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Enzyme dynamics during catalysis
Eisenmesser, Elan Zohar; Bosco, Daryl A; Akke, Mikael; Kern, Dorothee
Published in:
Science
DOI:
10.1126/science.1066176
2002
Link to publication
Citation for published version (APA):
Eisenmesser, E. Z., Bosco, D. A., Akke, M., & Kern, D. (2002). Enzyme dynamics during catalysis.
Science
,
295
(5559), 1520-1523. https://doi.org/10.1126/science.1066176
Total number of authors:
4
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Data (i) had to have small variance (as gauged by re-
ported SE) and (ii) had to be reported in units of kg of dry
weight per plant. A total of 385 species are represented
in the complete data set (including arborescent mono-
cots, dicots, and conifers). Data for the intraspecific
scaling of plant organ biomass were collected by B.J.E.
primarily from the agricultural and forestry literature
(25, 27, 28).
21.
M
L
, M
S
, and M
R
were each computed for an average plant
from each community or experimental manipulation
with the quotient of total plant biomass per site or
treatment and plant density. Model type II (reduced
major axis) regression analyses were then used to de-
termine scaling exponents and allometric constants (re-
gression slopes and y intercepts designated as ␣
RMA
and

RMA
, respectively), because functional rather than pre-
dictive relation were sought among variables that were
biologically interdependent and subject to unknown
measurement error (7). Because many authors failed to
report all of the necessary parameters required to assess
M
L
, M
S
, and M
R
, the sample size of regression analyses
varies across statistical comparisons.
22. Supplementary materials can be found on Science
Online at www.sciencemag.org/cgi/content/full/295/
5559/1517/DC1.
23. F. A. Bazzaz, in Plant Resource Allocation, F. A. Bazzaz,
J. Grace, Eds. (Academic Press, New York, 1997), pp.
1–37.
24. R. M. Callaway, E. H. DeLucia, W. H. Schlesinger,
Ecology 75, 147 (1994).
25. W. B. Smith, Allometric biomass equations for 98
species of herbs, shrubs, and small trees (North Cen-
tral Forest Experimental Station Research Note NC-
299, Forest Service U.S. Department of Agriculture,
Washington, DC, 1983).
26. B. J. Enquist, K. J. Niklas, Nature 410, 655 (2001).
27. W. H. Pearsall, Ann. Bot. 41, 549 (1927).
28. C. Monk, Bull. Torrey Bot. Club 93, 402.20 (1966).
29. We thank E. Charnov, A. Ellison, D. Ackerly, H.-C.
Spatz, L. Sack, and D. Raup for discussions or com-
ments on earlier drafts of this manuscript. This work
stems as an outgrowth of discussions from the Body
Size in Ecology and Evolution Working Group (F. A.
Smith, principle investigator) sponsored by The Na-
tional Center for Ecological Analysis and Synthesis.
B.J.E. was supported by NSF; K.J.N. was supported by
an Alexander von Humboldt Forschungspreis and
New York State Hatch grant funds.
19 September 2001; accepted 7 December 2001
Enzyme Dynamics During
Catalysis
Elan Zohar Eisenmesser,
1
Daryl A. Bosco,
1
Mikael Akke,
2
Dorothee Kern
1
*
Internal protein dynamics are intimately connected to enzymatic catalysis.
However, enzyme motions linked to substrate turnover remain largely un-
known. We have studied dynamics of an enzyme during catalysis at atomic
resolution using nuclear magnetic resonance relaxation methods. During cat-
alytic action of the enzyme cyclophilin A, we detect conformational fluctua-
tions of the active site that occur on a time scale of hundreds of microseconds.
The rates of conformational dynamics of the enzyme strongly correlate with
the microscopic rates of substrate turnover. The present results, together with
available structural data, allow a prediction of the reaction trajectory.
Although classical enzymology together with
structural biology have provided profound in-
sights into the chemical mechanisms of many
enzymes (1), enzyme dynamics and their rela-
tion to catalytic function remain poorly charac-
terized. Because many enzymatic reactions oc-
cur on time scales of micro- to milliseconds, it
is anticipated that the conformational dynamics
of the enzyme on these time scales might be
linked to its catalytic action (2). Classically,
enzyme reactions are studied by detecting sub-
strate turnover. Here, we examine enzyme ca-
talysis in a nonclassical way by characterizing
motions in the enzyme during substrate turn-
over. Dynamics of enzymes during catalysis
have previously been detected with methods
such as fluorescent resonance energy transfer,
atomic force microscopy, and stopped-flow flu-
orescence, which report on global motions of
the enzyme or dynamics of particular molecular
sites. In contrast, nuclear magnetic resonance
(NMR) spectroscopy enables investigations of
motions at many atomic sites simultaneously
(3, 4). Previous NMR studies reporting on the
time scales, amplitudes, and energetics of mo-
tions in proteins, have provided information on
the relation between protein mobility and func-
tion (5–15). Here, we have used NMR relax-
ation experiments to advance these efforts by
characterizing conformational exchange in an
enzyme, human cyclophilin A (CypA), during
catalysis.
CypA is a member of the highly con-
served family of cyclophilins that are found
in high concentrations in many tissues. Cy-
clophilins are peptidyl-prolyl cis/trans
isomerases that catalyze the interconversion
between cis and trans conformations of X-Pro
peptide bonds, where “X” denotes any amino
acid. CypA operates in numerous biological
processes (16, 17). It is the receptor for the
immunosuppressive drug cyclosporin A, is
essential for HIV infectivity, and accelerates
protein folding in vitro by catalyzing the
rate-limiting cis/trans isomerization of prolyl
peptide bonds (18, 19). However, its function
in vivo and its molecular mechanism are still
in dispute. X-ray structures of CypA in com-
plex with different peptide ligands show cis
X-Pro bonds (20, 21), except for a trans
conformation in the CypA/HIV-1 capsid
complex (22, 23). In each case, only one
conformer was observed in the crystal, even
though both isomers must bind to CypA for
catalysis of cis/trans isomerization to occur.
We characterized motions in CypA during
catalysis with the use of
15
N spin relaxation
experiments with and without the substrate
Suc-Ala-Phe-Pro-Phe-4-NA (24). Longitudinal
(R
1
) and transverse (R
2
) auto-relaxation rates,
transverse cross-correlated cross-relaxation
rates (
xy
), and {
1
H}-
15
N nuclear Overhauser
enhancements (NOE) were measured for all
backbone amides in CypA (25). Though all
parameters are sensitive to “fast” motions
( pico- to nanoseconds), only R
2
is sensitive to
“slow” conformational exchange (micro- to
milliseconds) (5–8). A progressive substrate-
induced shift for several CypA amide resonanc-
es (Fig. 1) indicates catalysis-linked motions. It
shows (i) that these amides experience different
magnetic environments in free CypA (E) and in
CypA bound to substrate (ES) and (ii) that the
1
Department of Biochemistry, Brandeis University,
Waltham, MA 02454, USA.
2
Department of Biophys-
ical Chemistry, Lund University, Post Office Box 124,
SE-221 00 Lund, Sweden.
*To whom correspondence should be addressed. E-
mail: dkern@brandeis.edu
Fig. 1. Chemical shift
changes of the amide
signals in CypA upon ti-
tration with the sub-
strate Suc-Ala-Phe-Pro-
Phe-4-NA. (A) At a con-
stant CypA concentra-
tion of 0.43 mM, spectra
were recorded at 0 mM
(blue), 0.38 mM (or-
ange), 1.01 mM (green),
and 2.86 mM (red) sub-
strate. The signal of R55 is progressively shifting upon addition of increasing amounts of substrate, indicating
fast conformational exchange during catalysis. The observed chemical shifts are population-weighted
averages of E and ES, and thus shift towards the position of the ES complex with increasing amounts of
substrate. In contrast, the signal of V139 is not affected by catalysis. (B) The chemical shift differences
between free CypA and in the presence of 2.86 mM substrate were mapped onto the structure (1RMH) (21)
with the use of a continuous color scale.
R EPORTS
22 FEBRUARY 2002 VOL 295 SCIENCE www.sciencemag.org1520
exchange rates between these states are faster
than the difference in their frequencies, i.e., the
time scale of exchange falls in the range of
10
⫺5
to 10
⫺3
s. In the presence of substrate, at
least three different states of CypA exist in
equilibrium (Scheme 1). Exchange between
these will increase the value of R
2
by an amount
R
ex
above that observed in free CypA, provided
that the nuclear spin experiences different
chemical shifts in at least two of the three states
(8). Direct measurements of R
2
revealed that
during catalysis CypA undergoes microsecond
conformational exchange in specific regions of
the protein. A significant increase in R
2
was
observed for 10 out of 160 backbone amide
nitrogens (26) (Figs. 2 and 3), which together
define a contiguous region in the structure (Fig.
3). The measured R
ex
for a particular amide
does not necessarily indicate motion of that
amide itself. R
ex
can be caused by slow time
scale motions of nearby atoms. Addition of
substrate induces only minor changes in pico-
to nanosecond dynamics, as evidenced by R
1
,
xy
, and {
1
H}-
15
N NOE (27). Thus, measure-
ment of R
2
identifies “hot spots” of micro- to
millisecond dynamics associated with either or
both of the processes involved in catalysis:
binding and isomerization.
Can the microscopic reaction steps be sep-
arated? The minimal reaction scheme (Scheme
1) consists of three microscopic reaction steps:
binding of (i) cis and (ii) trans isomers and (iii)
the catalytic step of substrate isomerization on
the enzyme. We separated the effects of binding
and isomerization by monitoring changes in R
2
as a function of substrate concentration. The
relative contributions to R
2
from exchange due
to binding and isomerization have different de-
pendencies on the total substrate concentration.
For most residues, R
2
first increases and then
decreases with the addition of substrate (Fig. 2,
D and E, and Fig. 4A). This pattern of maxi-
mum chemical exchange at intermediate sub-
strate concentrations—where E, ES
cis
, and ES-
trans
are all significantly populated—is diagnos-
tic of a predominant effect due to binding (see
Eq. 1). In contrast, R
2
increases monotonically
with substrate concentration for R55 (28) (Figs.
2B and 4B). This increase in R
2
with a concom-
itant increase in populations of ES
cis
and ES
trans
pinpoints a significant conformational ex-
change contribution to R
2
from interconversion
between ES
cis
and ES
trans
.
Do the exchange dynamics observed for the
enzyme correspond to the microscopic catalytic
steps of substrate turnover? To shed light on
this fundamental question, we determined rate
constants for the conformational changes on the
enzyme and compared them with rate constants
of substrate interconversion. For residues that
report only on binding, a simple two-state ex-
change model (including the free and a single
bound state) can be applied, enabling the use of
closed analytical formulae to determine the
binding constant, off-rate, and chemical shifts.
The exchange contribution (R
ex
)toR
2
may be
approximated as (29):
R
ex
⫽ P
E
P
ES
␦
2
k
ex
冉
1 ⫺
2
k
ex
cp
tanh
冋
k
ex
cp
2
册
冊
(1)
where P
E
and P
ES
are the fractional popula-
tions of the free (E) and bound (ES) states, ␦
is the chemical shift difference between E
and ES, k
ex
is the exchange rate, and
cp
is the
delay between refocusing pulses in the Carr-
Purcell-Meiboom-Gill (CPMG) experiment.
Expressing Eq. 1 in terms of the free substrate
concentration, [S
F
], and an effective dissoci-
ation constant, K
D
obs
⫽ P
E
[S
F
]/P
ES
⫽ k
off
/k
on
,
and making the substitution k
ex
⫽ k
off
⫹
k
on
[S
F
] ⫽ k
off
(1 ⫹ [S
F
]/K
D
obs
) ⫽ k
off
, one
gets
R
ex
⫽
␦
2
k
off
3
[S
F
]
K
D
obs
冉
1 ⫺
2
k
off
cp
tanh
冋
k
off
cp
2
册
冊
(2)
where [S
F
] is a function of K
D
obs
and the total
concentrations of substrate and enzyme, k
off
is
the off-rate, and k
on
is the on-rate (30). Hence,
K
D
obs
, k
off
, and ␦ can be estimated by nonlinear
regression (31). The data for residues K82, L98,
N102, and A103 are fit well by the two-state
model, yielding values of K
D
obs
between 0.95
and 1.20 mM, and values of k
off
between
10,700 and 14,800 s
-1
(Fig. 4A). These values
agree within uncertainties with those measured
from line shape analysis of the substrate (32).
Next, we obtained quantitative estimates of the
rate constants of protein dynamics by simulat-
ing the R
2
rates for the full three-state model of
Scheme 1. Excellent agreement between the
simulated and experimental data was obtained
with the use of the rate constants of cis/trans
isomerization determined separately from line
shape analysis (32) together with reasonable
chemical shift differences between the three
states (33) (Fig. 4). The results confirm the
qualitative evaluation of R
2
outlined above: for
Fig. 2. Comparison of
backbone
15
N R
2
relax-
ation between free CypA
and CypA during cataly-
sis. (A) R
2
rate constants
of CypA are shown for
samples with 0 mM (dark
blue), 1.01 mM (green),
and 2.86 mM substrate
(red). Four regions exhib-
iting changes in R
2
due to
steady-state turnover are
circled and are expanded
in (B to E), which include
also data for samples with
0.04 mM (light blue), 0.08
mM (magenta), and 0.45
mM substrate (gold).
Scheme 1. Three-state model of
CypA catalysis. E is the free en-
zyme, and ES
cis
and ES
trans
are the
two Michaelis-Menten complexes
with the substrate in the cis and
trans conformations, respectively.
K
D
is the dissociation constant, k
off
the off-rate, k
on
the on-rate, k
cat
ct
and k
cat
tc
are the rate constants of
isomerization. Superscripts cis and
trans identify the cis and trans iso-
mer, respectively. CypA catalyzes
the cis/trans isomerization of the
Phe-Pro peptide bond of the substrate used here with a turnover rate of several thousands per
second (43, 46).
R EPORTS
www.sciencemag.org SCIENCE VOL 295 22 FEBRUARY 2002 1521
most residues the binding processes dominate
the exchange contribution, whereas for others a
contribution from the isomerization step is re-
quired to yield satisfactory fits.
The backbone amide nitrogen of R55 clear-
ly experiences conformational exchange asso-
ciated with the isomerization and binding pro-
cess. Because the backbone amide group is
distant from the substrate binding site, it is
unlikely that the observed exchange merely
reflects a conformational change of the bound
peptide in a rigid active site. Rather, the results
provide evidence for conformational fluctua-
tions in the enzyme. Notably, R55 is essential
for catalysis (34). Its side-chain guanidino
group is hydrogen bonded to the prolyl nitrogen
of the substrate, and, thus, promotes isomeriza-
tion by weakening the double-bond character of
that peptide bond (20, 21, 23, 34). The good
agreement between the
15
N relaxation results
and those from substrate line enzyme shape
analysis implies that the observed conforma-
tional dynamics of the enzyme are strongly and
quantitatively correlated with the chemical
steps of substrate interconversion. Furthermore,
the conformational fluctuations of the enzyme
are likely to be collective motions, because
similar rate constants were inferred for several
backbone amides.
Interpretation of the results in light of struc-
tural data provides additional insights into
CypA catalysis. We focus here on residues that
show conformational exchange dynamics only
during catalysis, thus excluding the loop from
residues 68 –77 that also undergoes conforma-
tional exchange in resting CypA (35). Several
residues (101–103 and 109) with catalysis-
linked exchange are physically close to the
substrate bound in cis (Fig. 3A) (36 ). However,
L98 and S99 are distant from the peptide in the
cis conformation, yet undergo exchange during
the catalytic cycle and show chemical shift
changes upon titration with substrate that are
comparable to those observed for residues 101–
103 and 109 (Figs. 1 and 3). Though the struc-
ture of the enzyme in complex with the corre-
sponding trans isomer of the peptide is not
known, our results suggest that L98 and S99
may be interacting with the trans peptide. Thus,
we propose a reaction trajectory that involves a
rotation of the COOH-terminal peptide segment
around the prolyl peptide bond while the NH
2
-
terminal part remains bound to the enzyme
(37). This conformational rearrangement brings
all residues exhibiting exchange, except R55,
T68, and K82, into close proximity with the
peptide (Fig. 3B). K82 is located within a loop
remote from the active site (21). Our relaxation
data clearly indicate the involvement of K82 in
substrate binding, as confirmed by a reduction
in substrate affinity for the mutant Lys
82
3
Ala
82
(K82A) (38). As shown in Fig. 1B, ad-
ditional backbone amides show chemical shift
changes during turnover. However, these
changes are smaller than those observed for the
aforementioned residues; hence, their exchange
contribution to R
2
is below the present detection
limit (Eq. 1).
Taking all results together, we can envision
the enzymatic cycle of CypA as follows
(Scheme 1 and Fig. 3): the substrate exists in the
cis and trans conformations free in solution.
Both isomers can bind to CypA. For the cis
isomer, the areas around 101–103, 109, 82, and
55 are important in the binding process, which is
close to diffusion-controlled. After binding, the
enzyme catalyzes a rotation of the prolyl peptide
bond by 180°. For this to occur, the substrate tail
COOH-terminal to proline likely swings around
to contact the area around residues 98 and 99
(Fig. 3), whereas the NH
2
-terminal tail of the
substrate stays fixed. In other words, the enzyme
holds on to the substrate through this binding
interaction. The isomerization step takes place
with a rate constant of about 9000 s
-1
, and
motions of the protein coincide with the rate of
substrate rotation. The major player in catalysis
is R55, for which the observed changes in back-
bone conformation are likely to be coupled with
motions of the catalytically essential side chain.
After the isomerization step, the enzyme releas-
es the substrate, which is now in trans, with a
rate constant of about 13,000 s
-1
.
The approach outlined here allows the
identification of the dynamic “hot spots” dur-
ing catalysis and reveals that the time scales
of protein dynamics coincide with that of
substrate turnover. However, it does not pro-
vide a detailed physical picture of the mo-
tions during catalysis. To do this, side-chain
dynamics need to be included and, ultimately,
all the relaxation data need to be used in
molecular dynamics calculations. The appli-
cation of relaxation measurements during
substrate turnover promises to be of general
use in understanding the dynamic behavior of
enzymes and its relation to catalysis.
References and Notes
1. T. C. Bruice, S. J. Benkovic, Biochemistry 39, 6267
(2000).
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ence. A Guide to Enzyme Catalysis and Protein Folding
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Fig. 3. Residues in
CypA exhibiting mi-
crosecond time scale
dynamics during ca-
talysis. (A) Structure
of the cis conforma-
tion of the substrate
Suc-Ala-Phe-Pro-Phe-
4-NA (green) bound
to CypA, based on the
x-ray structure of
CypA complexed with
the cis form of Suc-
Ala-Ala-Pro-Phe-4-NA
(1RMH) (21). CypA
residues with chemical exchange in both the presence and absence of substrate are color coded in
blue (F67, N71, G74, S77, and S110). Residues in red exhibit chemical exchange only during
turnover (R55, K82, L98, S99, A101, N102, A103, and G109). Residues shown in magenta exhibit
chemical exchange in the absence of substrate, but increase in its presence ( T68 and G72). (B)
Suggested trajectory of the enzymatic pathway based on the dynamics results. CypA catalyzes
prolyl isomerization by rotating the part COOH-terminal to the prolyl peptide bond by 180° to
produce the trans conformation of the substrate. In this model, the observed exchange dynamics
for residues in strand 5 can be explained.
Fig. 4. Quantification of
exchange dynamics in
CypA during catalysis. R
2
rate constants are plotted
as a function of total sub-
strate concentration. (A)
R
2
data for K82. The con-
tinuous line indicates the
fitted Eq. 2, including con-
tributions only from bind-
ing. K
D
obs
⫽ 1.18 mM;
k
off
⫽ 11,100 s
⫺1
; ␦ ⫽
1450 s
⫺1
(3.8 ppm). (B) R
2
data for R55. The continuous line indicates a fit according to the full
three-state model, including contributions from both binding and isomerization (32); using K
D
obs
⫽
1.19 mM, then k
off
trans
⫽ 13,000 s
-1
; k
off
cis
⫽ 10,000; k
cat
ct
⫽ 9000s
-1
; k
cat
ct
⫽ 5100 s
-1
; ␦ ⫽ 440 s
⫺1
[1.2 parts per million (ppm)]; ␦
ct
⫽ 640 s
⫺1
(1.7 ppm). Of these parameters, only ␦
ct
and ␦
Et
were determined de novo, whereas for the others approximate values were known from the
two-site fitting and from line shape analysis.
R EPORTS
22 FEBRUARY 2002 VOL 295 SCIENCE www.sciencemag.org1522
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(2001).
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P. E. Wright, Biochemistry 40, 9846 (2001).
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Nature Struct. Biol. 8, 947 (2001).
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35, 3636 (1996).
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20. Y. Zhao, H. Ke, Biochemistry 35, 7362 (1996).
21.
㛬㛬㛬㛬 , Biochemistry 35, 7356 (1996).
22. T. R. Gamble et al., Cell 87, 1285 (1996).
23. Y. Zhao, Y. Chen, M. Schutkowski, G. Fischer, H. Ke,
Structure 5, 139 (1997).
24. The plasmid containing the gene for human CypA
was a generous gift from W. Sundquist. CypA was
expressed in BL21/DE3 cells in
15
N-labeled minimal
media. Cells were lysed in 25 mM MES, pH 6.1 and 5
mM -mercaptoethanol and were purified on a S-
Sepharose column equilibrated with the same buffer.
Remaining DNA was removed on a Q-Sepharose
column with 50 mM Tris, pH 7.8 and 5 mM -mer-
captoethanol. NMR sample conditions were 0.43 mM
CypA in 50 mM sodium phosphate buffer, pH 6.5, 3
mM dithiothreitol (DTT), 10% D
2
O, with peptide
concentrations between 0.04 and 2.6 mM. CypA re-
tained full activity during NMR data collection as
determined by the coupled chymotrypsin assay (39),
performed on samples before and after each series of
relaxation experiments.
25. Spectra were collected on Varian INOVA 600 spec-
trometers (Varian, Palo Alto, CA) at 25 ⫾ 0.1°C.
1
H-
15
N heteronuclear single-quantum correlation
(HSQC) spectra were acquired with the use of WA-
TERGATE (40).
15
N R
1
, R
2
,{
1
H}-
15
N NOE, and
xy
experiments were performed using pulse sequences
reported previously (41, 42). For each sample, at least
seven R
1
delays and eight R
2
delays were acquired,
ranging from 100 to 1000 ms and from 10 to 150 ms,
respectively. At least four relaxation delays were
acquired for the
xy
measurement, ranging from 30
to 120 ms. Spectra were acquired with 2048 and 128
complex points in the
1
H and
15
N dimensions, re-
spectively. Processing and analysis of the NMR spec-
tra were performed with the use of Felix (Accelrys,
San Diego, CA ). Uncertainties in peak heights were
estimated from duplicate spectra. Relaxation rates
were determined using the program CurveFit (A. G.
Palmer).
26. Additional residues show signs of exchange if the
CPMG pulse train is substituted for a single refocus-
ing pulse (12).
27. With addition of the peptide, several residues, includ-
ing residues N102 and A103, exhibit a decrease in R
1
,
whereas no significant change is observed for the
{
1
H}-
15
N NOE (see the supplementary material for
the complete R
1
and {
1
H}-
15
N NOE data, available on
Science Online at www.sciencemag.org/cgi/content/
full/295/5559/1520/DC1). Thus, for these residues,
pico- to nanosecond motions are more restricted in
the presence of peptide. A reduction in mobility on
this time scale has also been observed for residues
101–104 in the presence of the inhibitor cyclosporin
A(35).
28. Single-letter abbreviations for the amino acid resi-
dues are as follows: A, Ala; C, Cys; D, Asp; E, Glu; F,
Phe; G, Gly; H, His; I, Ile; K, Lys; L, Leu; M, Met; N, Asn;
P, Pro; Q, Gln; R, Arg; S, Ser; T, Thr; V, Val; W, Trp; and
Y, Tyr.
29. Z. Luz, S. Meiboom, J. Chem. Phys. 39, 366 (1963).
30. [S
F
] ⫽ (
公
(K
D
obs
⫺ [S
T
]⫹[E
T
])
2
⫹4[S
T
]K
D
obs
⫺ (K
D
obs
⫺ [S
T
] ⫹
[E
T
]))/2, where [S
T
] and [E
T
] are the total concentrations of
substrate and enzyme, respectively.
31. We first evaluated Eq. 2 in the limit k
ex
⬎⬎
cp
, which
amounts to fitting K
D
obs
and ␦
2
/k
off
. This yields a
good estimate of K
D
obs
(accurate to within approxi-
mately 10%), but a poorer estimate of ␦
2
/k
off
(ac-
curate to within approximately 25%). Knowledge of
K
D
obs
affords calculation of ␦ from the relation
obs
⫽ p
ES
␦⫹
E
, where
obs
and
E
are the
chemical shifts observed at a given concentration of
substrate and in free CypA, respectively. Note that
from the estimated K
D
obs
of 1.1 mM, p
ES
is calculated
to 0.67 for the highest substrate concentration used.
Subsequently, k
off
was estimated by fitting the full
Eq. 2 while keeping ␦ fixed. This approach circum-
vents problems that may otherwise arise when opti-
mizing the individual parameters k
off
and ␦, which
appear as a ratio in Eq. 2, simultaneously.
32. Line shape analysis was performed on the peptide in
the presence of catalytic amounts of CypA at 25°C
with the use of the method described (43) to deter-
mine the microscopic rate constants of catalysis.
33. The Bloch-McConnell equations (44) describing the
transverse relaxation in the full three-state system
(Scheme 1) were integrated numerically with the use
of Mathematica 4.0 ( Wolfram Research, Champaign,
IL), and monoexponential decays were fitted to the
resulting data sets, yielding transverse relaxation
rates for each residue at each concentration of sub-
strate. The three-state system is underdetermined by
the present number of experimental data points.
However, starting from parameter values determined
separately from line shape analysis of the substrate
during catalysis and from the two-site fitting, a sen-
sitivity analysis of the parameter space yields ranges
of possible values for the rate constants. The chem-
ical shift differences between the three states were
adjusted but kept within a reasonable range. The
chemical shift differences between the free and
bound states are related to the maximum chemical
shift change (␦) observed upon substrate addition
and were determined separately by: ␦⫽␦
ct
/(1 ⫹
K
eq
)–␦
Et
. K
eq
⫽ [ES
cis
]/[ES
trans
] is obtained from
line shape analysis, ␦
ct
is the chemical shift differ-
ence between ES
cis
and ES
trans
, and ␦
Et
is the chem-
ical shift difference between E and ES
trans
.
34. L. D. Zydowsky et al., Protein Sci. 1, 1092 (1992).
35. M. Ottiger, O. Zerbe, P. Guntert, K. Wuthrich, J. Mol.
Biol. 272, 64 (1997).
36. An initial structural model of CypA in complex with
Suc-Ala-Phe-Pro-Phe-4-NA was built from the crystal
structure of CypA in complex with the similar peptide
Suc-Ala-Ala-Pro-Phe-4-NA in the cis conformation
(PDB entry 1RMH) (21). All structures were viewed
and built using MOLMOL (45).
37. Chemical shift changes for this substrate upon bind-
ing to cyclophilin have previously been calculated by
line shape analysis, and these data also support the
proposed model for the conformational change (43).
38. D. Kern, unpublished data.
39. G. Fischer, H. Bang, C. Mech, Biomed. Biochim. Acta
43, 1101 (1984).
40. M. Piotto, V. Saudek, V. Sklenar, J. Biomol. Nucl.
Magn. Reson. 2, 661 (1992).
41. N. A. Farrow et al., Biochemistry 33, 5984 (1994).
42. N. Tjandra, A. Szabo, A. Bax, J. Am. Chem. Soc. 118,
6986 (1996).
43. D. Kern, G. Kern, G. Scherer, G. Fischer, T. Drakenberg,
Biochemistry 34, 13594 (1995).
44. H. M. McConnell, J. Chem. Phys. 28, 430 (1958).
45. R. Koradi, M. Billeter, K. Wuthrich, J. Mol. Graph. 14,
51 (1996).
46. The microscopic rate constants of substrate intercon-
version were determined separately from line shape
analysis of the peptide NMR spectrum using previ-
ously established methods (43).
47. We are grateful to C. Miller for assistance with the
quantitative analysis. Supported by NIH grant
GM62117 (D.K.) and VR grants K-650-19981661 and
S-614-989 (M.A.). Instrumentation grants were
awarded by the NSF and the Keck foundation to
(D.K.) and by the Knut and Alice Wallenberg founda-
tion (M.A.).
12 September 2001; accepted 11 January 2002
Role of Nucleoporin Induction
in Releasing an mRNA Nuclear
Export Block
Jost Enninga,
1
David E. Levy,
2
Gu¨nter Blobel,
1
*
Beatriz M. A. Fontoura
1,3
Signal-mediated nuclear import and export proceed through the nuclear pore
complex (NPC). Some NPC components, such as the nucleoporins (Nups) Nup98
and Nup96, are also associated with the nuclear interior. Nup98 is a target of
the vesicular stomatitis virus ( VSV ) matrix (M) protein–mediated inhibition of
messenger RNA (mRNA) nuclear export. Here, Nup98 and Nup96 were found
to be up-regulated by interferon (IFN). M protein–mediated inhibition of mRNA
nuclear export was reversed when cells were treated with IFN-␥ or transfected
with a complementary DNA (cDNA) encoding Nup98 and Nup96. Thus, in-
creased Nup98 and Nup96 expression constitutes an IFN-mediated mechanism
that reverses M protein–mediated inhibition of gene expression.
The Nup98 and Nup96 proteins are encoded
by a single gene. The primary transcript is
alternatively spliced, and the translation
products are autocatalytically proteolyzed at
one specific site (1–3). Nup98 interacts with
an intranuclear protein (4 ) and transport fac-
tors (5, 6). It is involved in nuclear import
and export of proteins and RNAs (7–11) and
is the target of the VSV M protein–mediated
inhibition of mRNA export (12). A cDNA
clone coding for part of the COOH-terminal
sequence of Nup96 has been detected among
mRNAs that were specifically induced by
IFN-␥ (13).
We found two classical elements, GAS
and ISRE, that mediate increased gene ex-
pression by IFN (14, 15). When U937 cells
were incubated with IFN-␥ for up to 12
R EPORTS
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