Received 1 May 1998/Returned for modification 29 June 1998/Accepted 15 July 1998
 |
INTRODUCTION |
Ribosomal protein synthesis is one
of the oldest and best-conserved processes taking place in a living
cell. Within this commonality, it is possible to distinguish between
the eubacterial and eukaryotic translation machineries, with the
members of the domain Archaea displaying an amazingly
heterogeneous mixture of eubacterial and eukaryotic features
(9). One of the distinguishing characteristics of the three
clades is the differential sensitivity to inhibitors of the translation
process (2).
Amino acid residues are added to the growing peptide chain in the
ribosome by an elongation process that involves two GTP-switched elongation factors, denominated EF1 and EF2 in eukaryotes. EF1-GTP brings the aminoacyl-tRNA (as the so-called ternary complex) to the
acceptor site on the ribosome. After the nascent protein chain is
transpeptidated to the newly arrived tRNA, EF2 catalyzes a conformational switch of the organelle, such that the newly generated peptidyl-tRNA is moved from the acceptor site to the peptidyl site,
liberating the former for a new round of elongation. EF2 is a large
(more than 800-residue), probably multifunctional, and remarkable
protein that apparently binds to the same ribosomal structures as the
EF1-GTP-aminoacyl-tRNA complex. Observations in bacteria indicate that
this can easily be accomplished, since the overall shape of the
bacterial EF2 homolog mimics that of the whole ternary complex
(18, 19, 25).
The main purpose of the work described here was to identify by genetic
means, in Saccharomyces cerevisiae, cellular components targeted by the family of compounds denominated FPS, for fungal protein
synthesis inhibitors, which are semisynthetic derivatives of the
natural product sordarin (6, 13, 23). These compounds inhibit translation elongation in fungal cells with a high degree of
selectivity (10, 16), despite the high degree of
conservation in the translation components within eukaryotes. One of
these compounds, denominated GM193663, was used to select for resistant mutants. We present genetic evidence indicating that EF2 is part of the
target to which this new family of antifungals bind. While the
manuscript was in preparation, a related compound was used to determine
that EF2 is the target of sordarins (15). The present work
independently confirms and extends those observations by investigating
the mechanism of resistance and by constructing a three-dimensional
model of EF2 which shows a possible binding site for the drug on the
protein. Furthermore, we present evidence showing that the EF2 function
does not represent the whole target for sordarins, because a second
complementation group of resistant mutants in S. cerevisiae
was detected.
| |
MATERIALS AND METHODS |
Strains, growth conditions, plasmids, and genomic library.
All yeast strains used in this study are derivatives of S. cerevisiae SEY6210 (MAT
ura3-52 leu2-3,112 his3
200
trp1901 lys2-81 suc29), S. cerevisiae SEY6211
(MATa ura3-52 leu2-3,112 his3200 trp1901
ade2-101 suc29) (S. Emr), and S. cerevisiae 373 (MATa ade2-101) (A. Jimenez).
Growth media and methods for tetrad analysis, gene disruption, and
allele recovery in yeasts were as described previously (12).
Yeast transformations were done by the lithium acetate method as
described by Ito et al. (14). Escherichia
coli DH5 [endA1 hsdR1 supE44 thi-1 recA1 gyrA9
relA1
lacU169(
80lacZM15)] was used for transformation and preparation of plasmid DNA. All DNA manipulations were carried out by standard procedures (3, 22).
Spontaneous FPS-resistant mutants were selected by plating them on
yeast extract peptone dextrose (YPD) medium plates containing either 6 or 100 µg of GM193663 per ml and incubating the plates at 30°C
until colonies appeared. To score the resistant phenotype after genetic
crosses, agar plates containing 1 µg of the inhibitor per ml were
used.
The genomic library from the resistant mutant FPR1-4 was
constructed by ligating partially digested (Sau3A-I) genomic
DNA into the BamHI site of the pRS316 vector (CEN6
URA3 Ampr) and transforming the ligation mixture into
E. coli DH5. Plasmid DNA from 15,000 independent primary
E. coli transformants with an average insert size of 13 kb
was pooled, and the pooled DNA was used to transform the yeast.
Disruption of EFT1 and EFT2 in the various
wild-type and mutant strains was carried out by allele replacement with
HIS3 by following standard techniques (3, 12).
The disrupted loci were checked by Southern blotting.
In vitro activity and binding assays.
The growth inhibitory
activities of FPS compounds were determined in 96-well microtiter
plates by the antibiotic twofold serial dilution technique (from 125 to
0.01 µg/ml). One hundred microliters of YPD was inoculated with
105 CFU per well. The MIC was defined as the lowest
concentration of compound that inhibited 95% of the control growth
after 24 h of incubation at 30°C. Stock solutions of sordarin
(Bioprocessing, Glaxo Wellcome, Stevenage, United Kingdom) and other
FPS derivatives (Glaxo Wellcome, S.A.) were made in dimethyl sulfoxide
at 5 mg/ml. The FPS compounds tested were sordarin, GR135402, GM160575,
GM163420, GM165119, GM193663, and GM237354. The chemical structures of
representative FPS compounds used in the main experiments described in
this work are shown in Fig. 1. Commercial
protein synthesis inhibitors were obtained from Sigma. Anisomycin,
cycloheximide, and hygromycin were prepared in water at 10 mg/ml.
Verrucarin A was dissolved at 2.5 mg/ml in water-dimethyl sulfoxide
(3:1). In vitro translation assays with S50 extracts (supernatant
fraction obtained by centrifugation of cell extracts at 50,000 × g for 30 min) from cells growing exponentially at 30°C
(optical density at 600 nm = 2) in YPD were carried out by
following the poly(U) (Sigma)-directed incorporation of
[3H]phenylalanine (Amersham) into acid-insoluble material
essentially as described previously (5).
View larger version (13K):
[in this window]
[in a new window]
|
FIG. 1.
Structures of the inhibitors used in this study.
Sordarin is a natural product; the other two are more potent
semisynthetic derivatives.
|
|
For the binding assays, it was necessary to disrupt the EFT1
gene in all mutant and parental strain pairs in order to reduce the
background binding due to the EF2 protein expressed from the EFT1 locus. [3H]sordarin, labeled at the
aldehyde group, was synthesized at 180 GBq/mmol by the Glaxo Wellcome
Isotope Chemistry Group (Stevenage, United Kingdom). Binding assays
were done with 2 mg of protein from an S50 fraction and 0.1 µg of
[3H]sordarin (36 kBq) in a final volume of 500 µl under
the same conditions used for poly(Phe) synthesis. After 15 min of
incubation at room temperature, 400 µl from each sample was applied
to a prepacked PD-10 Sephadex-G-25M column (Pharmacia) equilibrated with the binding buffer. [3H]sordarin bound to
macromolecules was measured by counting the radioactivity in the
excluded fractions. The radioactivity counts obtained in the presence
of a 100-fold excess of cold sordarin were subtracted from all datum
points. Assay points were always obtained in duplicate, and the values
were averaged. Independent experiments were performed with cell
extracts prepared on different days.
Molecular mapping of EFT2 mutations.
Single-strand
conformation polymorphism (SSCP) mapping of the resistance mutations
was carried out as described previously (20).
EFT2 from the mutant and wild-type strains was amplified as
six overlapping fragments of approximately 500 bp by standard PCRs with
the oligonucleotide pairs listed in Table
1.
Resistant alleles were rescued (12) from their chromosomal
locations with pRS316 plasmids carrying a wild-type EFT2
gene gapped by the removal of either of two internal regions of the coding sequence covering either amino acids 439 to 502 or amino acids
495 to 652, as appropriate. Plasmids were recovered from Ura+ stable transformants, retransformed into a wild-type
strain to check the resistant phenotype, and sequenced. Sequencing was
done with an ABI Prism 310 Sequencer Analyzer according to the
manufacturer's recommendations.
Three-dimensional modeling of EF2.
Computational and
structural details will be published elsewhere, but in essence, the
model resulted from homology mapping of EF2 onto the crystal structure
of EF-G from Thermus thermophilus (Brookhaven Protein Data
Bank [PDB] entries 1dar and 1elo) (1, 7). The model was
built in three main steps. First, the water of crystallization in EF-G
was removed. Second, EF2 residues were substituted for the
corresponding ones in EF-G by using the HOMOLOGY module of the BIOSYM
package (Molecular Simulations Inc., San Diego, Calif.). Third,
sequence stretches present in S. cerevisiae EF2 and absent
from T. thermophilus EF-G were modeled. For this last
purpose, protein fragments of known three-dimensional structure with
the same or a closely related sequence were obtained from the
PDB-select database, and they were fitted to the structure by using
HOMOLOGY. The complete structure was refined with DISCOVER CVFF and
AMBER force fields (8). The quality of modeling throughout the different steps of the process was monitored with the PROCHECK program (17). Molecular docking experiments were performed
by taking into account the molecular electrostatic potential minima of
GM185832, the complementarity of the atomic contact surface areas, and
the chemical properties of the contact atoms by using the DOCKING and
DELPHY modules of BIOSYM.
| |
RESULTS |
GM193663-resistant mutants fall into two complementation
groups.
A genetic approach was used to identify targets of
GM193663. Spontaneous resistant mutants in S. cerevisiae,
arising at frequencies of 10
7 to 108, were
selected at concentrations of GM193663 of 6 and 100 µg/ml on YPD
medium plates. All mutants were cross resistant to the members of the
FPS class tested (see Materials and Methods). No significant
cross-resistance was observed with other protein synthesis inhibitors:
anisomycin, cycloheximide, hygromycin, and verrucarin A.
Twenty-five independently isolated mutants were analyzed by making
genetic crosses, and at least 16 spore tetrads from each cross were
dissected to score the resistance phenotype of the segregants.
Resistance segregated in all cases as a single mutation in a Mendelian
fashion. It was observed in the backcrosses that the resistance of the
heterozygous diploids always fell between those of the two parents, but
the MICs for the diploids spread widely between the MIC for the
sensitive parental strain (0.2 µg/ml) and the MIC for the resistant
parental strain, indicating that resistance is a semidominant
phenotype. All 10 mutants obtained at the lower dose fell into a single
complementation group, which was named FPR1. From the 15 mutants isolated at the higher dose, 11 fell into FPR1 and
the remaining 4 defined a second complementation group,
FPR2. In summary, of 25 mutants analyzed, 21 fell into one
complementation group and 4 fell into a second one. This defines two
genetic loci capable of giving resistance to GM193663 when they are
mutated.
Poly(Phe) synthesis is resistant to GM193663 in mutant cell
extracts.
One of the most common mechanisms of resistance to
antimicrobials arises through changes in transport systems for the
inhibitory compounds. Since that is uninformative regarding the mode of
action of the inhibitor, the sensitivity of poly(Phe) synthesis to
GM193663 was tested with cell extracts from mutant strains. The 50S
supernatants were prepared from total cell homogenates obtained from
one of the most dominant mutants in each group (FPR1-5 and
FPR2-6). Poly(U)-directed synthesis of poly(Phe) was assayed
in the presence of increasing concentrations of GM193663. The
inhibition curves showed that resistance is also manifested in cell
extracts (Fig. 2), thus excluding
transport mechanisms and indicating that the mutations affect elements
involved in the mode of action of the inhibitor. It is also apparent
that the curve for the extract from the FPR2 mutant has a
complex shape. It could be interpreted to mean that there are at least
two cellular components with different sensitivities to GM193663.
Clarification of this issue will be facilitated by the identification
of the gene involved. Cloning and characterization of the gene mutated
in complementation group FPR1 are presented below. Group
FPR2 is currently being analyzed and will be described elsewhere.
View larger version (21K):
[in this window]
[in a new window]
|
FIG. 2.
Inhibition of poly(Phe) synthesis by GM193663 in
extracts from wild-type and resistant strains. Cell extracts from
mutants belonging to complementation groups 1 (FPR1) and 2 (FPR2) and from their corresponding parental strains were
programmed with poly(U) to synthesize poly(Phe) in the presence of
various concentrations of GM193663, as described in Materials and
Methods. Data from two independent experiments are represented
together.  , resistant mutant;  , wild-type parental strain.
|
|
The gene mutated in complementation group 1 is
EFT2.
Attempts to clone FPR1 by
transforming wild-type libraries into the least dominant mutant strain
failed. Therefore, a gene library on a centromeric vector was
constructed from one of the most dominant alleles, FPR1-4
(see Materials and Methods). Screening of 23,000 S. cerevisiae colonies transformed with the library DNA yielded six
GM193663-resistant (GM193663r) isolates. Plasmid DNA
recovered from the resistant transformants was analyzed to identify the
common regions and the minimum sequence capable of conferring
resistance. This was found to be a 3.5-kb EcoRI fragment,
which was then partially sequenced. The sequences obtained matched that
of EFT2, one of the two genes which encode the 842-residue
EF2 in S. cerevisiae (21). As a further check, EFT2 was disrupted in the FPR1-4 haploid strain.
The disruptant was viable due to the presence of an intact
EFT1 locus, but resistance was lost, confirming that the
mutation giving GM193663 resistance was on EFT2. In
agreement with this result, independent work with the pathogenic yeast
Candida albicans has shown that EF2 is the primary
sordarin-binding protein in cell extracts from this organism (11). GM193663 can therefore inhibit the growth of yeast
cells by interacting with EF2.
Most resistance mutations cluster on a 50-amino-acid segment of
EF2.
S. cerevisiae EF2 has 53% homology to bacterial
EF-G, and it presumably folds into the same structural and functional
domains (see below) (1, 7). Localization of resistance
mutations within the EF2 protein could give clues to the mechanism of
resistance and hence to the mode of action of GM193663. To avoid
sequencing this large gene from all mutants, EFT2 from
several mutant strains was amplified as a series of six overlapping PCR
fragments. SSCP techniques were used to map the mutations to individual
fragments. Fragments from 11 mutants displayed altered mobility
compared with that of the corresponding fragment from the parental
strain. In all cases, the mobility change affected fragment D or E only (Fig. 3A), indicating that changes in
just a small area of the protein can confer GM193663 resistance.
View larger version (61K):
[in this window]
[in a new window]
|
FIG. 3.
Mapping of the GM193663r mutations on the
primary structure of EF2. (A) Overlapping fragments of the
EFT2 gene used in the mapping by SSCP. Base pair 1 corresponds to the A in the first ATG codon. (B) Position and nature of
the amino acid change in the sequenced mutant alleles of
EFT2. (C) Protein sequence comparison of the region in EF2
from different organisms corresponding to the region where most
GM193663 resistance mutations were found in S. cerevisiae.
The mutated positions are boxed, and the amino acid residue found in
the resistant mutant is shown on top of each box. Fu, fungi; An,
animals; Pl, plants; Pr, protists.
|
|
Fragments displaying an altered mobility were cloned, and both strands
were completely sequenced, as was the corresponding region of the
original library clone, FPR1-4. A single base change was
detected in each case, leading to an amino acid substitution in the EF2
protein. Two mutants were found to have the same mutation. Two others
had different substitutions at the same position in the protein,
reducing to nine the total number of mutated positions. The nature of
the amino acid changes is shown in Fig. 3B. Six of the altered
positions clustered on a 50-amino-acid segment which maps to domain 3 of EF-G (see below); the remaining three changes map to positions
flanking this region. Although the clustering of mutations around the
one found in the genomic clone made it unlikely that they were PCR
artifacts, the possibility was checked by rescuing the mutations from
the genomes of several resistant isolates by using "gap repair"
techniques for allele rescue (see Materials and Methods). The repaired
plasmids were transformed into a eft1 eft2 S. cerevisiae 6210, and the region of interest was sequenced. They
were found to confer resistance and to have the expected base changes,
confirming that the observed amino acid replacements were responsible
for the resistance.
Figure 3C shows the six more tightly clustered mutations in the context
of the sequences of other eukaryotic EF2 proteins from widely divergent
groups. The nature of the amino acid substitutions does not give
obvious clues to the mechanism of resistance, but the sequence
alignment shows that they are located in a region bracketing a highly
conserved and hence potentially important region. It is therefore
surprising how radical an amino acid change EF2 can tolerate and yet
remain functional. The Q490E and Y521D mutations introduce an acidic
residue at a position where none is found naturally. Another
"unnatural" change is introduced by the A562P mutation. Yet, the
resistant mutants grow at a rate not significantly different from that
of the wild type in the absence of the drug (data not shown).
Resistant EF2 molecules have reduced affinity for GM193663.
Clustering of GM193663 mutations within a small area of the EF2 protein
strongly suggested that they might change the structure of a binding
site for the drug. The hypothesis was tested by preparing cell extracts
from the resistant mutants and measuring the level of binding of
[3H]sordarin to macromolecules by gel filtration. This is
an available labeled analog of GM193663 which competes for binding and
shows cross-resistance (data not shown). A background of binding
activity was expected, and indeed was observed, due to the presence of the EF2 protein expressed from the EFT1 gene, the second
gene encoding EF2 in S. cerevisiae (21, 24). To
simplify the system, EFT1 was interrupted in eight resistant
strains with unique mutations in EF2. Cell extracts containing cytosol
and ribosomes were prepared from the deletants, and the level of
[3H]sordarin binding was measured. It was observed that
GM193663 resistance mutations in EF2 eliminate the binding of the
labeled analog to cellular macromolecules, thus explaining the
resistance. Table 2 shows the results
from two independent binding experiments. It can be seen that when all
EF2 molecules in the cell carry a mutation for resistance to GM193663,
there is no significant sordarin binding to macromolecules.
View this table:
[in this window]
[in a new window]
|
TABLE 2.
Specific binding of [3H]sordarin to
macromolecules in cell extracts from wild-type and
GM193663r mutant strainsa
|
|
Mutated positions define a putative binding pocket in a
three-dimensional model of EF2.
Completely independent of the work
described above, a three-dimensional model of S. cerevisiae
EF2 was constructed by using as a scaffold the crystal structure of
EF-G from T. thermophilus (PDB entries 1dar and 1elo)
(1, 7) (see Materials and Methods). The values from the
Ramachandran plots obtained for the modeled EF2 structure (Table
3) allow a high degree of confidence in
the overall correctness of the model.
View this table:
[in this window]
[in a new window]
|
TABLE 3.
Quality tests of T. thermophilus EF-G crystal
structure and S. cerevisiae EF2 model by using the
PROCHECK programa
|
|
Next, a molecular docking experiment was performed with EF2 and another
member of the FPS family of compounds, GM185832 (see Materials and
Methods). The most favorable binding site that emerged was the one
shown in Fig. 4A, at a cleft in what
would be domain 3 in EF-G. Remarkably, this predicted binding site
coincided with the location on the model of the sequenced GM193663
resistance mutations. Seven of the 10 mutations which block binding
fall exactly on or around this predicted binding site (Fig. 4B). This area of the protein corresponds to a very flexible domain of EF-G, presumed to be a hinge connecting the G-protein portion of the molecule
to putative rRNA-binding domains (1, 7). According to this
model, FPS inhibitors could act as a molecular wedge, reducing the
flexibility of the factor around domain 3. It can clearly be seen in
Fig. 4B that the mutations would block access to or change the
structure of the proposed binding pocket.
View larger version (75K):
[in this window]
[in a new window]
|
FIG. 4.
Calculated three-dimensional model of S. cerevisiae EF2. (A) Site of most favorable interaction of GM185832
on EF2, obtained from molecular docking experiments (see Materials and
Methods for details). (B) Detail of the predicted contact region
between the protein and the inhibitor, with the addition of the side
chains of the amino acid residues present at the positions found to be
mutated in the different resistant alleles.
|
|
The gene mutated in complementation group 2 is not
EFT1.
Since mutations in EFT2, encoding
EF2, confer resistance to FPS compounds, EFT1, the second
expressed gene encoding this factor in S. cerevisiae
(21), was a natural candidate as an explanation for the
second complementation group of resistant mutants. Disruption of
EFT1 in a mutant from this group did not abolish resistance, however, negating the hypothesis. The result was confirmed by genetic
crosses and segregation analysis between FPR2-6 and an EFT1 locus genetically tagged with HIS3. Thus,
there is at least one more cellular component, apart from EF2, involved
in the mode of action of FPS compounds.
| |
DISCUSSION |
The ribosomal protein synthesizing machinery is thought to be
highly conserved among eukaryotic organisms, perhaps with the exception
of fungi, which have an additional soluble elongation factor (EF3) not
present in other analyzed eukaryotes (4). It was therefore
widely believed that protein synthesis inhibitors could make good
antibacterial compounds but were unlikely to be selective inhibitors of
eukaryotic microbes, again with the possible exception of drugs aimed
at fungal EF3. It therefore came as a surprise when our genetic
analysis showed that S. cerevisiae EF2, one of the most
ancient and conserved proteins throughout evolution, was a target of
Glaxo Wellcome's FPS family of selective antifungals, as evidenced by
the fact that single point mutations on the protein conferred
resistance to six different members of the class.
Eighty percent of the resistance mutations affected EF2. The fact that
7 of 10 alleles sequenced had changes clustered on a 50-amino-acid
segment of the EF2 protein, the same one predicted to fold into an
FPS-binding pocket in our modeling experiments, plus the fact that
mutants displayed negligible binding to FPS compounds, strongly
suggests that the mutated positions define the binding site for the
drug. Mutations outside of this pocket could reduce the binding
indirectly by affecting the folding of the binding site. Preliminary
results from cross-linking and protease digestion experiments are in
agreement with the hypothesis presented above. A detailed analysis by
mass spectroscopy will be undertaken to establish this point.
The proposed FPS binding site is located on what would be domain 3 of
EF2 by comparison with the X-ray structure of EF-G. Figure 3C shows
that some of these positions are highly conserved in EF2 proteins from
different organisms. Yet all these substitutions allow mutated EF2 to
catalyze translation elongation. This agrees with the notion, deduced
from the EF-G structure, that a precise conformation may not be
essential for the role of this protein domain, which may only require
some global physicochemical property, such as flexibility, to function
(1, 7). Impediment of EF-G's flexibility has been suggested
to underlie fusidic acid's mode of action (7). FPS
inhibitors could be doing exactly the same thing, acting also on this
hinge region, but from the side of the factor opposite the putative
binding site for fusidic acid. Recently communicated results, obtained
with cloned EF2 and a different sordarin derivative, found resistant
mutants with changes in parts of the protein not affected by the
mutations described here but corresponding to areas found to be mutated
in EF-G proteins resistant to fusidic acid. The mechanism of resistance
brought about by those mutations is unknown, but in agreement with the hypothesis presented above, some mutants were found to have
cross-resistance to fusidic acid (15).
It is noteworthy that 21 spontaneous mutants with changes on
EFT2 were found but that none had mutations on
EFT1. This may be due to the partial dominance of the
resistant phenotype and the fact that the EFT2 promoter
seems to be 2.5-fold more active than that of EFT1
(24). A highly dominant phenotype may be needed to detect
EFT1 mutants when EFT2 is being expressed at such
high levels.
The existence of a second complementation group of FPS-resistant
mutants indicates that the cellular function inhibited by FPS compounds
is not carried out by EF2 alone. The involvement of more than one
component in defining the functional target for sordarins could
contribute to the selectivity of these compounds, despite the high
degrees of homology between individual molecular components of the
target. Our genetic data show that the gene mutated in the second
complementation group does not encode EF2. Preliminary biochemical
experiments indicate that resistance in this group is associated with
ribosomes (data not shown), and in C. albicans,
high-affinity binding to EF2 requires the presence of ribosomes
(11), supporting a role for ribosomal components in the
interaction between the inhibitors and EF2. Given the number of
isolates analyzed so far, the mutant screen cannot be considered saturated. Thus, the pathway inhibited by sordarins could still contain
additional components, all presumably involved in the EF2 function and
hence ribosomal translocation.
New selective antifungal agents are sorely needed in the clinic.
Compounds from this family are very good candidates that could be used
to close some important gaps in the existing antifungal drug portfolio.
FPS inhibitors can also be useful tools in dissecting the mechanism of
the elongation cycle in eukaryotic ribosomes, including identification
of the ribosomal components involved, something that we hope to start
achieving once cloning of the FPR2 gene is accomplished.
We are indebted to M. J. Serramía for expert technical
assistance, M. Gómez for performing gene disruptions, J. M. Domínguez for measuring poly(Phe) synthesis, and our colleagues
in the Isotope Chemistry Group (Glaxo Wellcome) for providing tritiated
sordarin. The many scientific discussions and the critical reading of
the manuscript by J. P. G. Ballesta are also gratefully
acknowledged.
| 1.
|
AEvarsson, A.,
E. Brazhnikov,
M. Garber,
J. Zheltonosova,
Y. Chirgadze,
S. al Karadaghi,
L. A. Svensson, and A. Liljas.
1994.
Three-dimensional structure of the ribosomal translocase: elongation factor G from Thermus thermophilus.
EMBO J.
13:3669-3677[Medline].
|
| 2.
|
Amils, R.,
L. Ramirez,
J. L. Sanz,
I. Marin,
A. G. Pisabarro,
E. Sanchez, and D. Ureña.
1990.
Phylogeny of antibiotic action, p. 645-654.
In
W. E. Hill, A. Dahlberg, R. A. Garrett, P. B. Moore, D. Schlessinger, and J. R. Warner (ed.), The ribosome: structure, function and evolution. American Society for Microbiology, Washington, D.C.
|
| 3.
|
Ausubel, F. M.,
R. Brent,
R. E. Kingston,
D. D. Moore,
J. G. Seidman,
J. A. Smith, and K. Struhl (ed.).
1997.
Current protocols in molecular biology.
John Wiley & Sons, Inc., New York, N.Y.
|
| 4.
|
Belfield, G. P., and M. F. Tuite.
1993.
Translation elongation factor 3: a fungus-specific translation factor?
Mol. Microbiol.
9:411-418[Medline].
|
| 5.
|
Colthurst, D. R.,
P. Chalk,
M. Hayes, and M. F. Tuite.
1991.
Efficient translation of synthetic and natural mRNAs in an mRNA-dependent cell-free system from the dimorphic fungus Candida albicans.
J. Gen. Microbiol.
137:851-857[Medline].
|
| 6.
|
Coval, S. J.,
M. S. Puar,
D. W. Phife,
J. S. Terracciano, and M. Patel.
1995.
SCH57404, an antifungal agent possessing the rare sordaricin skeleton and a tricyclic sugar moiety.
J. Antibiot.
48:1171-1172[Medline].
|
| 7.
|
Czworkowski, J.,
J. Wang,
T. A. Steitz, and P. B. Moore.
1994.
The crystal structure of elongation factor g complexed with GDP, at 2.7 angstrom resolution.
EMBO J.
13:3661-3668[Medline].
|
| 8.
|
Dauber-Osguthorpe, P.,
V. A. Roberst, and J. Wolff.
1992.
Structure and energetics of ligand binding to proteins: E. coli dihydrofolate reductase-trimethoprim, a drug-receptor system.
Proteins
4:409-417.
|
| 9.
|
Dennis, P. P.
1997.
Ancient ciphers translation in Archaea.
Cell
89:1007-1010[Medline].
|
| 10.
|
Dominguez, J. M.,
V. A. Kelly,
O. S. Kinsman,
M. S. Marriott,
F. Gomez de las Heras, and J. J. Martin.
1998.
Sordarins: a new class of antifungals with selective inhibition of the protein synthesis elongation cycle in yeast.
Antimicrob. Agents Chemother.
42:2274-2278[Abstract/Free Full Text].
|
| 11.
|
Dominguez, J. M., and J. J. Martin.
1998.
Identification of elongation factor 2 as the essential protein targeted by sordarins in Candida albicans.
Antimicrob. Agents Chemother.
42:2279-2283[Abstract/Free Full Text].
|
| 12.
|
Guthrie, C., and G. R. Fink.
1991.
Methods in enzymology, vol. 194. Guide to yeast genetics and molecular biology.
Academic Press Inc., San Diego, Calif.
|
| 13.
|
Hauser, D., and H. P. Sigg.
1971.
Isolierung und Abbau von Sordarin. [Isolation and decomposition of sordarin.] Helv.
Chim. Acta
54:1178-1190.
|
| 14.
|
Ito, H.,
Y. Fukuda,
K. Murata, and A. Kimura.
1983.
Transformation of intact yeast cells treated with alkali cations.
J. Bacteriol.
153:163-168[Abstract/Free Full Text].
|
| 15.
|
Justice, M. C.,
M. J. Hsu,
B. Tse,
T. Ku,
J. Balkovec,
D. Schmatz, and J. Nielsen.
1998.
Elongation factor 2 as a novel target for selective inhibition of fungal protein synthesis.
J. Biol. Chem.
273:3148-3151[Abstract/Free Full Text].
|
| 16.
|
Kinsman, O. S.,
P. A. Chalk,
H. C. Jackson,
R. F. Middleton,
A. Shuttleworth,
B. A. M. Rudd,
C. A. Jones,
H. M. Noble,
H. G. Wildman,
M. J. Dawson,
C. Stylli,
P. J. Sidebottom,
B. Lamont,
S. Lynn, and M. V. Hayes.
1998.
Isolation and characterisation of an antifungal antibiotic (GR135402) with protein synthesis inhibition.
J. Antibiot.
51:41-49[Medline].
|
| 17.
|
Laskowski, R. A.,
M. W. MacArthur,
D. S. Moss, and J. M. Thornton.
1993.
PROCHECK: a program to check the stereochemical quality of protein structures.
J. Appl. Crystallogr.
26:283-291.
|
| 18.
|
Nissen, P.,
M. Kjeldgaard,
S. Thirup,
G. Polekhina,
L. Reshetnikova,
B. F. Clark, and J. Nyborg.
1995.
Crystal structure of the ternary complex of Phe-tRNAPhe, EF-Tu, and a GTP analog.
Science
270:1464-1472[Abstract/Free Full Text].
|
| 19.
|
Nyborg, J.,
P. Nissen,
M. Kjeldgaard,
S. Thirup,
G. Polekhina,
B. F. C. Clark, and L. Reshetnikova.
1996.
Structure of the ternary complex of EF-Tu: macromolecular mimicry in translation.
Trends Biochem. Sci.
21:81-82[Medline].
|
| 20.
|
Orita, M.,
H. Iwahana,
H. Kanazawa,
K. Hayashi, and T. Sekiya.
1989.
Detection of polymorphisms of human DNA by gel electrophoresis as single-strand conformation polymorphisms.
Proc. Natl. Acad. Sci. USA
86:2766-2770[Abstract/Free Full Text].
|
| 21.
|
Perentesis, J. P.,
L. D. Phan,
W. B. Gleason,
D. C. LaPorte,
D. M. Livingston, and J. W. Bodley.
1992.
Saccharomyces cerevisiae elongation factor 2. Genetic cloning, characterization of expression, and G-domain modeling.
J. Biol. Chem.
267:1190-1197[Abstract/Free Full Text].
|
| 22.
|
Sambrook, J.,
E. F. Fritsch, and T. Maniatis.
1989.
Molecular cloning: a laboratory manual, 2nd ed.
Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y.
|
| 23.
|
Schneider, G.,
H. Anke, and O. Sterner.
1995.
Xylarin, an antifungal Xylaria metabolite with an unusual tricyclic uronic acid moiety.
Nat. Prod. Lett.
7:309-316.
|
| 24.
|
Veldman, S.,
S. Rao, and J. W. Bodley.
1994.
Differential transcription of the two Saccharomyces cerevisiae genes encoding elongation factor 2.
Gene
148:143-147[Medline].
|
| 25.
|
Wilson, K. S., and H. F. Noller.
1998.
Mapping the position of translational elongation factor EF-G in the ribosome by directed hydroxyl radical probing.
Cell
92:131-139[Medline].
|
![[Feedback]](/icons/banner/feedbackACT.gif){kind=icon}
![[For Subscribers]](/icons/banner/subscriberACT.gif){kind=icon}
![[Archive]](/icons/banner/archiveACT.gif){kind=icon}
![[Search]](/icons/banner/searchACT.gif){kind=icon}
![[Contents]](/icons/banner/tocACT.gif){kind=icon}
Top
Abstract
Introduction
