DNA structural variations in the E. coli tyrT promoter

Cell, Vol. 37, 491-502,

June 1984, Copyright

0 1984 by MI’I

0092-8674/84/060491-l

2 $02.00/O

DNA Structural Variations in the E. coli ty17 Promoter Horace R. Drew and Andrew A. Travers Laboratory of Molecular Biology, Medical Research Council Centre Hills Road Cambridge, CB2 2QH, England

Summary X-ray studies have established that the structure of a right-handed, Watson-Crick double helix can change from place to place along its length as a function of base sequence. The base pairs transmit deformations out to the phosphate backbone, where they can then be recognized by proteins and other DNA-binding reagents. Here we have examined at single-bond resolution the interactions of three commonly used nucleases (DNAase I, DNAase II, and copper-phenanthroline) with a DNA of natural origin, the 160 bp fyr7 promoter. All three of these reagents seem sensitive to DNA backbone geometry rather than base sequence per se. Their sequence-dependent patterns of cleavage provide evidence for structural polymorphism of several sorts: global variation in helix groove width, global variation in radial asymmetry, and local variation in phosphate accessibility. These findings explain how sequence zones of a certain base composition, or purine-pyrimidine asymmetry, can influence the recognition of DNA by protein molecules. Introduction In order to understand the workings of promoters, enhances, and other DNA control regions, it will surely be necessary to learn something about their structure in solution. Unusual forms of DNA such as left-handed “z” and hairpins have been the focus of recent attention yet these forms are relatively unstable and have not been found in vivo (Courey and Wang, 1983; Hill and Stellar, 1983). Here we will be concerned with structural variations in normal right-handed DNA. X-ray and solution measurements have established that the conformation of a stable, right-handed double helix can change from place to place along its length with details of the base sequence (Dickerson and Drew, 1981; Lomonossoff et al., 1981; Calladine, 1982; Fratini et al., 1982; Klug et al., 1982; Dickerson, 1983a, 198313; Shakked et al., 1983). The problem is to determine, for any given sequence, what these structural variations might be. In a previous study, it was shown that DNA nucleases can exhibit well-defined specificities for certain kinds of DNA structure (Drew, 1984). DNAase I, for example, when presented with a hairpin loop and double helix of identical sequence, can distinguish one from the other according to how sugar-phosphate backbones are arranged; it cuts at a reasonable rate only where phosphates from opposing

strands face one another across the minor groove. DNAase Ii, on the other hand, cuts certain strands regardless of their helical environment and seems less sensitive to groove dimensions. A third reagent, copper-phenanthroline, probably seeks a base-pair step suitable for intercalation and so favors double helix over single strand. All three reagents produce sequence-dependent patterns of cleavage in duplex DNA without any obvious preference for a particular base or base-pair step. We now present a detailed analysis of nuclease cleavages in a duplex DNA of natural origin, the 160 bp E. coli tyrT promoter, and show that these patterns may be most easily interpreted in terms of sequence-induced variation in the structure of right-handed Watson-Crick DNA. Two sorts of variation in global helix structure, along with a single kind of variation in local helix structure, account for the digestion specificities of DNAase I, DNAase II, and copper-phenanthroline in their entirety. The results provide fresh insight into a wide range of biologically relevant phenomena, a few of which are discussed in detail. Results The tyrT promoter directs the synthesis of a major species of tyrosine tRNA, and bears a close resemblance to other E. coli promoters for transfer RNA and ribosomal RNA. It extends far upstream from the transcription start point, approximately 100 bp in the 5’ direction (Lamond and Travers, 1983; Travers et al., 1983). Nuclease Comparisons at 37% in H20 Shown in Figure 1, a-c are: (a) the nucleotide sequence of tyrT DNA and various time courses of digestion for its (b) Watson and (c) Crick strands (antisense and sense, respectively). In the numbering scheme used here, the transcription start point lies at position 100 on the Watson strand while the primary RNA polymerase binding site covers nucleotides 65-9.5. Symbols “W*C” and “WC*” indicate which of the two strands bears a radioactive 3’ end label; “DI,” “DII,” and “Cu” identify samples that were exposed for 1, 5, and 30 min to the reagents DNAase I, DNAase II, and copper-phenanthroline. All reactions were performed at 37°C in water, buffered at a suitable pH and ionic composition for each nuclease. The patterns of digestion shown in Figures 1 b and 1c are complex, but upon careful inspection they begin to yield information of an interpretable nature. First of all, certain regions of double helix, 5-10 bp in length, seem very resistant to nuclease attack of any sort. Several distinct “gaps” appear in DNAase I and copper-phenanthroline gel lanes near positions 30, 50, and 75 of the sequence on both strands; identical gaps are present, if less noticeable, in the DNAase II digest. Gaps 1 and 2 at positions 30 and 50 map out (Figure la) into several ATrich sequence blocks upstream of the RNA polymerase binding site, while gap 3 at position 75 lies in a thoroughly GC-rich tract well within the polymerase binding site.

Cell 492

(I

- Tyrosine

t RNA

Promoter

from

E. Coli

Figure 1. Cleavage

of tyrT DNA

(a) Sequence of the tyrosine RNA promoter from E. coli and its surrounding regions. The tRNA gene begins at position 100 in this numbering. Canonical RNA polymerase recognition signals TTTACA and TATGATG lie upstream at positions 68 and 90, respectively. (b, c) Digestion of the tyrosine tRNA promoter duplex by DNAase I, DNAase II, and copper-phenanthroline at 37°C in water. Both (b) Watson and (c) Crick strands were radioactively labeled at their 3’ ends prior to the addition of nuclease. Time in minutes (0, 1, 5, 30) after the addition of nuclease is shown at the top of each gel lane. Lanes labeled “G” are dimethylsulfate-piperidine markers specific for guanine. Numbers on the right-hand side of each gel refer to the numbering scheme shown in (a) for the Watson strand tyrT-W*C

tyrT-WC*

b

yggjjy

PPZ[

WP 3 [

Other regions of double helix seem much more susceptible to attack by DNAase II than by DNAase I or copperphenanthroline, although on one strand only. Several prominent “sites” of DNAase II cleavage lie at position 100 on the Crick strand (Figure 1c) and positions 115 and 130 on the Watson (Figure 1 b). Site 1 on the Crick strand at 100 maps out across from a long run of C and T bases, site 2 on the Watson at 115 spans a region of A mixed with G, while site 3 on the Watson at 130 covers a run of A ending in G. Within any gel lane, nearest-neighbor phosphates seem more uniformly susceptible to attack by copper-phenanthroline than by DNAase I or DNAase II. Around position 40 in Figure lc, for example, step-to-step variations in copper-phenanthroline band intensity are fairly smooth and regular, whereas stepwise variations in DNAase I and DNAase II intensity are somewhat abrupt and precipitous. These distinctions may surely be attributed to a difference in chemical mechanism. Copper-phenanthroline performs free-radical attack on DNA sugars, while DNAase I and DNAase II perform nucleophilic attack on DNA phosphates. The free-radical species, once formed, does not remain bound to the copper ion but may diffuse locally to react

with any of several nearby sugars (Pope et al., 1982; Uesugi et al., 1982; Drew, submitted). The protein-bound species, in contrast, may be targeted at a single bond: 03’-P in the case of DNAase I, or 05’.P for DNAase II (Bernardi, 1971; Laskowski, 1971). It would appear, then, that the modulation of cutting frequency along any strand in duplex DNA is influenced by at least two factors: the recognition or avoidance of particular sequence zones (“sites,” “gaps”) and the chemistry of bond cleavage. Quantitative Analyses of Nuclease Probability In order to gain a more accurate impression of nuclease specificities, the intensities from selected gel lanes in Figures 1b and 1c were measured by densitometry and converted to numerical probabilities of cleavage (Lutter, 1978). A typical result is shown in Figure 2a, where the intensities from DNAase I lanes, labeled “Dl,5” in Figure 1 b and “DI,i” in Figure Ic, have been converted to probabilities and superimposed on the known sequence. Individual data points are drawn as small black dots, which lie a variable distance above or below the baseline, depending upon the value of In (probability) at that point. The smooth

DNA Structure 493

a

ln (Probability

1 2II ‘..

b

in Solution

.

ln( Probability

of

cleavage)

: DNAase

:

:._

of

cleavage)

: DNAase

1

. _’

I average

actual

(...,.),

average(-)

Figure 2. DNAase

I 12

.:

t-1,

Copper-phenanthroline

6

line is a running three-bond average, calculated by averaging the logarithmic probability at each bond with those of its two nearest neighbors. Presented below are probability plots for all three nucleases. DNAase I, DNAase II, and copper-phenanthroline, under standard conditions of 37°C in water. The DNAase I digestion pattern may be usefully regarded as a combination of global and local components; the global function gives a good estimate of true double-stranded cutting rate, while the local does not. DNAase I is an enzyme that cuts just one strand at a time (Lomonossoff et al., 1981; Drew, submitted), yet probabilities of cleavage on opposite strands in Figure 2a are quite closely correlated. The averaged curves for strands Watson and Crick are approximate mirror images as drawn, and would match even more closely if the lower Crick strand were shifted laterally, a few bonds to the right, relative to the Watson. Gaps 1, 2, and 3 at positions 30, 50, and 75 provide the clearest evidence for a cross-strand relationship, while dips in the curve at positions 13-15, 97-100, 104-I 08, and 130-l 32 offer additional examples. Probabilities of cleavage at individual bonds vary more substantially in going across any base-pair step, and do not give a good estimate of overall DNAase I affinity. Near position 40, for example, individual values of probability on the lower strand are much more variable than on the upper. In a physical sense, the averaged function could describe the ability of DNAase I to bind the double helix.

(---I

I Speclflcltles

(a) Plot of actual versus averaged probabilities of cleavage for DNAase I at 37°C in water. Actual data points are drawn as small black dots giving the measured probability of cleavage for each bond, expressed In logarithmic units so as to approximate a relative measure of the activation energy for bond cleavage. Vertical scales (0, 2, 4. 6) on the left and right-hand sides of the duplex are in units of In (probability). The smooth line is a three-bond running average calculated by averaging the value of In at any bond with those of Its two nearest neighbors on the same strand. Radioactive nucleotides are circled. (b) Comparison of the DNAase I running average with the unaveraged copper-phenanthroline pattern, both at 37°C In water. The sequence on the lower strand has been shifted three bonds to the right so that nuclease curves for upper and lower strands may line up vertically.

6

Deviations from this curve up or down to the actual data points would then describe the ability of DNAase .I to cut a bond once it is bound. Given that the precut binding may be symbolized by an equilibrium constant K,, and the cutting step by a rate constant k,, then the probability of cleavage at bond i will be a product of terms: P, = (WI In(P,) = In(K)) + In(k,) We will refer to the slowly changing running average in Figure 2a as the DNAase I “global” function since it is associated with overall helix geometry. Single-bond differences from the running average also define a pattern that we will call the DNAase I “local” function, since it can change dramatically from one base step to the next. The DNAase I global function resembles the copperphenanthroline pattern in two respects: both are displaced by three bonds in the 3 ’ direction for cuts on one strand relative to cuts on the opposite strand and both exhibit minima at long runs of A/T or G/C. Compared in Figure 2b are the two sets of data. The lower strand of the sequence has been shifted three bonds to the right, along with all of its probability values, so that the staggered nature of cuts may be more easily visualized. It was mentioned above that the DNAase I global function moves up and down in unison on opposite strands, but with a slight displacement of the lower strand to the left. This can now be seen clearly in Figure 2b by noting the

Cell 494

o

Ln( Probability

of cleavage)

: DNAase

1

actual

(&-) , average

(“‘,,x,‘,,)

Figure 3. DNAase

II Specificities

(a) Plot of actual versus averaged probabilities of cleavage for DNAase II at 37°C in water. Radioactive nucleotides are circled. (b) Plot of local variation from the running average for DNAase II at 37% in water, superimposed on an identical plot for DNAase I. The vertical scale is in units of difference probability (+2, 0, -2) relative to the average curve. Three asterisks mark places where DNAase I and DNAase II local peaks coincide.

b

Local

variation

from

average

: DNAase

I

(-),

DNAase

II

close correspondence of the solid line directly above and below the diagonally skewed duplex throughout its length. Slight discrepancies appear at positions 52, 67, 83, and 124, but these are due in part to use of a three-bond average in calculating the global function. A five-bond average would smooth out some of the fluctuations. The copper-phenanthroline data are already smooth and require no averaging. As represented by the dashed line, they match (in a qualitative sense) even more closely above and below the skewed duplex than do the DNAase I data. Only at position 67 does a significant discrepancy occur, and in this case the dashed line for the upper strand dips while the line for the lower strand remains constant. Both sets of curves exhibit minima wherever there are long runs of A/T or G/C. Sequences AAAATTA, ATTTTT, GCGGCGCG, and GCGCCCCGC, the first three of which were labeled as gaps in Figure 2a, are particularly resistant to DNAase I and copper-phenanthroline; TTTAAT at position 11 and TAAAAA at position 127 are not so resistant at 37°C but become resistant at lower temperatures (data below). It seems wise to postpone any structural interpretation until all the facts are gathered, but it would be misleading not to note at this point that phosphates facing one another across the minor groove are displaced by three bonds in the 3’ direction relative to any base-pair step, just as are DNAase I global and copper-phenanthroline probabilities of cleavage. Where base pairs inside the

(,,‘,‘,,‘I

minor groove happen to be all A/T or all G/C, these probabilities fall towards zero on both strands.

The DNAase II digestion pattern is predominantly one-stranded, but still seems to require a separation into global and local components; global maxima lie at runs of A and G, with minima at runs of T and C. Shown in Figure 3a are logarithmic probabilities of cleavage for DNAase II at 37°C in water. The smooth curve is once again a running three-bond average of the actual data points, which are drawn as small black dots. If the DNAase II pattern were not a combination of global and local components, then the smooth curve would not exhibit such distinct maxima and minima. As it is, maxima on one strand (e.g., positions 100, 115, and 130) are always accompanied by minima on the other, very nearly across the same base-pair step. Probabilities of cleavage seem to be anticorrelated in a fashion suggestive of some asymmetry in double-helical structure or enzyme mechanism. Sites 1, 2, and 3 in Figure 3a are the most strongly cut clusters of bonds in this 160 bp molecule. Their sequences are all rich in A plus G: GGGAAGCGGGG, AAGGGAGCAGG, and AAAGCA. Three other moderately well-cut sites are also purine-rich: GTAAAGTG, AGCGGCG, and ACGGGG. Runs of A alone, such as those occurring at positions 30 and 50, are not sufficient to generate a DNAase II site; runs of A plus G, or of G

FsN$ Structure

in Solution

alone, seem to be necessary. The pyrimidine-rich partner strands to all six sites just mentioned are themselves exceptionally resistant to cleavage. Bonds in the sequence of C plus T opposite site 1, for example, are cut at a uniformly slow rate. A direct comparison of the DNAase II global function with the DNAase I global function is not shown, but the reader may scrutinize Figures 2a and 3a to learn that there is little resemblance between the two. Only at runs of G, where DNAase II cuts well on the G strand while DNAase I cuts poorly on both strands (positions 75, 98, 106, and 140), does a systematic relationship exist. The DNAase II local function typically exhibits a value opposite that of DNAase I locally. Neither is easily predictable from base sequence. As mentioned earlier, the local function at any bond is simply the difference between the actual probability at that point and the running three-bond average. Added together, global and local functions reconstruct the true pattern. Shown in Figure 3b are the local functions for DNAase I (solid line) and DNAase II (broken line), superimposed for purposes of comparison. Vertical axes are marked in units of (+2, 0, -2), which describe logarithmic distances from individual data points to the averaged curve. Measurable peaks and valleys show up less frequently in the DNAase II local function than in its DNAase I counterpart, simply because DNAase II cuts the double helix in a more selective fashion. Where it does appear, however, the DNAase II local function almost always has a value opposed to that of DNAase I locally. For example, in the sequence stretching from position 1 IO to position 135 on the upper strand (beginning AAGGG and ending GCATT) there are eight DNAase II peaks, six of which correspond to DNAase I minima. in the same region there are five major DNAase I peaks, four of which are DNAase II minima and a fifth which is DNAase II-neutral. At only three bonds in the entire sequence do strong DNAase I and DNAase II peaks coincide (positions 89, 94, and 113 on the lower strand). No hard and fast rules can be derived from these data regarding sequence versus rate. In the sequence of Figure 3b, for example, are 11 different CG sequences. Six of these (positions 17, 22, 35, 58, and 78) are cut strongly by DNAase I at the bond immediately following CG on each strand; four others (positions 73, 100, 107, and 141) are cut strongly at the CG bond itself; while the last (position 76) seems to be an intermediate case. Pyrimidinepurine steps CG, CA, TG, and TA do tend to be less susceptible to DNAase II attack than their nearest neighbors, examples being steps CA at position 118 (AGCAGG) and 133 (AGCATT), but other steps such as TA (position 33) and TG (position 90) are cut rather decisively. It is not difficult to imagine, at this juncture, how several simple kinds of variation in DNA structure could account for the specificities of all three nucleases in their entirety. DNAase I global and copper-phenanthroline rates vary inphase across the minor groove, DNAase II global rates

Cl

tyrT-WC?

DNAase

I

Figure 4. Gel Patterns, Temperature-DMSO Digestion of the tyrr promoter duplex by (a) DNAase I on the Crick strand or (b) DNAase II on the Watson strand, as a function of temperature or percent dimethylsulfoxide (DMSO) at 37°C. All reactions were performed at constant enzyme concentration but variable times in order to produce closely Identical extents of digestion. In (a) minutes of reaction were 120, 20, 3, 3, 4, and 8 at 4’, 23”, 37”, 50”, 60”, and 7O”C, respectively. Also in (a), minutes of reactions were 3, 3, 3, 4, and 8 for 0, 10, 20, 30, and 40% DMSO. In (b) times in minutes were 60, 6, 2, 0.5, 0.5, and 0.3 at 40 23”, 37”, 50”, 60”, and 70°C. Also in (b), times were 4, 1.5, 1.0, 1.0, and 0.5 for 0, 10, 20, 30, and 40% DMSO.

vary out-of-phase across any base-pair step, while local rates for DNAase I and DNAase II are typically out of phase at any phosphate. Before proceeding further in the direction of interpretation, however, let us present a final line of evidence on the subject. Changes Induced by Temperature and DMSO We know from the data of Depew and Wang (1975) and Lee et al. (1981) that both temperature and DMSO unwind DNA to a significant extent, the effect of DMSO being more pronounced. If the enzymes DNAase I and DNAase II are truly responsive to variations in DNA double-helical geometry, then their patterns of cleavage should change as the helix unwinds. Shown in Figure 4a are single time points for digestion of the Crick strand by DNAase I, over the temperature range 4°-70”C and in the presence of 0%-40% DMSO at 37°C. Five sets of bands, indicated by arrows on the right-hand side of the gel, increase somewhat in intensity by 70°C and dramatically so in 40%

Cell 496

a

b

DNAase

DNAase

I

II

averages:

averages

4” in H,O

: 4O in H,O

(-),

37”

(IIIIIIII) , 37’ in

in 40 % DMSO

40 % DMSO

DMSO. Similar increases are observed at identical positions on the Watson strand (gel not shown, data below). The effects of DMSO are so much more dramatic than those of temperature alone that it would seem unjustified to treat these data just in terms of helix unwinding; other sorts of structural variation may be equally important. Shown in Figure 4b are temperature-DMSO profiles of the DNAase II pattern for the Watson strand. Three sets of bands increase in going from water to 30%-40% DMSO, one of which is enhanced several thousand fold. Whereas the DNAase II sites in water show up on one strand only, these DMSO-induced sites appear on both strands (Crick strand gel not shown, data below). Once again, the DMSO effect far exceeds that of temperature alone. The DNAase I global function depends strongly on environmental factors. Minima near runs of A/T grow wider at 4°C but disappear at high temperature or in aqueous DMSO. Figure 2a supplies the reference curve from DNAase I at 37°C in water. In Figure 5a are shown DNAase I averaged probabilities at low temperature (4°C in water, dark line) and in aqueous DMSO (37°C in 40%, light line). At 4°C every long stretch of A/T pairs become a DNAase I global minimum. The two AT-rich gaps at positions 30 and 50 are even more extensive than at 37’C, covering just less than a full turn of helix. Three new gaps appear at positions 66, 82, and 129, the sequences of which are ACTTTACA, TCATTTGAT, and AGTAAAAA, respectively. These three predominantly AT-rich regions return to a normal cutting

(-)

(t,,m,c,,u )

Figure 5. Environmental

Effects

(a) Effect of temperature and percent DMSO on the DNAase I running average. Five stretches of A and T (marked with arrows) are cut much more rapidly at 37°C in 40% DMSO than at 4°C in water. Because bands in the 40% DMSO track of Figure 4a grow considerably more circular near the bottom of the gel lane, densitometer tracing from top to bottom gave an overestimate of their integrated intenatres. In order to correct for this, two scale factors were applied to the averages in 40% DMSO: bonds from 74-140 on the upper strand, and IO-74 on the lower, were plotted 1 .O log unit lower than the others. (b) Effect of temperature and percent DMSO on the DNAase II running average. Three stretches of alternating purinepyrimidrne (marked with arrows) are cut better at 37°C in 40% DMSO than at any temperature in water.

rate with an increase in temperature from 4” to 37°C but the first two runs of pure A/T persist as gaps to 70°C in water. They begin to be cut well only at high concentrations of DMSO. Two GC-rich minima at positions 75 and 98 remain basically unaffected by any environmental change. Other sequences that are an equal part A/T and G/C, particularly positions 60, 118, and 136, are not affected by temperature but go down somewhat in DMSO. The DNAase II global function also depends strongly on environmental factors. In aqueous DMSO new sites of affinity appear on both strands at runs of G and C, or of A and T. Figure 3a gives the reference curve for DNAase II at 37°C in water. In Figure 5b are shown DNAase II averaged probabilities for 4°C in water (thick line) and 37°C in DMSO (thin line). Temperature has little effect on the DNAase II pattern, but in 40% DMSO several new sites turn up on both strands at positions 76, 91, and 99. Sequences in these regions are GCGGCGCGT, TATGATGC, and CCCGC, respectively. Some of the new sites were weak maxima in water and have just grown stronger, such as the lower strand of position 91. Others were hardly cut in water and have increased by many logarithmic units, examples being both strands of position 76 and the upper strand of position 99. The three AG-rich sites discussed above (positions 100, 115, and 130) are still cut with about the same probabilities and remain one-stranded. No single-strand Sl nuclease-sensitive sites were detectable anywhere within the range O%-40% DMSO, so it

DNA Structure 497

in Solution

Table 1. Summary

CGCGAATTCGCG, center

9A

Fratini et al., 1982

-8”

11 A

Arnott et al., 1983

-6”

12A

Arnott and Hukins,

0”

12A

Fratini et at., 1982

GGCCGGCC

+I20

15A

Wang et al., 1982a

GGTATACC

+130

16A

Shakked

w?AdT)

“B-DNA CGCGAATTCGCG. ends

Figure 6. Extremes

tilt

= 0”

of Groove Wrdth in Right-Handed

tilt

= + 20’

DNA

Sugar-phosphate chains are drawn as ribbons, phosphates as dots on the ribbon, and base parrs as thin horizontal lines. On the left, the minor groove (double-ended arrow) is very narrow, only 9 A from phosphates on one strandto phosphates on the other. Since the radius of a phosphate group is 2.9 A, this leaves only 9.0 - 5.8 = 3.2 A empty space between strands; base pairs tilt by -10” to frt into such a narrow groove. In the center, phosphates Ire 12 A apart and base pairs stack perpendicular to the helix axis. On the right, minor-groove phosphates are separated by a nearmaximal 17 A, and base parrs tilt by +20” to make their connections. For idealized helices such as these, the sum of minor and major grooves remains constant at the product of pitch P times the cosine of pitch angle r, typically 30 cos (28”) = 26 A.

is highly probable that these variations in DNAase I and DNAase II rate are due entirely to changes in the shape of a right-handed double helix. Singleton et al. (1983) have shown that Sl sites always accompany the formation of left-handed DNA in plasmids. The variations cannot readily be attributed to a change in protein structure or hydrogenbonded specificity, since cutting rates in one part of the double helix often remain unperturbed while those in an adjoining region are strongly affected. Discussion The digestion data presented here for three n&eases, DNAase I, DNAase II, and copper-phenanthroline, may most easily be interpreted in terms of sequence-induced variation in the structure of right-handed, Watson-Crick DNA. Recent X-ray analyses of DNA structure in crystals and fibers have provided several examples of helical flexibility that account for the solution results in precise detail. Two Kinds of Global Helix Motion ‘Two different sorts of overall backbone motion have been observed in crystals, or proposed from fiber data as a reasonable stereochemical possibility. The first of these is shown in Figure 6. As the mean planes of base pairs (thin lines) tilt from the horizontal in a positive or negative sense, their attached sugar-phosphate chains (dotted ribbons) follow suit, moving up on one strand and down on the other. This motion opens up the minor groove from a width of 12 to 17A for +20” tilt (Figure 6, right) or closes it to 9A for -10” tilt (Figure 6, left). The spacing across the major groove, although not explicitly diagrammed, changes in a

Reference

-8”

PoWA).

= - lo0

Minor Groove

Tilt

Structure

tilt

of Recent DNA X-Ray Work

GGGGCCCC

-+I30

-16A

1972

et al., 1983

M. McCall and 0. Kennard, unpublished results

“/Y-DNA

+20”

17A

Arnott and Hukrns, 1972

r(GCG)TATACGC

+20”

17A

Wang et al., 1982b

Tilt is the angle of inclination of the pyrimidine CG-purine C8 vector to an Imaginary helix axis; minor-groove wrdth IS the closet approach of phosphorous atoms on the minor-groove side of a base pair. Values of tilt-groove width not available from original reports were kindly provided by Dr. R. E. Dickerson prior to their publication in Structures of Biological Macromolecules and Assemblres. (A. McPherson, ed.). The defined-sequence entries refer to single-crystal X-ray determrnatrons at 2.0-2.5 A resolution, for which the posrtions of atoms have been determined in space; the entries poly(dA). poly(dT), “B-DNA, and “A-DNA refer to fiber X-ray analyses in which the coordinates of a single nucleotide (or base pair) have been refined against the fiber data, and then extrapolated into an infinite helix.

reverse fashion: closed on the right for positive tilt, and open on the left for negative tilt. Not enough X-ray structures have been done to be sure of the relationship between base sequence and helix groove width; yet by looking at the existing structures in detail (Table 1) one can get an idea of how other sequences will behave. Runs of A/l, not containing the step TA, should exhibit a very narrow minor groove with base pairs tilted by - 10”. Only two short stretches of A/T are available from singlecrystal X-ray work, one with the step TA (Shakked et al., 1983) and one without (Fratini et al., 1982). Several authors have commented, however, that runs of A/T should have a narrow minor groove on account of their large “propeller twist” (angle of twist between A and T bases in the same base pair). Steps AA, AT, and TT can twist in a propellerlike fashion about their long axes to produce a narrow groove and negative tilt, while step TA is restricted in this motion by the close contact of A rings from opposing strands (Calladine, 1982; Fratini et al., 1982; Dickerson, 1983a, 1983b). Runs of G/C, containing few or no steps of the type GC, should exhibit a wide minor groove with base pairs tilted by +lO”. Just two DNA X-ray examples of GC-rich runs are available, both with wide minor grooves and few steps GC (Wang et al., 1982a; M. McCall and 0. Kennard, unpublished results), yet there is good reason to believe that this

Cell 498

sequence-related behavior may be rather general. Steps GG, CG, and CC are all forced open toward the minor groove by a close contact of bases from opposing strands, much as in the case of step TA above. Step GC, in contrast, opens weakly toward the major groove or not at all (roll-angle data from Dickerson, 1983a). We cannot be certain that these tentative sequencestructure rules are correct; they are merely the best descriptions currently available. For the dodecamer CGCGAATTCGCG, it has been reliably established by high-resolution NMR that the structure in solution is closely related to that in the crystal (Pate1 et al., 1982) but a more definite statement cannot yet be made. NMR methods are quite sensitive to local conformation, but are rather insensitive to global helix structure (Hare et al., 1983). In other respects, the AATT center of this sequence exhibits the same base-pair tilt (-8”) as does poly(dA). poly(dT) in the fiber (Table 1) and the same helical repeat (9.9 bp per turn) as poly(dA). poly(dT) in solution (Klug et al., 1982; Rhodes, 1982). For the octamers GGCCGGCC and GGGGCCCC, we can only note that both of these sequences twist at an approximate 10.8 bp per turn in the crystal, close to the 10.7 of poly(dG). poly(dC) in solution. DNAase I and copper-phenanthroline digestion patterns provide good evidence for sequence-induced groovewidth variation in solution. Both reagents cut at approximately equivalent rates for phosphates facing one another across the minor groove, parallel to the double-ended arrows shown in Figure 6. Furthermore, rates of attack fail off dramatically when base pairs inside the minor groove are either all A/T or G/C. Rates at A/T, in particular, fall off more noticeably when the step TA is absent (e.g., AAAATT, ATTTTT) than when it is present (TTTAAT, TAAAAA). In view of the sequence-structure correlations cited above, it would appear that DNAase I and copperphenanthroline cut rapidly when the width of the a7inor groove is at or near some intermediate value of 12 A, but more slowly as the spacing becomes either larger or smaller. While this paper was in the process of review, Keene and Elgin (1984) independently suggested that base-pair tilt might control copper-phenanthroline specificity. They did not, however, discuss the structural relationship between tilt and groove width, nor did they discuss why different sequences should prefer to adopt different values of tilt-groove width. We are indebted to Dr. C. R. Calladine for much of our understanding in this regard. A second kind of global helix motion, rather different from the first, is pictured in Figures 7a and 7b. As adjacent phosphate groups stretch out, like the folds of a pleated curtain, from 5.8 to 7.0 A (Figure 7a), the helical radius of a sugar-phosphate chain may increase from 8 to IO A (Figure 7b). One can imagine that the dotted ribbon lies on the surface of an expanding cylinder; its winding rate and height do not change. The important point here is that, since base pairs are fairly rigid and of fixed length, if one strand of a double helix shifts inward the helix axis

Figure 7. Extremes

of Strand Shape in Right-Handed

DNA

(a) the end-to-endOlength of any stacked single strand can vary from approximately 5.8 A (left) to 7.0 A (right) per phosphate; (b) hence, the helical radius of a strand can shrink to 8 A (left) or expand to IO A (riahtl without unduly straining the sugar-phosphate backbone. In a double helix, radii r, and r2 of strands 1 and 2 are related by the formula: r, cos a, + r2 cos (YP= D, where a, and CQ are the angles by which a level line drawn from strand 1 to strand 2 subtends the helix axis, and D is the level distance between strands, typically 17.5 A. I..,

I

(Figure 7b, left) then its partner strand must shift outward away from the axis (Figure 7b, right). This is currently only a model of DNA motion, not yet proved rigorously by single-crystal X-ray analysis but stereochemically plausible. It was proposed by Arnott et al. (1983) to explain certain anomalous features of the poly(dA). poly(dT) fiber X-ray pattern, and would seem to be compatible with other poly(purine). poly(pyrimidine) fiber results (Zimmerman and Pheiffer, 1981). In the Arnott model, it is the A strand that moves inward to a radius of 8 A and the T strand that moves out to IO A; there may be a general tendency, especially in regions of purinepyrimidine asymmetry, for purine bases to move inward so they can stack directly on top of one another up the helix axis (as in Figure 7b, left). DNAase II digestion patterns provide good evidence for sequence-induced radial asymmetry in solution. In water, as opposed to aqueous DMSO, rates of attack are anticorrelated across any base-pair step, with global maxima at runs of A plus G (or G alone) and global minima at runs of T plus C. The simplest interpretation might be that DNAase II cuts rapidly whenever the same-strand phosphate spacing is at or near an optimal 5.8 A, but more slowly as the spacing increases to 6.4 or 7.0 A. Thus runs of purine tend to be cut rapidly, while runs of pyrmidine tend to be cut not at all. Runs of purine containing only A, rather than a combination of A plus G, seem curiously resistant to DNAase II cleavage, yet possibly this is because the enzyme cannot gain access to its substrate. In the X-ray model for poly(dA). poly(dT) (emott et al., 1983) strands A and T are spaced by only 11 A across the minor

DNA Structure 499

in Solution

groove, and it seems conceivable that the two strands could protect one another from attack. Two groups have recently found that long runs of G, or A plus G, exhibit a marked sensitivity to single-strandspecific Sl nuclease in supercoiled plasmids (Nickel and Felsenfeld, 1983; Schon et al., 1983). At least 15-16 consecutive purine residues are a prerequisite for this sort of Sl sensitivity; runs of 10-l 1 are not sufficient. If we accept that there is a strong tendency for purine bases to stack among themselves within any strand, at the expense of stacking attraction between strands, then it seems reasonable that the double helix should come apart rather easily in these regions. Structural Changes Induced by Temperature and DMSO Random-sequence DNA unwinds from approximately 10.4 bp per turn at 37°C in water to an extrapolated 10.5 at 70°C or to 10.7 in 25% DMSO (Depew and Wang, 1975; Lee et al., 1981). In the absence of further structural information, it might be better to forgo a detailed analysis of the temperature-DMSO results presented above. We note, however, that an increase in the minor groove from 9 to 12 A in AT-rich tracts, as proposed by Fratini et al. (1982) would account for the increased sensitivity of these regions to DNAase I. A decrease *in the same-strand phosphate spacing from 6.6 to 5.8 A, as in the classical “B” to “A” transition (Arnott and Hukins, 1972), would account for the double-strand sensitivity of alternating purine-pyrimidine tracts to DNAase II. Local Helix Motion Sequence-induced variations in local helix conformation are important mainly to enzymes such as DNAase I and DNAase II, which perform nucleophilic attack on the phosphate group. By analogy with the mechanism worked out for micrococcal nuclease (Deiters et al., 1982) the nuclease-reactive species is probably a water molecule activated to OH- by its coordination to a bound metal ion or histidine residue. The lowest energy path of attack on the phosphate group is opposite to (i.e., in line with) the bond to be cut. Local rates of attack for DNAase I and DNAase II turn up strongly out of phase with one another at any phosphate; if DNAase I cuts the 03’-P bond well, then DNAase II cuts 05’.P poorly and vice versa. A likely explanation for this is diagrammed in Figure 8. Phosphate surfaces opposite 03’ and 05’ differ in orientation by a near-tetrahedral angle of 102”. Wherever bond 03’-P points outward so that DNAase I has easy access (top), bond 05’-P points upward, making DNAase II access difficult. Alternatively, where 05’.P points outward to give DNAase II easy access (bottom), 03’-P points downward to hinder DNAase I. Previously it was suggested that DNAase I recognizes local helix twist angle (Dickerson and Drew, 1981; Lomonossoff et al., 1981). Steps CG, for example, always have a low twist of 28”-30” and should therefore be resistant to

d

05’ I

-oP e

acce*s

Opposite

05’

0-R

sugar

Figure 8. Extremes

of Phosphate

Orientation

in Right-Handed

DNA

Phosphate rotations are induced by changes In the base-sugar arrangement, but for simplicity sugar rings are drawn here as if they were fixed objects. At top, the 03’.P bond points outward away from the helix axis while 05/-P points upward. In the center, neither bond points outward. At bottom, it is the 05’.P bond which points out while 0%P points down.

DNAase I (Dickerson, 1983a); yet many CG steps in the 160 bp sequence studied here are rather susceptible to DNAase I cleavage. From a structural point of view, phosphate orientation will depend on some combination of base-pair-step stacking parameters. To ascribe all of the stacking variability to twist alone may be sufficient for certain kinds of sequence, but now appears insufficient for the general case. Arguments against Hydrogen-Bonded Recognition of Base Pairs The three variations in helix geometry just presented, two global and one local, account for the digestion specificites of DNAase I, DNAase II, and copper-phenanthroline in their entirety. But could hydrogen-bonded recognition of DNA base pairs also play a role? In the case of copper-phenanthroline, definitely not: the complex of copper ion with two molecules of 1 ,lO-phenanthroline has no exposed donor or acceptor groups with which to participate in a hydrogen-bonding interaction. For the enzymes DNAase I and DNAase II, probably not: their DNA target sequences do not present any sort of consistent donor-acceptor surface to the protein. DNAase I cuts mixed-sequence DNA, avoiding extremes of base composition, while DNAase II cuts a mixture of A and G. The temperatureDMSO results argue even more forcibly against any sort of hydrogen-bonded recognition: in response to a change in solution environment, DNAase I and DNAase II cutting specificities change in some regions of the double helix, but not in others. This can only be attributed to a change in the structure of the DNA, rather than the protein.

Cell 500

Global Flexibility and Thermal Motion While it has been convenient in the present context to emphasize static aspects of DNA structure, some mention of DNA dynamics is clearly required. Levitt (1982) has found that, with respect to thermal motion, DNA is remarkably flexible in a global sense, much more so than a typical protein. He finds that large changes in helix groove width are thermally accessible at room temperature, and are closely related to the motions involved in DNA bending. This is not really surprising; if the small energies associated with base-sequence variation can perturb the structure in a way recognizable to proteins, then thermal motions should also be able to generate substantial fluctuations from time-averaged geometry. Biological Implications Two major forms of variation in DNA structure are now known to be possible. Under superhelical stress, the path of the double helix may be interrupted by cruciforms (Lilley, 1983) slippage loops (Mace et al., 1983) or left-handed “Z’-DNA (Peck and Wang, 1983). Under relaxed conditions, the double helix itself may assume a variety of sequence-dependent conformations that do not require the disruption of base pairs. What, if any, are the roles of these structural variations in biology? There presently exists little compelling evidence for the existence of cruciforms, slippage loops, or ‘Y-DNA in vivo (e.g., Courey and Wang, 1983; Hill and Stellar, 1983) and even less evidence for their involvement in gene regulation. In the one case where a particular biological role has been tested-that of a potential slippage loop 5’ to a Drosophila hsp70 gene-it was found that the sequences defining the structure can be deleted without affecting normal gene function (R. Dudler and A. Travers, unpublished data). We think that variations of the second kind, within the framework of a normal, right-handed double helix, may be of more biological relevance. These conformational differences are rather subtle, perhaps on the order of 2-4 kcal/ mole in free energy, yet small changes in free energy are often an important determinant of biological specificity. Here we have provided three examples, in the form of DNAase I, DNAase II, and copper-phenanthroline, where a protein or inorganic complex can recognize a specific helical conformation in the absence of any clear consensus sequence. It would be surprising if more examples were not forthcoming. Two proteins that come to mind in this regard are E. coli DNA gyrase (Morrison and Cozzarelli, 1981) and mouse glucocorticoid receptor (Payvar et al., 1983) neither of which exhibits a clear consensus sequence. Considerations of DNA structure may be of some value even for proteins that do exhibit a consensus sequence. If sequence is all-important, how then are we to rationalize the influence of flanking sequences on binding affinity? The restriction enzymes Eco RI and Hinf I provide a classic case in point, Both of these enzymes cut phosphates facing one another across the major groove, at sites

GAATTC and GANTC, respectively; their rates of cleavage, however, are not uniform for all sequences GAATTC and GANTC but depend rather strongly on the nature of sequences adjoining the protein binding site. In many instances, rates of cleavage are slowest when the restriction site lies amid runs of G/C base pairs; i.e., GGGGGAATTCCCCC for Eco RI (Armstrong and Bauer, 1983). Presumably the unusual groove width at G/C, wide in the minor and narrow in the major, carries over into the protein-binding site so as to disturb its structure. An easy test of this hypothesis would be to see if flanking runs of A/T can exert a similar effect, especially at low temperature. The E. coli tyrT promoter, used here as a digestion substrate, provides a more complex example of how variations in double-helical geometry could be of biological importance. In order to show optimal activity in vivo, this promoter requires sequences that extend far upstream of the primary RNA polymerase binding site, out to position 1 in Figure la (Lamond and Travers, 1983). The essential upstream region contains both a secondary polymerase binding site and two AT-rich blocks (positions 30 and 50) that are highly resistant to cleavage by DNAase I, DNAase II, and copper-phenanthroline (see also Travers et al., 1983). Other promoters regulated in a similar fashion to tyrTalso contain an upstream polymerase binding site and two Al-rich blocks, the positions but not the orientations of which are conserved (Travers, 1984). Hence, the essential feature of these sequence zones is probably their structure rather the sequence per se. We do not know how these sequences work but several tentative hypotheses have been suggested to us. One is that they provide an environmentally sensitive switch for gene regulation: since the conformation of these blocks seems rather sensitive to environmental factors such as temperature and DMSO, and since these blocks lie in positions that are contiguous to sequences defining the upstream RNA polymerase binding site, they might provide a means of regulating the affinity of polymerase for its upstream site. A second and entirely different mechanism would invoke the need for DNA bending around the polymerase; short runs of A or T, spaced at regular intervals along the double helix, can induce significant local curvature in the overall helix axis (Marini et al., 1982). Experiments are underway to test these and other possibilities,

Isolation and Purification of the Duplex Fragment Plasmid pKMA-98, a derivative of plasmid pBR322 containing the tyrosine tRNA promoter and its flanking sequences fused to the galactokinase gene (Lamond and Travers, 1983) was isolated from a culture of E. coli by lysis of the ceils at alkaline pH. It was purified by ultracentrifugatron in a cesium chloride/ethidium bromide densrty gradient, extracted with isopropanol to remove ethrdium bromide, dialyzed against low-salt buffer to remove cesium chloride, concentrated by ethanol precipitation, and redissolved in IO mM Tris. 0.1 mM EDTA, pH 8. The purified plasmid was next digested with restriction enzymes Ava I and Eco RI to leave a mixture of 160 bp promoter fragment and linearized plasmid DNA (Travers et al., 1983). From this mixture, the 160.mer fyrT

DNA Structure 501

in Solution

duplex could be isolated by electrophoresis on a 8% polyacrylamide gel. It was then eluted from a gel slice into 0.5 M NH,OAc, 1 mM EDTA, concentrated by ethanol precipitation, and redissolved in IO mM Tris, 0.1 mM EDTA at a concentration of 2 pg in 50 ~1. 3’-End-Labeling with Reverse Transcriptase The Ava I-Eco RI cleavage step left two sticky ends of unique sequence, whih could be filled in by the enzyme reverse transcriptase. Ava I recognizes the sequence CYCGRG (in this case CTCGGG) and leaves four unpaired nucleotides TCGG after cleavage. Incubation of the 160.mer duplex with reverse transcnptase, dGTP and n-32P-dCTP led to selective radioactive labeling of the 3’ end at the Ava I site. Eco RI, on the other hand, recognizes GAATrC and leaves four unpaired nucleotides AATT. Incubation with reverse transcnptase, dTTP, and a-?dATP led to selective labeling of the 3’ end at the Eco RI site. Positions of radioactive nucleotides in the labeled duplex are circled in Figures 2a and 3a above. Reverse transcriptase (Beard) and 0.1 pg DNA were incubated at 37°C for 2 hr in 100 mM Tris, IO mM MgClz, 6 mM mercaptoethanol, pH 8.0, with 50 pM dGTP or dTTP, and 5 ~1 a-32P-dCTP or dATP (3000 Ci/mmole, 10 mCi/ml). After heating to 65°C for 5 min to inactivate the enzyme, the reaction volume was mixed with an equal part 8 M NH,OAc, 100 @g/ml tRNA, then ethanol-precipitated three times to remove excess radioactive label, protein, and salt. The precipitant was redissolved in 60 ~1 of IO mM Tris, 0.1 mM EDTA, and stored frozen for later use. Nuclease digestion patterns were unaffected by the small amount of tRNA used in the precipitation step. Digestion Conditions and Electrophoresis Time courses of dlgestion were carried out at 37°C in water with allquots removed from the dlgestion mixture 1, 5, and 30 min after the addition of nuclease. DNAase I experiments included 0.3 U/ml enzyme (Sigma), 100 mM Tris, IO mM MgCl*, pH 7.6, and were stopped by the addition of 80% formamide, 0.1% bromophenol blue, 10 mM EDTA. DNAase II cuts included 3 U/ml enzyme (P. L. Biochemicals). 50 mM NH,,OAc, 1 mM EDTA, pH 5.4, and were frozen on dry ice immediately after addition of stop solution. Copper-phenanthroline digestlons contained 2 pM CuSO,, 20 PM 1 ,lOphenanthroline, 5 mM mercaptopropionlc acid, and 60 mM Tris, pH 7.6. Aliquots were mixed with an equal volume 8 M NH,OAc, 100 @g/ml tRNA, were twice ethanol-precipitated, then redissolved in 5 ~1 of 80% formamide, 0.1% bromophenol blue, 10 mM EDTA. Temperature-DMSO profiles of digestion were carried out at constant enzyme concentration and variable times, as Indicated in figure legends. All reactions were stopped by the addition of an equal volume 8 M NH,OAc, 100 *g/ml tRNA, were twice ethanol-preapitated, then redissolved in 5 ~1 of 80% formamide, 0.1% bromophenoi blue, 10 mM EDTA and heated at 100°C prior to electrophoresis. Products of digestlon were fractionated by electrophoresis on 8% polyacrylamide gels contalnlng 7 M urea and Tris-borate-EDTA buffer, pH 8.3. Following electrophoresis, gels were soaked in 10% HOAc for IO min, transferred to Whatman 3 MM paper, dried under vacuum at 7O”C, then subjected to autoradiography against an intensifying screen for IO-20 hr at -70°C. Densitometry and Data Processing Autoradiographs were scanned on a Joyce-Loebl microdensitometer to quantify the Intensities of gel bands. Chemical identities of digestion products were assigned using the known sequence and dimethyl sulfatepiperidine tracks specific for guanine. DNAase I and copper-phenanthroline products carry a 5’ phosphate and migrate in phase with guanine markers during electrophoresis. DNAase II products carry a 5’ hydroxyl and migrate 0.3-l .2 bonds more slowly than guanine markers, the difference In mobility becoming larger as the chains grow shorter. With reference to Figure 2b above, one should note that copperphenanthroline actually attacks sugar rings facing one another across the minor groove, two base-pair steps apart. When the sugar falls apart, cuts appear at either 03,-P or 05’.P positlons, depending on the choice of radioactlve end label. For a 3’ label, cuts on opposrte strands appear as 03,-P cleavages spaced three base-pair steps apart; for a 5’-label (Drew, submltted), cuts show up as 05,-P products spaced one step apart Probabllitles of cleavage were calculated by the method of Lutter (1978)

from data at single time points of digestron. In this procedure, the areas R, under bands in a gel lane are first summed to give the total area (or radioactivity) Rt, then converted to fractions of total cleavage f, = R,/R,. Probabilities of cleavage are derived from these fractional distributions by correcting for the incidence of cuts between bond i and the radioactive label. For example, if 0.40 of the total radioactivity lies between bond i and the radloactive label, then a cut at bond i WIII only appear as a chain of length i on the gel 0.60 fraction of time. The true cleavage probability in this case is P, = f,/O.60, or in general P, = f,/f,, where fa is the sum of f, over all bonds away from the label relative to bond i. Note that this sum should include both the fractional radioactivity in bond i itself and in the uncut starting material. It is desirable that the correction term be as small as possible, and for this reason time points of digestion were chosen at which the fraction of starting material left was as large as possible, typically 0.50 to 0.80. Sets of data collected from different gel lanes, under different solution conditions of temperature-DMSO, were placed on an approximate 1 :l scale by visual comparison of peak heights, both In the final probability plot and in the original gel patterns. Acknowledgments The authors thank Drs. Chris Calladine for extensive discussions of helix geometry Aaron Klug for critical comment, and Maxine McCall for continual encouragement. Dr. Angus Lamond generously provided plasmid pKMb 98. H. D. was the recipient of National Research Service Award CA 06971. 01 of the National Institutes of Health. Received

January

3, 1984; revised

February

17, 1984

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Lilley, D. (1983). Structural perturbation in supercoiled DNA: hypersensitivity to modification by a single-strand-selective chemical reagent conferred by inverted repeat sequences. Nucl. Acids Res. 77, 3097-3112. Lomonossoff, G. P., Butler, P. J. G., and Klug, A. (1981). Sequencedependent variation in the conformation of DNA. J. Mol. Biol. 149, 745760. Lutter, L. C. (1978). Kinetic analysis of deoxyribonuclease I cleavages in the nucleosome core: evidence for a DNA superhelix. J. Mol. Biol. 124, 391-420. Mace, H. A. F., Pelham, H. Ft., and Travers, A. A. (1983). Association of an Sl nuclease-sensitive region with short direct repeats 5’ of Drosophila heat shock genes. Nature 304, 555-557. Marini, J. C., Levene, S. D., Crothers, D. M., and Englund. P. T. (1982). A bent helix in kinetoplast DNA. Cold Spring Harbor Symp. Quant. Biol. 47, 279-283. Morrison, A., and Cozzarelli, N. R. (1981). Contacts between DNA gyrase and its binding site on DNA: features of symmetry and asymmetry revealed by protection from nucleases. Proc. Nat. Acad. Sci. USA 78, 1416-1420. Nickel, J. M., and Felsenfeld, G. (1983). DNA conformation of the chicken adult @-globin gene. Cell 35, 467-477.

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Rhodes, D. (1982). The helical periodicity of DNA in solution and in chromatin. In Topics In Nucleic Acid Structure, Part 2, S. Neidle, ed. (London: Macmillan Press), pp. 287-304 Schon, E., Evans, T., Welsh, J., and Efstratiadis, A. (1983). Conformation of promoter DNA: fine mapping of Sl-hypersensitive sites. Cell 35, 837848. Shakked, Z., Rabinovich, D., Kennard, O., Cruse, W. 8. T., Salisbury, S. A., and Viswamitra, M. A. (1983). Sequence-dependent conformation of an ADNA double helix: the crystal structure of the octamer d(GGTATACC). J. Mol. Biol. 166, 183-201. Singleton, C. K., Klysik, J., and Wells, R. D. (1983). Conformationalflexibility of junctions between contiguous B and Z-DNA in supercoiled plasmids. Proc. Nat. Acad. Sci. USA 80, 2447-2451. Travers, A. A. (1984). Conserved features moters. Nucl. Acids Res. 12, 2605-2618.

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Travers, A. A., Lamond, A. I., Mace, H. A. F., and Berman, M. L. (1983). RNA polymerase interactions with the upstream region of the E. coli tyrT promoter. Cell 35, 265-273. Uesugi, S.. Shida. T., Ikehara, M., Kobayashi, Y., and Kyogoku, Y. (1982). Identification of degradation productions of d(C-G) by a 1,lO-phenanthroline-copper ion complex. J. Am. Chem. Sot. 704. 54945495.

Zimmerman, S. B., and Pheiffer, B. H. (1981). A RNA-DNA hybrid that can adopt two conformations: an x-ray diffraction study of poly (rA).(dT) in concentrated solution or in fibers. Proc. Nat. Acad. Sci. USA 78, 78-82.