One characteristic of the CD-degrading enzymes is their additional N-terminal domain, which in neopullulanase from Thermoactinomyces vulgaris TVA-II [10], maltogenic amylase from Thermus
Trang 1Covalent and three-dimensional structure of the cyclodextrinase
Hanna B Fritzsche, Torsten Schwede and Georg E Schulz
Institut fu¨r Organische Chemie und Biochemie, Albert-Ludwigs-Universita¨t, Freiburg im Breisgau, Germany
Starting with oligopeptide sequences and using PCR, the
gene of the cyclodextrinase from Flavobacterium sp no 92
was derived from the genomic DN A The gene was
sequenced and expressed in Escherichia coli; the gene
pro-duct was purified and crystallized An X-ray diffraction
analysis using seleno-methionines with multiwavelength
anomalous diffraction techniques yielded the refined 3D
structure at 2.1 A˚ resolution The enzyme hydrolyzes
a(1,4)-glycosidic bonds of cyclodextrins and linear
malto-oligo-saccharides It belongs to the glycosylhydrolase family no 13
and has a chain fold similar to that of a-amylases,
cyclo-dextrin glycosyltransferases, and other cyclocyclo-dextrinases In
contrast with most family members but in agreement with
other cyclodextrinases, the enzyme contains an additional characteristic N-terminal domain of about 100 residues This domain participates in the formation of a putative D2 -sym-metric tetramer but not in cyclodextrin binding at the active center as observed with the other cyclodextrinases More-over, the domain is located at a position quite different from that of the other cyclodextrinases Whether oligomerization facilitates the cyclodextrin deformation required for hydro-lysis is discussed
Keywords: calcium-binding site; cyclodextrin degradation; glycosylhydrolase family no 13; oligomerization; X-ray analysis
Cyclodextrins (CDs) are cyclic malto-oligosaccharides of at
least six to generally eight glucosyl units linked via
a(1,4)-glycosidic bonds Their ability to form inclusion complexes
with numerous small hydrophobic molecules is used in
various applications such as microencapsulation of drugs [1]
and chromatographic separation of chiral compounds [2]
The increasing application of CDs has stimulated an interest
in the mechanisms of their degradation, particularly as they
are resistant to hydrolysis by most a-amylases
Several CD-degrading enzymes have been isolated [3–8]
They are from various bacterial sources and generally prefer
CDs but also accept other maltodextrins, converting them
to a broad spectrum of products It has been suggested that
these cyclodextrinases (EC 3.2.1.54) should be combined
with maltogenic amylases (EC 3.2.1.133) and
neopullula-nases(EC3.2.1.135)intoasingleenzymeclass[9]becausetheir
catalytic properties differ only partially and not distinctly
One characteristic of the CD-degrading enzymes is their
additional N-terminal domain, which in neopullulanase
from Thermoactinomyces vulgaris (TVA-II) [10], maltogenic amylase from Thermus sp (ThMA) [11] and cyclodextrinase from Bacillus sp I-5 (BaCD) [9] participates in dimer formation In these dimers, the N-terminal domain of one subunit contacts the active center of the other subunit and participates in CD binding Moreover, it constricts the active-center pocket, affecting substrate specificity, for instance, by excluding large molecules such as starch [11–13] Beyond the common dimerization, BaCD forms a hexamer of these dimers, i.e a dodecamer in solution [9] Here we investigate the cyclodextrinase from Flavobac-teriumsp no 92 (CDase) which hydrolyzes CDs and short linear malto-oligosaccharides at comparable rates The enzyme exhibits only minor hydrolytic activity on the a(1,4)-linkages of starch [14] and pullulan [15] but shows considerable transglycosylation activity [16] The sequence and 3D structure of the enzyme is presented and compared with related proteins
Experimental procedures
Isolation and sequencing of the gene The enzyme was purified from Flavobacterium sp no 92 as described [4] The N-terminal amino-acid sequence was determined to be AAPTAIEHMEPPFW using Edman degradation in a gas phase sequencer (Applied Biosystems) Furthermore the enzyme was cleaved with CNBr, and the sequences of the six resulting peptides were analyzed One of the fragments, with the sequence MPDRFANGDPSND, was selected because it showed 60% amino-acid sequence identity with several a-amylases and cyclodextrin glycosyl-transferases (CGTases) in the SWISSPROT Data Bank On the basis of these two peptides the following two primers were constructed (S denotes a C/G mixture and R stands for
Correspondence to G E Schulz, Institut fu¨r Organische Chemie und
Biochemie, Albertstr 21, Freiburg im Breisgau, D-79104, Germany.
Fax: + 49 761 203 6161, Tel.: + 49 761 203 6058,
E-mail: schulz@bio.chemie.uni-freiburg.de
Abbreviations: BaCD, cyclodextrinase from Bacillus sp I-5; CD,
cyclodextrin, i.e cyclic malto-oligosaccharide of six or more glucosyl
groups; CDase, cyclodextrinase from Flavobacterium sp no 92;
CGTase, cyclodextrin glycosyltransferase; TAKA, a-amylase from
Aspergillus oryzae; ThMA, cyclodextrin-degrading maltogenic
amylase from Thermus sp.; TVA-I, a-amylase 1 from
Thermoactino-myces vulgaris; TVA-II, neopullulanase from ThermoactinoThermoactino-myces
vulgaris.
(Received 28 January 2003, revised 21 March 2003,
accepted 2 April 2003)
Trang 2A/G): 5¢-GCSCCSACSGCSATCGAGCACATGGA-3¢
(residues 2APTAIEHME) and 3¢-TACGGSCTRGCSAA
GCGSTTG-5¢ (reverse, residues (113)MPDRFAN) Using
these two primers in a PCR amplification with genomic
DNA from the Flavobacterium as the template, a 350 bp
DNA fragment was produced Using the random primer
method [17], this fragment was taken as a template to
produce [32P]dCTP-labeled probes
Genomic DNA from Flavobacterium sp no 92 was
prepared using a slightly modified protocol of Sambrook
et al [18], partially digested with Sau3A and fractionated by
sucrose gradient centrifugation Fragments ranging from 7
to 12 kb were used to prepare a genomic library in kZAP
Express DNA (Stratagene) The recombinant phages were
packaged in vitro with Gigapack II Packaging Extract
(Stratagene) and plated on Escherichia coli XL1-Blue MRF¢
(Stratagene) to a final concentration of 5000 pfu per plate
(diameter 15 cm) As determined by blue/white selection,
the library contained 55 000 independent plaques including
10% wild-type phages without inserts Positive plaques were
identified by in situ hybridization with the radiolabeled
probe They were subcloned in vivo into pBK-CMV
phagemides (Stratagene) by coinfection with the helper
phage M13 Exassist (Stratagene), and then analyzed with
restriction enzymes A clone with the complete gene was
isolated With the use of the dideoxy method [19], the gene
sequence was determined by PAGE and by more advanced
methods (SeqLab, Go¨ttingen, Sweden) The complete DNA
sequence has been deposited in the EMBL Nucleotide
Sequence Database under accession code AJ489171
Expression, purification and crystallization
The CDase gene without the signal sequence was subcloned
into the expression vector pET22b+ (Novagene) using
restriction enzymes EcoRI and NdeI Thereby, the first
alanine of the mature enzyme was replaced by a methionine
The CDase gene was then expressed in E coli strain
BL21(DE3) Cells were grown at 25C in Luria–Bertani
broth supplemented with 100 lgÆmL)1 ampicillin and
induced at an A600of 0.4–0.5 by adding 0.1 mMisopropyl
thio-b-D-galactoside They were harvested 4.5 h after
induction, resuspended in buffer A (50 mM Hepes,
pH 6.5, 2 mMCaCl2), and disrupted using a French press
After centrifugation, the supernatant was diluted 1 : 1 with
buffer A and loaded on to a cation-exchange column
(Source 30S; Pharmacia) The enzyme was eluted from the
column at 120 mM within a 0–200 mM NaCl gradient in
buffer A and was identified using SDS/PAGE The main
fractions were pooled and concentrated to 6 mgÆmL)1
protein The yield of the purified protein was 8 mg per litre
of culture medium For the crystallization experiments, the
CDase was dialyzed against deionized water
For phasing the X-ray reflections with the
multiwave-length anomalous diffraction method, all methionines were
replaced with seleno-methionines For this purpose, the
CDase was expressed at 25C in the
methionine-auxo-trophic E coli strain B834(DE3) using a culture medium
containing seleno-D,L-methionine at a concentration of
50 mgÆL)1[20,21] The purification procedure was similar to
that of the wild-type enzyme, but 3 mMdithiothreitol was
added to all buffers and solutions to avoid oxidation of the
incorporated seleno-methionines The yield of Se-labeled CDase was 6 mg per litre of culture medium and thus only slightly lower than that of the wild-type enzyme
Crystallization was carried out by the hanging drop vapor diffusion method using a sparse matrix screen (Hampton Research, La Jolla, CA, USA)
optimiza-tion, the best crystal conditions for the wild-type enzyme were a 1 : 1 mixture of a 6 mgÆmL)1protein solution and the reservoir buffer containing 50 mMHepes, pH 7.5, and 3.8 MNaCl The Se-labeled CDase crystallized under the same conditions, except for the addition of 3 mM dithio-threitol Crystals appeared within 2 days and grew to final dimensions of 500· 200 · 100 lm3 For cryoprotection, 15% glycerol was added just before the crystals were mounted in a cryo-loop and shock-frozen The crystals of wild-type and Se-labeled CDase were isomorphous Structure determination, phasing and refinement Preliminary data for wild-type CDase crystals and heavy atom derivatives were collected on a wire-frame detector (X-1000; Bruker-Nicolet, Karlsruhe, Germany)
rotating anode (RU200B; Rigaku, Tokyo, Japan)
data, however, were collected with an Se-labeled CDase crystal at synchrotron beamline BW7A (EMBL-outstation, DESY Hamburg) at three different wavelengths, which were selected on the basis of an X-ray fluorescence spectrum taken from the same crystal Data were processed and scaled with the program suiteHKL[22] bringing Friedel pairs
to the same scale The positions of 48 selenium atoms were determined with program SHELX-D[23] Phases were cal-culated with SHELX-E [24] and in a second run also with programSHARP/AUTOSHARP[25] The two resulting density maps were of equal quality
The model was built by a combination ofARP/WARP[26] and manual operations using programO[27] The complete model was refined by simulated annealing with noncrystal-lographic symmetry (NCS) restraints using program CNS [28] Several refinement cycles with individual isotropic B-factors followed Water molecules were either automati-cally identified by program CNS or manually introduced using programO The final refinement was performed using theTLSapproach inREFMAC[29] without NCS restraints ProgramLSQMAN[30] was used for structural alignments Figures were produced withMOLSCRIPT[31] andRASTER3D [32] The co-ordinates and structure factors are deposited in the Protein Data Bank under accession code 1H3G
Results and discussion
DNA and polypeptide sequence The CDase gene consists of 1857 bases of which the first 54 bases code for a signal sequence for protein translocation into the periplasm The DNA sequence agreed with the independently established amino-acid sequences of seven peptides The derived amino-acid sequence is given in Fig 1 except for the 18-residue signal peptide The native mature protein consists of 601 residues with an Mrof 67 946 On the basis of sequence similarity, it belongs to the glyco-sylhydrolase family no 13 [33] The four conserved segments of family no 13 (Fig 1) represent the calcium
Trang 3site, Ca-I, and the three invariant catalytic acids, Asp311,
Glu340 and Asp418
The 102 N-terminal residues of CDase form a domain
that is missing in most other members of family no 13
However, it is also present in the other structurally
established CD-degrading enzymes neopullulanase TVA-II
[10], maltogenic amylase ThMA [11], cyclodextrinase BaCD
[9], and a second neopullulanase that resembles TVA-II, but
is not yet available from the Protein Databank [34]
Furthermore, it is present in the a-amylase TVA-I [13],
a trehalohydrolase [35], and an isoamylase [36] This
N-terminal domain is not present in the CD-producing
CGTases, which, however, contain about 150 additional
C-terminal residues that probably mediate starch binding
[37–39]
3D structure
The crystals of Se-labeled CDase belong to space group
R32 with unit cell dimensions a¼ b ¼ 181.3 A˚ and
c¼ 231.5 A˚ at 100 K and two CDase molecules in
the asymmetric unit They have a packing parameter of
2.6 A˚3/Da, which is 4% smaller than that of the
isomor-phous wild-type crystals at 100 K The wild-type crystals
failed to show a comparable diffraction quality and were
therefore not further analyzed Data collection statistics are
given in Table 1 The structural refinement yielded a
crystallographic R factor of 18.8% and an Rfreeof 22.3%
with over 90% of the residues in the most favored region of
a Ramachandran plot (Table 2) The resulting model is depicted in Fig 2 The Ca backbones of the two molecules can be superimposed, with an rmsd of 0.31 A˚, indicating conformational homogeneity Met1 is removed during expression in E coli; Ala2, Glu600 and Ala601 have no density The overall real space map correlation coefficient was 0.95 [30] The model includes about 0.6 water molecules per residue, which is appropriate at 2.1 A˚ resolution The
Bfactor plot for both molecules is shown in Fig 3 The peaks are almost exclusively in loop regions
Following the assignments in related enzymes, CDase was divided into four domains (Fig 2) As a member of the glycosylhydrolase family no 13, its chain fold is similar to that of known a-amylases consisting of a central TIM barrel [40] (domain A, residues 103–516), with a 60-residue insert after the third strand of the b-barrel (domain B, 223–282) and a C-terminal domain (domain C, 517–601) In addition, CDase contains an N-terminal domain (1–102), which assumes a characteristic b-sandwich structure composed of the antiparallel strands b1 to b8 The N-terminal domain is connected by an extended 10-residue linker to the TIM barrel It contacts the bulk of the molecule at helices a6, a7 and at domain B
Domain A harbors the active center at the C-termini of the TIM barrel b-strands The loops at the C-terminal barrel end connecting b-strands with the following a-helices are longer and more complex than the loops at the opposite barrel end The lengths of the b-strands in the barrel vary from two (b14)
to seven (b11) residues As in other enzymes of family no 13, the regularity of the CDase TIM barrel is broken by the a-helices after the sixth b-barrel strand where helix a9 extends in the direction of the preceding strand b14 and only the next helix a10 runs in the opposite direction (Fig 2) For historical reasons the large loop between the third strand of the TIM barrel (b11) and helix a6 is called domain
B (indicated in Fig 2) This inserted domain participates in substrate binding and is rather variable It is considered to play a role in determining the enzyme specificity [41] The end of the TIM barrel domain A is connected to domain C forming two antiparallel sandwiched b-sheets Between them, the sheet contacting the TIM barrel at helices a9, a10, a12 and a13 contains strands b17, b18, b19 and b24, whereas the solvent-exposed second b-sheet harbors strands b20, b21, b22 and b23 The b-sheet at the interface to domain A has lower B factors (Fig 3) and is structurally much better conserved within the family than the other
Fig 1 Amino-acid sequence of native mature CDase derived from the
DNA sequence Independently established peptide sequences are
underlined The four conserved segments at the calcium-binding site
Ca-I and at the three invariant catalytic residues (inverted) Asp311,
Glu340 and Asp418 are given in bold letters Residues shown in lower
case are not included in the model In the analyzed enzyme, Ala1 was
replaced by Met1 which, however, was removed during protein
expression in E coli.
Table 1 Data collection for phasing with multiwavelength anomalous diffraction.
Resolution a (A˚) 24–2.4 (2.5–2.4) 26.7–2.4 (2.5–2.4) 25–2.1 (2.18–2.08)
Unique reflectionsa 58931 (5892) 58981 (5898) 86572 (8625) Completeness a (%) 99.8 (99.8) 99.9 (99.9) 99.9 (99.9)
R sym-Ia(%) 7.7 (21) 5.1 (22) 6.0 (40)
a
Values in parentheses refer to the outermost shell.
Trang 4sheet, which supports the proposal that domain C stabilizes
the TIM barrel
Like most other members of family no 13, CDase
contains Ca2+ions One of the two Ca2+ions in CDase is
at site Ca-I, which is widely conserved forming the first of the four sequence fingerprints shown in Fig 1 [42] The removal of Ca-I was shown to promote proteolysis [43,44] Ca-I is co-ordinated by the side chains of Asp280 and Ser222 at the beginning and end of domain B as well as by the main-chain oxygens of Tyr315 (domain A) and Thr270 (domain B) and by two water molecules Ser222 of the Ca-I sequence fingerprint is specific for CDase, where it replaces
a highly conserved asparagine At its position between domains A and B, Ca-I stabilizes the conformation of domain B together with residues Tyr315, Phe274 and others that are directly or indirectly involved in substrate binding Ca-I is missing in the three CD-degrading enzymes TVA-II, ThMA and BaCD
The second and third calcium-binding sites of family no
13 enzymes show greater variation [42,43,45] The second calcium site of CDase is called Ca-II It is also present in the CD-producing CGTase [38] and in the CD-degrading
TVA-II [13], but not in the CD-degrading enzymes ThMA and BaCD nor in the majority of a-amylases Ca-II is located in the loop between a1 and a2 of domain A (Fig 2) and
Table 2 Refinement statistics Values in parentheses refer to the
outermost shell.
Resolution range (A˚) 21–2.1 (2.13–2.08)
Number of reflections 78164 (4817)
Average B factor (A˚ 2 ) 41
R cryst (%) 18.8 (21.9)
R free (%) (test set of 1991 reflections) 22.3 (27.8)
Rmsd bond lengths (A˚)/angles () 0.016/1.34
Ramachandran angles in:
Most favored region (%) 90.2
Allowed [generally allowed] region (%) 9.5 [0.3]
Fig 2 Stereoview of a ribbon plot of CDase showing the N-terminal domain (red), the TIM-barrel domain A (blue), the inserted domain B (green) and the C-terminal domain C (orange) The two Ca2+ions are represented by black spheres The active center is indicated by the three invariant catalytic residues (Fig 1) All a-helices and b-strands are labeled.
Fig 3 B-factor plots of the main chains of the two molecules of CDase in the asymmetric unit The averages of molecules A (solid line) and B (broken line) are 39 A˚2and 44 A˚2, respectively The a-helices and b-strands are given for reference Helices a1 through a14 and strands b9 through b16 comprise the TIM barrel Domain B is inserted between b11 and a6 The seven 3 -helices are not indicated.
Trang 5co-ordinated by the side chains of Asp125, Asp146, Asn119
and Asn124, the main-chain oxygens of Gly144 and
Asp121, and by a water molecule Ca-II stabilizes a surface
region far away from the active center The B factors of both
Ca2+ions in both CDase molecules are similar to those of
the surrounding atoms This indicates that the sites are fully
occupied in the crystal, even though the protein was
dialyzed against deionized water before crystallization and
the crystallization buffer lacked calcium
Oligomerization
In the crystal, CDase forms putative D2-symmetric
tetramers containing two noncrystallographic and one
crystallographic twofold axes The oligomerization was
derived from the large contact areas formed within each
tetramer and the small contacts between neighboring tetra-mers, making the crystal look like an assembly of tetramers As shown in Fig 4, the tetramer contact across the crystallographic twofold axis is 1590 A˚2in size, which
is in the normal range of oligomer interfaces The other internal contact is formed by the N-terminal domains and measures 520 A˚2, which exceeds an average crystal packing contact The large difference between these two interface areas classifies the tetramer as a weak dimer of strong dimers A preliminary size-exclusion chromato-graphy run (Sephacryl 300S, 100 mM Hepes, pH 7.5) at 0.15M N aCl compared with 3.8M NaCl in the crystals showed a dominating dimer mixed with other oligomers Similar runs under other conditions have yet to be performed to determine the detailed oligomerization pattern in solution
Fig 4 D 2 -symmetric tetramer structure of CDase in the crystal together with the symmetry axes (A) Front view placing the crystallographic twofold axis horizontally in the paper plane The crystallographic axis runs through the large interface and the vertical noncrystallographic axis runs through the small interface between the N-terminal domains One subunit is given in the colors and in an orientation similar to Fig 2 A b-CD (orange) derived from a superposition with the complex between b-CD and the homologous enzyme TVA-II [47] marks the active center (B) View from the left side of (A), which is along the crystallographic twofold axis, showing a smooth silhouette.
Trang 6The chain fold of the N-terminal domain of CDase is
similar to that of the related CD-degrading enzymes TVA-II
[10], ThMA [11] and BaCD [9] However, the positions of
these domains relative to the respective TIM barrel are
completely at variance as shown in Fig 5 The N-terminal
domains of TVA-II, ThMA and BaCD attach to the active
center of the other subunit and participate in substrate
selection [11,12] This dimer interface is not related to either
of the two interfaces in the CDase tetramer It seems very
unlikely that the deviating domain position in CDase is
a packing artefact caused by domain swapping during
crystallization because the interface between the N-terminal
domain and the protein remainder (domains A and B)
amounts to 1390 A˚2, which is much larger than a common
packing contact
Comparison with related enzymes
The relationships within the group of CD-degrading
enzymes were evaluated by a comprehensive chain-fold
comparison The comparison was extended to the
structur-ally related TVA-I [13], the CD-producing CGTases [38,39]
and a-amylase from Aspergillus oryzae (TAKA) [46], which
was taken as a well-known representative of family no 13
In principle, all comparisons could have been performed
with the amino-acid sequences alone, as the
glycosylhydro-lase families are defined by the sequences However, this method suffers from the rather arbitrary placing of the gaps Therefore, we took account of the geometry and first derived the group of structurally equivalent residues in a chain-fold superposition and then counted the number of identical residues within this group The results are given in Table 3
The most obvious result of this comparison is the close relationship between TVA-II, ThMA and BaCD, which can be almost completely structure-aligned, giving rise to about 50% identical residues As mentioned above, these three enzymes also form similar dimers (with a further hexameric association in BaCD) and have similar catalytic activities Therefore, we classify them as the TVA-II group When comparing CDase with this group, only 380 of the
600 residues can be structure-aligned, and only 28% of the aligned residues are identical This renders CDase an outlier among the structurally established CD-degrading enzymes As for the other enzymes, a-amylase TVA-I shows considerable structural similarity to the TVA-II group, although its function differs greatly (Table 3) Moreover, the data reveal that the CD-degrading enzymes are more similar to the CD-producing CGTase than to the a-amylase TAKA
A more obvious difference between CDase and the others
is the deviating spatial position of its N-terminal domain
Fig 5 Stereoview of the superposition of
CDase (colored as in Fig 2) with TVA-II
(black, Ca 2+ at Ca-II grey) given as
Ca-backbone plots The completely different
positions of the N-terminal domains and the
differences in domain B near Ca-I (right) are
clearly visible.
Table 3 Chain-fold comparisons within glycosylhydrolase family no 13 The upper right triangle shows the number of Ca atoms aligned within the
3 A˚ distance criterion in superpositions of the complete polypeptide chains using program LSQMAN (30) The numbers in parentheses are the percentages of identical residues in the aligned segments For CDase, CGTase and TAKA, only domains A, B and C could be superimposed with any of the other enzymes The lower left triangle shows the respective numbers for a separate superposition series involving only the N-terminal domains.
CDase 387 (29) 387 (28) 376 (26) 371 (25) 380 (26) 361 (25) TVA-II 47 (11) 547 (47) 552 (47) 428 (37) 378 (25) 369 (21) ThMA 47 (13) 121 (35) 570 (54) 472 (32) 366 (24) 368 (26)
Trang 7(Fig 5) A superposition restricted to the N-terminal
domains showed that those of the TVA-II group can be
almost fully structure-aligned, resulting in 40%
amino-acid residue identities (Table 3) TVA-I is somewhat outside
the TVA-II group, but can still be well aligned However,
the N-terminal domain of CDase aligns only with about
half of its residues, shows almost no sequence identity
(Fig 6 and Table 3), and clearly differs from the TVA-II
group with respect to sequence, chain fold, and position
The N-terminal domains of the trehalohydrolase [35] and
the isoamylase [36] have a similar chain fold to that of
CDase and the TVA-II group, but they are barely related to
any of them (data not shown) Interestingly, the general
location of the N-terminal domains of these two outliers
[35,36] corresponds to that of CDase
A superposition of the highly variable B domains, which
participate in the active center, is given in Fig 7 CDase has
a very long extension, whereas CGTase and TAKA have
intermediate ones In contrast, the TVA-II group and TVA-I have a much smaller B domain The large B domains
of CDase, CGTase and TAKA are fixed by Ca-I, which is absent in the TVA-II group with their small B domains The surprisingly large difference between CDase on one hand and the TVA-II group on the other corresponds to the different oligomeric structures CDase uses the long exten-sion of its B domain to make an intimate contact across the strong dimer interface with domains A and C of the other subunit In contrast, the TVA-II group dimer attaches the B domain to an N-terminal domain of the other subunit, which restricts the size of the B domain (Fig 7)
Active center The active center of CDase is depicted in Fig 8, which includes the superimposed structure of a TVA-II dimer with bound b-CD [47] Interestingly, the superposition causes a
Fig 6 Structural alignment of the N-terminal domain of CDase with those of TVA-II [10], ThMA [11], BaCD [9], TVA-I [13], a trehalohydrolase [35] and an isoamylase [36], which are the only structurally similar domains within glycosylhydrolase family no 13 CD-degrading activity has been reported for the top four enzymes The secondary structure of CDase is given, and every 10th amino acid residue is marked by a dot Residues 87–175 of the isoamylase have been omitted (marked #) All residues that superimpose within the 3 A˚ distance criterion of program LSQMAN [30] are underlined For reference, strand b9 of the TIM barrel has been included, and the alignments with the TIM barrels are given in bold.
Fig 7 Superposition of the inserted B domains
of CDase (green, His251 marked by a ball), TVA-II (red), CGTase (blue) and TAKA (grey)
as aligned on the TIM barrels TVA-II, ThMa and BaCD are so similar that only one of them was drawn out for clarity As TVA-I varies only slightly from TVA-II, it was omitted The chain direction is indicated by the N* and C* ends.
Trang 8clash between the long B-domain extension of CDase
(His251, Fig 7) and the N-terminal domain of the other
subunit of the TVA-II dimer (Tyr45¢) It has been suggested
that the N-terminal domain of the TVA-II group [9–11]
confers CD specificity because it covers one side of the bound
CD [47] In CDase, this role is fulfilled by the B domain of the
same subunit Therefore, it is sterically impossible for CDase
to form the same dimer as the TVA-II group
As the polypeptide superposition of Fig 8 places the
three catalytic residues of CDase (Fig 1) within less than
1 A˚ of the positions of those of TVA-II, and as the active
centers closely resemble each other, the CD molecule bound
to TVA-II can be expected to bind at a similar position in
the CDase structure The hydrolysis of CD should start by
Glu340 protonating a bridge oxygen of the cyclic substrate
However, the distance between Glu340 and the next bridge
oxygen is 6 A˚, which is much too long A similar distance to
a bound CD has been observed in CGTase [48], where,
however, it has been demonstrated that a linear
malto-oligosaccharide binds much deeper in the pocket at the
required 3 A˚ distance to the Glu340 equivalent [49]
Moreover, the conformation at the scissile bond in a CD
complex with CGTase showed a substantial deviation from
the circular symmetry [48] These observations indicate that
the observed CD-binding position in TVA-II is most likely
displaced by about 3 A˚ For catalysis, the CD molecule has
to be pushed 3 A˚ deeper into the active-center pocket and
deformed at its scissile bond [48] Such a CD position has
not yet been observed in any crystal structure It would
enable Phe274 of CDase (or Phe286 of TVA-II) to rotate
around its Ca–Cb bond and enter the CD hollow, as has
been implied for TVA-II [10] and for Tyr195 of CGTase
[49] Crystal experiments to clarify the CD position in
CDase are under way
The required induced fit and deformation of the bound
CD need energy, part of which may be derived from
co-operative effects in the CDase tetramer (or TVA-II
dimer) association This proposal is consistent with the
observation of a higher rate of CD hydrolysis for dimeric
ThMA than for the monomeric ThMA [12] Although such
an energy source is conceivable for the TVA-II group in
which the bound CD contacts the N-terminal domain of the other subunit, it is also possible for CDase in which the bound CD is very close to the long B-domain extension (Fig 7) as well as to A-domain and C-domain residues of the other subunit (Fig 4A) In fact, the interface mediating the strong dimer association would appear to explain the particularly long B-domain extension of CDase In con-clusion, the dimer association may help to overcome the conformational activation energy barrier during CD hydrolysis
Acknowledgements
We thank H Bender for drawing our attention to the enzyme, E Schiltz for amino-acid sequence analyses, M Ru¨ckels for preparing the initializing 350-bp fragment and C Vonrhein for help with SHARP / AUTOSHARP Moreover, we are grateful for the contributions of
S Thorspecken, A Dorowski, B Phillips, S Jelakovic, C Schleberger and M Mrosek at early stages of the analysis, and we thank the beamline staff of the EMBL-outstation (DESY Hamburg) for help with the data collection The project was supported by the European Commision under BIO4-98-0022 (AGADE) and by the Deutsche Forschungsgemeinschaft under GRK-434.
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