactions, 346 Cisplatin DNA binding assay, see Polymerase chain reaction, covalent DNA~lrug interaction assay mechanism, 358 volume change studies of DNA adducts, 164-165 Closed circular DNA~trug interactions binding constant and unwinding determination advantages of method, 65~56 calibration of micropipettes, 61 data analysis, 63-64 daunomycin as intercalator, 57-59 ethidium bromide as intercalator, 58, 60, 64 plasmids, 57 principles, 56 reaction conditions, 58-61, 65-66 troubleshooting faint bands, 66-67 resolution of gel, 67 topoisomerase I reaction and incomplete relaxation, 67-68 two-dimensional gel electrophoresis analysis casting, 61-62 image analysis, 62-63 running conditions, 62 staining, 62 linking difference, 51-52, 54, 63-64, 66 nearest neighbor exclusion model, 54-55 topoisomerase I relaxation reactions, 52-53 twist change on ligand removal, 52 unwinding angle, 51-52 ligand binding measurement equations, 54-56 writhe change on ligand removal, 52, 54

691

SUBJECT INDEX Cobalt, DNA binding conformational transition induction, 52O covalent modification, 520 restriction enzyme assay for site determination advantages, 528 agarose gel electrophoresis, 525 AseI linearization, 523-524 BamHI cleavage, 524, 527 binding reaction, 523 DraI leavage, 524, 527 EcoRV cleavage, 524-525,527 endonuclease concentration determination, 525-526 inhibition assessment, 528-528 selection, 522 HindlII cleavage, 524, 527 materials, 522-523 overview, 521 plasmid selection, 522 Pvull cleavage, 524-525,527 site specificity, 520-521 Competition dialysis assay, nucleic acid ligands advantages, 99 applications, 107-108 development, 100-101 dialysis conditions, 104-105 units, 102 interpretation of results binding constant determination, 107 ethidium as intercalator, 105-106 graphical representations, 106-107 nucleic acids concentration determination, 103 preparation, 102-103 quality control, 103-104 principles, 100-102 triplex DNA-drug interactions, 347-348 Cross-linking, DNA interstrand cross-link assays denaturing polyacrylamide gel electrophoresis, 398-399 digestion and chemical composition determination acid hydrolysis, 402-403 controlled depurination, 405-406 enzymatic digestion, 403-405 ethidium bromide assay calculations, 397-398

incubation conditions, 397 principles, 397-397 global structure determination, 406-407 hydroxyl radical footprinting for localization, 399-401 piperidine cleavage at cross-linked residues, 401-402 intrastrand cross-link analysis, 407 monoadduct analysis, 407 types of cross-links, 396 Crothers allosteric model, cooperativity, 16-19, 194

D Daunomycin-DNA interactions chemotherapy, 531 closed circular DNA interactions, 57-59 Crothers allosteric model for cooperativity, 16-19 dinucleotide binding model assumptions, 14-15 binding ratio equation, 12 development, 11-12 free energy equation, 13 unique dinucleotide steps, 12 validation, 12-14 DNA-binding site, 109 energetics of DNA binding, 532-533,535 free energy of ligation, 6 functional domains, 53 I Job plot, 4-5 5-methylcytosine substitution effects on binding, 500 modular drug design using anthracycline-based building blocks, see also WP631 combinatorial approach, 539,541, 544 overview, 530, 535-536 rational structure-based design, 536-537, 539 molecular modeling of (-)-daunorubicin intercalation B-DNA, 309-310 heating and equilibration, 312 hydration, 312 left-handed DNA preference, 308-309 starting model, 311 Z-DNA, 309-312

692

SUBJECT INDEX

neighbor exclusion models assumptions, 6-7 closed form equations, 7-8, 10 cooperativity inclusion, 15-16, 18-19 exclusion parameter, 8-9 fitting software, 8 Friedman-Manning model, 9-10 limitations, 10-11 McGhee-von Hippel model, 6-9 oligonucleotide binding studies, 20--21 Scatchard plot, 6-7 site specificity, 531 titration binding isotherm, 5-6 Diaminopurine, DNA substitution synthesis chemical synthesis, 489 nucleosides, 488-489 polymerase chain reaction, 489-491,502 purification, 491,493 melting temperatures of substituted duplexes, 493-494 protein-binding studies, 494-495,500 minor groove-binding drug studies, 495-496, 499 Diethylpyrocarbonate footprinting, s e e DNA footprinting Differential scanning calorimetry cooperativity analysis, 127-128 high-order DNA melting studies data acqusition, 129-130 data analysis, 130 DNA tetraplex stabilization by cations data acquisition, 144 melting transitions, 146-147 potassium versus sodium, 144-148 sample preparation, 144 thermodynamic parameters, 144-146 thrombin-binding aptamer, 144 sample preparation, 129 instrumentation, 127-129 thermodynamic parameter determination, 127-128 WP631 binding to DNA, 547-549 Dinucleotide binding model assumptions, 14-15 binding ratio equation, 12 development, 11-12 free energy equation, 13 unique dinucleotide steps, 12 validation, 12-14

Dissociation constant equation, 112 noncalorimetric determination for nucleic acid-ligand interactions, 125-126 DNA-binding proteins, s e e Zinc finger engineering DNA cross-linking, s e e Cross-linking, DNA DNA~trug interactions, s e e a l s o s p e c i f i c d r u g s binding modes, 76-77, 234, 466, 529 calorimetry, s e e Differential scanning calorimetry; Isothermal titration calorimetry chromatin, s e e Chromatin-drug interactions closed circular DNA unwinding, s e e Closed circular DNA~lrug interactions competition dialysis, s e e Competition dialysis assay, nucleic acid ligands cross-linking, s e e Cross-linking, DNA drug design criteria, 110-111 McGhee model for melting studies, s e e McGhee model, DNA melting with ligands modified DNA base probing base types, 488,502 groove structure, 486, 500 historical perspective, 485-486 melting temperatures of substituted duplexes, 493-494 minor groove-binding drugs with inosine or diaminopurine substitution, 495-496, 499 protein-binding studies, 494-495,500 synthesis containing inosine or diaminopurine chemical synthesis, 489 nucleosides, 488-489 polymerase chain reaction, 489-491, 502 purification, 491,493 uridine and 5-methylcytosine residues daunomycin binding effects, 500 DNase I sensitivity, 499-500 structural effects, 499-500 synthesis, 499 modular design using anthracycline-based building blocks, s e e a l s o WP631 combinatorial approach, 539, 541,544 overview, 530, 535-536 rational structure-based design, 536--537, 539

SUBJECT INDEX molecular modeling, s e e Molecular modeling, DNA--drug interactions nuclear magnetic resonance, s e e Nuclear magnetic resonance polyintercalators, s e e a l s o 1,4,5,8-Naphthalenetetracarboxylic diimide amino acid design considerations, 556-559 rationale for synthesis, 556 polymerase chain reaction assay, s e e Polymerase chain reaction, covalent DNA~lrug interaction assay scanning force microscopy, s e e Scanning force microscopy spectroscopy, s e e Circular dichroism; Linear dichroism stopped-flow studies, s e e Stopped-flow fluorescence, Hoechst 33258-DNA interactions telomeres, s e e Telomerase; Telomere therapeutic targeting, 109-110, 252, 290-291, 529, 556 transcription assays, s e e Transcription assay, DNA-clrug interactions triplex DNA, s e e DNA triplex volume change, s e e Volume change, nucleic acid-ligand interactions X-ray crystallography, s e e X-ray crystallography, DNA-drug interactions DNA duplex stability graph theory block sequence analysis, 190-192 D N A sequence representation adjacency matrix, 181- 183 Eulerian graph and trails, 184-186 submatrices, 183-184, 186 Eulerian trail finding, 186-187 isothermal sequence number finding decomposition of adjacency matrix, 187-188 different sequence restrictions, 188-189 mathematical nomenclature, 185-186 nearest neighbor model doublet format, 173-175 enthalpy change, 176-180 entropy change, 177 free energy change, 177-180 sequence-dependent end interactions, 171-172 singlet format, 172-175

693

sodium dependence correction, 176-177 stacking interactions, 169-170 transition temperature calculation, 175-176, 178-180 unique interactions, 171-172 rationale for study, 165-166 transition temperature calculation, 170, 175-176, 178-180 isothermal sequences, 166, 181 two-state melting theory dissociation constant, 167 enthalpy of transition, 168 entropy of transition, 168 external degrees of freedom, 168-169 internal degrees of freedom, 167-169 nucleation parameter, 168-169 DNA footprinting diethylpyrocarbonate footprinting echinomycin binding study, 427, 429 incubation conditions, 427 reaction specificity, 427 differential cleavage plots, 429 DNase I footprinting advantages and limitations, 418-4 19 cation dependence, 415 cleavage site, 416 Hoechst 33258 binding study, 419 incubation conditions for drug binding, 416-417 reaction conditions, 417-418 substrate specificity, 415-4 16, 452 WP631 effects on Spl binding, 555 DNase II footprinting, 419, 421 ethidium azide binding sites, 385 fragment selection and preparation, 413-4 15 hydroxyl radical footprinting Fenton chemistry, 399,422 Hoechst 33258 binding study, 424 incubation conditions, 423 interstrand cross-link localization, 399-401 limitations, 423-424 minor groove drug binding studies, 422-423 specificity of cleavage, 422 methidiumpropyl-EDTA-Fe(lI) footprinting cleavage specificity, 452-453 incubation conditions, 425 triplex DNA, 424

694

SUBJECT INDEX

micrococcal nuclease footprinting reaction conditions, 421-422 substrate specificity, 421 nucleosome~trug interactions actinomycin D binding, 515 DNase I footprinting, 511,513-515 gel electrophoresis, 513-514 hydroxyl radical footprinting, 513,515 netropsin binding, 514-515 radiolabeling of DNA, 510 reconstitution with radiolabeled DNA, 510-511 osmium tetroxide footprinting, 426 potassium permanganate footprinting, 425-426 principles, 412-4 13 probe comparison, 413 pyrrole-imidazole polyamide complexes, s e e Pyrrole-imidazole polyamides quantitatie analysis, 429-430 radiolabeling of DNA, 415 triplex DNA, 349, 353 DNA quadruplex differential scanning calorimetry of stabilization by cations data acquisition, 144 melting transitions, 146-147 potassium versus sodium, 144-148 sample preparation, 144 thermodynamic parameters, 144-146 thrombin-binding aptamer, 144 isothermal titration calorimetry sample preparation annealing, 118 buffers, 117 concentration determination, 118-119 melting behavior characterization, 119-120 purity, 117 stabilization by cationic porphyrin buffer conditions, 139 counterion effects, 142 oligonucleotides, 139 stoichiometry of binding, 141-142 thermodynamic parameters, 139-141 molecular modeling ligand specificity, 299 porphyrin intercalation 1:1 complexes, 321-323 2:1 complex, 323

3:1 complex, 322 hydration, 324, 326 starting models, 324 stoichiometry, 320-32 l simulations, 299 starting model generation, 301 spectroscopic studies of cationic stabilization 143 structure, 139, 142-143 telomerase inhibition, 137-138 telomere G-quadruplex targeting anthraquinone analogs, 586, 589 perylene, 590 PIPER, 590 porphyrins, 589 DNase footprinting, s e e DNA footprinting; Pyrrole-imidazole polyamides DNA triplex antigene approach applications, 312-313 improvement with stabilizers, 354 limitations, 313 therapeutic targeting, 130-132 base triples, 341-343 cleaving agent design and uses, 355-356 drug binding studies binding mode studies, 352 buffer conditions, 351-352 calorimetry, 348 circular dichroism, 346 competition dialysis, 347-348 fluorescence spectroscopy, 345-346 footprinting, 349, 353 gel-shift assays, 348-349 linear dichroism, 346-347 nuclear magnetic resonance, 347,352 rational drug design, 353-354 RNA-DNA hybrids, 352-353 sequence selection, 351 surface plasmon resonance, 350-351 thermal denaturation, 343-345,353 types and structures of drugs, 343 viscometric titration, 347 H-DNA stabilization by binding agents, 355 structure, 340-341 isothermal titration calorimetry sample preparation annealing, 118 buffers, 117

SUBJECT INDEX concentration determination, 118-119 melting behavior characterization, 119-120 purity, 117 triplex, 117-118 stabilization by intercalating drugs data acquisition, 134 drug types and specificities, 132-134. 137, 348 thermodynamic parameters. 135-136 titration curves, 134 molecular modeling antigene approach c - m y c P2 promoter targeting, 312, 314-316, 318 hydration, 318-319 RNA as third strand, 313-314 starting model, 318-319 ligand specificity, 298-299 naphtholflavone intercalation, 319-320 peptide nucleic acid complexes, 298 rational drug design, 349-350 starting model generation, 301 nuclear magnetic resonance of inducing oligonucleotides, 278,280 stability with RNA, 313-314 stabilization by proteins and drugs, 341,354 structure, 131-132 Doxorubicin, s e e Adriamycin DSC, s e e Differential scanning calorimetry

E Echinomycin, diethylpyrocarbonate footprinting, 427,429 E c o R I , scanning force microscopy of DNA interactions, 249 Elinafide. inosine or diaminopurine substitution effects on DNA binding, 496. 499 ELISA, s e e Enzyme-linked immunosorbent assay Enthalpy binding enthalpy determination, 3 calorimetric measurement of change, 113-114 nearest neighbor model, 176-180 two-state melting theory. 168 van't Hoffenthalpy, 126-128, 152 Entropy nearest neighbor model, 177 two-state melting theory, 168

695

Enzymeqinked immunosorbent assay, phage assay for zinc finger engineering applications, 595,604 binding conditions and analysis, 605 DNA target preparation, 604-4505 phage preparation, 604 scanning mutagenesis binding assays. 605, 607 Ethidium azides alkali lability of DNA adducts, 385,392 biophysical properties, 383 criteria for ideal photoaffinity analog, 395 DNA binding characteristics of monoazide, 379, 384, 386 footprinting studies and binding sites, 385 photolysis conditions, 394-395 photoreactive analog structures, 382 topoisomerase II-mediated DNA strand cleavage probing mechanism studies, 387-388 stimulation of cleavage reaction, 386-387 Ethidium bromide chromatin~lrug interactions, 509-510 competition dialysis assay. 105-106 DNA cross-linking assay calculations, 397-398 incubation conditions, 397 principles. 397-397 historical perspective of intercalation studies, 380-381 medical use, 38l unwinding assay, s e e Closed circular DNA-drug interactions Eulerian graph, s e e DNA duplex stability

F Factor for inversion stimulation, modified DNA base-binding studies, 494-495 FIS, s e e Factor for inversion stimulation Footprinting, s e e DNA footprinting; RNA footprinting Friedman-Manning model, neighbor exclusion model, 9-10

G Gal repressor, scanning force microscopy of DNA interactions, 249-250

696

SUBJECT INDEX

Gibbs free energy calculation of change, 3, 113 dinucleotide binding model, 13 ligation, 6 nearest neighbor model, 177-180 noncalorimetric determination for nucleic acid-ligand interactions, 125-126

H H-DNA, s e e DNA triplex Heat capacity, calculation of change, 113 Hexaammine cobalt(II1), McGhee model of DNA melting with ligand, 208 Hexadecyltrimethylammonium bromide, McGhee model of DNA melting with ligand, 203-207 High-mobility group proteins, modified DNA base-binding studies, 495 Histone, s e e Chromatin~lrug interactions HIV, s e e Human immunodeficiency virus Hoechst 33258 DNA binding fluorescence titration, 213, 231 mode, 212 stopped-flow studies, s e e Stopped-flow fluorescence, Hoechst 33258-DNA interactions structural studies, 212, 214 footprinting studies DNase I footprinting, 419 hydroxyl radical footprinting, 424 Human immunodeficiency virus integrase, s e e Integrase, HIV-1 packaging region RNA footprinting autoradiogram analysis, 439, 441-442 gel electrophoresis, 437, 439 kinetics of cleavage, 446 paromomycin binding, 433,435,444, 446-449 plot analysis of binding sites, 446-449 RNase I cleavage, 435,437,446, 448 RNase T1 cleavage, 446, 449 RNA synthesis, 435-437 structure, 442-444 therapeutic targeting, 433 pyrrole-imidazole polyamide targeting, 461 structure and derivatives, 212-213

Hydroxyl radical footprinting, footprinting

see

DNA

I Inosine, DNA substitution melting temperatures of substituted duplexes, 493-494 minor groove-binding drug studies, 495-496, 499 protein-binding studies, 494-495,500 synthesis chemical synthesis, 489 nucleosides, 488-489 polymerase chain reaction, 489-491,502 purification, 491,493 Integrase, HIV-1 assays disntegration reaction, 631 DNA-binding assays, 631,633 integration assay, 629 nucleophile selection for phosphodiester cleavage, 63 l oligonucleotide radiolabeling 3~-end, 628 5~-end, 628 reaction conditions, 628-629 strand transfer assay, 630-631 domains, 624 function, 624 recombinant protein purification from Escherichia

coli

amplification, 627 cell harvesting and lysis, 627 nickel affinity chromatography, 627~528 soluble mutant, 626 storage, 628 transformation, 626-627 therapeutic targeting, 624, 626, 633 Isothermal sequence, s e e DNA duplex stability Isothermal titration calorimetry binding constant range limitations, 122 instrumentation, 115-116 nucleic acid--drug interaction analysis advantages, 114 controls, 124 c value, 121 DNA sample preparation annealing, 118 buffers, 117

697

SUBJECT INDEX concentration determination, 118-119 melting behavior characterization, 119-120 purity, 117 tetraplex, 117-118 triplex, 117-118 excess-site method, 124-125 experimental variables, 121-123 interval between injections, 123 ligand concentration determination, 122-123 sample preparation, 120-121 temperature selection, 123 tetraplex DNA stabilization by cationic porphyrin buffer conditions, 139 counterion effects, 142 oligonucleotides, 139 stoichiometry of binding, 141-142 thermodynamic parameters, 139-141 trial experiments, 122 triplex DNA stabilization by intercalating drugs data acquisition, 134 drug types and specificities, 132-134, 137,348 thermodynamic parameters, 135-136 titration curves, 134 ITC, s e e Isothermal titration calorimetry

,J Job plot, daunomycin-DNA interactions, 4-5

K Ku, scanning force microscopy of DNA interactions, 248-249

L Linear dichroism absorption of polarized light, 69 DNA~lrug interactions applications, 68, 82, 92-93 binding rearrangement monitoring, 83, 86 cyanine dye binding, 86, 88 discrimination between intercalation and groove binding, 82-83

drug dimerization and DNA interactions, 82, 97-98 electric transition moment determination, 73-75, 93-95 exciton spectra, 97-98 modes of binding and spectra interpretation, 76-79 orientation of DNA electric field, 73 gel electrophoresis, 73 hydrodynamic flow, 70, 72 polyvinyl alcohol film, 72-73 site-directed linear dichroism of RecA, 88, 90 peptide nucleic acid formation, 90 principles, 70 signal origin, 68-70 triplex DNA-drug interactions, 346--347

M Magnesium, electrostatic interactions with DNA, 156-157 Magnetic suspension densimetry, volume change measurement, 153-154 Mass spectrometry N-methyl adriamycin, gas chromatography-mass spectrometry, 410-412 1,4,5,8-naphthalenetetracarboxylic diimide amino acid, 567 RNA~lrug interaction screening, 25 McGhee model, DNA melting with ligands binding free energy equation, 198 examples cationic lipid ligands applications, 203 cetyltrimethylammonium bromide, 206 hexadecyltrimethylammonium bromide, 203-207 hexaammine cobalt(Ill) as ligand, 208 linker histone binding, 209-210 netropsin and poly[d(AT)]melting, 202-203 helical base pair averages, 197 historical perspective, 193-195 Lifson model foundation, 196-197 ligand binding to both helix and coil, 201 melting temperature binding constant effects, 199 cooperativity effects, 199

698

SUBJECT INDEX

free ligand activity effects, 199 ligand size effects, 198-199 parameter prerequisites and limitations, 201-202, 211 program availability, 211 features, 197-198 titrations with constant total ligand concentration, 199-201 McGhee-von Hippel model cooperativity inclusion, 15-16, 18-19, 194, 247 neighbor exclusion model, 6-9 Melting temperature, s e e DNA duplex stability; McGhee model Methidiumpropyl-EDTA-Fe(ll) footprinting, s e e DNA footprinting; Pyrrole-imidazole polyamides 5-Methylcytosine, D N A substitution daunomycin binding effects, 500 DNase I sensitivity, 499-500 structural effects, 499-500 synthesis, 499 Micrococcal nuclease footprinting, s e e DNA footprinting Molecular modeling, DNA~trug interactions computing power, 290, 292 duplex DNA B-DNA simulation, 294-295 conformational transitions, 295-296 (-)-daunorubicin intercalation B-DNA, 309-310 heating and equilibration, 312 hydration, 312 left-handed DNA preference, 308-309 starting model, 311 Z-DNA, 309-312 hydration, 296-297 ion interactions, 297 peptide nucleic acid complexes, 296 starting model generation, 300-301 thermodynamics of ligand binding, 297-298 evaluation of simulations, 305-306, 308 force field development, 293-294 modular drug design using anthracycline-based building blocks, 537,539 molecular dynamics simulation, 304-305 molecular simulation protocol

equilibrium phase, 307 evaluation, 308 production phase, 307 starting model generation, 306-307 overview, 292 particle mesh Ewald methods, 290, 293-294 quadruplex DNA ligand specificity, 299 porphyrin intercalation 1:1 complexes, 321-323 2:1 complex, 323 3:1 complex, 322 hydration, 324, 326 starting models, 324 stoichiometry, 320-321 simulations, 299 starting model generation, 301 software, 300 starting model generation DNA-intercalator complex, 303-304 DNA-ligand complex, 302-303 ligands, 301-302 minor groove-binding ligands, 304 therapeutic application, 290-292, 312-313 triplex DNA antigene approach c - m y c P2 promoter targeting, 312, 314-316, 318 hydration, 318-319 RNA as third strand, 313-314 starting model, 318-319 ligand specificity, 298-299 naphtholflavone intercalation, 319-320 peptide nucleic acid complexes, 298 starting model generation, 301 validation, 293 visualization, 306 MS, s e e Mass spectrometry

N

1,4,5,8-Naphthalenetetracarboxylic diimide amino acid characterization high-performance liquid chromatography, 566 mass spectrometry, 567 ultraviolet-visible spectroscopy, 566-567 library synthesis, 565-566 polyintercalator design, 558-559

SUBJECT INDEX prospects binding affinity improvement, 568,570 sequence specificity, 570 threading versus nonthreading intercalation, 570 synthesis N-(2-tert-butoxycarbonylaminoethyl)-N'-

(2-carboxyethyl)- 1,4,5,8naphthalenetetracarboxylic diimide, 560 N-2-(Na-9-fluorenylmethoxycarbonyl-N etert-butoxycarbonyl)lysylaminoethylN'-(2-carboxyethyl)- l,4,5,8-

naphthalenetetracarboxylic diimide, 560-561 glycine adduct workup, 561 lysine adduct workup, 561 overview, 559-560 reversed-phase high-performance liquid chromatography, 564 solid-phase synthesis cleavage from resin and side chain deprotection, 562-564 coupling/deprotection cycle, 561-562 NB-506, topoisomerase I inhibition, 614, 622 NDI, see 1,4,5,8-Naphthalenetetracarboxylic diimide amino acid Nearest neighbor model, see DNA duplex stability Neighbor exclusion model assumptions, 6-7 closed circular DNA~lrug interactions, 54-55 closed form equations, 7-8, 10, 54 cooperativity inclusion, 15-16, 18-19 exclusion parameter, 8-9 fitting software, 8 Friedman-Manning model, 9-10 limitations, 10-11 McGhee-von Hippel model, 6-9 oligonucleotide binding studies, 20-21 Netropsin inosine or diaminopurine substitution effects on DNA binding, 496 McGhee model of DNA melting with ligand, 202-203 nucleosome-drug interactions, 514-515 volume changes on binding to model bent sequences, 161-162 NMR, see Nuclear magnetic resonance

699

Nogalamycin, nuclear magnetic resonance of DNA intercalation, 276-278 Nuclear magnetic resonance DNA~lrug interactions bisamidinium compound binding to minor groove berenil, 272,274 hydration analysis, 275-276 pentamidine, 275 propamidine, 272-275 structures, 271 titrations, 271 calculation strategies, 270-271 chemical shifts, 253-256 coupling constants cross-correlation effects, 257-258 Karplus equation, 257 nuclear Overhauser effect error function, 259 sugar conformations, 257-260 field strength, 280 hydration studies nuclear Overhauser effect spectroscopy, 266-268 principles, 265-266 rotating-frame Overhauser enhancement spectroscopy, 266-268 hydrogen bond scalar coupling, 281 instrumentation, 268-270 isotopic labeling, 281 line widths, 255 nogalamycin intercalation, 276-278 nuclear Overhauser effect spectroscopy, 264-265 overview, 252-253 rate constants, 254-256 relaxation times and dynamics 13 value, 260-261 carbon- 13,263 chemical shift anisotropy, 261-263 equations, 260 proton exchange, 263-264 resonance assignment, 253 sensitivity, 269-270 solvent suppression, 269 standards for referencing, 256-257 titrations, 254, 256, 268-269 triplex-forming oligonucleotides, 278,280 tubes, 269 WP631 binding, 549-550

700

SUBJECTINDEX

RNA-drug interactions, 23-24 triplex DNA~lrug interactions, 347,352 Nucleation parameter, two-state melting theory, 168-169 Nucleosome, s e e Chromatin~lrug interactions

O Osmium tetroxide footprinting, footprinting

see

DNA

P PARG, s e e Poly(ADP-ribose)glycohydrolase Paromomycin, s e e Human immunodeficiency virus, packaging region Particle mesh Ewald methods, molecular modeling, 290, 293-294 PCR, s e e Polymerase chain reaction Pentamidine, nuclear magnetic resonance analysis of minor groove binding, 275 Peptide nucleic acid assays for double-stranded DNA binding, 337-338 circular dichroism studies, 90, 92 design for double-stranded DNA binding, 338-339 DNA-binding modes double duplex invasion, 334-335 duplex invasion, 334 overview, 330-331 triplex binding in major groove, 331 triplex invasion, 331-334 invasion complex applications DNA-binding protein inhibition, 336 molecular biology, 337 transcriptional activation, 336 transcriptional arrest, 335-336 linear dichroism studies, 90 molecular modeling duplex DNA complexes, 296 triplex DNA complexes, 298 stability of duplex with DNA, 329-330 structure, 329-330 Perylene, telomere G-quadruplex targeting, 590 Phage display, s e e Zinc finger engineering Photoaffinity labeling of DNA, s e e 4 t -(9-Acridinylamino)methanesulfon-manisidide; 7-Azidoactinomycin D; Ethidium azides

Platinum, DNA modification sites, 519 PNA, s e e Peptide nucleic acid Poly(ADP-ribose)glycohydrolase, inhibition in telomere targeting, 591-592 Polymerase chain reaction modified base DNA synthesis, 489-491,502 radiolabeling of DNA, 4 5 5 Polymerase chain reaction, covalent DNA~lrug interaction assay antibody purification of DNA fragments binding reaction, 374 DNA preparation, 374 overview, 359, 371 protein A-Sepharose beads capture, 375 treatment, 374-375 sensitivity, 371,374 capture of biotinylated products, 366-367 damaging agent treatment cells monolayers, 362-363 suspensions, 362 isolated genomic DNA, 361-362 DNA isolation and digestion, 363-364 first-round amplification, 366 gel electrophoresis of products, 370-37 l ligation of double-stranded linker, 368 ligation-mediated polymerase chain reaction, 358 oligonucleotides end-labeling for third round, 369 melting temperature calculation, 365 phosphorylation, 365-366 sequences, 364-365 optimization, 364 overview of steps, 359-361 polymerase, 364 second-round amplication, 368-369 single-strand ligation polymerase chain reaction, 358-359, 364 terminal deoxynucleotidyltransferase ribo-tailing, 367-368 terminal transferase-dependent polymerase chain reaction, 359, 364 third-round amplication, 369-370 Potassium permanganate footprinting, s e e DNA footprinting Propamidine, nuclear magnetic resonance analysis of minor groove binding, 272-275 Pyrrole-imidazole polyamides

SUBJECT INDEX cell permeability, 450 design, 451 DNA binding specificity, 450 footprinting of DNA complexes association constant determination, 453, 460 DNase I footprinting binding reaction, 457 digestion, 457458 fractional occupancy calculation, 459 gel electrophoresis, 458-459 materials, 453-454 methidiumpropyl-EDTA-Fe(II) footprinting data analysis, 461 ImPy-[3-lmPy--y-ImPy-13-ImPy-[3-Dp characterization, 461,463-464 reaction conditions, 460 overview of probes, 451-453 radiolabeling of DNA gel purification, 456 polymerase chain reaction, 455 restriction digest and fill-in, 454--455 serial dilution of polyamides, 456-457

Q Quadruplex,

see

DNA quadruplex

R RecA, site-directed linear dichroism, 88, 90 Rev-responsive element solid-phase assay for ligand screening, 25-26 surface plasmon resonance conrol, 38-39 RNA~lrug interactions circular dichroism, 24-25 gel shift assays, 25 mass spectrometry screening, 25 nuclear magnetic resonance, 23-24 solid-phase assay for Rev-responsive element ligands, 25-26 surface plasmon resonance, s e e Surface plasmon resonance therapeutic prospects, 22-23 volume change, s e e Volume change, nucleic acid-ligand interactions RNA footprinting binding constant determination, 431432 human immunodeficiency virus packaging region

701

autoradiogram analysis, 439, 441-442 gel electrophoresis, 437,439 kinetics of cleavage, 446 paromomycin binding, 433,435,444, 446-449 plot analysis of binding sites, 446-449 RNase I cleavage, 435,437,446, 448 RNase T1 cleavage, 446, 449 RNA synthesis, 435-437 structure of RNA fragment, 442-444 therapeutic targeting, 433 plot analysis, 431 4 3 3 principles, 431 RNA polymerase DNA~lrug interaction assay, s e e Transcription assay, DNA~lrug interactions scanning force microscopy of DNA interactions, 250-251 RRE, s e e Rev-responsive element

S Scanning force microscopy (SFM) advantages and limitations in nucleic acid imaging, 234-235 components of instrument, 237 drug-DNA complex imaging buffer conditions, 241 groove binding assay, 247-248 immobilization of DNA, 240-242 intercalation assay binding affinity determination, 245 contour length determination, 242-243 exclusion number determination, 244-245 occupied site determination, 245-246 sample preparation, 243-244 Scatchard modeling, 246 site determination, 247 substrates, 240-241 modes for imaging contact mode, 235 intermittent contact mode, 237 noncontact mode, 236 protein-DNA complex imaging E c o R l studies, 249 Gal repressor studies, 249-250 Ku studies, 248-249 prospects, 251

702

SUBJECT INDEX

RNA polymerase studies, 250-251 sample preparation, 248 Trp repressor studies, 250 scanner calibration, 239 tip calibration, 237-239 forces, 239-240 Scatchard plot curvature, s e e Dinucleotide binding model; Neighbor exclusion model daunomycin-DNA interactions, 6-7 DNA ligand limitations, 194 Sedimentation velocity analysis, chromatin~lrug interactions, 515-516 Site-directed mutagenesis, linear dichroism of RecA, 88, 90 SN38, topoisomerase I inhibition, 620-622 Spl, WP631 effects on transcription DNase I footprinting, 555 gel retardation, 554-555 transcription assay, 553-554 SPR, s e e Surface plasmon resonance Stopped-flow fluorescence, Hoechst 33258-DNA interactions association constants, 212, 21 6-217 association rates, 214, 216-217 (A/T)4 sequence binding base-pair roll effects, 226 equilibrium data comparison, 225 minor groove width effects, 226 pseudo-first-order kinetics, 223-225 sequences in study, 223 data analysis, 214-215 derivative binding to isolated AATT site association kinetics, 217, 219 dissociation kinetics, 219-221 effect of m-OH and bis-m-OH phenyl substitutions, 222-223,232 equilibrium data comparison, 221 dissociation rates, 214 DNA types in study, 216 instrumentation, 21 4-215 precautions with dye, 215-216 sample preparation, 215 steady-state titrations, 213, 231 two-step binding models biexponential association kinetics, 228, 233 bimolecular association followed by isomerization, 228-230, 233

data collection, 227-228 rationale, 226 triexponential association kinetics, 231 two single-step associations, 230-231 Surface plasmon resonance advantages over conventional interaction analysis, 26, 50 injection modes, 37-38 principles, 27 RNA~lrug interactions controls, 37-39 data collection program, 39-40 data processing, 40-41 equilibrium analysis, 43-46 experimental design, 27-30 immobilization biotinylated RNA on streptavidin-coated sensor, 34-36 materials, 35 storage of RNA sensor chips, 36--37 strategies, 28-30 kinetic constant determination, 41,45-49 refractive index increment ratios, 42-43 regeneration of surface, 38 resonance units maximum predicted response, 42-43 steady state response, 45-46 RNA biotinylation, 33-34 sample concentrations, 38 sensor chip, streptavidin immobilization coupling reaction, 33 materials, 31-32 overview, 30-31 preconcentration of streptavidin, 32-33 thermodynamic analysis, 50 triplex DNA~lrug interactions, 350-351

T Tankyrase, inhibition in telomere targeting, 591-592 Telomerase antisense oligonucleotide targeting, 579 assays biotinylated primer extension assay, 584 cell extract preparation, 583 primer extension assay, 582-583 telomeric repeat amplification protocol, 582-584

SUBJECT INDEX assembly inhibition, 581 cancer therapy targeting, 577 components, 574 DNA tetraplex inhibition, 137-138 immortalization of cell cultures, 576 knockout mouse phenotype, 575-576 mechanism, 574-575 regulation of expression, 577 reverse transcriptase inhibitors, 578-579 RNA component targeting, 579-580 RNA/DNA hybrid targeting, 580 tumor expression, 137,574-575, 577 Telomere functions, 573 shortening in aging, 573 tandem repeats, 573 targeting cancer therapy rationale, 577 D-loop, 584, 590 t-loop anthraquinone analogs, 586, 589 duplex DNA, 585-586 perylene, 590 PIPER, 590 porphyrins, 589 sequence, 584-585 single-sranded DNA, 586, 589-590 t-loop binding proteins assays, 592 overview, 584-585 poly(ADP-ribose)glycohydrolase, 591-592 tankyrase, 591-592 Tetraplex, s e e DNA quadruplex Topoisomerase I camptothecin inhibition, 610, 622 cleavage reaction cleavage complex and inhibitor targeting, 610 filter-binding assay, 623 inhibitors BN80915, 620-622 indolocarbazoles, 622 SN38, 620-622 religation reaction distinguishing from cleavage ligation reaction, 617,619 oligonucleotide substrates, 617

703

restriction fragment substrates, 619~520 suicide substrates, 619 synthetic oligonucleotides as assay substrates, 6164517 DNA relaxation gel electrophoresis assay, 612-613 inhibitors ascididemin, 613 glycosylated indolocarbazoles, 614 NB-506, 614, 622 reactions, 52-53, 612 inhibitor classes, 61(P4511 sources, 611 unwinding assay, s e e Closed circular DNA~trug interactions Topoisomerase II 4'-(9-acridinylamino)methanesulfon-manisidide azido analog stimulation of DNA strand breaks, 388-389 inhibition of enzyme, 388 cancer role, 385-386 ethidium azide probing mechanism studies, 387-388 stimulation of cleavage reaction, 386-387 intercalator inhibitors, 386 Transcription assay, DNA~lrug interactions Adriamycin binding, 479, 483 bidirectional transcription footprinting applications, 483,485 principles, 482-483 reaction conditions and digestion, 485 dissociation kinetics, 479, 481 drug binding to initiated transcripts, 473, 475 gel elecrophoresis autoradiography, 476-477 phosphorimaging, 479 separation of blocked transcripts, 475-476 principles, 467 promoters ideal characteristics, 467 l a c U V 5 promoter fragment isolation, 468-470 types, 469 sequencing reaction preparation, 472-473 synchronization of transcription initiation complexes, 470472

704

SUBJECT INDEX

WP631 transcription inhibition assay basal transcription assay, 552-553 cell extracts, 552 overview, 550-551 Sp l-activated transcription assay, 553-554 vectors, 551-552 Transition temperature, see DNA duplex stability; McGhee model Triplex, see DNA triplex Trp repressor, scanning force microscopy of DNA interactions, 250

U Uridine, DNA substitution DNase 1 sensitivity, 499-500 structural effects, 499-500 synthesis, 499

V van't Hoff equation, 126 Volume change, nucleic acid-ligand interactions actinomycin D studies, 159-160 apparent molar volume, 154-155 DNA-binding mode effects intercalation and minor groove binding, 159-160 intercalation, 157-158 magnesium electrostatic interactions with DNA, 156-157 minor groove binding, 158-159 overview, 155-156 duplex formation and water uptake, 151 equilibrium constant dependence on pressure, 152 hydration and nucleic acid structure, 150 measurement techniques dilatometry, 152-153 indirect measurement, 153 magnetic suspension densimetry, 153-154 oscillating method, 154 molar volume flexibility of water, 150-151 molecular interpretation, 154-155 netropsin binding to model bent sequences, 161-162 oligomer duplex formation with covalently attached ligands

benzo[a]pyrene diol epoxide adducts, 163-164 cisplatin adducts, 164-165 oligonucleotides for study, 163 RNA intercalation studies, 160-161

W WP631 DNA-binding properties differential scanning calorimetry, 547-549 melting temperature shift, 546 overview, 539, 568 ultraviolet melting studies, 547 viscosity studies, 546 Sp 1 binding effect studies DNase I footprinting, 555 gel retardation, 554-555 structural studies nuclear magnetic resonance, 549-550 X-ray crystallography, 549-550 synthesis, 544-545 transcription inhibition assay basal transcription assay, 552-553 cell extracts, 552 overview, 550-551 Spl-activated transcription assay, 553-554 vectors, 551-552 WP760, melanoma toxicity, 541, 544

X Xenopus egg extract

advantages of system, 634 development of egg, 634 frog colony establishmet and maintenance, 635 preparation activation, 637 centrifugation, 637~638 egg collection, 635,637~638 jelly coat removal, 637 yields, 638 Xenopus sperm chromatin remodeling assays applications, 652-653 DNA replication assays deoxynucleotide incorporation, 6 5 0 ~ 5 2

SUBJECT INDEX overview, 648 radiolabeled deoxyribonucleotide incorporation, 650 electrophorettic analysis of proteins, 643--644 incubation conditions for drug testing, 641, 643 micrococcal nuclease digestion analysis, 644-645 nuclear assembly assays nuclear lamina assembly, 645, 647 nuclear membrane assembly, 645 overview, 645 protein import assay, 648 overview, 640-641 Xenopus sperm chromatin remodeling studies, see Xenopus egg extract nuclei preparation, 639-640 X-ray crystallography, DNA-drug interactions anisotropic diffraction, 288-289 crystal growth precipitants, 282-283 protein crystallization differences, 283 screening parameters, 283-284 toroid analogy, 282 data collection, 284, 288 dynamic range of intensity data, 284-289 overview of steps, 282 structure determination molecular replacement, 289 multiple isomorphous replacement, 289

705

multiple wavelength anomalous diffraction, 289 WP631,549-550

Z Zinc finger engineering bipartite complementary approach advantages, 607 automation, 607 cesium chloride gradient purification of replicative form DNA, 608 cooperating contacts in design, 594-595 coselection of complementary libraries, 603-604 electrocompetent cell preparation, 608-609 library construction, 599-600 materials, 607-608 overview, 595,597 phage enzyme-linked immunosorbent assay applications, 595,604 binding conditions and analysis, 605 DNA target preparation, 604-605 phage preparation, 604 scanning mutagenesis binding assays, 605,607 phage yield estimation, 609 premade libraries, 595 recombination of complementary libraries, 602-603 selection, 601-602 selective polymerase chain reaction, 597, 599 overview, 593 phage display techniques, overview, 594