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J. Phys. Chem. B 2006, 110, 25050-25058
Hexamerization of the Bacteriophage T4 Capsid Protein gp23 and Its W13V Mutant Studied by Time-Resolved Tryptophan Fluorescence Aike Stortelder,†,| Johnny Hendriks,‡,⊥ Joost B. Buijs,† Jaap Bulthuis,§ Cees Gooijer,† Saskia M. van der Vies,‡ and Gert van der Zwan*,† Laser Centre VU, Department of Analytical Chemistry and Applied Spectroscopy, Department of Biochemistry and Molecular Biology, and Department of Physical Chemistry, Vrije UniVersiteit, Faculty of Sciences, De Boelelaan 1083, 1081 HV Amsterdam, The Netherlands ReceiVed: July 31, 2006; In Final Form: September 27, 2006
The bacteriophage T4 capsid protein gp23 was studied using time-resolved and steady-state fluorescence of the intrinsic protein fluorophore tryptophan. In-vitro gp23 consists mostly of monomers at low temperature but forms hexamers at room temperature. To extend our knowledge of the structure and hexamerization characteristics of gp23, the temperature-dependent fluorescence properties of a tryptophan mutant (W13V) were compared to those of wild-type gp23. The W13V mutation is located in the N-terminal part of the protein, which is cleaved off after prohead formation in the live bacteriophage. Results show that W13 plays a role in the hexamerization process but is not needed to stabilize the hexamer once it is formed. Furthermore, besides the monomer-to-hexamer temperature transition (15-23 °C and 12-43 °C for wild-type and W13V gp23, respectively), we were able to observe denaturation of the N-terminus in hexameric wild-type gp23 around 40 °C. In addition, with the aid of a recently published homology model of gp23, the lifetimes obtained from time-resolved fluorescence measurements could tentatively be assigned to specific tryptophan residues.
Introduction The protein shells of bacteriophages are remarkably stable and yet dynamic structures. They protect the genome during transfer from one host cell to another and have to be able to withstand harsh environmental conditions and the high pressure of the condensed nucleic acid. The capsid of the dsDNA bacteriophage T4 is a prolate icosahedron that is composed of one major and four minor proteins.1 Bacteriophage T4 is a member of the MyoViridae family that infects the bacterium Escherichia coli. During infection of the host and propagation of the progeny phage, the capsid proteins are synthesized, folded, and assembled into the procapsid, the precursor of the mature capsid. During genome packaging and maturation, the capsid proteins undergo large conformational changes.2 The major capsid protein, gp23, consists of 521 amino acids, including four tryptophan residues at positions 13, 247, 309, and 345. The folding of gp23 requires the assistance of a specialized chaperonin folding machine. The chaperonins are a group of related proteins that belong to the larger family of molecular chaperones. For recent reviews on the structure and function of the different molecular chaperones, see refs 3-7. The chaperonin folding machines are involved in the folding and refolding of a wide variety of proteins. In E. coli, the GroEL-GroES chaperonin complex is responsible for the folding of about 10% of the newly synthesized proteins. Several bacteriophages utilize the GroEL-GroES system for the folding of their own protein * Author to whom correspondence should be addressed. E-mail: zwan@ few.vu.nl. † Department of Analytical Chemistry and Applied Spectroscopy. ‡ Department of Biochemistry and Molecular Biology. § Department of Physical Chemistry. | Current address: Heinrich-Heine-Universita ¨ t Du¨sseldorf, Institut fu¨r Molekulare Physikalische Chemie, Universita¨tsstrasse 1, 40225 Du¨sseldorf. ⊥ Current address: University of Amsterdam, Swammerdam Institute for Life Sciences, Nieuwe Achtergracht 166, 1018 WV, Amsterdam.
structures. Bacteriophage T4 forms an intriguing exception. For as yet unknown reasons, the GroEL-GroES chaperonin system is not able to aid the folding of gp23.8 To circumvent this problem, the phage produces a GroES-related protein, called gp31, which replaces GroES from the complex. This altered GroEL-gp31 complex is able to fold the capsid protein correctly. Recently, this was also demonstrated in vitro using purified proteins.9 We are interested to understand the molecular basis for this difference between the GroEL-gp31 and GroELGroES chaperonin systems. Consequently, the properties of the gp23 capsid protein are also of interest, as it is at the heart of the difference between the two chaperonin systems. This paper focuses on the properties of the gp23 protein structure. The mature bacteriophage T4 capsid contains 930 gp23 molecules arranged in 155 hexamers.1 Little is known about the association of the hexamers. In solution, gp23 can be present in several forms: as a monomer, as a hexamer, or as larger ordered aggregates, so-called polyheads. Hexamerization of wild-type gp23 is temperature-dependent: around 5 °C, gp23 is in monomeric form, but at room temperature the biologically important hexamer is formed.10 The chance of six monomers coming together at the same time is negligible, so dimer or trimer intermediates are expected to play a role as well. However, these may be observed only if their steady-state concentration is significant in a reasonable temperature range. In addition, since we make use of fluorescence spectroscopy, the fluorescence properties of these di- or trimers should also be sufficiently different from those of the monomer and hexamer. Conformational changes in proteins can in principle be monitored using the amino acid tryptophan as an intrinsic fluorescent probe.11-13 The emission characteristics of this amino acid are dependent on the microenvironment, so both lifetimes and emission wavelengths may change when the conformation
10.1021/jp064881t CCC: $33.50 © 2006 American Chemical Society Published on Web 11/16/2006
Tryptophan Fluorescence of Bacteriophage T4 gp23 of the protein changes.14 Whereas in a polar environment the emission maximum is around 360 nm, in apolar surroundings it may go down to 305 nm, for example, if tryptophan is located inside a protein.15,16 Free tryptophan molecules in aqueous solution show multiple fluorescence lifetimes17,18 probably because of the various rotameric conformations they may adopt.19 The wavelength and lifetime of the indole emission are assumed to be dependent on the distance between the amino group and the indole ring.20,21 In proteins, tryptophan residues experience a more rigid environment because of the bulky amino acid chain. In addition, the amino group has become part of the protein backbone through the formation of a peptide linkage and thus will not influence the indole emission. Still, multiple lifetimes have been observed, even in single tryptophan proteins. These seem to be the result of dynamic quenching via electron transfer from indole to a close-lying amide in the protein backbone.22,23 Depending on the distance and orientation of this amide and other closelying charged groups, such a charge-transfer mechanism may lead to a vast range of fluorescence lifetimes. Fluorescence lifetimes might also be affected because of different quenching characteristics of amino acids that surround the tryptophan residue such as tyrosine, histidine, or cysteine.24 The presence of such residues may cause strong differences even between spatially close-lying tryptophan residues. Furthermore, fluorescence lifetimes can be influenced by interactions with solvent molecules. As a result, when multiple tryptophan residues are present, the lifetime distribution becomes increasingly complicated and separate lifetimes for each tryptophan residue can generally not be resolved.25 A recently published model for the structure of gp23 may help with the interpretation.26 This model is based on the assumption that bacteriophage T4 gp24 and HK97 gp5 proteins are similarly folded. However, the sequence similarity of gp23 with these proteins is relatively low. Therefore, investigations of gp23 mutants where tryptophan residues have been removed are useful to gain additional insight in the protein’s structure. Here, a mutant of gp23 is studied, in which one of the four tryptophan residues is replaced by a valine (W13V), reducing the total number to three. This should simplify interpretation of the lifetime distribution and allow a study of the properties of the remaining residues and, indirectly, of the replaced residue as well. The replaced residue W13 is located in a part of the protein that is cleaved off during maturation of the phage and may hence be involved in hexamerization.10,27 In earlier experiments on wild-type gp23, four fluorescence lifetimes were found, and a crude assignment of these four lifetimes was made.28 In this paper, the lifetime assignments are further specified on the basis of steady-state and timeresolved fluorescence experiments of the wild-type and W13V mutant of gp23. The primary focus in this paper, however, is to gain insight into the gp23 monomer-hexamer equilibrium and the relevance for capsid maturation. Experimental Section Preparation of Mutants. For the preparation of bacteriophage T4 gp23 mutants, the gen23-gen31 fragment from the pET2331 plasmid (a gift from L. Black, University of Maryland, Baltimore) was placed in a pMPM-A4 vector.29 With the obtained plasmid p2331A4, gp23 is L-arabinose inducible. Mutagenesis was performed on the p2331A4 plasmid according to the QuikChange method by Stratagene. To make a semirandom mutation of gp23 W13, the necessary degenerate oligonucleotides were prepared by elongating the following
J. Phys. Chem. B, Vol. 110, No. 49, 2006 25051 complementary oligonucleotides: sense: 5′-ACTAAAGCTGAACTTTTGAACAAAVNSAAGCCATTACTGGAAGGTGAA G-3′ and antisense: 5′-CTTCACCTTCCAGTAATGGC3′. Obtained mutant plasmids were transformed to E. coli MC100930 for easy in-vivo screening. In-Vivo Test. Prepared gp23 mutants were screened for functionality by testing their ability to support growth of the bacteriophage T4 amber mutant B17 (T4 amB17).31 T4 amB17 is unable to produce gp23 in E. coli MC1009, as such T4 amB17 will only be able to grow if the gp23 mutant on the plasmid is functional. Protein Purification. Purification of gp23 was performed essentially as described previously.32 Unless otherwise noted, all centrifugation steps were performed at 20 °C. Cells containing gp23 polyheads were sonicated and centrifuged for 20 min at 22 000g. The pellet was resuspended in 100 mM KPO4 pH 7 buffer and was centrifuged again for 20 min at 22 000g. This pellet was resuspended in 100 mM KPO4 pH 7 buffer and was centrifuged for 8 min at 3000g. The turbid supernatant, which contains the gp23 polyheads, was centrifuged for 20 min at 22 000g. To dissociate the gp23 polyheads, the pellet was resuspended in dialysis buffer and was dialyzed for 3 h to 10 mM KPO4/boric acid pH 9 buffer at 4 °C. The dialyzed suspension was subsequently centrifuged at 4 °C for 15 min at 25 000g. The supernatant contains purified gp23. The purified gp23 was stored at -20 °C after glycerol was added to a final concentration of 10% (v/v). Fluorescence Sample Preparation. Samples of 1.5 µM purified gp23 W13V mutant were prepared by dilution of the 0.1 mM stock solution in a buffer containing 50 mM Tris pH 7.4, 10 mM KCl, 10 mM MgCl2, and 0.01% Tween-20. Experiments were performed at a range of temperatures between 5 °C and 60 °C. For quenching experiments, KI was added to obtain final concentrations in the range from 0 to 0.5 M. To avoid formation of I2, 10 mM Na2S2O3 was added. The ionic strength was kept constant by addition of KCl. Fluorescence Instrumentation. Time-resolved fluorescence experiments were performed using a laser system combined with time-resolved photon-counting detection. The laser system consisted of a Mira 900-P laser (Coherent, Santa Clara CA), emitting 3-ps pulses at a repetition rate of 76 MHz, pumped by a Verdi-8 diode laser (Coherent). The Mira 900-P is a modelocked titanium-sapphire laser, tunable from approximately 700 to 1000 nm. The laser emission was led through a pulse picker (Coherent), which reduces the repetition rate to 4.75 MHz to avoid double excitation of molecules. The output wavelength of 290 nm was generated by means of a frequency tripler (Coherent). This light was used to excite the protein sample in a cell with 1-cm path length (type 104F, Hellma GmbH &Co KG, Mu¨llheim, Germany), which was thermostated by a Peltier element at temperatures between 0 and 60 °C. For detection, an SPC-630 (Becker & Hickl GmbH, Berlin, Germany) system was used, for time-correlated single-photon counting (TCSPC) with a time-resolution of about 15 ps. A laser pulse focused on a photodiode provided the synchronization signal. Fluorescence emission was collected at 90°, dispersed by a monochromator (TVC JarrellAsh Monospec 18, Grand Junction CO), and was detected by a photomultiplier tube. Decays were recorded at wavelengths between 320 and 450 nm in 10-nm steps. Steadystate fluorescence emission and excitation spectra were recorded on an LS-50B instrument (PerkinElmer Inc., Wellesley, MA), with a Peltier-thermostated cuvette-holder. Data Analysis. Fluorescence decay curves were analyzed using a fit procedure on the basis of the Levenberg-Marquardt
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algorithm, as discussed in ref 28. This procedure uses the instrument response signal (obtained by recording scattered laser light) for deconvolution of the recorded decays and determination of lifetimes and wavelength-dependent amplitudes. Each decay curve is fitted by mono- or multiexponential curves:
I(t) )
( ) t
∑i Ai exp - τ
(1)
i
The quality of the fit (“goodness of fit”) was assessed on the basis of χ2, the covariance matrix, and the distribution of residuals. The set of time constants was kept as low as possible; normally, not more than three constants were needed to obtain a fit of sufficient quality. For measurements at a series of detection wavelengths, a global fit procedure was used, on the basis of the method described above. Lifetimes are assumed to be constant for the decays included, while the amplitudes were varied. This assumption is correct for proteins in many cases.33 Decay-associated spectra (DAS) provide information on the fluorescence emission per lifetime component. DAS are constructed by distributing the total intensity per decay curve over the lifetimes that make up the total intensity according to their ratio (amplitudes are obtained by global fit). The relative fluorescence intensity of lifetime component τj at wavelength λ, with amplitude A, can be expressed by the following equation:
Iτj(λ) )
Ajτj
∑i Aiτi
(2)
The decay-associated spectra were corrected for an emission around 420 nm because of a photodegradated species, observed mainly in the longest lifetime of W13V. To determine the spectral maxima, the emission spectra were fitted to Gaussian functions on a wavenumber scale. The spectra could be fitted nicely using a combination of two Gaussians. Temperature-dependent structural changes of the protein were treated as phase changes. For the general phase change from phase R to phase β, eq 3 can be derived (see Supporting Information), which describes the fraction of phase R that is present at a specific temperature.
fR )
1 ∆trsH 1 1 1 + exp R Tc T
[ ( )]
(3)
Here, fR is the fraction of phase R present at temperature T (K); ∆trsH (kJ‚mol-1) represents the transition enthalpy from phase R to β; R (kJ‚mol-1‚K-1) is the gas constant; Tc (K) is the temperature where phase R and β are present in equal amounts. As emission intensity is also temperature-dependent. Equation 3 is combined with a linear temperature dependence for each species of gp23 to describe the data. For the transition from monomeric to hexameric gp23, this results in eq 4.
I ) fm‚(am‚T + bm) + (1 - fm)‚(ah‚T + bh)
(4)
Here, fm represents the fraction monomeric gp23; am and ah represent the slope of the linear temperature dependence of monomeric and hexameric gp23, respectively; similarly, bm and bh represent the intercept of the linear temperature dependences. In case of an additional transition between two structurally different hexameric gp23 species, eq 4 was extended to eq 5:
I ) fm‚(am‚T + bm) + (1 - fm)‚fh1‚(ah1‚T + bh1) + ( 1 - fm)‚(1 - fh1)‚(ah2‚T + bh2) (5) Results Mutagenesis. Bacteriophage T4 gp23 contains four tryptophan residues in positions 13, 247, 309, and 345, respectively. For a better understanding of the tryptophan fluorescence, it would be useful to have mutants of the protein containing a range of different tryptophan replacements.13,20,28 The prime candidate for mutagenesis is tryptophan 13. This residue is located in the part of the protein that is cleaved off during the maturation process of the bacteriophage head.27 Initially, W13 was mutated to a phenylalanine, but in-vivo tests showed that this mutant was not able to support bacteriophage T4 growth. Since a nonfunctional mutant is not useful for our purposes, a semirandom mutagenesis strategy was employed to find a suitable mutant. For this, degenerate primers were used in the QuikChange site-directed mutagenesis strategy. Possible candidates were screened via an in-vivo test to determine gp23 functionality, after which the mutation was identified via sequencing. Two possible candidates were picked up, W13D and W13V. Both mutants are able to support bacteriophage T4 growth, but only W13V is able to form polyheads. This is a requirement for isolation of the capsid protein using the existing wild-type gp23 purification protocol; therefore, W13V was chosen for further experiments. Not all possible mutations were tested exhaustively, so there may be additional functional mutants. Steady-State Fluorescence. Wild-type gp23 is mainly present as monomers at low temperature and forms hexamers at higher temperatures.10 To determine the contribution of W13 to the overall tryptophan emission of monomeric wild-type gp23, the steady-state emission spectra of wild-type and W13V gp23 were compared at low temperature. The spectra were normalized on the basis of protein concentration, which was determined via the absorbance at 280 nm and the theoretical extinction coefficients on the basis of the primary sequence of the two proteins (wild-type 48480 and W13V 42790 M-1‚cm-1).34 Although the spectra were nearly identical in shape, the emission intensity of W13V gp23 was about 46% lower than that of wildtype gp23 (data not shown). Thus, almost half the emission intensity of wild-type gp23 stems from W13, indicating a relatively weak quenching environment for this residue. As stated before, the degree of oligomerization of gp23 is temperature-dependent: monomer at low temperature and hexamer at room temperature. To determine the transition temperature for hexamer formation more precisely, a comparison was made of steady-state emission spectra as a function of temperature for both wild-type and W13V gp23. On increasing temperature, a steady decrease in tryptophan emission intensity is expected as a result of thermal quenching, caused by intensified collisions with neighboring atoms and solvent molecules. In the case of free tryptophan in water, this decrease is more or less linear.35 However, in a protein, deviations from linearity may occur because of temperature-induced structural changes, such as the formation of multimers. In Figure 1A, the relative emission intensity of wild-type and W13V gp23 at various temperatures is shown. A clear deviation from linearity can be observed, especially for wild-type gp23. However, this deviation does not appear to occur between 5 and 25 °C, which is where the transition from monomer to hexamer is expected to occur. The emission maxima, which are an indication of tryptophan environment,12 also do not change with temperature (data not shown). Therefore, time-resolved fluorescence mea-
Tryptophan Fluorescence of Bacteriophage T4 gp23
J. Phys. Chem. B, Vol. 110, No. 49, 2006 25053 TABLE 1: Fluorescence Lifetimes (τ), Amplitudes (A), Emission Maxima (Emax), and Comparative Analysis Results for Wild-Type and W13V gp23a exp 0 τm (ns) τw (ns) Am(%) Aw (%) fW13 τm‚Am (%) τw‚Aw (%) fW13‚(τw‚Aw)b Emax,m (nm) Emax,w (nm)
exp 1
exp 2
exp 3
5 °C (wild type and W13V) (monomer) 0.29 (0.14) 2.4 (0.4) 6.6 (0.4) 0.028 (0.002) 0.37 (0.02) 2.31 (0.05) 5.6 (0.2) 0 48.4 (0.8) 38.5 (1.0) 13.1 (1.6) 80.6 (3.0) 6.7 (1.5) 9.1 (1.4) 3.6 (0.9) 1 0 0.41 0.50 7.3 47.9 44.8 4.9 5.4 45.8 43.9 4.9 0 19.0 21.8 333 341 345 345 333 339 348
25 °C (wild type) 40 °C (W13V) (hexamer) τm (ns) 0.37 (0.06) 2.4 (0.1) 6.5 (0.3) τw (ns) 0.032 (0.001) 0.39 (0.02) 2.13 (0.04) 6.8 (0.3) Am(%) 0 44.2 (1.0) 49.6 (0.6) 6.2 (0.5) Aw (%) 84.1 (0.6) 5.3 (0.6) 8.1 (0.3) 1.8 (0.3) fW13 1 0 0.27 0.59 τm‚Am (%) 9.2 68.7 22.1 τw‚Aw (%) 7.8 6.0 50.4 35.8 fW13‚(τw‚Aw)c 7.8 0 13.4 21.0 Emax,m (nm) 331 342 341 Emax,w (nm) 345 339 347 351
Figure 1. Temperature dependence of the relative emission intensity for wild-type (b) and W13V (O) gp23. Data are normalized on their intensity at 5 °C. Panel A: The straight lines show the extrapolated temperature dependence of the monomer (dashed line: wild type, dotted line: W13V). Panel B: Deviation from the monomer temperature dependence. Lines represent a fit with a single (W13V) and double (wild-type) temperature transitions (see Experimental Section, eq 3). The temperature transitions were fitted globally with data shown in Figure 2. Error bars indicate the standard error.
surements were done to distinguish between monomeric and hexameric gp23. Time-Resolved Fluorescence. We have previously shown that for wild-type gp23 the fluorescence decay can be described well by four exponential decays.28 Here, we report that the fluorescence decay of the W13V mutant is best described by three exponential decays. In Table 1, the observed lifetimes at 5 °C are presented for both wild-type and W13V gp23. Comparison of these lifetimes showed that for the mutant the fastest lifetime (τ0) has disappeared. This seems to prove that τ0 originates completely from W13. However, as it involves an extremely short lifetime at the edge of the instrumental resolution, it is also possible that a slight change in the protein conformation has caused this lifetime to become irresolvable with our system. The remaining three lifetimes show no major differences compared to the wild type. The relative contributions of each three/four fluorescence lifetimes to the total steady-state emission were calculated by multiplying the relative amplitude with the fluorescence lifetime (τ‚A). The temperature dependence of these contributions is shown in Figure 2. Clearly, the relative contributions of τ2 and τ3 are temperature-dependent. For wild-type gp23, a temperature transition was observed between 5 and 25 °C, where the transition from monomeric to hexameric gp23 was expected. In addition, wild-type gp23 data showed a second transition at higher temperature, whereas W13V data revealed only a single transition across the measured temperature range.
a Values in brackets indicate the standard error. Exp 0-3 represent the exponents in the multiexponential decay curves. The subscript m indicates the mutant W13V gp23, w applies to the wild-type protein. fW13 represents the fraction W13 contributes to the wild-type signal. b Sum of f W13‚(τw‚ Aw) is 45.7, i.e., 45.7% of the wild-type emission is caused by W13 in the monomer. c Sum of fW13‚(τw‚ Aw) is 42.2, i.e., 42.2% of the wild-type emission is caused by W13 in the hexamer.
The transition enthalpy (∆trsH) and transition temperature (Tc) of the different temperature transitions were calculated using eq 3. As ∆trsH and Tc should be the same for both the steadystate data from Figure 1A and the time-resolved data from Figure 2, a global fit of both data sets was performed. On the assumption that the steady-state emission intensity of a species usually depends linearly on temperature for small temperature ranges, eq 3 was combined with eq 4 (W13V gp23) or 5 (wildtype gp23) for the actual fit of the data. Since the relative contributions of the lifetimes to the total emission (Figure 2) are not expected to be linearly dependent on temperature, the a parameters in eqs 4 and 5 were set to 0 to fit those data. The resulting fitted curves are shown both in Figure 1A and 2. Additionally, the resulting linear temperature dependencies of wild-type and W13V monomeric gp23 are also included in Figure 1A. Furthermore, in Figure 1B this linear dependence was subtracted from the original data, making the transitions in the steady-state data more evident. A transition at 29 °C (∆trsH ) 102 kJ‚mol-1) was found for W13V gp23; for wild-type gp23, a transition at 19 °C (∆trsH ) 339 kJ‚mol-1) and at 40 °C (∆trsH ) 390 kJ‚mol-1) was found. For wild-type gp23, the hexamermonomer transition is expected to occur somewhere between 5 and 25 °C,10 and the observed 19 °C transition is likely the gp23 hexamer formation. On the basis of the behavior of the contributions of lifetimes τ2 and τ3 (Figure 2), the transition at 29 °C for W13V gp23 is likely to be hexamer formation as well. The W13V mutation therefore has a significant effect on gp23 hexamerization, that is, it lowers ∆trsH and causes a shift of 10 °C in Tc. Interestingly, a temperature transition at 46 °C has been observed previously in gp23 polyheads using differential scanning calorimetry.36 This temperature transition was attributed to denaturation of the N-terminal part of gp23, the part of the sequence that is cleaved off during maturation of the bacteriophage head. As such, the observed temperature transition
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Figure 3. Fractions of the gp23 species present at different temperatures. Monomer (continuous lines), hexamer (dotted lines), and hexamer with denatured N-terminus (dashed line, only for wild type). The fractions were calculated from the analysis of the data shown in Figures 1 and 2.
Figure 2. Relative contributions of the fluorescence lifetimes of (A) W13V and (B) wild-type gp23 to the total emission as a function of temperature. The different lifetime contributions are indicated by τ1 (4), τ2 (0), and τ3 (]). The contribution of τ0 for wild-type gp23 is not shown, as its contribution is negligible. Lines represent fits of the data. The temperature dependence of τ1 was fitted with a straight line; the temperature dependences of τ2 and τ3 were fitted with a combination of eqs 3 and 4 for W13V gp23 and a combination of eqs 3 and 5 for wild-type gp23 (see text). The temperature transitions were fitted globally with data from Figure 1.
at 40 °C in wild-type gp23 is likely the result of denaturation of the N-terminal part of gp23. This temperature transition was not observed in the mutant as expected: the tryptophan residue that allows us to monitor the denaturation of the N-terminus was replaced in the mutant. The fractions of each of the three gp23 species (monomer, hexamer, and hexamer with unfolded N-terminus) at temperatures between 5 and 60 °C are shown in Figure 3. For an evaluation of the role of W13 emission in monomeric gp23, the fluorescence data of wild-type and W13V gp23 at 5 °C were compared. The comparison for hexameric gp23 is a little more complex, as hexamer formation shows different temperature dependences for wild-type and W13V gp23. In addition, wild-type gp23 data of hexamers is influenced by denaturation of the N-terminal domain at 40 °C. Hence, for the evaluation of hexameric gp23, a comparison was made of wildtype gp23 at 25 °C and W13V gp23 at 40 °C. Using the fit of the temperature-dependent data, it was calculated that at 25 °C wild-type gp23 consists for 93% of hexamers, while W13V gp23 at 40 °C consists of 81% of hexamers (see also Figure 3). Such a comparison gives a good estimate of the contribution of W13 to the emission of hexameric gp23. Therefore, additional timeresolved measurements were performed at 5 and 25 °C for wildtype gp23 and at 5 and 40 °C for W13V gp23. From this more extensive data set, decay-associated spectra (DAS) of each species were calculated using eq 2 and are shown in Figure 4. The contributions of each lifetime were obtained by multiplying
the amplitudes at each wavelength with the lifetimes. The sum of the DAS, which is included in Figure 4 as well, represents the total emission spectrum and is normalized to 1 for each graph. Remarkably, all spectra in Figure 4 belonging to equivalent lifetimes are similar with regard to their emission maximum and relative contribution (see also Table 1, τ‚A values). Lifetimes τ0 and τ1 contribute little to the emission spectrum. Lifetimes τ2 and τ3 contribute equally in the monomer, whereas in the hexamer τ2 contributes significantly more than τ3. This trend applies both to wild-type and W13V gp23. Comparison of the information obtained from the more extensive time-resolved measurements allowed an estimation of the contribution of W13 to each of the wild-type gp23 DAS. Involved parameters are collected in Table 1, as well as other information obtained from this data set. Equation 6 was used to determine the contribution of W13 to each of the lifetimes:
Am,i )
Aw,i - fi‚Aw,i
∑i (Aw,i - fi‚Aw,i)
(6)
Here, Am,i and Aw,i are the amplitudes obtained from the timeresolved data for the mutant and wild-type gp23, respectively. fi is the fraction W13 contributes to amplitude Aw,i. As noted before, the shortest lifetime τ0 is absent in W13V gp23, and thus f0 is 1. The other values for fi were determined via a leastsquares analysis. For this, the SOLVER function from Microsoft Excel was used.37 Results are shown in Table 1. From these calculations, it is evident that W13 contributes to both τ2w and τ3w but not to τ1w. Calculation of the contribution of each of these lifetimes to the steady-state spectrum (fW13‚(τw‚Aw)) reveals that W13 contributes 46% to the monomer and 42% to the hexamer. This is in agreement with the 46% contribution of W13 to the steady-state emission spectra of the monomer found earlier (see above). The contribution of τ2 to the emission spectrum increases upon hexamer formation, whereas that of τ3 decreases. In contrast, the contribution of W13 to τ3 increases upon hexamer formation, whereas its contribution to τ2 deceases. In principle, the degree of exposure (or actually the polarity) of the tryptophan environment can be inferred from decayassociated spectra.38 However, care must be taken since dipolar relaxation of the protein environment may also play a role.19 This relaxation would cause a shift of the spectrum during the fluorescence lifetime resulting in a more red-shifted spectrum
Tryptophan Fluorescence of Bacteriophage T4 gp23
J. Phys. Chem. B, Vol. 110, No. 49, 2006 25055
Figure 4. Decay-associated spectra of the different forms of wild-type and W13V gp23. Shown is the relative contribution of each lifetime to the emission spectrum, i.e., amplitude × lifetime. τ0 (O), τ1 (4), τ2 (0), τ3 (]), and the sum of these (b). Lines are a fit of a single Gaussian in the energy domain through the data points of each DAS. Panel A: W13V gp23 at 5 °C (monomeric); panel B: W13V gp23 at 40 °C (hexameric); panel C: wild-type gp23 at 5 °C (monomeric); panel D: wild-type gp23 at 25 °C (hexameric).
for longer lifetimes. In fact, this effect has been observed for wild-type gp23,28 so it is likely to occur in W13V as well. Taking this into account, it was possible to discern two classes of tryptophan environment in the data for monomeric wild-type and W13V gp23: fully buried tryptophan for lifetime τ1 and moderately buried for lifetimes τ0, τ2, and τ3. For hexameric gp23, this distinction was not so clear: it seems τ1 represents a buried tryptophan in W13V but a moderately buried one in wild-type gp23. Since W13 does not contribute to this lifetime, it is an indication of its influence on the entire gp23 structure. To determine the degree of exposure of the tryptophan residues in W13V, iodide-quenching experiments were performed.39,40 The results were difficult to interpret using traditional Stern-Volmer relations since the emission from three or four fluorophores, each emitting with multiple lifetimes, needs to be taken into account. Qualitatively, it was concluded that the tryptophan residues in W13V gp23 are quenched only slightly more in the monomer than in the hexamer and thus the tryptophan residues are almost equally exposed in both forms of the capsid protein. Both major contributing lifetime components (τ2 and τ3) showed an identical trend, which indicates that most tryptophan residues emit with the same lifetimes, only with different amplitudes. Furthermore, lifetime τ1 appeared to be less influenced by quencher than the other lifetimes and was quenched to a similar degree in monomer and hexamer, indicating it originated from a buried tryptophan residue. The results for τ1 are comparable to wild-type gp23, but there τ2 and τ3 are quenched much less in the hexamer than in the monomer.28 Discussion The W13V mutant of the bacteriophage T4 capsid protein gp23 was studied using intrinsic tryptophan fluorescence. Especially, the role of W13 was assessed since it resides in a
part of the protein that is cleaved off after prohead formation during bacteriophage T4 biogenesis. The N-terminal part of the protein is not essential for phage formation, as a gp23 mutant that has its 65 N-terminal residues removed is still able to support bacteriophage T4 growth (data not shown). It is therefore surprising that it was difficult to find a suitable mutation for W13. Presumably, W13 is involved in a specific interaction, be it intra- or intermolecularly. Furthermore, the ability to form polyheads is not a prerequisite for a functional gp23 as is evident from the behavior of the W13D mutant, which supports bacteriophage T4 growth but does not form polyheads. Possibly, the N-terminus of gp23 interacts with another protein, maybe to help position the gp23 protein correctly in the phage head, thus increasing the efficiency of the phage maturation process. It is also possible that the cleavage of the N-terminus is a signal for conformational changes, DNA packaging, or other phagerelated processes. The importance of the N-terminus and more precisely W13 is further emphasized by the effect of the W13V mutation on the monomer-to-hexamer transition enthalpy of gp23. Where in wild type the ∆trsH is 339 kJ‚mol-1, in W13V gp23 it has dropped to 102 kJ‚mol-1. Also, the transition temperature has changed from 19 to 29 °C. The temperature range for gp23 to go from 90% monomer to 90% hexamer has broadened from 15 to 24 °C for wild type to 13-46 °C for W13V. Seemingly, W13 plays a stabilizing role in the hexamerization process. Once formed, W13 is no longer needed to keep the hexamer together and can be cleaved off. This is supported by the temperature transition at 40 °C in wild-type gp23, which represents the thermal denaturation of the N-terminus and has been indicated before on the basis of differential scanning calorimetry.36 It is likely that also in W13V gp23 the N-terminus denatures around 40 °C. However, because of removal of W13, it is no longer possible to monitor changes in the N-terminus via tryptophan
25056 J. Phys. Chem. B, Vol. 110, No. 49, 2006 fluorescence. As no transition around 40 °C is seen for W13V gp23, the other tryptophan residues are apparently not influenced by denaturation of the N-terminus. The relative instability of the N-terminus, compared to the rest of the protein, suggests it protrudes from the main protein structure. This would also make it easier for the N-terminus to become proteolytically cleaved during phage head maturation. The remainder of the protein thermally denatures around 65 °C.36 In fact, at these high temperatures, we were unable to obtain reliable results because of precipitation of the protein. As such, no reliable results could be obtained at those high temperatures. Whereas denaturation of the N-terminus can be easily visualized via steady-state spectra, this is not so straightforward for the hexamerization process. To visualize hexamerization via tryptophan fluorescence, time-resolved fluorescence experiments are necessary. As can be seen in Figure 2, the relative contributions of lifetimes τ2 and τ3 are clearly influenced by temperature. These lifetimes originate from multiple tryptophan residues, including W13. The contribution of W13 to these lifetimes was estimated via a comparison of wild-type and W13V gp23. As shown in Table 1, W13 contributes 41% to τ2 and 50% to τ3 in monomeric gp23, whereas in the hexamer its contribution to τ3 is higher (59%) and to τ2 lower (27%). Since the different lifetimes indicate different orientations of tryptophan residues, it appears that upon hexamer formation the orientations causing τ3 are favored. The influence of W13 on this effect is small, however, it was observed for both wildtype and W13V gp23. The largest part of this change in relative contribution is caused by changes in other tryptophan residues whose equilibrium shifts to orientations that cause the τ2 lifetime. A closer look at the temperature dependence of the rate constants and accompanying relative amplitudes confirms this (see Supporting Information). The solvent accessibility of the different tryptophan orientations represented by τ2 and τ3 is similar, as is also observed in iodide-quenching experiments (see Supporting Information). The same is suggested by the emission maxima of the decayassociated spectra of the different lifetimes, as the emission maxima of the different lifetimes of wild-type and W13V monomer show very little difference. The obvious exception here is of course the absence of lifetime τ0 for W13V. Possibly, its disappearance is caused by a shift out of the temporal range of our setup because of increased quenching by surrounding amino acids. Alternatively, this lifetime may be caused completely by W13. The latter interpretation is supported by comparison of the steady-state spectra of the monomer of wildtype and W13V gp23. There it is shown that W13 contributes 46% to the total emission; the same number is found on the basis of the time-resolved fluorescence data (see Table 1, τ0 contributes 5%). The decay-associated spectra of monomeric gp23 indicate that τ0, τ2, and τ3 represent tryptophan residues that are in a relatively polar environment and thus are likely to be exposed to solvent. The residue dominating τ1 on the other hand appears to be in a more hydrophobic environment, that is, completely buried in monomeric gp23. This residue is not W13 (see Table 1) but must be one of the other three tryptophan residues. The small differences in W13V monomer and hexamer with respect to the maxima of the decay-associated spectra and quenching behavior indicate that conformation of the subunits in the hexamer is similar to the monomer structure. However, in wild-type gp23, the subunits of the hexamer show significant differences compared to the monomeric form.28 This may be the cause for the difference in the temperature dependence of
Stortelder et al.
Figure 5. Representations of the gp23 homology model. Panel A: single subunit of the hexamer form of gp23, pictured sideways; panel B: complete hexamer, top view. Tryptophan residues are shown in red or are indicated by circles. The ribbons of alternating subunits have been given alternating colors (black and gold). The N-terminal part (1-73) is not shown. The two other unmodeled parts are shown as dashed lines (residues 229-250: green dotted line (right to left in panel A), residues 131-165: blue dotted line (left to right in panel A)). This figure was prepared using the program MOLMOL41 using the structure coordinate file deposited at the Protein Data Bank42 (http:// www.rcsb.org/pdb) with PDB ID: 1Z1U.26 The program POV-Ray (http://www.povray.org) was used to render the images.
hexamer formation. These observations provide a new insight into the role of the W13 residue, which appears to be involved in a significant structural change during hexamer formation. It seems plausible that this change in conformation prevents accumulation of di- or trimeric forms of gp23 and stimulates hexamerization. For the W13V mutant gp23, it is possible that di- and trimers accumulate (as evidenced by the large temperature range for hexamerization) even though they cannot be detected by fluorescence. Recently, a homology model of gp23 on the basis of the crystal structures of the bacteriophage T4 gp24 and HK97 gp5 proteins has become available.26 A representation of the model is shown in Figure 5. Sequence alignment and genetic data show that the folds of gp24 and gp23 are similar. Although gp24 is present as pentamers in the T4-head and gp23 as hexamers, it is likely they have similar structures and similar intermolecular interactions. This is supported by the observation that point mutations in gene 23 allow gp23 to form pentamers and substitute for gp24.2 In addition, gp24 shows strong structural similarity to the structure of the gp5 capsid protein of bacteriophage HK97. As such, it is likely the model closely represents the actual gp23 structure
Tryptophan Fluorescence of Bacteriophage T4 gp23 Figure 5A and B shows the homology model of the monomer and hexamer, respectively. For three stretches of the primary sequence (1-73, 131-165, and 229-250), no reasonable modeling information could be generated. The first region representing the N-terminal part containing W13 is not indicated; the other two parts are shown as dotted lines and include W247. Examination of the homology model shows that W309 is tightly packed inside the gp23 core but is still slightly exposed to solvent. It is the prime candidate for causing lifetime τ1 as observed in the fluorescence lifetime analysis. In the monomer, this residue should then be completely buried. However, since the environment of W247 is unclear because of its location in an unmodeled section, W247 is also a possible origin of lifetime τ1. The τ2 and τ3 lifetimes probably come from the other three tryptophan residues W13, W247, and W345. Since the contribution of W13 to these lifetimes (46%) has already been confidently assigned, this leaves the contribution of W247 and W345 to τ2 and τ3. Upon hexamer formation, the relative contribution of these tryptophan residues shifts from τ3 to τ2 (Figure 2). Solvent exposure is similar for both, but τ3 seems to become slightly more buried upon hexamerization. W345 seems to be quite exposed in the homology model of the monomer, whereas in the hexamer it is more confined but is still exposed to solvent and to the other five W345 residues. This will cause a quite polar environment. The situation of W247 is somewhat difficult to interpret as it is not included in the homology model. However, its approximate location suggests W247 may be involved in intersubunit interactions. As such, the W247 emission may also be influenced by the hexamerization process and changes most likely to that of a more buried and quenched state. It is hence likely that W345 will contribute mostly to τ2 and W247 mostly to the τ3 lifetime. Conclusions Steady-state and time-resolved fluorescence techniques, making use of the emission of the intrinsic fluorophore tryptophan, have provided interesting results regarding the monomerhexamer transition of the gp23 capsid protein of bacteriophage T4. Tryptophan emission of the wild-type gp23 has been described previously.28 In this study, the W13V mutant of gp23 was used for further examination of the monomer-hexamer equilibrium and a more detailed assignment of the fluorescence lifetimes. From temperature-dependent measurements, the hexamerization range was determined. This range turned out to be narrower for wild-type gp23 than for the W13V mutant (respectively, 15-24 °C and 13-46 °C) suggesting that the W13 residue has a stabilizing influence on intermediate structures and thereby accelerating hexamer formation. This result is in line with the difficulty in preparing a functional tryptophan mutant of this protein. In wild-type gp23, a second transition at 40 °C is found that is caused by denaturation of the N-terminus. The contribution of W13 to the total emission of gp23 is 46%, more than expected if the four tryptophan residues would contribute equally. The W13V mutant shows three fluorescence lifetimes (0.4, 2.4, and 6.5 ns), which are comparable to those found for the wild-type protein. The 32-ps lifetime found for the wild type is not observed in W13V, and it may be ascribed to the W13 residue. Furthermore, it could be inferred from the data revealed that W13 does not contribute to the 0.4-ns lifetime. A recently published homology model of gp23 enabled assignment of the fluorescence lifetimes to specific tryptophan residues. The 0.4-ns lifetime is probably caused by W309, a
J. Phys. Chem. B, Vol. 110, No. 49, 2006 25057 buried residue. The 2.4- and 6.5-ns lifetimes originate from W13 (27-59%, depending on the conformation of the protein), W247, and W345. The relative contributions to the total emission of the 2.4- and 6.5-ns lifetimes are temperature-dependent and most likely represent two different orientations of the tryptophan residues, of which one or the other state is favored at a particular temperature. Acknowledgment. The authors thank Ms. Esther van Duijn (Vrije Universiteit Amsterdam) for providing the N-terminally truncated gp23 mutant phage. A.S. and J.H. contributed equally to this work. Supporting Information Available: Derivation of a formula to fit the temperature-dependent data, table showing anisotropy results, figure showing the temperature dependence of the fluorescence lifetimes and corresponding amplitudes of wildtype and W13V gp23, and figure showing iodide quenching. This material is available free of charge via the Internet at http:// pubs.acs.org. References and Notes (1) Fokine, A.; Chipman, P. R.; Leiman, P. G.; Mesyanzhinov, V. V.; Rao, V. B.; Rossmann, M. G. Proc. Natl. Acad. Sci. U.S.A. 2004, 101, 6003-6008. (2) Black, L. W.; Showe, M. K.; Steven, A. C. In Molecular Biology of Bacteriophage T4; Karam, J. D., Ed.; ASM Press: Washington, DC, 1994; pp 218-258. (3) Deuerling, E.; Bukau, B. Crit. ReV. Biochem. Mol. Biol. 2004, 39, 261-277. (4) Nagradova, N. K. Biochemistry Moscow 2004, 69, 830-843. (5) Bhutani, N.; Udgaonkar, J. B. Curr. Sci. 2002, 83, 1337-1351. (6) Ben-Zvi, A. P.; Goloubinoff, P. J. Struct. Biol. 2001, 135, 84-93. (7) Slavonitek, A. M.; Biesecker, L. G. Trends Genet. 2001, 17, 528535. (8) Eiserling, F. A.; Black, L. W. In Molecular Biology of Bacteriophage T4; Karam, J. D., Ed.; ASM Press: Washington, DC, 1994; pp 209212. (9) Van Duijn, E.; Bakkes, P. J.; Heeren, R. M. A.; Van Den Heuvel, R. H. H.; Van Heerikhuizen, H.; Van Der Vies, S. M.; Heck, A. J. R. Nat. Methods 2005, 2, 371-376. (10) Bakkes, P. J. PhD-thesis, Vrije Universiteit, Amsterdam, 2005. (11) Beechem, J. M.; Brand, L. Annu. ReV. Biochem. 1985, 54, 43-71. (12) Eftink, M. R. Biochemistry Moscow 1998, 63, 276-284. (13) Engelborghs, Y. J. Fluoresc. 2003, 13, 9-16. (14) Callis, P. Methods Enzymol. 1997, 278, 113-150. (15) Lakowicz, J. R. In Principles of fluorescence spectroscopy; Lakowicz, J. R., Ed.; Plenum Press: New York, 1983; pp 341-379. (16) Vivian, J. T.; Callis, P. Biophys. J. 2001, 80, 2093-2109. (17) Szabo, A. G.; Rayner, D. M. J. Am. Chem. Soc. 1980, 102, 554563. (18) Stortelder, A.; Buijs, J. B.; Van Der Vies, S. M.; Gooijer, C.; Van Der Zwan, G. Appl. Spectrosc. 2004, 58, 705-710. (19) Lakowicz, J. R. Photochem. Photobiol. 2000, 72, 421-437. (20) Engelborghs, Y. Spectrochim. Acta 2001, 57, 2255-2270. (21) Petrich, J. W.; Chang, M. C.; McDonald, D. B.; Fleming, G. R. J. Am. Chem. Soc. 1983, 105, 3824-3832. (22) Liu, T.; Callis, P. R.; Hesp, B. H.; De Groot, M.; Buma, W. J.; Broos, J. J. Am. Chem. Soc. 2005, 127, 4104-4113. (23) Callis, P. R.; Liu, T. J. Phys. Chem. B 2004, 108, 4248-4259. (24) Chen, Y.; Barkley, M. D. Biophys. J. 1998, 81, 1765-1775. (25) Alcala, J. R.; Gratton, E.; Prendergast, F. G. Biophys. J. 1987, 51, 597-604. (26) Fokine, A.; Leiman, P. G.; Shneider, M. M.; Ahvazi, B.; Boeshans, K. M.; Steven, A. C.; Black, L. W.; Mesyanzhinov, V. V.; Rossmann, M. G. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 7163-7168. (27) Aijrich, L. G.; Kurochkina, L. P.; Mesyanzhinov, V. V. Biochemistry Moscow 2002, 67, 815-821. (28) Stortelder, A.; Buijs, J. B.; Bulthuis, J.; Gooijer, C.; Van Der Zwan, G. J. Photochem. Photobiol. B 2005, 78, 53-60. (29) Mayer, M. P. Gene 1995, 163, 41-46. (30) Aksoy, S.; Squires, C. L.; Squires, C. J. Bacteriol. 1984, 157, 363367. (31) Edgar, R. S.; Denhardt, G. H.; Epstein, R. H. Genetics 1964, 49, 635-648.
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