Altered Dynamics of DNA Bases Adjacent to a Mismatch: A Cue for Mismatch Recognition by MutS

doi:10.1016/j.jmb.2007.08.065

J. Mol. Biol. (2007) 374, 39–53

Altered Dynamics of DNA Bases Adjacent to a Mismatch: A Cue for Mismatch Recognition by MutS Nabanita Nag 1 , B. J. Rao 2 ⁎ and G. Krishnamoorthy 1 ⁎ 1

Department of Chemical Sciences, Tata Institute of Fundamental Research, Homi Bhabha Road, Mumbai 400005, India 2

Department of Biological Sciences, Tata Institute of Fundamental Research, Homi Bhabha Road, Mumbai 400005, India Received 17 April 2007; received in revised form 27 August 2007; accepted 27 August 2007 Available online 5 September 2007

The structural deviations as well as the alteration in the dynamics of DNA at mismatch sites are considered to have a crucial role in mismatch recognition followed by its repair utilizing mismatch repair family proteins. To compare the dynamics at a mismatch and a non-mismatch site, we incorporated 2-aminopurine, a fluorescent analogue of adenine next to a G.T mismatch, a C.C mismatch, or an unpaired T, and at several other non-mismatch positions. Rotational diffusion of 2-aminopurine at these locations, monitored by time-resolved fluorescence anisotropy, showed distinct differences in the dynamics. This alteration in the motional dynamics is largely confined to the normally matched base-pairs that are immediately adjacent to a mismatch/ unpaired base and could be used by MutS as a cue for mismatch-specific recognition. Interestingly, the enhanced dynamics associated with base-pairs adjacent to a mismatch are significantly restricted upon MutS binding, perhaps “resetting” the cues for downstream events that follow MutS binding. Recognition of such details of motional dynamics of DNA for the first time in the current study enabled us to propose a model that integrates the details of mismatch recognition by MutS as revealed by the high-resolution crystal structure with that of observed base dynamics, and unveils a minimal composite read-out involving the base mismatch and its adjacent normal base-pairs. © 2007 Elsevier Ltd. All rights reserved.

Edited by J. Karn

Keywords: mismatch DNA; mismatch repair; DNA dynamics; fluorescence lifetime; fluorescence anisotropy

Introduction DNA mismatch repair (MMR) system, an evolutionarily conserved biochemical pathway, plays an important role in regulating the genome by correcting base mismatches arising from either replication errors (error rate 10−6–10−7)1 or homologous recombination, thereby preventing recombination between DNA molecules that have a high level of sequence divergence (mismatches).2–4 Expectedly, inactivation of MMR genes leads to a significant increase in the spontaneous mutation rate, resulting in various genomic disorders.2,5–7 The most extensively studied adenine methyl-directed MMR pathway of Escherichia coli implicates the participation of several gene products in it, including MutS, MutL, *Corresponding authors. E-mail addresses: [email protected]; [email protected]. Abbreviations used: MMR, mismatch repair; IDL, insertion deletion loop; 2-AP, 2-aminopurine.

MutH, DNA helicase II, single-stranded DNAbinding protein, exonuclease I, exonuclease VII or RecJ exonuclease, DNA polymerase III holoenzyme and DNA ligase.2–8 MutS, the key protein in MMR, recognizes and binds to mismatched DNA basepairs/insertion-deletion-loop (IDL) with an affinity that is only several-fold higher than that of its binding to a normal matched DNA.9–12 However, a recent study13 using single-molecule imaging by atomic force microscopy has shown that the discrimination between specific and non-specific binding of MutS is dramatically higher than described in previous findings, which were based on bulk measurements. Furthermore, the binding affinities are found to be dependent on the type of mismatches,9,10,14 and the local sequence context.15–17 Amongst all the mispaired bases, the G.T mismatch is the most efficiently recognized and repaired,9–12,14 whereas the C.C mismatch is least efficiently repaired by MutS.9–12 After mismatch recognition in an ATP hydrolysisdependent manner, MutS with the assistance of MutL initiates the mismatch repair by activating

0022-2836/$ - see front matter © 2007 Elsevier Ltd. All rights reserved.

40 MutH, which nicks the newly synthesized, unmethylated GATC tract. This is followed by the concerted action of helicase, exonuclease, polymerase, and ligase functions, which restores the parental complementary sequence in the DNA strand.2,3,8,18–20 Currently, most efforts in mismatch repair studies are focused on addressing the finer mechanistic details of the pathway, using the E. coli system as a paradigm and apply the same for the newly discovered eukaryotic MMR. Several studies have addressed the following issues related to MutS. (i) How does the system achieve the specific recognition of mismatches and IDLs? (ii) What is the oligomeric status of MutS when bound to the mismatch site and when bound to the homoduplex stretch of DNA adjacent to the mismatch site? (iii) Following recognition of a mismatch, how is the signal of the mismatch recognition transmitted to a distant landmark site, the GATC tract, such that the downstream components, MutH-UvrD proteins, are activated to accomplish strand-specific correction. Structural studies,21–33 as well as molecular dynamics simulations,34,35 have revealed a sequencedependent deformation of the mismatched base-pair as compared to the matched base-pair. However there is very little perturbation in the global structure. 24,36,37 Introduction of unpaired bases induces a distinct alteration in the stacking properties of the neighboring bases,38,39 and thereby introducing a kink into the DNA helical axis.31,40 X-ray crystallography has shown that MutS makes specific contacts with the DNA helix in the vicinity of a mismatch and unpaired base, and generates a sharp kink (∼60°) in the DNA.41,42 Surprisingly, MutS binding has been shown to “unbend” the DNA at the site of mismatch in an atomic force microscopy study.43 The same study revealed that the MutS– DNA complexes exhibit two populations of DNA conformations, bent and unbent, at mismatch sites, suggesting significance of these conformational rearrangements for recognition and repair. Although the propensity of kinking of DNA seems to have a crucial role in the mismatch recognition process,41,42 any alteration in the dynamics of DNA at the mismatch site that relates to local DNA kink might affect mismatch recognition. NMR relaxation measurements have shown differences in dynamics of the normal and mismatched base-pairs.30 By virtue of its selectivity, sensitivity and large temporal range, fluorescence spectroscopy could be used for studying the structure and dynamics of both small and large biological macromolecules in solution.44–47 Here, we present several facets of site-specific dynamic properties of DNA as a consequence of a mismatch or an unpaired base. The hierarchy of dynamics in DNA can be classified by timescale and by molecular structure. We demonstrate the exquisiteness of site-specific fluorescence labeling of several types of DNA using picosecond time-resolved fluorescence techniques. We probed the dynamics of different bases using fluorescence techniques as an alternative approach in describing the base-pairing fluctuation

Altered Base Dynamics in Mismatch DNA

in DNA, which is usually depicted by NMR as the H-exchange rate,22,24 and the C-13 relaxation rate.30 To compare the dynamics at a mismatch and a nonmismatch site, we used 2-aminopurine (2-AP), a fluorescent analogue of adenine that forms normal Watson–Crick base-pairs with thymine,48,49 as a reporter next to either a G.T or a C.C mismatch or a T-insertion loop, and at several other non-mismatch positions. The design in this setting is such that 2-AP remains paired with its complementary base T and reports changes in strand dynamics as a function of neighboring local mismatch/insertion loop in cis. The alterations in the stacking and motional dynamics of the bases near mismatched and unpaired thymine were determined using time-domain fluorescence measurements. Our results show that the motional dynamics of bases proximal to either a G.T mismatch or a T-insertion site are enhanced. Interestingly, in the same set of sequence context, the level of enhancement next to a C.C mismatch that is inefficiently repaired by MMR is rather low and insignificant. How this altered dynamics of the bases play a crucial role along with the structural variations in the context of MutS binding is the subject of this work. The results are discussed in the light of current models of MutS recognition of base mismatches/insertion loops.

Results Studies on mismatched (G.T) and unpaired base (T-bulge)-containing DNA Steady-state fluorescence We used 25mer DNA duplexes having a G.T mismatch at the center and 2-AP placed next to the thymine of the G.T pair and two or three nucleotides away from the G.T mismatch (Table 1). Firstly, we observed a red shift in the emission peak of 2-AP when placed next (sample GT(±1)) to the G.T mismatch (379 nm) as compared to other sites (374 nm) in the same duplex DNA (Table 2). A similar difference was seen when comparing a mismatched (sample GT(+1)) and a normal (sample AT(+1)) DNA. This suggests that the environment around the base next to the mismatch is more polar when compared to other positions. Interestingly, the emission peak of 2-AP that is placed next to C.C mismatch (373 nm) was comparable to that of the normal DNA (sample AT(±1); Table 2), suggesting equivalence in their polarities. The uniqueness in the environment of the base next to the mismatch was further probed through its accessibility to solvent by quenching the fluorescence of 2-AP by acrylamide. The bimolecular quenching constant, kq estimated from standard Stern-Volmer plots (not shown) reveals that 2-AP placed next to the G.T mismatch or T-bulge(±1, ±2 and ±3) has a significantly higher level of solvent exposure when compared to equivalent locations next to matched pairs (Table 2), confirming the uniqueness of the base-pairs in the

41

3.20

376 1.25

1.00 3.50 0.152 0.80

374

2.05

374 1.64

1.43 2.60 0.143 0.58

373

2.10

374 1.66

1.40 2.90 0.142 0.53

373

4.15 2.90 1.56

372 4.83 371 4.80 372 1.80

4.50 8.35 0.075 0.80 4.50 7.60 0.065 0.61 1.25 5.06 0.111 0.42

374 374 377

1.80 2.20

373 1.90 371 1.75

1.30 5.50 0.133 0.48 1.26 6.50 0.123 0.65

377 375

2.06 1.45 2.41

373 2.54 372 2.53 374 1.64

2.00 5.34 0.140 0.78 2.00 5.10 0.131 0.65 1.25 6.14 0.117 0.49

375 375 379

374 1.70

2.30 2.20

1.20 6.01 0.113 0.43 1.10 5.05 0.121 0.67

373 1.50

379 375

λemmax (nm)a Fluorescence Intensity (a.u)b kq X109 (M−1s−1)c rdss ϕmean

λemmax (nm)a Fluorescence Intensity (a.u.)b kq X109 (M−1 s−1)c

−MutS

+MutS

λem, maxima of fluorescence emission spectra; kq, bimolecular quenching constant, rss, steady-state fluorescence anisotropy; ϕmean, average rotational correlation time. Maximum error: a ±2 nm, b b5%, c ∼10%, d ∼2%.

3.00

375 1.28

1.02 3.45 0.154 0.85

375

T A* T A* T A* T A* T A* T A* T A*

T A*

G T

T A*

T A*

T A*

T 5′ 3′…..T A…3′ 5′…..A

T A*

T A*

____ T

ΔT(+1) ΔT(+2) ΔT(+3) ΔT(−2) ΔT(−1)

3′ T 5′…A Sequences

where r0 is the initial anisotropy. By using the mean fluorescence lifetime (τm in Table 3) we have calculated ϕmean from Perrin's equation (Table 2). ϕmean provides an overall measure of motional freedom of 2-AP. A significantly shorter value of ϕmean obtained for 2-AP placed next to the G.T mismatch (Table 2) indicates an increased level of dynamics when compared to other positions. Thus, we see that all the observations, namely fluorescence emission peak red-shift, acrylamide quenching, steady-state anisotropy (rss) and rotational correlation time (ϕmean), mentioned above indicate the special nature of the base next to the G.T mismatch (GT(±1)). We further note that this special nature of the base next to the mismatched pair was independent of

GT(+1) GT(+2) GT(+3)

rss ¼ r0 =ð1 þ H =/Þ

GT(−2) GT(−1)

vicinity of the G.T mismatch. 2-AP placed next to a C.C mismatch showed reduced solvent accessibility compared to similar locations in G.T mismatch/ Tbulge (Table 2). Next, we used fluorescence anisotropy to infer the position-dependent motional freedom of 2-AP. We observed that the steady-state anisotropy (rss) was significantly lower (0.117) when 2-AP was placed next to the G.T mismatch as compared to other positions in the same duplex (0.13–0.15, Table 2). This might indicate an enhanced level of rotational dynamics of 2-AP when located next to the G.T mismatch. However, we recognize that rss is controlled by rotational correlation time (ϕ) and fluorescence lifetime (τ) through Perrin's equation:

Sample names

A*, 2-aminopurine; ___, no nucleotide.

Table 2. Steady-state fluorescence parameters associated with the mismatch or unpaired base-containing DNA

T…5′ 3′..T A…3′ 5′..A

CC(−1)

C C

CC(+1)

T...5′ 3′..T A ..3′ 5′..A

AT(−1)

A T

AT(+1)

Table 1. Sequences of oligonucleotides used in this study

T A*

T...5′ A ..3′

Altered Base Dynamics in Mismatch DNA

42 Table 3. Parameters associated with the decay of fluorescence intensity and fluorescence anisotropy of 2-AP in G.T and C.C mismatch, unpaired T and matched A.T pair containing DNA Sample name

GT(−2) GT(−1) 3′…T 5′…A

T A*

τ1 α1 τ2 α2 τ3 α3 τ4 α4

0.15 0.59 0.65 0.19 2.20 0.17 7.20 0.05

Mean lifetime (ns)c τm ϕ1 β1 ϕ2 β2

Fluorescence lifetime, τi (ns)a Normalised amplitudes, αib

Rotational correlation times, ϕi (ns)d Normalised amplitudes, βie

G T

T A*

T A*

T A*

0.09 0.67 0.80 0.18 2.00 0.10 7.60 0.05

0.07 0.72 0.28 0.14 2.00 0.08 7.50 0.06

0.12 0.67 0.90 0.15 2.70 0.13 7.40 0.05

0.94

0.75

0.71

0.35 0.62 3.20 0.38

0.20 0.65 2.80 0.35

0.12 0.70 2.80 0.30

T...5′ 3′…..T A…3′ 5′…..A

T A*

T A*

0.12 0.53 0.50 0.20 2.20 0.20 7.00 0.07

0.14 0.70 0.56 0.14 2.56 0.10 8.10 0.06

0.93

1.01

0.26 0.62 2.80 0.38

0.28 0.62 2.80 0.38

ΔT(+1) ΔT(+2) ΔT(+3) ____ T

CC(−1) T…5′ 3′..T A…3′ 5′..A

T A*

CC(+1) C C

T A*

AT(−1) T...5′ 3′..T A ..3′ 5′..A

T A*

AT(+1) A T

T A*

T A*

T A*

T A*

0.10 0.64 0.60 0.22 2.50 0.08 8.00 0.06

0.07 0.53 0.28 0.29 2.00 0.11 8.13 0.07

0.10 0.45 0.55 0.21 2.20 0.15 9.60 0.17

0.15 0.41 0.60 0.20 2.20 0.23 9.20 0.17

0.08 0.64 0.46 0.23 2.20 0.10 7.20 0.03

0.09 0.55 0.48 0.36 2.19 0.08 7.50 0.03

0.18 0.45 0.46 0.39 2.11 0.11 7.05 0.05

0.14 0.64 0.41 0.20 2.20 0.10 6.25 0.06

0.92

0.88

0.92

2.17

2.25

0.60

0.64

0.81

0.77

0.35 0.60 3.20 0.40

0.20 0.60 2.80 0.40

0.18 0.62 2.80 0.38

0.30 0.57 3.80 0.43

0.28 0.55 3.80 0.45

0.34 0.60 4.00 0.40

0.40 0.56 3.80 0.44

0.35 0.60 3.40 0.40

0.28 0.60 3.40 0.40

T...5′ A ..3′

A*, 2-AP. τm = ∑αiτi, where mean lifetime αi is the amplitude associated with the fluorescence lifetime τi.. Fluorescence anisotropy at time t, r(t) = r0{β1 exp(–t/ϕ1)+β2 exp(–t/ϕ2)}, where r0 is the initial anisotropy, βi is the amplitude of the ith rotational correlation time ϕi. Maximum errors are ∼10% in the values of τ'sa, α'sb and ϕ'sd and β'se and b5% for τmc (see ‘Methods’).

Altered Base Dynamics in Mismatch DNA

T A*

Sequences

ΔT(−2) ΔT(−1)

GT(+1) GT(+2) GT(+3)

Sample name

GT(−2) GT(−1) 3′…T 5′…A

T A*

T A*

τ1 α1 τ2 α2 τ3 α3 τ4 α4

0.15 0.55 0.60 0.20 2.40 0.15 7.50 0.10

Mean lifetime (ns)c τm ϕ1 β1 ϕ2 β2

Sequences Fluorescence lifetime, τi (ns)a Normalised amplitudes, αib

Rotational correlation times, ϕi (ns)d Normalised amplitudes, βie

ΔT(−2) ΔT(−1)

GT(+1) GT(+2) GT(+3) G T

T A*

T A*

T A*

0.06 0.54 0.60 0.15 2.20 0.15 7.50 0.16

0.07 0.60 0.50 0.15 2.30 0.13 6.60 0.12

0.10 0.60 0.50 0.16 2.20 0.14 7.20 0.10

1.31

1.65

1.21

0.35 0.60 N15.0 0.40

0.20 0.50 N15.0 0.50

0.15 0.45 N15.0 0.55

T...5′ 3′…..T A…3′ 5′…..A

T A*

T A*

0.14 0.50 0.57 0.20 2.30 0.20 6.70 0.10

0.15 0.50 0.50 0.23 2.40 0.12 9.50 0.15

1.17

1.31

0.30 0.55 N15.0 0.45

0.30 0.60 N15.0 0.40

ΔT(+1) ΔT(+2) ΔT(+3) ____ T

CC(−1) T…5′ 3′..T A…3′ 5′..A

T A*

CC(+1) C C

T A*

AT(−1) T...5′ 3′..T A ..3′ 5′..A

T A*

AT(+1) A T

T A*

T A*

T A*

T A*

0.15 0.35 0.65 0.35 2.40 0.15 9.00 0.15

0.12 0.45 0.50 0.22 2.70 0.13 7.50 0.20

0.10 0.40 0.80 0.20 2.80 0.20 9.00 0.20

0.13 0.40 0.80 0.20 2.80 0.20 8.50 0.20

0.05 0.54 0.41 0.22 2.43 0.13 7.05 0.12

0.25 0.68 0.85 0.19 3.15 0.08 7.76 0.04

0.10 0.67 0.82 0.20 2.80 0.09 7.60 0.05

0.10 0.60 0.40 0.20 2.00 0.15 7.60 0.05

1.90

1.99

2.01

2.56

2.48

1.10

0.94

0.86

0.82

0.35 0.60 N15.0 0.40

0.20 0.50 N15.0 0.50

0.15 0.45 N15.0 0.55

0.30 0.58 N15.0 0.42

0.30 0.60 N15.0 0.40

0.35 0.45 10.0 0.55

0.45 0.40 10.0 0.60

0.30 0.60 10.0 0.40

0.30 0.60 10.0 0.40

T...5′ A ..3′

Altered Base Dynamics in Mismatch DNA

Table 4. Parameters associated with the decay of fluorescence intensity and fluorescence anisotropy decay of 2-AP in G.T and C.C. mismatch, unpaired T and A.T pair-containing DNA, when bound to MutS

A*, 2-AP. τm = ∑αiτi, where mean lifetime αi is the amplitude associated with the fluorescence lifetime τi. Fluorescence anisotropy at time t, r(t) = r0{β1 exp(–t/ϕ1)+β2 exp(–t/ϕ2)}, where r0 is the initial anisotropy, βi is the amplitude of the ith rotational correlation time ϕi. Maximum errors are ∼10% in the values of aτ, bα, dϕ, and eβ; and b5% for cτm (see Materials and Methods).

43

44 whether 2-AP was positioned on the 3′ side or on the 5′ side of the G.T mismatch (Table 2). The enhanced level of motional freedom exhibited by 2-AP when placed next to a G.T mismatch was also replicated when it was placed next to an unpaired thymine (unpaired T samples ΔT(±n), Table 1). The value of ϕmean for 2-AP placed next to the unpaired T was significantly smaller (0.42 ns) when compared to other locations (Table 2), indicating increased flexibility. The observed red shift in the emission maximum for the sample ΔT(+1) when compared to ΔT(+2) and ΔT(+3) also supports the model of increased polarity of bases next to the unpaired base similar to the G.T mismatch. The bimolecular quenching constant kq, which is associated with quenching of fluorescence by acrylamide, was estimated from standard Stern-Volmer plots and shows that 2-AP placed next to the unpaired T (sample ΔT(+1)) is associated with significantly higher values of kq when compared to 2-AP placed next to A.T (Table 2). This indicates that the base next to an unpaired T is associated with an increased level of solvent exposure when compared to a base next to a matched base-pair such as an A.T pair. Furthermore, in the case of the unpaired T, 2-AP placed even two or three bases away showed an enhanced level of solvent exposure (samples ΔT(+2) and ΔT(+3) in Table 2). This could be the result of the extrahelical nature of bases near a T-bulge seen as hyperactive sites when probed by 1,10-phenanthroline footprinting.50 NMR studies have shown that, unlike that of the A-bulge that stacks into a double helix irrespective of sequence context,38 a T-bulge exhibits significant extrahelical character.39 Time-resolved fluorescence Fluorescence intensity decay kinetics of 2-AP in all the double-stranded DNAs studied here showed the presence of at least four lifetime components with a mean lifetime (τm) of ∼1 ns (Table 3). The shortest lifetime component in the range of 0.05–0.2 ns, which is absent from single-stranded DNAs, has been, in general, taken as an indicator of basepairing of 2-AP with thymine.45,47,51–53 An increase in the level of stacking with neighboring bases concomitant with base-pairing is argued as the source of this lifetime component, and the value of this lifetime and its amplitude are taken as indicators of the strength of these interactions.45,47,51 Furthermore, electron and hole transfer from 2-AP to other bases in the sequence also modulates its excited state lifetime, and the charge transfer efficiency depends upon both the identity and distance of other bases from 2-AP in the sequence.54 In the majority of constructs used in this study, 2-AP is flanked by adenine bases on both sides. Stacking with adenine was seen to alter the 2-AP photophysics due to extensive mixing of their ground states,55 which complicates the analysis of the timeresolved components in terms of base dynamics. A study using Streak Camera detection56 has shown the presence of an ultrafast lifetime of ∼10 ps, which

Altered Base Dynamics in Mismatch DNA

was hypothesized to be due to hole transfer from 2AP to nearby guanine bases. Thus, in mixed sequences such as those used in this work, interpretation of changes in the lifetime is far from unique and hence we refrain from interpreting them except in some special situations (see below). In fact, the observed uncorrelated variation of the steadystate fluorescence intensity and the mean lifetime τm (Tables 2 and 3) is a clear indication of the presence of unresolved lifetime component(s). The value of the shortest lifetime component, observed in our photon counting set-up, which has a time resolution of ∼30 ps, was ∼70 ps in the samples GT(±1) (2-AP next to a G.T mismatch), CC(±1) (2-AP next to a C.C mismatch) and ΔT(±1) (2-AP next to an unpaired T) as opposed to 120–150 ps in other samples (see Table 3). (See Materials and Methods for validation of estimation of time constants in the range 50–200 ps.) In the previous section, we showed that, in GT(±1) and ΔT(±1), 2-AP is more dynamic and associated with increased exposure to solvent when compared to normally matched samples. Thus, it is tempting to correlate the shorter lifetime to the special nature of 2-AP in these samples. The reduction in values of both the shortest lifetime and the mean lifetime (τm) in GT(+1) when compared to GT(+2) and GT(+3) could have arisen from an enhanced level of motional dynamics. The significant increase in the value τm of ΔT(+2) and ΔT(+3) when compared to that of ΔT(+1) (Table 3) warrants some explanation. Since this increase is largely due to the high amplitude of the longest lifetime component (which is taken to correspond to the population of solvent-exposed probe), this observation could reflect unstacking of 2-AP with the neighboring adenine bases due to the presence of the nearby unpaired T. Motional dynamics of bases in mismatch or T-bulge-containing DNA as revealed by time-resolved fluorescence anisotropy Picosecond timescale motional dynamics of bases were measured by monitoring the decay of fluorescence anisotropy of 2-AP located at various positions. Anisotropy decay kinetics could be fit satisfactorily to a sum of two exponentials. The faster rotational correlation time (ϕ1 or ϕ fast) represents the local motional freedom of the probe with respect to the DNA strand and the slower correlation time (ϕ2 or ϕslow) represents a combination of segmental mobility of the DNA strand with respect to the overall length of DNA and the global tumbling dynamics of the entire chain.44,45,47 Rotational diffusion of 2-AP placed next to a G.T mismatch (either side) showed enhanced dynamics as compared to A.T base-pair (Figure 1(a)). In contrast, the bases next to the C.C mismatch were seen to be as rigid as A.T base-pair (Figure 1(a)). The enhancement in the dynamics of 2-AP decreased significantly as we move two or more bases away from the G.T mismatch (Figure 1(b)). Analyses of the decay kinetics revealed an enhancement in the rate of

45

Altered Base Dynamics in Mismatch DNA

local motion of 2-AP placed next to the G.T mismatch (ϕ1 ≈ 0.12 ns) when compared to that of 2-AP located two or three bases away from the mismatch (ϕ1 ≈ 0.3 ns) (Table 3). (See Materials and Methods for validation of estimation of time constants in the range 50–200 ps.) Furthermore, the amplitude (β1) associated with the shorter correlation time (ϕ1) for 2-AP placed next to a G.T mismatch was found to be slightly higher (70% as compared to 62%) when compared to other locations, indicating reduced level of restriction of internal dynamics.57,58 In contrast, bases next to a C.C mismatch were not seen to have enhanced flexibility (ϕ1 = 0.34–0.40 ns) when compared to the bases next to an A.T pair (ϕ1 = 0.35–0.28 ns) (Table 3). This indicates differences in the dynamics of the neighboring bases of different mismatches. The longer correlation time (ϕ2) was found to be ∼3 ns for all the samples, and this is likely to represent the overall tumbling dynamics of double-stranded DNA. An analogous increase in flexibility was found in bases neighboring an unpaired T (Figure 1(c)). 2-AP positioned on either side of an unpaired T showed a shorter value of ϕ1 (∼0.2 ns) when compared to other locations (ϕ1 ∼ 0.3 ns, Table 3). Thus, we get an unambiguous picture of the enhanced level of internal dynamics of 2-AP when placed next to either a G.T mismatch or an unpaired T. This interpretation is consistent with NMR studies involving 13 C relaxation studies, where it was shown that displacement of the wobble-paired G in a G.T mismatched duplex causes dynamic perturbations in its flanking nucleotides.30 Dynamics of MutS–DNA complexes

Figure 1. Decay kinetics of fluorescence anisotropy of 2-AP in (a) A.T, G.T and C.C base-pair-containing DNA (AT(+1), green; GT(+1), red; CC(+1), blue). (b) G.T mismatch-containing DNA (GT(+1), red; GT(+3), blue). (c) Unpaired thymine-containing DNA (ΔT(+1), red; ΔT(+3), blue).

After establishing the motional dynamics of bases in mismatch-containing DNA, we proceeded further to explore the dynamics of bases in MutS–DNA complexes. Preformed DNA duplexes were allowed to interact with excess of MutS (1:5 stoichiometry) and then these complexes were subjected to steadystate as well as time-resolved fluorescence analysis. The steady-state fluorescence intensity of 2-AP increased considerable on binding to MutS (Table 2). The percentage increase in the intensity was found to be highest for 2-AP placed next to either the G.T mismatch or the unpaired T (Table 2). This observation was further supported by time-resolved fluorescence studies, as seen from the extent of increase in τm (Tables 3 and 4). Thus, we infer that MutS interacts with the bases near the mismatches in a specific manner. Quenching of fluorescence by acrylamide showed that values of the bimolecular quenching constant kq decreased by a factor of 2–3 for all the DNAs with a mismatch or an unpaired base following binding of MutS (Table 2), indicating protection offered by the bound MutS. Similar to the naked DNA, the MutS-bound DNA showed a four exponential decay profile for fluorescence intensity of 2-AP (Table 4). Heterogeneity of MutS–DNA complexes seen in atomic force microscopy images43 could also have contributed to the

46

Altered Base Dynamics in Mismatch DNA

heterogeneity observed in fluorescence decay kinetics. MutS binding-induced enhancement of the mean lifetime (τm) could be explained mainly from the increase in the amplitude (α4) of the longest lifetime (τ4). This increase was found to be significant when 2-AP was placed next to either the G.T mismatch (from 0.07 to 0.16) or the T-bulge (from 0.07 to 0.20, Table 4). The extent of increase in τm was found to be higher for the base placed 5′ to the G.T mismatch when compared to the 3′ side. Increase in the excited state lifetime could be the result of inhibition of internal motion-induced non-radiative decay process apart from a reduction in the level of stacking with neighboring bases subsequent to the binding of MutS. Such a model is supported by the MutS-induced reduction in the amplitude associated with the internal motion of 2-AP monitored through fluorescence anisotropy decay (see below). However, the observed increase in the amplitude (α4) of the longest lifetime component suggests that the increase in τm could be the result of the base acquiring extra-helical character. Time-resolved fluorescence anisotropy In order to determine the effect of binding of MutS on the motional dynamics of the bases and the backbone of DNA, we monitored fluorescence anisotropy decay of 2-AP. The mismatch (G.T and C.C) or unpaired T-containing DNA was allowed to interact with excess MutS, and depolarization of fluorescence anisotropy of the complex was measured following a pulse excitation. Typical traces of fluorescence anisotropy decay are shown in Figures 2 and 3. Upon MutS binding, we observed striking changes in the amplitude β1 associated with the local motion (represented by ϕ1). There was a significant reduction in β1 for 2-AP placed next to either the G.T mismatch or the unpaired T (Figures 2 and 3; Tables 3 and 4). The extent of MutS binding-induced decrease in β1 was gradual as the position of 2-AP was moved away from either the G.T mismatch or the unpaired T. The decrease in β1 became negligible when 2-AP was positioned just two bases away from either the G.T mismatch or the unpaired T similar to normally matched samples (Figures 2 and 3; Tables 3 and 4). Even though the bases next to C.C were found to be less dynamic (ϕ1 = 0.34 ns as compared to the G.T mismatch (ϕ1 = 0.12 ns), there was significant reduction in the amplitude (β1) associated with the local motion (ϕ1) of the bases in the MutS–DNA complex (Table 4). This indicates, once again, that MutS has specific interaction with the base at the mismatch site very locally, although the binding footprint is 20 base-pairs wide.9,10,16,50 Furthermore, it is interesting to note that, despite the decrease in the value of β1, the rate of internal motion (ϕ1) is almost unchanged. The reduction in the amplitude (β1) of the local motion is a reflection of the hindrance around 2-AP following the binding of MutS to the mismatch site. The increase in the value of the longer correlation time (ϕ2) from ∼3 ns (Table 3) to N15 ns (Table 4), upon binding of MutS, is due to the increase in the

Figure 2. Decay kinetics of fluorescence anisotropy of 2-AP in G.T mismatch-containing DNA upon MutS binding. (a) GT(+1), red; GT(+2), blue; GT(+3), green. (b) GT(−1), red; GT(−2), blue; AT(−1), green.

overall size and the concomitant reduction in the rate of global tumbling dynamics. The limit of ∼15 ns arises due to the time-window limited by the excited state lifetime. There was no significant change in the motional dynamics of 2-AP in the presence of ATP (data not shown). This might indicate that ATP hydrolysis-induced sliding of MutS along the DNA strand8,11 has no significant effect on the nanosecond timescale dynamics of the DNA bases, but may have influence on much slower motions in the system.

Discussion Altered dynamics around the site of a mismatch/unpaired base The main aim of this work was to search for a plausible mechanism by which MutS senses the presence of either base mismatches or an unpaired

Altered Base Dynamics in Mismatch DNA

Figure 3. Decay kinetics of fluorescence anisotropy of 2-AP in unpaired thymine-containing DNA upon MutS binding. (a) ΔT(+1), red; ΔT(+2), blue; ΔT(+3), green. (b) ΔT(−1), red; ΔT(−2), blue; AT(-1), green.

base (bulge/loop) amongst several hundred normal base-pairs. This seemingly needle-in-a-haystack kind of search is likely to be aided by alterations in either the structure or dynamics or both, localized at or around the site of a mismatch or unpaired base loop. In this work, we have used various steadystate and time-domain fluorescence parameters to gain insights into the dynamics of 2-AP located close to either a G.T mismatch or a T-bulge, taken as representative sites that are repaired efficiently by MMR and contrasted the same with that of C.C, which is poorly repaired. The overall picture, as justified by the discussion below, is that the internal dynamics at/around the site of a mismatch or an unpaired base is enhanced significantly compared to matched base-pairs, and MutS could very well use this as the cue in the search for mismatches and deletion/insertion loops. This theme is highly consistent with the observations that MutS has a higher propensity to bind any distortions in the

47 DNA helix and the same may in fact stem from the enhanced motional dynamics associated with such targets. The classical, simple example of this notion being the ends of normal duplex DNA to which MutS binds avidly,13 and what has been shown by us recently as highly dynamic part of a normal Watson–Crick duplex DNA.47 Introduction of a G.T mismatch (but not a C.C mismatch) or an unpaired T was found to change the stacking property and the dynamics of bases adjacent to the site of mismatch/bulge. This conclusion is the result of several observations, such as fluorescence emission spectral peak position, fluorescence intensity, mean fluorescence lifetime, solvent accessibility inferred from bimolecular quenching constant. and steady-state and time-resolved fluorescence anisotropy (Tables 2 and 3; Figures 1–3). Of all the observations listed above, steady-state and time-resolved fluorescence anisotropy could be taken as the most direct and revealing in their information content. Enhancement of the angular excursion of the internal motion of 2-AP on either side of the mismatch/bulge is shown clearly by the fluorescence anisotropy data (Tables 2 and 3; Figure 1). Enhancement of the dynamics of bases adjacent to the site of mismatch/bulge could imply a corresponding enhancement in the dynamics of the mismatched base-pair itself. An earlier study of this issue used 2-AP itself as a mimic of a mismatch base and found significant changes in the dynamics when 2-AP was presented with a mismatch in the opposite strand.59 However, interpretation of this result is limited by the fact that that 2-AP.T pair itself is treated as a mismatch by MutS.60–62 In the present work, we address the question of whether the enhanced dynamics caused by the mismatch/bulge is localized. Our data show clearly that enhancement is highly localized and decays rapidly as we move by two base-pairs either side of either the G.T mismatch or the unpaired T (Figure 1(b) and (c); Tables 2 and 3). However, the case of an unpaired T is more complex when compared to a G.T mismatch. Alteration in both motional dynamics (scored by steady-state and time-resolved fluorescence anisotropy) and solvent exposure (inferred from the bimolecular quenching constant kq) of base is highly localized near the G.T mismatch. In contrast, in the samples with an unpaired T, the motional dynamics is highly localized, similar to a G.T mismatch, whereas the level of solvent exposure remains high for several bases away from the site of the unpaired T (Table 2). These observations are strikingly similar to the observation of hypersensitivity of an unpaired T (but not a G.T mismatch) to cleavage by either 1,10-phenanthroline-copper or osmium tetroxide.50 NMR studies of DNA containing G.T and other mismatches led to the conclusion that the alteration in the structure is subtle, and is localized to the site of the mismatch, but subtly dependent on sequence context effects.21–32 This renders the recognition rules of mismatches by MutS rather complex, although a variety of models have been put forward

48 to rationalize the same.33,41,42,63 One of the most paradoxical aspects of such models was the notion that more efficiently recognized/repaired mismatched base-pairs by the MutS system appeared to be thermodynamically more stable,2,64 thereby implying that localized stable rather than dynamic distortion in base-mismatch provides the required cue for recognition by MutS-MutL. Our results try to assuage and perhaps even clarify this paradox by showing that in addition to localized structural distortions in a mismatch, enhanced dynamics of normal base-pairs in the vicinity of a mismatch might be relevant for such modulations in MutS recognitions. In this scenario, alterations in the base dynamics at the site of a mismatch/bulge found in the present work could be used to build a realistic model of mismatch recognition. The absence of such enhanced dynamics around a C.C mismatch (Figure 1(a); Table 3), which is repaired less efficiently, strongly endorses the relevance of the base dynamics associated with adjacent base-pairs in mismatch recognition. In general, recognition of a specific binding site on DNA by proteins is hypothesized to result from a one-dimensional search along the DNA using the weak and non-specific binding of the protein.65,66 This reduction in the dimensionality (from 3D to 1D) would enhance the rate of search dramatically.67 MutS could easily fit such a model, due to its ability

Altered Base Dynamics in Mismatch DNA

to bind and track DNA through relatively weak and non-specific interactions.9–11 Thus, we could propose a hypothesis wherein MutS scans the DNA through a 1D tracking mode and the enhanced dynamics of bases around the site of mismatch/ bulge would herald the cue of a mismatch. We speculate that stacking of the conserved residue Phe36/Phe39 of MutS (E. coli/Thermus aquaticus) with the mismatched base-pair (Figure 4) as observed in the reported crystal structures41,42,33 could have been further aided by the enhanced dynamics associated with mismatched bases (see later). In addition, the enhanced flexibility of the base might give room to facilitating the observed hydrogen bonding pattern of Glu38/Glu41 (E. coli/ T. aquaticus) with one of the mispaired bases,10,63 thereby inducing the ATPase activity of DNAbound MutS.63 The ultrafast (100–200 ps) timescale of the internal dynamics of bases could match with rapid conformational fluctuations in the DNAbinding pocket of MutS protein, which in turn might enable rapid search for its specific binding site. Motional dynamics of DNA bases have been implicated in functions such as long-range charge transfer in DNA,68 and DNA–protein interactions.69 The efficiency of mismatch repair is higher when mismatches such as G.T are flanked by GC-rich sequences.14 It would be interesting to see whether such counterintuitive results have their origins in

Figure 4. Structure of mismatch DNA-binding site of MutS (E. coli), highlighting the interacting amino acid residues (Phe36 and Glu38) in yellow. DNA with a G.T mismatch is presented according to strand polarity. The π-stacking and H-bonding interactions are indicated by light blue and red dotted lines, respectively. The structure is drawn using RAS-MOL software and the PDB file with accession code 1E3M.

49

Altered Base Dynamics in Mismatch DNA

any correlation between the level of base dynamics and the GC content of the flanking sequence. Such possibilities are to be seriously considered now in view of recent novel findings demonstrating the occurrence of sequence-dependent DNA dynamic changes.47 More of such studies are in progress. MutS binding causes changes in the dynamics at the mismatch site The crystal structure of the DNA–MutS complex revealed that only one subunit of the MutS dimer has specific mismatch-binding contacts, involving exclusively a Phe-X-Glu motif at the N-terminal mismatch-recognition domain (Figure 4). 10 The phenylalanine (Phe36) was observed to stack with thymine of either the G.T mismatch or the T-bulge at the 3′ side. This thymine is unstacked out from its neighboring bases and rotated out of the minor groove by ∼3 Å.41,42,69–71 As a result, the protein with Phe36 is wedged into the minor groove, where the mismatch resides and kinks the DNA duplex by 60° towards the major groove, creating a discontinuity in the DNA duplex.41,42,69 Does the interaction of bases with MutS occur in a localized fashion around the mismatch or encompass the entire binding domain? This question becomes relevant and interesting when we recognize that the binding domain of MutS is ∼20 base-pairs, as shown by footprinting studies,50 and by crystal structures.41,42 Our observations based on both the increase in the fluorescence lifetime and hindrance of the internal motion of 2-AP subsequent to binding of MutS to either mismatched or unpaired T DNA show clearly that the changes are highly localized to the site of mismatch/bulge (Tables 3 and 4). Furthermore, MutS-induced hindrance to the internal motion of 2-AP was more pronounced at the 3′ side of the mismatch/bulge when compared to the 5′ side, which is in line with the observed stacking of Phe36 at the 3′ side (Figure 4).41,42 Similarly, the observed asymmetry in the MutS-induced increase in τm of 2AP placed on either side of the G.T mismatch (samples GT(+1) and GT(−1) in Tables 3 and 4) could be the result of the base at the 5′ side of the thymine of G.T mismatch becoming extra-helical. Again, the insertion of Phe36 of MutS into the 3′ side of thymine could be the origin of this effect. Overall, we get a picture of specific interaction between MutS and the bases around the mismatch/bulge, despite the fact that the protein covers a region of ∼20 basepairs centered on the mismatch/bulge. It is striking to note that the gradation in the solvent exposure (with 2-AP adjacent to the mismatch/bulge being more exposed when compared to even one or two positions away) observed in the absence of MutS (Table 2) was totally wiped out (all the positions being protected from solvent) in the presence of MutS in spite of differential interaction of MutS with the mismatch (Table 4). We propose that intrinsic dynamics prevalent as a sharp gradient in the vicinity of a mismatch base-pair along with structural distortion of a mismatch might serve as a

“composite cue” for initial MutS recognition, following which protein stably covers about 20 base-pairs in the recognition complex. Binding of MutS to the normal duplex results in definite changes in at least some of the fluorescence parameters, such as the long correlation time, which is indicative of protein binding (Tables 3 and 4). This could be due to nonspecific binding of MutS apart from possible recognition of T.2-AP as a mismatch. Several studies have implicated that mismatchbound MutS exists as a tetramer, while nonspecifically bound MutS remains as a dimer.72–74 Such a structural switch requires conformational changes in MutS that is on a mismatch base-pair. Specific interaction between a mismatch/bulge and MutS scored in our observations could provide the conformational changes needed for dimer to tetramer conversion. An altered conformation of MutS while bound to a mismatch/bulge could lead to polymerization of MutS on DNA, as required in one of the models of communication between mismatch recognition and DNA excision by MutH bound at a distal site.75,76

Materials and Methods Materials Bradford reagent was from Bio-Rad (Hercules, CA, USA), 2-AP was from Sigma (USA). Oligonucleotides were from DNA Technology (Aarhus C., Denmark). The Sep-Pak C-18 cartridge was from Waters Corporation (Massachusetts, USA). DNA substrate All the oligonucleotides used in this study were purified by electrophoresis in a denaturing 10% polyacrylamide gel containing 8M urea. The full-length oligonucleotide was excised from the gel and eluted into autoclaved buffer (10 mM Tris–HCl (pH 8.0), 1 mM EDTA) by diffusion, followed by desalting through a Sep-pak C-18 cartridge.77 The final concentration of purified DNA was determined by measuring the absorbance of an aliquot at 260 nm. The concentrations expressed pertain to that of molecules. The DNA substrates used in all the assays were a single G.T mismatched or thymine unpaired 25mer duplex (T-bulge) with 2-AP at different positions, the names and their corresponding sequences are given in Table 1. Complementary strands (see Table 1) were annealed at a ratio of 1:1 (50 μM each strand in a total volume of 50 μl) in 20 mM Tris–HCl (pH 7.5), 10 mM MgCl2 by heating the sample for 5 min at 90 °C, followed by slow-cooling to room temperature. Analysis of an aliquot of annealed sample by native PAGE revealed that the annealed duplex was well resolved from the singlestranded controls and annealing was achieved with an efficiency N90%. Protein purification The MutS clone was obtained from Dr Leroy Worth, NIEHS. The mutS gene is in the His-tag expression vector

50

Altered Base Dynamics in Mismatch DNA

pQE30. The protocol followed to purify MutS has been described.78 The His tag was not cleaved from the protein, as it does not seem to alter the biochemical properties of MutS.76,79 Protein concentrations were measured using the Bradford reagent. All the protein concentrations expressed pertain to protein dimers. MutS–DNA complex formation The MutS–DNA complexes were formed by adding 0.2 μM duplex DNA to 1.0 μM MutS in buffer A (20 mM Tris–HCl (pH 7.5), 50 mM KCl, 10 mM MgCl2, 1 mM DTT), followed by incubation of the sample for 10 min at room temperature (∼22 °C). Fluorescence measurements and data analysis The DNAs were taken up in buffer A (20 mM Tris–HCl (pH 7.4), 10 mM Mg2+, 50 mM K+) and excited with 310 nm light. The fluorescence emission maximum was ∼380 nm. Steady-state fluorescence measurements were carried out using a SPEX fluorolog FL111 fluorimeter (SPEX Industries Inc, NJ, USA) in T-format, with the excitation wavelength set at 310 nm monitoring the changes in fluorescence intensity at 380 nm. Fluorescence anisotropy was measured by monitoring the emission at polarizations parallel with and perpendicular to that of excitation simultaneously by the use of the T-format optical arrangement. The steady-state anisotropy is defined as: rss ¼

II  I8 II8 þ 2I8

ð1Þ

where III and I⊥ are the fluorescence intensity measured with the emission polarizers kept parallel with and perpendicular to the excitation polarizer, respectively. Fluorescence quenching measurements Solvent exposure of 2-AP was estimated by monitoring the quenching of fluorescence by acrylamide. SternVolmer plots (I0/I versus [Q]) were fit to the following equation: I0 ¼ ð1 þ KSV ½QÞ I

ð2Þ

where I0 is the fluorescence intensity of the 2-AP in DNA in the absence of quencher, I is the fluorescence intensity of the sample in the presence of a concentration [Q] of the quencher, KSV is the Stern-Volmer constant (association constant) and could be obtained from the slope of the curve. The Stern-Volmer constant is equal to τ0kq, where kq is bimolecular quenching constant and τ0 is fluorescence lifetime of the fluorophore in the absence of quencher. [Q] was varied in the range of 0–300 mM acrylamide. Time-resolved fluorescence measurements Time-resolved fluorescence intensity and anisotropy of 2-AP in DNA was measured by employing a CW passively mode-locked frequency-doubled Nd-YAG laser (Vanguard, Spectra Physics, USA) driven rhodamine 6G dye tunable laser, which generates pulses of width ∼1 ps

(repletion rate 4 MHz). 2-AP in DNA was excited by using the second harmonic output (310 nm) of an angle-tuned KDP crystal. Fluorescence decay curves were obtained by using a time-correlated, single-photon counting setup, coupled to a micro-channel plate photomultiplier (model 2809u; Hamamatsu Corp.). The instrument response function was obtained at 310 nm using a dilute colloidal suspension of dried non-dairy coffee whitener. The halfwidth of the instrument response function was ∼40 ps, and the time per channel was 40 ps. The samples were excited at 310 nm, and the fluorescence emission was collected through a 340 nm cut-off filter followed by a monochromator at 380 nm with a collection bandwidth of 10 nm. The cut-off filter was used to prevent scattering of the excitation beam from the samples. The number of counts in the peak channel was at least 10,000. In fluorescence lifetime measurements, the emission was monitored with the polarizer at the magic angle (54.7°) to eliminate any contribution from the decay of anisotropy. In time-resolved fluorescence anisotropy measurements, the emission was collected at directions parallel with (III) and perpendicular to (I⊥) the polarization of the excitation beam. The anisotropy was calculated as: r ð tÞ ¼

III ðtÞ  I8 ðtÞGðEÞ III ðtÞ þ 2I8 ðtÞGðEÞ

ð3Þ

where G(λ) is the geometry factor at the wavelength λ of emission. G(λ) of the emission collection optics was determined in separate experiments using a standard sample of 2-AP for which the rotational correlation time was 0.1 ns and the fluorescence lifetime was 12 ns. Time-resolved fluorescence decay data were fit to a function that is a sum of discrete exponentials: X IðtÞ ¼ ai expðt=si Þ ð4Þ i

where ∑αi = 1, by the iterative deconvolution method.80 The correction factors for αi and τi were determined the Marquardt method in non-linear least-squares analysis.81 Numerical calculation of the convolution integrals for intensity and partial derivatives were done with the Grinvald-Steinberg recursion equations.80 The mean lifetime, τm = ∑αiτi, gives us information on the average fluorescence yield of the system. Errors in the values of time-resolved fluorescence parameters arise in several ways: (i) measurements on several samples; (ii) several measurements on a sample; and (iii) several sets of parameters obtained from analysis of a single measurement. The values are given in the Tables. In many cases, uncertainty in the estimated values arises largely due to the inability to select a given set of recovered parameters from other sets when all the sets satisfy the fitness criteria (such as χ2, residual distribution, etc) more or less in a similar manner. This is true especially in multiparameter fits of decay curves with relatively low values of signal/noise. In such situations, the mean lifetime (τm) varies significantly less when compared to individual decay parameters such as lifetime and amplitude, since τm represents the area under the Intensity versus Time curve. In other words, one could get the same value of mean lifetime for various sets of individual decay parameters as a result of compensating the variation of one parameter with respect to another. This explains why the uncertainty in the mean lifetime is significantly less when compared to the variation of individual parameters.

51

Altered Base Dynamics in Mismatch DNA T ime-resolved anisotropy decays were analyzed with the following equations. 1 Ill ðtÞ ¼ IðtÞ½1 þ 2rðtÞ ð5Þ 3

1 I8 ðtÞ ¼ I ðtÞ½1  rðtÞ 3

ð6Þ

rðtÞ ¼ r0 fh1 expðH =f1 Þ þ h2 expðs=f2 Þg

ð7Þ

where r0 is the initial anisotropy, βi is the amplitude of the ith rotational correlation time (ϕi) such that ∑βi = 1. The shorter component ϕ1, representing the internal motion of 2-AP, could be modeled as due to hindered rotation.51,52 We have estimated, in several samples, lifetimes and rotational correlation times in the range of 0.1–0.3 ns. Since these values are very close to both the width of the instrument response function and the time/channel used, both of which are ∼40 ps, their reliability might seem uncertain. In order to check the reliability of estimation of such short time constants, we measured the rotational correlation time (ϕ) of a small molecule, N-acetyltryptophanamide in glycerol/water mixtures with viscosity (η) in the range 1–5 cP. The values of ϕ estimated were 0.061(±0.025) ns, 0.123(±0.036) ns, 0.226(±0.028) ns, and 0.3119(±0.027) ns when the solvent viscosity was 1.0 cP, 2.1 cP, 4.4 cP, and 5.4 cP, respectively, as expected from the Stokes–Einstein relationship (ϕ = ηV/kT), where ϕ is the rotational correlation time of the sphere of volume V and η is the viscosity of the medium, T is the temperature and R is the gas constant= ηV/kT), where ϕ is the rotational correlation time of the sphere of volume V and η is the viscosity of the medium, T is the temperature and R is the gas constant”. –>. Thus, these data attest to the reliability of estimation of the time constants in the range 0.1–0.3 ns (Tables 3 and 4).

Acknowledgements We thank Professor N. Periasamy for providing us with the home-developed software used in the analysis of the time-resolved fluorescence data. This work was funded by the Tata Institute of Fundamental Research, and by the Department of Science and Technology, Government of India.

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