From the Immunochemistry Laboratory, National Institute of Immunology, Aruna Asaf Ali Road, New Delhi 110067, India
Received for publication, March 19, 2001, and in revised form, April 25, 2001
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Mammalian ribonucleases interact very
strongly with the intracellular ribonuclease inhibitor (RI). Eukaryotic
cells exposed to mammalian ribonucleases are protected from their
cytotoxic action by the intracellular inhibition of ribonucleases by
RI. Human pancreatic ribonuclease (HPR) is structurally and
functionally very similar to bovine RNase A and interacts with human RI
with a high affinity. In the current study, we have investigated the involvement of Lys-7, Gln-11, Asn-71, Asn-88, Gly-89, Ser-90, and
Glu-111 in HPR in its interaction with human ribonuclease inhibitor.
These contact residues were mutated either individually or in
combination to generate mutants K7A, Q11A, N71A, E111A, N88R, G89R,
S90R, K7A/E111A, Q11A/E111A, N71A/E111A, K7A/N71A/E111A, Q11A/N71A/E111A, and K7A/Q11A/N71A/E111A. Out of these, eight mutants,
K7A, Q11A, N71A, S90R, E111A, Q11A/E111A, N71A/E111A, and
K7A/N71A/E111A, showed an ability to evade RI more than the wild type
HPR, with the triple mutant K7A/N71A/E111A having the maximum RI
resistance. As a result, these variants exhibited higher cytotoxic
activity than wild type HPR. The mutation of Gly-89 in HPR produced no
change in the sensitivity of HPR for RI, whereas it has been reported
that mutating the equivalent residue Gly-88 in RNase A yielded a
variant with increased RI resistance and cytotoxicity. Hence, despite
its considerable homology with RNase A, HPR shows differences in its
interaction with RI. We demonstrate that interaction between human
pancreatic ribonuclease and RI can be disrupted by mutating residues
that are involved in HPR-RI binding. The inhibitor-resistant cytotoxic
HPR mutants should be useful in developing therapeutic molecules.
Mammalian ribonucleases constitute a ubiquitous superfamily of
proteins with a high level of structural and functional divergence. These include a group of homologous proteins isolated from many mammalian, avian, reptilian, and amphibian sources and are collectively known to be part of the RNase A superfamily (1). The current resurgence
of interest in RNases is the result of the discovery of RISBASES
(RNases with Special Biological Actions), which have been identified to
influence tumor cell growth, neurological development, and biological
differentiation (2). An important biological function of mammalian
RNases may be host defense, as has been observed in the case of the two
eosinophil RNases, eosinophil cationic protein and
eosinophil-derived neurotoxin, which exhibit antiviral, antibacterial,
antiparasitic, and neurotoxic activities (3-7). Also, frog RNase
onconase and bovine seminal ribonuclease (BS-RNase)1 exhibit antitumor
activity (8-11).
Human pancreatic ribonuclease (HPR) is secretory in nature and has been
considered as a counterpart of bovine pancreatic RNase A (12, 13).
Although HPR shares 70% homology with RNase A and possesses similar
key structural and catalytic residues, it displays some unique features
(14). HPR possesses substantial activity against double-stranded RNA,
contains a higher proportion of basic residues, its activity is
differentially influenced by ionic strength and divalent ions, and
compared with RNase A it has a carboxyl-terminal extension of four
residues, EDST (15-17). We have reported earlier that deletion of the
carboxyl-terminal EDST extension enhances the RNase activity and
thermostability of HPR (18). Although both RNase A and HPR catalyze RNA
degradation efficiently, they have not been associated with any special
biological action, and hence their physiological role, especially that
of HPR, is not clearly defined. RNases have much potential as
chemotherapeutics. Onconase is presently undergoing phase III human
clinical trials for the treatment of malignant mesothelemia (19) and
has also been shown to inhibit human immunodeficiency virus type I
replication in chronically infected human cells (20). A majority of
pancreatic RNases, with the exception of BS-RNase and onconase, are not
cytotoxic. The major apparent reason for this poor cytotoxicity is the
neutralization of the ribonucleolytic activity of RNases by the
cytosolic RNase inhibitor (RI). Hence, affinity of a RNase for the
intracellular RI could play an important role in defining its cytotoxic
potency. HPR holds tremendous promise as a therapeutic agent for
humans, and compared with other RNases it is likely to be less
immunogenic and thus more efficacious. When the RI-sensitive
"noncytotoxic" RNases are injected directly into Xenopus
oocytes, which lack strong inhibitors to mammalian RNases, they display
cytotoxic activity comparable to ricin and diphtheria toxin (21).
Moreover, the two classes of RNases with anticancer activity, onconase
and BS-RNase, are found to be resistant to cytosolic RI protein, and their cytotoxic activities appear to be a consequence of their abilities to escape inactivation by RI. (22, 23). BS-RNase is a
naturally occurring homodimer that is stabilized by two intersubunit disulfide bridges. The BS-RNase monomer is highly homologous to RNase
A; however, the dimeric form has a much lower affinity for RI than the
free monomer (23). Onconase evades RI as a monomer because of a lack of
amino acid residues that are responsible for making contact with RI.
Only three residues that contact RI in RNase A are conserved in
onconase (19, 24). RI is a 50-kDa protein that constitutes 0.01% of
the total cytosolic protein and is a highly conserved protein in
various mammalian species (25, 26). RI forms a 1:1 noncovalent complex
with RNase A; and as seen from the three-dimensional structure of
pRI·RNase A complex, one-third of the enzyme, including its
active site, sits within the horseshoe-shaped structure of the
inhibitor (27). A similar interaction has been observed with angiogenin
and human RI (hRI); however, the intermolecular contacts in the
RI·RNase complex differ because of the differences in the sequences
of the two RNases (28). Recently, the crystal structure of a variant of
HPR, having five amino-terminal residues replaced by those in the
BS-RNase, has been determined (29). The structure of this HPR variant
shares the overall size and characteristic V shape of the other RNases
of its family; however, it differs significantly from RNase A in
various loop regions (29).
It has been demonstrated recently that RNase A can be engineered as a
cytotoxin by mutating the specific contact residue Gly-88, which led to
a decrease in its susceptibility to RI inactivation (30). Because the
potency of a cytotoxic RNase can be defined in terms of its affinity
for the RI, it is possible that HPR could be transformed into a
cytotoxin by lowering its sensitivity to RI inactivation. In the
current study we have investigated the role of four plausible contact
residues in HPR in its interaction with the hRI with an aim to generate
cytotoxic HPR variants. We have generated variants of HPR which have
enhanced resistance toward inactivation by the hRI and are more cytotoxic.
Construction of HPR Mutants--
HPR is a protein consisting of
128 amino acid residues. pHPR, a plasmid containing the 384- base pair
HPR gene, cloned downstream of a T7 promoter (18), was used as template
to mutate the target residues Lys-7, Gln-11, Asn-71, and Glu-111 to
Ala. Similarly, the residues Asn-88, Gly-89, and Ser-90 were mutated to
arginine. Except for K7A, all of the mutations were carried out by
oligonucleotide-mediated site-directed mutagenesis (31). Uracil
containing DNA template was prepared by infecting CJ236 strain of
Escherichia coli with the recombinant phage and growing it
in the presence of uridine and chloramphenicol (31). Mutagenesis was
performed using DNA primers JKB 8, JKB 9, JKB 10, JKB 11, JKB 15, JKB
16, and JKB 29 containing the mutations G89R, N88R, K7A, Q11A, N71A,
E111A, and S90R, respectively. Sequences of various primers used are shown in Table I. Primer extension
products were transformed into E. coli strain DH5
For constructing the double mutants K7A/E111A, Q11A/E111A, and
N71A/E111A, mutants K7A, Q11A, N71A, and E111A were digested with
NheI (present at the 5'-end) and KpnI (present at
position 292 in the HPR gene), and the 290-base pair fragment released from the mutants was ligated with the NheI-KpnI
vector fragment obtained from the E111A mutant. The triple mutant
K7A/N71A/E111A was prepared by mutating the Lys-7 to Ala by polymerase
chain reaction as mentioned above, using the double mutant N71A/E111A as the template. The triple mutant Q11A/N71A/E111A was created by
oligonucleotide-mediated site-directed mutagenesis using the double
mutant N71A/E111A as the template and primer JKB 11. For the
construction of the quadruple mutant K7A/Q11A/N71A/ E111A, the triple
mutant Q11A/N71A/E111A was used as the template, and the K7A mutation
was introduced by polymerase chain reaction. All mutations were
confirmed by DNA sequencing using the dideoxy chain termination method
(32).
Expression and Purification of the Recombinant Proteins--
HPR
has been overexpressed earlier in E. coli and purified from
the inclusion bodies to obtain functionally active enzyme (18). The HPR
mutant proteins were prepared similarly from the cultures of E. coli strain BL21( Structural Characterization by Circular
Dichroism--
CD-spectra of purified proteins were recorded using a
Jasco J720 (Easton) dichrograph in the far UV region (190-250 nm).
Each protein, 33 µg/ml in 10 mM sodium phosphate, pH 7.0, was used in a cell with a 1-cm optical path to record the spectra. The spectra were acquired at a scan speed of 50 nm/min with a sensitivity of 50 mdeg and response time of 1 s. The spectra measured were an
average of 10 accumulations, and the results are presented as mean
residual ellipticity values.
Assay of Ribonucleolytic Activity of HPR and Mutants--
The
ribonucleolytic activity of various mutants was assayed on substrates
poly(C), poly(U), yeast tRNA, and poly(A·U) as described by Bond
(34). Each substrate (40 µg) was incubated separately with different
concentrations of the wild type HPR or its mutants in 100 mM Tris-HCl, pH 7.5, for 1 h at 37 °C. The
undigested large molecular weight RNA was precipitated with perchloric
acid and uranyl acetate on ice and removed by centrifugation at
15,000 × g for 10 min. The acid-soluble product was
quantitated by measuring the absorbance at 260 nm.
The hydrolytic activity of HPR and its mutants on cyclic CMP was
assayed according to the method of Crook et al. (35). In a
reaction buffer consisting of 0.2 M Tris-HCl, pH 7.5, and
0.02 M EDTA, 0.1 mg/ml cyclic CMP was mixed with 40 µg/ml
of the enzyme, and the reaction was monitored spectrophotometrically at
284 nm at 25 °C.
The RNase activity of various proteins on dinucleotide substrates CpA,
UpA, and UpG was measured by using the procedure of Witzel and Barnard
(36). The appropriate substrate, 50 µM in 100 mM Tris-HCl buffer, pH 7.0, was incubated with HPR or its mutants (final concentration 5 µM) at 25 °C. The
change in absorbance at 284 nm was monitored spectrophotometrically.
RI Binding Assays--
The HPR mutants were screened for
ribonucleolytic activity in the presence of hRI by using an agarose
gel-based assay (30). Briefly, in a total volume of 10 µl, 10 ng of
enzyme was mixed with 4 µg of total rat liver RNA and 20 units of
recombinant hRI in 100 mM Tris-HCl, pH 7.5, containing10
mM dithiothreitol. The mixture was incubated for 10 min at
37 °C; the reaction was stopped by the addition of 2 µl of gel
loading buffer containing 10 mM Tris-HCl, pH 7.5, 50 mM EDTA, glycerol (30% v/v), xylene cyanol FF (0.25%
w/v), and bromphenol blue (0.25% w/v) and subjected to electrophoresis
on a 1.5% agarose gel containing ethidium bromide.
The RNase activity of the mutants, in the presence of RI, was studied
quantitatively by assaying their activity on the most preferred RNA
homopolymer substrate, poly(C), in an assay described above (34).
The inhibition constants (Ki) for the RI-HPR mutant
interactions were determined by measuring the steady-state rate of
poly(C) cleavage in the presence of RI. Reactions were performed in 100 mM Tris-HCl, pH 7.5, containing 2.8 nM enzyme
and 50-300 µM poly(C). RI concentrations in the range
300-700 pM were used, and the initial velocity data were
used to prepare Lineweaver-Burk plots, from which Ki
was calculated.
Cytotoxicity Assays--
The cytotoxicity assays were performed
on five different cell lines: U373MG (human glioblastoma), J774A.1
(mouse monocyte-macrophage), K562 (human erythroleukemia), A431 (human
epidermoid carcinoma), and A549 (human lung carcinoma). Cytotoxicity
was evaluated by measuring [3H]leucine incorporation into
newly synthesized protein. Cells were incubated with RNases for 40 h, followed by a 3-h pulse with 0.75 µCi/well
[3H]leucine. The cells were then harvested onto glass
fiber filters using a cell harvester. The filters were dried, and
counts were taken using a liquid scintillation counter. The
ID50 values represent the concentration of the RNase which
inhibited the cellular protein synthesis by 50%.
Design of HPR Mutants
The aim of the study was to investigate whether the affinity of
HPR for RI could be reduced by mutating specific contact residues, presumably involved in the binding of HPR to RI. We selected target residues in HPR based on two criteria. First, the residue must be
involved in binding of HPR with RI by either forming a hydrogen bond or
van der Waal contact with RI, as defined by the crystal structure of
the RI·RNase complex. Second, the target residue must not be
involved in the active site of HPR.
On the basis of homology studies with the residues involved in the RI
binding of several RNases, especially RNase A, we selected four target
residues in HPR: Lys-7, Gln-11, Asn-71, and Glu-111. We replaced these
residues in HPR with alanine to yield mutants K7A, Q11A, N71A, and
E111A. Alanine was chosen because it eliminates the side chain beyond
the In RNase A mutation of Gly-88 to Arg has been shown to decrease its
sensitivity to RI inactivation and consequently increase the cytotoxic
potency by many fold (30). Gly-88 in RNase A is homologous to Gly-89 in
HPR. In the current study we individually mutated Gly-89 and also
Asn-88 and Ser-90 to Arg, yielding three single mutants, N88R, G89R,
and S90R.
Expression and Purification of HPR Mutants
The mutants were expressed in E. coli, and the
overexpressed proteins, isolated from the inclusion bodies, were
purified to homogeneity through a two-step purification scheme
comprised of cation exchange and gel filtration chromatography. The
purified HPR mutants migrated on SDS-polyacrylamide gel electrophoresis as single bands corresponding to their expected molecular weights (Fig.
1A). A polyclonal antibody
against HPR reacted with all 13 mutant proteins equally well as shown
by the Western blots (Fig. 1B). Typical final yields of the
purified recombinant proteins were in the range of 10-20 mg/liter of
culture.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
by
standard methods. The mutant K7A was constructed by polymerase chain
reaction using pHPR as the template, JKB 10 as the forward primer,
and a universal EcoRI reverse primer. The primers were
designed such that the amplified HPR fragment carrying the mutation K7A
had recognition sites for NheI at the 5'-end and
EcoRI at the 3'-end. The amplified fragment was digested with NheI and EcoRI, purified by gel
electrophoresis, and cloned into a T7 promoter based-E. coli
expression vector pVEX11 restricted with the same enzymes. pVEX11 is a
pUC-based vector that has a phage T7 promoter, multiple cloning sites,
and a T7 transcription terminator.
Sequence of primers used for mutagenesis of the putative residues
DE3), transformed with appropriate plasmid,
and grown in superbroth containing 100 µg/ml ampicillin (18). All HPR
mutants were found to accumulate in cytoplasmic inclusion bodies that
were processed further as described (33). The solubilization of the
inclusion body pellet was achieved in 6 M guanidium HCl.
Renaturation of the solubilized protein was done by diluting the
protein in a refolding buffer containing L-arginine and
oxidized glutathione. The renatured protein, after dialysis, was loaded
on an S-Sepharose cation exchange column. The protein was eluted with a
gradient of 0-1 M NaCl and purified further by gel
filtration chromatography (18).
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-carbon without altering the main conformation. We combined the
single mutations to form three double mutants, K7A/E111A, Q11A/E111A,
and N71A/E111A; two triple mutants, K7A/N71A/E111A and Q11A/N71A/E111A;
and a quadruplet mutant, K7A/Q11A/N71A/E111A.
View larger version (38K):
[in a new window]
Fig. 1.
SDS-polyacrylamide gel electrophoresis and
Western blot analysis of HPR mutants. The mutants were expressed
in BL21( DE3) cells of E. coli and purified from the
inclusion bodies by cation exchange and gel filtration chromatography.
The recombinant proteins were analyzed by 12.5% SDS-polyacrylamide gel
electrophoresis under reducing conditions followed by Coomassie Blue
staining (panel A). Western blot analysis of the mutants was
done using a polyclonal antibody raised against HPR (panel
B). The different lanes in panel B
correspond to the same proteins in panel A.
Structural Characterization of the Proteins by Circular Dichroism
Structural characterization was carried out by CD spectral
analysis to study the effect of mutations on the overall conformation of HPR. As shown in Fig. 2, the
recombinant HPR appears to be folded compactly with an +
conformation. The CD spectra of the mutant proteins K7A, Q11A, N71A,
and E111A (Fig. 2A); K7A/E111A, Q11A/E111A, and N71A/E111A
(Fig. 2B); K7A/N71A/E111A, Q11A/N71A/E111A, and
K7A/Q11A/N71A/E111A (Fig. 2C); and N88R and G89R (Fig.
2D) indicated a modest alteration; however, the overall
structure appears to be similar to that of the wild type protein. The
conformation of S90R (Fig. 2D) was found to be altered
compared with the wild type HPR, showing a significant decrease in the
-helical content of the mutant.
|
Enzymatic Activity of HPR and Its Mutants
The RNase activity of the seven single alanine mutants was assayed on three different RNA substrates: poly(C), yeast tRNA, and cyclic CMP. Pancreatic RNases have a preference for pyrimidine-rich RNA substrates. On the single-stranded, pyrimidine homopolymer substrate, poly(C), the mutants K7A, Q11A, N71A, and E111A displayed a similar or higher activity compared with wild type HPR (Table II). On yeast tRNA, the mutant K7A showed 78% activity compared with the wild type enzyme, whereas the mutants Q11A, N71A, and E111A displayed about 50% lower activity (Table II). The hydrolytic activity of HPR and its mutants was studied by spectrophotometrically monitoring the breakdown of the cyclic CMP to 3'-monophosphate. The mutants K7A, Q11A, N71A, and E111A were found to possess hydrolytic activity similar to that of the wild type HPR (Table II).
|
The mutants K7A, Q11A, N71A, and E111A exhibited activity similar to the wild type enzyme on the most favored dinucleotide substrate CpA (Table II). On UpA, the mutant K7A showed activity similar to that of HPR, but there was a significant loss in activity of the mutants Q11A, N71A, and E111A (Table II). On the dinucleotide substrate UpG, like HPR, mutants K7A and Q11A also were found to be inactive, whereas mutants N71A and E111A displayed RNase activity, with N71A being more active (Table II).
On poly(C), compared with HPR the mutant N71A/E111A was found to have a 3-fold higher activity (Table III). The activity of the mutants N88R, G89R, S90R, K7A/E111A, and K7A/N71A/E111A was found to be similar to that of the wild type HPR on poly(C), whereas the mutants Q11A/E111A, Q11A/N71A/E111A, and K7A/Q11A/N71A/E111A displayed very poor RNase activity (Table III).
|
Interaction of HPR Mutants with RI
Agarose Gel-based Assay--
The agarose gel-based assay is a
visual qualitative assay in which the extent of RNA degradation
observed in the gel is an indication of the effect of RI on HPR
activity. In the absence of RI, a progressive increase in total rat
liver RNA degradation was observed with increasing amount of HPR (Fig.
3A). The RNase shows a
preference for the 28 S rRNA. The mutants K7A, Q11A, N71A, and E111A
also displayed RNase activity similar to wild type HPR (Fig.
3A). In the presence of RI (20 units), the activity
of HPR was significantly inhibited, indicating its high sensitivity to RI (Fig. 3A). The single alanine mutants, however, were able
to degrade the RNA substrate even in the presence of RI at all enzyme concentrations used, except at a very low concentration of 0.1 ng,
implying a decreased inhibitory effect of RI on these mutants (Fig.
3A).
|
The double mutants Q11A/E111A and N71A/E111A and the triple mutant K7A/N71A/E111A also showed a higher resistance to RI, and they were able to degrade rRNA to a greater extent than HPR in the presence of inhibitor (Fig. 3B). Because the mutant N71A/E111A was enzymatically 2.5-fold more active, and Q11A/E111A 3-fold less active, in the agarose gel-based assay 2.5 times lower amount of N71A/E111A and 3 times higher amounts of Q11A/E111A were also included to equalize their enzymatic activity with HPR (Fig. 3B).
The extent of RNA degradation observed with N88R, G89R, and S90R in the absence or presence of RI was similar to that observed for HPR, implying that these residues are not crucial for HPR-RI binding (Fig. 3B).
Assay of Enzymatic Activity of the Mutants in the Presence of
RI--
The RNase activity of the mutants, in the presence of RI, was
quantitated by assaying their activity on the most preferred RNA
homopolymer substrate, poly(C) (Fig. 4).
For these studies mutants K7A, Q11A, N71A, E111A, Q11A/E111A,
N71A/E111A, and K7A/N71A/E111A displaying greater resistance to RI were
taken. As shown in Fig. 4A, at a fixed enzyme concentration
of 0.8 ng and RI concentration of 0.25 unit, HPR, K7A, and N71A showed
30% activity, whereas the mutants Q11A and E111A exhibited 60-80%
activity. However, when the enzyme concentration was increased to 1.6 ng, all the four mutants showed 100% activity even in the presence of
0.25 unit of RI, whereas HPR exhibited only 50% activity (Fig.
4B). These results demonstrate that the four mutants K7A,
Q11A, N71A, and E111A are less sensitive to RI than HPR, and among
these the mutants Q11A and E111A appear to be more resistant to RI than K7A and N71A (Fig. 4A).
|
Similarly, the mutants Q11A/E111A, N71A/E111A, and K7A/N71A/E111A exhibited 65-95% activity at an enzyme concentration of 0.8 ng and RI concentration of 0.25 unit/rxn, whereas wild type HPR showed only 30% activity (Fig. 4C). On further increasing the RI concentration to 0.5 unit/rxn, the three mutants still showed 50-70% activity compared with only 10% of HPR (Fig. 4D).
Inhibition Constants-- The inhibition constants (Ki) for the RI-HPR mutant interactions were determined by measuring the steady-state rate of poly(C) cleavage in the presence of RI. Initial velocity data were used to prepare Lineweaver-Burk plots, from which Ki was calculated. As shown in Table IV, the Ki values for the mutants were significantly higher than that of HPR. The Ki value of the triple mutant K7A/N71A/E111A was 25-fold higher than that of HPR, followed by the mutants N71A/E111A and E111A, which had an 18-fold higher value. Both Q11A and Q11A/E111A had a 10-fold higher Ki value than HPR, whereas those for K7A and N71A were 8- and 5-fold higher, respectively.
|
Cytotoxic Activity of HPR Mutants
To study the effect of mutations on the interaction of HPR with the intracellular RI, cytotoxic activity of HPR and its mutants was assayed on five different cell lines, U373MG, J774.A1, K562, A431, and A549. All mutants except Q11A/N71A/E111A, N88R, and G89R displayed a higher cytotoxic activity than the wild type HPR, as depicted in their lower ID50 values (Table V). The mutants K7A/N71A/E111A, E111A, and Q11A displayed the maximum cytotoxic activity. Out of the five cell lines used in the study, U373MG and J774A.1 were the most sensitive to these mutants. The triple mutant K7A/N71A/E111A was found to be the most potent. It exhibited at least a 10-fold higher cytotoxic activity compared with HPR on U373MG cell line, a 4-fold higher activity on J774A.1, and 2-3-fold higher activity on K562, A431, and A549. Similarly, the cytotoxic activity of the mutants E111A and Q11A varied from at least 2- to 3-fold more than that of HPR (Table V). The mutants N71A/E111A, Q11A/E111A, K7A, N71A, and S90R were found not to be as potent; however, they exhibited up to a 2-fold higher cytotoxic activity than HPR depending on the cell line. The mutants Q11A/N71A/E111A, N88R, and G89R showed cytotoxic activity similar to that of HPR on all cell lines studied (Table V).
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
An important prerequisite for a RNase to act as a cytotoxic molecule is its ability to escape inactivation by RI, present in the cytosol of mammalian cells, which functions to preserve the integrity of cellular RNA (22, 25, 37). Onconase, although a much weaker RNase than RNase A, is highly cytotoxic because it evades RI exceptionally well (19, 24). In RNase A replacing Gly-88 with arginine or aspartic acid has been shown to result in 103-104 fold higher resistance to hRI, and the mutants exhibit a potent toxic effect on K562 cells (30).
In this study, with an aim to investigate the HPR-hRI interaction and to generate a cytotoxic HPR mutant(s), we have mutated, either individually or in combination, seven residues in HPR which are presumably involved in its interaction with RI. The individual mutations of the four residues Lys-7, Gln-11, Asn-71, and Glu-111 were not detrimental to the activity of HPR on poly(C); however, these mutants had reduced activity on yeast tRNA. These residues appear to be involved in substrate binding in HPR, similar to that in RNase A. In RNase A, Lys-7 is present in the phosphate binding subsite (38-41), whereas Asn-71 and Glu-111 are present in the base binding subsite (41-43). The primary role of Gln-11, a conserved residue that donates a hydrogen bond to the reactive phosphoryl group of the bound substrate, is to prevent the nonproductive binding of the substrate (44). Using dinucleotide monophosphates as substrates, Witzel and Barnard (36) showed that the rate constant of RNase A is higher when the base at 5' position is a purine, the order being A > G > C > U (36). The mutations N71A and E111A produced a change in the substrate specificity of HPR. The four individual mutations were combined further to prepare mutants K7A/E111A, Q11A/E111A, N71A/E111A, K7A/N71A/E111A, Q11A/N71A/E111A, and K7A/Q11A/N71A/E111A. However, only K7A/E111A, N71A/E111A, and K7A/N71A/E111A had full enzymatic activity. An interesting inference from these inactive mutants is that the presence of either Gln-11 or Glu-111 in HPR appears to be absolutely essential for the full ribonucleolytic activity of the enzyme, and simultaneous mutation of both these residues is detrimental to the activity of HPR.
The mutation of Lys-7, Gln-11, Asn-71, and Glu-111 to Ala resulted in a decrease in the sensitivity of HPR to RI inactivation. There was a further augmentation in resistance to RI on combining these individual mutations, as seen in the case of the active double mutant N71A/E111A and triple mutant K7A/N71A/E111A. The triple mutant K7A/N71A/E111A showing the maximum RI resistance had a Ki value 25-fold greater than that of HPR. The greater ability of the mutants to escape RI inactivation was also reflected in their improved cytotoxic activity on a variety of cell lines. The triple mutant K7A/N71A/E111A was found to be most potent, displaying a minimum 10-fold higher cell killing ability than HPR on the glioma cell line U373MG. The double mutant Q11A/E111A, despite being 60% less active than HPR, was found to be almost 2-fold more cytotoxic than the wild type enzyme. The 10-fold higher RI resistance of the double mutant Q11A/E111A, compared with HPR, appears to be responsible for its enhanced cytotoxic activity. A similar result was observed in RNase A by Bretscher et al. (45). They found a double mutant K41R/G88R of RNase A to be enzymatically less active than the single G88R mutant but more cytotoxic. The double mutant showed a very low affinity for RI, which apparently accounts for the enhanced cytotoxicity. In contrast, another double mutant K41A/G88R of RNase A, which has the same affinity for RI but is a much weaker RNase compared with K41R/G88R mutant, is not cytotoxic. These data suggest that for a variant of RNase A to be cytotoxic, it is necessary to maintain sufficient ribonucleolytic activity (45). In the current study equivalent mutations in HPR did not produce similar results, and the mutations of Asn-88 and Gly-89 to Arg had no effect on the interaction of HPR with RI. Only the mutant S90R displayed a higher cytotoxicity than HPR. Pous et al. (29) have also reported similar results with a N88R/G89R double mutant of HPR. Our study clearly demonstrates that even though RNase A and HPR share a very close homology, with the key structural and catalytic residues identified in the bovine analog retained in the human enzyme, the observations with RNase A cannot be fully extended in HPR.
Recently the crystal structure of a variant of HPR has been determined
(29). The variant has 5 residues in the first 20 residues in its amino
terminus replaced by the equivalent residues in the BS-RNase. The
structure exhibits three helices (,
2, and
3) and seven
strands (
1-
7). Strand
1 is positioned between the helices
2 and
3, and the rest of the strands are located in sequence
after
3. Strands
3+
4 and
5+
6 run antiparallel and form a
twisted
sheet defining the V-shaped cleft where the active site is
located. The core structure of the HPR variant is very similar to that
of RNase A; however, it differs in the loop regions. The active site
cleft shows an architecture similar to that of RNase A with essential
amino acids occupying the equivalent positions. However, remarkable
differences are found at loops
4
5 (residues 90 and 91), and
2
1 (residues 37 and 38). The loop
2
3 in the HPR variant
also has a different conformation compared with that in RNase A
(residue 67). From the three-dimensional structure of pRI·RNase A and
angiogenin·hRI complexes it is apparent that the contact surface
mainly involves
2
1,
2
3, and
4
5 in RNase A and loops
2
1 and
4
5 in angiogenin, apart from the residues belonging
to their respective active sites (24, 27, 28). The loop comprising
amino acids 87-89 has been shown to be highly exposed in HPR structure
with a different conformation; accordingly, it is proposed that regions
2
1 (residues 33-43) and
2
3 (residues 64-71) might be
involved significantly in the interaction of HPR with RI (29). Our
results also support this proposal as we have observed an increased
resistance in HPR variants containing Asn-71 mutation, which lies in
the
2
3 region. Based on the crystal structures of the RI·RNase
A complex and hot spot mutagenesis in hRI, residues Lys-7 and Gln-11 of
RNase A interact with Ser-460; Asn-71 with Tyr-437; and Glu-111 with
Tyr-437, Trp-438, Ser-439, and Glu-460 of the inhibitor (46-48). In
this study with HPR we also observed increased resistance in HPR
mutants where these residues were mutated.
In conclusion, we have demonstrated that residues Lys-7, Gln-11, Asn-71
and Glu-111 in HPR are involved in its interaction with the hRI, and
mutants with these residues replaced by alanine have higher resistance
toward inactivation by RI. Further investigation of contact residues
might prove useful in developing much more potent cytotoxic variants of
HPR.
![]() |
FOOTNOTES |
---|
* This work was supported in part by grants to the National Institute of Immunology from the Department of Biotechnology, Government of India, and Grant SP/SO/D-44/96 from the Department of Science and Technology, Government of India (to J. K. B.).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.
Recipient of a senior research fellowship from the Council of
Scientific and Industrial Research, India.
§ Present address: Mouse Cancer Genetics Program, Mammalian Genetics Laboratory Bldg. 538, Rm. 133, NCI-Frederick Cancer Research and Development Center, Frederick MD 21702.
¶ To whom correspondence should be addressed. Tel.: 91-11-616-3009 or 616-2281; Fax: 91-11-616-2125 or 610-9433; E-mail: janendra@nii.res.in or jkbatra{at}yahoo.com.
Published, JBC Papers in Press, May 7, 2001, DOI 10.1074/jbc.M102440200
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: BS-RNase, bovine seminal ribonuclease; HPR, human pancreatic ribonuclease; RI, ribonuclease inhibitor; hRI, human ribonuclease inhibitor.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Beintema, J. J., Breukelman, H. J., Carsana, A., and Furia, A. (1997) in Ribonucleases: Structures and Functions (D'Alessio, G. , and Riordan, J. F., eds) , pp. 245-269, Academic Press, New York |
2. | D'Alessio, G. (1993) Trends Cell Biol. 3, 106-109[CrossRef] |
3. | Domachowske, J. B., and Rosenberg, H. F. (1997) J. Leukocyte Biol. 62, 363-368[Abstract] |
4. |
Lehrer, R. I.,
Szklarek, D.,
Barton, A.,
Ganz, T.,
Hamann, K. J.,
and Gleich, G.
(1989)
J. Immunol.
142,
4428-4434 |
5. |
Rosenberg, H. F.
(1995)
J. Biol. Chem.
270,
7876-7881 |
6. | Durack, D. T., Ackerman, S. J., Loegering, D. A., and Gleich, G. J. (1981) Proc. Natl. Acad. Sci. U. S. A. 78, 5165-5169[Abstract] |
7. | Newton, D. L., Walbridge, S., Mikulski, S. M., Ardelt, W., Shogen, K., Ackerman, S. J., Rybak, S. M., and Youle, R. J. (1994) J. Neurosci. 14, 538-544[Abstract] |
8. | Nitta, K., Ozaki, K., Ishikawa, M., Furusawa, S., Hosono, M., Kawauchi, H., Sasaki, K., Takayanagi, Y., Tsuiki, S., and Hakomori, S. (1994) Cancer Res. 54, 920-927[Abstract] |
9. | Mikulski, S. M., Grossman, A. M., Carter, P. W., Shogen, K., and Costanzi, J. J. (1993) Int. J. Oncol. 3, 57-64 |
10. | Vescia, S., Tramontano, D., Augusti-Tocco, G., and D'Alessio, G. (1980) Cancer Res. 40, 3740-3744[Abstract] |
11. | Laccetti, P., Portella, G., Mastronicola, M. R., Russo, A., Piccoli, R., D'Alessio, G., and Vecchio, G. (1992) Cancer Res. 52, 4582-4586[Abstract] |
12. | Beintema, J. J., Wietzes, P., Weickmann, J., and Glitz, J. J. (1984) Anal. Biochem. 136, 48-64[Medline] [Order article via Infotrieve] |
13. | Seno, M., Futami, J., Kosaka, M., Seno, S., and Yamada, H. (1994) Biochim. Biophys. Acta 1218, 466-468[Medline] [Order article via Infotrieve] |
14. | Weickmann, J. L., Elson, M., and Glitz, D. G. (1981) Biochemistry 20, 1272-1278[Medline] [Order article via Infotrieve] |
15. | Bardon', A., Sierakowska, H., and Shugar, D. (1976) Biochim. Biophys. Acta 438, 461-473[Medline] [Order article via Infotrieve] |
16. | Sorrentino, S., and Libonati, M. (1994) Arch. Biochem. Biophys. 312, 340-348[CrossRef][Medline] [Order article via Infotrieve] |
17. |
Sorrentino, S.,
Glitz, D. G.,
Hamann, K. J.,
Loegering, D. A.,
Checkel, J. L.,
and Gleich, G. J.
(1992)
J. Biol. Chem.
267,
14859-14865 |
18. | Bal, H. P., and Batra, J. K. (1997) Eur. J. Biochem. 245, 465-469[Abstract] |
19. |
Ardelt, W.,
Mikulski, S. M.,
and Shogen, K.
(1991)
J. Biol. Chem.
266,
245-251 |
20. |
Saxena, S. K.,
Gravell, M.,
Wu, Y.-N.,
Mikulski, S. M.,
Shogen, K.,
Ardelt, W.,
and Youle, R. J.
(1996)
J. Biol. Chem.
271,
20783-20788 |
21. |
Saxena, S. K.,
Rybak, S. M.,
Winkler, G.,
Meade, H. M.,
McGray, P.,
Youle, R. J.,
and Ackerman, E. J.
(1991)
J. Biol. Chem.
266,
21208-21214 |
22. |
Wu, Y. N.,
Mikulski, S. M.,
Ardelt, W.,
Rybak, S. M.,
and Youle, R. J.
(1993)
J. Biol. Chem.
268,
10686-10693 |
23. | Murthy, B. S., and Sirdeshmukh, R. (1992) Biochem. J. 281, 343-348[Medline] [Order article via Infotrieve] |
24. | Kobe, B., and Diesenhofer, J. (1995) Nature 374, 183-186[CrossRef][Medline] [Order article via Infotrieve] |
25. | Lee, F. S., and Vallee, B. L. (1993) Prog. Nucleic Acids Res. 44, 1-30[Medline] [Order article via Infotrieve] |
26. | Hofsteenge, J. (1997) in Ribonucleases: Structures and Functions (D'Alessio, G. , and Riordan, J. F., eds) , pp. 621-658, Academic Press, New York |
27. | Kobe, B., and Diesenhofer, J. (1996) J. Mol. Biol. 264, 1028-1043[CrossRef][Medline] [Order article via Infotrieve] |
28. |
Papageorgiou, A. C.,
Shapiro, R.,
and Acharya, K. R.
(1997)
EMBO J.
16,
5162-5177 |
29. | Pous, J., Canals, A., Terzyan, S. S., Guasch, A., Benito, A., Ribo, M., Vilanova, M., and Coll, M. (2000) J. Mol. Biol. 303, 49-59[CrossRef][Medline] [Order article via Infotrieve] |
30. |
Leland, P. A.,
Wayne Schultz, L.,
Kim, B. M.,
and Raines, R. T.
(1998)
Proc. Natl. Acad. Sci. U. S. A.
95,
10407-10412 |
31. | Kunkel, T. A., Roberts, J. D., and Zakour, R. A. (1987) Methods Enzymol. 154, 367-382[Medline] [Order article via Infotrieve] |
32. | Sanger, F., Niklen, S., and Coulson, A. R. (1977) Proc. Natl. Acad. Sci. U. S. A. 74, 5463-5467[Abstract] |
33. | Buchner, J., Pastan, I., and Brinkmann, U. (1992) Anal. Biochem. 205, 263-270[Medline] [Order article via Infotrieve] |
34. | Bond, M. D. (1988) Anal. Biochem. 173, 166-173[Medline] [Order article via Infotrieve] |
35. | Crook, E. M., Mathias, A. P., and Rabin, B. R. (1960) Biochem. J. 74, 234-238[Medline] [Order article via Infotrieve] |
36. | Witzel, H., and Barnard, E. A. (1962) Biochem. Biophys. Res. Commun. 7, 295-299[Medline] [Order article via Infotrieve] |
37. | Kawanomoto, M., Motojima, K., Sasaki, M., Hattori, H., and Goto, S. (1992) Biochim. Biophys. Acta 1129, 335-338[Medline] [Order article via Infotrieve] |
38. | Richardson, R. M., Pares, X., and Cuchillo, C. M. (1990) Biochem. J. 267, 593-599[Medline] [Order article via Infotrieve] |
39. |
Boque, L.,
Coll, M. G.,
Vilanova, M.,
Cuchillo, C. M.,
and Fita, I.
(1994)
J. Biol. Chem.
269,
19707-19712 |
40. |
Boix, E.,
Nogues, M. V.,
Schein, C. H.,
Benner, S. A.,
and Cuchillo, C. M.
(1994)
J. Biol. Chem.
269,
2529-2534 |
41. |
Fontecilla-Camps, J. C.,
de Liorens, R.,
le Du, M. H.,
and Cuchillo, C. M.
(1994)
J. Biol. Chem.
269,
21526-21531 |
42. |
Zegers, I.,
Maes, D.,
Dao-Thi, M.,
Poortmans, F.,
Palmer, R.,
and Wyns, L.
(1994)
Protein Sci.
3,
2322-2339 |
43. | Pavlovsky, A. G., Borisova, S. N., Borisov, V. V., Antonov, I. V., and Karpeisky, M. Y. (1978) FEBS Lett. 92, 258-262[CrossRef][Medline] [Order article via Infotrieve] |
44. | delCardayre, S. B., Ribo, M., Yokel, E. M., Quirk, D. J., Rutter, W. J., and Raines, R. T. (1995) Protein Eng. 8, 261-273[Abstract] |
45. |
Bretscher, L. E.,
Abel, R. L.,
and Raines, R. T.
(2000)
J. Biol. Chem.
275,
9893-9896 |
46. | Shapiro, R., Ruiz-Gutierrez, M., and Chen, C.-Z. (2000) J. Mol. Biol. 362, 487-519 |
47. |
Chen, C.-Z.,
and Shapiro, R.
(1997)
Proc. Natl. Acad. Sci. U. S. A.
96,
1761-1766 |
48. | Chen, C.-Z., and Shapiro, R. (1999) Biochemistry 38, 9273-9285[CrossRef][Medline] [Order article via Infotrieve] |