 |
INTRODUCTION |
The genes encoding the variable domains of immunoglobulins or T
cell receptors are generated during lymphocyte differentiation by a
somatic recombination reaction known as V(D)J recombination (1). The
reaction is initiated by DNA double strand breaks created at the
junction between two coding segments (termed V, D, or J) and their
flanking recombination signal sequences
(RSSs).1 Cleavage is followed
by a complex repair process that results in imprecise joining of the
two coding segments and typically precise joining of the two RSSs. The
RSSs consist of two conserved sequence elements, the heptamer
(consensus 5'-CACAGTG-3') and the nonamer (consensus 5'-ACAAAAACC-3'),
separated by a poorly conserved spacer sequence of either 12 or 23 base
pairs. Efficient recombination occurs only between a 12-RSS and a
23-RSS, a phenomenon known as the 12/23 rule (for review, see Ref. 2).
Recognition of 12/23-RSSs and concerted cleavage at the RSS-coding
sequence border is performed by a complex of the RAG1 and RAG2 proteins (for reviews, see Ref. 3 and 4), the lymphoid-specific products of the
recombination-activating genes RAG1 and RAG2 (5,
6). Binding and cleavage of DNA by the RAG1·RAG2 complex is
facilitated by the ubiquitously expressed architectural DNA-binding
proteins HMG1 and -2 (7, 8).
Deletion mutagenesis has established the minimal "core" domains of
murine RAG1 (residues 384-1008 of the 1040 aa RAG1 protein; Fig.
1A) and RAG2 (residues 1-383 of the 527 aa RAG2 protein) required for recombination activity in transfected nonlymphoid cell
lines (9-12). Most biochemical studies of V(D)J recombination have
been performed with the core RAG proteins because of their enhanced
solubility compared with the full-length proteins.
Surface plasmon resonance (13) and in vivo one-hybrid (14)
experiments demonstrated specific recognition of the nonamer by the
RAG1 nonamer binding domain and a less significant contribution of the
heptamer to binding. In the one-hybrid experiments, RAG2 only modestly
enhanced RAG1 DNA binding (14), implying that RAG1 by itself was
capable of significant, sequence-specific DNA binding. Subsequently,
numerous studies making use of electrophoretic mobility shift assays
(EMSAs) and, in most cases RAG proteins fused to large N-terminal tags
(glutathione S-transferase (GST) or maltose-binding protein
(MBP)), led to the identification of a stable RAG1·RAG2·RSS complex
termed the SC (for signal complex) (15-18). These studies also
indicated that RAG1 alone binds the RSS with low specificity and that
RAG2 substantially enhances the affinity and specificity of the
interaction. Such findings have led to the suggestion that the RAG1-RSS
interaction is unlikely to be biologically significant in the absence
of RAG2 (15). Only weak, nonspecific DNA binding by RAG2
alone has been reported (17), but recent evidence supports a direct
interaction between RAG2 and DNA in the RAG1·RAG2·RSS complex (19).
Footprinting experiments (15, 20, 21) reveal that RAG1 generates
protection over the nonamer and the flanking portion of the spacer,
whereas with both RAG proteins, protein-DNA interactions extend
throughout the spacer and heptamer (21). All footprinting experiments
utilized MBP-RAG1 core fusion proteins.
The stoichiometry of the RAG proteins in the SC is controversial. Some
experiments indicated the presence of a dimer of RAG1 (18, 22, 23, 24),
whereas others argue for the presence of three or four RAG1 subunits
(25, 26). RAG2 is present in the SC either as a monomer (18) or as a
dimer (23, 26). The nonamer binding domain of RAG1 is thought to
contact the nonamer, whereas a central domain of RAG1 (aa 528-760) has
been shown to display some specificity for the heptamer (27). In
addition, a C-terminal domain of RAG1 has been shown to cross-link to
the coding flank (28). Recent experiments indicate a division of labor
between RAG1 monomers in the SC; one contacts the nonamer, whereas
another engages the heptamer at the site of cleavage (25, 29).
We currently do not know the equilibrium constant that governs the
association of RAG1 monomers into dimers. Furthermore, since most DNA
binding studies have been performed by EMSA, we do not know the
equilibrium constants governing reactions involving RAG1 alone or RAG1
in conjunction with RAG2 and HMG proteins in forming complexes with the
RSS in solution. The equilibrium dissociation binding constant for the
interaction of a MBP-RAG1 core (aa 384-1008) fusion protein with 12- or 23-RSS oligonucleotides was determined by EMSA to be close to 100 nM (22). In this study, however, the MBP-RAG1 fusion
protein was capable of forming several protein-DNA complexes with
various protein stoichiometries and various DNA binding affinities, and
the determined dissociation constant reflected the overall binding
without specification of the individual components of reaction.
We have now used fluorescence spectroscopy and a RAG1 protein lacking a
bulky tag to study the RAG1-RSS interaction in solution, thereby
overcoming some of the limitations inherent in the EMSA methodology and
any complications caused by a large fusion partner. We find that the
RAG1-RSS interaction is of higher affinity and specificity than
previously reported. In addition, we document a substantial
conformational change in RAG1 induced by interaction with the RSS.
Based on these findings, we propose that the RAG1-RSS interaction may
play an important role in vivo, especially in light of the
observation that for a portion of the cell cycle, developing
lymphocytes express RAG1 in the virtual absence of RAG2 protein
(30).
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EXPERIMENTAL PROCEDURES |
DNA Oligonucleotides--
Unlabeled and fluorescently labeled
deoxyoligonucleotides were synthesized and high performance liquid
chromatography-purified by Integrated DNA Technologies Inc.
(Coralville, IA). The 12-RSS substrate was made by annealing (top
strand) 5'-GTCGACCACAGTGCTACAGACTGGAACAAAAACCCTGCAG-3' with its complement. The nonspecific DNA substrate was made by annealing (top strand)
5'-GTCGACTGGCCATCTACAGACTGGAGCGGCCGCACTGCAG-3' with its
complement. The fluorescein fluorophore was attached at the 5' ends of
the top strands using a 5'-fluorescein phosphoramidite that attaches
6-carboxyfluorescein to the oligonucleotide via a C6 chain linker.
Annealing of oligonucleotides to generate double-stranded DNA was
performed in binding buffer (BB; 10 mM Tris-HCl (pH 7.5), 50 mM NaCl, 5 mM MgCl2) by heating
the complementary oligonucleotides mixed in equimolar amounts for 5 min
at 95 °C followed by slow cooling to room temperature.
Expression and Purification of Strep-RAG1 (StrRAG1)--
Murine
core RAG1 was expressed as a Strep-tag II fusion in the context of the
pASK-IBA5 vector (Sigma Genosys). The catalytic core of murine RAG1
from aa 377-1008 was amplified using primers that created an
NdeI site at the 5' end of the gene and a XhoI site at the 3' end. These PCR products were subcloned into the corresponding sites in pASK-IBA5, placing the Strep tag at the N
terminus of the protein.
The fusion protein was expressed in the BL21 strain of
Escherichia coli. Expression cultures were started by adding
20 ml of overnight culture per 1 liter final culture volume and grown at 25 °C with rapid shaking (>200 rpm) until the
A600 was ~1. Expression was induced by
the addition of anhydrotetracycline to a final concentration of ~1
µM and ZnCl2 to a final concentration of 0.1 mM. Cultures were grown for an additional 12-16 h, at
which point the A600 was usually ~5
absorbance units. Pelleted bacteria were resuspended in ice-cold lysis
buffer (20 mM Tris (pH 8.0), 100 mM NaCl, 0.5 mM EDTA, 5 mM 2-mercaptoethanol, 2 µM ZnCl2, 10% glycerol, and protease
inhibitor mixture (phenylmethylsulfonyl fluoride, pepstatin A,
aprotinin, leupeptin). Cells were lysed by sonication on ice, and
debris and insoluble material were pelleted by ultracentrifugation at
26,000 rpm for 1 h at 4 °C in a SW-41 rotor (Beckman). Cleared
lysate was added to Fast-Flow Q-Sepharose (Amersham Biosciences)
50-75-ml bed volume column preequilibrated in the lysis buffer before
the addition of lysate. After loading, the column was washed with two
bed volumes of lysis buffer containing 200 mM NaCl, and
RAG1 was eluted with 3 bed volumes of lysis buffer containing 500 mM NaCl. Q-Sepharose elution fractions were applied to a
7-10-ml bed volume Streptactin-Sepharose column (Sigma Genosys) pre-equilibrated in lysis buffer containing 500 mM NaCl,
and after washing with lysis buffer containing 500 mM salt,
Strep-RAG1 was eluted in lysis buffer (500 mM salt)
containing 2.5 mM desthiobiotin. The eluted Strep-RAG1
protein was typically >90-95% pure. Aggregated Strep-RAG1 was
separated from dimers and other oligomers using a Superdex 200HR column
(Amersham Biosciences). Fractions containing the dimeric protein were
dialyzed against BB. MBP-RAG1 core (aa 384-1008) and
glutathione S-transferase-RAG2 (aa 1-383) protein expression and purification were performed as described in (22, 31).
Coupled Cleavage Assay--
Coupled cleavage reactions (50-µl
final volume) contained 10 ng of body-labeled substrate containing both
a 12-RSS and a 23-RSS, 100 ng of each RAG protein, 30 ng of HMG2, and 5 mM MgCl2 and were incubated for 2 h at
37 °C using buffer conditions described previously (32). Reaction
products were resolved on a 4% native Tris borate EDTA
polyacrylamide gel.
EMSA--
To create the EMSA probe, 2 µM 12-RSS
double-stranded oligonucleotide DNA was 5' end-labeled using T4
polynucleotide kinase (New England Biolabs) and [
-32P]
ATP (3000Ci/mmol) (PerkinElmer Life Sciences) and purified on a 5%
native polyacrylamide gel. StrRAG1 or MBP-RAG1 was incubated with
radioactively labeled 12-RSS oligonucleotide in the presence or absence
of unlabeled 12-RSS or nonspecific DNA in BB supplemented with 10 ng/ml
heparin and 0.5% glycerol at 25 °C for 20 min, and samples were
resolved on 4% native polyacrylamide gels in 0.5× Tris borate (45 mM Tris (pH 8.9), 45 mM borate) at room
temperature. For better separation of the multimeric species formed by
MBP-RAG1, a discontinuous 3.5-8% polyacrylamide gel was used. Gels
were dried and quantitated using a PhosphorImager (Molecular Dynamics). The fraction bound at saturation, maximal fraction bound,
FMAX = 1, is set constant, whereas we expressed
the normalized fraction bound as the ratio between the fraction bound
at each data set and the actual constant value of the fraction bound at
saturation. The data were fit to a Hill binding isotherm with
n = 2.
|
(Eq. 1)
|
[StrRAG1] represents the concentration of free protein
expressed as monomeric protein.
Steady-state Fluorescence Measurements--
The StrRAG1
intrinsic emission fluorescence spectra were recorded using a SLM-8000
L format spectrofluorometer equipped with a 750-watt Xenon arc lamp.
All fluorescence measurements were performed in 100-µl quartz
cuvettes (Starna, Atascadero, CA) at 25 °C constant temperature
maintained by a circulating water bath. The StrRAG1 intrinsic emission
fluorescence spectra were recorded with an excitation wavelength of 280 nm using an 8-nm band pass for both the monochromator and the emission
long pass. All fluorescence emission spectra were recorded between 300 and 420 nm using 1-nm steps and 2-s integration times. Dilution of
StrRAG1 samples was done in ice-cold BB buffer filtered twice through
0.2-µm sterile filters (Gelman Laboratory) followed by a 10-min
incubation at 25 °C. For static experiments, mixing of StrRAG1 with
DNA or acrylamide was done inside the quartz cuvette followed by a
10-min incubation at 25 °C constant temperature before recording.
Each spectrum was corrected (33) for background, photobleaching
(5-15%), and dilution relative to a control sample for which the same
amount of protein was incubated in parallel with a corresponding volume of buffer. Photobleaching was always reversible, indicating that no
photo products are generated in the reaction. The fluorescence emission
intensity of all spectra were expressed as a ratio relative to a
standard rhodamine reference excited with the same source and whose
emission output was a constant
46,500. If sample absorption at
excitation wavelength exceeded 0.001, inner filter corrections were
also applied according to Lakowicz (34). For the time course experiment
shown in Fig. 7b, DNA was mixed with protein rapidly with a
long capillary tip with the cuvette in the holder and an automatic
shutter opened immediately for recording. The emission at 327 nm was
recorded at 1-s intervals.
Steady-state Fluorescence Polarization
Measurements--
Polarization anisotropy measurements were performed
on a PTI C-61 T-format fluorometer equipped with UV-transmitting
Glan-Thompson plane polarizers and a circulating water bath to control
the cell temperature. All measurements were performed using the same
quartz cuvette used for emission scans and temperature set at 25 °C. For tryptophan fluorescence, the samples were excited at 295 nm, and
emission was collected at 327 nm, whereas for fluorescein the
excitation wavelength was set at 492 nm, and emission was recorded at
520 nm. The recorded intensities were corrected for variations in lamp
intensity, detector, and monochromator wavelength-dependent sensitivity. G-factor corrections were included in all calculated anisotropy values. For the time course experiment shown in Fig. 7a, fluorescence intensities were recorded at 1-s intervals.
A 1-ml quartz cuvette with a round bottom was used to allow magnetic stirring. To the solution containing 50 nM 12-RSS
fluorescein-labeled DNA, either a concentrated StrRAG1 solution or BB
buffer supplemented with 50% glycerol (control reaction) was added in
a volume less than 1/20 of the total sample, without mixing. The
unmixed components were incubated together for 100 s, and then
mixing was initiated by rapid stirring.
Circular Dichroism Measurements--
Circular dichroism spectra
either of DNA or DNA incubated with StrRAG1 was recorded using a JASCO
J715 spectropolarimeter. The buffer used for incubation was 10 mM Tris-Cl (pH 7.5), 10 mM NaCl, 40 mM KCl, 5 mM MgCl2. The spectra
were recorded at 25 °C between 200 and 300 nm. For analysis of the
spectra of DNA and StrRAG1 mixtures, the separately recorded DNA
spectra were subtracted, and then the spectra were converted to molar
ellipticity based upon the molar concentration of StrRAG1 present in
each sample.
Time-resolved Fluorescence Measurements--
Time-resolved
fluorescence intensity and anisotropy decay kinetics were measured in
the frequency domain using an SLM-AMINCO 48000 multiharmonic Fourier
(MHF) spectrofluorometer (Spectronic Instruments). An argon-ion laser
(Coherent, Innova 90-6) operating at 488.0 nm was used as the
excitation source. An interference filter (Oriel) was placed in the
excitation beam path to minimize extraneous plasma tube super radiance
from reaching the detection system. The sample fluorescence was
monitored in the typical L-format after passing through a 515-nm
longpass filter. The Pockels cell modulator was operated at a 5 MHz
base repetition rate. Typically, data were acquired for 60 s
between 5 and 150 MHz (30 total frequencies), and at least 10 discrete
multifrequency data sets were acquired for a given sample. For the
excited-state intensity decay measurements, we used a dilute solution
of rhodamine 6G dissolved in water as the reference lifetime standard
(3.85 ns). Magic angle polarization conditions were used for all
excited-state intensity decay measurements to eliminate bias arising
from fluorophore rotational reorientation. The excited-state
fluorescence lifetimes and rotational reorientation times were
recovered from the frequency-domain data using a commercially available
non-linear least squares software package (Globals Unlimited) wherein
the true uncertainty in each datum was used as the
frequency-dependent weighting factor. The data were fitted
for single double and triple exponential decay with limiting anisotropy
set nonvariable to 0.4 (35), but only the double exponential
fitting gave more than 98% correlation with our recorded data points
and confident
2 values between
0.05-0.2.
Fluorescence Data Analysis--
For every set of anisotropy
data, a minimum of three experiments was performed under identical
conditions, and the data points shown are average values. Each averaged
data point was considered only if the sample S.D. was less than 5-7%
with respect to the calculated mean. The fitting of averaged data
points was performed using a typical Hill binding isotherm (36),
|
(Eq. 2)
|
where robs is the observed anisotropy of
the reporter species, rb is the basal anisotropy of
the reporter species in the absence of ligand,
rsat is the maximal anisotropy reached at
saturation, [L] is the free ligand concentration varied in the
reaction, expressed as monomeric species, n is the Hill
coefficient expressing the degree of cooperative ligand binding to the
reporter species, and KD is the apparent
dissociation constant comprising cooperative interaction factors, which
reflects the ligand concentration at half-maximal anisotropy. To fit
our data values, Equation 2 was written as a function of
[L]tot, total concentration of the ligand species, and
[R]tot, total concentration of the reporter species in
solution.
|
(Eq. 3)
|
The real solutions fc =
[R(L)n]/[R]tot of Equation 3 were substituted
into the fluorescence anisotropy expression to be fitted to,
|
(Eq. 4)
|
The best fit for StrRAG1 binding curve to fluorescein-labeled
12-RSS DNA was obtained with the Hill coefficient n = 1.8. The solution calculations and fitting were performed using Maple 7.0 and Origin 6.0 software.
For experiments in which tryptophan anisotropy was monitored while
adding unlabeled 12-RSS, because StrRAG1 quantum yield changes with DNA
binding we calculated the correction factor R = F/F0, where F0
is the relative quantum yield (derived from the integral value of
fluorescence corrected spectra) of free StrRAG1, and F is
the relative quantum yield of DNA bound StrRAG1 at saturation.
|
(Eq. 5)
|
Using Equation 5, which relates the fraction bound
protein to anisotropy (34, 37), combined with Equation 2 for DNA
binding, we obtained the equation used for fitting our tryptophan
anisotropy data in Fig. 5 where L is the unbound DNA
([DNA] = [L]) (Equation 6).
|
(Eq. 6)
|
The calculations of the apparent molecular weight of protein-DNA
complexes using the rotation reorientation time were performed using
the original Perrin equation,
|
(Eq. 7)
|
where
= 0.94 centipoise P, the viscosity of the
aqueous medium at 25 °C, Mr is molecular
weight,
= 0.73 ml/g, the specific volume for the
anhydrous protein, h is the degree of hydration (h = 0 for an anhydrous sphere, h = 0.2-0.4 ml/g for the typical hydration of most protein molecules),
T = 298 K, and R = 8.31 × 107 erg K
1mol
1; one can predict
based on the rotation reorientation time
Global = 48.03 ± 10.05 ns the size of the StrRAG1·12-RSS complex to be between
175 kDa (using h = 0) and 120 kDa (using
h = 0.4).
Acrylamide quenching data represented as Stern-Volmer plots show an
upward curvature concave towards the y axis. This is a characteristic feature for fluorophores quenched both by collisional and static quenching mechanisms. The fractional remaining fluorescence (F0/F) versus acrylamide
concentration [Q] was fit to a second-order function (38, 39),
|
(Eq. 8)
|
where KD and KS are the
apparent dynamic and static quenching constants that characterize the
ensemble of fluorophores (Trp and Tyr) present in StrRAG1.
 |
RESULTS |
Catalytically Active, Dimeric RAG1 Protein Lacking a Bulky
Tag--
In our study, we used a bacterially expressed and purified
murine RAG1 protein (aa 377-1008) fused at its N terminus to a short
(8 aa) Strep-II tag (Sigma Genosys, Inc.) that allows affinity purification with Streptactin-coupled Sepharose (Fig.
1a). Three purification steps
result in a StrRAG1 protein of greater than 98% purity (Fig.
1b). The elution profile from the final gel filtration column reveals a prominent peak at the position expected for a StrRAG1
dimer (compare Fig. 2, a and
c; the predicted molecular mass of the monomer is 72 kDa).
When the purified dimeric protein was reanalyzed on the gel filtration
column, only the dimer peak was observed, indicating that the protein
remains stably associated in dimeric form (Fig. 2b). The
intrinsic polarization anisotropy of the purified StrRAG1 at
concentrations from 25 to 500 nM is constant, consistent
with the existence of protein as a dimer at all concentrations within
this range (data not shown). In a standard coupled cleavage assay, the
StrRAG1 protein was approximately as active as highly purified, dimeric
MBP-core RAG1 (aa 384-1008) (Fig. 1c).

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Fig. 1.
The StrRAG1 protein. a, diagram of
the murine core RAG1 protein used in these studies showing the
N-terminal Strep tag, the nonamer binding domain (NBD), and
the DDE motif (three acidic amino acids involved in catalysis by the
RAG1/2 complex). b, diagram of the three steps of StrRAG1
purification and the 8% SDS-polyacrylamide gel stained with Coomassie
Blue showing the protein obtained from the final purification step.
c, StrRAG1 (aa 377-1008) displays cleavage activity
comparable with that of MBP-RAG1 (aa 384-1008) is active for coupled
cleavage in vitro. RAG and HMG proteins were added as
indicated above the lanes. Substrate and products
are depicted schematically to the left, with open
and filled triangles indicating the 12- and 23-RSSs,
respectively, and circles indicating the hairpin coding
ends.
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Fig. 2.
StrRAG1 exists in solution as a dimer.
a, gel filtration chromatography (Superdex 200HR) profile
from the last purification step. Approximately 70% of the protein is
present in dimeric form. b, fractions 13 and 14 corresponding to the dimeric protein in a were combined, and
after concentration, the protein was subjected again to gel filtration
chromatography. Essentially all of the protein remains in dimeric form.
c, gel filtration chromatogram of protein calibration
markers separated on the same Superdex 200 HR column. IgG,
goat purified immunoglobulin G; BSA, bovine serum albumin;
Ribo A, ribonuclease A.
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|
By EMSA, StrRAG1 Binds Specifically to the RSS, Forming a Single
Protein-DNA Complex--
We first tested the DNA binding ability of
StrRAG1 by EMSA. Increasing amounts of StrRAG1 (or for comparison,
MBP-RAG1) were incubated with a constant amount of labeled DNA bearing
a consensus 12-RSS sequence. The binding buffer used throughout this
study contains a moderate salt concentration (50 mM NaCl
and 5 mM Mg2Cl) and was chosen to minimize
nonspecific RAG1-DNA interactions (13, 15). Under these binding
conditions, StrRAG1 (Fig. 3a)
gives rise to one predominant shifted species, whereas MBP-RAG1 (Fig. 3b) yields at least three complexes of distinct mobility,
consistent with a previous study (22). Binding by StrRAG1 was
quantified, and the data points were fit to a single Hill binding
isotherm equation (Equation 1; "Experimental Procedures"), giving a
dissociation constant of 92 ± 12 nM. To assess
binding specificity, a competition assay was performed. 200 nM StrRAG1 samples were individually incubated with 75 nM labeled 12-RSS probe followed by the addition of either
unlabeled 12-RSS DNA or an oligonucleotide of the same length but
lacking both nonamer and heptamer consensus sequences (nonspecific
DNA). The 12-RSS is an effective competitor, whereas greater than a
6-fold excess of nonspecific competitor results in only an
30%
decrease in the shifted complex (Fig. 3c). This 30%
decrease may be the result of a fraction of StrRAG1 that binds non-specifically to DNA as well as the presence of a high concentration of polyanionic charge contributed by the phosphate groups of DNA, which
may affect the overall stability of the complex in the electrophoretic field within the gel matrix. Gel-sieving effects combined with prolonged exposure of protein-DNA complexes to an electrophoretic field
(3 h) and electrophoretic buffer may affect the results obtained using
the EMSA methodology (40, 41). These perturbations may especially
affect the interactions between two macromolecular species when
opposite charge attraction is a major factor in the stabilization of
the complex. Such interactions have been implicated in the RAG1-RSS
interaction (13, 14).

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Fig. 3.
By EMSA, StrRAG1 binds specifically to the
12-RSS, forming a single protein-DNA species. a, EMSA using
75 nM labeled 12-RSS double-stranded oligonucleotide
(40-mer) and increasing amounts of StrRAG1, as indicated
above the lanes. The inset shows the
saturation curve obtained from quantifying the bands corresponding to
free probe and 12-RSS·StrRAG1 complex. The fraction bound represents
the fraction of 12-RSS·StrRAG1 complex formed normalized relative to
the maximal binding obtained at saturation. M, molecular
mass standards. b, EMSA using 75 nM 12-RSS probe
and increasing amounts of MBP-RAG1. A discontinuous 3.5%/8%
polyacrylamide gel was used, as indicated at the right.
c, competitive EMSA to assess specificity. 200 nM StrRAG1 was incubated with 75 nM labeled
12-RSS in the absence (lane 2) or presence of increasing
concentrations of unlabeled 12-RSS (lanes 3-9) or
nonspecific (nsp) DNA (lanes 10-14). Labeled
probe and competitor DNA were added simultaneously to the StrRAG1
protein.
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|
In Solution, StrRAG1 Binds with Higher Affinity and Specificity to
the 12-RSS Than Detected by EMSA--
To circumvent the disadvantages
associated with EMSAs, we used fluorescence polarization to investigate
interactions between StrRAG1 and DNA in solution. Fluorescence
anisotropy provides an indirect measure of the rotational diffusion of
molecules in solution, which is in turn dependent on their size and
shape (larger molecules have lower tumbling rates and have higher
anisotropy values) (34). The first experiments made use of
double-stranded 12-RSS DNA or nonspecific oligonucleotides labeled at
the 5' end of one strand with fluorescein (12-RSS-Fl,
nonspecific-DNA-Fl), which allowed measurement of the anisotropy of the
DNA molecules (excitation wavelength 492 nm, emission recorded at 520 nm). 12-RSS-Fl or nonspecific-DNA-DNA-Fl (50 nM) was
incubated with increasing amounts of StrRAG1, and the fluorescein
polarization anisotropy was determined (Fig.
4a). The curve for 12-RSS-Fl
has a sigmoidal shape with increasing anisotropy values that reach
saturation around 100 nM StrRAG1 (monomeric protein
concentration) and an increase of anisotropy of 90-100% over the
value in the absence of protein. The best fit of the data points
(correlation = 96%; sum of squares = 0.65 × 10
4) was achieved using a Hill binding isotherm with
n = 1.8 (Equations 3 and 4), yielding an apparent
KD = 41 ± 8 nM (reflecting monomeric protein concentration). This value for KD
corresponds to an affinity of interaction higher than that determined
by EMSA experiments. The increase in 12-RSS DNA anisotropy observed in this experiment is due solely to the formation of protein-DNA complexes, which increase the molecular size reported by the
fluorophore attached to the DNA. In Fig. 4b the same
anisotropy data are represented as a function of molar ratio of
StrRAG1:12-RSS, which should yield an inflection point indicating
binding stoichiometry (37). The inflection point observed indicates
dimeric protein binding to 12-RSS DNA. In the case of
nonspecific-DNA-DNA-Fl there is a reduced, non-saturable linear
increase in anisotropy, corresponding to nonspecific binding of StrRAG1
to DNA. The specificity of the StrRAG1-DNA interaction was also
examined by anisotropy competition experiments. First, 50 nM 12-RSS-Fl was incubated with 100 nM StrRAG1,
and then either specific 12-RSS or nonspecific unlabeled DNA was added
in increasing amounts to the mixture (Fig. 4c). Similar to
the results of EMSA competition experiments, only the 12-RSS was
capable of dissociating the 12- RSS-Fl·StrRAG1 complex and reducing
the anisotropy to values characteristic of 12-RSS-Fl DNA alone.
Even a 100-fold excess of nonspecific DNA resulted in only a slight
reduction (12-15%) in the anisotropy of the complex.

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Fig. 4.
StrRAG1 binds as a dimer to the 12-RSS with
KD 41 nM. a, fluorescence polarization anisotropy of
50 nM 12-RSS-Fl fluorescein labeled DNA (black
squares) incubated in solution with increasing concentrations of
StrRAG1. Each data point represents an average of five individual
determinations. Using a Hill equation with n = 1.8 generates a fit with 96% correlation and a sum of squares = 0.65 × 10 4 (Equations 3 and 4, see "Experimental
Procedures"). Open triangles show results obtained with
fluorescein-labeled nonspecific DNA (nsp-DNAFl) incubated
with the same concentrations of StrRAG1. Each data point represents an
average of three individual determinations. b, determination
of the stoichiometry of StrRAG1-DNA binding by fluorescence anisotropy.
The anisotropy data of panel a are plotted against the molar
ratio [StrRAG1]/[12-RSS-Fl] present in the reaction (calculated for
monomeric StrRAG1). The intersection of the linear-fitting of the slope
and plateau yields a molar ratio of two StrRAG1 monomers per 12-RSS-Fl
DNA molecule (37). c, the 12-RSS·StrRAG1 complex is
resistant to competition with nonspecific DNA. 50 nM
12-RSS-Fl was incubated with 100 nM StrRAG1, and after
DNA-protein complex formation, either unlabeled 12-RSS (filled
squares) or nonspecific DNA (open triangles) were added
in increasing concentrations. Each data point represents the average of
three individual determinations. The arrow points the level
of anisotropy of free 12-RSS-Fl. The inset depicts the same
data points with the concentration of unlabeled competitor DNA
represented on a logarithmic scale.
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In conclusion, polarization anisotropy experiments using fluorescently
labeled 12-RSS show that StrRAG1 binds relatively tightly (KD = 41 ± 8 nM) and specifically
to DNA in solution. The inflection point in our anisotropy data
strongly suggest a molar ratio of protein to DNA of 2:1 in the
StrRAG1·RSS complexes.
DNA Rotational Reorientation Times (
) Are Consistent with
Binding of a StrRAG1 Dimer to the 12-RSS--
To further investigate
the stoichiometry of StrRAG1 in protein-DNA complexes in solution, we
used frequency domain fluorescence spectroscopy (FD). Frequency domain
fluorescence measures the fast changes in either phase or modulation
that occur in the fluorescent light emitted a short time after the
fluorophore is excited with a coherent source of light (34). These
changes allow direct calculation of the fluorophore life-time
and
the rotational reorientation time
(when the emitted fluorescence
comes through a vertically oriented polarizer), which is the inverse
value of the rotational diffusion coefficient of the molecule (tumbling rate of molecule).
We first measured the lifetime of the fluorescein fluorophore attached
to the 12-RSS or nonspecific DNA (Table
I) and found that it was not
significantly changed by the addition of StrRAG1 or HMG2 (a nonspecific
DNA binder) from the standard 3.6-4.1-ns value reported for
fluorescein alone in pH 7.5 aqueous solutions (42). These important
control measurements demonstrate that neither DNA bases nor protein
amino acid side groups interact with fluorescein or significantly
change the fluorophore environment when these proteins interact with
the DNA.
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Table I
Lifetime and rotation reorientation time values measured by frequency
domain spectroscopy
Fluorescein life-time ( ) measured by FD spectroscopy either for DNA
alone or DNA in the presence of various concentrations of StrRAG1 or
HMG2.* The short life time for 12-RSS-FI reflects the presence of
free, uncoupled fluorescein. The FD anisotropy decay traces were fitted
by Global analysis to a double exponential decay (35)
from which the local rotation reorientation time local
(corresponding to the fluorophore) and global rotation reorientation
time global (corresponding to the entire molecule or
ensemble of molecules) could be obtained.
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We then used frequency domain differential phase and polarized
modulation ratio measurements to determine the time-resolved anisotropy
decay of the fluorescein-labeled 12-RSS or nonspecific DNA in the
presence or absence of StrRAG1 or HMG2 (Table I). Fitting of the
anisotropy decay curves was performed using a double exponential
equation (35, 43) (see "Experimental Procedures"). In this
equation, one exponent describes the rotational diffusion of the
fluorophore with respect to the whole molecule (local rotational reorientation time,
local), and the other describes the
global rotation of the macromolecules (global rotational reorientation time,
global). In all cases,
local is
almost unchanged from the 0.35-0.4-ns value, confirming that the
fluorophore does not interact with the bases or the proteins during
protein-DNA complex formation.
global values change
considerably upon protein binding, indicating significant changes in
the size of 12-RSS-Fl DNA in complexes with StrRAG1 under conditions of
partial binding (50 nM protein) or near binding saturation
(100 nM StrRAG1) (44, 45). Importantly, much lower global
rotational reorientation times are reported for nonspecific DNA in the
presence of StrRAG1, either at 50 or 100 nM protein
concentrations, confirming the lower representation of protein-DNA
species that form non-specifically in solution in our assay conditions
(Table I). From the calculated KD, one can estimate
that more than 85% of the 50 nM 12-RSS DNA is present in
protein-DNA complexes in the mixture with 100 nM StrRAG1
(saturation conditions). Hence, the global rotation reorientation time
obtained in this reaction predominantly represents that of the
protein-DNA complex. Using the rotation reorientation time
Global = 48.03 ± 10.05 ns obtained under
saturation binding conditions and the original Perrin equation
(Equation 7; see "Experimental Procedures"), we can estimate the
size of the StrRAG1·12-RSS complex to be between
175 kDa (using
h = 0 ml/g, an anhydrous sphere) and 120 kDa (using
h = 0.4 ml/g, the maximal degree of hydration) (46,
47). Given the apparent molecular masses of the protein (72 kDa) and
12-RSS-Fl DNA (31 kDa), we conclude that StrRAG1 is most likely present
as a dimer in these complexes.
Tryptophan Fluorescence Anisotropy of StrRAG1 Increases in the
Presence of the 12-RSS--
We wanted to monitor by fluorescence
anisotropy the effect of adding DNA to StrRAG1 having as a reference
the protein present in solution. This set of polarization anisotropy
experiments took advantage of the intrinsic fluorophores in the StrRAG1
protein (primarily its 6 tryptophan residues) to serve as the
fluorescent reporters (excitation wavelength 295 nm, emission recorded
at 327 nm). Hence, these experiments measured changes in the size of
the protein complexes whether or not they were bound to DNA. StrRAG1
(250 nM) was incubated with increasing concentrations of
unlabeled 12-RSS, and tryptophan anisotropy was measured (Fig. 5). With the 12-RSS, anisotropy increased
abruptly and reached a saturation value about 20% greater than that of
the protein alone. The data fit a rectangular hyperbola, with an
apparent KD = 16.5 ± 7 nM. This
apparent KD value reflects the 12-RSS DNA
concentration present in solution at half-maximal increase in
anisotropy. Because the protein binds as a dimer, this
KD value is expected to be approximately one-half of
the KD value obtained from the data points in Fig. 4a, which reflects the concentration of monomeric StrRAG1
present in the reaction under similar conditions. The increase in
tryptophan anisotropy is predominantly due to an increase in the
molecular size of a significant population of StrRAG1 molecules in the
presence of 12-RSS DNA. This increase can be due either to formation of protein-DNA complexes or to formation of higher order protein-protein oligomers not associated with DNA. In the presence of nonspecific DNA,
there is an increase in anisotropy to levels 5-6% above the value for
the free protein, and the increase is linear and nonsaturable.

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Fig. 5.
Intrinsic tryptophan fluorescence
polarization anisotropy of 250 nM StrRAG1 incubated with
increasing amounts of 12-RSS DNA (dark diamonds) or
nonspecific DNA (open triangles). Each data point
represents the average of five individual determinations. The 12-RSS
data points were fit with a rectangular hyperbola ( 2 = 5 × 10 4, R2 = 0.89, yielding
an apparent KD = 16.5 ± 7 nM,
taking into account the quantum yield corrections for the bound
fraction (see Equation 6, "Experimental Procedures"), whereas the
nonspecific (nsp)-DNA data were fit to a linear
function.
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Binding of StrRAG1 to DNA Causes Major Changes in the Protein
Intrinsic Fluorescence Spectrum--
We next wanted to ask whether DNA
alters the conformation of the StrRAG1 protein. First, we tested if the
intrinsic emission fluorescence spectrum (excitation at 280 nm,
emission at 350 nm) of StrRAG1 protein is altered by DNA. Upon
incubation of 100 nM StrRAG1 with increasing amounts of
12-RSS oligonucleotide, a drastic decrease of emission fluorescence
intensity (fluorescence quenching) occurred, saturating at 50-60% of
the fluorescence in the absence of DNA (Fig.
6a). As with the increase in
tryptophan polarization anisotropy (Fig. 5), this decrease in
fluorescence caused by 12-RSS DNA can be fit to a rectangular hyperbola
(Fig. 6c), but the effect does not quantitatively parallel
the DNA binding phenomenon. It is important to note that since the
quenching effect occurs at concentrations of DNA lower than those
expected to yield a significant amount of protein-DNA complexes, it is
possible that interaction with the 12-RSS induces a change in protein
configuration (see below). The decrease in fluorescence
intensity induced by 12-RSS is associated with a slight shift of the
maximum emission intensity (
max) toward the red portion
of the spectrum from 335 to 340 nm. A small decrease in fluorescence
emission intensity was obtained in the presence of nonspecific DNA
(Fig. 6, b and c), but the quenching was not
saturable, and there are no detectable spectral changes of
max in this case. As a control for the quenching
potential of the amine groups of DNA bases, we also used an amount of
bovine serum albumin that gives a fluorescence intensity similar to
that of 100 nM StrRAG1; the addition of increasing amounts
of 12-RSS DNA resulted in minimal quenching (Fig. 6c).

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Fig. 6.
Steady-state emission fluorescence of StrRAG1
in the absence or presence of DNA. a, intrinsic emission
fluorescence spectra of 100 nM StrRAG1 in the absence or
presence of increasing amounts of unlabeled 12-RSS DNA. b,
intrinsic emission fluorescence spectra of 100 nM StrRAG1
in the absence or presence of increasing amounts of unlabeled
nonspecific DNA. c, relative decrease in emission intrinsic
fluorescence ( F/F) plotted as a
function of DNA concentration. Black diamonds, StrRAG1 plus
12-RSS (curve represents the fitting of data points to a rectangular
hyperbola); open triangles, StrRAG1plus nonspecific
(nsp) DNA; shaded circles, 300 nM
bovine serum albumin plus 12-RSS. In the latter two cases, lines were
obtained by fitting the data points to a linear function.
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Next, we performed kinetic experiments to determine whether quenching
and binding occur simultaneously. First, 50 nM 12-RSS labeled with fluorescein was mixed with 50 nM StrRAG1, and
the change in DNA anisotropy was measured with time (Fig.
7a). 10-15 s after mixing,
anisotropy reached levels expected from the static equilibrium
experiment (Fig. 4a) and then remained unchanged for 13 min.
This indicates that protein-DNA complexes are formed very rapidly. In
the second experiment, 50 nM StrRAG1 was incubated with 10 nM 12-RSS or with nonspecific DNA, and intrinsic StrRAG1 fluorescence was measured with time. The 12-RSS induced a complex, slow
multiphasic quenching of StrRAG1 fluorescence, quite distinct from that
induced by nonspecific DNA. The first phase, which lasts ~200 s, was
characterized by a substantial (17%) decrease in fluorescence. In the
second phase, lasting between 200 and 700 s, fluorescence increased transiently, perhaps because of transient changes in fluorophore environment caused by protein conformational changes. Surprisingly, StrRAG1 intrinsic fluorescence quenching continued even
after 800 s but at a much reduced rate. Although the kinetic behavior of StrRAG1 intrinsic fluorescence will require further study,
it is clear that DNA binding (Fig. 7a) occurs much more rapidly than fluorescence quenching (Fig. 7b). We conclude
that fluorescence quenching predominantly reflects conformational
changes induced by the 12-RSS DNA.

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Fig. 7.
Kinetics of StrRAG1 interaction with 12-RSS
DNA. a, time course of anisotropy of 50 nM 12- RSS fluorescein-labeled DNA mixed after 100 s (arrow
indicates the time of mixing) either with 50 nM StrRAG1
(dark lines) or with a similar volume of control binding
buffer (gray lines) (see "Experimental
Procedures"). b, time course of intrinsic emission
fluorescence of 50 nM StrRAG1 after mixing
(arrow) either with 10 nM 12-RSS (dark
squares) or with 10 nM nonspecific DNA (gray
squares).
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Acrylamide Quenching Indicates That 12-RSS Binding Causes StrRAG1
Fluorophores to Become More Exposed to Solvent--
To investigate the
mechanism by which 12-RSS DNA acts as a quencher of StrRAG1 intrinsic
fluorescence, we tested the effect of acrylamide on the StrRAG1
fluorescence in the presence or absence of DNA. Acrylamide is a small
soluble molecule that quenches both tryptophan and tyrosine
fluorescence and has access to regions of a protein where a
macromolecule like DNA does not. Therefore, adding acrylamide after
protein-DNA complex formation tests whether the intrinsic protein
fluorophores become more exposed to the solvent interface (in the case
of a conformational change) or less exposed to solvent molecules (if
they are involved in stable complexes either with DNA bases or with
other surrounding amino acids) relative to the protein fluorophores in
the absence of DNA. Increasing concentrations of acrylamide were added
to 600 nM StrRAG1 in the absence or presence of 150 nM specific 12-RSS or nonspecific DNA, and protein
intrinsic emission fluorescence was measured. In Fig.
8, the ordinate represents the
ratio between the original sample fluorescence in the absence of
acrylamide (Fo) and the sample fluorescence
(F) at the indicated concentration of acrylamide on the
abscissa (Stern-Volmer plot) (38). In Table
II, we present the values of the
quenching constants derived from fitting our data to the quadratic
Equation 8 (see "Experimental Procedures" and "Discussion").
The upward curvature of the Stern-Volmer plots indicates the combined
effect of dynamic and static quenching (see "Experimental
Procedures"). The slope of the curve, however, is increased by the
presence of DNA (and more by 12-RSS than nonspecific DNA), indicating
that DNA causes some of the amino acid fluorophores to become more
accessible to the quencher, as reflected in the increased values of
their dynamic quenching constants (Table II). This result is also
consistent with the red shift of the spectra that occurs upon the
addition of DNA. StrRAG1 has a wealth of fluorophores (6 Trp and 20 Tyr residues), and it is, therefore, difficult to address where in StrRAG1
quenching occurs. In summary, these experiments indicate an important
role for dynamic (solvent mediated) quenching of intrinsic fluorophores
of StrRAG1 upon interaction with DNA, and that this effect is stronger
with specific than nonspecific DNA.

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Fig. 8.
Stern-Volmer plots for acrylamide quenching
of StrRAG1 (600 nM) intrinsic emission fluorescence in the
absence (dark squares) or presence of 150 nM 12-RSS (open triangles) or 150 nM nonspecific DNA (open circles).
Data points were fit with 95% confidence interval to a second order
parabola with ordinate intercept = 1.
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Table II
The static KS and dynamic KD quenching constants
expressed as mM 1, derived from fitting of the
acrylamide quenching data (Fig. 8) to Equation 8 (see "Experimental
Procedures")
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Circular Dichroism Provides Further Support for a DNA-induced
Conformational Change in StrRAG1--
We then asked if the
12-RSS-induced conformational change in StrRAG1 observed by
fluorescence could also be detected by circular dichroism (CD). Fig.
9 presents the close and far UV CD
spectra of 500 nM StrRAG1 protein alone or in the presence
of increasing amounts of 12-RSS DNA. The prominent minima peaks at 210 and 222 nm are indicative of significant
helical content in StrRAG1 (48). Upon 12-RSS addition, the molar ellipticity at both of these
characteristic wavelengths decreases dramatically, saturating at a DNA
concentration of 50-60 nM (Fig. 9b). When
nonspecific DNA was used, the spectral changes observed were smaller
(Fig. 8, c and d; compare b with
d).

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Fig. 9.
Circular dichroism (CD) spectra displaying
molar ellipticity (Mol. ) of StrRAG1
versus wavelength (nm) in the absence or presence of
DNA. a, CD spectra of 500 nM StrRAG1 in the
absence or presence of increasing concentrations of 12-RSS DNA.
b, molar ellipticity of StrRAG1 as a function of 12-RSS
concentration, recorded at 210 nm (open circles), 222 nm
(filled squares), and 245 nm (open triangles).
c, CD spectra of 500 nM StrRAG1 in the absence
or presence of increasing concentrations of nonspecific DNA.
d, molar ellipticity of StrRAG1 as a function of nonspecific
DNA concentration, recorded at 210 nm (open circles), 222 nm
(filled squares), and 245 nm (open
triangles).
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Together, the StrRAG1 fluorescence and CD spectral changes induced by
12-RSS DNA lead to the conclusion that RSS sequences induce major
conformational changes in StrRAG1. In addition, because these changes
are much greater with 12-RSS than nonspecific DNA, sequence-specific
interactions apparently underlie much of the observed changes.
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DISCUSSION |
RSS Binding by RAG1 in the Absence of RAG2--
In an early study
using an in vivo one-hybrid assay, core RAG1 was found to
interact specifically with a tandem array of eight 12-RSS elements, and
RAG2 appeared to enhance binding only modestly (14). Based on this and
other findings, it was suggested that RSS recognition might be a
two-step process consisting of RAG1 binding followed by recruitment of
RAG2 (13, 14). The results of subsequent in vitro
experiments provided at least two types of arguments against the
general validity of this model. First, it was found that the initial
rate of DNA cleavage could be enhanced by preincubating MBP-RAG1 with
RAG2 but not by preincubating MBP-RAG1 with the DNA (49). Second,
studies relying primarily on EMSA methodology and RAG1 proteins with a
bulky tag suggested that the RAG1-RSS interaction was not sufficiently
specific to be physiologically relevant (see the Introduction). The
only study to investigate the equilibrium binding of highly purified,
dimeric RAG1 core protein used EMSA methodology and documented an
approximate 10-fold specificity for the RSS and a reasonably high
affinity (KD = 100 nM) (22). However, as
confirmed here, the MBP-RAG1 core protein used in those experiments
forms three species with the RSS in which the protein is present in
various oligomeric states.
We show here that a RAG1 core protein with only a short eight amino
acid tag is soluble and forms stable dimers in solution in the absence
of DNA. Both EMSA and solution DNA binding assays demonstrate that this
protein binds the RSS as a dimer and that it does so specifically and
with moderately high affinity (in solution, apparent
KD = 41 ± 8 nM). Furthermore, the complex that forms with the 12-RSS is relatively resistant to competition with nonspecific DNA (Fig. 4c) and by a number
of different measures is distinct from that formed upon interaction with nonspecific DNA. These findings lead us to reconsider the relevance of the two-step model for RSS recognition noted above.
In developing lymphocytes in the G1 phase of the cell
cycle, RAG1 and RAG2 are co-expressed, with RAG2 apparently in molar excess (50). In the S, G2, and M phases of the cell cycle,
however, RAG2 is rendered unstable by phosphorylation by a
cyclin-dependent kinase (51, 52), and RAG2 levels drop
dramatically, whereas RAG1 levels are almost unchanged (30). We would
like to propose that in S, G2, and M phase cells, RAG1 can
bind to RSSs in the absence of RAG2. Furthermore, we propose that as
these cells complete mitosis and enter G1 and RAG2 levels
begin to rise, RAG2 can be recruited to RSSs by pre-bound RAG1,
creating a catalytically competent complex. This does not exclude the
possibility that some and perhaps the majority of RSS recognition is
performed by RAG1·RAG2 complexes. Instead, our results lead to the
idea that a pattern of RAG1-RSS interactions laid down in
S/G2/M may be relevant to the subsequent targeting of the
recombination reaction.
A DNA-induced Conformational Change in RAG1--
It has been
proposed previously that RAG2 can induce a conformational change in
RAG1 that facilitates binding of RAG1 to the RSS and perhaps also the
appropriate folding of the RAG1 DDE active site (3, 18, 27). Until the
experiments reported here, however, there have been no studies of
conformational changes in the RAG proteins either induced by one
another or by DNA. We have been able to study the interactions of
StrRAG1 with DNA in solution by monitoring protein intrinsic
fluorescence, circular dichroism spectra, and fluorescence
polarization. These studies would likely have been considerably less
informative if RAG1 had been attached to bulky MBP or glutathione
S-transferase tags due to the spectral contributions of the
fusion partners.
Two pieces of evidence support the conclusion that RSS DNA induces a
conformational change in RAG1. The addition of RSS DNA to StrRAG1
results in (i) strong quenching of intrinsic StrRAG1 protein
fluorescence (Fig. 6) and (ii) a dramatic alteration in the CD spectrum
of the protein (Fig. 9). We consider each of these in turn. The
decrease in fluorescence intensity and red shift of
max
observed when StrRAG1 interacts with 12-RSS DNA could have two, not
mutually exclusive, explanations.
(i) Dynamic quenching could occur if some of the main fluorophores (Trp
and Tyr residues) of the protein undergo a change in environment from
the hydrophobic core to the solvent interface of the macromolecule,
where they are subject to dynamic quenching by solvent molecules. The
red shift of
max when StrRAG1 was incubated with 12-RSS
DNA is in itself an indication that the dielectric constant
of the
environment of some of the protein fluorophores increases, consistent
with the red shift reported for the indole ring in cyclohexane
versus water (53). This can only be caused by a
conformational change in the protein structure. Such changes would also
increase accessibility of the protein fluorophores to the small
molecule quencher acrylamide, and hence, one would expect stronger
acrylamide quenching for strRAG1 plus DNA than for strRAG1 alone. This
is exactly what is observed in Fig. 8.
(ii) Static quenching could occur if some of the internal fluorophores
of StrRAG1 became involved in direct complexes with amines of DNA
bases. Alternatively, the DNA molecule could be an indirect static
quencher if upon binding to StrRAG1, it induces a protein
conformational change that causes charged amino acid residues to form
direct complexes with Trp or Tyr fluorophores (the charge transfer that
occurs in such complexes forbids the decay of the excited state and
prevents photon emission). Direct static quenching by DNA does not
necessarily involve a protein conformational change. In these cases,
stronger acrylamide quenching should be observed for StrRAG1 alone than
for the 12-RSS·StrRAG1 complex since fluorophores involved in static
quenching cannot be quenched further by acrylamide. This scenario,
however, is in conflict with the data that we present in Fig. 8. This
reasoning was applied in assigning the static and dynamic acrylamide
quenching (KS, KD)
values presented in Table II, which were simply derived as solutions of
the quadratic Equation 8 (see "Experimental Procedures").
The kinetics of 12-RSS-induced quenching of StrRAG1 fluorescence shows
a complex pattern (Fig. 7b) and occurs
considerably more slowly than protein-DNA complex
formation. Quantitatively, the 12-RSS specifically and efficiently
quenches StrRAG1 intrinsic emission fluorescence, with the effect
reaching saturation at DNA concentrations where only a fraction of the
protein is engaged in forming StrRAG1·DNA complexes (Fig.
6c). Together, our observations are best explained by a
model in which dimers that dissociate from the DNA either retain the
newly acquired conformation or further modify their configuration (as
the transient increase in fluorescence in Fig. 7b suggests)
and, hence, are as susceptible or even more susceptible to solvent
quenching than those bound to DNA. Therefore, it is possible that the
12-RSS acts as a "catalyst" to induce major conformational changes
in StrRAG1 whether bound to the DNA or not.
We note that the oligomerization state of other DNA-binding proteins
such as
repressor (54) and human immunodeficiency virus integrase
(55) has been shown to be influenced by the presence of DNA. In the
case of human immunodeficiency virus integrase, time-resolved
anisotropy measurements of its tryptophan rotation reorientation times
show that integrase exists in various oligomeric states in solution,
and Mg2+ and DNA cause dissociation of the tetrameric form
to the monomeric form that is found in complex with DNA (55, 56).
The addition of 12-RSS DNA and to a lesser extent nonspecific DNA
results in a dramatic change in the CD spectrum of StrRAG1 (Fig. 9). A
possible explanation for the DNA-induced decrease in both 209- and
222-nm peaks (indicative of
helicity (57)) is the movement of the
peptide groups from an orientation almost parallel to the helix axis
towards a tilted configuration with carbonyl groups pointing outwards,
due to a drastic change in the protein backbone. Such a reorientation
of the peptide groups has been predicted to decrease the amplitude of
long wavelength CD peaks of proteins with
helix content (58).
The stoichiometry of pre-cleavage RAG1·RAG2·RSS complexes is
controversial, with two reports suggesting that these complexes contain
more than two (presumably four) monomers of RAG1 (25, 26), and another
suggesting that they contain only a single dimer of RAG1 (24). If the
former model is correct, then since DNA-bound StrRAG1 dimers show no
propensity to associate with other StrRAG1 dimers, it is likely that
stable association of two dimers of the RAG1 core with an RSS requires
RAG2. The situation might be different for the full-length RAG1
protein, which contains an additional zinc-dependent
dimerization domain (59).
In conclusion, we demonstrate that the core RAG1 protein containing a
small epitope tag binds the RSS as a dimer and undergoes a
conformational change upon doing so. Our results provide a starting point for future studies of possible RAG2-induced conformational changes in RAG1.