Fluorescence-Based Kinetic Analysis of the Interaction between Avidin and Singlet Oxygen Damaged Lambda Phage DNA

ABSTRACT

Oxidative damage of DNA, especially damage induced by singlet oxygen, correlates with a host of disease processes. Due to the link between oxidative DNA damage and disease, a great deal of effort has been put forth to develop methods to detect and quantify DNA damage based on the presence of oxidation-specific biomarkers. Two oxidation products, 8-oxoguanine and 8-oxoadenosine have become important biomarkers of oxidative damage. This study describes a two solution-based methods for monitoring the binding of a fluorescein-labelled avidin to photodamaged lambda phage DNA. The first method, fluorescence polarization (FP), measured changes in the emission of polarized light originating from the fluorescein label that resulted from avidin binding to DNA. The second method, measured the quenching of tryptophan fluorescence resulting from formation of the avidin-DNA complex. This fluorescence-based approach proved to be very sensitive, allowing detection of the avidin-DNA complex when the concentration of avidin and DNA were 10-9 M and 10-12 M, respectively. Competition experiments using either biotin, or 8-oxoguanosine, demonstrated that binding of avidin to photodamaged DNA could be disrupted, thus providing evidence for the involvement of the biotin binding site. Kinetically, the binding of avidin to DNA is a complex process, the rate of complex formation being dependent on the extent of damage and the structure of the DNA (i.e., native or denatured). Based on these results, a three-step kinetic model is presented.

INTRODUCTION

Characteristics and Origin of Singlet Oxygen

Reactive oxygen species (ROS) are molecules that impose oxidative damage on proteins, lipids, and nucleic acids (Cooke et al., 2003; Davies, 2004; Piette, 1991). Reactions between ROS and biomolecules can lead to mutations, and the inhibition of cellular processes and biomolecular function (Cooke et al., 2003). ROS are by-products of cellular metabolism and other biological processes. Examples include, but are not limited to, generation via reactions of the electron transport chain, enzymatic production, and formation and release by activated phagocytic cells engaged in destruction of bacteria and viruses (Cooke et al., 2003). Increased production of ROS have been associated with a wide variety of noncancerous diseases including neurodegenerative diseases like Alzheimer's disease, Huntington's disease, and Parkinson's disease (Cooke et al., 2003). These reactive molecules have also been implicated in the chronic inflammatory disease systemic lupus erythematosus and atherosclerotic plaque development (Cooke et al., 2003). Increased levels of ROS have been correlated with age and are therefore thought to play a key role in aging and age-related diseases (Cooke et al., 2003).

Singlet oxygen, 1O2, is a highly reactive member of the ROS family of molecules. Electronically, 1O2 is an excited stated form of molecular oxygen. Ground state O2 exists in a triplet electronic configuration designated 3O2, with two unpaired electrons in the highest occupied molecular orbital (Foote and Clennan, 1995). Two forms of singlet oxygen exist. The first excited state, designated 1Deltag, is characterized by having paired electron spins in a single orbital, leaving the other vacant. The second excited state is a more energetic singlet state designated 1Sigmag+. The electrons in this state have opposite spins and occupy different orbitals. The energy difference between the first excited state and the ground state is 22 kcal/mole. The energy difference between the first and second excited state is 15 kcal/mole. Studies have concluded that the 1Sigmag+ state has a significantly shorter lifetime than the 1Deltag state, due in part to relaxation to the 1Deltag state before the chemical reaction can occur (Cooke et al., 2003; Foote and Clennan, 1995). As a result of this relaxation event, the chemical reactivity of 1O2, which is oxidative, is most often associated with the first excited 1Deltag state.

Formation of 1O2 may occur via chemical or photochemical processes. Examples of chemical reactions that form 1O2 include reactions involving H2O2 with HOCl, sodium molybdate, or peroxynitrite (Davies, 2004). Thermal decomposition of endoperoxides such as 3,3'-(1,4-naphthalyidene) dipropionate leads to the efficient in situ production of 1O2 (Khan et al., 1992). Photogenerated 1O2 results from interaction of 3O2 with a molecule residing in an excited singlet state due to absorption of light energy. Interaction between 3O2 and the excited molecule results in the transfer of excited state energy from the molecule (donor) to O2 (acceptor) which returns the donor to the ground state and elevates O2 to a singlet excited state. This type of photochemical process is known as photosensitization. Common photosensitizing agents include, porphyrins, methylene blue, rose bengal, and riboflavin. In fact, the mechanistic basis of photodynamic therapy lies in photosensitized formation of 1O2 (Kawanishi et al., 1986; Niedre et al., 2005).

Deleterious Effects of Singlet Oxygen-Mediated DNA Damage

Oxidative damage to DNA is an important area of research that has received considerable attention. Exposure of DNA to 1O2 can lead to strand breakage and base damage (Kawanishi et al., 1986; Khan et al., 1992). Oxidative damage to DNA results in the formation of numerous products (Aruoma et al., 1989). The nucleosides 8-oxodeoxyguanosine and 8-oxoadenosine are two of the most common products (Struthers et al., 1998). Reaction of 1O2 with guanosine or adenosine generates an unstable oxetane type adduct that decomposes into an enolic 8-hydroxynucleoside. The 8-hydroxy product exists in equilibrium with the keto or oxo product. Model studies have shown that the nucleotide 8-oxodeoxyguanosine triphosphate causes mutations by pairing with adenine during DNA replication (Minnick et al., 1994), inducing a 46-fold increase in the number of A•T→ C•G transversions (Minnick et al., 1994). The other oxidized base products also produce mutations, but the oxidized nucleoside 8-oxodeoxyguanosine is the most popular subject of these studies (Cooke et al., 2003). Increased frequency of mutation resulting from oxidative damage to DNA bases, may play a key role in the initiation, promotion, and progression of carcinogenesis (Cooke et al., 2003).

Modified Nucleosides as Biomarkers of Oxidative Stress

The diverse array of diseases associated with increased ROS levels indicates that ROS have the propensity to wreak havoc on many different processes in the body. Therefore, detection, and quantification of ROS is of great importance, as it allows an avenue to monitor and track the abundance of ROS within certain tissues. Further research in this area could allow the attribution of these molecules to a disease with greater certainty. One can indirectly determine the abundance of ROS and oxidation caused by ROS by measuring the oxidized nucleoside 8-oxodeoxyguanosine, the most commonly used biomarker of oxidative damage to DNA (Cooke et al., 2003). Gas chromatography coupled to mass spectrometry (GC-MS) and high performance liquid chromatography coupled with electrochemical detection (HPLC-EC) are the most common methods for detecting oxidative base damage (Cadet et al., 1997). These methods are extremely sensitive but pose many limitations. For example, they produce highly variable estimates of the oxidative damage in the same sample of DNA, indicating that they may induce oxidative damage in the sample (Cadet et al., 1997). In addition, these assays are time-consuming and difficult to perform. Assays of oxidized DNA bases based on the use of antibodies, purified repair enzymes, single-cell gel electrophoresis, or ligation-polymerase chain reaction techniques are highly sensitive, but lack the specificity required to be a suitable assay (Cadet et al., 1997).

Use of Avidin to Detect Singlet Oxygen-Mediated Damage of Nucleic Acids

Due to its high affinity and specificity for biotin, as well as, the stability of the avidin-biotin complex, avidin is used extensively in a diverse number of applications ranging from immunoassays, to affinity purification of biotinylated proteins and nucleic acids (Schetters, 1999; Struthers et al.,1998; Wilchek et al., 2006). Avidin is a tetrameric protein with four identical subunits (combined MW = 67kDa). The protein is soluble in water, stable over a wide range of pH values, and the avidin-biotin interaction is the strongest noncovalent interaction known in biological systems with a dissociation consant, Kd, of 10-15 M. A novel use of avidin arose from the observation that avidin binds to the nuclei of oxidatively damaged cells (Struthers et al., 1998). Results of this work revealed that formation of the avidin-DNA complex involves recognition of 8-oxodeoxyguanosine. Competition experiments using biotin resulted in inhibition of avidin binding to DNA, thus confirming the specificity of the interaction with respect to the biotin binding site (Struthers et al., 1998). The DNA base guanine and nucleoside dGMP exhibit very little structural similarity to biotin, however, sufficient structural similarity exists between biotin, 8-oxoguanine, and 8-oxodeoxyguanosine to allow binding. The structures of biotin, 8-oxodeoxyguanosine, and 8-oxodeoxyadenosine, a derivative also shown to be recognized by avidin (Conners et al., 2006), are shown in Figure 1.

Figure 1.  Structures of biotin, and oxidation products formed upon reaction of 1O2 with guanosine and adenosine.  Highlighted atoms define the common ureido functionality required for avidin binding.

Figure 1. Structures of biotin, and oxidation products formed upon reaction of 1O2 with guanosine and adenosine. Highlighted atoms define the common ureido functionality required for avidin binding.

The highlighted atoms represent the key recognition feature, the ureido functionality that is present in biotin (Green, 1975; Struthers et al., 1998). The ureido group forms stabilizing hydrogen bonds with the avidin binding site, and is responsible for optimal stabilization of the complex (Conners et al., 2006; Green, 1975; Struthers et al., 1998).

In this investigation, we present a simple, specific, and sensitive, solution-based method to detect oxidized DNA that utilized the interaction between avidin and 8-oxo forms of deoxyguanosine and deoxyadenosine. Experiments were designed to test the hypothesis that binding of avidin to DNA containing oxidized guanine and adenine bases would increase the molecular weight of avidin, thus making it possible to detect complexes using fluorescence polarization. Secondarily, it was hypothesized that changes in trpytophan fluorescence resulting from the interaction of avidin with DNA would be indicative of complex formation. The results of this investigation indicate that fluorescence polarization is a suitable method for monitoring avidin-DNA interactions. Additionally, changes in tryptophan fluorescence were observed upon formation of avidin-DNA complexes.

Materials and Methods

Photosensitized Oxidation of DNA

Oxidation reactions took place in 10 mM sodium phosphate buffer (pH 7.4) containing 100 mM NaCl. The reaction mixture (150 µl) contained 100 µM rose Bengal (RB) (Aldrich), 5 µg Lambda phage DNA (Sigma). A quartz microcuvette containing the reaction mixture was irradiated for 30 minutes in a Photon Technologies International QM-2 spectrofluorometer at the wavelength of 547 nm (Houseman, 2005). After 15 minutes, the solution was mixed with a micropipette to ensure adequate aeration. Micro Bio-Spin 6™ columns (BioRad) were used to separate the DNA from rose bengal following the standard protocol provided by the manufacturer. The volume of the eluted sample was approximately equal to the volume added to the column, and this indicated that little dilution of the DNA occurred. A sample of eluate containing damaged DNA was denatured at 95C for 10 minutes and quenched on ice.

Fluorescence Polarization Binding Assays

A Beacon 2000 Fluorescence Polarization (FP) Analyzer (Panvera) outfitted with fluorescein excitation (excitation = 490 nm) and emission (emission = 520 nm) filters was used for this work. Avidin labeled with fluorescein isothiocyanate (Avidin-FITC) was used as the fluorescent reporter. A protocol with a 2-second delay that averaged five intensity readings parallel and perpendicular was used to produce the observed FP value. For each assay, a blank solution was prepared by placing 600 µl of buffer (10 mM phosphate, pH 7.4, 0.1 M NaCl) in a 2 ml glass tube. The sample was prepared by placing 600 µl of buffer (10 mM phosphate, pH 7.4, 0.1 M NaCl) and 0.5 µl of 0.4 µg/µl avidin-FITC (Sigma) in a 2 ml glass test tube. After measuring the FP of this DNA free sample, 1.0 µl of a DNA containing solution was added and the contents mixed. The polarization was recorded at 1 minute intervals. The concentration of DNA in the assay was 0.055 ng/µl ~ 2 pM, and avidin-FITC concentration was 0.333 ng/µl ~ 5.0 nM. For the competitive binding studies, the required amount of inhibitor (i.e., biotin (Sigma) or 8-oxodeoxyguanosine (Sigma)) was added to a solution of avidin-FITC before the addition of the DNA. The FP of the samples was collected as previously described. Polarization values are given as milliunits (mP).

Tryptophan Fluorescence Assays

A reaction mixture that contained 600 µl of buffer (10 mM phosphate, pH 7.4, 0.1 M NaCl) and 0.2 µl of 5 µg/µl native, fluorescein-free avidin (Sigma) was prepared in a quartz cuvette. The fluorescence emission spectrum of the sample was recorded using the aforementioned spectrofluorometer. The sample was excited with 280 nm light, and the emission recorded from 290 to 450 nm. To this avidin containing solution, a 1µl aliquot of oxidized DNA was added and after rapid mixing, the emission spectrum recorded. Subsequent readings were taken at five and ten minutes after the addition of DNA. For the time-based studies, a reaction mixture that contained 600 µl of buffer (10 mM phosphate, pH 7.4, 0.1 M NaCl) and 0.2 µl of avidin (5 µg/ µl) was prepared in a quartz cuvette. A 1 µl aliquot of oxidized DNA was added to the cuvette and rapidly mixed. The cuvette was placed immediately in the fluorometer, and the time-based readings were initiated. The time-based samples were excited at 280 nm, and the emission was recorded at the wavelength of maximum tryptophan fluorescence (340 nm).

RESULTS

Changes in Fluorescence Polarization Correlate to Formation of Avidin-DNA Complexes

Fluorescence polarization assays have been used for more than a twenty years in a diverse variety of research fields to study the mobility of membrane lipids, the domain movement of proteins, and biomolecular interactions (Jameson and Sawyer, 1995; Lundblad et al., 1996). In this investigation, FP was used to determine if fluorescein-labeled avidin forms a complex with Lambda-phage DNA. Figure 2 shows a representative FP assay for three types of DNA.

Figure 2.  Time-based fluorescence polarization curves of avidin-FITC-lambda DNA solutions.  RB-treated denatured DNA (circles), RB-treated native DNA (squares), and native untreated DNA (triangles).  Denatured DNA was prepared after exposure to RB and isolation of the photodamaged DNA.  Data were collected at 23C, in the presence of buffer (10 mM sodium phosphate, pH 7.4, 0.1 M NaCl).  The concentration of DNA and avidin-FITC were approximately 2 pM and 5 nM, respectively.  The data shown and the time to reach maximum polarization (TmaxP) values represent one trial out of three independent trials. Initial polarization values (t = 0):  RB-treated denatured DNA = 152.5 mP; RB-treated native DNA = 180.2 mP, and untreated native DNA = 160.7 mP.  TmaxP values: RB-treated denatured DNA = 5 min; RB-treated native DNA = 8 min, and untreated native DNA = 90 min.  Average TmaxP values: RB-treated denatured DNA = 5 + 1 min; RB-treated native DNA = 9 + 2 min, and untreated native DNA = 88 + 6 min.

Figure 2. Time-based fluorescence polarization curves of avidin-FITC-lambda DNA solutions. RB-treated denatured DNA (circles), RB-treated native DNA (squares), and native untreated DNA (triangles). Denatured DNA was prepared after exposure to RB and isolation of the photodamaged DNA. Data were collected at 23C, in the presence of buffer (10 mM sodium phosphate, pH 7.4, 0.1 M NaCl). The concentration of DNA and avidin-FITC were approximately 2 pM and 5 nM, respectively. The data shown and the time to reach maximum polarization (TmaxP) values represent one trial out of three independent trials. Initial polarization values (t = 0): RB-treated denatured DNA = 152.5 mP; RB-treated native DNA = 180.2 mP, and untreated native DNA = 160.7 mP. TmaxP values: RB-treated denatured DNA = 5 min; RB-treated native DNA = 8 min, and untreated native DNA = 90 min. Average TmaxP values: RB-treated denatured DNA = 5 + 1 min; RB-treated native DNA = 9 + 2 min, and untreated native DNA = 88 + 6 min.

The results of these trials indicate that avidin interacts with DNA, and that this interaction is sensitive to DNA structure. Based on the data shown on Figure 2, avidin binding affinity increases in the order damaged, denatured > damaged, native > undamaged, native. Undamaged, denatured DNA also formed complex, and was faster than undamaged, native DNA, but slower than damaged, native DNA (data not shown). Oxidized, denatured DNA displayed the fastest rate of polarization change reaching a maximum polarization of 310 mP within 5 minutes (TmaxP). Native, damaged DNA reached a maximum of 300 mP within 8 minutes, while untreated, native DNA displayed the slowest rate of polarization increase; reaching a maximum of 290 mP in approximately 90 minutes. For all samples, a sharp decrease to a stable minimum polarization ranging between 210 and 220 mP was observed. The binding of avidin to native undamaged DNA is consistent with the presence of oxidized bases as a result of natural oxidation (Struthers et al., 1998).

To ascertain whether the binding between avidin and DNA involved the biotin binding site, a competition experiment using damaged, native DNA was performed. The presence of 2 µM biotin (squares) increased TmaxP on average by a factor of two compared to the sample with no biotin (circles). Upon reaching TmaxP , the polarization of each sample rapidly decreased to a minimum polarization of approximately 210 mP. In general, the kinetic behavior of this system in the presence of biotin paralleled that observed in its absence. To obtain additional evidence for the involvement of the biotin binding site and confirm that recognition of photodamaged DNA by avidin involved this nucleoside, a second competition experiment was performed using 8-oxodeoxyguanosine (8-oxodG) as the competitor. The data (Figure 3) exhibit parallel kinetic behavior and associated increase in TmaxP that was observed in the presence of biotin.

Figure 3.  Competitive binding assay.  DNA alone (triangles), DNA + 2 uM biotin (squares), DNA + 3 uM 8-oxo-dG (circles).  Data were collected at 23C, in the presence of buffer (10 mM sodium phosphate, pH 7.4, 0.1 M NaCl ).  The concentration of damaged, native DNA and avidin-FITC were approximately 2 pM and 5 nM, respectively. The data shown and TmaxP values represent one trial out of three independent trials.  TmaxP values:  DNA alone = 10 min; DNA + biotin = 19 min, and DNA + 8-oxo-dG = 21 min.  Average TmaxP values: DNA alone = 10 + 1 min, DNA + biotin = 20 + 3 min, DNA + 8-oxo-dG = 20 + 2 min.

Figure 3. Competitive binding assay. DNA alone (triangles), DNA + 2 uM biotin (squares), DNA + 3 uM 8-oxo-dG (circles). Data were collected at 23C, in the presence of buffer (10 mM sodium phosphate, pH 7.4, 0.1 M NaCl ). The concentration of damaged, native DNA and avidin-FITC were approximately 2 pM and 5 nM, respectively. The data shown and TmaxP values represent one trial out of three independent trials. TmaxP values: DNA alone = 10 min; DNA + biotin = 19 min, and DNA + 8-oxo-dG = 21 min. Average TmaxP values: DNA alone = 10 + 1 min, DNA + biotin = 20 + 3 min, DNA + 8-oxo-dG = 20 + 2 min.

The results of these competition experiments are consistent with previous observations of diminished avidin binding to photodamaged DNA in fixed tissue samples using fluorescence microscopy (Struthers et al., 1998). Moreover, these results are also consistent with a recent study which measured the dissociation constant (kd) for avidin binding to biotin ( upper limit: kd < 0.04 µM), 8-oxodeoxadensine (kd = 24 µM), and 8-oxodeoxyguanosine ( kd = 117 µM) (Conners et al., 2006). Significantly, this study also measured the binding of avidin to single-stranded oligonucleotides containing a single 8-oxodeoxyguanosine or 8-oxodeoxyadenosone base (shown in bold) (Conners et al., 2006). The sequences of these oligonucleotides from 5' to 3' are as follows: CTCGTCT and CTCATCT. Sequence analysis of Lambda phage DNA for the presence of these model sequences with the modified bases replaced by its unmodified counterpart revealed the presence of one CTCG TCT sequence and eight CTCATCT sequences. It is interesting to note that both oligonucleotides exhibited similar affinity toward avidin, having kd values of 14 and 12 µM, respectively. Thus, the observed affinity of avidin toward photodamaged Lambda DNA is consistent with this reported result.

Binding of Avidin to Photodamaged DNA Quenches Tryptophan Fluorescence

Steady-state fluorescence emission spectroscopy was used to detect tryptophan (Trp) fluorescence. Avidin contains several Trp residues in or near the biotin binding site. It has been shown that formation of the avidin-biotin complex causes quenching of Trp fluorescence (Conners et al., 2006; Kurzban et al., 1989). Based on the results of the biotin and 8-oxodG competition experiments, which support involvement of the biotin binding site, we hypothesized that formation of an avidin-DNA complex should alter Trp fluorescence. To determine if Trp fluorescence is altered upon complex formation, a series of steady-state measurements were made (Figure 4).

Figure 4.  Steady emission spectrum of avidin trpytophan fluorescence as a function of time in the absence and presence of damaged, native DNA.  Data were collected at 23C, in the presence of buffer (10 mM sodium phosphate, pH 7.4, 0.1 M NaCl ).  Tryptophan florescence was excited at a 275 nm.  The concentration of DNA and avidin were approximately 2 pM and 5 nM, respectively.  The data shown represent one trial out of two independent trials.

Figure 4. Steady emission spectrum of avidin trpytophan fluorescence as a function of time in the absence and presence of damaged, native DNA. Data were collected at 23C, in the presence of buffer (10 mM sodium phosphate, pH 7.4, 0.1 M NaCl ). Tryptophan florescence was excited at a 275 nm. The concentration of DNA and avidin were approximately 2 pM and 5 nM, respectively. The data shown represent one trial out of two independent trials.

The spectra measured immediately after addition of DNA (Avidin + DNA curve), and at times of 5 and 10 minutes post-addition of DNA, reveal decreased Trp fluorescence relative to the spectrum collected in the absence of DNA (Avidin curve). The observation of DNA-induced quenching of Trp fluorescence further supports binding of DNA to the biotin binding site.

The time-dependent quenching of Trp fluorescence is shown in Figure 5.

Figure 5.  Time-dependent change in tryptophan fluorescence induced by complex formation between avidin and photodamaged native DNA.  Tryptophan florescence was excited at 275 nm and detected at 340 nm.  The concentration of DNA and avidin-FITC were approximately 2 pM and 5 nM, respectively. Data was collected at a rate of 1 point/second.  The approximate boundary between the fast and slow kinetic phases is marked by the dashed vertical line at t = 120 seconds.  The data shown represent one trial out of two independent trials.

Figure 5. Time-dependent change in tryptophan fluorescence induced by complex formation between avidin and photodamaged native DNA. Tryptophan florescence was excited at 275 nm and detected at 340 nm. The concentration of DNA and avidin-FITC were approximately 2 pM and 5 nM, respectively. Data was collected at a rate of 1 point/second. The approximate boundary between the fast and slow kinetic phases is marked by the dashed vertical line at t = 120 seconds. The data shown represent one trial out of two independent trials.

It is interesting to note the appearance of a fast phase during the first 120 seconds, followed by a slow phase. The observed kinetic behavior of this system is not described by a one-component first-order kinetic model having the form y = Aoexp(-kt), which would be expected considering the concentrations of avidin and DNA are 5 nM and 2pM, respectively. If this system exhibited classic first-order kinetics, the data as plotted in Figure 5 would be linear with a negative slope. The observed kinetic behavior is characteristic of a system having multiple (at least two) first-order components. To retrieve these components, the slow and fast phases of the data were analyzed separately using the curve-fitting program, TableCurve 2D (Systat, Inc.) (Figure 6).
Figure 6.  Separation of kinetic components associated with avidin-DNA complex formation. Data represent the fast and slow phases of the kinetic data shown in Figure 4.  Fast component 0 to 120 seconds (inset), k = 25.1 x 10-3 sec-1 (r = 0.980).  Slow component 120 to 1800 seconds, k = 1.95 x 10-3 sec-1 (r = 0.995).

Figure 6. Separation of kinetic components associated with avidin-DNA complex formation. Data represent the fast and slow phases of the kinetic data shown in Figure 4. Fast component 0 to 120 seconds (inset), k = 25.1 x 10-3 sec-1 (r = 0.980). Slow component 120 to 1800 seconds, k = 1.95 x 10-3 sec-1 (r = 0.995).

Based on the value of the regression coefficient and visual inspection, the data is best described by the kinetic equation y = Feq + (Fo – Feq)exp(-kt), where y represents the fluorescence intensity at any given time, Fo and Feq represent the, initial and equilibrium fluorescence intensities, respectively; k, the net first-order rate constant, and t, time. Analysis of each phase resulted in the recovery of the first-order kinetic constants kfast and kslow. The fast phase which spans the first 120 seconds (see Figure 5 and inset, Figure 6) is characterized by kfast = 25.1x10-3 sec-1 (r = 0.980). The slow phase which spans 120 to 1800 seconds is characterized by kslow = 1.95x10-3 sec-1 (r = 0.995). Multiple independent trials exhibit the same kinetics, although the duration of the fast and slow phases appears variable. Future work will seek to characterize these kinetic phases.

DISCUSSION AND CONCLUSION

The goal of this work was to investigate the interaction of avidin with lambda phage DNA, a model DNA having 48,502 base pairs. Using fluorescence-based detection it was possible to detect and monitor the formation of avidin-DNA complexes in solution, in real-time. Most significantly, due to the sensitivity of the detection method, it was possible to observe complex formation in a system containing picomolar and nanomolar concentrations of DNA and avidin, respectively. Time-based steady-state fluorescence polarization experiments confirmed the formation of avidin-DNA complex based on a change in rotational diffusion, as well as, revealed a transition hypothesized to be indicative of a change of state from a mobile form of DNA to a condensed form.

Fluorescence polarization experiments carried out in the presence of antagonist ligands (i.e., biotin or 8-oxodeoxyguanosine) provide strong evidence for the involvement of the biotin binding site when avidin forms a complex with DNA. Steady-state fluorescence experiments of intrinsic tryptophan fluorescence revealed time-dependent quenching of fluorescence which is indicative of complex formation involving the biotin binding site. Furthermore, time-based experiments revealed complex behavior that could be deconvolved into two first-order components. Based on the results obtained from these experiments, a preliminary model based is presented. It is intriguing to hypothesize that these two distinct kinetic phases may be indicative of one or more of the following: 1) the initial formation of avidin-DNA interactions involving primarily electrostatic interactions, 2) the binding of avidin to accessible nucleoside bases, and 3) differential binding to 8-oxodeoxyadenosine (kd = 24 µM) or 8-oxodeoxyguanosine ( kd = 117 µM) (Conners et al., 2006). The first point that of electrostatic binding, is based on the fact that avidin is a basic protein having a pI of 8.5. As a result, it will carry a net positive charge under the conditions of these experiments (pH 7.4), which would favor electrostatic attraction and binding to the negatively charged phosphate ester backbone of the DNA. Specific binding of avidin to a modified adenosine or guanosine will be dependent on the accessibility of the base. Accessibility will be dependent on the conformation of the DNA, as well as, thermodynamic stability of the nucleotide sequence containing the modified base. DNA is a dynamic macromolecule undergoing numerous conformational changes in solution (Georghiou et al., 1996; Kang et al., 2005; Kidoaki and Yoshikawa, 1996). Regions having low stability will have a greater tendency to assume alternate conformations compared to regions of higher stability. If destabilization is associated with local denaturation, then the modified base would become more accessible, which in turn would increase the probability of recognition by avidin.

The results obtained from polarization and fluorescence quenching experiments have been used to develop a three-step kinetic model of the avidin-DNA interaction. The proposed model is illustrated in Figure 7.

Figure 7.  Proposed kinetic model of avidin-DNA complex formation based on fluorescence polarization and time-dependent fluorescence quenching data. The first-order rate constant kfast corresponds to initial electrostatic binding and recognition of accessible bases.  The second rate constant, and kslow corresponds to avidin-induced alterations in DNA structure and conformation, respectively.  The formation of condensed DNA is associated with a sudden decrease in polarization because of depolarizing light scattering which is observed at time = TmaxP (See Figures 2 and 3).

Figure 7. Proposed kinetic model of avidin-DNA complex formation based on fluorescence polarization and time-dependent fluorescence quenching data. The first-order rate constant kfast corresponds to initial electrostatic binding and recognition of accessible bases. The second rate constant, and kslow corresponds to avidin-induced alterations in DNA structure and conformation, respectively. The formation of condensed DNA is associated with a sudden decrease in polarization because of depolarizing light scattering which is observed at time = TmaxP (See Figures 2 and 3).

The first step is rapid and involves electrostatic binding and recognition of accessible bases by avidin. Significantly, all FP assays exhibit a rapid increase in polarization during the first 60 – 120 seconds, which coincides with the initial fast phase observed in the fluorescence quenching experiment (Figure 5, and inset, Figure 6). This increase in polarization is consistent with an increased molecular weight, and hence, slower rotational diffusion of avidin-FITC. The second step is slow, involving structural alterations of both DNA and avidin, which result in increased avidin-DNA interactions. The increased interactions between avidin and DNA serve to decrease of the rotational diffusion of the bound avidin-FITC further. Significantly, the FP assays exhibit a slower rate of polarization change up to a maximum polarization (TmaxP). The duration of this phase is dependent on the amount of photodamage, as well as, the structure of the DNA. Recall that polarization of avidin-FITC changed most rapidly when measured in the presence of denatured, damaged DNA (Figure 2). The third step, which leads to collapse or condensation of the DNA, is rapid. Although the fluorescence quenching experiments did not reveal any abrupt change after a long period, FP experiments consistently revealed a transition from a value of maximum polarization to a stable minimum. This equilibrium polarization is designated as TmaxP. This transition is analogous to that observed when a system undergoes a change from one form to another (i.e., a two-state transition). Examples include, but are not limited to, denaturation of proteins and nucleic acids, micelle formation, and polymer aggregation. Significantly, the formation of avidin-DNA "nanoparticles" has been observed using the method of dynamic light scattering (Morpurgo, 2004). In the case of this work, a drastic structural change, such as condensation of the avidin-DNA complex, would result in increased light scattering, which would depolarize the emitted fluorescence and cause a decrease in the observed FP.

In closing, the results of this investigation not only confirmed earlier reports of the high affinity that avidin displays toward DNA, but reveal the importance of time-based measurements with respect to revealing complex nature of the interaction between avidin and DNA.

ACKNOWLEDGEMENTS

The authors wish to thank Stetson University for providing support and an ideal environment for pursuing intellectual development.

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