Hydrogen exchange as a probe of the dynamic structure of DNA

J. Mol. Bid. (1970) 59,297-316

Hydrogen Exchange as a Probe of the Dynamic Structure of DNA I. General Acid-Base Catalysis BMJCE McCoNmmt

AND PETER H. VON HIPPEL

Institute of .&f&c&~ University

Biology and Departnwn.4of Chemisty of Oregon, Eugene, Oregon97403, U.B.A.

(Received31 October1969, and in rev&d

fimn 16February 1970)

In this paper it is shown that positively-charged proton trsnsfer agents can catalyze the exchange-out from native DNA of the hydrogens involved in internucleotide hydrogen-bonding. The negatively-charged catalysts tested did not increase the rate of exchange. The effectiveness of hydronium ion catalysis is decreesed by inc reasing concentrations of sodium ion, while that of hydroxide ion is imxeas4. The measured rates of proton transfer in the presence of various catalysts can be quantitatively accounted for in terms of a Bronsted-type relationship involving the pK, values of both the catalyst and the titratible sites of DNA, plus a superimposed electrostatic effect of the DNA phosphates which decreases the effectiveness of negatively-charged catalysts aud facilitates catalysis by positively-chmrged species. The two types of chemically distinct DNA proton exchange sites (amino and imino groups), do not participate independently in exchange, since the addition of catalysts affects all kinetic classes of hydrogens equally. These results support the hypothesis (prints & von Hippel, 1968) that exchsnge of the inter&&u hydrogens of DNA proceeds wio a local, transiently-open (non-hydrogen bonded) conformational St&e which is maintained (“propped” open) by the presence of two (at acid pH) or no (at alkaline pH) hydrogens at the locus of the N-H . . . N internucleotide hydrogen bond. The over-all rate of the exchange process appears to depend on the product of the equilibrium constant for the formation of open (propped) sites, and the rate constant for the initial transfer of protons to and from the open DNA sites exposed in this process.

1. Introduction Under physiologiosl conditions and temperatures, DNA exists primarily in the doubkhelioal hydrogen-bonded Watson-Crick structure. However this “native” conformation, like all ordered structures, has only finite stability and thus is subject to continuous, though transient, thermshy-induced local fluctuations and distortions. Some of the types of trsnsient distortions (“breathing” modes, or “dynamic” structure) which might be conceived sre illustrsted crudely in F’igure 1, and include: unstecking of adjacent base pairs without hydrogen-bond breakage (Pig. l(b)); hydrogen-bond breaksge without unstacking (Fig. 1(c)) ; snd local meltii (hydrogenbond breakage ati unstacking) accompsnied by psrtisl (Fig. l(d)) or complete t Present eddress: Department of Biochemistry end Biophysios, University of Hawaii, Honolulu, Hawaii 96822, U.S.A. 297

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(4

(e)

FICA 1. Various possible transient local distortions of the native DNA structure. (a) Native structure, staoked, hydrogen bonded and helical. (b) Chains untwisted, bases hydrogen bonded but unstacked. (c) Chains untwisted, bases stacked but not hydrogen bonded. (d) Structure “melted”, brtaes unstacked and not hydrogen bonded, partial ohain separation. (e) Struoture totally “melted”, chains separated.

(Fig. 1(e)) chain separation. Some of these transient conformational modes may have a controlling role in biologically significant processes such as intercalation of dyes and mutagens, and the recognition of specific base sequences by protein repressors, polymerases and perhaps by complementary polynucleotide strands in DNA recombination or DNA-RNA hybrid formation. Thus it has seemed worthwhile to attempt to define some of these conformational forms, and to measure their relative concentrations under a variety of environmental conditions. The usual optiaal methods, e.g. ultraviolet hypochromism and optical rotatory dispersion-circular dichroism, are not well suited to the task since they weigh each conformation only in rough proportion to the ooncentration present. The various distorted species pictured in F’igure 1 doubtless involve muoh less than 1y. of the base pairs at any time under conditions where the native structure predominates, due to the co-operative character of the melting transition. Conversely, methods which sense only the rate of “opening” of base pairs, or the fraction of base pairs open at any time, should be well suited for such studies, Among such methods is hydrogen exchange, which can in principle be used to monitor the rates and extents of any transconformation reaction in DNA which involves the transient making and breaking of the interchain hydrogen bonds (for example, the conformations depicted in Fig. l(o), (d) and (e)). However, before one can put a struatural interpretation to changes in the hydrogen exchange kinetics of DNA, one must know how these kinetics are affected by the various ohemiaal and physioal factors which alter the stability of the native structure as characterized by other, more conventional techniques. In this paper, and that which follows (MoConnell & von Hippel, 1970), as well as in Printz $ von Hippel(1968), we attempt to provide suoh a foundation for structural interpretations of hydrogen exahange measurements on DNA. The interstrand hydrogen-bonded hydrogens of the DNA diuble-helix exchange with solvent (water) hydrogens lo6 to lo7 times more slowly than do the corresponding hydrogens of free nucleotides under the same conditions (e.g. compare Printz & von Hippel, 1965; and Marshall & Grunwald, 1969). A simple model (Linderstrom-Lang, 1955) which incorporates oonsideration of dynamic changes in DNA secondary structure into the kinetic interpretation of exchange, has been used to account for the slow exohange of ordered DNA (van Hippel & pnintz, 1965; Printz t von Hippel, 1968).

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This model postulates a conformational step followed by a chemical reaction, and may be written for any potentially exchangeable site : 4 ka closed z open + exchanged ka

(1)

giving

km =

W, k, + k, + k,

where koas is the observed apparent first-order rate constant for the disappearanoe of isotopic hydrogen from a particular class of exchange sites in the fully-labeled macromolecule. In DNA, kobs is conveniently expressed in units of hydrogens per nucleotide-pair per second. The open form (equation (1)) can be visualized as a structure in which potentially exchangeable hydrogens have been made accessible to solvent and to proton exchange catalysts by some rapidly reversible conformational process, as, for example, a thermally-induced localized separation of complementary strands. If the open form occurs in a small steady-state concentration, i.e. k, 9 k,, equation (2) may be reduaed to one of two limiting kinetic expressions, depending on the relative magnitudes of the rate constant for the closing reaction, k,, and the overall rate constant for the structurally unencumbered chemical exchange process, k,. If ka % kB: kobs = $ k, = Rk,

(3)

koas= k,.

(4)

a

andifk,

Q k,:

Studies of DNA hydrogen exchange are ultimately intended to help define and characterize the transient conformational modes available to DNA under conditions where the native form is the dominant species. To do this within the framework of the soheme represented by equation (1) requires explicit recognition of the assumptions underlying this formulation. These are : (1) that exchange proceeds only through open forms; i.e. the closed form does not exchange directly with solvent and all events associated with the chemical step must follow the opening event; (2) that exchange proceeds primarily through one type of open conformation under particular experimental conditions, and not through other open conformations with which the open form dominating exchange may be in equilibrium; and (3) that during the periods a given site is open, the chemical exchange reaction (k,) proceeds as in simple “fullyopen” model compounds. The mechanisms which lead to equations (3) and (4) differ kinetically, and these differences provide the basis for both the experimental discrimination of these alternatives and for testing the over-all adequacy of equation (1) as a description of the kinetics of DNA hydrogen exchange. The rate-limiting step in the Rk, mechanism (equation (3)) is bimolecular, involving an effective encounter between an open DNA site and a proton transfer agent. Thus the observed rate is a function of the effective concentration of both thebe species. The kl mechanism (equation (4)) involves the monomolecular opening event as the rate-limiting step, with exchange going to completion prior to closing. Accordingly the rate of hydrogen exchange in this scheme

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is independent of both the rate of the chemical exchange process and of the equilibrium concentration of open sites. In this paper we describe the effects of a number of general acid-base catalysts on the observed kinetics of DNA hydrogen exchange. These catalysts were used at concentrations too low to affect the stability of DNA secondary structure as monitored by melting profiles, with the expectation that the results would bear primarily on the chemical step of the exchange process. A lack of response of DNA hydrogen exchange to the addition of catalyst would favor mechanisms of the k, type and in general exclude any mechanism in which a chemical step precedes a rate-limiting conformational reaction. We find that the addition of various catalysts does increase the exchange rate, but that these increases are not explicable in terms of a simple Kk, mechanism (equation (3)). These results, together with previous studies on the effects of changes of pH and salt concentration on the rate of DNA hydrogen exchange (Printz BEvon Hippel, 1968) can be interpreted most simply by adopting the following view of the formation of open (potentially exchangeable) sites on DNA : (1) opening is a locally co-operative process involving sequences of DNA several base pairs in length; (2) only a small fraction of the opening events result in measurable exchange; and (3) the normally short lifetime of these open segments can be increased greatly by an initial proton transfer event mediated by solvent catalysts. Effective exchange proceeds primarily through these long-lived protonated or deprotonated (propped) regions.

2. Materials and Methods In general, hydrogen exchange experiments were conducted as described in earlier publications (Printz BEvon Hippel, 1966,1968). Only the parts of the procedures which modify or supplement the earlier descriptions are discussed here and in the following paper. (a) MateTial8 All reagents were American Chemical Society certified or reagent grade, spectrally pure. Solutions were made with double distilled water and included 0.001 ar-di-sodium EDTA (pH 7) to complex possible contaminating metal ions. Hydrogen exchange buffers for the pH range 6-3 to 7.6 involved sodium cacodylate as the buffer salt, and were adjusted to the required pH at room temperature (Radiometer pH 26 meter). The pH values of the borate buffers were established by calibrating the pH meter against 0.01 n-borax (National

Bureau of Standards primary standard) at room temperature, followed by correction to 0°C (Bates, 1964). Procedures described in the following paper were employed for the characterization of all DNA samples, and for the isolation of DNA from Microcoecus Zy&&&u.s and Eecherichia c&i. Calf thymus DNA (Worthington) was used as supplied after routine checks for protein, RNA and denatured DNA (see following paper),

In the two-column gel filtration procedures for macromolecular hydrogen zz tritium exchange (Englander, 1963; Printz & von Hippel, 1966), the first column is used to separate the bulk of the solvent tritium from the DNA. The labeled DNA is then pooled and incubated under controlled solvent conditions, and fInally samples are passed through a second column to remove the remainder of the solvent tritium plus that which exchangedout from the DNA during the incubation. Normally both columns sre pre-equilibrated with the same solvent. However, since measurement of DNA concentration (Ace,,) and tritium content (scintillation counting) are carried out on effluents from the second column, the solvent emerging with the DNA from the second column cannot contain components which absorb strongly at 260 rnp, which alter the extinction co&lcient of DNA, or wbioh 081188severe quen&ing in &ntillation counting.

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In the present study (see also the following paper, McConnell & von Hippel (1970)) many of the buffer components used do interfere with absorbsnce and radioactivity measurements (e.g. aniline and the halogenated acetates). Moreover the high viscosities of concentrated solutions of destabilizing salts (following paper) greatly decreases flowrates through the columns. Therefore, only the first columns were equilibrated with solvents containing hydrogen exchange catalysts or destabilizing salts, and the second columns were all equilibrated with a common buffer (0.1 ar-NaCl, 0.01 M-sodium cacodylate, pH 76) which established a minimum rate of hydrogen exchsnge for DNA (Printz & von Hippel, 1906). Thus, in addition to separating DNA from the tritium which had exchangedout in the pool, the second columns also served to separate the DNA from the various components of the original solvent and provided a uniform, non-interfering environmenb for data collection. Using this mod&d technique, the DNA in the pooled effluent of the first column loses its tritium to solvent at a rate dictated by the nature of the buffer used to equilibrate the first column. This rate in all csaes wss equal to or greater than that characteristic of the “slow-exchange” medium of the second columns. Since total exchangetimes generally exceeded 400 set in these studies, no corrections were made for the less than 60 set the DNA samples spend in the slow-exchange environment of the second columns. Columns were prepared by pouring a slurry of G25 Sephadex sufficient to form the entire ( N 16 cm) Sephadex bed into siliconized 2-cm columns, adding water almost to the top and inverting several times before allowing the beads to settle. After the ilrst 2 cm had settled onto the coarse scintered glses disk, flow wae begun and continued at maximum hydrostatic head pressure until the bed wae fully packed. This procedure resulted in good separation without the use of the layering techniques previously employed (Printz & von Hippel, 1968). A sharkskin illter paper disk (Schleicher & Schull Co.) was allowed to settle on the Sephadex bed, providing a stable surface for the rapid layering of liquid directly onto the top of the gel. After the initial packing, the columns were used, rinsed and stored in the cold-box (0%) at all times to prevent bubble formation. Void volumes were determined with Blue Dextrsn 2000 (Pharmacia). At times, air-pressure was applied to speed the passage of highly viscous salt solutions through the first column (McConnell & von Hippel, 1970). We found that pressures exceeding 4.6 lb/ins. generally resulted in poor separation of DNA from solvent tritium in these columns. (c) HyoTrogen ezckmge procedures Sonicated DNA (Printz & von Hippel, 1965) samples of total volume 2.6 to 3 ml. and DNA content N 2 mg/ml. were dialyzed into the appropriate exchange buffer and brought to a tritium level of N 20 me/ml. by adding N 60 & of tritiatedwater (1000 me/ml.). Samples were incubated at 0°C for 1 hr ( > 10 half-times) to permit exchange-in to go to completion, and then placed onto a 2 cm x 16 cm G26 Sephadex “6rst” column equilibrated with the same exchange buffer. The first (front) half of the DNA peak wse collected in a graduated tube immersed in ice water. This fraction served as the pool and usually showed a 1OS-fold reduction in solvent tritium. The appropriate elution volume was determined with Blue Dextrsn and checked by counting the pooled eluate. One-ml. samples of the pool were then applied after various exchange-out times to 2 cm x 12 cm “second” columns equilibrated with slow-exchange buffer. Several 1.2~ml. fractions were collected from each column, and the collection times recorded. DNA concentrations were determined for each sample (Aas,,), scattering corrections for the Sephadex particles which had come through the scintered glass disk were applied if necessary (Englander & Epstein, 1967) and O-2 ml. of the samples were transferred to a scintillation vial containing 6 ml. of Bray’s solution (Bray, 1960) for counting in a Packard Tri-Garb scintillation counter. The experiments were conducted under constant refrigeration and forced air circulation in a cold box especially designed for chromatographic hydrogen exchange experiments. Although manipulation of columns and glassware within the box required frequent opening of access ports, constant monitoring of temperature during exchange experiments showed that the temperature generally did not vary by more than f 1 deg. C from 0°C in any part of the box. The internal temperature of each column was measured before and after use, and showed that temperatures within the gel bed also did not vary by more than f 1 deg. C during the experiment.

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3. Results (a) Effects of catalysts The observation that Tris greatly increases the rate of DNA hydrogen exchange, together with the elucidation of the effect of pH on exchange rates (Printz & von Hippel, 1968), suggested that proton donors and acceptors might affect DNA hydrogen exchange. To this end, the catalytic effectiveness of a number of organic acids and bases differing in charge type, pK, and general molecular structure were tested on calf thymus DNA (Figs 2 and 3). In all cases it was shown by melting experiments that these concentrations of potential catalyst had no measurable effect on DNA stability as monitored by melting profile (T,) determinations. The effects of amines, consisting of mixtures of neutral bases and positivelycharged conjugate acids at the exchange pH (~7), are shown in Figure 2. Each experiment was conducted in slow exchange medium, i.e. at pH 7, 0.01 M-cacodylate buffer, and a total cation molarity of approximately0.3, consisting in each case of the

Time (set) FIQ. 2. Tritium exchange-out curves for native calf thymus DNA at 0°C in 0.10 M-N&~ plus various partially postively-oharged general acid-bese catalysts at 0.20 ra total concentration. (0) 0.10 wNeC1 only, pH 7.0; (0) guenidine-HCl, pH 7.0; (A) Tris (M. ~y8odaikticu-9 DNA), pH 7.0; (0) Tris, pH 7.0; (0) triethanolamine, pH 7.0; (a) aniline, pH 7.3 ; (0) aniline, pH 6.8 ; (w) imidctzole, pH 7.0.

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positively-charged conjugate acid present at pH 7 in a total catalyst concentration of O-2 M, and added NaCl. The top curve of Figure 2 shows exchange in the absence of added catalyst. (At this pH and ionic strength the exchange rate is relatively unaffected by moderate changes in pH or ionic strength; see Printz & von Hippel, 1968.) With the exception of aniline (pKi = 5.2), the other compounds shown in Figure 2 exist substantially in the positively-charged (proton donor) form at pH 7, and can be ranked in order of increasing catalytic effectiveness under the conditiona as : imidazole (pK6 = 7.7) a triethanolamine (pKi = 8-4) N Tris (pKi = 8.9) 4 guanidinium chloride (pK = 14). The values of pKi (in parentheses) are calculated for 0°C and ionic strength N O-2 (Bates, 1964). Clearly there exists a qualitative correlation between increasing pKI, and decreasing catalytic effectiveness for these compounds. Aniline is approximately as effective as Tris and triethanolamine at pH 7.3, but appreciably more effective at pH 6.8 (Fig. 2). The correlation between the effectiveness of these organic bases as proton transfer agents (as measured by pKi), and as catalysts of DNA hydrogen exchange, suggests that they exercise their catalytic function by transferring protons to and from the potentially titratible groups on the DNA bases involved in interchain hydrogen bonding. Figure 3 shows that the largely negatively charged proton transfer agents : cacodylate (pKi = 6.2) at various concentrations and acetate (pKi = 4+3), (also phosphate, PK;,, = 7.3, C. W. Lees, personal communication), are not effective DNA hydrogen exchange catalysts at neutral pH.

0.06 0

III

400

I 800

I

I I200

I

11 1600

11 2000

1 2400

Time kec) FIG. 3. Tritium exchenge-out curves for native calf thymus DNA at O”C, pH 7.6 plus various partially uegetively-aherged general aoid-bese oatiysts 8t 0.20 M total conoentration. (0) 0.01 ~-sodium caoodylate; (0) 0.10 af-sodium oaoodylete, (A) 1.0 rd-sodium ceoodylate; (0) 3.4 Bwmdium aoetate.

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(b) Analysis of the catalytic effects These results can be subjected to quantitative analysis. The chemical portion of the exchange-out process (equation (1)) involves the transfer to the solvent of a labeled proton which had been involved in interchain hydrogen bonding prior to the conformational opening reaction. This process is written aa an irreversible reaotion in scheme (l), since the probability of back-exchange of tracer tritium under the exchange-out conditions used is negligible. Of course reversible transfer of unlabeled hydrogens occurs continuously. However, we are interested in the initial proton transfer rate (k,) included in k,, since we can assume that it is at this level that the rate-limiting process affected by catalyst occurs (see Discussion). The initial proton transfer (i.e. protonation of the amino group and endocyclic nitrogens of cytosine and adenine and deprotonation of the ring nitrogens of guanine and thymine) can be divided into pH-dependent and pa-independent processes :

k, = k, + k,,o+ VW+1 + km - W-1

+ Z&-m?[HAPI + XL; [A;1 I 1

(5)

where k, is a pseudo fist-order rate constant in units of hydrogens (per mole nucleotide pair) per second, k0 represents the rate constant for proton transfer to and from DNA sites by pHindependent processes (e.g. concerted water catalysis mechanisms), + and k,,are the second-order rate constants (in I./mole-set) for proton k tzisfer to and from DNA sites by H,O+ and OH-, respectively, and kaAb and kA; are the second-order rate constants for proton transfer to and from DNA site; by other proton donors (positively charged or neutral) and acceptors (neutral or negatively charged) added to the system at concentrations [HAP] and [A:], respectively. (For further discussion of the rates of proton transfer reactions in simple systems see Eigen, 1964.) We have no theoretical basis for estimating k,,, though this term has been shown experimentally to be negligible in hydrogen exchange measurements on free purine (Marshall & Grunwald, 1969), various small amide compounds (e.g. see Berger, Lowenstein & Meiboom, 1959) and a number of proteins and polypeptides (see Hvidt & Nielsen, 1966; Englander, 1967). In any case, we may assume that this term makes a constant contribution in experiments such as those of Figures 2 and 3, in which changes in catalyst type (pKJ represent the only experimental variable. This assumption permits us to write:

-do

= k,,o+ I&O1 + km- [OH-I + i+,$

VW% + 7b.y [@I

63)

where is the expected pseudo first-order rate constant for all the pH-dependent proton transfer reactions. Using this expression, the various catalysts can be compared quantitatively in terms of their ability to increase the observed rate of hydrogen exchange, since rate constants for proton transfer between catalyst and DNA can be estimated from a knowledge of the pKi values of the proton donors and acceptors involved (Eigen, 1964). If the pKX of the donor is less than that of the acceptor, proton transfer to the acceptor will occur with every sterically satisfactory collision and the reaction will proceed at a diffusion-controlled rate (k CY10” I./mole-set). If pKaCCePtOr < pKdonor, proton transfer to the acceptor will be successful only once in lOARKcollisions (QpK = pKdonor -~Kscceptor)> resulting in a forward rate constant of N lOlo x 19-APK I./mole-sec.

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In addition to information about the pKi values of the various proton-transfer agents used, we must assign pKi values to the relevant titratible sites of DNA in order to calculate values of for various solutions. Since these hydrogen exchange measurements were made on native DNA, we utilize pKi values of 3 and 12, determined by titration of native calf thymus DNA at an ionic strength of O-2 and extrapolated to 0°C (Printz & von Hippel, 1968). These pK; values correspond to the protonation of the endocyclic nitrogens N1 of adenine and N, of cytosine (and N, of guanine) at pH 3, and to the protonation of the N, of guanine and the N, of thymine at pH 12 (e.g. see Steiner & Beers, 1961; Marsh, 1968). Although titration studies show no protonation of the exocyclic amino groups, these equilibrium studies do not preclude the possibility that the latter can accept proton8 from solvent as an initial event in proton transfer (see Discussion). Rate constants for the various solution catalysts are then calculated as illustrated in the following example for an O-2 M-imidazole solution at pH 7-O. The solution contains the following catalysts : lo- 7 M-H,O + and 0.167 Irz-imidazole conjugate acid (ImH+) as proton donors (calculated using pKi = 7.7 for imidazole under these conditions) ; and 10T8 M-OH- (K, 11 lo-l5 at OY!) and O-034 M-imidazole base (Im) as proton acceptors. The endocyclic nitrogens of adenine and cytosine (pKk N 3 from titration curves of native DNA) will accept proton8 from H,O+ (pK N -2) at the diffusion-limited rate ( - lOlo I./mole-set) and from ImH+ at (103-‘.‘) (lOlo) =106’3 l./mole-aec (per mole nucleotide pair). By similar reasoning, the endocyclic nitrogens of guanine and thymine (pKi~12) will donate protons to OH- (pK N 16) at m lOlo I./mole-set and to Im at (IO”’ -la ) (lOlo) =106” I./mole set (per mole nucleotide pair). Substitution of these rate constants and the catalyst concentration8 into equation (6) produces a calculated value for of lOlo (lo-‘) + lOlo (lo-*) + 105’3 (0~167)+105’7 (O-034) = 6.16 x IO* hydrogen8 per nucleotide pair per second or log = 4.7. That such a method of prediction based on the ability of donor and acceptor sites on DNA to exchange protons with catalyst is reasonable is demonstrated in Figure 4, where log obtained from equation (6) for all the compound8 tested is plotted against the observed rates of exchange (kobson a logarithmic ordinate). Values of k,,,,s were obtained by drawing tangents to the experimental kinetic curves at a constant value of hydrogens/nucleotide pair remaining (O-2) to obtain half-times, which were then converted to first-order rates by division into O-69. (See further discussion of this procedure in connection with Table 1.) The limits of experimental error (shown in Fig. 4), and the crudeness of some of the estimate8 of the parameters applied in equation (6), do not obscure the obvious relationship between catalyst pKi and the rate of DNA hydrogen exchange. Among the compounds tested only the amines, which can serve as neutral proton acceptor8 and positively-charged proton donors, were found to be effective. The results with aniline show unambiguously that the positively-charged donor species is an important active principle in hydrogen exchange catalysis. This follows because the only appreciable calculated contribution to log for aniline (pKi=5*2) arises from the small amount of conjugate acid present at pH 7. If the overwhelming excess of conjugate base were the only active species, then the value of log due to aniline would be small relative to that due to H,O + and OH-, and no increase in hydrogen exchange rate would be expected. Furthermore, if the assumptions on which these calculations are based were in error, and the conjugate base of aniline were an 20

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Aniline, pH 6.8

/ Triethanolamine 2.0

I

Tris

1.5 I.0

Guanidine -yI$

/

00

Aniline, pH 7.3

Cacodylate gl14

/

dition I I

I 5

3,4 M-acetate I

I 6

-

7

log FIG. 4. The observed rete of tritium-hydrogen exchange (plotted on a logarithmic ordinate) v~8u.n the logarithm of the expected proton transfer rate cahxhted as in text. The observed rates were estimated from tangents to kinetic curves at 0.2 hydrogen/nucleotide pair (see text). Error limits include variation in half-time estimation from tangents as well as experimental error.

effective catalytic species which dominated exchange, then there should also be little or no difference between the rates observed in the presence of aniline at pH 6.8 and 7.3, since the difference in concentration of conjugate base at these two pH values is negligible. Whether the neutral conjugate bases can act as proton acceptors to catalyze exchange is not as easy to determine unambiguously. The fact that the marginal value of log for guanidine is accompanied by no measurable catalysis (Fig. 4) may be a weak indication that neutral proton acceptors are less effective than the positively charged donors (the value of log for guanidine is due entirely to conjugate base). However, an indication that the neutral forms of the amine bases are active catalysts is obtained from the observation that aniline and imidazole do not fall on the same line (Pig. 4). Aniline appears to be a weaker catalyst than imidazole by about I.5 orders of magnitude (aniline at pH 6*8), and at least half an order of magnitude weaker than Tris and triethanolamine (aniline, pH 7.3). Differences of the same magnitude and direction between Bronsted catalytic rate constants for primary amines (e.g. aniline), and heterocyclic and tertiary amines are encountered in studies on reactions catalyzed by the neutral bases (Bell & Trotman-Dickenson, 1949; Trotman-Dickenson, 1949). Steric considerations related to accessibility of the amine nitrogen are not necessary to explain these effects, and actually run counter to the observation that triethanolamine (a tertiary amine carrying bulky substituents) is as good a catalyst as Tris and a better catalyst than aniline (see also the above references for similar observations). The deviations from linearity in Figure 4 are better explained by the fact that the measured pK, of aniline in aqueous solution represents an overestimate of its true basicity, since the primary anilinium ion is stabilized to a greater extent than the secondary or tertiary ammonium ions in a hydrogen-bonding solvent (Pearson & Vogelsong, 1968). Although we measure a given amount of stabilized conjugate acid for aniline from pK, measurements in water, the true ability of the neutral conjugate

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base to remove a proton from substrates other than water (e.g. a DNA nitrogen) would be estimated more closely by pK, measurements in non-hydrogen bonding solvents (Bell & Bales, 1962). These arguments, based on the deviations from linearity seen in Figure 4, suggest that the conjugate base is an active catalyst. If only the conjugate acid were active, quite different deviations would be seen. (c) Electrostatic effects On the other hand Figures 3 and 4 show that neutral proton donors in equilibrium with negatively-charged acceptors are ineffective as catalysts, since the calculated increases in log for cacodylate and acetate arise predominately from the small amounts of conjugate acid present. The negatively-charged conjugate base makes a sensible contribution to log for 1-Or,r-caoodylate (and, of course, for phosphate), proving that this species is also not catalytically effective at the concentrations tested. These results show that the effectiveness of a psrticular hydrogen exchange catalyst depends not only on the pK differences considered in equation (6), but also on the effective charge of the catalyst, with negatively-charged compounds preferentially excluded and positively-charged species preferentially attracted by the negative charge of the backbone phosphates of the DNA molecule. This suggests that at pH values where hydroxide ion catalysis is dominant, increasing the counterion (Na+) concentration should increase the exchange rate, while at pH values where hydronium ion catalysis is dominant, exchange rates should be decreased by increasing the Na+ concentration. Figures 6 and 6 show that these expectations are fully borne out for E. wli DNA, con6rming similar results obtained earlier with calf thymus DNA (Printz & von Hippel, 1968). Figure 4 shows that caoodylate (and phosphate) at high concentrations should greatly increase the exchange rate, so the fact that these species are ineffective shows that the polyanionic exclusion effect due to the DNA phosphates modifies the apparent effectiveness of other negatively-charged catalysts in addition to hydroxide ion.

Timek.ec) FIG. 6. High pH tritium exchange-out curves for native 1. cdi DNA 0.1 ar-N&l, pH 7.6; -A-A--, concentrations of NaCl. -O-O-, --A-A-, 1-O M-N&I, pH 9.4.

at 36°C and various 0.1 wNaC1, pH 9.4;

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of different exchmqe c.!u.m?s

As may be seen in Figures 2, 3, 5 and 6, the hydrogen6 of DNA exchange-out as more than one kinetic class (von Hippel & F’rintz, 1965), since the loas of bound tritium follows a curved rather than a straight-line course in the semilogarithmic representation. The structural or chemical origin of these different classes is not yet clear (Printz & von Hippel, 1968). However an important result of this study is that the various hydrogen exchange catalysts seem to affect all the exchange classes equally within experimental error. This can be demonstrated quantitatively by showing that any curve of Figure 2 can be generated from any other curve by multiplying the time scale (x-axis) by a constant factor. For example, the time required for the Tris curve to reach any particular value of hydrogens/nucleotide pair is just twice that required for the imidazole curve to reach the same value. In Table 1 several of the kinetic curves of Figure 1 are compared in this way, and we may see that the effects of all catalysts on the various parts of the exchange curve are constant, except for the pH 6.8 aniline

O.Olo

’

’ 400

’

I 800

I

I 1200

I

I 1600

I

I 2000

1 2400

Time bed FIG. 6. Low pH tritium exchange-out curves for native E. coli DNA at 36%’ and various conoentrationa of N&I. -O-O--, 0.10 M-NaCl, pH 76; -A-A-, 0.10~.NaCl, pH 6.3; (-A-A0.01 M-NaCl, pH 6.3. TABLE

1

Catalyst effects on various parts of the exduznge-out curvest Ratio Hydrogens/nucleotide pair remaining

1.0 0.4 0.2 0.1 0.06

Guanidine Tris 2.53 2.8 23 -

Tria Imidazole

of times (+ 0.1) Aniline

2.03 2.0$ 2*1$ 2.1 2.0

t Taken from data of Fig. 2. $ Taken from a smooth curve drawn between the experimental gens/nucleotide pair)0 value of 2.4.

Tris (pH 6.3) 2.03 1.91 1.9 1.8 1.6

Aniline (pH 7.3) Imidazole 2.01 2.1$ 2.13 2.1 2.0

points and the assumed (hydro-

HYDROGENEXCHANGE

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309

curve in which some deviations appear at very low hydrogens/nuoleotide pair values. It should be pointed out specifically that the actual experimental points which support this conclusion for some of the more effective catalysts apply only to the last 10 to 16% of the exchangeable hydrogens, since the rest exchange-out too rapidly to be followed by the techniques used here (see Fig. 2). However it is clear from their absence that exchange of the remaining hydrogens is speeded to at least a comparable extent by the added catalysts. Thus there is no indication that hydrogen-exchange catalysts affect the kinetic classes of each kinetic curve to a different extent, or that there is a gross change in the number of different kinetic classes following catalyst addition. This observation constitutes justification for the manner in which the rates of hydrogen exchange were compared and exhibited in the ordinate of Figure 4 (see were obtained from tangents drawn at the same value of above). Values of k,,,,:obs hyd.rogens/nucleotide pair (0.2) for each kinetic curve. This level was chosen so that comparison could be made in regions containing actual experimental points. On the other hand, if the tangents had been taken at any other value of hydrogens]nucleotide pair, the rate constants would be different, but would hold the same order and produce the same ratios for the different kinetic curves. In addition, this result further justifies our treatment of the exchange phenomenon in terms of only one DNA proton acceptor (pKl, N 3) and one DNA proton donor (pK1 N 12) (equation (6) ), and provides support for a structural (conformational) role for the proton transfer process (see Discussion). 4. Discussion The results described here, together with earlier findings, have shown that hydrogen exohange involves proton transfer to and from DNA nitrogens as a rate-limiting step. Furthermore the rate of this proton transfer, which depends primarily on the basicity of the catalysts, is also strongly affected by electrostatic interactions between the catalysts and the negatively charged sugar-phosphate backbone chains of DNA?. Positively charged proton donors are the best catalysts. Comparable (equation (6)) neutral proton acceptors are somewhat less effective, while no catalysis haa been observed with negatively charged species (except OH-) at the concentrations tested. St&c considerations pertaining to the molecular structure of the catalyst are not required to explain the observed deviations from the predictions of equation (6) (Fig. 4), though such effects may also play a minor role. Differences in DNA base composition do not alter the effectiveness of a given catalyst (e.g. Fig. 2 shows that Tris has approximately equal effects on M. ZysodeiWcw DNA (72 mole y0 G + C) and calf thymus DNA (42 mole + G + C); see also McConnell & von Hippel, 1970). All these observations can be viewed as effects on a simple rate-limiting chemical (proton transfer) step (k3 in equation (l)), occurring subsequent to a structural (opening) event. On the other hand, there are a number of facts which are not explicable on the basis of a simple catalytic effect on individual open exchange sites. It has been shown that catalysis of the various chemically-different exchangeable interchain hydrogens of DNA does not occur independently, since the exchange rate of all are increased equally by a particular catalyst (Table 1). After equilibration with tritiated water, the groups t Bimiler chcqe-dependent &e&s have been observed on the rates of ma&ion of charged compounds with groupa loceted within charged micelles (e.g. see Winters & Crunwald, 1966). We are grataful to Dr Orunwald for pointing this out to us.

310

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on DNA that carry the isotope for times measurable by the Sephadex procedure are the amino groups of cytosine, adenine and guanine, and the endocyclic nitrogens (imino groups) of guanine (N,) and thymine (Nc). Protonation of the former, and deprotonation of the latter, could serve as the chemical (proton transfer) step that initiates exchange. The pKI, values involved make deprotonation of these amino groups, or additional protonation of these imino groups, a very unlikely initial chemical event. If exchange of the various DNA hydrogens proceeded independently after an initial opening event, with k, (equation (1)) as an independent variable made up of a sum of chemical rates for each exchangeable hydrogen (equation (5)); then a catalyst operating as a proton donor would increase the exchange rate of the amino groups only, and a proton acceptor catalyst would affect only the endocyclic imino groups. For a given catalyst the separate contributions of its conjugate species to can be quite different (equation (6)), especially in the case of aniline and Tris at pH near neutrality. This unequal effect is not observed; all exchangeable hydrogens sxe catalyzed by all the catalysts tested. Furthermore it is the endocyclic nitrogens of adenine (NJ and cytosine (N3), which carry 1~)hydrogens at neutral pH (Fig. 7) and therefore cannot be directly involved in the exchange process, that appear to serve as proton acceptors in the initial proton transfer, since it is the pKi values of these groups that provide the correlation seen in Figure 4 for the particularly effective positively charged proton donors?. Thus the initial proton transfer step mediated by solvent components (Ha0 + or OH-) or by added catalyst appears to affect the rate of exchange of other sites not directly involved in the transfer by affecting the secondary structure of the DNA helix. A simple model that accounts for these facts can be formulated by postulating the continual occurrence of local short-lived, thermally-induced disruptions of the DNA helix, involving concerted rupture (opening) of the interbase hydrogen bonds, (e.g. see Fig. 1). (Discussion of possible opening mechanisms and of the size of the open units produced by these distortions will be deferred to the following paper.) The lifetime of these disrupted regions is presumably short, but during the time a given base pair is open, the groups that arenormallyinvolved ininterchain hydrogen-bonding, and thus inaccessible to solvent, are available for reaction with proton transfer agents. And if proton transfer to or from the DNA sites does take place during the open phase, we postulate that this interferes with the normal reversal of the opening event, prolonging the existence of the disrupted region and thus allowing more time for exchange at vi&al sites. Therefore we propose the existence of a “propped conformation” (Printz t von Hippel, 1968) which permits one proton transfer event to catalyze exchange at several chemical sites. This mechanism can be more specifically visualized with the aid of Figure 7. The initial proton transfer event involves the protonation or deprotonation of the imino nitrogens normally involved in hydrogen bonding, resulting in the A-T or G-C base pair structures depicted in Figure 7(a), (b), (e) and (f). Under these conditions the base pair involved (and perhaps neighboring base pairs as well, if the co-operative unit of t From equilibrium data it is known that these nitrogens have a greater b&city than do the amino groups of the unprotonuted base+ and thus that they should aocept protons more readily than the amino groups (Dove, Wellaoe & Davidson, 1969; MassouliB, Michelson & Pochon, 1966; Marsh, 1968). Furthermore since the endocyclic nitrogens of guenine (N,) and thymine (N3) are clearly involved in the catalytic process as proton donors, the same event which exposes these groups to proton transfer agents in the solvent should also make available the opposite nitrogen of edenine and cytosine.

Zr /

I-Z

F

0 k=

-2

=\ Z-X

\I z: < \

0

-

z

=5-t

=v= Y

/

-

B. MCCONNELL

312

AND

P. H.

VON

HIPPEL

exchange is larger than a single base pair) would be unable to close properly until the distribution of hydrogens required for normal hydrogen bonding had been restored (Fig. 7(c) and (d)). This state, in which a potential N-H.. . .N hydrogen bond contains 2 hydrogens (Fig. 7(a) and (b)), or none (Fig. 7(e) and (f)) has been called the “propped” state, and during this phase exchange of all the potentially exchangeable interchain hydrogens oan proceed by the usual proton transfer mechanisms. This proposal can be explicitly formulated as follows: ” kl +

(7)

where C is a closed (non-exchangeable) site, 0 is an open site in which probably more than one base pair is made accessible to solvent, P represents a proton transfer agent (Ha0 + , OH-, HA&, A’), 0 l represents the intermediate or propped state containing at least one of the exposed base pairs with two or no hydrogens between the endocyclic nitrogens, and Ex is the exchanged site. The rate constants k,, k, and k, indicate the reactions that apply to equation (1) to delineate the extension of the original model. The rate constants k’; and kg are included, since now we have no operational measure of 0 (see below).? The rate constants for the formation of O* from C (ky), and for its conversion to Ex (kj) or reversal to C (kg), can be estimated from equation (6). At 0°C and ,,J= 0.2, assuming k, (equation (5)) to be negligible and H,O+ and OH- to be the only effective proton transfer species present, both k; and kj (equation (7)) are effectively equal to (k) (see Results) and thus lo3 hydrogens/nucleotide pair-sec. On the other hand, the reversal of the propped state (k!!) will proceed primarily by protonation or deprotonation of water. Thus for reprotonation: k!! = k, o + [H,O+]

+ k,,, [H,O] = lOlo (10-“5) + 1O1O(10-5) (55) = 102’5 + 55 x 106

N 5 x lo6 hydrogens/nucleotide

pair-set;

for deprotonation: k; = k,,-

[OH-I + b,o [Ha01 = 1010 (10-7’5) + 1O’O(10-5) (55)

N 5 x lo6 hydrogens/nucleotide

pair-sec.

In both cases the contribution of H,O+ and OH- to the rate of reversal of the “propped” state to the open form is negligible compared to that of water, and k! & kj or k’;. (The above estimate of k, represents a maximum value, because it is calculated t For completeness it should be stated that an alternative pathway to O* could be written: C + P ~)t C * ~fc 0 l , where C, P and 0 * have the same significance as ebove and C l is a closed site beering a protonated or deprotonated exocyclic amino group. Thii site might then open and rearrange to 0 l by some combination of amino-imino and keto-enol tautomerization of the base pair. Such a pathway is not definitively excluded by the availeble evidence, though we consider it unlikely largely because of the unfavourable pKk values which probably apply to the initial titretion event.

HYDROGEN

EXCHANGE

OF DNA.

I

313

for a single protonated or deprotonated site. The closing rate constant for the entire propped region may be smeller since it involves a number of potentially protonatible or deprotonatible sites which can react with the solvent and continue to hold open the region despite the reversal of the original proton&ion or deprotonation event; see below.) Since k; and ki are small compared to k;, the observed rate of hydrogen exchange is proportional to the product of kj end the equilibrium concentration of O-. Then if we assume that general acid-base catalysts operate on the exchange rate V&Zthe same kinetic psthway, they can affect the observed rate by increasing k3 (equation (6)) and rtlso perhaps by increasing [O*]. The effective general acid-base catalytic species studied here were mostly positively-charged proton donors, which, as indicated above, operate primarily by protonating the endocyclic nitrogens of sdenine and cytosine which carry no hydrogens at neutral pH and thus are not directly involved in the chemical exchange step characterized by the rate constant kj. Thus these proton transfer agents can only affect the over-all rate of exchange (as represented by equation (7)) by increasing the concentration of Of (“propping”). Such a structural explanation is in keeping with the observation that these agents increase the rates of exchange of all the classes of exchangeable hydrogens equally. Other catalysts might also have selective effects on the chemical exchange rates of certain groups (kb) and thus for these catalysts the effect on the various exchange groups might not be constant. How, then, do these exchange catalysts increase [Of]? This concentration is a function only of pH and of the pKi of the DNA sites subjected to proton&ion. At constant pH and pKi general acid-base catalysts can increase the rate of formation of 0 f from 0 (increase ki), but the principle of microscopic reversibility would appear to require that the reverse process be catalyzed equally, so [0*] should remain constant. This, however, may not be the case since the entire propped region comprises more than a single exchsngeable site and during the time required for the reversal of the state of protonation of the initial site, other exchangeable sites in the propped region may have become titrated. Thus the exchange rates appear to be a function of the secondary structure of DNA, and it is for this reason that the description of the intermediate as the propped configuration, rather than merely as a titrated site, provides a number of interpretive sdvantages. First, as a specific means to account for the observed interdependence of exchange sites, propping is the most simply conceived description from the stsndpoint of our existing knowledge of DNA structure and proton transfer. It is useful in this regard to mention that the propped configuration can be visualized in two ways: either a single base pair is made accessible in the reaction C e 0, so that proton transfer at the exchange site must be followed by a structural change that exposes more sites: i.e. Of is not the same as 0 structually, but represents a more extended structural disruption. Alternatively, we may view the formation of 0 as a reaction that initially exposes several exchange sites to solvent, so that the formation of Of by proton transfer to or from one of these sites merely increases the lifetime of this conformation: i.e. 0 and Of are structuctlly the same. For the present we favor the second view, both because it is simpler, and because of considerations described in the following paper (McConnell & von Hippel, 1970). A second advantage of describing the intermediate as a propped configuration is that it provides a reasonable mechanism for varying the concentration of intermediate (extent of titration of DNA) at constant pH. Changes in proton binding et constant

314

B. MCCONNELL

AND

P. H.

VON

HIPPEL

pH are associated with changes in the pKA of the tit&able groups. DNA helical structure appears to be the chief factor governing the required pK1; changes, since it is well known that the structural stabilization of the double helix provided by the normal base pairing patterns of DNA is responsible for the greater sharpness and more extreme pH values of the acid and alkaline limbs of the native DNA titration curve relative to the corresponding features of the denatured curve. By increasing the lifetime of the open configuration (through propping), a shift towards the “denatured” pKA value is produced which is associated with a greater extent of titration, i.e. greater proton&ion of acceptor groups and deprotonation of donor groups at neutral pH. Thus the initial proton&ion exposes a number of other sites which can continue to maintain the propped state, and thus maintain the Of site, even when the initial proton&ion or deprotonation event has been reversed. It is this structural (locally co-operative) aspect of the interaction of DNA sites with proton transfer agents which in this model leads to an increase in [0*] with catalyst basicity (Fig. 4). Equation (7) divides the chemical reaction labeled k, in equation (1) into two consecutive steps: an initial proton transfer to or from an exchange site within the “open” region, followed by exchange of the sites conformationally affected by this initial process. Since we hypothesize that exchange proceeds primarily from propped sites, the increased exchange rates accompanying the addition of catalyst are due to the increased probability of converting an 0 site to a propped (0 *) site in response to the larger number (per unit time) of initial proton transfer events resulting from the presence of the added catalyst (equation (6) ). Thus in this view the equilibrium concentration of open sites (0) does not depend on the concentration and effectiveness of the proton transfer agents present, but the fraction of 0 sites which are converted into 0 l sites and thus become exchangeable, does. We conclude, then, that the over-all rate of exchange (kobs)=K’kj, (where K’ is defined as 0*/C or K’;/ki). On this basis, the ratio of propped sites to closed sites can be calculated. Of/C or KY/K: is equal to K,,,/k~ (all in units of hydrogens/nucleotide pair-see) which is approximately 10d3/103 = 10e6 at neutral pH, TV~0.2, 0°C and with no added catalyst. Thus approximately one nucleotide pair in lo6 is “propped” under these minimal exchange rate conditions. It should be pointed out explicitly that the remaining assumption in this calculation is that kj has the magnitude expected for an exchangeable hydrogen located on a mononucleotide free in solution. In a recent preliminary communication, Hanson (Biophys. Sot. Meeting Feb. 1969. Abstr SAM-D9) has reported that by using special rapid methods of column manipulation he can detect a faster kinetic class than the internucleotide hydrogen-bonded hydrogena observed in these and previous studies. He suggests that this da88 may correspond to the non-hydrogen-bonded amino hydrogens of adenine, guanine end cytosine, which if conflrmed would mean these hydrogen8 exchange much more slowly in the environment of the grooves of native DNA than in free solution. Carrying the argument further this would also suggest that the internucleotide hydrogen-bonded hydrogena transiently exposed to exchange in the propped region might also exchange more slowly than expected, and thus the value of kj used above would be too large and the calculated value of K’ therefore too small. Thus the estimate of [0 *] cited above should be viewed as an upper limit value. Some additional conclusions can be drawn from the data presented here. The Bronsted relationship for general acid-base catalysis can be written: log k, = y log K, + G, where y and G are constants peculiar to the reaction system and k, and K,

HYDROGENEXCHANGE

OFDNA.1

315

are the catalytic rate and catalyst dissociation constants respectively. Although we cannot separate the contributions of different catalyses or those of their acid or basic forms, an analogous expression can be written for OIIJ data: log kc = y log

+ B.

(8) The slope y is estimated from a line drawn closer to the imidazole point than to aniline (see Fig. 4), since we assume that the pK, of imidazole is a closer estimate of true basicity than that obtained for aniline. The constant y is then approximately 0.5, which is not very different from corresponding slopes determined for the base-catalyzed deuterium exchange of model amides (Klotz & Frank, 1965) or base-catalyzed hydrolysis reactions (Bell & Trotman-Dickenson, 1949). Moreover, this is the expected value of the slope if both the initial proton transfer rtnd its reversal from the titrated form of the catalyst are affected (Frost, & Pearson, 1961). Thus, there is nothing chemically unusual about the catalysis of DNA hydrogen exchange by amine proton transfer agents which would invite interpretations involving direct effects of the catalysts on the secondary structure of DNA other than through proton transfer mechanisms modified by electrostatic effects. While the addition of catalyst can increase exchange by increasing [0*] (and kj), the concentration of 0 is maintained presumably by thermal processes sensitive to other features of the solvent environment. The observation that DNA hydrogen exchange may show a larger temperature dependence than expected from chemical considerations alone (Printz & von Hippel, 1968) suggests that the concentration of 0 (and of Of) might be controlled by factors that govern the stability of DNA against thermal melting. It has been verified in this study that the addition of 0.2 M-CatdySt does not lower the T, of DNA. On the other hand, in postulating a propped intermediate, we predict that the addition of hydrogen exchange catalyst should increase the over-all disruption of the helix, since the lifetime of the open conformation is extended. Thus the manipulation of compositional and environmental factors related to the thermal stability of DNA might influence the kinetics of exchange. That this is not the case at temperatures well below T, is demonstrated in the following paper, in which we consider the effects of the T,-related variables of base composition and destabilizing salts on the kinetics of DNA hydrogen exchange. This investigation was supported by U.S. Public Health Service Research Grants AM-03412, AM-12216 and GM-15792 and Research Career Program Award GM-K3-5479 to one of us (P. H. vonH.) and Post-Doctoral Fellowship 6-F2-GM-29,835 to the other author (B. M.).

REFERENCES Bates, R. G. (1964). Determiwtion of pH: Theory and Practice, 2nd ed. New York: Wiley. Bell, R. P. BEBales, J. W. (1962). J. Chem. Sot. p. 1618. Bell, R. P. & Trotman-Dickenson, A. F. (1949). J. Chem. Sot. p. 1288. Berger, A., Lowenstein, A. & Meiboom, S. (1969). J. Amer. Chem. Sot. 81, 62. Bray, G. A. (1960). Amdyt. Biochem. 1, 279. Dove, W. F., Wallace, F. A. t Davidson, N. (1959). Biochem. Biophys. Rea. Comm. 1, 312. Eigen, M. (1964). Angew. Chem. 8, 1. Englander, S. W. (1963). Biochemdy, 2, 798. Englander, S. W. (1967). In BioZogicaZ Macromolecules, ed. by G. Fasman, vol. I, p, 339. New York: Marcel Dekker. Englander, S. W. & Epstein, H. T. (1967). Arch Biochem. Biophys. 68, 144.

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Frost, A. A. & Pearson, R. G. (1901). Kinetics and Me&anti, 2nd edn, p. 218. New York: Wiley. von Hippel, P. H. & Prima, M. P. (1906). Fed. Proc. 24, 1468. Hvidt, A. & Nielsen, S. 0. (1906). Advanc. Protein Chem. 21, 288. Klotz, I. M. & Frank, B. H. (1966). J. Amer. Chem. Sot. 87, 2721. Linderstrom-Lang, K. U. (1965). Chem. Sot. Spec. Publ. 2, 1. Marsh, R. E. (1968). In Structural Chemistry and Molecular Biology, ed. by A. Rich & N. Davidson, p. 484. San Francisco: Freemen. Marshall, T. H. & Grnnwald, E. (1969). J. Amer. Chem. Sot. 91, 4641. Massoulie, J., Michelson, A. M. & Pochon, F. (1966). Biochim. biophye. Acta, 114, 16. McConnell, B. & von Hippel, P. H. (1970). J. Mol. Biol. 50, 317. Pearson, R. G. & Vogelsong, D. C. (1958). J. Amer. Chem. Sot. 80, 1038. Printz, M. P. & von Hippel, P. H. (1966). Proc. Nut. Acud. Sk., Wwh. 53, 363. Printz, M. P. & von Hippel, P. H. (1968). Biochembtry, 7, 3194. Steiner, R. F. & Beers, R. F., Jr. (1961). PoZynudeotidea. Amsterdam: Elsevier. Trotman-Dickenson, A. F. (1949). J. Chem. Sot. p. 1293. Winters, L. J. BEGrunwald, E. (1966). J. Amer. Chem. Sot. 87, 4608.