From the Department of Biochemistry, McGill University, Montreal, Quebec H3G 1Y6, Canada
Received for publication, October 16, 2000, and in revised form, February 1, 2001
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
2',3'-Cyclic nucleotide 3'-phosphodiesterase
(CNP; EC 3.1.4.37) catalyzes in vitro hydrolysis of
3'-phosphodiester bonds in 2',3'-cyclic nucleotides to produce
2'-nucleotides exclusively. N-terminal deletion mapping of the
C-terminal two-thirds of recombinant rat CNP1 identified a region that
possesses the catalytic domain, with further truncations abolishing
activity. Proteolysis and kinetic analysis indicated that this domain
forms a compact globular structure and contains all of the
catalytically essential features. Subsequently, this catalytic fragment
of CNP1 (CNP-CF) was used for chemical modification studies to identify
amino acid residues essential for activity.
5,5'-Dithiobis-(2-nitrobenzoic acid) modification studies and kinetic
analysis of cysteine CNP-CF mutants revealed the nonessential role of
cysteines for enzymatic activity. On the other hand, modification
studies with diethyl pyrocarbonate indicated that two histidines are
essential for CNPase activity. Consequently, the only two conserved
histidines, His-230 and His-309, were mutated to phenylalanine and
leucine. All four histidine mutants had kcat
values 1000-fold lower than wild-type CNP-CF, but
Km values were similar. Circular dichroism studies demonstrated that the low catalytic activities of the histidine mutants
were not due to gross changes in secondary structure. Taken together,
these results demonstrate that both histidines assume critical roles
for catalysis.
2',3'-Cyclic nucleotide 3'-phosphodiesterase
(CNP1; EC 3.1.4.37) is one of
the earliest myelin-related proteins to be expressed in differentiating
oligodendrocytes and Schwann cells (1-3). CNP is associated
exclusively with these glial cells in the nervous system, and
constitutes 4% of the total myelin proteins in the central nervous
system; it is also present at lower levels in photoreceptor cells and
several nonneural cells, notably lymphocytes (3, 4). In
oligodendrocytes, this enzyme is found throughout the cell body but is
much more abundant within the process extensions, as well as in the
outer cell periphery, in close apposition to the plasma membrane (5,
6). Although its biological function is unknown, numerous independent
studies suggest a role for this protein in migration and/or expansion
of membranes during myelination (7-12). CNP is expressed as two
isoforms with an apparent molecular mass of 46 kDa (CNP1) and 48 kDa (CNP2), differing only by the 20-amino acid extension at the
N terminus (3, 13, 14). Both isoforms are modified by isoprenylation at
the C terminus (12, 15).
One of the more intriguing aspects of CNP is the in vitro
enzymatic activity that it possesses. CNP specifically catalyzes the
irreversible hydrolysis of 2',3'-cyclic nucleotides to produce exclusively 2'-nucleotides (2, 3). In addition to 2',3'-cyclic mononucleotides, other molecules possessing this cyclic moiety, such as
2',3'-cyclic oligonucleotides and 2',3'-cyclic NADP, can also serve as
substrates in vitro (16, 17). Unlike other cyclic nucleotide
phosphodiesterases, CNP does not hydrolyze 3',5'-cyclic nucleotides (2,
3). Although physiologically relevant substrates with 2',3'-cyclic
termini have not yet been elucidated for CNP, numerous cyclic
phosphate-containing RNAs, such as U6 small nuclear RNA (18), tRNA
(19), mRNA (20), and rRNA (21), that are generated as intermediate
products from various splicing mechanisms exist within eukaryotic
cells. In addition to CNP, other enzymes that are capable of
hydrolyzing 2',3'-cyclic nucleotides to 2'-nucleotides include both
tRNA ligase (22, 23) and plant cyclic phosphodiesterase (24-26).
However, the lack of any significant sequence resemblance, as well as
differences in substrate specificity and enzymatic efficiencies
in vitro, suggest that CNP is unrelated to these enzymes.
These observations have collectively led to the speculation that the
in vitro enzymatic activity of CNP may not contribute to the
function of this protein in vivo; consequently, this aspect of CNP has largely been ignored.
Interestingly, the recently discovered protein, regeneration-induced
CNPase homolog (RICH) from goldfish (27, 28) and zebrafish (29),
possesses the same catalytic activity as CNP with similar kinetic
constants (28). Sequence comparisons between CNP and RICH revealed
substantial homologies entirely within the C-terminal two-thirds of
both proteins (27-29). The CNPase catalytic domain is located within
this region, since its sequence matched that of a 30-kDa enzymatically
active proteolytic fragment generated from elastase digestion of bovine
CNP from myelin (30). In addition, similar to CNP, RICH also contains
an isoprenylation motif at its C terminus (27-29). Although RICH is
expressed by retinal ganglion cells of the optic nerve, particularly
during nerve regeneration (29, 31), in contrast to expression of CNP by
oligodendrocytes, both cells share some biological features, such as
process extension and rapid, abundant membrane synthesis. These
similarities suggest that CNP and RICH may operate by similar
physiological mechanisms involving this enzymatic activity.
As a prerequisite in our overall strategy to fully address the
physiological relevance for CNPase activity in oligodendrocytes using
CNPase inactive mutants for cell biological studies, we sought to
identify amino acid residues critical for enzymatic activity. Although
nothing is known about the active site structure and the catalytic
mechanism of CNP to suggest candidate residues, recent site-directed
mutagenesis studies of zebrafish RICH suggested that a conserved
histidine residue in both CNP and RICH proteins was critical for
enzymatic activity (29). Furthermore, there was evidence to suggest
that cysteine(s) may be essential for activity, based on the
observation that CNP was inactivated by inorganic mercurials (2, 3).
Accordingly, in this paper, we have undertaken a study of shared motifs
that comprise part of the catalytic domains of both CNP and RICH
proteins and of specific cysteines and histidines within them,
employing chemical modification and site-directed mutagenesis. We
report here that, whereas cysteines are not essential for enzymatic
activity, two conserved histidine residues in CNP and RICH proteins,
corresponding to His-230 and His-309 in rat CNP1, are essential for catalysis.
Materials--
Pure 2',3'-cyclic NADP (cNADP) was prepared as
described by Sogin (17) using NADP (disodium salt) from Roche Molecular
Biochemicals and 1-ethyl-3-(3-dimethyl-aminopropyl)-carbodiimide-HCl
from Sigma. DEPC, DTNB, MMTS, 2'-AMP, D-glucose
6-phosphate (monosodium salt), hydroxylamine, ampicillin, and
phenylmethylsulfonyl fluoride were purchased from Sigma. Lysozyme (hen
egg white), glucose-6-phosphate dehydrogenase (yeast), pancreatic
elastase (pig), and thrombin (human plasma) were obtained from
Calbiochem. Potassium cyanide (KCN) was from Fisher, and
isopropyl- Plasmid Description--
All recombinant CNP expression vectors
were generated from the rat CNP1 and CNP2 cDNA clones (14, 32).
First, a BamHI site, 17 nucleotides upstream of the ATG
start codon of CNP1, was previously engineered by polymerase chain
reaction to generate the vector, pBS/CNP1 (12). The
BamHI-HindIII fragment derived from pBS/CNP1 was
isolated and ligated to BamHI-HindIII-linearized SK/CNP2 (14), creating the plasmid, SK/CNP (BamEco). This new CNP
plasmid contains the complete coding sequence of CNP1 with its entire
3' noncoding region.
For expression of full-length CNP1 in Escherichia coli as
GST- or His6-tagged fusion proteins, the
BamHI-EcoRI CNP1 cDNA fragment from SK/CNP
(BamEco) was subcloned in frame between the BamHI and
EcoRI sites of pGEX-3X (Amersham Pharmacia Biotech) and
pTrcHisC (InVitrogen, Carlsbad, CA) vectors.
Deletion and Site-directed Mutagenesis--
GST-tagged rat CNP1
N-terminal deletion (GST-CNP1 ND) mutants were constructed using
convenient restriction sites or polymerase chain reaction-based
strategies. Each CNP1 deletion mutant was subcloned into pGEX-3X. These
constructs encode the following residues of rat CNP1: CNP1 ND-150,
residues 150-400; CNP1 ND-164, residues 164-400; CNP1 ND-184,
residues 184-400; CNP1 ND-214, residues 214-400; and CNP1 ND-255,
residues 255-400. The structure of these plasmids was verified by
restriction and sequence analysis to ensure that the reading frame was maintained.
For chemical modification studies, a fragment of the CNP1 cDNA,
corresponding to ND-150 (the last 250 C-terminal residues), was
inserted into the pET-15b vector (Novagen, Madison, WI) to generate
pET-15b/CNP-CF. This plasmid encodes the CNP catalytic fragment
(CNP-CF) and was used to create all plasmids containing mutated
sequences of CNP-CF. The H230F, H230L, C231A, C231S, C236A, C236S,
H309F, H309L, C314A, C314S, and C397S CNP-CF mutants were created by
overlap extension polymerase chain reaction (33), using Tli
DNA polymerase (Promega, Madison, WI). The mismatched oligonucleotide
sequences used to generate the histidine mutants were as follows
(underlined sequences correspond to the mutated codon): H230F (CAC to
TTC), 5'-CCA GGC GTG CTG TTC TGT ACA ACC AAA-3' (sense) and
5'-TTT GGT TGT ACA GAA CAG CAC GCC TGG-3' (antisense); H230L (CAC to CTA), 5'-CCA GGC GTG CTG CTA TGT ACA ACC
AAA-3' (sense) and 5'-TTT GGT TGT ACA TAG CAG CAC GCC
GGT-3' (antisense); H309F (CAC to TTC), 5'-AGC CGA GCT TTC
GTC ACC CTA GGC-3' (sense) and 5'-GCC TAG GGT GAC GAA AGC
TCG GCT-3' (antisense); H309L (CAC to CTA), 5'-AGC CGA GCT
CTA GTC ACC CTA GGC-3' (sense) and 5'-GCC TAG GGT GAC
TAG AGC TCG GCT-3' (antisense). To create the cysteine mutants the primers used were as follows: C231A (TGT to GCT), 5'-GGC
GTG CTG CAC GCT ACA ACC AAA TTC-3' (sense) and 5'-GAA TTT GGT TGT AGC GTG CAG CAC GCC-3' (antisense); C231S (TGT to
AGT), 5'-GTG CTG CAC AGT ACA ACC AA-3' (sense) and 5'-TT
GGT TGT ACT GTG CAG CAC-3' (antisense); C236A (TGT to GCT),
5'-ACA ACC AAA TTC GCT GAC TAC GGG AAG-3' (sense) and
5'-CTT CCC GTA GTC AGC GAA TTT GGT TGT-3' (antisense);
C236S (TGT to AGT), 5'-ACC AAA TTC AGT GAC TAC GG-3'
(sense) and 5'-CC GTA GTC ACT GAA TTT GGT-3' (antisense);
C314A (TGT to GCT), 5'-TC ACC CTA GGC GCT GCA GCC GAC GT-3'
(sense) and 5'-AC GTC GGC TGC AGC GCC TAG GGT GA-3' (antisense); C314S (TGT to TCT), 5'-A GGC TCT GCA GCC GAC
GTG C-3' (sense) and 5'-G CAC GTC GGC TGC AGA GCC T-3'
(antisense). The CNP1 C397S mutant was generated as previously
described (12). The authenticity of the substitutions and absence of
any undesired mutations were confirmed by sequence analysis.
Protein Expression and Purification--
pGEX-3X and pTrcHisC
CNP constructs were transformed into E. coli BL21 competent
cells using standard protocols. For pET-15b CNP constructs, E. coli BL21 Gold (DE3) (Stratagene, La Jolla, CA) cells were used
for transformation. All proteins were expressed in the following
manner. A single colony was transferred to LB medium containing
100 µg/ml ampicillin and shaken overnight at 37 °C. 2× YT medium
supplemented with 100 µg/ml ampicillin was inoculated with
one-hundredth of its volume of overnight culture and grown at 37 °C
to an A600 of ~1.0. Protein expression was induced by adding 0.1 mM
isopropyl-
All operations described below were carried out at 4 °C unless
otherwise indicated. GST-tagged CNP1 N-terminal deletion (GST-CNP1 ND)
mutants were purified as follows. Cell pellet was resuspended in
one-twentieth of the culture volume of lysis buffer (phosphate-buffered saline containing 1 mM EDTA, 0.5 mg/ml lysozyme, and 0.1 mg/ml phenylmethylsulfonyl fluoride) and stirred for 30 min. DTT (5 mM) and Triton X-100 (1%) were added to the cell
suspension, followed by sonication on ice (30-s bursts, each separated
by a 1-min cooling period) until the cell suspension was no longer
viscous. The lysate was centrifuged at 14,000 × g for
30 min, and the soluble extract was incubated with one-twenty-fifth of
its volume of glutathione-Sepharose 4B for 1 h with agitation.
Resin was transferred to an empty column and washed extensively with
phosphate-buffered saline. Bound proteins were eluted by the addition
of 3× 1 bed volume of 100 mM Tris-HCl, pH 8.0, 150 mM NaCl, and 20 mM reduced glutathione.
Fractions with high protein content were pooled and dialyzed overnight
against phosphate-buffered saline. CNPase activity of full-length
GST-CNP1 and GST-CNP1 ND mutants were measured.
Purified full-length CNP1 and CNP-CF (wild type and mutants) were
obtained as follows. Cell pellet was resuspended in one-twentieth of
the culture volume of lysis buffer (50 mM sodium phosphate, pH 7.5, 500 mM NaCl, 20 mM imidazole, 0.5 mg/ml
lysozyme, and 0.1 mg/ml phenylmethylsulfonyl fluoride) and stirred for
30 min. Triton X-100 (1%) and
All purified protein preparations were analyzed by Coomassie Blue
staining after separation of proteins by SDS-polyacrylamide gel
electrophoresis. The identity of the proteins was confirmed by
appropriate Western blot analysis for GST tag, 6× His tag, or CNP.
Enzyme and Protein Assays--
CNPase activity was measured
using cNADP as substrate, according to the spectrophotometric coupled
enzyme assay procedure described previously (17). This assay measures
the rate of hydrolysis of cNADP to NADP, which is coupled to the
dehydrogenation of glucose 6-phosphate catalyzed by glucose-6-phosphate
dehydrogenase. Briefly, the assay mixture (1 ml) consisted of 100 mM MES, pH 6.0, 30 mM MgCl2, 5 mM D-glucose 6-phosphate, 5 µg of
D-glucose-6-phosphate dehydrogenase, and 2.5 mM
cNADP. After the addition of CNP variants to initiate the reaction, the
assay was carried out at 25 °C using a Beckman DU-600
spectrophotometer fitted with thermostatically controlled cuvette
holders. CNPase activity was determined by monitoring the formation of
NADPH at 340 nm ( Proteolysis of Full-length and Truncated Fragment of
CNP1--
Purified full-length CNP1 and CNP-CF were dialyzed against
50 mM sodium phosphate, pH 7.0, and 150 mM
NaCl. Final protein concentrations were adjusted to 1.0 mg/ml with the
same buffer. Both proteins (100 µg) were subjected to proteolysis by
pancreatic elastase at a substrate/protease ratio of 100:1 (w/w) at
22 °C for 18 h. Aliquots withdrawn from both protein samples,
including control samples that lacked elastase, were analyzed by
electrophoresis on 10% SDS-polyacrylamide gels. The extent of
proteolysis was assessed by comparing the gel band patterns visualized
by Coomassie Blue staining. CNP-CF proteolytic fragments (wild type and
cysteine mutants) used for DTNB modification studies were generated
from CNP-CF digestion with elastase, under the same conditions
described above, and were extensively dialyzed against 50 mM sodium phosphate, pH 7.0.
DTNB Inactivation--
CNP-CF (3 to 4 µM) in 50 mM sodium phosphate, pH 7.0, and 1 mM EDTA was
incubated with various concentrations of DTNB at 25 °C. Control
samples were incubated under the same conditions except that DTNB was
omitted. At timed intervals after the addition of DTNB, aliquots of the
reaction mixture were assayed for residual activity. Quantification of
DTNB-modified cysteines was carried out in parallel by monitoring the
time-dependent increase in the absorbance at 410 nm using
an extinction coefficient of 13,600 M MMTS Inactivation--
Modification of wild-type and cysteine
mutants CNP-CF (3 µM) with 0.25 mM MMTS were
performed in 50 mM sodium phosphate, pH 7.0, and 1 mM EDTA at 25 °C. At timed intervals, aliquots of the reaction mixture were removed, and enzymatic activity was immediately assayed.
Cyanolysis of DTNB-inactivated CNP-CF--
Wild-type and
cysteine mutants CNP-CF (3 µM) were treated with 0.2 mM DTNB under the same conditions described above, and the
inactivation was allowed to go to completion. Following exhaustive dialysis against 50 mM sodium phosphate, pH 7.0, inactivated CNP-CF was incubated with 80 mM KCN in the same
buffer or 10 mM DTT at 25 °C. Residual activities and
absorbance at 410 nm were routinely monitored until no further changes
were observed. Following this, 0.2 mM DTNB was added to the
treated samples, and its effect on CNPase activity was monitored. The
addition of KCN to native CNP-CF did not have any detrimental effects
on enzymatic activity.
DEPC Inactivation--
DEPC was freshly prepared prior to each
experiment by diluting the stock solution with anhydrous ethanol. DEPC
concentration was determined by reaction with imidazole and monitoring
increase in the absorbance at 240 nm using an extinction coefficient of 3400 M Hydroxylamine Reversal--
Hydroxylamine stock solution was
initially adjusted to pH 7.0 using NaOH. Reactivation of
DEPC-inactivated enzyme with hydroxylamine was assessed by incubating
CNP-CF (2 µM) in 100 mM sodium phosphate buffer, pH 6.5, with 1 mM DEPC for 2.5 min at 22 °C
until enzyme activity decreased to 9% of its original activity. The
reaction was rapidly quenched with 10 mM imidazole, pH 7.0. Hydroxylamine was then added to a final concentration of 0.5 M. Aliquots were removed at every half hour, and residual
enzyme activity was measured. In the control reaction using unmodified
enzyme, hydroxylamine did not affect enzyme activity.
Mapping the CNPase Catalytic Domain--
Two separate lines of
evidence indicated that the CNPase catalytic domain is located within
the C-terminal region comprising two-thirds of the polypeptide.
Previous sequence alignment analyses of CNP and RICH proteins revealed
that both proteins are highly homologous only within this region (28,
29) (Fig. 1). This also matched the amino
acid sequence of a CNPase active ~30-kDa proteolytic fragment
generated by pancreatic elastase digestion of bovine CNP1 from myelin
(30) (see Fig. 1). To map further the catalytic domain of CNP within
its primary amino acid sequence, various rat CNP1 N-terminal deletion
(ND) mutants were generated and expressed as recombinant GST fusion
proteins in E. coli (Fig. 2).
Purified full-length and mutant proteins were assayed for CNPase
activity. As expected, GST-CNP1 ND-150, which corresponds to the
C-terminal two-thirds region, exhibited activity identical to the
full-length enzyme. However, further truncations after residue 164 resulted in complete loss of activity, indicating that the conserved
subregion (residues 165-173) (Fig. 1) is required for activity and is
part of the CNPase catalytic domain.
Rationale for Using a Truncated Form of CNP1 for Chemical
Modification Studies--
Both deletion mutants, GST-CNP1 ND-150 and
ND-164, exhibited enzymatic activities identical to the full-length
protein, corroborating previous observations that the activity
exhibited by the ~30-kDa proteolytic fragment generated by elastase
digestion was similar to that prior to protease treatment (38-40). In
addition, limited proteolysis of CNP using other proteases (trypsin and
pronase) also yielded CNPase active fragments of similar sizes compared with the elastase-generated fragment (40). The fact that the C-terminal
two-thirds of CNP is enzymatically active and resistant to limited
proteolysis suggested that this region forms a tightly folded
globular structure, which contains all of the necessary molecular
components for enzymatic activity. Accordingly, the recombinantly
expressed truncated fragment of CNP could then be used for chemical
modification studies to simplify analysis. In order to address this
question, purified CNP1 ND-150 and full-length CNP1 were treated with
pancreatic elastase (1:100, w/w) for 18 h at 22 °C, and the
peptide fragments were analyzed by SDS-polyacrylamide gel
electrophoresis (Fig. 3). Complete
proteolysis was attained under these conditions, since longer
incubation did not alter the gel band patterns. Proteolysis of
full-length and truncated CNP1 both yielded identical ~30 kDa
polypeptides. The proteolytic fragment was slightly smaller than CNP1
ND-150, since extraneous residues at the C terminus were removed, based
on the amino acid sequence of elastase-generated bovine CNP1 fragment
(30) (see Fig. 1). As shown in Table I,
Km and kcat values were similar for both protein samples prior to and after elastase treatment. These results confirm that the carboxyl-terminal two-thirds of CNP is
both a catalytic and a compact structural domain. Consequently, this
truncated fragment of CNP1, herein referred to as CNP-CF (for CNP
catalytic fragment), was expressed recombinantly and was used for
chemical modification analyses.
Kinetics of CNPase Inactivation by DTNB--
It was previously
reported that incubation of CNP with inorganic mercurials resulted in
complete inactivation, implicating the potential involvement of
cysteine residue(s) in CNPase activity (16, 41-43). To determine if
these residues are important for enzymatic activity, inactivation
kinetics, using DTNB as a cysteine-modifying agent, were characterized.
DTNB inactivated CNP-CF in a time- and dose-dependent
manner, and the rate of inactivation followed pseudo-first-order
kinetics (Fig. 4A). Loss of
enzymatic activity was fully reversed by excess DTT or
Recombinant CNP-CF contains four cysteines (Cys-231, Cys-236, Cys-314,
and Cys-397), one of which (the C-terminal proximal cysteine, Cys-397)
is located outside the CNPase active proteolytic fragment generated by
elastase digestion (see Fig. 1). Furthermore, this cysteine is the site
of isoprenylation in native CNP in eukaryotes (7, 8, 12, 15); hence, it
would not assume any enzymatic role. DTNB titration under denaturing
conditions resulted in the modification of all four cysteines,
indicating that recombinant CNP-CF does not contain disulfides.
However, in the native state, only three cysteines were modified.
Correlation between the loss of enzymatic activity and the number of
cysteines modified showed that complete inactivation was observed when
all three cysteines were modified (Fig.
5A, filled
circles). Interestingly, there was no observable activity
loss after the first cysteine residue was rapidly modified (~5 s);
instead, inactivation occurred with the modification of the other two
slower reacting cysteines. To determine whether the faster reacting
cysteine is Cys-397, the elastase-generated proteolytic fragment of
CNP-CF was used for the same experiment. In this case, the faster
reacting cysteine was absent, and only two cysteines were modified when
the enzyme was completely inactivated (Fig. 5A;
open circles).
To determine whether DTNB modification is active site-directed, the
ability of 2'-AMP, a product inhibitor (45, 46), to protect against
DTNB inactivation was tested. A high concentration of 2'-AMP was used
to obtain maximum occupancy of the active site, since DTNB also acts as
a reversible and competitive inhibitor with a KI
value similar to that of 2'-AMP (KI = 500 µM). Incubation of elastase-digested CNP-CF with 2'-AMP afforded extensive protection against DTNB inactivation and cysteine modification (Fig. 5B). Rate constants for inactivation and
modification were both reduced 5-fold. The fact that one less cysteine
was modified in the presence of 2'-AMP suggested that this cysteine may
be located in or near the active site and could be essential for activity.
Site-directed Mutagenesis of Cysteines in CNP-CF and Chemical
Modification of Mutants--
To confirm results of the chemical
modification studies and to identify essential cysteine(s), Cys-231,
Cys-236, and Cys-314 were individually mutated to both serine
(stucturally similar to cysteine) and alanine (absence of hydrophilic
functional group). Recombinant CNP-CF mutant proteins were expressed,
purified, and assessed for enzymatic activity in the same manner as the
wild-type protein (Table II). Contrary to
the chemical modification results, none of the mutants exhibited
sufficient differences in its kinetic parameters compared with the
wild-type enzyme to indicate that cysteines assumed an important role
for enzymatic activity.
In an effort to identify the two DTNB-sensitive cysteines, purified
cysteine-to-serine CNP-CF mutants were digested with elastase to
generate the slightly smaller proteolytic resistant fragment for DTNB
titration experiments. Under denaturing conditions, two free sulfhydryl
groups were determined for all of the mutants (Table
III). In the native state, approximately
only one cysteine residue could be detected for the C236S and C314S
mutants, suggesting that the two DTNB-sensitive cysteines must be
Cys-314 and Cys-236, respectively. Of these two modifiable cysteines,
Cys-236 is protected from DTNB inactivation in the presence of 2'-AMP,
since only 0.3 cysteine was modified in the C314S mutant. DTNB
modification under the conditions of inactivation for wild-type CNP-CF
was carried out for C236S and C314S mutants to determine whether
one or both DTNB modified cysteines were responsible for inactivation
(Table IV). After 30 min, both mutants
were inactivated to constant levels, albeit the C236S mutant was
inactivated to a lesser extent than C314S mutant or wild-type enzyme.
This indicated that modification of either cysteine caused loss of
enzymatic activity. CNPase activity was fully restored by treatment
with DTT.
Loss of enzymatic activity by DTNB could be caused by incorporation of
a large functional group, such as TNB, at a cysteine within or near the
active site, thereby sterically hindering substrate binding and/or
catalysis. Additionally, modification may induce conformational changes
leading to the formation of a less active enzyme. Recovery of enzymatic
activity following replacement of the TNB moiety with a smaller
functional group, such as a cyanide or thiomethyl group, has previously
been used to demonstrate the nonessential role of cysteines and steric
sensitivity to DTNB modification (47-51). Reaction of DTNB-inactivated
wild-type and mutant CNP-CF with 80 mM KCN resulted in
significant recovery of activity (Table IV). Further addition of DTNB
did not cause loss of activity, indicating that all of the
surface-exposed cysteines were stable thiocyano-derivatives (data not
shown). Complete chemical modification of wild-type and mutant CNP-CF
with the smaller thiol-reacting reagent, MMTS, resulted in minor
activity losses, ranging from 20 to 42% of initial activity (Table
IV). These results demonstrate that the loss of CNPase activity is
attributable to the steric effects of DTNB modification of
Cys-236 and Cys-314.
Kinetics of CNPase Inactivation by DEPC--
It was recently
reported that histidine(s) may play an important role in CNPase
activity; mutation of a histidine in the zebrafish CNP homolog,
z-RICH, corresponding to His-309 in rat CNP1, completely abolished enzymatic activity (29). Consequently, selective chemical modification with DEPC was pursued to examine the potential involvement of histidine(s) in CNP.
CNP-CF incubated with DEPC at pH 6.5 and 22 °C resulted in a
time-dependent loss of CNPase activity (Fig.
6A). However, DEPC is unstable
in aqueous solutions, and in order to correct for its decomposition,
the inactivation data were fitted using Equation 2 (52),
Although DEPC reacts selectively with histidine residues, it can also
react with other nucleophilic amino acid residues, such as cysteine and
tyrosine, as well as with primary amino groups (37, 54, 55).
Consequently, it was necessary to rule out the possibility that
modification of a residue other than histidine causes inactivation
under the conditions used for the DEPC inactivation experiment.
Treatment of DEPC-inactivated CNP-CF with 0.5 M
hydroxylamine at pH 7.0 and 22 °C resulted in complete recovery of
CNPase activity within 1 h. This suggested that DEPC inactivation
was not due to the modification of reactive lysine, arginine, or
cysteine residues, since hydroxylamine cannot cleave carbethoxy adducts from these residues. In addition, inactivation is not attributable to
irreversible conformational changes or biscarbethoxylation of
histidines, since neither possibility can be reversed by hydroxylamine. Difference spectra of native and DEPC-treated CNP-CF revealed a peak
with an absorption maximum at 240 nm, characteristic of N-carbethoxyhistidines (data not shown). Furthermore,
O-carbethoxylation of tyrosine residues can be excluded,
since no decrease in absorbance at 278 nm was noted (56). The data
collectively prove that DEPC inactivation is solely due to the
modification of histidine residue(s). Partially inactivated CNP-CF with
10% residual activity had a Km value for cNADP
similar to that of the native enzyme. This ruled out the possibility
that the loss of activity was attributable to large increases in
Km and suggested, instead, that one or more
histidines have a catalytic role. Moreover, DEPC modification did not
cause marked secondary structural changes in the protein. CD spectra
obtained from 190 to 250 nm of native and modified CNP-CF were not
significantly different (data not shown).
To assess whether the chemical modification by DEPC is active
site-directed, 2'-AMP was used to protect the active site from inactivation (Fig. 7A). When
CNP-CF was preincubated with 2'-AMP, the rate of inactivation was
significantly reduced. This suggests that the modification of one or
more reactive histidine residue(s) at or near the active site is
responsible for loss of CNPase activity. In order to establish the
number of essential histidines, the relationship between the number of
modified histidines and loss of activity was examined (Fig.
7B, filled circles). Although a total
of two out of five histidine residues were modified per monomer,
prolonged incubation indicated that a maximum of three residues were
modified (data not shown). Since the method of extrapolating the linear
portion of the curve down to complete loss of activity to derive the
number of essential residues is rarely accurate (57, 58), we used
Tsou's statistical method to determine the number of essential
residues, using Equation 3 (57),
Site-directed Mutagenesis of Conserved Histidines in the CNPase
Catalytic Domain--
Results of the DEPC modification studies suggest
that two histidines are essential for enzymatic activity. The
C-terminal catalytic domain of rat CNP1 contains five histidines, two
of which (His-230 and His-309) are conserved in all CNP and RICH catalytic domain sequences (Fig. 1). It seemed probable, therefore, that these two histidines are essential for enzymatic activity. To test
this idea, both histidines in CNP-CF were individually mutated to
phenylalanine and leucine. Both sterically conservative residues are
hydrophobic and electrically neutral and thus are incapable of acting
as acids or bases. All four mutant enzymes were expressed in E. coli and purified by the same procedures used for the wild-type
protein. No significant differences were noticed in the expression
level of the mutants or in the final yield after its purification
compared with the wild-type enzyme, suggesting that the mutation did
not affect protein stability. In order to compare the catalytic
capabilities of mutant and wild-type CNP, kinetic parameters were
measured for the enzymes (Table II). Substitution of His-230 and
His-309 with phenylalanine and leucine almost completely eliminated
CNPase activity. Despite the low enzymatic activity detected, all
histidine mutants were amenable to kinetic analysis. Mutants exhibited
very low substrate turnover; kcat values were
roughly 1000-fold lower than those of wild-type CNP-CF. In contrast,
all mutants had slightly lower Km values than the
wild-type enzyme.
To determine whether the absence of CNPase activity in these mutants
was due to gross conformational changes resulting from single amino
acid substitutions, purified wild-type and mutant enzymes were
subjected to circular dichroism spectral analysis (data not shown). Far
UV spectral comparisons between wild-type and mutant proteins revealed
no significant secondary structural differences, indicating that the
mutation did not induce any gross conformational changes. Taken
together, these results demonstrate that both conserved residues,
His-230 and His-309, are essential for CNPase activity and may play a
role in catalysis.
Despite prior literature on CNP (reviewed in Refs. 1-3), there is
a lack of kinetic and structural information that could illuminate the
biological role of this enzymatic activity. In our strategy to address
the physiological relevance of its in vitro enzymatic
activity, we used chemical modification and site-directed mutagenesis
to identify essential residues critical for enzymatic activity, in order to ultimately generate dominant negative mutants for
future expression studies in oligodendrocytes.
Two separate lines of evidence suggested that the catalytic domain
of CNP might be located within the C-terminal two-thirds of the
protein: 1) sequence alignment of CNP and RICH showed extensive homology only within this region (28, 29), and 2) its amino acid
sequence corresponded to the enzymatically active ~30-kDa peptide
fragment generated by proteolysis (30). An updated full-length sequence
alignment of known CNP and RICH proteins, including the recently
discovered RICH protein from zebrafish (29), reveals significant
homology between both proteins, in addition to a conserved C-terminal
isoprenylation motif (Fig. 1). In an attempt to further delineate the
primary amino acid sequence of the CNPase catalytic domain, we found
that removal of the first highly conserved segment (residues 165-173)
completely abolished enzymatic activity, indicating that the C-terminal
two-thirds of CNP is the minimal C-terminal fragment that is
enzymatically active. This region of CNP appears to contain all of the
necessary molecular components for enzymatic activity, since CNP1
ND-150 and the ~30-kDa proteolytic fragment generated by elastase
digestion had Km and kcat
values similar to those of full-length CNP1. Furthermore, because this truncated fragment is resistant to proteolysis by elastase, as well as
to a lesser degree by other proteases (40), the catalytic domain of CNP
is likely to have a tightly folded, compact globular structure. We
therefore used this truncated fragment of CNP for chemical modification studies.
Based on previous reports, CNPase activity was effectively inhibited by
inorganic mercurials (2, 3). We investigated the importance of
cysteine(s) for enzymatic activity using DTNB, a sulfhydryl-specific
reagent, which allows simple and direct quantitation of modified
cysteines. Incubation of CNP-CF with DTNB resulted in complete loss of
enzymatic activity. Stoichiometric analysis of the CNP-CF elastase
proteolytic fragment suggested that two cysteines are modified, one of
which may be in or near the active site. Although these results suggest
that one or both cysteines may be essential for CNPase activity, three
observations contradict this: 1) mutation of each of the three
candidate cysteines in CNP-CF (Cys-231, Cys-236, and Cys-314) to serine
and alanine residues generated no substantial changes in kinetic
parameters as compared with the wild-type enzyme; 2) treatment of
TNB-modified wild-type and mutant CNP-CF with KCN resulted in recovery
of enzymatic activity; and 3) wild-type and mutant CNP-CF were much
less inactivated by MMTS, a smaller sulfhydryl-modifying reagent.
Stoichiometric studies of the mutants showed that Cys-236 and Cys-314
are modified by DTNB, of which Cys-236 could be located in or near the
active site. These results demonstrate that the addition of a large and bulky TNB group to the side chains of Cys-236 and Cys-314 caused conformational changes of the enzyme and/or steric obstruction of the
active site, resulting in CNPase inactivation.
We examined whether histidines assumed key roles for CNPase activity,
since mutagenic studies of z-RICH revealed that mutation of His-334 to
alanine resulted in loss of CNPase activity (29). Recombinant CNP-CF
treated with DEPC resulted in complete inactivation. The ability of
hydroxylamine to restore activity and the observed spectral changes
from DEPC modification provided evidence that histidine(s) were
specifically modified, as opposed to other nucleophilic residues such
as tyrosine, lysine, and cysteine. Complete inactivation was
concomitant with the modification of three histidines, two of which
were indicated to be essential. Protection against DEPC inactivation by
2'-AMP suggested that one or both histidines are present in or near the
active site. Based on these results, we expected two histidines to be
invariant in the CNP and RICH proteins for which the primary sequences
have been elucidated. Sequence alignment analysis shows that two
residues in rat CNP1, His-230 and His-309, meet this criterion. Not
surprisingly, this latter histidine corresponds to His-334 in z-RICH,
which further strengthens the notion that the active sites of CNP and
RICH proteins are structurally and functionally similar (see Fig. 1).
Replacement of His-230 or His-309 with phenylalanine and leucine
resulted in substantial loss of enzymatic activity, corroborating the
conclusions drawn from chemical modification studies. All histidine
mutants displayed dramatic decreases in catalytic efficiency
(kcat/Km) by about 500-fold
compared with wild-type CNP-CF, suggesting a catalytic role for both
His-230 and His-309. On the other hand, the observed loss of activity
may be due to conformational changes caused by the histidine
substitutions. However, the fact that the histidine mutants exhibited
identical parameters in Km as wild-type CNP-CF and
that the circular dichroism spectra for wild-type and mutant enzymes
were identical indicates that the loss of enzymatic activity is not
attributed to marked secondary structural changes. This conclusion is
further supported by NMR analysis (determination of three-dimensional
structure of CNP-CF by NMR is in
progress).2
Although there are no mechanistic studies available on CNP, RICH, or
other enzymes that cleave the 3'-phosphodiester bonds of 2',3'-cyclic
nucleotides to suggest specific roles for the two critical histidine
residues in CNPase activity, their function may be inferred from the
known catalytic mechanism of ribonuclease A (59). Although ribonuclease
A catalyzes hydrolysis of cyclic phosphodiester bonds of 2',3'-cyclic
phosphate RNA intermediates, this protein, unlike CNP and RICH, cleaves
the phosphate ring to generate the 3'-nucleotides as end products
instead. It is interesting to note that although there are no obvious
sequence similarities between ribonuclease A and CNP, ribonuclease A
contains two essential catalytic histidine residues: one that functions as an acid catalyst and the other as a base catalyst. Further kinetic
studies are required to investigate the catalytic roles of His-230 and
His-309 in CNPase activity. It will also be of interest to elucidate
the structure of CNP and to compare its active site structure with
those of other phosphodiesterases that cleave 2',3'-cyclic nucleotides.
A variety of observations call attention to the potential significance
of CNP in the process of myelin formation. Earlier studies reported
strong indications that CNP is associated with cytoskeletal elements of
oligodendrocytes (7, 8, 60-62). The ability of CNP, expressed in
transfected nonglial cells, to promote major alterations of cellular
morphology, with the appearance of networks of filopodia and large
processes reminiscent of those elaborated by oligodendrocytes, is
further evidence for the interaction of CNP with cellular proteins that
are normally responsible for such surface features. Moreover, we have
recently shown that CNP overexpression in transgenic mice induced
aberrant oligodendrocyte and myelin membrane formation during early
stages of oligodendrocyte differentiation and myelination, with the
appearance of redundant myelin membrane and intramyelinic vacuoles in
later stages of development (9, 10). While the function of CNP is
unknown, these observations suggest that CNP is part of a molecular
complex that regulates and/or modulates the oligodendrocyte surface
membrane expansion and migration, which lead to myelin formation.
It is intriguing that the CNPase catalytic domain of CNP and RICH
proteins is highly conserved among evolutionary divergent vertebrates.
In addition to the conserved regions, which presumably collectively
contribute to the CNPase active site, both proteins contain a
C-terminal isoprenylation motif that is conserved. Although isoprenylation has only been studied in CNP (7, 8, 12), it is probable
that this motif is also functional in RICH proteins, and the function
of both proteins is dependent on proper membrane and/or cytoskeletal
localization. Furthermore, expression of RICH and CNP occurs prior to
specific cellular events that share common characteristics, in that
both retinal ganglion cells and oligodendrocytes undergo process
extensions accompanied by membrane assembly during optic nerve
regeneration and myelin formation, respectively. Consequently, these
commonalities suggest that both proteins have similar functions that
include CNPase activity.
In conclusion, we have begun to identify specific residues that are
essential for CNPase activity. We have demonstrated, through chemical
modification studies and site-directed mutagenesis, essential roles for
His-230 and His-309 and shown that both are critical for catalysis.
These mutants will subsequently be used for more extensive kinetic
studies to determine the specific roles of the histidines for
catalysis. More importantly, the availability of these CNPase inactive
mutants now provides us with an opportunity for the first time to
evaluate the physiological relevance of this enzymatic activity. We are
currently investigating the functional consequence of
expressing these mutants as dominant negatives in oligodendrocytes.
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
-D-thiogalactopyranoside was from Diagnostic
Chemicals (Charlottetown, Prince Edward Island, Canada).
Glutathione-Sepharose 4B was supplied by Amersham Pharmacia Biotech,
while Ni2+-nitrilotriacetic acid-agarose was from Qiagen
(Valencia, CA). All other reagents were of the highest available grade.
-D-thiogalactopyranoside, and the culture was
grown for an additional 6 h at 37 °C. Cells were harvested by
centrifugation at 5000 × g for 15 min, and the cell
pellet was stored at
80 °C until ready for use.
-mercaptoethanol (10 mM)
were added to the cell suspension, followed by sonication on ice (30-s
bursts, each separated by a 1-min cooling period) until the cell
suspension was no longer viscous. The lysate was centrifuged at
14,000 × g for 30 min, and the soluble extract was
incubated with one-fiftieth of its volume of
Ni2+-nitrilotriacetic acid-agarose for 1 h with
agitation. Resin was transferred to an empty column and washed
extensively with 50 mM sodium phosphate, pH 7.5, 500 mM NaCl, and 20 mM imidazole. Bound proteins
were eluted by the addition of 7 × 1 bed volume of 50 mM sodium phosphate, pH 7.5, 500 mM NaCl, and
250 mM imidazole, and fractions with high protein content
were pooled. His6-tagged full-length CNP1 was dialyzed
against 50 mM sodium phosphate, pH 7.0, and 150 mM NaCl, before it was used for elastase proteolysis experiments and Km and Vmax
determinations. Purified wild-type and mutant CNP-CF were further
treated in the following manner. To cleave the His6 tag,
eluate from the Ni2+-nitrilotriacetic acid column was
dialyzed against thrombin cleavage buffer (10 mM Tris, pH
7.5, 500 mM NaCl, 5% (v/v) glycerol, and 2.5 mM CaCl2) in the presence of thrombin (10 units
per 1 liter of culture preparation). Following overnight cleavage, the
protein sample was passed through the Ni2+-nitrilotriacetic
acid column to capture the His6 tag. Flow-through fractions
containing cleaved CNP-CF were pooled and dialyzed overnight against
storage buffer (50 mM Tris, pH 7.5, 150 mM
NaCl, 20% (v/v) glycerol, 1 mM EDTA, and 1 mM
DTT) and stored at
80 °C.
= 6.22 mM
1
cm
1). One unit of enzyme activity is defined
as the amount of enzyme that can hydrolyze 1 µmol of cNADP/min. For
Km and Vmax determinations,
50 ng of full-length CNP1, 10 ng of elastase-treated enzymes, 10 ng of
CNP-CF wild type and cysteine mutants, and 20 µg of CNP-CF
histidine mutants were used. cNADP concentration was varied between
0.02 and 2.5 mM. Initial velocity data obtained were fitted
to the Michaelis-Menten equation using the computer program GraFit,
version 3.0 (Leatherbarrow). Molar concentration of CNP-CF (wild type
and mutants), elastase-digested CNP1 and CNP-CF, and
His6-tagged full-length CNP1 were estimated using subunit
mass values of 28, 26, and 45 kDa, respectively. For the GST-tagged
CNP1 N-terminal deletion mutants, initial rates were determined and
normalized for molar concentrations using subunit mass values of 71, 54, 52, 50, 46, and 41 kDa, for GST-CNP1 wild type, ND-150, ND-164,
ND-184, ND-214, and ND-255, respectively. Protein concentration was
estimated using both the Bio-Rad protein assay kit (based on the
dye-binding method of Bradford (34)) with bovine serum albumin as a
standard and absorbance measurement at 205 nm (35).
1 cm
1
for the released TNB chromophore (36). The recorded absorbance was
corrected for the blank (buffer and DTNB only). For protection studies,
CNP-CF was incubated at 25 °C with or without 50 mM
2'-AMP in 50 mM sodium phosphate, pH 7.0, and 1 mM EDTA, for 10 min prior to the treatment with 0.025 mM DTNB. For quantitation of free sulfhydryls under
denaturing conditions, proteins were prepared in the same buffer
containing 2% SDS.
1
cm
1 for the formation of the reaction
product, N-carbethoxyimidazole (37). For histidine
modification, CNP-CF (2 µM) in 100 mM sodium phosphate buffer, pH 6.5, was incubated with DEPC (0.5-3.1
mM) at 22 °C. The final concentration of ethanol in
reaction mixtures never exceeded 4% (v/v) of the total volume and was
shown not to have any effect on enzymatic activity. At various time
intervals, aliquots were removed, and residual CNPase activity was
measured as described above except that the enzyme assay mixture
additionally contained 20 mM imidazole to quench unreacted
DEPC. To examine substrate protection against DEPC inactivation, CNP-CF
was preincubated with or without 50 mM 2'-AMP in 100 mM sodium phosphate buffer, pH 6.5, for 10 min prior to
inactivation with DEPC. Stoichiometry of N-carbethoxylation
of histidines in CNP-CF (22 µM), treated with 0.5 mM DEPC, was determined by monitoring the
time-dependent increase in absorbance at 240 nm. The
control containing the same components but without DEPC was used to
blank the absorbance. The number of modified histidines was calculated
using an extinction coefficient of 3400 M
1 cm
1
(37). The stoichiometry of histidine modification was correlated with
enzyme activity by monitoring in a parallel experiment the time-dependent loss of activity.
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
View larger version (102K):
[in a new window]
Fig. 1.
Amino acid sequence alignment of rat CNP1
with other known CNP and RICH proteins. Multiple sequence
alignment was performed with the ClustalW program using the complete
multiple alignment protocol with default parameters. Alignment was
improved manually using the GENEDOC software. All known CNP and RICH
amino acid sequences used for the alignment are as follows: rat (14,
32), human (63, 64), bovine (30), mouse (65), chicken (66), bullfrog
(66), g-RICH68 (27), g-RICH70 (28), and z-RICH (29). Amino acid
positions are shown on the right, and dashed
lines in the sequences correspond to alignment gaps. Invariant and
variant amino acids conserved in all of the sequences are indicated
below the alignments by amino acid single-letter
codes and colons, respectively. Conserved residues
within the C-terminal two-thirds of CNP and RICH proteins are shown as
gray-shaded letters. Black-shaded letters show the histidine
and cysteine residues that were mutated in rat CNP1. Numbers
above the alignments denote the N-terminal residue of the
GST-CNP1 deletion mutants, where 1, 2,
3, 4, and 5, correspond to GST-CNP1
ND-150, ND-164, ND-184, ND-214, and ND-255, respectively. The
dashed line above the alignments, from
1 to 6, delineates the primary amino acid
sequence of the ~30-kDa proteolytic fragment generated from
pancreatic elastase digestion of bovine CNP from residue 150 to 385 (30).
View larger version (16K):
[in a new window]
Fig. 2.
Mapping the CNPase catalytic domain by
comparing relative enzymatic activities of various GST-tagged CNP1
N-terminal deletion mutants. A series of GST-tagged CNP1
N-terminal deletion mutants used to map the catalytic domain are shown
schematically. Numbers refer to the amino acid position
along the rat CNP1 primary sequence. Wild-type and deletion mutants
were expressed and purified, and their enzymatic activities were
determined as described under "Experimental Procedures." Enzymatic
activities of the deletion mutants were compared relative to wild-type
CNP1 and scaled as + or , denoting CNPase active or inactive mutants,
respectively.
View larger version (46K):
[in a new window]
Fig. 3.
Proteolysis of full-length CNP1 and CNP1
ND-150. Purified full-length CNP1 (I) and CNP1 ND-150
(II) were digested with pancreatic elastase at a mass ratio
of 1:100 of elastase/protein at room temperature. Proteolysis of both
proteins resulted in the formation of an ~30-kDa peptide
(III). Lane 1, full-length CNP1; lane
2, full-length CNP1 after 18-h digestion; lane 3, CNP1
ND-150; lane 4, CNP1 ND-150 after 18-h digestion. Four µg
of each sample were electrophoresed on a 10% SDS-polyacrylamide gel,
and the gel pattern was visualized by Coomassie Brilliant Blue.
Molecular weight marker sizes are indicated on the
left.
Kinetic parameters of full-length and truncated CNP1 prior to and after
proteolysis
-mercaptoethanol, indicating that inactivation was due to
modification of cysteine(s). Linearity of the double reciprocal plot of
pseudo-first-order rate constant (kinact)
against DTNB concentration indicated initial reversible binding of DTNB
prior to irreversible inactivation by covalent modification (Fig.
4B). Using Equation 1 (44),
View larger version (14K):
[in a new window]
Fig. 4.
Inactivation of CNP-CF by DTNB.
A, CNP-CF (3-4 µM) in 50 mM
sodium phosphate, pH 7.0, and 1 mM EDTA was incubated at
25 °C with either 0 ( ), 0.10 (
), 0.25 (
), 0.50 (
), 0.75 (
), or 1.0 (
) mM DTNB. Aliquots were withdrawn at
various time intervals for determining residual activities. Results
were plotted as logarithm of residual activity versus time,
yielding straight lines, the slopes of which represent the
pseudo-first-order rate constants of inactivation
(kinact). B, double reciprocal plot
of kinact versus DTNB concentration
from which KI and k2 values
were calculated.
where KI
(k
(Eq. 1)
1/k1) is the
dissociation constant for the noncovalent complex and
k2 is the first-order rate constant of
inactivation for the noncovalent complex, KI and
k2 were determined to be 0.350 mM
and 0.171 min
1, respectively.
View larger version (17K):
[in a new window]
Fig. 5.
Stoichiometry of DTNB-mediated inactivation
and protection by 2'-AMP. A, correlation between number
of cysteines modified by DTNB and residual activity of CNP-CF ( ) and
CNP-CF elastase proteolytic fragment (
). Both proteins (3-4
µM) were treated with 0.025 mM DTNB at
25 °C. Aliquots were removed at intervals and assayed for enzymatic
activity. The number of modified cysteines was calculated from the
differential absorbance at 410 nm in a parallel experiment under
identical conditions. B, protection of CNP-CF elastase
proteolytic fragment from DTNB inactivation and modification. CNP-CF
elastase proteolytic fragment (4 µM) was preincubated for
10 min at 25 °C in the absence (
) or presence (
) of 50 mM 2'-AMP, prior to DTNB inactivation.
Kinetic parameters of the catalytic domain mutants of CNP1
Stoichiometry of DTNB modification in wild-type and CNP-CF cysteine
mutants
Effect of KCN, DTT, and MMTS treatment on the enzymatic activities of
wild-type and CNP-CF cysteine mutants
where A/Ao is the residual activity
at time t, Io is the initial
concentration of DEPC, k1 is the
pseudo-first-order rate constant for the reaction of CNP-CF with DEPC,
and k' is the first-order rate constant for DEPC hydrolysis.
A plot of the natural log of residual activity against effective time
[(1
(Eq. 2)
e-k't/k')]
at different DEPC concentrations yielded straight lines, indicating
that inactivation followed pseudo-first-order kinetics. The
pseudo-first-order rate constant varied linearly as a function of DEPC
concentration with a second-order rate constant of 0.41 mM
1
min
1 (Fig. 6B). The linear curve
intersected at the origin, indicating that the chemical modification is
the result of a simple, irreversible bimolecular process (53).
View larger version (13K):
[in a new window]
Fig. 6.
Inactivation of CNP-CF by DEPC.
A, CNP-CF (2.0 µM) in 100 mM
sodium phosphate, pH 6.5, was incubated with either 0 ( ), 0.5 (
),
1.0 (
), 1.5 (
), or 3.1 (
) mM DEPC at 22 °C.
Samples were withdrawn at various time intervals to determine residual
activities. Inactivation data were fitted to Equation 2 to obtain the
pseudo-first-order rate constants of inactivation
(kinact). B, plot of the
concentration dependence of kinact for the
inactivation of CNPase activity.
where p represents the total number of modifiable
residues, m is the number of modified residues at a given
time point, a is the residual activity when m
residues have been modified, and i is the number of
essential residues for enzymatic activity. As shown in Fig.
7B, using p = 3, a straight line is obtained only when i = 2 (open circles),
indicating that two histidine residues are essential for CNPase
activity.
(Eq. 3)
View larger version (17K):
[in a new window]
Fig. 7.
Protection of CNP-CF against DEPC
inactivation by 2'-AMP and stoichiometry of inactivation.
A, protection by 2'-AMP against inactivation of CNP-CF by
DEPC. CNP-CF (2.0 µM), treated in the absence ( ) or in
the presence (
) of 50 mM 2'-AMP for 10 min, was
inactivated with 1.9 mM DEPC. At fixed time intervals,
aliquots were withdrawn from the mixture and assayed for CNPase
activity. B, relationship between residual activity and the
number of histidine residues modified by DEPC. CNP-CF (22 µM) was modified with 0.5 mM DEPC at 22 °C
as described under "Experimental Procedures." At various time
intervals, aliquots of the reaction mixture were withdrawn and assayed
for enzyme activity. In a parallel experiment, the increase in
absorbance at 240 nm was measured, from which the number of histidine
residues modified were calculated. The data were analyzed in the form
of a Tsou plot (57), where i = 1 (
),
i = 2 (
), and i = 3 (
).
DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
![]() |
ACKNOWLEDGEMENTS |
---|
We greatly appreciate the comments and suggestions made by Dr. J. Turnbull in the preparation of the manuscript. We also thank Vicky Kottis for expert technical assistance.
![]() |
FOOTNOTES |
---|
* This work was supported by a grant from the Medical Research Council of Canada (MRC).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 studentship from the MRC and the Fonds pour la
Formation de Chercheurs et l'Aide à la Recherche.
§ These authors contributed equally to this work.
¶ To whom correspondence should be addressed: Dept. of Biochemistry, McGill University, 3655 Promenade Sir William Osler, Montreal, Quebec H3G 1Y6, Canada. Tel.: 514-398-7281; Fax: 514-398-7384; E-mail: braun@med.mcgill.ca.
Published, JBC Papers in Press, February 5, 2001, DOI 10.1074/jbc.M009434200
2 G. Kozlov, J. Lee, M. Gravel, P. E. Braun and K. B. Gehring, unpublished data.
![]() |
ABBREVIATIONS |
---|
The abbreviations used are: CNP or CNPase, 2',3'-cyclic nucleotide 3'-phosphodiesterase; RICH, regeneration-induced CNPase homologue; GST, glutathione S-transferase; ND, N-terminal deletion; CNP-CF, CNP catalytic fragment; DEPC, diethylpyrocarbonate; DTNB, 5,5'-dithiobis-(2-nitrobenzoic acid); TNB, 5-thio-2-nitrobenzoate; cNADP, 2',3'-cyclic NADP; 2'-AMP, adenosine 2'-monophosphate; DTT, dithiothreitol; MMTS, methyl methanethiosulfonate; PMSF, phenylmethylsulfonyl fluoride; MES, 4-morpholineethanesulfonic acid; KCN, potassium cyanide.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1. | Vogel, U. S., and Thompson, R. J. (1988) J. Neurochem. 50, 1667-1677[Medline] [Order article via Infotrieve] |
2. | Sprinkle, T. J. (1989) CRC Crit. Rev. Neurobiol. 4, 235-301 |
3. | Tsukada, Y., and Kurihara, T. (1992) in Myelin: Biology and Chemistry (Martenson, R. E., ed) , pp. 449-480, CRC Press, Inc., Boca Raton, FL |
4. | Weissbarth, S., Maker, H. S., Raes, I., Brannan, T. S., Lapin, E. P., and Lehrer, G. M. (1981) J. Neurochem. 37, 677-680[Medline] [Order article via Infotrieve] |
5. | Trapp, B. D., Bernier, L., Andrews, S. B., and Colman, D. R. (1988) J. Neurochem. 51, 859-868[Medline] [Order article via Infotrieve] |
6. | Braun, P. E., Sandillon, F., Edwards, A., Matthieu, J. M., and Privat, A. (1988) J. Neurosci. 8, 3057-3066[Abstract] |
7. | De Angelis, D. A., and Braun, P. E. (1996) J. Neurochem. 67, 943-951[Medline] [Order article via Infotrieve] |
8. | De Angelis, D. A., and Braun, P. E. (1996) J. Neurochem. 66, 2523-2531[Medline] [Order article via Infotrieve] |
9. | Gravel, M., Peterson, J., Yong, V. W., Kottis, V., Trapp, B., and Braun, P. E. (1996) Mol. Cell. Neurosci. 7, 453-466[CrossRef][Medline] [Order article via Infotrieve] |
10. | Yin, X., Peterson, J., Gravel, M., Braun, P. E., and Trapp, B. D. (1997) J. Neurosci. Res. 50, 238-247[CrossRef][Medline] [Order article via Infotrieve] |
11. | Staugaitis, S. M., Bernier, L., Smith, P. R., and Colman, D. R. (1990) J. Neurosci. Res. 25, 556-560[Medline] [Order article via Infotrieve] |
12. | De Angelis, D. A., and Braun, P. E. (1994) J. Neurosci. Res. 39, 386-397[Medline] [Order article via Infotrieve] |
13. | Douglas, A. J., and Thompson, R. J. (1993) Biochem. Soc. Trans. 21, 295-297[Medline] [Order article via Infotrieve] |
14. | Gravel, M., DeAngelis, D., and Braun, P. E. (1994) J. Neurosci. Res. 38, 243-247[Medline] [Order article via Infotrieve] |
15. | Braun, P. E., De Angelis, D., Shtybel, W. W., and Bernier, L. (1991) J. Neurosci. Res. 30, 540-544[Medline] [Order article via Infotrieve] |
16. | Olafson, R. W., Drummond, G. I., and Lee, J. F. (1969) Can. J. Biochem. 47, 961-966[Medline] [Order article via Infotrieve] |
17. | Sogin, D. C. (1976) J. Neurochem. 27, 1333-1337[Medline] [Order article via Infotrieve] |
18. | Lund, E., and Dahlberg, J. E. (1992) Science 255, 327-330[Medline] [Order article via Infotrieve] |
19. | Peebles, C. L., Gegenheimer, P., and Abelson, J. (1983) Cell 32, 525-536[Medline] [Order article via Infotrieve] |
20. |
Gonzalez, T. N.,
Sidrauski, C.,
Dörfler, S.,
and Walter, P.
(1999)
EMBO J.
18,
3119-3132 |
21. | Hannon, G. J., Maroney, P. A., Branch, A., Benenfield, B. J., Robertson, H. D., and Nilsen, T. W. (1989) Mol. Cell. Biol. 9, 4422-4431[Medline] [Order article via Infotrieve] |
22. |
Phizicky, E. M.,
Schwartz, R. C.,
and Abelson, J.
(1986)
J. Biol. Chem.
261,
2978-2986 |
23. |
Pick, L.,
Furneaux, H.,
and Hurwitz, J.
(1986)
J. Biol. Chem.
261,
6694-6704 |
24. |
Tyc, K.,
Kellenberger, C.,
and Filipowicz, W.
(1987)
J. Biol. Chem.
262,
12994-13000 |
25. |
Culver, G. M.,
Consaul, S. A.,
Tycowski, K. T.,
Filipowicz, W.,
and Phizicky, E. M.
(1994)
J. Biol. Chem.
269,
24928-24934 |
26. |
Genschik, P.,
Hall, J.,
and Filipowicz, W.
(1997)
J. Biol. Chem.
272,
13211-13219 |
27. | Ballestero, R. P., Wilmot, G. R., Leski, M. L., Uhler, M. D., and Agranoff, B. W. (1995) Proc. Natl. Acad. Sci. U. S. A. 92, 8621-8625[Abstract] |
28. |
Ballestero, R. P.,
Wilmot, G. R.,
Agranoff, B. W.,
and Uhler, M. D.
(1997)
J. Biol. Chem.
272,
11479-11486 |
29. | Ballestero, R. P., Dybowski, J. A., Levy, G., Agranoff, B. W., and Uhler, M. D. (1999) J. Neurochem. 72, 1362-1371[CrossRef][Medline] [Order article via Infotrieve] |
30. |
Kurihara, T.,
Fowler, A. V.,
and Takahashi, Y.
(1987)
J. Biol. Chem.
262,
3256-3261 |
31. | Wilmot, G. R., Raymond, P. A., and Agranoff, B. W. (1993) J. Neurosci. 13, 387-401[Abstract] |
32. | Bernier, L., Alvarez, F., Norgard, E. M., Raible, D. W., Mentaberry, A., Schembri, J. G., Sabatini, D. D., and Colman, D. R. (1987) J. Neurosci. 7, 2703-2710[Abstract] |
33. | Higuchi, R. (1990) in PCR Protocols: A Guide to Methods and Applications (Innis, M. A. , Gelfand, D. H. , Sninsky, J. J. , and White, T. J., eds) , pp. 177-183, Academic Press, Inc., San Diego |
34. | Bradford, M. M. (1976) Anal. Biochem. 72, 248-254[CrossRef][Medline] [Order article via Infotrieve] |
35. | Scopes, R. K. (1987) Protein Purification Principles and Practice , 2nd Ed. , pp. 280-282, Springer-Verlag, New York |
36. | Habeeb, A. F. S. A. (1972) Methods Enzymol. 25, 457-464 |
37. | Miles, E. W. (1977) Methods Enzymol. 47, 431-442[Medline] [Order article via Infotrieve] |
38. | Nishizawa, Y., Kurihara, T., and Takahashi, Y. (1980) Biochem. J. 191, 71-82[Medline] [Order article via Infotrieve] |
39. | Kurihara, T., Nishizawa, Y., Takahashi, Y., and Odani, S. (1981) Biochem. J. 195, 153-157[Medline] [Order article via Infotrieve] |
40. | Müller, H. W., Clapshaw, P. A., and Seifert, W. (1981) J. Neurochem. 36, 2004-2012[Medline] [Order article via Infotrieve] |
41. | Sprinkle, T. J., and Knerr, J. R. (1981) Brain Res. 214, 455-459[Medline] [Order article via Infotrieve] |
42. |
Drummond, G. I.,
Iyer, N. T.,
and Keith, J.
(1962)
J. Biol. Chem.
237,
3535-3540 |
43. | Domanska-Janik, K., and Bourre, J. M. (1987) Neurotoxicology 8, 23-32[Medline] [Order article via Infotrieve] |
44. |
Kitz, R.,
and Wilson, I. B.
(1962)
J. Biol. Chem.
237,
3245-3249 |
45. | Hugli, T. E., Bustin, M., and Moore, S. (1973) Brain Res. 58, 191-203[Medline] [Order article via Infotrieve] |
46. | Starich, G. H., and Dreiling, C. E. (1980) Life Sci. 27, 567-572[Medline] [Order article via Infotrieve] |
47. |
Birchmeier, W.,
Wilson, K. J.,
and Christen, P.
(1973)
J. Biol. Chem.
248,
1751-1759 |
48. |
Degani, Y.,
Veronese, F. M.,
and Smith, E. L.
(1974)
J. Biol. Chem.
249,
7929-7935 |
49. | Fujioka, M., Takata, Y., Konishi, K., and Ogawa, H. (1987) Biochemistry 26, 5696-5702[Medline] [Order article via Infotrieve] |
50. |
Padgette, S. R.,
Huynh, Q. K.,
Aykent, S.,
Sammons, R. D.,
Sikorski, J. A.,
and Kishore, G. M.
(1988)
J. Biol. Chem.
263,
1798-1802 |
51. | Salleh, H. M., Patel, M. A., and Woodard, R. W. (1996) Biochemistry 35, 8942-8947[CrossRef][Medline] [Order article via Infotrieve] |
52. | Gomi, T., and Fujioka, M. (1983) Biochemistry 22, 137-143[Medline] [Order article via Infotrieve] |
53. | Church, F. C., Lundblad, R. L., and Noyes, C. M. (1985) J. Biol. Chem. 260, 4936-4940[Abstract] |
54. | Lundblad, R. L. (1995) Techniques in Protein Modification , pp. 110-124, CRC Press, Inc., Boca Raton, FL |
55. | Christendat, D., and Turnbull, J. (1996) Biochemistry 35, 4468-4479[CrossRef][Medline] [Order article via Infotrieve] |
56. | Melchior, W. B., Jr., and Fahrney, D. (1970) Biochemistry 9, 251-258[Medline] [Order article via Infotrieve] |
57. | Tsou, C. L. (1962) Sci. Sin. 11, 1535-1558 |
58. | Horiike, K., and McCormick, D. B. (1979) J. Theor. Biol. 79, 403-414[Medline] [Order article via Infotrieve] |
59. | Deakyne, C. A., and Allen, L. C. (1979) J. Am. Chem. Soc. 101, 3951-3959 |
60. | Dyer, C. A., and Benjamins, J. A. (1989) J. Neurosci. Res. 24, 201-211[Medline] [Order article via Infotrieve] |
61. | Wilson, R., and Brophy, P. J. (1989) J. Neurosci. Res. 22, 439-448[Medline] [Order article via Infotrieve] |
62. | Braun, P. E., Bambrick, L. L., Edwards, A. M., and Bernier, L. (1990) Ann. N. Y. Acad. Sci. 605, 55-65[Medline] [Order article via Infotrieve] |
63. | Kurihara, T., Takahashi, Y., Nishiyama, A., and Kumanishi, T. (1988) Biochem. Biophys. Res. Commun. 152, 837-842[Medline] [Order article via Infotrieve] |
64. | Douglas, A. J., Fox, M. F., Abbott, C. M., Hinks, L. J., Sharpe, G., Povey, S., and Thompson, R. J. (1992) Ann. Hum. Genet. 56, 243-254[Medline] [Order article via Infotrieve] |
65. | Monoh, K., Kurihara, T., Sakimura, K., and Takahashi, Y. (1989) Biochem. Biophys. Res. Commun. 165, 1213-1220[Medline] [Order article via Infotrieve] |
66. | Kasama-Yoshida, H., Tohyama, Y., Kurihara, T., Sakuma, M., Kojima, H., and Tamai, Y. (1997) J. Neurochem. 69, 1335-1342[Medline] [Order article via Infotrieve] |