 |
INTRODUCTION |
Human factor XI (FXI)1
is a plasma glycoprotein (~5% carbohydrate) involved in the
intrinsic pathway of blood coagulation (1). FXI exists as a covalently
linked two-chain homodimer, thus making it unique among coagulation
factors. Dimeric FXI circulates in plasma as a zymogen in a 1:2
stoichiometric complex with the cofactor high molecular weight
kininogen at plasma concentrations in the range of 4-6 µg/ml (~30
nM) (2, 3). Each subunit of the dimer contains 607 amino
acids and has a molecular mass of ~80,000 Da based on reducing SDS
gel electrophoresis (4). The activation of this zymogen to an active
serine protease can be carried out in the presence of a polyanionic
surface by activated factor XII and high molecular weight kininogen,
thrombin, or activated FXI (FXIa) (4, 5). Activation is dependent upon
cleavage of the peptide bond between Arg369 and
Ile370 (6). Each subunit of FXIa is composed of a heavy
(50,000 Da) and light (30,000 Da) chain held together via disulfide
bonds (4). The light chain functions as a serine protease involved in
the activation of coagulation FIX (7). The heavy chain of FXIa contains
four tandem repeat sequences called Apple domains consisting of 90-91
amino acids each (8). Each of the four Apple domains contains three
internal disulfide bonds, and the first and fourth Apple domains each
contain one additional cysteine residue (9). Cys321 in the
A4 domain has been implicated in the formation of a covalent FXI dimer
(9). The first Apple domain (A1) contains distinct binding sites for
cofactor high molecular weight kininogen, for thrombin, and for the
Kringle 2 domain of prothrombin (10-12), whereas the second (A2)
contains a FIX-binding site (13); the third (A3) has been implicated in
interactions with platelets, with heparin, and with FIX (14-17); and
the fourth (A4) contains an activated factor XII-binding site along
with its ability to mediate dimer formation (9, 18, 19).
One of the more provocative questions surrounding FXI is its
homodimeric structure and how that relates to its physiologic function
in blood coagulation. FXI is activated on the platelet surface, and the
resulting FXIa subsequently activates FIX (7). The recombinant A3
domain of FXI binds to the activated platelet surface with the same
stoichiometry (~1500 sites/platelet) as that found for full-length
dimeric FXI, suggesting that one A3 domain/dimer is responsible for the
interaction with the platelet surface (16). Since it is difficult to
rationalize one A3 (17) and/or A2 (13) domain of FXI binding to both
FIX and to the platelet surface simultaneously, it has been
hypothesized that the A3 domain from one subunit of the dimer binds the
platelet, while the A2 and/or A3 domain of the opposite subunit
interacts with FIX. Gailani et al. (20) recently
demonstrated that recombinantly derived FXI monomers in which the A4
domain of FXI was replaced with that of prekallikrein (FXI/PKA4) in
comparison with their dimeric counterparts (FXI/PKA4-Ala326
and wild-type FXI) were functionally defective in a partial
thromboplastin time assay utilizing platelets as a surface, but normal
with phospholipids. These results are consistent with the
aforementioned hypothesis since the monomeric chimera bound to the
platelet surface lacks an additional free A2 and/or A3 domain for
interaction with FIX and would therefore be deficient in its ability to
activate FIX. Additionally, FXI dimerization may be important for its
ability to activate FIX since cleavage occurs at two spatially distinct sites. A detailed kinetic analysis of the activation of FIX by FXIa
revealed that FIX is activated via a processive reaction mechanism
without release of a singly cleaved intermediate (21). These findings
suggest the possibility that each active site within the dimer carries
out a single cleavage without the need for reorientation of the FIX
molecule between successive cleavages, further suggesting that it may
be functionally advantageous for FXI to exist as a dimeric molecule.
To fully understand the functional significance of FXI homodimer
formation, we aim to define the structural determinants and mechanism
involved in this process. Meijers et al. (19) demonstrated that at least part of the molecular information required for mediating dimer formation between the two identical subunits appears to reside
within the A4 domain of FXI. Chimeric tissue plasminogen activator
molecules with the A4 domain of FXI substituted for the finger and
growth factor domains exist as dimers, as shown by gel filtration
analysis (19). In the same chimeras as well as the full-length FXI
protein, the replacement of Cys321 with serine results in
molecules that also exist as dimers, as determined by gel filtration
(19). These results strongly suggest that the A4 domains mediate
noncovalent interactions resulting in dimer formation, which is
subsequently stabilized by a covalent linkage between cysteine residues
in each monomer at position 321 (9, 19). We have therefore prepared the
rA4 domain of FXI as well as a C321S mutant to obviate covalent dimer
formation and to permit determinations of binding constants under
conditions of varying pH values and ionic strength. We employed a rapid
gel filtration procedure similar to that described by Manning et
al. (22) for the determination of natural and recombinant
hemoglobin dissociation constants (KD).
 |
EXPERIMENTAL PROCEDURES |
Materials--
Pfu polymerase, DNA markers (
DNA-HindIII digest, pBR322 DNA-MspI digest, and
X174 DNA-HaeIII digest), restriction enzymes BamHI and PstI, and NEB buffer 2 were from New
England Biolabs Inc. (Beverly, MA). All reagents used for SDS-PAGE were
purchased from National Diagnostics, Inc. (Atlanta, GA). Ammonium
persulfate and
-mercaptoethanol were purchased from Bio-Rad.
Bacto-Tryptone, Bacto-yeast extract, LB broth base (Lennox L
broth base), T4 DNA ligase, dNTPs, and prestained protein molecular
mass markers (low range) were from Life Technologies, Inc.
DIAFLO® ultrafiltration membranes (3000-Da cutoff) and the
Amicon 8200 stirred cell ultrafiltration apparatus were from Amicon,
Inc. (Beverly, MA). Isopropyl-
-D-thiogalactopyranoside
(IPTG) was purchased from LabScientific, Inc. (Princeton, NJ). HEPES,
CAPS, MES, Tris, cysteine, urea, guanidine, dithionitrobenzoate, and the MW-GF-70 gel filtration molecular mass marker kit were from Sigma.
M15(pREP4) cells, the QIAexpress® pQE-9
vector, nickel-nitrilotriacetic acid (Ni2+-NTA) resin, the
QIAEX II gel extraction kit, and the QIAquick polymerase chain reaction
(PCR) purification kit were from QIAGEN Inc. (Chatsworth, CA). The
Wizard® Plus miniprep DNA purification system was from
Promega (Madison, WI). Bovine FXa was purchased from Hematologic
Technologies, Inc. (Essex Junction, VT) or Enzyme Research
Laboratories, Inc. (South Bend, IN). The enzymes lysozyme, trypsin, and
endoproteinase Lys-C were from Calbiochem. HiTrap metal-chelating,
Superose 12 (10 × 300 mm), and Superdex 75 (16 × 600 mm)
columns were from Amersham Pharmacia Biotech. The
Ultrafree®-15 centrifugal filter device, the Millex-GV
0.22-µm filter unit, and the Millex-HV 0.45-µm filter unit were
from Millipore Corp. (Bedford, MA). The BCA protein assay reagent and
the Slide-A-Lyzer® 3.5K dialysis cassettes were
from Pierce. Sequenase Version 2.0 and reagents were supplied by United
States Biochemical Corp. Spectrapor® dialysis membrane
(9.3 ml/cm) was purchased from Spectrum Medical Industries, Inc.
(Laguna Hills, CA). The chromogenic substrate S-2765 for measurement of
FXa activity was purchased from Chromogenix (Mölndal, Sweden).
The apolar probe 4,4'-dianilino-1,1'-binaphthyl-5,5'-disulfonic acid
dipotassium salt (bis-ANS) was from Molecular Probes, Inc. (Eugene, OR).
Expression of Human FXI Apple 4 Domain and Mutants in Escherichia
coli--
The rA4 domain construct contains sequences that code for
Phe271-Glu361. The construct was generated by
utilizing PCR and FXI cDNA as a template (a 2.1-kilobase
pair EcoRI fragment containing the complete FXI
coding sequence, a generous gift from Drs. Dominic W. Chung, Kazuo
Fujikawa, and Earl W. Davie, Department of Biochemistry, University of
Washington, Seattle, WA) to create a 318-base pair insert that is
identical to that coding for the A4 domain of FXI.
Primers that flank either end of the A4 domain coding sequence were
prepared by Life Technologies, Inc. The 5'-upstream primer contains an
engineered BamHI restriction site (GGATCC) along
with a FXa cut site (ATCGAAGGTAGA) that, when translated
(Ile-Glu-Gly-Arg), was used to efficiently cleave an N-terminal
6-histidine tag. The 3'-downstream primer contains an engineered
PstI restriction site (CTGCAG) along with a stop
codon (TAA) to ensure transcription termination. The sequence of the
upstream primer (5'-BamHI) is as follows:
5'-CGCGGATCC(ATCGAAGGTAGA)TTCTGCCATTCTTCATTTTAC. The sequence of the downstream primer (3'-PstI) is as follows:
5'-AAAACTGCAG(TTA)CTCATTATCCATTTTACACAA. The PCR was set up
as follows: 10 ng of FXI cDNA as template, a 1 µM
concentration of both the upstream (5'-BamHI) and downstream (3'-PstI) primers, 2.5 units of cloned Pfu DNA
polymerase, 10 µl of 10× Pfu reaction buffer, and dNTPs
at 0.2 mM brought to a final volume of 100 µl. The
reaction mixture was then overlaid with 100 µl of mineral oil to
prevent evaporation. The reaction mixtures were placed in a PerkinElmer
Life Sciences DNA thermal cycler. The reactions were heated to 94 °C
for 7 min, followed by 40 cycles at 94 °C for 1 min, 50 °C for 1 min, and 75 °C for 3 min. The reaction was completed by incubation
at 75 °C for 7 min and a final incubation at 4 °C. The PCR was
subsequently purified with the QIAquick PCR purification kit.
PCR was also performed in the generation of the C321S mutant
(rA4-C321S). The PCR-based mutagenesis protocol was adapted from Picard
et al. (23). Template plasmid DNA was obtained using a
QIAGEN plasmid purification kit (~0.3-kilobase pair human FXI A4
domain subcloned into a pQE-9 expression vector). The mutagenesis reaction follows a three-step protocol. Step 1 involves synthesis of a
megaprimer using a mutagenic primer and a downstream primer (3'-PstI). The mutagenic primer is as follows:
5'-CCAAGCATCC(A)GCAACGAAGG. The 11th base in the primer was switched
from thymine in the wild-type sequence to adenine, thus allowing
Cys321 to be replaced by serine in the mutant amino acid
sequence. The reactions contained 10 ng of DNA template, 10 pmol (100 nM) each of the mutagenic and 3'-PstI primers,
2.5 units of cloned Pfu DNA polymerase, 10 µl of 10×
Pfu reaction buffer, and dNTPs at 0.2 mM brought
to a final volume of 95 µl. 10 cycles of amplification were performed
for each step (94 °C for 1 min, 48 °C for 1 min, and 72 °C for
2 min). The reaction was completed by incubation at 75 °C for 5 min
with a final incubation at 4 °C. Addition of 50 pmol of the upstream
primer (5'-BamHI) to the aqueous phase begins step 2, and
addition of 50 pmol of the downstream primer (3'-PstI)
initiates step 3. The same amplification protocol as employed in step 1 is used in steps 2 and 3. The insert was cut with BamHI and
PstI restriction enzymes using the protocols supplied by New
England Biolabs Inc. The insert was ligated to the QIAexpress pQE-9
vector, and transformations were carried out using competent E. coli K12-derived M15(pREP4) cells. Expression from the
pQE-9 vector was induced by the addition of IPTG.
Expression, Purification, and Folding of the rA4
Domain--
Large-scale expression cultures (1 liter) were propagated
in LB broth in the presence of 100 µg/ml ampicillin and 50 µg/ml kanamycin at 37 °C. The large-scale cultures were inoculated from small-scale growth cultures (1:40, v/v). The cultures were grown until
A600 reached 0.6-0.7. Induction of protein
expression required the addition of 0.5 mM IPTG, followed
by further incubation and shaking at 37 °C for 3 h. Cells were
harvested by centrifugation (4000 × g for 20 min) and
then stored as cell pellets at
70 °C. The concentration of protein
was determined by absorbance at 280 nm employing an extinction
coefficient of 12,420 M
1
cm
1 (per dimer). SDS-PAGE analysis was used
to determine purity and to assure noncovalent association of subunits.
Harvested cell pellets stored at
70 °C were resuspended in Buffer
A (6 M guanidine hydrochloride, 0.1 M
NaH2PO4, and 0.01 M Tris-HCl, pH
8.0) at 5 ml of buffer/g of cells and allowed to stir at room
temperature for 1 h. The supernatant was collected from the cell
lysate following centrifugation at 10,000 × g for 15 min at 4 °C. The crude extract was then mixed with a
Ni2+-NTA resin slurry (50% Ni2+-NTA and 50%
Buffer A, 10-ml total volume). The mixture was then allowed to stir at
room temperature for 1 h. The crude extract/resin mixture was
loaded onto a 10-ml polypropylene column (QIAGEN Inc.). A series of
wash steps then ensued: 5 column volumes of Buffer B (8 M
urea, 0.1 M NaH2PO4, and 0.01 M Tris-HCl, pH 8.0), a wash with Buffer C (8 M
urea, 0.1 M NaH2PO4, and 0.01 M Tris-HCl, pH 6.3) until the eluant absorbance at 280 nm
was <0.01, and 20 ml of Buffer D (8 M urea, 0.1 M NaH2PO4, and 0.01 M
Tris-HCl, pH 5.9). The FXI rA4 domain was then eluted with 20 ml of
buffer E (8 M urea, 0.1 M
NaH2PO4, and 0.01 M Tris-HCl, pH
4.5). Fractions containing protein were visualized by 15%
SDS-PAGE.
The protein was then refolded using a protocol that makes use of a
thiol/disulfide exchange in which cysteine is utilized as the low
molecular mass reducing molecule (24). Protein concentration was held
in the range of 200-250 µg/ml. A Spectrapor dialysis membrane (9.3 ml/cm) with a 3500-Da cutoff was used. The starting buffer consisted of
2 M urea, 20 mM Tris-HCl, 100 mM
NaCl, and 2 mM cysteine at pH 9.0. A series of seven buffer
changes was used to reduce the urea and cysteine concentrations (1 M urea and 1 mM cysteine; 0.5 M
urea and 0.5 mM cysteine; 0.25 M urea and 250 µM cysteine; 0.1 M urea and 100 µM cysteine; and 0 urea and 0 cysteine). Upon completion
of dialysis, the FXI rA4 domain was then concentrated via an Amicon
stirred cell ultrafiltration apparatus using DIAFLO ultrafiltration
membranes (3000-kDa cutoff). The protein solution was typically
concentrated down to a volume of 2-4 ml, and the protein concentration
(5.0 to 2.5 mg/ml) was determined. SDS-PAGE under reducing conditions
was used throughout to monitor purification and under nonreducing
conditions to detect correctly oxidized FXI rA4 domain.
The N-terminal 6-histidine tag was removed by cleavage with bovine FXa.
Bovine FXa cleaves proteins at the C-terminal side of the recognition
sequence Ile-Glu-Gly-Arg, which is identical to that engineered at the
N terminus of the FXI rA4 domain. Cleavage buffer consisted of 20 mM Tris-HCl and 100 mM NaCl, pH 9.0. A 1:50
molar ratio of FXa to the FXI rA4 domain was used. The reaction mixture
was incubated at 37 °C for 16-18 h. Upon completion of the
reaction, success of cleavage was determined by SDS-PAGE analysis. Following the cleavage reaction, separation of the His-tagged FXI rA4
domain from the non-His-tagged FXI rA4 domain was carried out using the
HiTrap metal-chelating column (5 ml). The HiTrap resin was charged with
Ni2+ ions by applying 2.5 ml of a 0.1 M
NiSO4 solution. The cleavage reaction was applied to the
column, followed by a 5-column volume (25 ml) wash step (20 mM Tris-HCl and 100 mM NaCl at pH 9.0). The
desired non-His-tagged FXI rA4 domain along with FXa was then eluted
with 20 mM imidazole in the above buffer system.
The final step in the purification made use of gel filtration
chromatography to separate the FXI rA4 domain from bovine FXa and to
obtain protein that formed productive dimers. A Superdex 75 gel
filtration column was equilibrated in 20 mM HEPES and 100 mM NaCl, pH 7.4. The proteins were resolved at a flow rate
of 1 ml/min, and protein elution was monitored by absorbance at 280 nm.
Fractions that eluted at a retention time representative of the dimeric
FXI rA4 domain were pooled and concentrated with a Millipore
concentrator. To determine the multimeric state of the eluted protein,
data compiled for a standard curve were generated using the MW-GF-70
gel filtration molecular mass marker kit. To assess purity, all
fractions were assayed for the presence of FXa using the chromogenic
substrate S-2765. The protein was subsequently examined by
SDS-PAGE.
The rA4 domain was examined by HPLC and matrix-assisted laser
desorption ionization time-of-flight mass spectroscopy (analysis conducted by the Protein Chemistry Laboratory at the University of
Pennsylvania, Philadelphia). The data demonstrated the presence of a
single homogeneous species of protein with a molecular mass of
19,836 Da, which is consistent with the calculated molecular mass of
the rA4 domain, which is 19,888 Da. Ellman's reagent
(5,5'-dithiobis(2-nitrobenzoic acid)) was used to determine free
sulfhydryl groups (25), which indicated <0.04 mol of free sulfhydryl
group/mol of rA4 domain.
Size Exclusion Chromatography--
An Amersham Pharmacia Biotech
fast protein liquid chromatography (FPLC) system was used along with an
Amersham Pharmacia Biotech FPLC high resolution Superose 12 column
(10 × 300 mm) to determine monomer-dimer
KD for the FXI rA4 domain C321S mutant. A Superdex
75 column was also employed in purification and confirmation of dimer
formation. To determine the multimeric state of the eluted proteins,
data for a standard curve were compiled at varying pH values and salt
concentrations using the MW-GF-70 gel filtration molecular mass marker
kit. To construct standard curves, four proteins were employed with
molecular masses as follows: albumin, 66,000 Da; carbonic anhydrase,
29,000 Da; cytochrome c, 12,400 Da; and aprotinin, 6500 Da.
The effects of varying protein concentration, pH, and salt
concentration were determined. All buffer solutions used to study pH
effects contained 20 mM buffering reagent, 100 mM NaCl, and 1 mM EDTA. Buffering reagents for
the various pH values tested were as follows: CAPS, pH 10.0; Tris-HCl,
pH 9.0 and 8.0; HEPES, pH 7.0 and 7.4; and MES, pH 6.0. All buffer
solutions used to study salt effects contained 20 mM HEPES,
the appropriate NaCl concentration (0.025-2.0 M), and 1 mM EDTA, pH 7.4. For all experimental studies, the proteins
were dialyzed against the appropriate buffer for 16-18 h in
Slide-A-Lyzer dialysis cassettes. The resultant protein solution was
cleared of any precipitate by centrifugation, and protein concentration
was determined spectrophotometrically by absorbance at 280 nm with the
previously stated extinction coefficient. Samples were then prepared by
dilution of concentrated stock solutions and incubated for
3 h at
room temperature to ensure that equilibrium was established. Samples of
100 µl at various protein concentrations were injected onto the
pre-equilibrated column. Elution of resolved protein species was in the
same buffer as the sample equilibration at a flow rate of 1 ml/min. The
elution profile was followed by absorbance at 280 nm. The chromatograms were analyzed using FPLC Director software (Amersham Pharmacia Biotech). The retention times were determined by comparison of elution
peaks at peak half-width and peak half-height. The calculated area
under the elution peaks was used to determine the amount and percentage
of dimeric and monomeric species present in the sample. A plot of the
percent dimeric rA4-C321S as a function of the total protein
concentration was used to determine dimer KD. All
experimentally determined KD values (KD(app)) were divided by a dilution
factor of 6.5 ± 0.27. The dilution factor was employed since
dilution of the protein sample occurs as the protein is eluted through
the column. The justification for this correction can be found under
"Results" and in Refs. 22 and 36. The dilution factor equals the
peak width at half-height (milliliter) divided by the sample load
volume (100 µl) (22, 36).
Conversion of KD to the Change in Binding Energy
(
G0)--
The Gibbs free energy of dissociation was
calculated using the equation
G0 =
RT ln KD, where R is the gas
constant (1.987 cal × mol
1 × K
1), T is the absolute temperature
at which experiments were done (298 K), and KD is
the dissociation constant or the concentration at which equal amounts
of dimer and monomer exist in solution.
Fluorescence Spectroscopy--
Fluorescence measurements were
performed on an SLM-AMINCO/Bowman Series 2 luminescence
spectrometer. The stock solutions of bis-ANS were filtered, and
the concentration was determined by absorbance at 385 nm using an
extinction coefficient of
385 = 16,790 cm
1 M
1
(30). Varying amounts of the apolar probe were used to titrate the rA4
domain while monitoring extrinsic probe fluorescence. Protein
concentrations were held constant at 0.5 µM. The samples were excited at 385 nm, and emission was monitored at 497 nm. The
excitation and emission slit widths were set to 4 and 8 nm, respectively. All results were corrected with respect to spectra of
buffer and protein background.
The establishment of dissociation kinetics was followed in a
time-dependent manner by monitoring the increase in bis-ANS
fluorescence as well. For dissociation kinetics, the protein was
diluted to a concentration of 0.5 µM in the presence of
20 µM bis-ANS. The data were evaluated using the program
KaleidagraphTM (Abelbeck Software) and fitted using a
first-order equation with a single rate constant. The
association rate constant (kon) for the
conversion of monomer to dimer was calculated using the following equation, KD = koff/kon, where
KD represents the dissociation constant determined
by size exclusion chromatography and koff is the
dissociation rate constant determined by fluorescence spectroscopy.
 |
RESULTS |
Expression and Purification of the rA4 Domain of FXI--
Since
the FXI rA4 domain is expressed as insoluble inclusion bodies, its
recovery requires solubilization, folding, and oxidation. To facilitate
the purification of the rA4 domain, we generated an N-terminal
6-histidine-tagged fusion protein expressed in an E. coli
K12-derived strain (M15(pREP4)). DNA corresponding to the
A4 domain of FXI was PCR-amplified and placed in the pQE-9 expression
vector. The vector contains a regulatable promoter/operator element
allowing expression to be controlled by the addition of IPTG. Fig.
1A shows the expression of the
rA4 domain compared with uninduced cells and cells induced with
IPTG.

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Fig. 1.
SDS-PAGE analysis of FXI rA4 domain
purification. A, small-scale expression cultures were
analyzed by 15% SDS-PAGE for their ability to express the FXI rA4
domain. Cells were grown to mid-log phase (A600 ~ 0.6-0.7), induced with 0.5 mM IPTG, and incubated at
37 °C for an additional 3 h. Lane 1 represents a
sample taken prior to the addition of IPTG, and lane 2 is
representative of a sample induced with IPTG. Samples exposed to IPTG
expressed the expected ~11-kDa protein, as shown by the appearance of
a protein band not present in the "uninduced" sample. The uninduced
sample (lane 1) demonstrates positive regulation of protein
expression since no corresponding protein band was detected in the
sample prior to the addition of IPTG. B, lane
1 represents the final protein product following gel
filtration chromatography on a Superdex 75 column. The sample loaded in
lane 1 was ~40 µg, indicating the high level of purity
achieved.
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|
Solubilization and purification took place in the presence of the
denaturants 6 M guanidine hydrochloride and 8 M
urea. We utilized the N-terminal 6-His tag as a molecular handle to
facilitate binding of the recombinant protein by a high affinity
interaction for the Ni2+-NTA resin (KD = 10
13 M at pH 8.0). Wash steps
with subsequent stepwise decreases in pH allowed for the removal of
contaminating proteins (Buffers B-D). The protein of interest was
eluted at pH 4.5 (Buffer E), a condition favoring the protonation of
the 6-His tag and thereby diminishing the interaction made with the
Ni2+-NTA resin. The eluant was collected in 1-ml fractions
and analyzed by both absorbance at 280 nm and SDS-PAGE. The FXI rA4
domain eluted in the first 10 ml of buffer E. This first step proved to
be quite advantageous, allowing us to purify the fusion protein from
the crude extract in one step with considerable purity. Purity was
assessed by resolving the fractions by 15% SDS-PAGE and staining with
Coomassie Blue (Fig. 1). The typical yield after the Ni2+
affinity chromatography step from a 500-ml culture was ~10 mg of
denatured fusion protein.
The protein was then refolded using a protocol that makes use of a
thiol/disulfide exchange in which cysteine is utilized as the low
molecular mass reducing molecule (24). By stepwise removal of the urea,
the protein slowly renatures, allowing the correct cysteine residues to
come in close proximity of one another. Furthermore, the presence of
cysteine molecules allows for a rapid disulfide exchange or reshuffling
of incorrect disulfide bonds, which ultimately allows for formation of
the most energetically favorable disulfide linkages. Even though the
protein concentration during the folding step was held relatively low
(200-250 µg/ml), a considerable amount of protein precipitate was
observed at this step due to nonspecific aggregate formation.
Approximately 30-50% of the protein was lost, making it the most
inefficient step in the purification scheme. Protein folding,
especially dependent upon disulfide bond formation, is highly sensitive
to fluctuations in pH, temperature, and protein concentration (25, 26),
which may account for the variability between preparations and the
substantial loss of protein at this step. The protein that remained
soluble was separated from the precipitate by centrifugation and
ultrafiltration. Success of folding was examined by SDS-PAGE analysis
under reducing and nonreducing conditions (Fig.
2) and gel filtration. Despite the
considerable loss of protein at this step, protein that remained soluble was capable of forming productive dimers exclusively, as
evidenced by distinct bands resolved by SDS-PAGE and single peaks on
gel filtration chromatograms. Exclusive dimer formation at this step
provides some evidence that protein folding was successful since
improperly folded rA4 domains would conceivably produce a wide range of
multimers. Mixed disulfide-bonded species tend to resolve as diffuse
bands and multimeric ladders upon nonreducing SDS-PAGE as well as broad
elution peaks upon gel filtration chromatography (25, 26). Total
recovery of protein post-folding based on yields achieved from the
initial Ni2+-NTA chromatography step were in the
neighborhood of 50-70% or 5-7 mg.

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Fig. 2.
SDS-PAGE of nonreduced and reduced samples of
FXI rA4 and the rA4-C321S mutant. The rA4 domain migrated as a
disulfide-linked dimer under nonreducing conditions and as a monomer
under reducing conditions. The C321S mutant resolved as a monomer under
both conditions, as expected. Lane M contains molecular mass
markers.
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The removal of the 6-His tag plus four amino-terminal residues
(Met-Arg-Gly-Ser) was accomplished by cleavage with bovine FXa. Success
of cleavage includes the removal of the 6-His tag plus the four amino
acids (Ile-Glu-Gly-Arg) composing the FXa cut site. The molecular mass
of the protein was reduced by 1854 Da to give a calculated molecular
mass of 9943 Da/monomer. The cleavage reaction was monitored by
SDS-PAGE analysis.
To separate the cleaved FXI rA4 domain (without the 6-His tag) from the
noncleaved FXI rA4 domain, the digested sample was applied to a HiTrap
metal-chelating column charged with Ni2+ ions. The digested
sample contained primarily three species: bovine FXa, the 6-His-tagged
A4 domain, and the non-6-His-tagged A4 domain. Following an initial
wash step (20 mM Tris-HCl and 100 mM NaCl, pH
9.0), all three species remained on the column, which was somewhat
unexpected since FXa and the nontagged A4 domain are devoid of
histidine tags. However, both FXa and the A4 domain are capable of
binding to a Mono QTM anion exchange column (data not
shown) at pH 9.0, suggesting that these proteins are somewhat
negatively charged at this pH. This may account for a nonspecific
charge interaction with the positive Ni2+ ions bound to the
column. Elution of the nontagged A4 domain was accomplished by addition
of 20 mM imidazole to the buffer. Once again, 1-ml
fractions were collected and assessed by absorbance at 280 nm and
SDS-PAGE. Bovine FXa and the nontagged A4 domain coeluted in fractions
10-20 (10-20 ml). As a final step in the purification, the remaining
protein was resolved on a Superdex 75 gel filtration column. The gel
filtration step accomplished two goals: removal of bovine FXa (~45
kDa) and purification of only the A4 domain protein (~20 kDa) that is
capable of forming productive dimers. Fractions that eluted at volumes
corresponding to the dimer form of the FXI rA4 domain (~76 ml) were
collected. Elution volumes were compared with a standard curve of log
molecular mass versus elution volume generated by
chromatography of the gel filtration molecular mass marker kit (data
not shown). The collected fractions were also assayed with the
chromogenic substrate S2765 to confirm separation of FXa from the A4
domain. Final purity was determined by SDS-PAGE analysis. It was
conservatively estimated from the gel that the A4 domain was purified
to >95% homogeneity (Fig. 1B). Based on the protein yield
achieved after the Ni2+-NTA chromatography step, a final
yield of ~1-2 mg (10-20% recovery) of pure FXI rA4 domain remained
per 500 ml of expression culture.
Reducing and Nonreducing SDS-PAGE--
The SDS-PAGE results
provided additional confirmation for both covalent (rA4) and
noncovalent (rA4-C321S) dimer formation (Fig. 2). Size comparisons were
made against low molecular mass markers (Life Technologies, Inc.).
Samples of both rA4 and rA4-C321S (±
-mercaptoethanol) were resolved
by SDS-PAGE. Both reduced samples resolved corresponding to monomer
molecular masses migrating between markers of 14 and 6 kDa (expected
size of 9944 Da). The nonreduced rA4 domain resolved as a dimer
migrating between molecular mass markers of 29 and 18 (expected size of
19,888 Da). The nonreduced rA4-C321S mutant resolved as a
monomer identical to that of the reduced samples (Fig. 2). The results
indicate that the wild-type rA4 domain exists as a dimer and that dimer
formation is mediated by a disulfide linkage, as would be expected. The
rA4-C321S mutant dimer does not contain a cysteine residue at position
321, but was still capable of forming dimers under native conditions,
as seen from the gel filtration results (Fig.
3). Under nonreducing conditions, SDS was
capable of disrupting the C321S dimer interaction, confirming that the
mutant dimer interaction is mediated via noncovalent intersubunit amino
acid interactions.

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Fig. 3.
Representative dissociation curve and elution
profile for rA4-C321S. Shown is a plot of percent dimer
versus total rA4-C321S concentration (micromolar). 100%
dimer represents a theoretical condition at infinite concentration of
rA4-C321S. The conditions of the experiment include 20 mM
HEPES, 100 mM NaCl, and 1 mM EDTA, pH
7.4. An apparent dissociation constant (KD) of 1.49 µM (uncorrected for dilution of the sample on the column)
was determined for the experimental conditions. The inset
represents elution profiles of varying total protein concentrations
(0.25, 0.5, 1.0, 2.0, and 10 µM) injected onto a
pre-equilibrated Superose 12 column. The protein was diluted to the
appropriate concentration in 20 mM HEPES, 100 mM NaCl, and 1 mM EDTA at pH 7.4 and incubated
at room temperature (~25 °C) for >3 h to establish equilibrium.
The elution profiles were followed by the absorbance at 280 nm, and the
amounts of protein present as monomer and dimer were assessed by peak
area determined by FPLC Director software. With a flow rate of 1 ml/min, the retention times are equivalent to elution volume in
milliliters. The peak labeled Dimer refers to the
elution of dimer, and that labeled Monomer refers to that of
monomer.
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Dimerization Assay--
A high resolution Superose 12 column
(10 × 300 mm) was used to develop a dimerization assay to
determine monomer-dimer KD for mutant rA4 domains of
FXI. Similar assays have been described by both Manning et
al. (22) and Gallagher and Huber (27) to measure
tetramer-dimer dissociation constants of natural and recombinant hemoglobins and the monomer-dimer equilibrium of M15
-galactosidase from E. coli, respectively. To determine the multimeric
state of the eluted protein, data for a standard curve were compiled under varying conditions using the MW-GF-70 gel filtration molecular mass marker kit (see "Experimental Procedures"). Peak area,
determined by FPLC Director software, was used to calculate the percent
dimeric and monomeric species for each sample. A plot of percent
dimeric rA4-C321S as a function of the total rA4-C321S concentration
prior to application to the column gave a hyperbolic curve. A
representative dissociation curve for the C321S mutant along with
elution profiles (chromatograms) are presented in Fig. 3. The elution
volumes of the two peaks labeled dimer (14.4 ml) and monomer
(16.7 ml) in Fig. 3 correspond to molecular masses of 21,060 and 8291 Da, respectively. These masses correspond well to the theoretical
calculated monomer (9944 Da) and dimer (19,888 Da) forms of the
protein. Under conditions approximating physiologic salt concentration
and pH (20 mM HEPES, 100 mM NaCl, and 1 mM EDTA, pH 7.4), it was determined that the monomer-dimer
equilibrium was characterized by a
KD(app) of 1.5 ± 0.2 µM. This value of KD(app)
was corrected for a 6.5-fold dilution factor during the gel filtration
procedure (22, 36) to give a KD value of 229 nM. The justification for this correction is given in the
last paragraph under "Results" and in Refs. 22 and 36.
The results indicate that the monomer-dimer equilibrium of rA4-C321S is
reversible, as evidenced by a shift in the ratio of dimer to monomer as
a function of decreasing protein concentration. The reversibility of
the equilibrium was also demonstrated by subjecting the dimer peak to a
second round of chromatography and observing the presence of two peaks
corresponding to dimer and monomer (data not shown). The equilibrium
can be driven to completion, generating either nearly all dimeric or
monomeric species, indicating that all the protein present in the
reaction is functional and participates in the equilibrium process. The elution times were highly reproducible for both varying amounts of the
dimeric and monomeric species, as evidenced by peak width at
half-height determinations. All elution profiles resolved as highly
symmetric peaks, indicative of a mixture of monomeric and dimeric
rA4-C321S domains in slow equilibrium between species (28). The lack of
peak broadening also rules out potentially nonspecific interactions
with the column material.
These results indicate that the A4 domain by itself is sufficient to
cause dimerization and that the interchain disulfide bond at C321S is
not necessary for dimer formation. These findings are consistent with
the results of Meijers et al. (19) for the Cys321 mutant of full-length FXI, which migrated as a
dimeric molecule upon gel filtration. Conclusively, the analysis
indicates that the C321S mutant exists in a reversible monomer-dimer
equilibrium that can be measured and used to determine the
KD(app) for this interaction under
varying conditions.
A time course for dissociation using the same size exclusion
chromatography method was examined (Fig.
4). A stock solution of the rA4-C321S
domain (~25 µM) was diluted to 2.0, 0.75, and 0.35 µM in buffer containing 20 mM HEPES, 100 mM NaCl, and 1 mM EDTA, pH 7.4. Aliquots were
then resolved on a Superose 12 column at various times post-dilution.
The data were subjected to linear regression analysis. We observed that
the equilibrium was essentially stable over the time scale tested,
demonstrating that equilibrium had already been achieved prior to our
measurements. The final monomer concentration for all three trials
(2.0, 0.75, and 0.35 µM) after equilibrium was
established and was internally consistent with our experimentally
determined KD(app) of 1.5 µM. A solution of rA4-C321S at concentrations of 2.0, 0.75, and 0.35 µM should be ~40, 65, and 80%
monomeric, respectively. This is exactly what we observed. This
observation leads us to conclude that all the protein participates in
the equilibrium process. However, this method of analysis does not
allow us to effectively study the dissociation kinetics of the dimer
interaction.

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Fig. 4.
Time course for dimer dissociation measured
by size exclusion chromatography. Starting at ~25
µM, the dimer (rA4-C321S) was dissociated by dilution
with buffer to 2, 0.75, or 0.35 µM. At different times
after dilution, aliquots were taken and analyzed on a Superose 12 gel
filtration column. All reactions were performed at 25 °C in 20 mM HEPES, 100 mM NaCl, and 1 mM
EDTA, pH 7.4. The monomeric species was detected by peak area of the
elution profiles as described under "Experimental Procedures." The
time scale does not reflect the true time post-dilution, and it was
estimated that 100 s had elapsed before the column separation was
initiated. The 100 s can be accounted for by sample dilution and
mixing, sample application to the FPLC system, and injection onto the
column. The data were fit by linear regression analysis.
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Dimer Dissociation (rA4-C321S) as a Function of pH--
To analyze
the dimer interaction, size exclusion chromatography at various
concentrations of rA4-C321S was used to determine KD
as a function of pH in a range between 6.0 and 9.0. A composite of the
dissociation curves is represented in Fig. 5. The results are presented in Table
I. The tightest interaction (KD ~ 229 nM) for the dimer was seen
in the acidic pH range tested (6.0-7.4). From pH 6.0 (KD = 283 ± 32 nM,
G = 8.9 kcal/mol) to pH 7.4 (KD = 229 ± 26 nM,
G = 9.1 kcal/mol),
little or no change (
KD = 54 nM,

G = 0.2 kcal/mol) was observed for the binding
interaction. In contrast, a marked change was observed when comparing
pH 7.4 with pH 8.0 (KD = 229 ± 26 nM and
G = 9.1 kcal/mol
versus KD = 614 ± 146 nM and
G = 8.5 kcal/mol). An ~2.7-fold
increase in KD was observed, followed by a further
1.5-fold increase in KD at pH 9.0 (KD = 955 ± 122 nM,
G = 8.2 kcal/mol). Conclusively, an increase in pH
causes a general destabilization of the rA4-C321S dimer interaction,
reflected in an overall ~4.0-fold increase in KD
(
G = 0.8 kcal/mol) from pH 6.0 to 9.0. The
destabilization trend observed as a function of pH provides evidence
for a charge interaction at the dimer interface that aids in mediating
dimer formation. A marked shift in binding affinity between pH 7.4 and
8.0 suggests the involvement of amino acid residue(s) with a
pKa of ~8.0.

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Fig. 5.
Composite of dissociation curves illustrating
pH effects on dimer dissociation. Plots of percent dimer
versus total C321S protein concentration were used to
determine dimer dissociation constants (KD) at a
range of pH values from 6.0 to 9.0. Total [C321S] represents the
concentration of dimer that would be present if the protein were all
dimer. Percent dimer is the percentage of protein that is actually
dimer at various C321S concentrations.
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Table I
Effect of pH on dimer dissociation
Shown is the effect of pH on dimer dissociation represented in terms of
KD (µM) and Gibbs free energy
( G, kcal/mol). The KD values were
determined from plots of percent dimer versus [C321S]
utilizing Kaleidagraph software and were corrected for 6.5-fold
dilution of samples on the column as described under "Experimental
Procedures." The Gibbs free energy of dissociation was calculated
using the following equation: G = RT ln
KD, where R is the gas constant (1.987 cal × mol 1 × K 1), T is the
absolute temperature (298 K), and KD is the dimer
dissociation constant experimentally determined.
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Chromatography was also attempted for pH values below 6.0 and above
10.0 that are outside the range shown in Table
II. The resultant chromatograms revealed
evidence of peak broadening and inconsistencies in peak elution times.
Therefore, it was not possible to interpret the results. However, these
results suggest that the A4 domain (rA4-C321S) may be somewhat
conformationally unstable in pH conditions below 6.0 and above 10.0. These results suggest three possibilities: global structural changes in
the protein and/or changes in the rate of dissociation or interaction
with the column material.
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Table II
Effect of salt on dimer dissociation
Shown is the effect of salt on dimer dissociation represented in terms
of KD (µM) and Gibbs free energy
( G, kcal/mol). The KD values were
determined from plots of percent dimer versus [C321S]
utilizing Kaleidagraph software and were corrected for 6.5-fold
dilution of samples on the column as described under "Experimental
Procedures." The Gibbs free energy of dissociation was calculated
using the following equation: G = RT ln
KD, where R is the gas constant (1.987 cal × mol 1 × K 1), T is the
absolute temperature (298 K), and KD is the dimer
dissociation constant experimentally determined.
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Dimer Dissociation (rA4-C321S) as a Function of Salt
Concentration--
To analyze the dimer interaction, size exclusion
chromatography at various concentrations of rA4-C321S was used to
determine dissociation constants as a function of salt concentration
between 0.025 and 2.0 M. A composite of the dissociation
curves is represented in Fig. 6. The
results are presented in the Table II. Salt concentrations of 0.025 and
0.05 M exhibited the tightest binding interactions, with
KD values equal to ~163 nM
(
G = 9.3 kcal/mol). At 0.1 M NaCl,
KD = 229 ± 26 nM and
G = 9.1 kcal/mol were observed. A 15-fold increase
in salt concentration (1.5 M NaCl) resulted in an
~2.9-fold decrease in binding affinity (KD = 662 ± 169 nM,
G = 8.4 kcal/mol). A
salt concentration of 1.0 M NaCl (KD = 255 ± 66 nM,
G = 9.0 kcal/mol) did
not reflect the overall trend and may be indicative of an aberrant observation. In general, an increase in salt concentration can cause a
destabilization of the dimer interaction, as demonstrated by the
~4-fold decrease in binding affinity as the NaCl concentration was
increased from 0.025 to 2.0 M. The results obtained while examining the effects of salt on the dimer interaction are consistent with the notion that electrostatic interactions contribute to FXI rA4
domain dimerization. Salt concentrations tested below 25 mM
produced aberrant results, i.e. an increase in retention times for both the dimeric and monomeric species. These findings indicate that protein is being retarded by the column material as a
result of nonspecific interactions with the column matrix or global
changes in conformation.

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Fig. 6.
Composite of dissociation curves illustrating
salt effects on dimer dissociation. Plots of percent dimer
versus total C321S protein concentration were used to
determine dimer dissociation constants (KD) at a
range of salt concentrations from 0.025 to 2.0 M. Total
[C321S] represents the concentration of dimer that would be present
if the protein were all dimer. Percent dimer is the percentage of
protein that is actually dimer at various C321S concentrations.
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Bis-ANS Titration--
We conducted studies to assess the role of
hydrophobic interactions in A4 dimer formation. The apolar probe
bis-ANS was used to titrate the A4 domain while monitoring extrinsic
probe fluorescence. Upon binding to a hydrophobic moiety or hydrophobic
cluster on the surface of a protein, one observes a severalfold
increase in bis-ANS fluorescence intensity (29). In comparison with
bis-ANS emission maximum (553 nm) when the dye is unbound and in
aqueous medium, one also observes a blue-shifted emission maximum upon binding of the dye to a hydrophobic moiety (30). The extent of both the
fluorescence intensity increase and the shift in emission maxima is
strictly dependent upon the environment of the dye-binding site (31).
An increase in the bis-ANS fluorescence and a shift in the emission
maximum from 533 to 497 nm were observed upon addition of saturating
levels of bis-ANS (10 µM) to rA4-C321S (0.5 µM) (data not shown). When higher concentrations of dye
were added, we observed no further shift in the emission maximum, which is suggestive of a single dye-binding site (32). As a control, the
fluorescence intensity of 10 µM bis-ANS in buffer (in the absence of protein) was negligible compared with the fluorescence of
dye in the presence of protein (data not shown). We compared the
wild-type rA4 domain (0.5 µM) and the rA4-C321S domain
(0.5 µM) in the presence of increasing amounts of bis-ANS
and noted the change in relative bis-ANS fluorescence (Fig.
7). Binding of the probe to the mutant
rA4-C321S domain showed saturable binding, along with a marked increase
in probe fluorescence, as evidenced by a plot that was hyperbolic in
character (Fig. 7). The hyperbolic nature of the curve is indicative of
a single dye-binding site with a KD equal to ~2
µM. In contrast, the covalently linked, fully dimeric
wild-type rA4 domain showed little or no significant ability to bind
the apolar probe, as evidenced by the absence of a detectable increase
in relative bis-ANS fluorescence. These results suggest the absence of
hydrophobic sites on the surface of the covalently linked rA4 domain.
To test the specificity of the dye-binding site on the rA4-C321S
domain, a denatured rA4-C321S domain incubated in 6 M
guanidine hydrochloride was titrated with bis-ANS. The results
demonstrated a small amount of nonspecific binding of the dye, as
evidenced by the linear plot shown in Fig. 7. The dye-binding site
exposed on the surface of the partially dissociated native rA4-C321S
domain can be disrupted by exposure of the protein to denaturant. These
findings suggest that the hydrophobic site exposed only in the presence
of the dissociated dimer must contain some conformational
specificity.

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Fig. 7.
Titration of the A4 domain with bis-ANS.
Shown are the results of titration of the A4 domain at different
concentrations of the apolar probe bis-ANS. The increase in bis-ANS
fluorescence (arbitrary units) was measured at a fixed emission
wavelength of 497 nm. The C321S mutant under both denaturing (guanidine
hydrochloride (GuHCl)) and nondenaturing conditions and the
covalently linked wild-type (WT) A4 domains were at
concentrations of 0.5 µM. The covalently linked dimeric
wild-type A4 domain and the denatured C321S mutant showed no
significant change in fluorescence compared with the freely dissociable
C321S mutant. We were able to detect a change in the exposure of
hydrophobic patch(es) due to increased bis-ANS binding as result of A4
dimer (C321S) dissociation.
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In conclusion, these results give clear indication that a saturable
hydrophobic site(s) is exposed only upon dimer dissociation and that
this site(s) can be disrupted by denaturation of the protein. A
hydrophobic patch present only on the monomeric subunit gives credence
to the argument that a hydrophobic component exists at the homodimer
interface, which may be in part responsible for mediating A4 dimer formation.
Kinetics of rA4-C321S Dissociation--
The change in the
fluorescence of bis-ANS upon binding to the dissociated rA4-C321S
domain was useful for monitoring the kinetics of dissociation (Fig.
8). Concentrated stock solutions (25-100 µM) of both the rA4-C321S and rA4 domains were diluted to
a final concentration of 0.5 µM in the presence of 10 µM bis-ANS and monitored for increases in bis-ANS
fluorescence. We observed an increase in bis-ANS fluorescence for
rA4-C321S, which follows a single exponential reaction with a
dissociation rate constant (koff) of 4.3 × 103 s
1 for the dissociation of
dimer to monomer (Fig. 8). Utilizing the previously determined
KD of 229 ± 26 nM (size exclusion chromatography), we were able to calculate an association rate constant
(kon) for the conversion of monomer to dimer of
1.9 × 104 M
1
s
1 (see "Experimental Procedures"). The
determination of the kinetic parameters associated with the rA4 domain
monomer-dimer equilibrium provides a detailed look at the kinetic
mechanism behind the dimer interaction of the rA4 domain as well as the
noncovalent interaction of FXI as a whole. It is of critical importance
to point out that this assay is based on the assumption that bis-ANS
binds to the dissociated monomer at a much faster rate compared with
the rate of dissociation of the dimer. We can make this statement since the rate of encounter of a small molecule with a large molecule for a
diffusion-controlled reaction gives a second-order rate constant in the
range of 109-1011
M
1 s
1
(33). If we hypothesize that the true rate of association for the
apolar probe were altered by 3 orders magnitude (106
M
1 s
1),
accounting for changes in protein conformation upon association of the
probe with the protein, the rate of binding of the probe (104 s
1 at 10 µM
bis-ANS) to the protein would still be much faster compared with our
observed rate for dimer dissociation of 4.3 × 10
3 s
1. Therefore,
in this experiment, we are observing the dissociation of the rA4-C321S
dimer and not the binding of bis-ANS to the monomer. Moreover, the
covalently linked wild-type protein (rA4) does not bind the apolar
probe, demonstrating the specificity of the probe for the dimer
interface. These observations are in accordance with the bis-ANS
titration experiments, in which we observed no increase in bis-ANS
fluorescence in the presence of the wild-type rA4 domain.

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Fig. 8.
Kinetics of rA4-C321S dissociation as
monitored by the binding of bis-ANS. rA4-C321S and the rA4 domain
were diluted to 0.5 µM in the presence of 10 µM bis-ANS. The kinetics of dissociation was monitored by
following the increase in bis-ANS due to the binding of the probe to
the dissociated monomers (rA4-C321S). The covalently linked dimeric
wild-type rA4 domain showed no significant change in fluorescence
compared with the freely dissociable rA4-C321S mutant. The increase in
bis-ANS fluorescence was measured at a fixed wavelength of 497 nm
(excitation at 385 nm). The curve was fit according to a first-order
reaction.
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The findings of both the fluorescence assay (kinetics of dissociation)
and the gel filtration assay are mutually consistent. Based on the
experimentally determined rate of dissociation
(koff = 4.3 × 10
3 s
1), we
calculated a half-life (t1/2 for the dissociation
reaction of 160 s. In the time scale of the gel filtration
experiment in which the majority of the determinations were made 10 min
(3.7 half-lives) after dilution, we would expect at least 92% of the
equilibrium to have been achieved (Fig. 4). This provides an
explanation as to why in this experiment we observed no change in
monomer concentration over time and also provides a justification for
correcting values of KD(app) to account for the 6.5-fold dilution of protein samples during the gel filtration dimerization assay (see "Experimental Procedures").
 |
DISCUSSION |
The results presented in this paper demonstrate the feasibility of
studying the noncovalent dimer interaction of FXI by use of a
recombinantly generated A4 domain. Herein we report the
KD (~230 nM) and the Gibbs free energy
of dissociation (
G ~ 9 kcal/mol) for the
rA4-C321S noncovalent mutant under physiologic conditions, as
determined by size exclusion chromatography. We also report a
koff of 4.3 × 10
3 s
1 and a
calculated kon of 1.9 × 104
M
1 s
1
using fluorescence spectroscopy.
The FXI rA4 domain and the rA4-C321S mutant have been purified and
characterized, including their capacity to form both covalent and
noncovalent dimers, DNA sequencing of the recombinant DNA, mass
spectroscopic analysis of the protein product, and N-terminal sequence
analysis. Considering the enormous number of disulfide bond
combinations (135,135) that are theoretically possible in a protein
like the rA4 domain with a total of seven cysteines (25, 26), we took
great pains in characterizing the rA4 domains. The single most
important line of evidence that these proteins are conformationally
intact is the fact that they maintain their capacity to form dimers.
Moreover, the rA4 domain exists as a single species upon reverse-phase
HPLC, mass spectroscopy, and gel filtration, supporting the conclusion
that the protein is homogeneous and conformationally intact (34). We
also tested our proteins for their susceptibility to protease cleavage.
Both trypsin and endoproteinase Lys-C were unable to cleave the rA4 domains, as monitored by SDS-PAGE (data not shown). While elucidating the disulfide bond arrangement in FXI, McMullen et al. (9) similarly observed resistance to trypsin proteolysis for an isolated fragment containing the entire A4 domain
(His267-Met358). In the native conformation, a
majority of proteins possess some protease resistance, and proteolysis
typically takes place in regions that are typically exposed or
disordered (35). Therefore, one might expect that if the rA4 domain
were folded incorrectly, cleavage by the previously mentioned proteases
would occur more readily. We conclude that all of the rA4 domain
polypeptides exist in native conformation.
Molecular sieve chromatography or size exclusion chromatography has
been well described in the literature and has become a valuable tool
for determining equilibrium constants for reversibly associating
systems (28, 36, 37). Values obtained by this technique have been shown
to be reliably comparable to sedimentation equilibrium and fluorescence
depolarization assays, among other techniques designed to scrutinize
multimer equilibria (22, 27). The results presented in Figs. 3-6
provide evidence for a process in which a mixture of both dimeric and
monomeric rA4 domains is unaffected by the chromatography process and
exists in a slowly exchanging equilibrium (28). In a slowly exchanging
system like ours, the separation process (column chromatography) is
faster than the reversible exchange of dimer and monomer. This explains why we observed two well resolved, highly symmetrical peaks in our
study (28, 37).
We examined the influence of pH on the dissociation equilibrium using
the freely dissociable mutant (rA4-C321S) and observed an increase in
KD at the more alkaline pH values tested (Table I).
Although the pH effect could be due to deprotonation of an ionizable
group (or groups) on the protein resulting in a structural distortion
of the protein, we observed a high degree of peak uniformity and
consistency of retention times, suggesting that the proteins are
behaving uniformly both in respect to their Stokes radii and lack of
interaction with the column matrix (28). We can therefore theorize that
the pH effects are attributed to a deprotonation of a critical amino
acid side chain involved in the dimer interaction. The most significant
changes in KD occur in the pH range between 7.4 to
8.0 (~2.7-fold increase in KD), suggesting
deprotonation of a side chain with a pKa value of
~7.4-8.0. There are only two possible candidates: cysteine
(pKa = 8.5) and the N-terminal amino group
(pKa = 8.0) (38, 39). Since all cysteine residues
within the A4 domain in both the full-length FXI molecule (9) and our
own protein constructs (see "Results") are engaged in disulfide
linkages, there are no free sulfhydryl groups to mediate dimer
formation. The N-terminal amino group of the A4 domain
(Phe271) is engaged within FXI in a peptide bond with its
neighboring residue (Val270). Moreover, both the FXI rA4
and rA4-C321S domains are capable of forming dimers with and without
the histidine tag present (data not shown). Furthermore, if the N
terminus were located at the dimer interface, bovine Xa probably would
not have steric access to the cleavage site, and this would present a
problem for removal of the 6-His tag. Therefore, it is highly unlikely
that the N terminus of the rA4 subunit plays a role in the dimer
interaction. It is difficult to predict accurately the ionization
behavior of any one group without additional information. However, we
can conclude from our analysis that an increase in pH affects the ability of the A4 domain to associate, suggesting the possibility that
deprotonation of an amino acid side chain with a perturbed pKa due to its microenvironment directly weakens the monomer interactions. The obvious implication from this analysis is
that an electrostatic component mediates dimer formation; whether we
are observing a specific side chain contribution or global charge
repulsion remains to be seen. In any event, it is this type of
information that will provide the basic knowledge about the
oligomerization of proteins within a cell and the rationale for future
experimentation designed to answer questions about the nature of the
FXI dimer interface.
Electrostatic complementarity has been proposed to play a significant
role in the formation of a large number of protein complexes (43). For
example, homodimers of MyoD are subsequently destabilized by increasing
salt, which, in turn, confirmed an electrostatic contribution to
subunit interactions first suggested by crystallographic studies (44,
45). The study of cytochrome P-450 and P-450 reductase has revealed an
interaction that is also held together by complementary charged
residues (46-48). Electrostatic interactions play an important role in
the dimerization of FXI mediated by the A4 domain. The shift in
equilibrium toward monomer at the high concentrations of salt (Table
II) is thought to be a result of charge shielding by the addition of
ions and a reduction in the productive charge interactions at the
interface (49, 50). When small diffusible ions such as Na+
and Cl
are present in an aqueous environment, the ions
tend to concentrate in the vicinity of charges that are opposite in
sign (positive versus negative) (51). Therefore, the
increase in salt concentration would gradually neutralize the charge
interactions by subsequently weakening the resultant protein-protein
interaction at the A4 dimer interface. Under conditions of high salt,
water molecules tend to be excluded at protein interfaces (51, 52),
intensifying an existing hydrophobic interaction resulting in lower
KD values. Since we did not observe a large change
in KD or
G due to increasing salt
concentrations, we can speculate that the increase in salt may
intensify an existing hydrophobic interaction at the dimer interface
compensating for the destabilizing effect seen due to shielding of
charges at the interface.
The specificity of the bis-ANS probe for hydrophobic moieties on the
surface of proteins (53) has led to many studies, including the
characterization of protein interface contacts between subunits found
in coagulation cofactor VIII (32). The probe permitted the
identification of a surface-exposed hydrophobic site on dissociated rA4-C321S as well as the absence of hydrophobic sites on the covalently linked rA4 domain. The presence of a site only on the dissociated subunits is suggestive of its involvement in homodimer formation. These
results provide clear evidence of a hydrophobic component present at
the A4 dimer interface and demonstrate that the bis-ANS probe binds
only to a single affinity dye-binding site. These findings are also in
agreement with the salt-dependent changes seen in binding
affinity, which suggest that the hydrophobic component can compensate
for the charge destabilization caused by increasing salt concentrations.
Our study has clear implications for the assembly mechanism of FXI as
well as for the study of oligomeric proteins in general. Our results
are consistent with published studies of homodimer interfaces that
indicate that the overall proportions of nonpolar interactions (van der
Waals and hydrophobic) and polar interactions (hydrogen bonds) tend to
vary greatly from one oligomer interface to another (54). The majority
of the interfaces surveyed contained a mixture of small hydrophobic
patches, polar interactions, and water molecules scattered over the
entire interface (55).
This study provides an initial characterization of the interaction
between the two monomeric subunits of FXI and provides seminal
information that will guide future investigations of
platelet-associated coagulation reactions and function of FXI (56).
Since the dimeric nature of FXI may be critical for its ability to
function properly on the platelet surface (16, 20, 21), it is important
to understand the mechanism of FXI dimerization. A future goal is to
compare full-length FXI with the rA4 proteins to determine whether all
of the binding information resides within the A4 domain. It will be
interesting to compare the rA4 domain with the full-length FXI C321S
mutant to determine whether another region of the FXI molecule could
harbor additional binding energy. Such studies would be facilitated by
the development of dimerization assays such as fluorescence
polarization (44, 57) and isothermal titration calorimetry (58),
solution-based methods that provide a true equilibrium measurement that
does not require separation of free and bound species. Such studies,
together with structural information from NMR and x-ray crystallography
combined with site-directed mutational analysis, will allow us to
examine individual amino acid contributions made at the dimer interface.