|
|
Proc Natl Acad Sci U S A. 2003 July 8; 100(14): 8113–8118. | PMCID: PMC166191 |
Copyright © 2003, The National Academy of
Sciences Chemistry Water at DNA surfaces: Ultrafast dynamics in minor groove
recognition Samir Kumar Pal, Liang Zhao, and Ahmed H. Zewail* Laboratory for Molecular Sciences, Arthur Amos Noyes Laboratory of Chemical Physics, California Institute of Technology, Pasadena, CA 91125 Water molecules at the surface of DNA are critical to its equilibrium
structure, DNA–protein function, and DNA–ligand recognition. Here
we report direct probing of the dynamics of hydration, with femtosecond
resolution, at the surface of a DNA dodecamer duplex whose native structure
remains unperturbed on recognition in minor groove binding with the
bisbenzimide drug (Hoechst 33258). By following the temporal evolution of
fluorescence, we observed two well separated hydration times, 1.4 and 19 ps,
whereas in bulk water the same drug is hydrated with time constants of 0.2 and
1.2 ps. For comparison, we also studied calf thymus DNA for which the
hydration exhibits similar time scales to that of dodecamer DNA. However, the
time-resolved polarization anisotropy is very different for the two types of
DNA and clearly elucidates the rigidity in drug binding and difference in DNA
rotational motions. These results demonstrate that hydration at the surface of
the groove is a dynamical process with two general types of trajectories; the
slowest of them (≈20 ps) are those describing dynamically ordered water.
Because of their ultrafast time scale, the “ordered” water
molecules are the most weakly bound and are accordingly involved in the
entropic (hydration/dehydration) process of recognition. Hydration of DNA plays important role in its structure, conformation, and
function. Of significance to the function is the selective recognition by DNA
of small molecules (ref. 1 and
references therein). X-ray crystallography, NMR, dielectric relaxation, and
molecular dynamics simulation studies have shown that a significant amount of
water molecules are bound to DNA (for reviews, see refs.
2– 6).
For example, measurements of dielectric relaxation caused by water molecules
bound to DNA in mixed water-ethanol solutions have found that 18–19
water molecules per nucleotide are present in B-DNA, but only 13–14
water molecules are bound in A-DNA
( 5). The study also suggested
that a structural transition of poly(dG-dC)·poly(dG-dC) DNA from its B
to Z form takes place on the removal of the bound water molecules,
preferentially from the phosphate groups. The molecular picture of hydration in the minor groove of B-DNA is unique.
An x-ray crystallographic investigation
( 7) followed by solution NMR
study ( 8) on a model dodecamer
B-DNA duplex (for the sequence of A/T tracts, CGCGAATTCGCG) showed that the
minor groove is hydrated in an extensive and regular manner, with a zigzag
“spine” of first- and second-shell hydration along the floor of
the groove. In contrast, hydration within the major groove is principally
confined to a monolayer of water molecules. The conformational energy
calculation suggested that the presence of the spine of hydration is the prime
reason for the further narrowing of minor groove
( 9). The influence of drug binding on DNA hydration is striking. Acoustic and
densimetric studies have shown that a fraction (not total) of the water
molecules is released on recognition
( 10,
11). Hence, the balance
between enthalpic and entropic changes in determining the overall free energy
of recognition. The degree of order in water dynamics is determined by the
time scale of the motion and is critical to the hydrophobic contribution.
Recently, studies of solvation dynamics have been reported for an extrinsic
chromogenic probe, inserted into DNA either by covalent adduction of coumarin
dye ( 12) or hydrophobic
intercalation of acridine dye
( 13). The fluorescence results
( 12) give two relaxation time
constants of 300 ps (47%) and 13 ns (53%), both measured with 100-ps time
resolution and attributed to the local reorganization (by DNA and/or water) in
the modified DNA. The results from femtosecond-resolved transient absorption
( 13) suggest an ultrafast
(within 200 fs) “repolarization” of nuclear degrees of freedom of
the DNA pocket. However, the lack of information on structures for both
complexes keeps unknown the extent of perturbation on DNA. Moreover, neither
study gave the hydration dynamics in the DNA grooves. Using the natural DNA
bases to probe hydration is hampered by their ultrafast lifetimes (ref.
14 and references
therein). In this article, we present our study of DNA hydration dynamics, with
femtosecond resolution, in the minor groove
(), using the dodecamer
B-DNA duplex d(CGCAAATTTGCG) whose x-ray structure without and with the drug
Hoechst 33258 ( H33258) has been reported
( 15,
16); the site for recognition
is the minor groove and remains unchanged on binding. We follow the time
evolution of the Stokes shift of the fluorescence of the drug, bound to DNA,
and obtain the decay of the hydration correlation function. The function
reflects the rotational and translational motions of water molecules in the
minor groove. The drug, which is in a class of antimicrobial agents
( 17), has high affinity to DNA
( Kd ≈ 10 -8 M); its fluorescence intensity
increases when complexed to DNA, relative to that in aqueous solution, and is
accompanied by a significant blue shift
( 18– 20).
We also report our study of calf thymus DNA (CT-DNA) on binding to the same
drug ( H33258) to compare with the dodecamer DNA. Finally, to probe the
rigidity of the drug and the time scale of its motion with DNA we examined the
time-resolved polarization anisotropy, r( t). Time-Resolved Studies. All of the transients were obtained by using
the femtosecond-resolved fluorescence up-conversion technique. The detailed
experimental setup is described elsewhere
( 21). For the studies reported
here, a fs excitation pulse (200 nJ) was used at 350 nm. To measure
time-resolved anisotropy, r( t), and to construct hydration
correlation function, C( t), we followed the procedure
detailed in our previous publications
( 21– 23). Sample Preparation. CT-DNA (Sigma) and Hoechst 33258 (Molecular
Probes, 99% pure) were used without further purification. The drug was in the
form of trihydrochloride, pentahydrate, and at neutral pH it only has one
positive charge (see below, ). The dodecamer DNA with sequence CGCAAATTTGCG was synthesized,
purified by reverse-phase cartridge, annealed to make duplex, and supplied by
Gene Link (Hawthorne, NY). The purity of the duplex was checked by urea PAGE;
essentially one band was characterized by the supplier. Aqueous DNA solutions
were prepared in a Tris buffer (0.01 M Tris/0.001 M EDTA) at pH 7.4 in water
from a Nanopure (Dubuque, IA) purification system. The DNA–drug complexes were prepared by mixing the drug H33258 (96
μM) with the duplex (144 μM) in the aqueous buffer solutions with
continuous stirring for 4 h. The yield of the complexes is >99% (1:1
complex) at our drug concentration, because the equilibrium constant is
≈10 8 M -1
( 24). The procedure for making
CT-DNA aqueous solution is similar to that of ref.
20. Solid CT-DNA (1 mg/ml) was
dissolved into buffer solutions. The DNA solutions were sonicated for 30 s to
reduce the DNA chain length and stirred for 1 h. The ratio of absorption at
260 and 280 nm for the DNA solutions gave a value 1.84, in accord with the
limit of 1.8–1.9 for highly purified preparation of DNA
( 25). For the dodecamer duplexes, the complexes are 1:1, given the equilibrium
constant and the relative concentrations. For CT-DNA, the drug sites are far
apart because of the concentration used. The nucleotide concentration was
determined by absorption spectroscopy of CT-DNA using the average extinction
coefficient per nucleotide of CT-DNA (6,600
M -1·cm -1 at 260 nm)
( 20) and found to be 2.5 mM,
i.e., 1.25 mM for base pairs. A known concentrated drug solution was added
dropwise to the CT-DNA solutions with continuous stirring for 2 h to achieve a
final concentration of 50 μM for the drug. Accordingly, we estimate that 25
bp of CT-DNA are available for each drug molecule. Drug in Bulk Water. The steady-state fluorescence spectra of the
drug in three neat solvents are presented in
Upper. The
spectra show a large red solvatochromic effect. The fluorescence maximum
changes from 436 nm in dioxane to 477 nm in ethanol and to 510 nm in aqueous
buffer solutions. The effect of DNA binding on the fluorescence intensity and
maxima is shown in
Lower. In water, the fluorescence is quenched dramatically; the quantum yield is
0.015 in water and increases to 0.5 in ethanol
( 19). It is known
( 20,
26) that two major processes
are involved in the deactivation of the excited state. At high pH (>7.0)
the excited molecule undergoes intramolecular proton transfer from the phenol
(see below) to the closest benzimidazole nitrogen, resulting in a keto
structure. In polar solvents, the fluorescence intensity decreases and is
dominated by emission from the keto form. The deactivation at low pH values
(<7.0), and on the complexation with DNA, mainly involves rotation along
the bisbenzimide axis ( 20),
similar to other cases involving charge transfer
( 27). The solvent's pH,
polarity, and rigidity determine the wavelength and yield of emission. We have
studied the lifetime of the drug in bulk water and found it to be multiple
exponentials (ps and ns ranges) and to depend on its concentration. The
enhancement of the ps component, characteristic of high pH decay
( 20) at high concentration,
may suggest the presence of ground-state conformations whose equilibrium
changes with concentrations and/or aggregation; a subset of those
conformations is more poised for twisting/proton transfer. In contrast, in DNA
the fluorescence decay is dominated by ns components and does not change with
concentration of DNA studied (μM to mM); DNA/drug remains at 1.5. Femtosecond-resolved fluorescence up-conversion transients of the drug in
bulk water (buffer) are presented in Upper Left for three characteristic wavelengths; we
have studied at least 13 of these transients at different wavelengths. The fs
transients are typical of those observed for other chromophores in water
( 22,
23). On the blue edge of the
spectrum, the signal is seen to decay (≈1.5 ps), whereas on the red edge it
rises on a similar time scale. The signal decays with time constants of ≈40
and 500 ps in the time window studied, up to 200 ps. From the transients it is
clear that the contribution of ultrafast hydration is well separated from the
nonradiative processes. Note that hydration is essentially independent of
details of the solute fluorophore (see, e.g., refs.
21– 23
and references therein). From this family of transients we constructed the time-resolved emission
spectra (TRES) shown in
Upper Right and the hydration correlation function
C( t) given in Lower. C( t) shows an apparent biexponential
decay with time constants of 195 fs (33%) and 1.2 ps (67%). This behavior is
typical of hydration of a molecular probe in bulk water
( 22,
28,
29). The overall spectral
shift we observed is 3,184 cm -1; any sub-100-fs component in the
dynamics would be unresolved. To complete the picture regarding bulk dynamics
of the drug in water and in a less polar solvent (for comparison with DNA
environment), we made similar studies of the drug in ethanol (data not shown).
In this case, decays are slower than those in water and consistently show the
time scale of solvation for ethanol [the average solvation time is 16 ps
( 30)]. To elucidate the time scale for the rotational diffusion of the drug and
its complex we measured the time-resolved anisotropy, r( t),
using single-photon counting techniques
( Lower Inset).
At t = 0, r( t) ≈0.4 and decays to the base line
(properly corrected for polarization/intensities) with τ = 530 ps.
According to the Stokes–Einstein–Debye hydrodynamics theory, and
assuming a prolate shape for the drug molecule
( 31,
32), the rotational times in
water range from 1 to 5 ns, considering the boundary conditions. Our measured
value of 530 ps must include the twisting process, which changes the
anisotropy. Note that the r( t) decay in water is on a much
longer time scale than that of hydration. Thus rotational diffusion and
conformational changes are separable in their time scales from that of
hydration. Vibrational relaxation is insignificant given the calculated
0—0 transition in water (316.5 nm) and for the reasons given in ref.
23. Drug–DNA Dodecamer Complex.
Upper Left
shows three characteristic transients obtained from up-conversion experiments
on the drug–DNA dodecamer complex in aqueous buffer solutions; other
wavelengths are not shown. On the blue edge of the spectrum the signals decay
on different time scales depending on wavelengths, whereas on the red edge the
signal is seen to rise. The constructed TRES are shown in
Upper Right,
indicating that hydration is complete within 100 ps; the spectrum at 100 ps
reaches the equilibrium spectrum obtained from steady-state fluorescence. The C(t) function, as shown in
Lower, is a sum
of two exponentials with the time constants of 1.4 ps (64%) and 19 ps (36%);
any sub-100-fs components in these dynamics are unresolved. The net spectral
shift observed is 1,304 cm-1, consistent with the behavior depicted
in . To ascertain the
degree of orientational rigidity of the drug in the complex we obtained the
fluorescence anisotropy, r(t), decay at 510 nm by using
single-photon counting. The r(t) is observed
( Lower Inset)
to decay with a time constant of ≈5.5 ns (the estimated hydrodynamic
rotational relaxation times of the complex are in the range 10–50 ns).
The one order of magnitude lengthening of the drug anisotropy decay in
dodecamer, compared with that in the bulk, is consistent with a rigid drug
binding in the minor groove. Drug–CT-DNA Complex. In
Upper Left, we
show the up-conversion signals at three characteristic wavelengths; other
wavelengths are not shown. The time scales of the decay in the blue edge and
corresponding rise in the red edge are similar to those observed in the
dodecamer duplex. The constructed TRES in
Upper Right
indicate that at 100 ps the equilibrium of the fluorescence, as observed from
steady-state emission spectrum, has been reached. The C(t) function for the CT-DNA in
Lower can be
fitted to a biexponential decay with time constants of 1.1 ps (60%) and 19 ps
(40%); any sub-100-fs components in these dynamics are unresolved. The net
spectral shift observed is 1,582 cm-1. The time constants along
with their contributions are similar to those observed in the dodecamer
duplex. But, the fluorescence anisotropy at 510 nm decays with a different
time constant: ≈55 ns (major component), compared with that of ≈5.5 ns
in the duplex. This finding is consistent with the fact that natural CT-DNA is
much longer than the duplex DNA (12 pairs of bases only), but the effect of
binding to sites of genomic DNA is another factor to consider. Summarizing our observations we can make the following points: (i)
Hydration dynamics in the minor grooves of both types of DNA (dodecamer duplex
and calf thymus) are similar but differ substantially from those in bulk
water; (ii) for both types of DNA, and within our time resolution, a
“bimodal” hydration behavior with two distinct time constants,
≈1 ps (60%) and ≈20 ps (40%), was observed, reflecting the presence of
two types of water, bulk-type, labile water and weakly bound, ordered water;
and (iii) rotational diffusion of the drug–DNA complexes is
much slower than that of the drug in bulk water (530 ps), but is different for
the two types of DNA studied; 5.5 ns for dodecamer duplex and 55 ns for calf
thymus. These results indicate the rigidity caused by binding in the minor
grooves and the disparity in time scales of hydration and rotational
diffusion. The observed bimodality () in surface hydration of DNA(s) are in fact consistent with our
previous reports
( 21– 23)
on hydration dynamics at protein surfaces. The DNA hydration time of 19 ps
(40%) for the weakly bound water is in line with those observed for the
protein surface hydration: Subtilisin Carlsberg (38 ps, 39%),
Monellin (16 ps, 54%), and α-chymotrypsin (28 ps, 10%). Relating these
times of the correlation functions to residence times in the water layer
identifies the effect of first-shell polarization caused by the restricted
motions of water molecules by rotational and translational diffusion
( 33). These residence times
are for a layer of nm thickness
( 22) and for the weakest-bound
water because our measurements span the earliest possible (fs) time scale. The
persistence of time scales in DNA and proteins is consistent with solvation by
water (not internal groups), as discussed elsewhere
( 23). Considering the equilibrium between water molecules and the DNA sites, a
residence time of minor grooves with koff ≈ 5 ×
10 10 s -1 and diffusion controlled
kon ≈ 10 10
M -1·s -1
( 34) gives
Kd ≈ 5 M, so that each accessible site on the average
would be in contact with water for >90% of the time; the site has very high
occupancy at equilibrium. Dynamically speaking, however, the site is in
exchange with bulk, destroying the order on the ps time scale, and this is the
origin of bimodality ( 22,
33). The residence time for ordered water relative to other time constants of
DNA is of significance to the stability and recognition. First, we must
consider the time scale of making and breaking bonds of the dynamically
ordered water, τ DOW, relative to that of structural
conformational changes, τ CC, by bending and twisting (ref.
35 and references therein).
The value of τ CC is important to, for example, the change of
the B form of DNA to A and Z forms whose relative stabilities depend on the
water content and sequence; B form predominates in aqueous solution. If
τ DOW is shorter than τ CC then recognition is an
effective process with structural integrity. The loss of order on the ps time
scale is significant in increasing the entropy and it is possible that this
contribution to the free energy is governed by the change in the rotations of
water molecules. It is interesting that for this drug and DNA sequence, the
entropic contribution is indeed dominant
( 36). Repeating these
experiments for different drugs or sequences should correlate the
thermodynamics
( 37– 39)
with the dynamics. Second, it is important to compare the residence time of weakly bound
water, τDOW, with the time of breaking/making hydrogen bonds,
τHB, in bulk water. With a few kcal/mol barrier, kinetically
τHB is on the order of a few picoseconds, and for an effective
recognition, τDOW should not be orders of magnitude longer than
the value of τHB, so that the efficiency becomes optimum. If
τDOW/τHB ≈ 1, then the degree of order is
that of the bulk. Lastly, the time scale for the motion of the drug in the
groove, by orientational diffusion, τOD, relative to that of
τDOW. For the drug studied here, τOD is much
longer than τDOW for both types of DNA, as evidenced by the
anisotropy, ensuring a well defined geometry, certainly on the time scale of
dynamically ordered water. The robustness of the range of values for the hydration times in DNA (and
proteins) is indicative of the nature of the layer, being ordered on the
molecular scale even in the presence of the drug. This picture is consistent
with the results of NMR study
( 34) on the hydration and
solution structure of the duplex sequence we have used and its complex with a
minor groove binding drug, propamidine. It was found that complexation with
the drug has little effect on the residence times for water molecules bound
either in the major groove or at the sites in the minor groove. The range of
residence times was found to be ≈0.2–0.4 ns for surface water at
grooves; the residence times of water molecules in the major groove are an
order of magnitude shorter than for the most long-lived waters in the minor
groove. From the above discussion two points should be emphasized. Our
above-mentioned results of hydration dynamics in the two DNA systems studied
were obtained by using the time window from zero and up to 200 ps. Longer-time
Stokes shifts may be present, reflecting the influence of more rigid water
structure on the time scale indicated by NMR studies
( 4,
40). However, the resemblance
of the evolving spectra to the maximum of steadystate fluorescence spectrum in
100 ps [ν max(∞)] indicates that most of the dynamics are
complete within our time window, but a fraction of strongly bound water may
still be present with a time scale of sub-ns or longer [we measured the
lifetime of the drug in the dodecamer and found it to be ≈1.5 and 4 ns,
consistent with the values in the literature
( 20)]. We note that the ns
decays are similar for all DNA concentrations studied, which indicates that
the dodecamer remains as a double strand, and not as a hairpin that is
possible for central AT and terminal GC bases of a single strand; NMR confirms
the double-strand structure at 1.1 mM
( 34). Because the ordered water is probed here around the drug in the grooves,
one must not ignore such water in the study of recognition processes and drug
design strategies ( 36). In
fact, studies using densimetric and ultrasonic measurements have shown that a
minor groove binding drug, netropsin, displaces, depending on the base
sequence, a significant number of water molecules on complexation with a DNA
duplex ( 10). We note that
thermodynamic measurements of release of water is concerned with essentially
all waters occupied in the grooves whereas the dynamics experiments can detect
only a small subpopulation of a certain net of interfacial water molecules
that contribute to ligand binding. The dynamics of the ordered water and its
loss by, for example, rotational diffusion contribute to the entropic process
involved. From a structural point of view, the drug studied here
() binds in the minor
groove covering the sequence AATTT of the central A tract, with the piperazine
group close to one of the GC region. It makes two three-centered hydrogen
bonds from the nitrogen atoms of benzimidazole rings to the N (A18) and O (T7,
T8, T19) atoms of the DNA bases. This hydrogen bonding (and
electrostatic/dispersion interactions) is facilitated by the presence of
ordered water (entropic) around the drug; if ordered water is involved in
direct binding of the drug then enthalpic contributions must be included.
Hydrogen bonding is also possible for the drug
(; nitrogens opposite
to N1 and N3) with water near the surface of the groove, which is more of a
bulk type in our bimodal distribution of hydration. The most weakly bound
water molecules are of critical importance to biological function
( 41) and are unlikely to be
seen in crystal structures or by NMR
( 41). They are part of total
hydration, which influences structural and biological activities
( 42,
43). The residence times of
weakly bound water are only an order of magnitude different from that of bulk,
and they are the ones that have to be probed with femtosecond resolution. In conclusion, the study presented here characterizes, with femtosecond
resolution, the dynamics of hydration/dehydration at the DNA surface of known
local structure and with a drug in the minor groove. The fact that the water
is dynamically ordered at the surface of DNA without spatial averaging or
position inhomogeneity of the drug (known structure) allow us to observe the
earliest processes of hydration dynamics. Recognition of minor grooves by
charge and shape complementarities, and using directional hydrogen bonds,
cannot be fully understood from a static structure without including the role
of water and the time scale for the loss of the order. It may turn out that
this dynamically ordered water is also crucial for interfacial recognition,
not only of drugs but also between macromolecules. We thank Drs. Jorge Peon and Spencer Baskin for helpful discussion and
Profs. Kenneth J. Breslauer and Peter B. Dervan for careful reading of the
manuscript and helpful suggestions. This work was supported by the National
Science Foundation. 1. Dervan, P. B. (2001) Bioorg. Med.
Chem. 9,
2215-2235. [PubMed] 2. Feig, M. & Pettitt, B. M. (1998)
Biopolymers 48,
199-209. [PubMed] 3. Schneider, B., Cohen, D. & Berman, H. M. (1992)
Biopolymers 32,
725-750. [PubMed] 4. Halle, B. & Denisov, V. P. (1998)
Biopolymers 48,
210-233. [PubMed] 5. Umehara, T., Kuwabara, S., Mashimo, S. & Yagihara, S.
(1990) Biopolymers
30,
649-656. [PubMed] 6. Duan, Y., Wilkosz, P., Crowley, M. & Rosenberg, J. M.
(1997) J. Mol. Biol.
272,
553-572. [PubMed] 7. Drew, H. R. & Dickerson, R. E. (1981)
J. Mol. Biol. 151,
535-556. [PubMed] 8. Leipinsh, E., Otting, G. & Wuthrich, K. (1992)
Nucleic Acids Res. 20,
6549-6553. [PMC free article] [PubMed] 10. Chalikian, T. V., Plum, G. E., Sarvazyan, A. P. & Breslauer, K.
J. (1994) Biochemistry
33,
8629-8640. [PubMed] 11. Chalikian, T. V., Volker, J., Srinivasan, A. R., Olson, W. K. &
Breslauer, K. J. (1999) Biopolymers
50,
459-471. [PubMed] 12. Brauns, E. B., Madaras, M. L., Coleman, R. S., Murphy, C. J. &
Berg, M. A. (1999) J. Am. Chem. Soc.
121,
11644-11649. 13. Hess, S., Davis, W. B., Voityuk, A. A., Rosch, N., Michel-Beyerle,
M. E., Ernsting, N. P., Kovalenko, S. A. & Lustres, J. L. P.
(2002) Chemphyschem.
3, 452-455. [PubMed] 14. Pal, S. K., Peon, J. & Zewail, A. H. (2002)
Chem. Phys. Lett. 363,
57-63. 15. Edwards, K. J., Brown, D. G., Spink, N., Skelly, J. V. &
Neidle, S. (1992) J. Mol. Biol.
226,
1161-1173. [PubMed] 16. Vega, M. C., Saez, I. G., Aymami, J., Eritja, R., Marel, G. A. V.
D., Boom, J. H. V., Rich, A. & Coll, M. (1994)
Eur. J. Biochem. 222,
721-726. [PubMed] 17. Denham, D. A., Suswillo, R. R., Rogers, R., McGreevy, P. B. &
Andrew, B. J. (1976) J. Helminthol.
50,
243-250. [PubMed] 18. Latt, S. A. & Stetten, G. (1976) J.
Histochem. Cytochem. 24,
24-33. [PubMed] 19. Gorner, H. (2001) Photochem.
Photobiol. 73,
339-348. [PubMed] 20. Cosa, G., Focsaneanu, K.-S., McLean, J. R. N., McNamee, J. P. &
Scaiano, J. C. (2001) Photochem.
Photobiol. 73,
585-599. [PubMed] 21. Peon, J., Pal, S. K. & Zewail, A. H. (2002)
Proc. Natl. Acad. Sci. USA
99,
10964-10969. [PMC free article] [PubMed] 22. Pal, S. K., Peon, J. & Zewail, A. H. (2002)
Proc. Natl. Acad. Sci. USA
99,
1763-1768. [PMC free article] [PubMed] 23. Pal, S. K., Peon, J. & Zewail, A. H. (2002)
Proc. Natl. Acad. Sci. USA
99,
15297-15302. [PMC free article] [PubMed] 24. Haq, I., Ladbury, J. E., Chowdhry, B. Z., Jenkins, T. C. &
Chaires, J. B. (1997) J. Mol. Biol.
271,
244-257. [PubMed] 25. Gallagher, S. R. (1994) in Current
Protocols in Molecular Biology, eds. Ausubel, F. M., Brent, R.,
Kingston, K. E., Moore, D. D., Seidman, J. G., Smith, J. A. & Struhl, K.
(Greene and Wiley-Interscience, New York), Suppl. 28, Appendix 3D. 26. Kalninsh, K. K. & Pestov, D. V. (1994)
J. Photochem. Photobiol. A
83, 39-47. 27. Zhong, D., Pal, S. K. & Zewail, A. H. (2001)
Chemphyschem. 2,
219-227. 28. Jarzeba, W., Walker, G., Johnson, A. E., Kahlow, M. A. &
Barbara, P. F. (1988) J. Phys. Chem.
92,
7039-7041. 29. Jimenez, R., Fleming, G. R., Kumar, P. V. & Maroncelli, M.
(1994) Nature
369,
471-473. 30. Horng, M. L., Gardecki, J. A., Papazyan, A. & Maroncelli, M.
(1995) J. Phys. Chem.
99,
17311-17337. 31. Hu, C. M. & Zwanzig, R. (1974) J. Chem.
Phys. 60,
4354-4357. 32. Baskin, J. S. & Zewail, A. H. (2001) J.
Phys. Chem. A 105,
3680-3692. 33. Pal, S. K., Peon, J., Bagchi, B. & Zewail, A. H.
(2002) J. Phys. Chem. B
106,
12376-12395. 34. Lane, A. N., Jenkins, T. C. & Frenkiel, T. A.
(1997) Biochim. Biophys. Acta
1350,
205-220. [PubMed] 35. Millar, D. P., Robbins, R. J. & Zewail, A. H.
(1982) J. Chem. Phys.
76,
2080-2094. 36. Haq, I. (2002) Arch. Biochem.
Biophys. 403,
1-15. [PubMed] 38. Breslauer, K. J., Remeta, D. P., Chou, W., Ferrante, R., Curry, J.,
Zaunczkowski, D., Snyder, J. G. & Marky, L. A. (1987)
Proc. Natl. Acad. Sci. USA
84,
8922-8926. [PMC free article] [PubMed] 39. Marky, L. A. & Breslauer, K. J. (1987)
Proc. Natl. Acad. Sci. USA
84,
4359-4363. [PMC free article] [PubMed] 40. Denisov, V. P., Carlstrom, G., Venu, K. & Halle, B.
(1997) J. Mol. Biol.
268,
118-136. [PubMed] 41. Rand, R. P. (1992) Science
256,
618-618. 42. Mrevlishvili, G. M., Carvalho, A. P. S. M. C., da Silva, M. A. V.
R., Mdzinarashvili, T. D., Razmadze, G. Z. & Tarielashvili, T. Z.
(2001) J. Therm. Anal. Cal.
66,
133-144. 43. Robinson, C. R. & Sligar, G. (1993) J.
Mol. Biol. 234,
302-306. [PubMed] 44. Teng, M., Usman, N., Frederick, C. A. & Wang, A. H. J.
(1988) Nucleic Acids Res.
16,
2671-2690. [PMC free article] [PubMed]
|
This article has been 
