From the Department of Biochemistry, Stanford University School of Medicine, Stanford, California 94305
Received for publication, December 4, 2000, and in revised form, February 20, 2001
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ABSTRACT |
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Crystal structures of the myosin motor
domain in the presence of different nucleotides show the lever arm
domain in two basic angular states, postulated to represent prestroke
and poststroke states, respectively (Rayment, I. (1996) J. Biol. Chem. 271, 15850-15853; Dominguez, R., Freyzon, Y.,
Trybus, K. M., and Cohen, C. (1998) Cell 94, 559-571). Contact is maintained between two domains, the relay and the
converter, in both of these angular states. Therefore it has been
proposed by Dominguez et al. (cited above) that this
contact is critical for mechanically driving the angular change of the
lever arm domain. However, structural information is lacking on whether
this contact is maintained throughout the actin-activated myosin ATPase
cycle. To test the functional importance of this interdomain contact,
we introduced cysteines into the sequence of a "cysteine-light"
myosin motor at position 499 on the lower cleft and position 738 on the
converter domain (Shih, W. M., Gryczynski, Z., Lakowicz, J. L., and Spudich, J. A. (2000) Cell 102, 683-694).
Disulfide cross-linking could be induced. The cross-link had minimal
effects on actin binding, ATP-induced actin release, and
actin-activated ATPase. These results demonstrate that the
relay/converter interface remains intact in the actin strongly
bound state of myosin and throughout the entire actin-activated myosin
ATPase cycle.
Myosins are molecular motors that transduce the chemical energy of
ATP hydrolysis to mechanical work in the form of the vectorial translocation of substrate actin filaments. In the swinging lever arm
model of actin-myosin motor action, myosin binds to the actin with its
globular catalytic domain and then rotates its carboxyl-terminal lever
arm domain (reviewed in Ref. 5). The anchoring of the end of the lever
arm domain results in the translocation of the catalytic domain and the
attached actin filament.
Crystal structures of the motor domain of myosin show the lever arm
domain in two basic angular classes, which have been postulated to
represent prestroke and poststroke states (2, 3, 6). Scallop myosin
complexed with ADP has been crystallized in a third angular state,
proposed as a myosin·ATP actin-detached state (7). Recent dynamic
studies of the lever arm position using steady state and time-resolved
fluorescence energy transfer measurements support a swing of the lever
arm from a prestroke state to a poststroke state through an angle of
more than 70 degrees (4). The lever arm domain consists of a
disc-shaped "converter domain" from which a long We have used a cysteine engineering approach to address the question as
to whether the relay/converter contact is maintained in the
actin strongly bound state. We constructed myosinII alleles containing
cysteine-light mutations (C49S, C312Y, C442S, C470I, C599L, and C678Y)
(4) and substituted cysteine codons into positions corresponding to
either residue 499 or 738 or both (Fig. 1). We show that the mutant
myosins are functional in vivo and in vitro. The
cross-linked myosin (containing cysteines at both positions 499 and
738) retained the ability to bind to actin in the absence of ATP as
well as the ability to be released from actin in the presence of ATP.
The cross-linked myosin also retained actin-activated ATPase activity.
Nomenclature--
The changes made in a given mutant myosin are
described within parentheses. For example, myosin(CL, I499C) refers to
a full-length myosin gene with the cysteine-light
(CL)1 mutations (C49S, C312Y,
C442S, C470I, C599L, and C678Y) (4) and the mutation I499C.
Mutagenesis and Subcloning--
Subcloning procedures were
carried out using standard protocols (8). myosin(CL, I499C), myosin(CL,
R738C), and myosin(CL, I499C, R738C) were generated by splice overlap
extension mutagenesis (9) using myosin(CL) as a template. Myosin genes
were subcloned into the expression vector pTIKL-Myo. The introduced
mutations were verified by dideoxy-DNA sequencing. The S1 gene
fragments were then subcloned into pTIKLOES1, an expression vector for
producing S1 with a carboxyl-terminal His6 tag on
the heavy chain.
The following oligonucleotides were used for mutagenesis: I499C-F,
5'-TATCTTAAAGAGAAATGTAATTGGACTTTCATC-3'; I499C-R,
5'-GATGAAAGTCCAATTACATTTCTCTTTAAGATA-3'; R738C-F,
5'-GATCCAGAACAATATTGTTTCGGTATCACCAAG-3'; and R738C-R, 5'-CTTGGTGATACCGAAACAATATTGTTCTGGATC-3'.
Transformation into Dictyostelium
Cells--
Dictyostelium cells were grown in HL-5 medium as
described previously (10). Cells were grown at 22 °C in HL-5
supplemented with 17% FM medium (Life Technologies, Inc.), 100 units/ml penicillin, and 100 units/ml streptomycin. Transformations
were performed as described previously (11). The mhcA null
cell line HS1 was transformed with 10 µg of each of the pTIKL-Myo
plasmids bearing wild type or mutant versions of the full-length
myosin, whereas the AX3-ORF+ cell line was transformed with 10 µg of
each of the pTIKLOES1 plasmids bearing wild type or mutant versions of
the S1 fragment of myosin. Clonal cell lines that grew in the presence of HL-5 supplemented with penicillin, streptomycin, and 8 µg/ml G418
were isolated, and these cell lines were further characterized.
Growth in Suspension Assay--
Cells were grown on plates to
near confluence before they were transferred to shaking flasks. Cells
were diluted to 4 × 104 cells/ml in 25 ml of total
volume HL-5 in 125-ml Erlenmeyer flasks and shaken at 200 rpm at
22 °C for 6 days. A small aliquot was removed at regular intervals,
and the number of cells was counted using a hemocytometer.
Protein Purification--
Dictyostelium S1
His6 was expressed in Dictyostelium
AX3-ORF+ cells (grown in suspension) and purified as described
(12).
Actin-activated ATPase Assay--
S1 ATPase activities were
measured as the release of labeled Pi using
Cross-linking Assay--
Cross-linking was induced by the
addition of 25 µM 5,5'-dithiobis(nitrobenzoic
acid) to S1 at a concentration of 1-2 µM. The buffer
conditions were 25 mM HEPES, pH 7.0, 25 mM
NaCl, and 10 mM MgCl2. The cross-linking
reaction was quenched by the addition of 1 mM
dithiothreitol. (The Cys499-Cys738 disulfide
cross-link is not reduced by 1 mM dithiothreitol under nondenaturing conditions.) Disulfide cross-linking was assayed by mobility shift in SDS-PAGE behavior in the absence of reducing agent. Confirmation of disulfide bond formation was made by SDS-PAGE analysis in the presence of reducing agent reversing the mobility shift.
Actin Cosedimentation Assay--
The buffer conditions used were
25 mM HEPES, pH 7.0, 25 mM NaCl, and 10 mM MgCl2. For the noncompetitive assay, S1 (0.8 µM final concentration) was mixed with F-actin (3.0 µM final concentration) for 10 min, and then the mixture
was centrifuged at 100,000 × g for 10 min. The
supernatant and the resuspended pellet were examined by SDS-PAGE to
determine whether the S1 cosedimented with the F-actin. The same assay
was repeated in the presence of 2 mM Mg-ATP. For the
competitive assay, the same procedure was used but with S1 at a final
concentration of 1.3 µM and F-actin at a concentration of
0.5 µM.
The Cysteine Mutant MyosinII Is Functional in
Vivo--
Dictyostelium cells that lack the myosinII gene
are unable to divide in suspension, instead becoming large and
multinucleate before eventually lysing and dying (13, 14). Thus the
transformation of the mutant myosinII gene into these myosinII null
cells can lead to a simple assay for in vivo function
(assaying for the rescue of the growth in suspension defect). The
ability to rescue an in vivo defect serves as a useful
benchmark to demonstrate that mutant myosins behave in a functional
manner. The design of this experiment relies on the un-cross-linked
double-cysteine mutant myosin to behave in a functional manner and then
to assay for biochemical differences upon specific cross-linking.
Arg738 (with its
Fig. 2 shows a growth curve examining the
growth of Dictyostelium cells that were missing the genomic
copy of myosinII but were supplied with another copy on an
extrachromosomal plasmid. All of the mutant myosinII genes introduced
(myosin(CL), myosin(CL, I499C), myosin(CL, R738C), and
myosin(CL, I499C, R738C)) rescued growth in suspension to a rate
comparable with that of the wild type. The parent strain lacking a copy
of myosinII, however, failed to grow in suspension. Therefore it
appears that the introduction of either cysteine, both individually or
in tandem, is well tolerated by the structure of myosin.
A Specific Cross-link Is Inducible between Introduced Cysteines at
Positions 499 and 738--
According to the crystal structures
available, the side chains of residues at positions 499 and 738 are in
close proximity (2, 15). Previous studies have shown that cysteines
placed at nearby positions in a structure usually can be
induced to form a disulfide cross-link, which is catalyzed either by
ambient oxygen or by a disulfide exchange reagent (16). The
cross-linking of two residues in the structure of a protein that are
separated by a large number of residues results in a covalently closed
large loop within the primary sequence. Denatured proteins containing such a loop might be expected to exhibit a different mobility during
gel electrophoresis; examples have been found where cross-linking induces either a gel mobility increase (17) or a gel mobility decrease
(18). Fig. 3 shows that treatment with
the disulfide exchange reagent dithionitrobenzoate induces an SDS-PAGE
mobility increase in over 85% of a cysteine-light myosin S1
only when cysteines are introduced at both positions 499 and 738 but
not when the cysteines are introduced individually. This gel mobility
shift is reversible if the protein is loaded onto the gel in the
presence of a reducing agent such as
Curiously, the disulfide cross-link that is formed is quite robust
against reduction under native conditions. Overnight treatment in the
presence of 1 mM dithiothreitol, 1 mM
Strong Actin Binding (in the Absence of ATP) and ATP
Disruption of This Strong Binding State Are Not Perturbed by
Cross-linking the Introduced Cysteines--
Wild type S1 precipitates
with F-actin in the absence of ATP during centrifugation at sufficient
speeds. Small, unattached proteins sediment much more slowly and
therefore remain in the supernatant. This assay can be used to
investigate the binding of S1 to actin. The property of the
cross-linked form of S1 (that it migrates more rapidly during SDS-PAGE,
relative to the un-cross-linked form) lends itself as a special
advantage in the F-actin cosedimentation assay because this assay
allows for the analysis of the supernatant (actin-detached) and pellet
(actin-attached) fractions using SDS-PAGE. Thus the behavior of the two
forms can be analyzed simultaneously in the same sample volume.
Fig. 4A shows S1 binding to
F-actin in the presence of excess F-actin and in the absence of
ATP. Both the cross-linked and un-cross-linked forms of S1 cosediment
with the F-actin. Fig. 4A also shows S1 binding to F-actin
in the presence of excess F-actin and ATP. Both the cross-linked and
un-cross-linked forms of S1 are similarly released from the actin by
the ATP. Fig. 4B shows S1 binding to F-actin in the presence
of limiting F-actin. If the cross-linked form of S1 had a significantly
different binding affinity to F-actin as compared with the
un-cross-linked form of S1, then the ratio of the un-cross-linked to
cross-linked bands should be significantly different in the supernatant
and pellet lanes. However, Fig. 4B shows that no significant
difference is evident. Thus the cross-link does not perturb the binding
of myosin to F-actin in its strong affinity mode.
Cross-linking the Introduced Cysteines Results in a Modest Decrease
in the Actin Activation of the ATPase of Myosin--
The rate-limiting
step in the ATPase cycle of myosin in the absence of F-actin is the
product release (19). The presence of F-actin stimulates the myosin to
release its products more rapidly (20). We examined the actin-activated
ATPase activity of the cross-linked form versus the
un-cross-linked form to see if the actin stimulation of product release
might be impaired. Fig. 5 shows that the
actin activation of the cross-linked species is decreased by 29% and
that the Km for the actin activation is decreased by
29%. The basal rate of ATP hydrolysis (i.e. in the absence
of F-actin) is the same in the cross-linked and un-cross-linked forms.
Thus phosphate release in the cross-linked myosin is still stimulated
by F-actin, although the amount of the stimulation is slightly
decreased.
The crystal structures of the Dictyostelium myosinII
catalytic domain have been solved in complex with a number of
nucleotide analogues, including ADP beryllium fluoride
(BeFx), ADP aluminum fluoride
(AlF The lever arm extends off from a 60-residue globular domain, often
referred to as the converter domain. This converter domain shares in
both structures a hydrophobic interface with the back end of the lower
jaw of the large cleft of the catalytic domain, referred to as the
relay helix, despite a 70° rotation of the converter domain between
the two structures. Fig. 6 shows this interface in the two structures, as well as in an intermediate structure that was generated as an interpolation between the crystal structure models (4). Inspection of the crystal structures suggests a
mechanical picture describing the sequence of forces that are behind
the power stroke. It has been proposed that changes in the nucleotide
binding site, triggered by the formation of a bond between the backbone
amide of switch II residue Gly457 with the
INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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-helix,
bound by two calmodulin-like light chains, emerges. In both crystal
states, the converter domain maintains a contact on a face of its
radial edge to a rigid helix extending from the lower domain of the
large cleft, referred to as the relay helix (3). This helix has to
undergo a small conformational change, primarily a rigid body
translation and rotation, to accommodate the angular rotation of the
converter domain. Because this domain-domain interface is maintained
between the two angular states, it has been proposed that this
interface is important for mechanically driving the angular change.
However, structural information is lacking as to whether this contact
is maintained throughout the actin-activated myosin ATPase cycle,
including in an actin strong binding state.
MATERIALS AND METHODS
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MATERIALS AND METHODS
RESULTS
DISCUSSION
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-32P, as reported previously (12). The plotted
points and error bars of Fig. 5 represent measurements
from three independent trials for each of two different protein preparations.
RESULTS
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and
carbons) and
Ile499 form part of the hydrophobic interface between the
relay helix and the converter domain, respectively (Fig.
1). Disruption of this hydrophobic
interface thus could potentially be destabilizing for the myosin.
Cysteines are relatively hydrophobic, however, and should be good
candidates for replacement side chains for packing in the
interface.
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Fig. 1.
Introduction of two cysteines into the
structure of Dictyostelium myosinII. The
structure of the Dictyostelium myosinII catalytic domain
complexed with ADP·BeFx is depicted (1mmd in the
PDB data base). The lower jaw of the large cleft is colored
red, and the converter domain is colored yellow.
The SH1-SH2 helix is colored green. The actin binding face
is on the right, and the lever arm would extend from the
structure on the lower left. Ile499 from the
relay helix and Arg738 from the converter domain are
rendered in the Corey-Pauling-Koltun space filling model. These two
positions were mutagenized to cysteines for this study.
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Fig. 2.
Dictyostelium growth in
suspension. Plus signs, wild type myosin; open
circles, myosin(CL); dark diamonds, myosin(CL, I499C);
dark boxes, myosin(CL, R738C); tetrasected boxes,
myosin(CL, I499C, R738C); and open triangles, myosinII null.
All myosinII alleles tested in this study rescued the
growth-in-suspension defect exhibited by the myosinII null
Dictyostelium cells.
-mercaptoethanol. Thus a
specific cross-link is formed between introduced cysteines at positions 499 and 738.
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Fig. 3.
Effect of disulfide cross-linking on the
electrophoretic mobility of cysteine-engineered myosin motor
domains. Purified S1 proteins were treated with 25 µM 5,5'-dithiobis(nitrobenzoic acid) and then analyzed by
SDS-PAGE. Only the S1 with cysteines at both positions 499 and 738 exhibited increased mobility upon treatment with the cross-linking
reagent. This mobility shift was reversed when the S1 was
electrophoresed in the presence of the reducing agent
-mercaptoethanol. Cyslite, cysteine light.
-mercaptoethanol, or 1 mM tricarboxyethylphosphine
followed by passage through a Sephadex spin column (to remove the
dithiothreitol) fails to significantly reduce the disulfide, as
analyzed by SDS-PAGE (data not shown). Only after SDS denaturation does
the disulfide become accessible to reduction. In the absence of the
disulfide exchange reagent and reducing agents, the un-cross-linked
protein is converted slowly to the cross-linked form over a period of
weeks when stored at 4 °C, presumably catalyzed by ambient dissolved
oxygen (data not shown).
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Fig. 4.
Actin cosedimentation assays. The
SDS-PAGE lanes containing supernatant samples are labeled
Sup, whereas the gel lanes containing pellet samples are
labeled Pel. A, both the cross-linked and
un-cross-linked forms of S1(CL, I499C, R738C) bind to F-actin and are
released by ATP, in the presence of excess F-actin. B, the
cross-linked and un-cross-linked forms of the S1(CL, I499C, R738C) have
a similar affinity to F-actin, as analyzed by actin cosedimentation
using substoichiometric amounts of F-actin.
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Fig. 5.
Actin-activated ATPase assays. The
cross-linked and un-cross-linked forms of the S1(CL, I499C, R738C)
exhibit similar basal ATPase rates (0.24 s 1). The
cross-linked S1 exhibits an actin-activated ATPase
(Vmax = 0.82 s
1) that is 30% less
than the activity of the un-cross-linked S1
(Vmax = 1.06 s
1). The cross-linked
S1 exhibits a Km for the actin activation
(Km = 4.6 µM) about 30% lower than
that shown by the un-cross-linked S1 (Km = 6.5 µM).
DISCUSSION
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-phosphate in
the prestroke state, are amplified into a larger change in the position
of the relay helix. This change drives a rotation and translation of
the lever arm through its converter domain (3, 15).
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Fig. 6.
Conformational changes in the relay helix to
converter domain interface. Shown here are three successive
frames, going from left to right,
depicting the myosin S1 conformational change in transforming from the
putative poststroke angle state (based on 1mmd in the PDB data
base) to the putative prestroke angle state (based on 1vom in the PDB
data base). The second frame was generated as described (4).
The relay helix (colored red), which forms an interface with
the converter domain (colored yellow), undergoes a rotation
and translation. This change drives the attached converter domain to
rotate and translate. This results in a large movement upward of the
lever arm -helix (attached to the converter domain and also colored
yellow). The action is reminiscent of a hand turning a
steering wheel. Residues 499 and 738 are colored in
cyan.
These structures, however, are thought to represent myosin in conformations that have a weak affinity to actin (21). Myosin undergoes a conformational change that allows it to bind to actin with a much higher affinity; this conformational change in myosin can be monitored by the pyrene labeling of Cys374 on actin (22) or with mant-ADP in the active site of the myosin (23). These spectroscopic probes, however, give no structural details of the nature of this conformational change. Therefore currently there is no experimental evidence that allows one to build a reliable model of myosin in its strong affinity to actin state.
One proposal has been that the large cleft completely closes to achieve this strong affinity to actin state (1, 6). One possible extension of this model is that the complete closure of the large cleft causes the relay helix to slip away from the converter domain. The converter domain, after its release from the relay helix, snaps back to its poststroke conformation, thus completing the power stroke. In this model, the lever arm behaves as a torque spring that is wound by 70° by the back end of the lower jaw and then suddenly released once actin strong binding has been achieved.
Another model is that the interface between the converter domain and the relay helix is maintained in the strong actin affinity state of myosin as well. In this case, the relay helix may mechanically transmit the changes to the lever arm both in the actin-detached stages (the recovery stroke) and in the actin-attached stages (the power stroke).
If the first model were correct (that the interface must slip), then the cross-linking of the interface would prevent that slipping and should prevent the myosin from achieving its actin strong binding affinity state. On the other hand, if the second model were correct (that the interface is maintained during the whole cycle), then the cross-linking of the interface should have little or no effect on the transition to the strong affinity to actin state.
Our experiments demonstrate that cross-linking these two domains
together through a cysteine at position 499 on the relay helix and a
cysteine at position 738 on the converter domain does not inhibit
myosin from achieving a strong affinity to actin state nor does it
inhibit the effect of ATP in shifting the myosin back to the weak
affinity to actin state. Thus our results are consistent with the model
that the interface is maintained both in the actin-detached and
actin-attached stages of the actomyosin ATPase cycle.
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ACKNOWLEDGEMENTS |
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We thank András Málnási-Csizmadia and Clive Bagshaw for an advance copy of their manuscript and discussions. We thank Wen Liang for assistance with generating the growth curves, and we thank Doug Robinson for helpful discussions.
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FOOTNOTES |
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* This work was supported by Grant AR42895 from the National Institutes of Health (to J. A. S.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Supported by a Howard Hughes Predoctoral Fellowship.
§ To whom correspondence should be addressed: Dept. of Biochemistry, Beckman Center B400, Stanford University School of Medicine, Stanford, CA 94305-5307. Tel.: 650-723-7634; Fax: 650-723-6783; E-mail: jspudich@cmgm.stanford.edu.
Published, JBC Papers in Press, February 21, 2001, DOI 10.1074/jbc.M010887200
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ABBREVIATIONS |
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The abbreviations used are: CL, cysteine-light; PAGE, polyacrylamide gel electrophoresis.
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