| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
J. Biol. Chem., Vol. 275, Issue 32, 24484-24489, August 11, 2000
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
From the Departments of
Received for publication, March 13, 2000, and in revised form, May 18, 2000
Small subunit (16 S) rRNA from the archaeon
Haloferax volcanii, for which sites of modification were
previously reported, was examined using mass spectrometry. A census of
all modified residues was taken by liquid chromatography/electrospray
ionization-mass spectrometry analysis of a total nucleoside
digest of the rRNA. Following rRNA hydrolysis by RNase T1,
accurate molecular mass values of oligonucleotide products were
measured using liquid chromatography/electrospray ionization-mass
spectrometry and compared with values predicted from the corresponding
gene sequence. Three modified nucleosides, distributed over four
conserved sites in the decoding region of the molecule, were
characterized: 3-(3-amino-3-carboxypropyl)uridine-966, N6-methyladenosine-1501, and
N6,N6-dimethyladenosine-1518
and -1519 (all Escherichia coli numbering). Nucleoside
3-(3-amino-3-carboxypropyl)uridine, previously unknown in rRNA, occurs
at a highly conserved site of modification in all three evolutionary
domains but for which no structural assignment in archaea has
been previously reported. Nucleoside
N6-methyladenosine, not previously placed in
archaeal rRNAs, frequently occurs at the analogous location in
eukaryotic small subunit rRNA but not in bacteria.
H. volcanii small subunit rRNA appears to reflect the
phenotypically low modification level in the Crenarchaeota kingdom and
is the only cytoplasmic small subunit rRNA shown to lack pseudouridine.
Although posttranscriptional modification of RNA in general serves
to modulate regional structural features (1, 2), the observed and often
conserved patterns of modification (mononucleotide structure and
sequence location) result from the dual influences of phylogenetic
position and environmental factors such as temperature of growth. For
example, modifications used for structural stabilization in tRNAs of
bacterial thermophiles exhibit notable differences compared with those
found in archaeal thermophiles (3, 4). Understanding of the relative
importance of these influences and of the functional roles of these
modifications at the single nucleotide level requires detailed and
accurate knowledge of modification sites in a suitable number of
diverse organisms. A more narrowly focused question is the extent to
which the modification site, but not necessarily the modified
nucleotide structure, may be conserved. Such instances may point to
functionally important sites of RNA and to the molecular mechanisms
served by modification; the best example of this is the influence of
modification in the first position of the anticodon in tRNA on the
regulation of codon recognition (5). Knowledge of detailed rRNA
modification sites also ultimately bears on a number of related issues,
including, for example, the number and sequence specificity of
modification enzymes (see discussion in Ref. 6), the utilization (or
not) of small nucleolar RNAs as guides for ribose methylation (7, 8) or
other modifications (6), and the extent to which RNA modification
systems may be subject to lateral gene transfer, a prospect first
raised by Woese et al. (9).
Whereas information concerning modification sites is readily available
for tRNA as a consequence of the relatively large number of reported
tRNA sequences (10), much less is known concerning the diversity of
ribosomal RNA modifications (11), in part because of the greater
experimental complexity of mapping modification sites in large RNAs
(discussed in Refs. 8 and 12) compared with tRNA. Reliable rRNA data
consists of complete or extensive modification maps of
SSU1 rRNAs from Xenopus
laevis, yeast, human (11), and Escherichia coli (Ref.
13 and references therein) and LSU rRNAs of human (11) and
E. coli (Ref. 14 and references therein) and
extensive surveys of pseudouridylation sites in rRNA (6), particularly those from the LSU (15). To these studies can be added a number of
early reports, e.g. Ref. 16, especially those by Woese and co-workers (17, 18) of modifications in RNase T1 fragments obtained in the course of phylogenetic cataloging, even though the
structural identities of modified residues, as well as placement of the
modified T1 fragment in the RNA sequence (which predated the now common availability of rDNA sequences), were not known in many cases.
We report here an investigation of the modified sites in 16 S rRNA of
Haloferax volcanii using LC/ESI-MS. The locations of five
modified sites had earlier been reported based on partial RNA
sequencing in conjunction with oligonucleotide cataloging (19). Of
particular interest was modification at position 966 (E. coli numbering), inferred from the corresponding gene sequence to
be a U derivative, a conserved site of modification in the decoding
region of the rRNA that has been cross-linked to C-32 of the anticodon
loop of P-site tRNA (20). In various eukaryotes, including human
(21), X. laevis (21), and yeast (21, 22), the modified
residue has been established as
1-methyl-3-(3-amino-3-carboxypropyl)pseudouridine, a hypermodified
nucleoside (23) regarded as phenotypically eukaryotic (11, 12).
In E. coli, however, modification in this region occurs as
N
2-methylguanosine-966, 5-methylcytidine-967 (24), with
no apparent structural correlation with the eukaryotic modification. No
modification structure assignments in any archaeal SSU rRNA have
previously been made at this site although the occurrence of
modification has been documented in Sulfolobus solfataricus
(25, 26) and in RNase T1 catalogs showing the appropriate
conserved sequence for this loop (NCAACG) in Sulfolobus
acidocaldarius, Thermoproteus tenax (25),
Methanococcus jannaschii (27), and in fifteen other
methanogens (17); the modification motif is thus highly conserved.
Isolation and Enzymatic Hydrolysis of rRNA--
H.
volcanii (ATCC 29605) was grown as reported earlier (28). 30 S
ribosomal subunits were prepared (29), and the 16 S rRNA was isolated
by extraction with phenol and chloroform (29). Purity of the RNA was
assessed by gel electrophoresis (1% agarose) using an Applied
Biosystems 230A micropreparative electrophoresis system. 16 S rRNA was
hydrolyzed to nucleosides using nuclease P1 (Sigma), venom
phosphodiesterase I (E.C. 3.1.15.1, Sigma), and bacterial alkaline
phosphatase (E.C. 3.1.3.1, Calbiochem) (30). Digestion of rRNA (10 pmol/µl in 20 mM Tris hydrochloride plus 1 mM
EDTA, pH 7) by RNase T1 (E.C. 3.1.27.3) (Ambion,
Austin, TX) was carried out for 30 min at 37 °C at a concentration
of 1000 units/50 pmol of RNA.
LC/ESI-MS of Nucleosides from Total Digestion of H. volcanii 16 S
rRNA--
A Quattro II mass spectrometer with MassLynx version 3.1 data system (Micromass, Beverly, MA) interfaced to an HP 1090 liquid chromatograph with diode array detector (Hewlett-Packard, Palo Alto, CA) was used for all LC/MS studies. Two hundred picomoles (~100
µg) of rRNA hydrolysate was injected directly onto a 250 × 2.1-mm Supelco LC-18S column fitted with a matching 15 × 2.1-mm precolumn (Supelco, Bellefonte, PA). The column was eluted at a flow
rate of 300 µl/min using an ammonium acetate/acetonitrile gradient as
described previously (31), except the concentration of ammonium acetate
was decreased to 0.005 M for compatibility with
electrospray ionization. Diode array UV absorbance data were acquired
from 240-320 nm.
The chromatographic effluent was conducted without splitting into the
mass spectrometer, using the standard megaflow inlet. The ion source
was 180 °C. Capillary and lens voltages were 3.1 and 0.24 kV,
respectively, for measurement of positive ions. Data were
acquired in "centroid" mode over the mass range 105-450 in 0.9 s (with a 0.1-s interscan delay, for a cycle time of 1 s). The "Cluster" algorithm from the MassLynx software was used to interrogate the data set for the presence of unknown nucleosides; cluster values were 132 and 146 units for normal and
2'-O-methylated nucleosides, respectively.
LC/ESI-MS of Oligonucleotides from RNase T1 Digests
of H. volcanii 16 S rRNA--
The mass spectrometer and liquid
chromatograph are described in the preceding section; a Z-spray
interface was available for these studies. Fifty pmol (~25 µg) of
rRNA hydrolysate was injected directly onto a 300 × 1-mm Supelco
LC-18S column (Supelco) with a 15 × 1-mm OptiGuard C-18
precolumn cartridge (Optimize Technologies, Oregon City, OR). The
solvent system consisted of 0.8 M
1,1,1,3,3,3-hexafluoro-2-propanol (J. T. Baker Inc.) adjusted to
pH 7.0 with triethylamine, half of which was diluted 1:1 with
HPLC-grade water (Buffer A) whereas the other half was diluted 1:1 with
methanol (Burdick and Jackson, Muskegon, MI) (Buffer B) (32). The
column was eluted using a linear gradient of 0-100% Buffer B in
Buffer A plus Buffer B over 50 min at a flow rate of 60 µl/min. Diode
array UV data were acquired from 240-320 nm.
The chromatographic effluent was conducted without splitting into the
mass spectrometer. The ion source and desolvation temperatures were
140 °C and 300 °C, respectively. Capillary and lens voltages were
The posttranscriptional modification status of H. volcanii rRNA was examined using a combination of LC/MS-based
methods (33) involving analysis of mixtures of nucleosides produced by
total enzymatic hydrolysis (31) and of oligonucleotides from RNase T1 digestion. The latter analysis, carried out directly on
the total rRNA digest, provides accurate molecular mass values for oligonucleotide products, which can in turn be converted to base compositions (34) and correlated with specific oligonucleotide sequences in the rRNA through comparison with the corresponding gene
sequence (35).
A chromatogram based on UV detection from LC/MS analysis of a total
nucleoside digest of 16 S rRNA is shown in Fig.
1 and indicates the presence of three
modified nucleosides, acp3U, m6A, and
m26A. The assignments shown are based on
HPLC retention times compared with tabulated values for RNA nucleoside
standards (31) and on mass values for the protonated molecule and the
protonated base produced as a fragment ion (31). The molar ratio of
m26A:m6A is approximately 2, based on chromatographic peak areas by UV detection using mixture
standards (data not shown). The presence of acp3U was
unexpected, but of particular interest because it previously was known
to occur only in tRNA (36). Both the masses of the protonated
nucleoside and base (346 and 214, respectively) and the relatively
early retention time of 6.4 min are highly distinctive compared with
values for other RNA nucleosides (31, 36). Examination of the mass
spectra recorded during HPLC elution of A, U, G, and C provided no
evidence for additional modified nucleosides that might have co-eluted
with the major nucleosides and not have been evident in the UV
detection chromatogram.
To assign each modified nucleoside to the rRNA sequence, molecular
masses of oligonucleotides from an RNase T1 digest of the 16 S rRNA were determined by LC/ESI-MS for comparison with calculated masses of (unmodified) T1 oligonucleotides predicted from
the gene sequence (35). Shown in Fig. 2
are the traces for absorbance at 260 nm (extracted from photodiode
array data; panel A) and for base fragment ions of the
modified nucleosides (acp3U, m6A,
m26A) expected from results of the
initial modification screen shown in Fig. 1 (extracted from the high
cone voltage scan function; panels B-D). The
chromatographic profiles generated by these three ions
(m/z 212, 148, 162) thus mark the elution times
of oligonucleotides that contain them. These mass channels were time
aligned with total ion current profiles from a normal cone voltage scan
function, recorded in the same analysis, from which full mass spectra
of the modified oligonucleotides were derived.
Identities and Phylogenetic Comparisons of Posttranscriptional
Modifications in 16 S Ribosomal RNA from Haloferax
volcanii*
§,¶
Biochemistry and
¶ Medicinal Chemistry, University of Utah, Salt Lake City,
Utah 84112
ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
2.75 and 0.50 kV, respectively, for measurement of negative ions. Two
alternating scan functions were used for data acquisition. The first
one, used to determine oligonucleotide molecular masses, utilized a
44-V cone setting, a typical value for generating mass spectra with
minimal fragmentation. Data were acquired in continuum mode over the
mass range 480-1380 in 3 s (with a 0.2-s interscan delay, for a
cycle time of 3.2 s). The second scan function utilized a cone
setting of 130 V to fragment the oligonucleotides. Data were acquired
in continuum mode over the mass range 100-350 in 0.4 s (with a
0.1-s interscan delay, for a cycle time of 0.5 s).
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
View larger version (20K):
[in a new window]
Fig. 1.
LC/MS analysis of nucleosides from a
total enzymatic digest of H. volcanii 16 S rRNA.
Detection by UV absorbance at 260 nm. Shown below in panels
A-C are nucleosides whose identities were established from HPLC
retention times and mass spectra recorded every 0.5 s:
A, acp3U; B, m6A;
C, m26A.
View larger version (16K):
[in a new window]
Fig. 2.
LC/MS analysis of oligonucleotides from RNase
T1 hydrolysis of H. volcanii 16 S rRNA
showing the region of 4-mers and longer
oligomers. A, UV detection at 260 nm. B, mass channel 212 representing acp3U base
ion. C, mass channel 148, m6A base.
D, mass channel 162, m26A
base.
Shown in Fig. 3 are five summed mass
spectra across the apex of the peak eluting at 25.8 min (Fig. 2,
B and C), which is expected to include
m6A- and acp3U-containing T1
oligonucleotides. Although the coincident elution times of the two base
fragment ions suggests that they may belong to the same oligonucleotide
component, the corresponding molecular mass values dictate their
presence in different oligonucleotides, as follows. Comparison of the
measured masses of the three oligonucleotides A, B, and C with all
masses predicted from the gene sequence (19) allowed component B
(Mr 2574.5) to be readily assigned as
unmodified 1093-UACAUUAGp-1100 (calculated Mr
2574.5). The relative masses of the remaining oligonucleotides
(A, Mr 2305.5 and C, Mr
2673.8) are not present in the calculated T1 catalog, so
each one contains one of the two modified nucleotides. To derive the
T1 oligonucleotide to which each belongs, the residue mass
of each of the modified residues (14 Da for methyl in m6A;
101 Da for aminocarboxypropyl in acp3U) was subtracted in
turn from the molecular masses of oligonucleotides A and C. Allowable T1 compositions were obtained only for
Mr 2305.5 14 and for
Mr 2673.8 101. The m6A is
therefore confined to the oligonucleotide UAACAGp, whereas the
acp3U can be accommodated within either of two
(A3,C3,U)Gp sequences. The base composition of
the acp3U-containing RNase T1 fragment inferred
from the measured molecular mass was independently confirmed by
isolation of the 8-mer oligonucleotide by anion exchange and
reversed-phase chromatographies (35), digestion to nucleosides, and
LC/MS analysis. These results (data not shown) confirm the composition
acp3U plus (A3,C3)GP (derived from
chromatographic peak heights) with no unmodified uridine. The published
sequence (19) indicated unspecified modified A and U in the sequences
UAACAAGp and ACUCAACGp, respectively; the mass
data in the present study define the corresponding modified
oligonucleotides as 1498-UAm6ACAAGp-1505 and
964-ACacp3UCAACGp-971 (E. coli numbering).
|
The remaining modified nucleotide,
m26A, is present in an oligonucleotide
component eluting at 30.7 min (Fig. 2D); five summed mass
spectra spanning the apex of this peak are shown in Fig. 4. Subtraction of the modification
element CH3 × 2 (28 Da for one
m26A) from the indicated molecular mass
of the single oligonucleotide in this peak does not yield a mass value
allowed from the gene sequence-based T1 catalog.
Subtraction of CH3 × 4 (56 Da), however, yields an allowed
composition of (A2,C,U2)Gp, represented in five oligonucleotides in the RNA. Reference to the original RNase
T1 catalogs (19) allows assignment of this modified
oligonucleotide to one of three AAUCUGp oligonucleotides:
1518-m26Am26AUCUGp-1523
(E. coli numbering).
|
In summary, the experiments described revealed the presence of three
posttranscriptionally modified species, each of which was localized to
a specific sequence location within four sites in the RNA. Structure
assignments and sequence locations (E. coli numbering) are
summarized in Table I. No evidence was
found for an additional modified C assigned from RNase catalog data,
1401-GCCCGp-1405 (19) (see "Discussion"), as a result
of two key experiments: failure to observe additional modified
nucleoside(s) in the total nucleoside digest (Fig. 1) and failure to
find additional base ions released during analysis of RNase
T1 digestion products as in Fig. 2,
B-D. Specific attention was paid to mass values
corresponding to the known modified C bases in RNA (36) and to any
other low mass fragment ions that could be candidates for new bases of
unknown or unexpected structure, which would have been revealed in the analysis shown in Fig. 2. In addition, no oligonucleotide molecular mass values were found within the 
3-mer products (represented by Fig.
2A) that were unassignable based on expected RNA masses calculated from the gene sequence. However, this latter experiment alone is not considered conclusive because of the difficulty in making
Mr measurements in the (mass spectrally) complex
3- and 4-mer elution region in the chromatogram. Finally, no modified nucleosides or sites were found that might have evaded detection in
small oligonucleotides (e.g. NGp) from partial RNase
sequencing and cataloging (19).
|
| |
DISCUSSION |
|---|
|
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
|
|---|
In the present study three different modified nucleotide species were structurally identified and placed at four sites in the 16 S rRNA sequence. These modification sites (Table I) correlate with those reported from RNase T1 catalogs in conjunction with the corresponding gene sequence (19), but a modified cytidine (shown in the reported sequence as 1401-GCCCG-1405 (19)) was not found in the total nucleoside digest, as base fragment ion in LC/MS analysis of the RNase T1 digest, or in oligonucleotide molecular masses reflecting incremental mass additions to Mr 1278.8 (unmodified CCCGp). Were such a modified C present (and sufficiently stable to survive isolation and digestion protocols), it would have to elute underneath one of the seven nucleosides apparent in Fig. 1, would have to have a base fragment coincident in mass with a limited number of ubiquitous sugar-phosphate backbone fragment ions (present in significant excess in the analysis shown in Fig. 2), and in any instance would have a previously unknown structure.
The finding of acp3U in H. volcanii 16 S rRNA
(as 964-ACacp3UCAACGp-971) was unexpected. This
hypermodified uridine derivative (37, 38) was previously unknown in
rRNA. It occurs in bacterial tRNAs, where it is restricted to position
47 in the extra loop (21 of 45 reported sequences in which
acp3U occurs) or positions 20, 20a, or 20b in the
dihydrouridine loop (24 sequences) (10). Although there are 23 different modifications reported in rRNA from all sources (36), the
structures of only two uridine derivatives, pseudouridine and
2'-O-methyluridine, were previously established in
archaeal SSU rRNAs (39). The SSU rRNA position analogous to the
location of acp3U in H. volcanii (E. coli 966) appears to be nearly universally modified in other
phylogenetic domains (e.g. bacteria (24) and human and yeast
(11)), although the extent of conservation between domains of the
modified structure itself is limited but intriguing. As shown in Fig.
5 the 966 position in E. coli
is occupied by N2-methylguanosine, adjacent to a
second modification, 5-methylcytidine-967. Oligonucleotide catalog data
from other bacterial SSU rRNAs (Halobacteroides halobus (40)
and Brevibacillus brevis (41)) support the same tandem
modification pattern of 966-GC-967. However, in eukaryotes (11) N-966
is usually a conserved U that appears to be commonly modified to the
structurally related compound
1-methyl-3-(3-amino-3-carboxypropyl)pseudouridine (23) (structure in
Fig. 5).
|
RNase T1 catalogs of SSU rRNAs have been published by Woese and co-workers for S. solfataricus, S. acidocaldarius, and T. tenax (25), for M. jannaschii (27), and for 15 additional methanogens (17) in addition to H. volcanii (19). In all of these archaea the position analogous to 966 is designated as an unknown modified residue N in the conserved sequence 966-NCAACG. The extent to which the assignment of N-966 as acp3U is common to archaea must await study of a broader range of organisms.
In tRNA, the side chain of nucleoside acp3U was found to be biosynthesized from S-adenosylmethionine, based on 14C and 3H labeling (42). Although the occurrence of acp3U in tRNA, and possibly rRNA, is conserved, its biological function is presently unknown. The effect of side chain substitution by the 3-amino-3-carboxy moiety at N-3 of uridine was studied by nuclear magnetic resonance spectroscopy and found to result in only a small influence on the C3'-endo/C2'-endo ribose conformer population (43). However, evidence from nuclear magnetic resonance spectroscopy that acp3U binds Mg2+ in small oligonucleotides suggests a possible role for acp3U in the stabilization of regional RNA structure (44).
Assignment of m6A in the sequence 1498-UAm6ACAAGp-1505 constitutes the first known sequence placement of m6A in archaeal rRNA; its presence at a level of 1-2 residues was previously reported in S. solfataricus 16 S rRNA (39). The only previously established placement of m6A in SSU rRNA in other phylogenetic domains was in eukaryotes: X. laevis and human (11) and Rattus norvegicus (16, 45), at positions analogous to that in which it occurs in H. volcanii, in the highly conserved mRNA decoding region of the SSU rRNA. Interestingly, this nearly universally conserved A (position 1500 in the E. coli numbering system) appears from the limited data available to be commonly modified in other archaea (various methanogens (17, 27) and sulfur-dependent thermophiles (25)) but not in bacteria (18, 24, 40, 41, 46), yeast (11, 47), or Dictyostelium discoideum (48). The extent to which the modified A at this location in archaea is specifically m6A (an uncommon modification in both rRNA and tRNA) remains to be determined.
Two m26A nucleotides could have been assigned to any of five (A2,C,U2)Gp sequences present in H. volcanii SSU rRNA. Their assignment as tandem m26As at positions 1518-1519 at the interface of the ribosomal subunits (49) is expected in view of the very highly conserved nature (50) and functional importance of this tandem pair of modifications (51).
With the exception of acp3U-966 (which is otherwise a
bacterial tRNA modification), the modification structures and sites
found in H. volcanii 16 S rRNA are decidedly more eukaryotic
than bacterial in nature (Fig. 5 and Table I). A similar conclusion was
reached earlier concerning base and sugar modification motifs found in archaeal tRNA (4). However, this characteristic does not extend to
modification levels; the H. volcanii 16 S rRNA is the least modified cytoplasmic SSU rRNA of which we are aware. For example, in
eukaryotes the numbers range from about 20-80 modifications per rRNA
(yeast, ~22 and X. laevis, ~44 (11); R. norvegicus, ~77 (16)), and there are 11 in E. coli
(24). It is possible that lower modification levels are a
phenotypic characteristic of Euryarchaeota (52) from the archaeal
domain. RNase T1 catalogs (generally 5-mers) derived from
SSU rRNAs of mostly mesophilic methanogens suggest levels of ten or
fewer modified sites (17). Catalogs from hyperthermophiles from
the Crenarchaeota show about twice those levels (25), whereas a more
accurate estimate from a total nucleoside digest of S. solfataricus 16 S rRNA of ~38 residues was reported (39). The
high modification levels in S. solfataricus P2 (optimal
growth temperature, 75-80 °C) were attributed in part to a
role in structural stabilization, particularly from the
prevalence of 2'-O-methylated nucleotides (39).
However, earlier RNase catalog data for M. jannaschii
(optimal growth temperature, ~85 °C (27)) from the Euryarchaeota
domain showed only seven modifications (27), reflecting a total of
perhaps 10-12 in the molecule, still comparatively low. Taken together
with the modification levels of H. volcanii SSU rRNA (Ref.
19 and the present study), the intermediate level of modification
exhibited in M. jannaschii (27) may reflect a combination of
influences in the latter organism from phenotypically lower
modification levels in the Crenarchaeota, coupled with the functional
utility of modifications in the stabilization of rRNA in thermophiles.
| |
ACKNOWLEDGEMENT |
|---|
We are grateful to Jef Rozenski for access to a collection of SSU rRNA modification data currently being compiled.
| |
FOOTNOTES |
|---|
* This work was supported by NIGMS Grant GM29812 from the National Institutes of Health.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.
§ Present address: Laboratory of Neurotoxicology, National Institute of Mental Health, Bldg. 10, Rm. 3D40, Bethesda, MD 20892-1262.
To whom correspondence should be addressed. Tel.:
801-581-5581; Fax: 801-581-7457; E-mail:
james.mccloskey@m.cc.utah.edu.
Published, JBC Papers in Press, May 18, 2000, DOI 10.1074/jbc.M002153200
2 M. Schenk (Dalhousie University), personal communication.
| |
ABBREVIATIONS |
|---|
The abbreviations used are: SSU, small subunit rRNA (e.g. 16-18 S); LSU, large subunit rRNA (e.g. 23-28 S); LC/ESI-MS, directly combined liquid chromatography/electrospray ionization-mass spectrometry; acp3U, 3-(3-amino-3-carboxypropyl)uridine; m6A, N6-methyladenosine; m26A, N6,N6-dimethyladenosine.
| |
REFERENCES |
|---|
|
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
|
|---|
| 1. | Agris, P. F. (1996) Prog. Nucleic Acids Res. Mol. Biol. 53, 79-129 |
| 2. | Davis, D. R. (1998) in Modification and Editing of RNA (Grosjean, H. , and Benne, R., eds) , pp. 85-102, ASM Press, Washington, D. C. |
| 3. | Watanabe, K., Shinma, M., and Oshima, T. (1976) Biochem. Biophys. Res. Commun. 72, 1137-1144 |
| 4. | Edmonds, C. G., Crain, P. F., Gupta, R., Hashizume, T., Hocart, C. H., Kowalak, J. A., Pomerantz, S. C., Stetter, K. O., and McCloskey, J. A. (1991) J. Bacteriol. 173, 3138-3148 |
| 5. | Yokoyama, S., and Nishimura, S. (1995) in tRNA: Structure, Biosynthesis, and Function (Söll, D. , and RajBhandary, U. L., eds) , pp. 207-223, ASM Press, Washington, D. C. |
| 6. | Ofengand, J., and Fournier, M. J. (1998) in Modification and Editing of RNA (Grosjean, H. , and Benne, R., eds) , pp. 229-253, ASM Press, Washington, D. C. |
| 7. | Tollervey, D. (1996) Science 273, 1056-1057 |
| 8. | Bachellerie, J.-P., and Cavaillé, J. (1998) in Modification and Editing of RNA (Grosjean, H. , and Benne, R., eds) , pp. 255-272, ASM Press, Washington, D. C. |
| 9. | Woese, C. R., Gutell, R., Gupta, R., and Noller, H. F. (1983) Microbiol. Rev. 47, 621-669 |
| 10. | Sprinzl, M., Horn, C., Brown, M., Ioudovitch, A., and Steinberg, S. (1998) Nucleic Acids Res. 26, 148-153 |
| 11. | Maden, B. E. H. (1990) Prog. Nucleic Acid Res. Mol. Biol. 39, 241-303 |
| 12. | Limbach, P. A., Crain, P. F., and McCloskey, J. A. (1994) Nucleic Acids Res. 22, 2183-2196 |
| 13. | Bakin, A., Kowalak, J. A., McCloskey, J. A., and Ofengand, J. (1994) Nucleic Acids Res. 22, 3681-3684 |
| 14. | Kowalak, J. A., Bruenger, E., Hashizume, T., Peltier, J. M., Ofengand, J., and McCloskey, J. A. (1996) Nucleic Acids Res. 24, 688-693 |
| 15. | Ofengand, J., and Bakin, A. (1997) J. Mol. Biol. 266, 246-268 |
| 16. | Choi, Y. C., and Busch, H. (1978) Biochemistry 17, 2551-2560 |
| 17. | Balch, W. E., Fox, G. E., Magrum, L. J., Woese, C. R., and Wolfe, R. S. (1979) Microbiol. Rev. 43, 260-296 |
| 18. | Stackebrandt, E., Fowler, V. J., and Woese, C. R. (1983) Syst. Appl. Microbiol. 4, 326-337 |
| 19. | Gupta, R., Lanter, J. M., and Woese, C. R. (1983) Science 221, 656-659 |
| 20. | Brimacombe, R., Mitchell, P., Osswald, M., Stade, K., and Bochkariov, D. (1993) FASEB J. 7, 161-167 |
| 21. | Maden, B. E. H. (1986) J. Mol. Biol. 189, 681-699 |
| 22. | Raué, H. A., Klootwijk, J., and Musters, W. (1988) Prog. Biophys. Mol. Biol. 51, 77-129 |
| 23. | Saponara, A. G., and Enger, M. D. (1974) Biochim. Biophys. Acta 349, 61-77 |
| 24. | Brosius, J., Palmer, M. L., Kennedy, P. J., and Noller, H. F. (1978) Proc. Natl. Acad. Sci. U. S. A. 75, 4801-4805 |
| 25. | Woese, C. R., Gupta, R., Hahn, C. M., Zillig, W., and Tu, J. (1984) Syst. Appl. Microbiol. 5, 97-105 |
| 26. | Olsen, G. J., Pace, N. R., Nuell, M., Kaine, B. P., Gupta, R., and Woese, C. R. (1985) J. Mol. Evol. 22, 301-307 |
| 27. | Jones, W. A., Leigh, J. A., Mayer, F., Woese, C. R., and Wolfe, R. S. (1983) Arch. Microbiol. 136, 254-261 |
| 28. | Gupta, R. (1984) J. Biol. Chem. 259, 9461-9471 |
| 29. | Moazed, D., Van Stolk, B. J., Douthwaite, S., and Noller, H. F. (1986) J. Mol. Biol. 191, 483-493 |
| 30. | Crain, P. F. (1990) Methods Enzymol. 193, 857-865 |
| 31. | Pomerantz, S. C., and McCloskey, J. A. (1990) Methods Enzymol. 193, 796-824 |
| 32. | Apffel, A., Chakel, J. A., Fischer, S., Lichtenwalter, K., and Hancock, W. S. (1997) Anal. Chem. 69, 1320-1325 |
| 33. | Crain, P. F., Ruffner, D. E., Ho, Y., Qiu, F., Rozenski, J., and McCloskey, J. A. (1999) in Mass Spectrometry in Biology and Medicine (Burlingame, A. L. , Carr, S. A. , and Baldwin, M. A., eds) , pp. 531-551, Humana Press, Totowa, NJ |
| 34. | Pomerantz, S. C., Kowalak, J. A., and McCloskey, J. A. (1993) J. Am. Soc. Mass Spectrom. 4, 204-209 |
| 35. | Kowalak, J. A., Pomerantz, S. C., Crain, P. F., and McCloskey, J. A. (1993) Nucleic Acids Res. 21, 4577-4585 |
| 36. | Rozenski, J., Crain, P. F., and McCloskey, J. A. (1999) Nucleic Acids Res. 27, 196-197 |
| 37. | Ohashi, Z., Maeda, M., McCloskey, J. A., and Nishimura, S. (1974) Biochemistry 13, 2620-2625 |
| 38. | Friedman, S., Li, H. J., Nakanishi, K., and Van Lear, G. (1974) Biochemistry 13, 2932-2937 |
| 39. | Noon, K. R., Bruenger, E., and McCloskey, J. A. (1998) J. Bacteriol. 180, 2883-2888 |
| 40. | Oren, A., Weisburg, W. G., Kessel, M., and Woese, C. R. (1984) Syst. Appl. Microbiol. 5, 58-70 |
| 41. | Kop, J. A., Kopylov, A. M., Magrum, L., Siegel, R., Gupta, R., Woese, C. R., and Noller, H. F. (1984) J. Biol. Chem. 259, 15287-15293 |
| 42. | Nishimura, S., Taya, Y., Kuchino, Y., and Oashi, Z. (1974) Biochem. Biophys. Res. Commun. 57, 702-708 |
| 43. | Smith, W. S., Nawrot, B., Malkiewicz, A., and Agris, P. F. (1992) Nucleosides Nucleotides 11, 1683-1694 |
| 44. | Stuart, J. W., Basti, M. M., Smith, W. S., Forrest, B., Guenther, R., Sierzputowska-Gracz, H., Nawrot, B., Malkiewicz, A., and Agris, P. F. (1996) Nucleosides Nucleotides 15, 1009-1028 |
| 45. | Chan, Y. L., Gutell, R., Noller, H. F., and Wool, I. G. (1984) J. Biol. Chem. 259, 224-230 |
| 46. | Woese, C. R., Blanz, P., and Hahn, C. M. (1984) Syst. Appl. Microbiol. 5, 179-195 |
| 47. | Ofengand, J., Bakin, A., Wrzesinski, J., Nurse, K., and Lane, B. (1995) Biochem. Cell Biol. 73, 915-924 |
| 48. | McCarroll, R., Olsen, G. J., Stahl, Y. D., Woese, C. R., and Sogin, M. L. (1983) Biochemistry 22, 5858-5868 |
| 49. | Mitchell, P., Osswald, M., and Brimacombe, R. (1992) Biochemistry 31, 3004-3011 |
| 50. | van Knippenberg, P. H., Van Kimmenade, J. M., and Heus, H. A. (1984) Nucleic Acids Res. 12, 2595-2604 |
| 51. | van Knippenberg, P. H. (1985) in Structure, Function, and Genetics of Ribosomes (Hardesty, B. , and Kramer, G., eds) , pp. 412-424, Springer-Verlag New York Inc., NY |
| 52. | Woese, C. R., Kandler, O., and Wheelis, M. L. (1990) Proc. Natl. Acad. Sci. U. S. A. 87, 4576-4579 |
| 53. | Fellner, P. (1969) Eur. J. Biochem. 11, 12-27 |
| 54. | Youvan, D. C., and Hearst, J. E. (1981) Nucleic Acids Res. 9, 1723-1741 |
| 55. | Tautz, D., Hancock, J. M., Webb, D. A., Tautz, C., and Dover, G. A. (1988) Mol. Biol. Evol. 5, 366-376 |
CiteULike 
Complore 
Connotea 
Del.icio.us 
Digg 
Reddit 
Technorati What's this?
This article has been cited by other articles:
|
|
 |
|
  P. V. Sergiev, A. A. Bogdanov, and O. A. Dontsova Ribosomal RNA guanine-(N2)-methyltransferases and their targets Nucleic Acids Res., April 1, 2007; 35(7): 2295 - 2301. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 R. Guymon, S. C. Pomerantz, J. N. Ison, P. F. Crain, and J. A. McCloskey Post-transcriptional modifications in the small subunit ribosomal RNA from Thermotoga maritima, including presence of a novel modified cytidine RNA, March 1, 2007; 13(3): 396 - 403. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 D. V. Lesnyak, J. Osipiuk, T. Skarina, P. V. Sergiev, A. A. Bogdanov, A. Edwards, A. Savchenko, A. Joachimiak, and O. A. Dontsova Methyltransferase That Modifies Guanine 966 of the 16 S rRNA: FUNCTIONAL IDENTIFICATION AND TERTIARY STRUCTURE J. Biol. Chem., February 23, 2007; 282(8): 5880 - 5887. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
M. Hossain and P. A. Limbach Mass spectrometry-based detection of transfer RNAs by their signature endonuclease digestion products RNA, February 1, 2007; 13(2): 295 - 303. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. Mengel-Jorgensen and F. Kirpekar Detection of pseudouridine and other modifications in tRNA by cyanoethylation and MALDI mass spectrometry Nucleic Acids Res., December 1, 2002; 30(23): e135 - e135. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 T.-H. Tang, J.-P. Bachellerie, T. Rozhdestvensky, M.-L. Bortolin, H. Huber, M. Drungowski, T. Elge, J. Brosius, and A. Huttenhofer Identification of 86 candidates for small non-messenger RNAs from the archaeon Archaeoglobusfulgidus PNAS, May 28, 2002; 99(11): 7536 - 7541. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 J. OFENGAND, A. MALHOTRA, J. REMME, N.S. GUTGSELL, M. DEL CAMPO, S. JEAN-CHARLES, L. PEIL, and Y. KAYA Pseudouridines and Pseudouridine Synthases of the Ribosome Cold Spring Harb Symp Quant Biol, January 1, 2001; 66(0): 147 - 160. [Abstract] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| All ASBMB Journals | Molecular and Cellular Proteomics |
| Journal of Lipid Research | ASBMB Today |
ion of component A. Inset, Mr values calculated from
these mass spectra.
) in the 966 region (E. coli numbering) of SSU
rRNAs: E. coli (
,
1-methyl-3-(3-amino-3-carboxypropyl)pseudouridine.