Báo cáo khoa học: Solution structure of the catalytic domain of RICH protein from goldfish pot

Solution structure of the catalytic domain of RICH protein
from goldfish
Guennadi Kozlov, Alexey Y. Denisov, Ekaterina Pomerantseva, Michel Gravel, Peter E. Braun
and Kalle Gehring
Department of Biochemistry, McGill University, Montreal, Quebec, Canada
Axonal injuries in the mammalian central nervous sys-
tem do not cause any significant regeneration response
due to inhibitory signaling suppressing axon outgrowth
and low trophic response [1–5]. In contrast, the axons
of teleost fish regenerate upon nerve injury and have
been used as a model system to study nerve regener-
ation in the central nervous system [6]. A better under-
standing of molecular processes leading to axonal
regeneration in teleost fish could find important appli-
cations for treatment of human central nervous system
injuries.
Previous studies have identified numerous axonal
growth-associated proteins, which are induced during
nerve regeneration in teleost fish [6–9]. Regeneration-
induced CNPase homologs (RICH) proteins are axonal
growth-associated proteins that were originally discov-
ered in the studies of regenerating optical nerve in
goldfish and were termed p68 ⁄ 70 based on their appar-
ent molecular weight [10]. RICH proteins are induced
in the retinal ganglion cells during axonal regrowth
upon the optic nerve crush, and also expressed in the
germinal neuroepithelium of retina, which generates
new neurons throughout the lifespan of the fish [11].
The cloning of the RICH proteins from goldfish
(gRICH68 and gRICH70) and zebrafish (zRICH)
revealed significant homology with mammalian brain

2¢,3¢-cyclic nucleotide 3¢-phosphodiesterases (CNPases)
[12–14]. CNPases hydrolyze 2¢,3¢-cyclic nucleotides
in vitro and are abundant in oligodendrocytes and
Schwann cells [15]. Recent studies on CNPase-null
Keywords
CNPase; 2H phosphoesterase superfamily;
NMR solution structure; RICH protein; tRNA
splicing
Correspondence
K. Gehring, Department of Biochemistry,
McGill University, 3655 Promenade Sir
William Osler, Montreal, Quebec, Canada
H3G 1Y6
Fax: +1 514 3987384
Tel: +1 514 3987287
E-mail:
Website: />department/faculty/gehring/
(Received 6 December 2006, revised 16
January 2007, accepted 17 January 2007)
doi:10.1111/j.1742-4658.2007.05707.x
Regeneration-induced CNPase homolog (RICH) is an axonal growth-
associated protein, which is induced in teleost fish upon optical nerve
injury. RICH consists of a highly acidic N-terminal domain, a catalytic
domain with 2¢,3¢-cyclic nucleotide 3¢-phosphodiesterase (CNPase) activity
and a C-terminal isoprenylation site. In vitro RICH and mammalian brain
CNPase specifically catalyze the hydrolysis of 2¢,3¢-cyclic nucleotides to
produce 2¢-nucleotides, but the physiologically relevant in vivo substrate
remains unknown. Here, we report the NMR structure of the catalytic
domain of goldfish RICH and describe its binding to CNPase inhibitors.
The structure consists of a twisted nine-stranded antiparallel b-sheet sur-

rounded by a-helices on both sides. Despite significant local differences
mostly arising from a seven-residue insert in the RICH sequence, the active
site region is highly similar to that of human CNPase. Likewise, refinement
of the catalytic domain of rat CNPase using residual dipolar couplings
gave improved agreement with the published crystal structure. NMR titra-
tions of RICH with inhibitors point to a similar catalytic mechanism for
RICH and CNPase. The results suggest a functional importance for the evo-
lutionarily conserved phosphodiesterase activity and hint of a link with
pre-tRNA splicing.
Abbreviations
CNPase, 2¢,3¢-cyclic nucleotide 3¢-phosphodiesterase; RICH, regeneration-induced CNPase homolog.
1600 FEBS Journal 274 (2007) 1600–1609 ª 2007 The Authors Journal compilation ª 2007 FEBS
mutant mice revealed that the absence of CNPase cau-
ses axonal swelling and neuronal degeneration pointing
to a role for CNPase in the maintenance of the mye-
lin–axonal interface [16]. While the enzymatic activity
of CNPase has been well characterized, its physiologi-
cal substrate remains a mystery [17].
Structurally, RICH protein consists of three regions:
a glutamate- and aspartate-rich N-terminal domain, a
catalytic phosphodiesterase domain, and a C-terminal
isoprenylation site (Fig. 1). Recent studies showed that
the catalytic domain is fully sufficient for the in vitro
activity of RICH [18], as previously shown for mamma-
lian CNPase [19,20]. The catalytic domains of RICH
and CNPase share a pair of conserved sequence motifs
H-X-(T ⁄ S)-X (Fig. 1B) with three other groups of
enzymes: fungal ⁄ plant RNA ligases, bacterial RNA
ligases and fungal ⁄ plant cyclic phosphodiesterases.
Together, the catalytic domains of these proteins form a

superfamily of so-called 2H enzymes, which occur in
evolutionary kingdoms ranging from bacteria to
mammals [21]. The RNA ligases are involved in tRNA
splicing. In bacteria and archaea, they join tRNA half-
molecules containing 2¢,3¢-cyclic phosphate and
5¢-hydroxyl termini. Plant and yeast cyclic phosphodi-
esterases (CPD or CPDase) hydrolyze ADP-ribose
1¢,2¢-cyclic phosphate to yield ADP-ribose 1¢-phosphate
(at least one of these latter enzymes also hydrolyzes
nucleoside 2¢,3¢-cyclic phosphates). CPDases are also
thought to play a role in the tRNA splicing pathways.
NMR titrations with CNPase inhibitors and mutagen-
esis studies of rat CNPase [20] in combination with the
high-resolution crystal structure [22] of human CNPase
catalytic domain have been used to propose a cata-
lytic mechanism involving the catalytic H-X-(T ⁄ S)-X
motifs.
Here, we report the structure of the catalytic domain
from goldfish RICH determined by NMR. We show
that its structure and its active site are highly similar
to that of the mammalian CNPase. NMR titrations
with CNPase inhibitors identified the residues in RICH
involved in inhibitor binding and suggest the proteins
use a similar catalytic mechanism. These findings
underline the importance of the evolutionarily con-
served phosphodiesterase activity of 2H proteins and
suggest that a not yet understood link exists between
RNA metabolism and axon growth and maintenance.
RICH
H-T H-T

plant tRNA ligase
H-T H-TGIPGxAKS
T4 Pnk
H-T H-T
GCPGSGKS
mammalian CNPase
H-T H-T
GLPGSGKS
fish CNPase
H-T H-T
GLPGSGKS
yeast tRNA ligase
H-T H-TGCGKT
CNPase domain
acidic domain
A
B
isoprenylation site
Fig. 1. (A) Domain organization of RICH and
the related 2H proteins. Domains are repre-
sented by rectangles with functional motifs
added. Cyclic phosphodiesterase domains
are shown in purple. RICH and CNPase
contain a C-terminal isoprenylation motif
shown in red, domains with experimentally
confirmed polynucleotide kinase and aden-
ylyltransferase activity are in cyan and
green, respectively. The negatively charged
low-complexity N-terminal domain of RICH
is in magenta. (B) Sequence alignment of

catalytic domains of goldfish RICH
(gRICH68), human and rat CNPase (hCNP
and rCNP, respectively) and a homologous
protein from puffer fish (Gi:47207595). The
secondary structure elements refer to the
solution structure of goldfish RICH. The con-
served catalytic H-X-(T ⁄ S)-X motifs are
shown in bold.
G. Kozlov et al. Solution structure of the RICH catalytic domain
FEBS Journal 274 (2007) 1600–1609 ª 2007 The Authors Journal compilation ª 2007 FEBS 1601
Results
Structure of the RICH catalytic domain
We determined the structure of the 24 kDa catalytic
fragment of goldfish RICH protein (Fig. 2). The previ-
ously determined resonance assignments [23] were used
to assign NOEs from
15
N- and
13
C-edited 3D NOESY
experiments (Fig. S1). The 10 structures with the low-
est energy and least number of restraint violations
were chosen to represent the final ensemble (Fig. 2A).
The structural statistics are shown in Table 1.
The folded domain extends from Leu175 to Phe386
and presents its N- and C-termini together close in
space. This would position the N-terminal domain of
full-length RICH protein relatively close to its
Fig. 2. Structure of goldfish RICH catalytic domain. (A) Stereo view of the backbone superposition of 10 lowest energy structures of the
RICH catalytic domain. The superposition was carried out using regions Pro174–Glu208 and Leu233–Phe386. (B) Ribbon representation of

the RICH catalytic domain. Secondary structure elements and the N- and C-termini are labeled. (C) Backbone overlay of catalytic domains of
goldfish RICH (in cyan) and human CNPase (in purple) showing overall similarity of the structures. The lowest-energy structure from the
RICH NMR ensemble is used for the overlay. (D) The surface of the RICH catalytic domain shows several negatively charged patches of res-
idues. The catalytic site itself is not charged. Positive charges are shown in blue, negative charges are in red.
Solution structure of the RICH catalytic domain G. Kozlov et al.
1602 FEBS Journal 274 (2007) 1600–1609 ª 2007 The Authors Journal compilation ª 2007 FEBS
C-terminal isoprenylation site. This feature could be
responsible for the smaller degree of association with
the plasma membrane observed for RICH compared
with CNPase [24] as the negative charge of the N-ter-
minal domain of RICH should lead to electrostatic
repulsion with the membrane.
The catalytic domain of RICH is composed of a
highly twisted antiparallel b-sheet consisting of nine
b-strands (b1–b9) (Fig. 2B). Both sides of the b-sheet
are covered with a-helices. The twisted nature of the
b-sheet creates two extended grooves on the opposite
sides of the protein, which are occupied by the longest
helices a1 and a9. A number of short helical fragments
group together in the vicinity of the N-terminus. This
helical patch is the most basic part of the molecule
and a potential interaction surface for the preceding
acidic N-terminal domain.
Structural comparison with CNP
A structural similarity search using the Dali database
[25] showed that the best hit (Z ¼ 21.0) was the cata-
lytic domain of CNPase (PDB code 1WOJ) with an
rmsd of 2.8 A
˚
over 197 residues (Fig. 2C). The struc-

tural similarity to 2¢-5¢ RNA ligase (PDB code 1VDX)
is much weaker with an rmsd of 5.3 A
˚
over 123 resi-
dues.
As noticed previously [22], the NMR structure [20]
of the rat CNPase catalytic domain contained an erro-
neously positioned helix. This was caused by the sparse
number of NMR constraints in this part of the mole-
cule. To address this, we measured residual dipolar
couplings for the catalytic domain of rat CNPase using
the C
12
E
5
⁄ hexanol liquid crystalline medium (data not
shown). Analysis of these residual dipolar couplings
added invaluable information about this region of the
rat CNPase structure and allowed us to identify several
misassigned NOE constraints. The structure was recal-
culated with the addition of residual dipolar coupling
constraints and deposited to the RCSB database under
the accession code 2ILX (supplementary Table S1).
The corrected solution structure is in a good agree-
ment with the crystal structure of human CNPase cata-
lytic domain (rmsd of 2.3 A
˚
over 205 residues).
Sequence alignment of the catalytic domain of
RICH and related proteins (Fig. 1B) identifies the big-

gest difference between RICH and CNPase as the
seven-residue insert in the helical region between b4
and b5 strands of RICH. This insert results in addi-
tional a-helical structure in RICH comprising helices
a6 through a8 and causes this to be the most structur-
ally dissimilar region when comparing the two pro-
teins. The functional significance of this difference is
unclear but, of note, the recently identified, CNP-
related protein from the puffer fish (gi:47207595) also
contains a long 34-residue insert, on this side of the
molecule, between helices a4 and a5 (Fig. 1B).
The catalytic domains of RICH and CNPase differ
significantly in their surface charges. This changes the
overall highly positive charge of the CNPase catalytic
domain to a surface dominated by negatively charged
patches in RICH (Fig. 2D). Interestingly, the region
around the catalytic H-X-(T ⁄ S)-X motifs is relatively
neutral in both RICH and CNP. Thus, it is likely that
the overall charge difference between RICH and CNPase
is more related to protein–partner interactions and less
related to their catalytic activity on physiological
substrate(s).
We measured heteronuclear NOEs for the RICH cat-
alytic domain to identify mobile regions of the structure.
Besides the unstructured N-terminus, the most flexible
part of the protein fragment is the internal loop immedi-
ately following the helix a2 (Fig. 3). This is highly remi-
niscent of CNP, where the corresponding region in the
primary sequence, Gly208 to Lys214, produced the most
Table 1. Structural statistics for RICH protein

Restraints for structure calculations
Total restraints used 1789
Intraresidue NOEs 664
Sequential NOEs 502
Medium and long range NOEs 222
Hydrogen bonds 79
Backbone angles 322
Final energies (kcalÆmol
)1
)
E
total
190.1 ± 3.0
E
bond
7.1 ± 0.3
E
angle
91.4 ± 3.0
E
impr
9.1 ± 0.4
E
repel
39.6 ± 1.4
E
NOE
34.0 ± 2.0
E
cdih

9.0 ± 1.7
rmsd from idealized geometry
Bond (A
˚
) 0.0014 ± 0.0001
Bond angles (°) 0.30 ± 0.01
Improper torsions (°) 0.18 ± 0.01
rmsd for experimental restraints
a
Distances (A
˚
) 0.017 ± 0.001
Dihedral angles (°) 0.48 ± 0.05
Coordinate rmsd from average structure (A
˚
)
b
Backbone atoms (N,C
a
,C¢) 0.61 ± 0.05
All heavy atoms 1.15 ± 0.07
Ramachandran analysis (%)
Residues in most favored regions 84.5 ± 1.8
Residues in additional allowed regions 12.3 ± 1.6
Residues in generously allowed regions 3.0 ± 2.6
a
Calculated structures had 3–8 dihedral angle violations >2° and
three distance violations >0.2 A
˚
.

b
For residues 174 : 208 and
233 : 386.
G. Kozlov et al. Solution structure of the RICH catalytic domain
FEBS Journal 274 (2007) 1600–1609 ª 2007 The Authors Journal compilation ª 2007 FEBS 1603
intense peaks in the
15
N–
1
H heteronuclear single-quan-
tum correlation spectroscopy spectrum indicative of
backbone flexibility [20]. This region is far from the nuc-
leotide binding site (vide infra) and unlikely to play a
role in catalysis.
Binding to CNPase inhibitors
To obtain more information about the active site of
RICH, we titrated
15
N-labeled catalytic domain of
RICH with orthophosphate and the CNPase inhibitor,
adenosine-3¢-monophosphate (3¢ -AMP). The titrations
were followed by
1
H–
15
N heteronuclear single-quan-
tum correlation spectra and the shifts of amide signals
as a function of ligand addition recorded. These sig-
nals act as a fingerprint to identify amino acid residues
affected by binding and to measure the binding affinity

(Fig. 4).
Titration of the catalyti c domain of RICH with 3¢-AMP
resulted in chemical sh ift changes of roughly 20 backbone
amides, indicating binding to the protein. The biggest
chemical shift changes were o bserved for T hr322
(0.67 p.p.m.), Thr236 (0.60 p.p.m.), Asp241 (0.36 p.p.m. ),
Val332 (0.35 p.p.m.), Gly335 (0.27 p.p.m.), Phe239
(0.27 p.p.m.) and Ala319 (0.27 p.p.m.) (Fig. 5A). Thr236
and Thr322 are part of the H-X-(T ⁄ S)-X motifs, which are
essential for the catalytic activity of the related CNPase
Fig. 3. Identification of the mobile regions in the RICH catalytic
domain. (A) Plot of heteronuclear NOEs identifies the a2–a3 loop
(Gly215–Val221) as the most mobile place in the RICH catalytic
fragment. Secondary structure and the two catalytic motifs (*) are
shown. (B) Representation of flexibility in the solution structure of
the RICH catalytic domain. The width of the sausage is reversely
proportional to the heteronuclear NOE values. The figure was gen-
erated with
MOLMOL [46].
Fig. 4. NMR titration of the RICH catalytic domain with 3¢-AMP.
(A) Overlay of six heteronuclear single-quantum correlation spectra
of the
15
N-labeled RICH catalytic domain at different concentrations
of 3¢-AMP. The color changes from cyan (unliganded RICH) to dark
blue (9.3 : 1 ratio of 3¢-AMP to RICH). The most shifted amides are
labeled. (B) Determination of the dissociation constant of
4.6 ± 0.3 m
M for 3¢-AMP binding from the amide chemical shift
changes of Gly335.

Solution structure of the RICH catalytic domain G. Kozlov et al.
1604 FEBS Journal 274 (2007) 1600–1609 ª 2007 The Authors Journal compilation ª 2007 FEBS
[19,20]. Mapping of the chemical shift chan ges on the
RICH catalytic domain structure (Fig. 5B) shows that all
the changes are closely grouped in space, thereby unam-
biguously identifying this region as the catalytic site of the
protein (Fig. 6). A similar pattern of chemical shift changes
was observed for the rat CNPase catalytic domain [20],
which confirms the structural and catalytic relatedness of
the two protein s.
The binding of orthophosphate results in chemical
shift changes very similar to those observed upon bind-
ing of 3¢-AMP (Fig. 5C). As previously shown for
CNPase [20,22], phosphate binds in the active site.
Fewer residues are affected by phosphate binding,
which reflects the smaller size of the phosphate group
leading to a more local effect (Fig. 5D). Comparison
of the 3¢-AMP and orthophosphate titrations allowed
us to identify the residues of RICH affected by binding
of the adenine group (Fig. 5E). Located in the loop
between strand b2 and helix a2, Phe239 and Asp241
appear to be in a proximity of adenine base in the
RICH ⁄ 3¢-AMP complex (Fig. 5F).
A
081
022
062
003
043
0

83
0.1
0.3
0.5
0.7
Residue Number
Δ p.p.m.)(tfihs.mehc
T236
F239
D241
T322
A319
V332
G335
081
022
062
003
0
4
3
083
Residue Number
0.1
0.3
0.2
Δ p.p.m.)(tfihs.mehc
F239
D241
081

022
062
00
3
04
3
08
3
Residue Number
0.1
0.3
0.5
0.7
Δ p.p.m.)(tfihs.mehc
T236
T322
V332
G335
T236
T322
V332
T236
T322
V332
F239
D241
F
E
D
C

B
Fig. 5. Chemical shift perturbation plot of
the
15
N-labeled catalytic domain of RICH
upon titration with 3¢-AMP (A) and ortho-
phosphate (C) and mapping of the chemical
shift changes upon binding of 3¢-AMP (B)
and orthophosphate (D) on the RICH cata-
lytic domain structure. The color representa-
tion is white for no change to red for the
maximum change. (E) The difference
between chemical shift changes from titra-
tion with 3¢-AMP and orthophosphate identi-
fies orientation of 3¢-AMP when bound to
the catalytic domain of RICH. (F) Mapping
of the chemical shift changes due to the
adenine group of 3¢-AMP on the RICH
structure.
Fig. 6. Catalytic site of RICH. Histidines and threonines from the
catalytic motifs and other residues affected by inhibitor binding are
shown as sticks and labeled.
G. Kozlov et al. Solution structure of the RICH catalytic domain
FEBS Journal 274 (2007) 1600–1609 ª 2007 The Authors Journal compilation ª 2007 FEBS 1605
The NMR titration experiments also allowed us to
estimate binding affinities. Under conditions of weak
binding and fast exchange, the shifts of the signals in
heteronuclear single-quantum correlation spectra can
be used to measure the amount of inhibitor bound.
These shifts can be fitted using a simple equation,

assuming K
d
 [protein], to estimate the dissociation
constant (K
d
). The resulting values were 4.6 ± 0.3 and
12 ± 2 mm for 3¢-AMP and orthophosphate, respect-
ively, binding to RICH (Fig. 4B and Fig. S2). In com-
parison, orthophosphate binding to CNPase shows an
identical binding affinity (K
d
of 12 ± 2 mm), while
3¢-AMP binds CNPase with a much better K
d
of
0.57 ± 0.04 mm [20]. The physiological significance
of the relatively poor affinity of catalytic domain of
RICH for 3¢-AMP is unclear, since 3¢-AMP is not a
substrate of CNPase activity.
Discussion
The role of 2H proteins in myelination and nerve
growth remains mysterious. RICH shows highest struc-
tural similarity with CNPase in its catalytic domain
and catalytic site, which suggests that the conserved
2¢,3¢-cyclic phosphodiesterase activity is important for
the in vivo function of both proteins. The best-charac-
terized members of the 2H protein superfamily are
involved in RNA-processing pathways, specifically
tRNA splicing and ligation. This leads to speculation
about possible physiological substrate(s) for RICH

and CNP.
One of the mechanisms of tRNA splicing involves
endonuclease cleavage of an intron-containing tRNA
at two exon–intron borders, yielding 2¢,3¢-cyclic phos-
phates and 5¢-OH termini. Following cleavage, three
reactions are required to put the ends of fragmented
tRNA together: first, the 2¢,3¢-cyclic phosphate is
hydrolyzed by a cyclic phosphodiesterase; secondly,
the 5¢-OH terminus is phosphorylated by an NTP-
dependent polynucleotide kinase; and thirdly, the
modified ends are joined by an ATP-dependent RNA
ligase [26–31]. RNA ligases in plants and fungi con-
sist of a single polypeptide chain with three domains.
Despite the low sequence similarity, the domain
organization of plant and fungal ligases is very sim-
ilar: an N-terminal adenylyltransferase ⁄ ligase domain,
followed by a polynucleotide kinase domain and a
C-terminal cyclic phosphodiesterase domain (Fig. 1).
In multicellular animals, all three domains are still
essential, but are not necessarily in the same
polypeptide [31].
The C-terminal domains in all these ligases contain
two H-X-(T ⁄ S)-X motifs, which identify them as 2H
proteins. The central GTP-dependent polynucleotide
kinase domain of yeast ⁄ plant tRNA ligases contains
an NTP-binding P-loop consensus sequence of
GxxGxGKS that is critical for function. The sequence
of the putative P-loop in the N-terminal domain of
CNPase (
37

GLPGSGK
44
S) is strikingly similar to that
of plant tRNA ligases (GIPGSAKS for Arabidopsis
thaliana) (Fig. 1; [32]), which reinforces the connection
between CNPase and tRNA maturation. While
CNPase is missing a ligase domain, this activity could
be performed by another protein. In T4 bacteriophage,
tRNA ligation is carried out by two different enzymes.
The bifunctional enzyme T4 Pnk, which contains a
P-loop (GCPGSGKS) almost identical to CNP, pre-
pares the 3¢ and 5¢ ends of the cleaved tRNA, and T4
Rnl1 ligase reconnects the ends [33–35]. This advocates
the hypothesis that CNPase is a functional homolog
of T4 Pnk and participates in tRNA splicing and
maturation. Intriguingly, the fish homolog of CNPase
(gi:47207595) contains an additional N-terminal
domain, which could potentially possess an adenylyl-
transferase activity (Fig. 1). While showing higher
sequence homology to the fish CNPase (56% identity
versus 47% identity to human or rat CNPase), RICH
appears to lack both the adenylyltransferase and kin-
ase domains; little is known about the function of its
acidic N-terminal segment.
The cellular localization of CNPase does not contra-
dict its involvement in pre-tRNA splicing. Recent stud-
ies revealed that the yeast tRNA splicing endonuclease
mainly localizes on mitochondria and this localization
is important for its function [36]. Interestingly, one
CNPase isoform (CNP2) is specifically targeted to

mitochondria [37]. More intriguingly, RICH and
CNPase may be involved in other RNA splicing
events. XBP1 mRNA, in humans, and HAC1 mRNA,
in yeast, undergo cytoplasmic splicing as part of the
unfolded protein response that regulates the endoplas-
mic reticulum volume and protein composition [38].
While no 2¢,3¢-cyclic phosphate intermediates have
been identified in these reactions or in regulation by
micro RNAs, it is not impossible that the evolutionari-
ly ancient phosphoesterase activity of 2H proteins is
involved in regulating membrane biogenesis in oligo-
dendrocytes or neurons via RNA.
In conclusion, RICH proteins have been less char-
acterized than mammalian CNPases, but the strong
structural similarity of these proteins suggests a sim-
ilar function. The structure of RICH and nucleotide
binding studies presented here represent another step
towards understanding of CNP ⁄ RICH function and
suggest new avenues to study these still enigmatic
proteins.
Solution structure of the RICH catalytic domain G. Kozlov et al.
1606 FEBS Journal 274 (2007) 1600–1609 ª 2007 The Authors Journal compilation ª 2007 FEBS
Experimental procedures
Protein expression and purification
The catalytic domain of goldfish gRICH68 protein (residues
172–389) was subcloned into pET15b (Novagen, Inc.,
Madison, WI, USA) and expressed in the Escherichia coli
expression host BL21 (DE3) (Stratagene, La Jolla, CA,
USA) as a His-tagged fusion protein. The protein was puri-
fied by immobilized metal affinity chromatography on

Ni
2+
-loaded chelating sepharose column (Amersham Phar-
macia Biotech, Piscataway, NJ, USA). Isotopically labeled
RICH was prepared from cells grown on minimal M9
media containing
15
N ammonium chloride and
13
C glucose
(Cambridge Isotopes Laboratory, Andover, MA, USA).
The N-terminal His-tag was cleaved from RICH by over-
night dialysis with thrombin (Amersham Pharmacia Bio-
tech, Piscataway, NJ, USA) at 1 unit per mg fusion protein
at room temperature. Benzamidine sepharose and Ni
2+
-loa-
ded chelating sepharose were used to remove thrombin and
the His-tag peptide from RICH. The resulting 222 amino
acid protein contained four N-terminal extraneous residues
(GSHM) from the His tag. The sequence composition of
purified RICH was confirmed by mass spectrometry.
NMR spectroscopy
NMR resonance assignments of the catalytic domain of
RICH were determined previously [23]. All NMR experi-
ments were recorded at 307 K. NMR samples were 1 mm
protein in 50 mm 4-morpholineethanesulfonic acid buffer,
0.15 m NaCl, 1 mm dithiothreitol at pH 6.0. NMR spectra
were processed with nmrpipe [39] and xwinnmr software
version 3 (Bruker Biospin) and analyzed with xeasy [40].

For titrations, 3¢-AMP and Na
2
HPO
4
were purchased
from Sigma (Saint-Louis, MO, USA) and used without any
additional purification. Titrations were monitored by
15
N-
1
H
heteronuclear single-quantum correlation spectra following
addition of inhibitors to
15
N-labeled RICH (172–389) on a
Bruker DRX 600 MHz spectrometer. The experiments were
recorded with 128 increments using 4–8 scans and lasted for
10–20 min. Chemical shift changes were calculated as
(DHN
2
+ (0.2*DN)
2
)
1 ⁄ 2
in p.p.m. Samples contained 50 mm
4-morpholineethanesulfonic acid, 0.15 m NaCl and 1 mm di-
thiothreitol at pH 6.0 and 0.4–0.5 mm RICH at 307 K. The
inhibitor concentrations ranged from 0.1 to 60 mm depend-
ing on the affinity and solubility of the inhibitor. The hetero-
nuclear single-quantum correlation spectra of complexes

were assigned by monitoring chemical shift changes upon
addition of the substrate, since the binding takes place in the
fast exchange. The pH of the NMR samples was monitored
during the titrations and adjusted as needed. Chemical shift
changes for individual residues were fit to a one-site binding
equation: d ¼ d
max
Æ [L] ⁄ (K
d
+ [L]) where d is the chemical
shift change, [L] is the total ligand concentration (uncor-
rected for binding to RICH), and K
d
is the dissociation con-
stant. The fitting was carried out using the computer
program grafit, version 3.0 (Ericathus Software, Horley,
UK) to determine d
max
and the K
d
of binding.
Structure calculation
NOESY constraints for the structure determination were
obtained from
15
N-edited NOESY (mixing time 80 ms) and
13
C-edited NOESY (mixing time 80 ms) 3D experiments
acquired on a Varian Unity Inova 800 MHz spectrometer
at the Quebec-Eastern Canada High-Field NMR Facility.

For the structure determination, a set of ARIA-assigned
[41] and manually verified 1388 NOEs were collected from
15
N- and
13
C-edited NOESY spectra of RICH (172–389).
Three hundred and twenty-two backbone angles resulted
from chemical shift index using the TALOS database [42].
Hydrogen bonds were predicted from NOE analysis. The
starting structure was generated with modeller 6v2 [43]
using human CNPase crystal structure (PDB code 1WOJ)
and was in agreement with manually assigned NOEs. One
hundred and fifty structures were calculated and refined
using standard protocols in cns v.1.1 [44]. procheck-nmr
[45] was used to check the protein stereochemical geometry.
The structural statistics for 10 structures are shown in
Table 1. The coordinates have been deposited in the
Research Collaboratory for Structural Bioinformatics
(RCSB) under PDB accession code 2I3E and the NMR
assignments under BMRB accession number 7167.
Acknowledgements
We acknowledge Dr. M. D. Uhler (University of
Michigan, USA) for a gift of gRICH68 cDNA. We
thank T. Sprules for help in running NMR experi-
ments at the Quebec-Eastern Canada High-Field
NMR Facility. This work was funded by operating
grant MOP-43967 to KG and PB from the Canadian
Institutes of Health Research.
References
1 Bahr M & Bonhoeffer F (1994) Perspectives on axonal

regeneration in the mammalian CNS. Trends Neurosci
17, 473–479.
2 Fawcett JW & Asher RA (1999) The glial scar and central
nervous system repair. Brain Res Bull 49, 377–391.
3 Horner PJ & Gage FH (2000) Regenerating the
damaged central nervous system. Nature 407, 963–970.
4 Spencer T, Domeniconi M, Cao Z & Filbin MT (2003)
New roles for old proteins in adult CNS axonal regen-
eration. Curr Opin Neurobiol 13, 133–139.
5 Schwab ME (2004) Nogo and axon regeneration. Curr
Opin Neurobiol 14, 118–124.
G. Kozlov et al. Solution structure of the RICH catalytic domain
FEBS Journal 274 (2007) 1600–1609 ª 2007 The Authors Journal compilation ª 2007 FEBS 1607
6 Bernhardt RR (1999) Cellular and molecular bases of
axonal regeneration in the fish central nervous system.
Exp Neurol 157, 223–240.
7 Skene JH (1989) Axonal growth-associated proteins.
Annu Rev Neurosci 12, 127–156.
8 Caroni P (1997) Intrinsic neuronal determinants that
promote axonal sprouting and elongation. Bioessays 19 ,
767–775.
9 Plunet W, Kwon BK & Tetzlaff W (2002) Promoting
axonal regeneration in the central nervous system by
enhancing the cell body response to axotomy. J Neu-
rosci Res 68, 1–6.
10 Heacock AM & Agranoff BW (1982) Protein synthesis
and transport in the regenerating goldfish visual system.
Neurochem Res 7 , 771–788.
11 Wilmot GR, Raymond PA & Agranoff BW (1993) The
expression of the protein p68 ⁄ 70 within the goldfish vis-

ual system suggests a role in both regeneration and
neurogenesis. J Neurosci 13, 387–401.
12 Ballestero RP, Wilmot GR, Leski ML, Uhler MD &
Agranoff BW (1995) Isolation of cDNA clones encoding
RICH: a protein induced during goldfish optic nerve
regeneration with homology to mammalian 2¢,3¢-cyclic-
nucleotide 3¢-phosphodiesterases. Proc Natl Acad Sci
USA 92, 8621–8625.
13 Ballestero RP, Wilmot GR, Agranoff BW & Uhler MD
(1997) gRICH68 and gRICH70 are 2¢,3¢-cyclic-nucleo-
tide 3¢-phosphodiesterases induced during goldfish optic
nerve regeneration. J Biol Chem 272, 11479–11486.
14 Ballestero RP, Dybowski JA, Levy G, Agranoff BW &
Uhler MD (1999) Cloning and characterization of
zRICH, a 2¢,3¢-cyclic-nucleotide 3¢-phosphodiesterase
induced during zebrafish optic nerve regeneration.
J Neurochem 72, 1362–1371.
15 Sprinkle TJ (1989) 2¢,3¢-Cyclic nucleotide 3¢-phosphodi-
esterase, an oligodendrocyte-Schwann cell and
myelin-associated enzyme of the nervous system.
Crit Rev Neurobiol 4, 235–301.
16 Lappe-Siefke C, Goebbels S, Gravel M, Nicksch E, Lee
J, Braun PE, Griffiths IR & Nave KA (2003) Disrup-
tion of Cnp1 uncouples oligodendroglial functions in
axonal support and myelination. Nat Genet 33, 366–374.
17 Braun PE, Lee J & Gravel M (2004) 2¢,3¢-Cyclic nucleo-
tide 3¢-phosphodiesterase: structure, biology and func-
tion. In Myelin Biology and Disorders, Vol. 1 (Lazzerini,
RA, ed.), pp. 499–521. Elsevier, Academic Press, New
York.

18 Challa M, Chapa GR, Govindaraju S, Gonza
´
lez-Garcı
´
a
M & Ballestero RP (2006) Characterization of the
domains of zRICH, a protein induced during optic
nerve regeneration in zebrafish. Brain Res 1100, 42–54.
19 Lee J, Gravel M, Gao E, O’Neill RC & Braun PE
(2001) Identification of essential residues in 2¢,3¢-cyclic
nucleotide 3¢-phosphodiesterase. Chemical modification
and site-directed mutagenesis to investigate the role of
cysteine and histidine residues in enzymatic activity.
J Biol Chem 276, 14804–14813.
20 Kozlov G, Lee J, Elias D, Gravel M, Gutierrez P, Ekiel
I, Braun PE & Gehring K (2003) Structural evidence
that brain cyclic nucleotide phosphodiesterase is a mem-
ber of the 2H phosphodiesterase superfamily. J Biol
Chem 278, 46021–46028.
21 Mazumder R, Iyer LM, Vasudevan S & Aravind L
(2002) Detection of novel members, structure-function
analysis and evolutionary classification of the 2H
phosphoesterase superfamily. Nucleic Acids Res 30,
5229–5243.
22 Sakamoto Y, Tanaka N, Ichimiya T, Kurihara T &
Nakamura KT (2005) Crystal structure of the catalytic
fragment of human brain 2¢,3¢-cyclic-nucleotide 3¢-phos-
phodiesterase. J Mol Biol 346, 789–800.
23 Denisov AY, Kozlov G, Gravel M, Sprules T, Braun
PE & Gehring K (2006)

1
H,
13
C and
15
N resonance
assignments of the catalytic domain of the goldfish
RICH protein. J Biomol NMR 36 (Suppl. 5), 75.
24 Leski ML & Agranoff BW (1994) Purification and char-
acterization of p68 ⁄ 70, regeneration-associated proteins
from goldfish brain. J Neurochem 62, 1182–1191.
25 Holm L & Sander C (1995) Dali: a network tool for
protein structure comparison. Trends Biochem Sci 20,
478–480.
26 Abelson J, Trotta CR & Li H (1998) tRNA splicing.
J Biol Chem 273, 12685–12688.
27 Greer CL, Peebles CL, Gegenheimer P & Abelson J
(1983) Mechanism of action of a yeast RNA ligase in
tRNA splicing. Cell 32, 537–546.
28 Apostol BL, Westaway SK, Abelson J & Greer CL
(1991) Deletion analysis of a multifunctional yeast
tRNA ligase polypeptide: identification of essential and
dispensable functional domains. J Biol Chem 266, 7445–
7455.
29 Phizicky EM, Consaul SA, Nehrke KW & Abelson J
(1992) Yeast tRNA ligase mutants are nonviable and
accumulate tRNA splicing intermediates. J Biol Chem
267, 4577–4582.
30 Westaway SK & Abelson J (1995) Splicing of tRNA
precursors. In Trna: Structure, Biosynthesis and Function

(Soll, D & RajBhandary, UL, eds), pp. 79–92 American.
Society for Microbiology, Washington, DC.
31 Sawaya R, Schwer B & Shuman S (2003) Genetic and
biochemical analysis of the functional domains of yeast
tRNA ligase. J Biol Chem 278, 43928–43938.
32 Koonin EV & Gorbalenya AE (1990) Related domains
in yeast tRNA ligase, bacteriophage T4 polynucleotide
kinase and RNA ligase, and mammalian myelin
2¢,3¢-cyclic nucleotide phosphohydrolase revealed by
amino acid sequence comparison. FEBS Lett 268, 231–
234.
33 Amitsur M, Levitz R & Kaufmann G (1987) Bacter-
iophage T4 anticodon nuclease, polynucleotide kinase
Solution structure of the RICH catalytic domain G. Kozlov et al.
1608 FEBS Journal 274 (2007) 1600–1609 ª 2007 The Authors Journal compilation ª 2007 FEBS
and RNA ligase reprocess the host lysine tRNA. EMBO
J 6, 2499–2503.
34 Cameron V & Uhlenbeck OC (1977) 3¢-Phosphatase
activity in T4 polynucleotide kinase. Biochemistry 16,
5120–5126.
35 Galburt EA, Pelletier J, Wilson G & Stoddard BL
(2002) Structure of a tRNA repair enzyme and molecu-
lar biology workhorse: T4 polynucleotide kinase. Struc-
ture (Camb) 10, 1249–1260.
36 Yoshihisa T, Yunoki-Esaki K, Ohshima C, Tanaka N
& Endo T (2003) Possibility of cytoplasmic pre-tRNA
splicing: the yeast tRNA splicing endonuclease mainly
localizes on the mitochondria. Mol Biol Cell 14, 3266–
3279.
37 Lee J, O’Neill RC, Park MW, Gravel M & Braun PE

(2006) Mitochondrial localization of CNP2 is regulated
by phosphorylation of the N-terminal targeting signal
by PKC: implications of a mitochondrial function for
CNP2 in glial and non-glial cells. Mol Cell Neurosci 31,
446–462.
38 Bernales S, Papa FR & Walter P (2006) Intracellular
signaling by the unfolded protein response. Annu Rev
Cell Dev Biol 22, 487–508.
39 Delaglio F, Grzesiek S, Vuister GW, Zhu G, Pfeifer J &
Bax A (1995) NMRPipe: a multidimensional spectral
processing system based on UNIX pipes. J Biomol
NMR 6, 277–293.
40 Bartels C, Xia T-H, Billeter M, Gu
¨
ntert P & Wu
¨
thrich
K (1996) The program XEASY for computer-supported
NMR spectral analysis of biological macromolecules.
J Biomol NMR 6, 1–10.
41 Nilges M, Macias MJ, O’Donoghue SI & Oschkinat H
(1997) Automated NOESY interpretation with ambigu-
ous distance restraints: the refined NMR solution
structure of the pleckstrin homology domain from
beta-spectrin. J Mol Biol 269, 408–422.
42 Cornilescu G, Delaglio F & Bax A (1999) Protein back-
bone angle restraints from searching a database for che-
mical shift and sequence homology. J Biomol NMR 13,
289–302.
43 Sali A & Blundell TL (1993) Comparative protein mod-

elling by satisfaction of spatial restraints. J Mol Biol
234, 779–815.
44 Bru
¨
nger AT et al. (1998) Crystallography & NMR sys-
tem: a new software suite for macromolecular structure
determination. Acta Crystallogr D Biol Crystallogr 54,
905–921.
45 Laskowski RA, Rullmannn JA, MacArthur MW,
Kaptein R & Thornton JM (1996) AQUA and PRO-
CHECK-NMR: programs for checking the quality of
protein structures solved by NMR. J Biomol NMR 8,
477–486.
46 Koradi R, Billeter M & Wu
¨
thrich K (1996) MOLMOL:
a program for display and analysis of macromolecular
structures. J Mol Graph 14, 51–55.
Supplementary material
The following supplementary material is available
online:
Fig. S1. Plot of NOE number per residue.
Fig. S2. Determination of the dissociation constant of
12.0 ± 1.9 m
M for orthophosphate binding from the
amide chemical shift changes of Gly335.
Table S1. Structural statistics for rCNP catalytic
domain.
This material is available as part of the online article
from

Please note: Blackwell Publishing is not responsible for
the content or functionality of any supplementary
materials supplied by the authors. Any queries (other
than missing material) should be directed to the corres-
ponding author for the article.
G. Kozlov et al. Solution structure of the RICH catalytic domain
FEBS Journal 274 (2007) 1600–1609 ª 2007 The Authors Journal compilation ª 2007 FEBS 1609