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Cell Motility and the Cytoskeleton 36:246–252 (1997)
Physical Properties of Dystrophin
Rod Domain
Edith Kahana,1* G. Flood,2 and W.B. Gratzer1
1Medical
Research Council Muscle and Cell Motility Unit, King’s College, London
2National Centre for Macromolecular Hydrodynamics,
University of Leicester, England
We have prepared two fragments of the human dystrophin rod domain, each
containing eight spectrin-like repeating units, by expression in Escherichia coli.
The first corresponds to the central portion of the rod, the other to three repeats
from the N-terminal end, fused to five repeats from the C-terminal end. The latter
makes up the entire mutant rod, found in a patient with mild (Becker-type)
muscular dystrophy. Both fragments were found to possess an ordered, stable
structure, and had the form of short rod-like particles in the electron microscope.
Molecular weight determinations by sedimentation equilibrium revealed that both
polypeptides were monomeric in solution, suggesting that the dystrophin rod
domain is incapable of forming an antiparallel homodimer. This supports the
inference from sequence analyses [Winder et al., 1995: FEBS Lett. 369:27–33,
1996: Biochem. Soc. Trans. 24:2805] that the dystrophin rod domain lacks the
arrangement of sites required for lateral self-association, and that dystrophin,
unlike the other known proteins of the spectrin superfamily, may thus exist as a
monomer. Cell Motil. Cytoskeleton 36:246–252, 1997. r 1997 Wiley-Liss, Inc.
Key words: dystrophin; spectrin repeats
INTRODUCTION
The manner in which dystrophin functions in stabilising the sarcolemma is unclear. It serves as a link
between the transmembrane glycoprotein complex, which
in turn connects the membrane to the extracellular matrix,
and the cortical actin cytoskeleton [Ahn and Kunkel
1993]. The actin-binding site is in a globular N-terminal
domain and the glycoprotein-binding site is in the Cterminal domain. The latter is preceded in the sequence
by a cysteine-rich domain with unknown, but apparently
essential function, and between this and the actin-binding
domain is a rod domain, comprising 24 degenerate
spectrin-like repeating units. Dystrophin is mainly located in specialised zones of the sarcolemma, the costameres, surrounding the Z-discs of the myofibrils [Minetti
et al., 1992; Porter et al., 1992; Straub et al., 1992]. These
probably represent the sites of mechanical coupling
between the sarcolemma and the contractile system. The
dystrophin is presumed to protect the membrane against
the shearing stresses to which it is subjected, in much the
same way as the spectrin lattice stabilises the red cell
r 1997 Wiley-Liss, Inc.
membrane. Spectrin is responsible for the elasticity of the
membrane with which it associates, and the dystrophin
rod domain, with its spectrin-related sequence [Koenig
and Kunkel, 1990] and conformation [Kahana et al.,
1994], must be supposed to exert a similar effect.
Whereas total absence of dystrophin leads to severe
(Duchenne) muscular dystrophy (DMD), many types of
mutation results in much milder or intermediate forms of
the disease, known as Becker muscular dystrophy (BMD).
Thus, whereas deletions of the C-terminal or cysteinerich domains invariably cause DMD, loss of the actinbinding domain or of a large proportion of the rod are
associated with BMD [Ahn and Kunkel, 1993; Ohlendieck, 1996]. It appears therefore that the membrane can
survive and the muscle continue to function, even if in
*Correspondence to: E. Kahana, Department of Hematology, Tel Aviv
Sourasky Medical Center and Sackler Faculty of Medicine, 6 Weizmann Street, Tel-Aviv 64239, Israel.
Received 2 August 1996; accepted 11 October 1996.
Dystrophin Rod Domain
impaired form, despite a presumed loss in membrane
elasticity.
Little is known about the organisation of the
dystrophin-containing complex on the membrane, but
most published models have explicitly or implicitly
assumed that the dystrophin rod forms an antiparallel
dimer, like all other known proteins of the spectrin class.
This would allow a continuous structure to be generated
at the membrane surface. The evidence for dimer formation is equivocal, however [Pons et al., 1990], and a
recent analysis of the rod domain sequence concludes that
dimerisation, as it occurs in the spectrins and a-actinin, is
unlikely [Winder et al., 1995, 1996]. We have attempted
here to address this problem by examining the properties
of large fragments of the rod domain and also to analyse
the conformational characteristics of the rod, about which
little was previously known.
MATERIALS AND METHODS
Dystrophin rod fragments were prepared by expression in Escherichia coli. We selected a segment from the
centre of the rod, comprising repeats 8–15, and the
shortened rod domain from a patient with BMD [England
et al., 1990]. In the latter, repeats 4 to 18 and half of repeat
19 are deleted, leaving a fusion of the first three and the
last five repeats of the rod. The cDNA for both protein
sequences was kindly provided by Professor K.E. Davies.
We term the products RFM (rod fragment, middle) and
RFE (rod fragment, extremities).
RFM begins at met-1141 in the dystrophin sequence and ends with gln-1973. The following primers
were synthesised: sense-strand nt 3629 to 3659, with an
added phased NdeI restriction site; antisense-strand nt
26127 to 26107, with an added stop codon and a HindIII
restriction site. The restriction enzymes did not cut the
cloned fragment. DNA fragments were prepared by 15
cycles of PCR, using thermostable Vent DNA polymerase. The cloning vector was pMW172 [Way et al., 1990].
The dephosphorylated vector and the PCR products were
purified by gel electrophoresis. The fragment was ligated
into the cloning vector and transformed into a nonexpressing (TG1) strain of E. coli and recombinants were
selected by PCR, using the T7 vector sense and antisense
primers. The DNA was introduced into the expression
strain (BL21/DE3) and the bacteria were grown to late
log phase at 37°C in LB medium, containing 0.1 mg ml21
ampicillin. Sequences from the 38 and 58 ends of the
recombinant DNA were determined to confirm its identity.
RFE begins at asp-329 and ends at lys-3117,
residues 665 to 2367 being deleted. Expression was based
on DNA from the BMD patient. A vector, pT79, modified
from pMW172 by Dr P.J. Marsh to embody additional
247
restriction sites, was employed. A sense primer, corresponding to nt 1187 to 1216, with an added NdeI
restriction site, was synthesised, as well as an anti-sense
primer, corresponding to nt 29559 to 29539, with an
added stop codon and a NotI restriction site. Cloning and
expression were performed as above.
Both RFE and RFM were found in inclusion bodies,
as well as in the soluble fraction of the bacteria. The
inclusion bodies were collected, dispersed in 6 M guanidinium chloride, 5 mM dithiothreitol, and the protein
was recovered by chromatography in the same solvent on
a Sephacryl S-300 (Pharmacia, Piscataway, NJ) column
of 2.5 3 100 cm. The product at this stage was
contaminated with nucleotide material. This was eliminated by precipitating the protein with an equal volume of
saturated ammonium sulphate. It was redissolved and
dialysed against 50 mM sodium chloride, 20 mM sodium
phosphate, pH 7.6, and rechromatographed on Superose
12 in a Pharmacia FPLC system. Fractions were screened
for purity by gel electrophoresis in SDS. N-terminal
sequencing was carried out on the products to establish
their identities. (We are indebted for these determinations
to A.C.M. Harris, National Institute for Medical Research, London.) Concentrations were measured spectrophotometrically, using calculated specific absorptivities
[Perkins, 1986].
For chemical cross-linking, proteins at a concentration of 60 to 170 µg ml21 in 0.1 M triethanolamine, pH
8.5, on ice were treated with 1 mg ml21 dimethylsuberimidate [Davis and Stark, 1970] and examined by SDS-gel
electrophoresis in 5% acrylamide gels. Red cell spectrin
was used as a control.
Molecular weight determinations by sedimentation
equilibrium were performed in a Beckman (Palo Alto,
CA) Optima XL-A analytical ultracentrifuge, using absorption optics at 280, 234, or 220 nm on solutions in 0.15
M sodium chloride, 50 mM sodium phosphate, pH 7.6.
Sedimentation velocities were measured in the same
instrument at 42,000 rpm. Partial specific volumes were
calculated from the amino acid compositions [Perkins,
1986].
For electron microscopy (performed by Dr. A.P.R.
Brain, King’s College London Electron Microscopy Unit)
solutions of protein at 50–200 µg ml21 in 50 mM sodium
chloride, 5 mM imidazole, pH 7.6, containing 60% (v/v)
glycerol, were sprayed onto mica, shadowed at an angle
of 6° with platinum, and coated with carbon. The replicas
were transferred to grids and examined in a Philips
(Mahwah, NJ) 301G instrument.
Circular dichroism spectra were measured in a
Jobin-Yvon CD6 Dicrograph (Longjumeau, France), using cells of 0.5–2 mm path-length in a thermostated
housing. For thermal denaturation profiles the temperature was varied in steps of 5°C.
248
Kahana et al.
RESULTS
The purified expressed dystrophin rod segments,
RFM and RFE, were at least 95% pure, as judged by gel
electrophoresis in SDS. The apparent molecular weights
were about 97,000 and 125,000, respectively, in accordance with expectation for these fragments. Circular
dichroism showed both polypeptides to be highly structured. The molar residue ellipticities at 222 nm, averaged
in each case from data on several column fractions, were
222,200 and 223,900 degrees cm2 dmol21, respectively;
these correspond, in the absence of other ordered structures, to about 62 and 67% a-helix [Greenfield and
Fasman, 1969], or fitting the spectra with a basis-set of
standard proteins by the CONTIN programme [Provencher
and Glöckner, 1981], average a-helix contents of 77 and
75% are obtained, with 3 and 4% b-structure, respectively. To determine whether the polypeptide chains have
entered their stable native fold, thermal denaturation
profiles were determined by circular dichroism (Fig. 1).
These show an extended plateau region, in which the
structure remains essentially unchanged, followed by a
sigmoidal transition to an unfolded state. The conformation is thus stable below the transition region. The
transition itself is broader than those of a single repeating
unit from the rod domain [Kahana et al., 1994; Kahana
and Gratzer, 1995], indicating that the unfolding process
is not fully cooperative along the rod in either case.
Chemical cross-linking with dimethylsuberimidate
gave no evidence of the formation of species of higher
molecular weight than the monomer by either polypeptide. Spectrin dimers under the same conditions were
totally cross-linked to the covalent dimer (Fig. 2).
Sedimentation equilibrium distributions of RFM at
concentrations of 40, 70, and 100 µg ml21, scanned at 234
and 220 nm, were well fitted by a monodisperse, ideal
solute with molecular weight (based on a calculated
partial specific volume of 0.740 ml g21 ) of 110,000 6
8,000 (n 5 5) (theoretical value, 97,000). RFE, measured
at concentrations of 120, 170, and 230 mg ml21 and
scanned at 280 and 234 nm, similarly gave an equilibrium
distribution consistent with a monodisperse solute of
molecular weight 130,000 6 5000 (n 5 6) (theoretical
value, 126,300). Results are summarised in Table I. The
precision of these measurements was probably limited by
the rather low protein concentrations, and does not
exclude fits that allow the presence of some aggregated
material; it leaves no doubt, however, that both polypeptides are monomeric, rather than dimeric.
Sedimentation velocity measurements gave corrected sedimentation coefficients (s020,w) of 4.9S for RFM
and 4.6S for RFE. Corresponding frictional ratios are
1.15 and 1.25, respectively, implying a relatively low
degree of asymmetry for both particles. That both are
Fig. 1. Thermal denaturation profiles, measured by circular dichroism,
of middle rod fragment, RFM (W) and rod with central deletion,
RFE (M).
significantly asymmetric and/or flexible is nevertheless
clear, since globular proteins of the same molecular
weight would sediment at about 6.0S and 7.2S, respectively [Halsall, 1967].
Electron microscopy of the shadowed preparations
showed the presence of elongated particles in both
polypeptides. The estimated lengths were about 33 6 7
nm (100 particles measured) for RFM and about 38 6 6
nm (105 particles) for RFE (Fig. 3). In the latter case there
appears to be some tendency towards end-to-end association. As Figure 3 shows, however, globular particles were
also distributed throughout the fields.
DISCUSSION
The expressed fragments were shown by conformational criteria to have entered the native protein fold. Both
were monomeric in solution. By electron microscopy
they had the form of short, flexible rods, although
globular particles were also present and probably represented polypeptides that had undergone conformational
collapse under the shearing stresses encountered during
drying. Globular objects can similarly be discerned in
electron micrographs of intact dystrophin [Pons et al.,
1990].
The lengths of the rods were in the range of 30–40
nm. Judged by the extended length of erythroid spectrin,
as measured in the electron microscope by the same
procedure [Shotton et al., 1979], a length of about 30 nm
would be expected for both fragments. On the other hand,
the observed length of the very stable and rigid a-actinin
rod, comprising four spectrin-like repeats, was 25 nm
[Imamura et al., 1988] or 22.5 nm [Flood et al., 1995].
Dystrophin Rod Domain
Fig. 2. Cross-linking of rod fragments with dimethylsuberimidate. Analysis of products by gel
electrophoresis in SDS. A: RFM (middle rod). Lanes a, unreacted rod fragment; b, protein after treatment
with cross-linking reagent at protein concentrations of 170 µg ml21; c, protein reacted at 90 µg ml21; d,
untreated erythroid spectrin; e, spectrin after treatment with reagent at protein concentration of 170 µg
ml21; f, spectrin treated at 90 µg ml21. B: RFE (rod fragment with central deletion). Lanes a, untreated
protein; b, protein treated with cross-linking reagent; c, untreated erythroid spectrin; d, spectrin treated
with cross-linking reagent. Left-hand tracks in both panels contain molecular weight markers.
249
250
Kahana et al.
Fig. 3. Shadowing electron microscopy of rod fragments, RFM (A) and RFE (B). Bar 5 50 nm.
TABLE I. Sedimentation Equilibrium of Dystrophin Fragments,
RFM and RFE
Fragment
Middle rod (RFM)
Concentration
(µg ml21 )
Wavelength
(nm)
Molecular
weighta
40
220
234
220
234
234
114,500
98,000
107,700
108,000
118,600
234
280
234
280
234
280
127,200
126,700
129,200
137,000
125,600
136,600
70
100
Fused rod extremities
(RFE)
120
170
230
aBest
fit to monodisperse ideal solute.
This value would lead to a predicted length of 45–50 nm
for RFE and RFM. The length of a single Drosophila
spectrin repeat in the crystal is close to 5 nm [Yan et al.,
1993], and on this basis a length of approximately 40 nm
would be expected for the fragments. Considering especially the variability of sequence in the dystrophin rod
repeats, a difference in contour length and stiffness can
certainly be envisaged. The asymmetry of the hydrodynamic particle in each case is relatively low, implying that
both chains are far from rigid. The rigidity might be
expected to be lower for a monomer than for a dimer of
laterally associated subunits; nevertheless, sedimentation
coefficients of spectrin and fodrin tetramer, spectrin
dimer, a-actinin rods of four and three repeats and of the
dystrophin fragments RFE and RFM, all adhere quite
closely to the same sedimentation coefficient-molecular
weight relation, namely s ~ M 0.44.
The expressed dystrophin rod fragment, RFM,
corresponds to the central portion of the rod domain,
beginning 7 residues before the start of repeat 8 and
terminating at the end of repeat 15. The fragment, RFE,
derived from a natural dystrophin mutant [England et al.,
1990], contains repeats 1 to 3 and the 11 amino acid
residues preceding, fused to a segment which starts in the
middle (centre of the B-helix) of repeat 19 and terminates
77 residues beyond the end of the rod domain; it thus
contains 8 repeats, like RFM. The identification of the
repeat boundaries is based on the alignment of Winder et
al. [1995] (which is almost identical to that of Koenig and
Kunkel [1990]). In the other proteins of the same
superfamily, spectrin and a-actinin, hetero- and homodimers, respectively, are formed by pairwise association between the repeats of antiparallel subunits, the first
(N-terminal repeat) with the last (C-terminal repeat), the
second with the last-but-one, and so on. Thus, if this
simple scheme applies to dystrophin, RFM has the
possibility of self-associating through three antiparallel
pairs of repeats and RFE through 6 such pairs.
In the case of a-actinin, the four repeats of the
isolated rod domain are sufficient to form an extremely
tight dimer [Imamura et al., 1988; Kahana and Gratzer,
1991], but in spectrin the situation is more complex. For
erythroid spectrin Speicher and his colleagues [Speicher
et al., 1992; Ursitti et al., 1996] found that lateral, antiparallel association of the a- and b-chains was a function
of the four repeats nearest to the N-terminus of the
b-chain and a complementary set at the C-terminus of the
a-chain. These were envisaged to nucleate the interaction,
bringing the remaining, more weakly interacting, pairs of
repeats into register. The first of the b-chain repeats was
found to be necessary but insufficient for association and
the smallest b-chain element that bound detectably to the
a-chain comprised repeats 1 and 2; addition of repeats 3
and 4 increased the association constant by successive
factors of 5 [Ursitti et al., 1996]. A feature of the strongly
interacting repeats is an N-terminal insert of eight amino
acids, also present in all a-actinin repeats.
Dystrophin Rod Domain
Requirements for inter-chain binding were also
defined by Viel and Branton [1994] for Drosophila
spectrin. Two repeats in each chain were found to be
essential, each of them containing the eight-residue
insert. In addition some residues in the adjoining nonhomologous, terminal sequences were found to contribute to the association, although this was not the case in
human spectrin [Ursitti et al., 1996].
A search of the dystrophin sequence revealed no
eight-residue stretches homologous to the inserts in the
a-actinin and terminal spectrin repeats. Allowing up to 3
amino acid differences, four related eight-residue sequences were found, but none was close to the Nterminus of a repeat in the internal alignment.
The only direct experimental data that bear on the
association state of native, intact dystrophin stem from
electron microscopy. Pons et al. [1990] concluded that
their preparations contained mainly monomers with some
dimers. Later data [Sato et al., 1992] should probably be
disregarded because of the authors’ subsequent doubts
concerning the identity of the material [Kake et al., 1995].
Winder et al. [1995] suggest that the poor sequence
conservation of surface residues in the dystrophin rod
repeats and the presence of loops between coiled-coil
elements argues strongly against lateral association. Moreover, the presumptive self-association sites, present in all
erythroid spectrin repeats, occur in only a few dystrophin
repeats, and of these all but two are out of register in an
antiparallel alignment [Winder et al., 1996].
Our results demonstrate that neither the middle
portion of the rod nor the ends, which would interact by
pairs in an antiparallel dimer, are able to self-associate
measurably. The rod domain is thus unlike those of
erythroid spectrin and a-actinin, for example, in its
behaviour. We have not proved that dystrophin is incapable of self-associating in other, unrelated modes, for
example end-to-end. Madhavan and Jarrett [1995] have
generated a fragment of the C-terminal domain, which on
immunoblots exhibits binding to intact dystrophin, but
there are no data on the affinity of this interaction. To
establish definitively whether dystrophin has any capacity to self-associate in a specific manner is likely to
require studies on the intact native protein.
ACKNOWLEDGMENTS
This work was made possible through the support
of the Muscular Dystrophy Group of Great Britain. We
are grateful to P.J. Marsh, Randall Institute, King’s
College, for help and advice on expression of the
proteins, to A.P.R. Brain of the Electron Microscopy Unit
of King’s College for electron microscopy, and to A.C.M.
Harris, National Institute for Medical Research, London,
for N-terminal sequencing.
251
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