Rational design, synthesis, evaluation and enzyme - Substrate structures of improved fluorogenic substrates for family 6 glycoside hydrolases

Article abstract of DOI:10.1111/febs.12060

Methylumbelliferyl-β-cellobioside (MUF-G2) is a convenient fluorogenic substrate for certain β-glycoside hydrolases (GH). However, hydrolysis of the aglycone is poor with GH family 6 enzymes (GH6), despite strong binding. Prediction of the orientation of the aglycone of MUF-G2 in the +1 subsite of Hypocrea jecorina Cel6A by automated docking suggested umbelliferyl modifications at C4 and C6 for improved recognition. Four modified umbelliferyl-β-cellobiosides [6-chloro-4-methyl- (ClMUF); 6-chloro-4-trifluoromethyl- (ClF3MUF); 4-phenyl- (PhUF); 6-chloro-4-phenyl- (ClPhUF)] were synthesized and tested with GH6, GH7, GH9, GH5 and GH45 cellulases. Indeed the rate of aglycone release by H. jecorina Cel6A was 10-150 times higher than with MUF-G2, although it was still three orders of magnitude lower than with H. jecorina Cel7B. The 4-phenyl substitution drastically reduced the fluorescence intensity of the free aglycone, while ClMUF-G2 could be used for determination of k_cat and K_M for H. jecorina Cel6A and Thermobifida fusca Cel6A. Crystal structures of H. jecorina Cel6A D221A mutant soaked with the MUF-, ClMUF- and ClPhUF-β-cellobioside substrates show that the modifications turned the umbelliferyl group 'upside down', with the glycosidic bond better positioned for protonation than with MUF-G2. 2012 The Authors Journal compilation

Full text of DOI:10.1111/febs.12060

Rational design, synthesis, evaluation and enzyme–
substrate structures of improved fluorogenic substrates
for family 6 glycoside hydrolases
Miao Wu1, Wim Nerinckx2, Kathleen Piens1,2,*, Takuya Ishida1,†, Henrik Hansson1, Mats Sandgren1
1
ꢀ
and Jerry Stahlberg
1 Department of Molecular Biology, Swedish University of Agricultural Sciences, Uppsala, Sweden
2 Laboratory for Protein Biochemistry and Biomolecular Engineering, Department of Biochemistry and Microbiology, University of Gent,
Gent, Belgium
Keywords
Methylumbelliferyl-b–cellobioside (MUF–G2) is a convenient fluorogenic
substrate for certain b–glycoside hydrolases (GH). However, hydrolysis of
the aglycone is poor with GH family 6 enzymes (GH6), despite strong
cellobiohydrolase; cellulase; exo-anomeric
effect; Hypocrea jecorina;
methylumbelliferyl-b–cellobioside
binding. Prediction of the orientation of the aglycone of MUF–G2 in the
+1 subsite of Hypocrea jecorina Cel6A by automated docking suggested
Correspondence
umbelliferyl modifications at C4 and C6 for improved recognition. Four
modified umbelliferyl-b–cellobiosides [6–chloro-4–methyl- (ClMUF);
6–chloro-4-trifluoromethyl- (ClF3MUF); 4–phenyl- (PhUF); 6–chloro-4–
phenyl- (ClPhUF)] were synthesized and tested with GH6, GH7, GH9,
GH5 and GH45 cellulases. Indeed the rate of aglycone release by H. jecori-
na Cel6A was 10–150 times higher than with MUF–G2, although it was
still three orders of magnitude lower than with H. jecorina Cel7B. The
4–phenyl substitution drastically reduced the fluorescence intensity of the
free aglycone, while ClMUF–G2 could be used for determination of kcat
and KM for H. jecorina Cel6A and Thermobifida fusca Cel6A. Crystal
structures of H. jecorina Cel6A D221A mutant soaked with the MUF-,
ClMUF- and ClPhUF-b–cellobioside substrates show that the modifica-
tions turned the umbelliferyl group ‘upside down’, with the glycosidic bond
better positioned for protonation than with MUF–G2.
ꢀ
J. Stahlberg, Department of Molecular
Biology, Swedish University of Agricultural
Sciences, PO Box 590, SE-75124 Uppsala,
Sweden
Fax: +46 18 536971
Tel: +46 18 4714590
E-mail: jerry.stahlberg@molbio.slu.se
Present address
*Oxyrane Belgium, Gent-Zwijnaarde,
Belgium
†Department of Biomaterials Sciences,
The University of Tokyo, Japan
M. Wu and W. Nerinckx contributed equally
to this study
Database
(Received 31 August 2012, revised 31
Structural data have been submitted to the Protein Data Bank under accession numbers pdb
October 2012, accepted 5 November 2012)
4AU0, 4AX7, 4AX6
doi:10.1111/febs.12060
Structured digital abstract
• http://mint.bio.uniroma2.it/mint/search/interaction.do?interactionAc=MINT-7260296
• Cel6A and Cel6A bind by x-ray crystallography (View Interaction: 1, 2)
Abbreviations
CBH, cellobiohydrolase; ClF3MUF, 6–chloro-4-trifluoromethylumbelliferone; ClF3MUF–G2, 6–chloro-4-trifluoromethylumbelliferyl-
b–cellobioside; ClMUF, 6–chloro-4–methylumbelliferone; ClMUF–G2, 6–chloro-4–methylumbelliferyl-b–cellobioside; ClPhUF, 6–chloro-4–
phenylumbelliferone; ClPhUF–G2, 6–chloro-4–phenylumbelliferyl-b–cellobioside; EG, endoglucanase; GH, glycoside hydrolase; MUF,
4–methylumbelliferone; MUF–G2, 4–methyl-umbelliferyl-b–cellobioside; MUF-G4, 4–methylumbelliferyl-b–cellotetraoside; PhUF,
4–phenylumbelliferone; PhUF–G2, 4–phenylumbelliferyl-b–cellobioside.
184
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M. Wu et al.
Fluorogenic substrates for GH6
Introduction
The filamentous ascomycete fungus Hypocrea jecorina
(previously known as Trichoderma reesei) has served as
a model organism for research on the mechanism, speci-
ficity and synergistic cooperation of cellulose-degrading
enzymes. It is also the predominant source of cellulase
enzymes for industrial use [1]. The genome of H. jecori-
na has been sequenced and transcriptome analysis
identified ꢀ 35 genes coding for known or putative
biomass-active proteins that were upregulated on cellu-
lose-containing media [2]. Major components are two
cellobiohydrolases (CBH), Cel7A (40–50%) and Cel6A
( ꢀ 20%), characterized by having their active sites
enclosed in a cello-oligosaccharide binding tunnel [3,4],
and the ability to processively cleave off several cellobi-
ose units from the end of a cellulose chain before
substrate release [5–7]. The enzymes cooperate synergis-
tically and act preferentially from opposite ends of cellu-
lose chains, Cel7A from the reducing end and Cel6A
from the nonreducing end. They also utilize different
reaction mechanisms. Cel7A hydrolyses glycosidic
bonds with net retention of the b-anomeric configura-
tion, whereas Cel6A is an inverting enzyme [8].
been divided into tunnel-type cellobiohydrolases (CBH)
and endoglucanases (EG) that have their active sites in a
more open cleft and show a higher tendency to cut inter-
nal bonds in the cellulose polymer. X–Ray structures are
available of both types, five CBHs: H. jecorina Cel6A
(PDB code 1CBH) [4], Humicola insolens Cel6A (PDB
code 1BVW) [24], Coprinopsis cinerea Cel6A (PDB code
3VOH) [25], C. cinerea Cel6C (PDB code 3A64) [26] and
Chaetomium thermophilum Cel6A (PDB code 4A05) [27],
and three EGs: H. insolens Cel6B (PDB code 1DYS)
[28], Thermobifida fusca Cel6A (PDB code 1TML) [29]
and Mycobacterium tuberculosum Cel6A (PDB code
1UP0) [30].
Enzyme mechanism of family 6 glycoside
hydrolases
Structure studies of ligand binding in GH6 CBHs have
provided detailed insights into the enzyme mechanism
[25,30–32]. Six D–glucose units of the nonreducing end
of a cellulose chain bind in subsites ꢁ2 to +4 with the
cellobioside to be cleaved off in subsites ꢁ2 to ꢁ1.
Tryptophan residues provide glucosyl-binding plat-
forms in subsites ꢁ2, +1, +2 and +4. An aspartic acid
residue, D221 in H. jecorina Cel6A, acts as the cata-
lytic acid and protonates the scissile glycosidic bond
between subsites ꢁ1 and +1. A water molecule is held
in position for nucleophilic attack on the anomeric
carbon of the D-glucose unit in subsite ꢁ1, but is not
in direct contact with any carboxylate side chain.
Instead it is hydrogen bonded to a second water mole-
cule, which in turn binds to the side chain of an aspar-
tate residue, D175 in H. jecorina Cel6A, that is
believed to act indirectly as the catalytic base, in a
so-called Grotthus mechanism. This has also been sup-
ported by molecular dynamics simulations [23]. Dual
conformations have been observed for both catalytic
residues. In several structures, they form a hydrogen
bond to each other. This hydrogen bond must be bro-
ken and both side chains must be in their ‘active’ con-
formations to catalyse the reaction. Furthermore, one
loop near the catalytic centre is flexible and needs to
swing in to a closed conformation to isolate the two
water molecules near the anomeric carbon [23,31].
Aryl saccharides have been used as chromogenic
substrates in cellulase research for over 50 years
[9–11]. Chromogenic and fluorogenic cellobiosides and
lactosides have been very helpful tools in research on
H. jecorina Cel7A and the homologous endoglucanase
Cel7B, as well as on GH7 enzymes from other organ-
isms, ranging from simple applications such as expres-
sion assays and monitoring of protein purification, to
high-throughput screening, pH- and temperature-
dependence profiling, enzyme kinetics and inhibition
studies [12–20]. However, similar applications have not
been readily available for H. jecorina Cel6A and other
GH6 enzymes, because of poor hydrolysis of the
heteroglycosidic bond and/or preferential cleavage at
homoglycosidic bonds in the substrate.
During the 1980s, Claeyssens and co-workers synthe-
sized 4–methylumbelliferyl (MUF) cello-oligosaccha-
rides to use as analytical tools for cellulase enzymes
[21]. Although the compounds did not work as fluoro-
genic substrates with H. jecorina Cel6A, the MUF
group served well as a reducing end label in studies of
bond cleavage preference within the saccharide part.
Furthermore, because release of the fluorophore
(4–methylumbelliferone; 1) was negligible, despite
strong binding at the active site, they could be used as
probes in direct and displacement fluorescence quench-
ing titrations for ligand-binding studies [22,23].
Mechanistic itinerary of GH6
A glycosidic bond substitution appears to subject the
involved glycon ring to a conformational change
[33–35]. Within the crystal structures of the GH6
CBHs (Cel6A) from H. jecorina (PDB codes 1QK2
GH6 cellulases are common in both cellulolytic
fungi and bacteria (http://www.cazy.org/) and have
FEBS Journal 280 (2013) 184–198 ª 2012 The Authors Journal compilation ª 2012 FEBS
185

Fluorogenic substrates for GH6
M. Wu et al.
and 1QJW) [31], and H. insolens (PDB codes 1GZ1
and 1OC5) [32] that are in Michaelis complex with a
nonhydrolysable thio-oligosaccharide, the 2SO confor-
mation is observed for the sugar moiety occupying the
ꢁ1 subsite. Also, in the 3D structures of H. insolens
Cel6A and M. tuberculosis Cel6 in complex with a
gluco-isofagomine cellobioside analogue (resp. PDB
codes 1OCN [36] and 1UP2 [30]), the isofagomine
moiety occupying the ꢁ1 subsite shows a conforma-
Closer inspection of ꢁ1/+1 subsite-spanning ligands
in crystal structures reveals
a further intriguing
corollary [38]. In anti-protonating GH enzymes, the
scissile anomeric bond (often as thio-analogue)
always shows a dihedral angle φ (O5′–C1′–[O,S]x–Cx)
that is in the lowest-energy synclinal (gauche)
position. A minus-synclinal dihedral angle φ for an
equatorial glycosidic bond, or plus synclinal for an
axial bond [39], allows for hyperconjugative overlap
of the C1′–O5′ antibonding orbital with an antiperi-
planar-oriented lone pair orbital lobe of the glyco-
sidic oxygen, thereby creating partial double-bond
character and stabilization of the glycosidic bond by
2
2
tion that is close to a S5 (equivalent to a SO in the
pyranoside numbering). The apparent preference for a
b-2SO sugar conformation has been interpreted as
being indicative of a synperiplanar lone pair hypothe-
sis compliant [b-2SO pre-TS ? 2,5B TS ? nonground
state a-5S1] hydrolysis mechanism itinerary [34,36], but
may just as well be indicative of a standard antiperi-
planar lone pair hypothesis compliant [b-2SO or nearby
4–5 kcalꢂmolꢁ1
; the so-called ‘exo-anomeric effect’
[40,41]. However, anti protonation occurs precisely on
the oxygen’s antiperiplanar lone pair, which removes
the stabilizing effect, thus providing an additional
enzyme-strategy for lowering the activation barrier
(Fig. 1 centre).
4
b-1S3 pre-TS ? E3 or nearby H3 TS ? ground state
a-4C1] itinerary [33,34].
Syn protonators show a different strategy [38]. In
many ligand complexes the dihedral angle φ of the
scissile bond has been substantially twisted away from
its lowest-energy synclinal position: clockwise to
minus-anticlinal or antiperiplanar for beta-equatorial;
counterclockwise to plus-anticlinal or antiperiplanar
for alpha-axial anomeric bonds. This removes the
hyperconjugative overlap and thus the stabilizing exo-
anomeric effect. Because of this twisting, the lone pair
of the glycosidic oxygen is pointing into the syn half-
space, ready to be protonated by the syn-positioned
proton donor (Fig. 1 right). GH6 enzymes are syn
protonators and in several subsite ꢁ1/+1 spanning
ligand complexes the dihedral angle of the (thio)glyco-
sidic bond is in minus-anticlinal position (e.g.
φ2 = ꢁ125° in H. jecorina PDB-entry 1QK2, φ2 =
ꢁ132° in H. insolens 1GZ1 or φ2 = ꢁ140° in T. fusca
2BOF).
Anti versus syn protonation and the
exo-anomeric effect
The concept of semi-lateral glycosidic oxygen proton-
ation was introduced in a seminal article by Height-
man and Vasella [37]. It was originally only described
for beta-equatorial glycoside hydrolases, but appears
to be equally applicable to enzymes acting on an
alpha-axial glycosidic bond [38]. When dividing subsite
ꢁ1 by the Ox, C1′ and H1′ plane of the ꢁ1 glycoside,
many ligand-complexed structures show that the pro-
ton donor is positioned either in the syn half-space,
near the ring-oxygen of the ꢁ1 glycoside, or in the anti
half-space at the opposite side of the ring-oxygen.
Members of the same GH family are always syn or
always anti protonators and this specificity appears to
be preserved within clans of families.
Anti protonation
Syn protonation
Beta glycosidic bond
Syn
Anti Syn
Anti
5'
O
2'
O
C2'
C2'
O5'
= minus-synclinal
C4
O5'
O
O
H
O
O
O
O
4
H
H
O
O
H1'
turned clockwise
to minus-anticlinal
or antiperiplanar
C4
H1'
H1'
subsite –1 subsite +1
Alpha glycosidic bond
turned counterclockwise
to plus-anticlinal
or antiperiplanar
O
Syn
Anti Syn
Anti
5'
H1'
H1'
C4
O
C4
= plus-synclinal
O5'
H1'
2'
O
H
O
O
O
4
O5'
C2'
C2'
subsite –1
subsite +1
Fig. 1. Newman projections, with the glycosidic oxygen as the proximal atom and the anomeric carbon as the distal atom, showing anti
(centre) versus syn (right) semi-lateral protonation in beta-equatorial (upper) and alpha-axial (lower) glycoside hydrolases. / is the dihedral
angle for O5′–C1′–O4–C4.
186
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M. Wu et al.
Fluorogenic substrates for GH6
tion towards possibly better recognized variants that
can be easily synthesized: (a) the 4–methyl group of
the umbelliferone points towards the +2 subsite which
can be filled by introduction of a 4–phenyl group; and
(b) the C6–H6 bond of the umbelliferone points
towards an empty local zone in subsite +1, which in
the original (Glc)2–S–(Glc)2–OMe complex is occupied
by the hydroxymethyl group of the resident D–gluco-
side. Introduction of a chlorine at C6 would occupy
this zone. This is an electron-withdrawing group that
will increase the leaving capacity of the aglycone, and
because it is close to the anomeric oxygen it may also
decrease the exo-anomeric effect. Concomitant incor-
poration of a 4–trifluoromethyl group can further
increase the leaving capacity of the aglycone.
Consequences for GH6 substrates
Summarizing from the considerations above, with the
syn-protonating GH6 enzymes the productive pose of
a substrate is expected to possess a ring conformation
close to a 2SO skew for the b-D-glucoside moiety in
subsite ꢁ1, whereas the dihedral angle φ of the scissile
anomeric bond is in the minus-anticlinal position. This
is a double deformation away from the ground state,
and the combined energetic burden (4–5 kcalꢂmolꢁ1
for the exoanomeric effect [40,41] plus 5–6 kcalꢂmolꢁ1
for the 4C1 into 2SO glucoside deformation [42]) will
have to be compensated for by favourable enzyme–
substrate interactions. We suspected that 4–methylum-
belliferyl-b–cellobioside (MUF–G2, 2) fails to be a
good substrate for GH6 enzymes because the 4–meth-
ylumbelliferyl moiety cannot make enough favourable
interactions with the +1 subsite to help the substrate
to be induced into the productive pose, after all, this
subsite has been evolved to specifically recognize a
D-glucoside, not a 4–methylumbelliferyl group. By con-
trast, the also syn-protonating GH7 enzymes do recog-
nize MUF–G2 (2) as a good substrate [21,22] probably
from a lucky subsite +1 match.
Vina dockings with ClMUF–G2 (7), ClF3MUF–G2
(8), PhUF–G2 (9) and ClPhUF–G2 (10) indeed indi-
cate a better recognition of the productive pose, each
Table 1. Glycosidic bond dihedral angles /2 and w2 at the ꢁ1/+1
subsite junction in the docking and in crystal structures of GH6
enzymes, and distance between the glycosidic oxygen/sulfur and
the catalytic acid.a
We thus decided to use automated docking with
MUF–G2 (2), with its central D-glucoside in the 2SO
skew conformation, into the active site of H. jecorina
Cel6A in order to obtain information on how its
umbelliferyl group would be oriented versus the +1
subsite and how the aglycone may be modified
towards better positioning and more efficient hydroly-
sis. Four modified umbelliferyl b–cellobiosides were
designed, synthesized and tested as substrates for GH6
and other enzymes, and enzyme-substrate complex
structures were determined by X-ray crystallography.
Pose
nrb
Dihedral
c
Dihedral
c
Calculated
affinityd
Docked ligand
/
w2
2
MUF–G2 (2)
11
3
ꢁ114°
ꢁ113°
ꢁ115°
ꢁ102°
ꢁ104°
ꢁ108°
ꢁ114°
ꢁ128°
ꢁ121°
ꢁ115°
ꢁ8.6
ꢁ9.1
ClMUF–G2 (7)
ClF3MUF–G2 (8)
PhUF–G2 (9)
4
ꢁ9.3
2
ꢁ10.0
ꢁ10.3
ClPhUF–G2 (10)
2
c
c
w2
e
ꢀ
Crystal structure
+1 group
/
Distance (A)
2
MUF–G2, B-mol
MUF–G2, D-mol
MUF–G2, A-mol
MUF–G2, C-mol
ClPhUF–G2, A-mol
ClPhUF–G2, B-mol
1QK2, A-molf
1QK2, B-molf
1GZ1g
MUF
MUF
Glc
ꢁ88°
+21°
+21°
3.7
3.9
2.5
2.8
3.0
2.9
2.5
2.4
2.6
3.3
ꢁ90°
ꢁ123°
ꢁ122°
ꢁ105°
ꢁ103°
ꢁ125°
ꢁ125°
ꢁ132°
ꢁ140°
ꢁ155°
ꢁ150°
ꢁ137°
ꢁ133°
ꢁ148°
ꢁ149°
ꢁ158°
ꢁ161°
Glc
Results and Discussion
ClPhUF
ClPhUF
4S-Glc
4S-Glc
4S-Glc
Glc
Predictions from automated docking and
rationale for substrate development
2BOFh
Docking with AutoDock-Vina [43] of MUF–G2 (2,
central D-glucoside in 2SO) into H. jecorina Cel6A
structure 1QJW did yield the productive pose occupy-
ing subsites ꢁ2 to +1 with a minus-anticlinal anomeric
bond: φ2 (O5′–C1′–O7umb–C7umb) = ꢁ114°, albeit only
as the 11th found pose with a calculated affinity of
ꢁ8.6 kcalꢂmolꢁ1 (Table 1). The latter must be an over-
estimation because the docking program does not take
anomeric effects into account. More importantly, the
orientation of the 4–methylumbelliferyl group within
a
Vina dockings of umbelliferyl-b–cellobioside variants into the
active site of H. jecorina Cel6A 1QJW, with the central D–glucoside
in a 2SO skew conformation; /1 (O5″–C1″–O4′–C4′) was set as un-
b
rotatable at ꢁ71°, whereas /2 was kept rotatable. Pose number
c
of the correctly docked productive pose. Dihedral angles /2 (O5′–
C1′–O7umb–C7umb) or (O5′–C1′–O/S4Glc–C4Glc) and w2 (C1′–O7umb
–
d
C7umb–C6umb
)
e
or (C1′–O/S4Glc–C4Glc–C5Glc).
Calculated affinity
(kcalꢂmolꢁ1). Glycosidic O or S distance to the ‘active’ rotamer of
f
D221 from the B chain of PDB-entry 1HGW [23]. H. jecorina
g
Cel6A wt/Glc2–S–Glc2 complex [31]. H. insolens Cel6A D416A/
h
subsite +1 (reflected by w2 (C1′–O7umb–C7umb
–
Glc2–S–Glc2 complex [32].
T. fusca Cel6A Y73S/cellotetraose
complex [44].
C6umb) = ꢁ108°) suggests two positions for substitu-
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187

Fluorogenic substrates for GH6
M. Wu et al.
predicting a near-identical orientation of the umbellif-
erone moiety in subsite +1 (reflected by similar φ2 and
w2 values; Table 1); this prompted us to carry out
their synthesis. Our H. jecorina Cel6A crystal structure
in complex with ClPhUF–G2 (10) revealed a near-
identical pose as that predicted by the Vina-docking
(Fig. S2).
sion spectra demonstrate that the background fluores-
cence of the substrate ClMUF–G2 (7) is very low,
even at 0.4 mM concentration and pH 10.2, compared
with 16 lM of the free fluorophore ClMUF (3) (Fig.
S1).
Monitoring hydrolysis of fluorogenic substrates
on UV table
Substrate synthesis
Different H. jecorina and T. fusca cellulases were
assayed for activity against the fluorogenic cellobioside
substrates by monitoring the fluorescence on a UV
table (Fig. 3). ClF3MUF–G2 (8) was less soluble and
had to be used at lower concentration (0.1 mM) than
the other substrates (0.5 mM). The substrates them-
selves did not show any visible fluorescence as seen in
the top row.
The four modified umbelliferyl b–cellobiosides were
synthesized in a short-step straightforward route with
good overall yields (Fig. 2). The 6–chloro-4–methyl-, 6
–chloro-4-trifluoromethyl-, 4–phenyl- and 6–chloro-4–
phenyl-umbelliferones (respectively 3, 4, 5 and 6) were
synthesized by Pechmann condensation in trifluoroace-
tic acid [45]; an alternative preparation of 6–chloro-4–
methylumbelliferone (3) by condensation in sulfuric
acid has been described previously, but without analyt-
ical data [46]. Aglycone coupling with acetobromo cel-
lobiose (11) to the respective peracetates 12, 13, 14
and 15 was Knorr-type in the presence of silver car-
bonate and 2,4,6–collidine [47], which was followed by
deacetylation under mild acidic conditions yielding the
respective b–cellobiosides 7, 8, 9 and 10.
The free fluorophores ClMUF (3) and ClF3MUF
(4) showed similar intensity as MUF (1) under UV
light (pH 5.0), but the fluorescence was much weaker
with PhUF (5) and ClPhUF (6). Both GH6 enzymes,
H. jecorina Cel6A (CBH) and T. fusca Cel6A (endo-
glucanase) showed clear activity against ClF3MUF–
G2 (8), while fluorescence was barely visible with
ClMUF–G2 (7), and not at all with MUF–G2 (2). The
fluorescence was weak with PhUF–G2 (9) and
ClPhUF–G2 (10), but may still indicate significant
activity due to the low intensity of the corresponding
free fluorophores. H. jecorina Cel7A (CBH) and Cel7B
(endo), and T. fusca Cel9B (endo) could hydrolyse all
substrates and gave very strong fluorescence, indicating
high degree of substrate consumption. No activity was
detected against any substrate with T. fusca Cel9A
(endo-processive).
Fluorescence properties of synthesized
compounds
The fluorescence spectra of ClMUF (3), PhUF (5) and
ClPhUF (6) showed excitation (Ex) and emission (Em)
maxima at about the same wavelengths as MUF (1,
ꢀ 360 and ꢀ 450 nm), whereas the trifluoro-methyl
group of ClF3MUF (4) caused a shift to longer wave-
lengths (385 and 495 nm; Table 2). All fluorophores
showed higher fluorescence intensity at pH 10.2 than
at pH 5.0. ClMUF (3) and ClF3MUF (4) gave
approximately three and four times higher intensity at
pH 5.0, and somewhat lower intensity at pH 10.2
( ꢀ 70% and ꢀ 20%, respectively) than MUF (1),
whereas the phenyl substituted ClPhUF (6) and espe-
cially PhUF (5) gave weaker fluorescence at pH 5.0,
and even more so at pH 10.2, than MUF (1). Emis-
Initial hydrolysis rates
Initial hydrolysis rates ([product]/[enzyme]ꢂminꢁ1) with
0.5 mM fluorogenic substrate [0.1 mM with ClF3MUF–
G2 (8)] were determined for H. jecorina Cel6A and
Cel7B and T. fusca Cel6A by following the increase of
fluorescence at 37 °C (Table 3). The GH6 enzymes did
actually hydrolyse MUF–G2 (2) at detectable, albeit
OAc
OH
Ag2CO3
2,4,6-collidine
CH2Cl2
HO
X
OH
OAc
O
OH
AcO
O
HO
O
HCl
MeOH
AcO
O
X
O
Y
O
HO
HO
O
X
O
Y
O
HO
X
O
Y
O
TFA
O
OAc
AcO
Br
O
O
O
Y
OEt
OAc
O
OAc
OH
AcO
O
AcO
AcO
OAc
OH
O
X = H, Me : 1
X = H, Y = Me : 2
X = Cl, Y = Me : 3, 60%
X = Cl, Y = CF3 : 4, 30%
X = H, Y = Phe : 5, 74%
X = Cl, Y = Phe : 6, 28%
X = Cl, Y = Me : 12, 99%
X = Cl, Y = CF3 : 13, 50%
X = H, Y = Phe : 14, 78%
X = Cl, Y = Phe : 15, 92%
X = Cl, Y = Me : 7, crude
X = Cl, Y = CF3 : 8, crude
X = H, Y = Phe : 9, 72%
X = Cl, Y = Phe : 10, 83%
O
X = H. Cl
Y = Me, CF3, Phe
OAc
OAc
11
Fig. 2. Synthesis of MUF–G2 (2), ClMUF–G2 (7), ClF3MUF–G2 (8), PhUF–G2 (9) and ClPhUF–G2 (10).
188
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M. Wu et al.
Fluorogenic substrates for GH6
Table 2. Fluorescence properties of the umbelliferone variants.
Wavelength
Fluorescence intensity
(a.u.)
maximum (nm)
No enzyme
Fluorophore
Ex
Em
pH 5.0
pH 10.2
MUF (1)
365
360
385
360
360
450
450
495
450
450
7
26
20
3
460
327
82
5
ClMUF (3)
ClF3MUF (4)
PhUF (5)
Hj Cel6A
ClPhUF (6)
0.2
2
Tf Cel6A
Hj Cel7A
Table 3. Initial hydrolysis rates of 0.5 mM fluorogenic b–cello-
biosides at 37 °C.a
Hydrolysis rate (minꢁ1 b
)
Hj Cel7B
Tf Cel9A
Tf Cel9B
H. jecorina
T. fusca
H. jecorina
Substrate
Cel6A
Cel6A
Cel7B
MUF–G2 (2)
0.001
0.078
0.176
0.024
0.012
0.006
0.192
n.d.
46
204
250
26
ClMUF–G2 (7)
ClF3MUF–G2 (8)c
PhUF–G2 (9)
0.018
0.012
ClPhUF–G2 (10)
10
10 μM
20 μM
a
pH 5.0 with H. jecorina Cel6A and Cel7B; pH 5.5 with T. fusca
b
Cel6A. Mol of fluorophore released per mol of enzyme per min-
c
ute.
The concentration of ClF3MUF–G2 (8) was limited to
ꢀ 0.1 mM due to low solubility.
100 μM
500 μM
very low, rates. In all cases, hydrolysis of the modified
umbelliferyl substrates was faster. ClF3MUF–G2 (8)
gave the highest rate increase compared with MUF–G2
(2) (150 times with H. jecorina Cel6A) followed by
ClMUF–G2 (7). HPLC analysis revealed that < 1%
glucose was released after > 10% consumption of
ClMUF–G2 (7) or ClPhUF–G2 (10), demonstrating
preferential cleavage of the heteroglycosidic bond to
the aglycone. The hydrolysis rates for the GH6
enzymes were still approximately three orders of mag-
nitude lower than with H. jecorina Cel7B. Interestingly,
also with this enzyme ClMUF–G2 (7) and ClF3MUF–
G2 (8) were hydrolysed much faster than MUF–G2 (2)
(approximately five times). Preliminary tests indicated
increased hydrolysability of the synthesized substrates
also with H. jecorina Cel5A and Cel45A, but the activ-
ity was too low for accurate determination of hydroly-
sis rates (data not shown).
ClPhUF PhUF
Fig. 3. Fluorescence on a UV table of the umbelliferone variants,
and corresponding b–cellobiosides after 1 h incubation at 40 °C with
enzymes H. jecorina Cel6A, 2 lM; Cel7A, 1 lM; Cel7B, 0.1 lM
(pH 5.0); T. fusca Cel6A, 2 lM; Cel9A, 1 lM; Cel9B, 0.1 lM (pH 5.5).
product was released within 20 min for determination
of kcat and KM for both H. jecorina Cel6A at 37 °C and
T. fusca Cel6A at 50 °C (Table 4). A lower KM value
Table 4. Enzyme kinetics constants for release of fluorophore from
ClMUF–G2 (7).a
ꢁ1
Enzyme
kcat (minꢁ1
)
KM (lM)
kcat/KM (sꢁ1ꢂM
)
Enzyme kinetics
H. jecorina Cel6Ab
T. fusca Cel6Ac
0.11 ꢃ 0.01
0.20 ꢃ 0.02
45 ꢃ 7.9
21 ꢃ 5.7
41 ꢃ 1.7
162 ꢃ 27
ClMUF–G2 (7) was chosen as the most suitable sub-
strate for measurement of enzyme kinetics, and was
used at substrate concentrations well above and below
the KM values. By stopping the reaction and enhancing
fluorescence with addition of pH 10.2 buffer, enough
a
After 20 min incubation with 1 lM enzyme, 1/5 volume of 1 M
glycine buffer, pH 10.2, was added and the fluorescence intensity
b
measured. pH 5.0 and 37 °C with H. jecorina Cel6A. pH 5.5
and 50 °C with T. fusca Cel6A.
c
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189

Fluorogenic substrates for GH6
M. Wu et al.
for T. fusca Cel6A indicates about twice as strong bind-
ing as H. jecorina Cel6A. The kcat values are not directly
comparable because of different temperatures.
A
Crystal structures of H. jecorina Cel6A D221A
substrate complexes
X–Ray crystallographic studies were with the catalytic
acid mutant D221A of H. jecorina Cel6A to avoid
rapid hydrolysis of the substrates [23]. Soaking experi-
ments were successful with MUF–G2 (2), ClMUF–G2
(7) and ClPhUF–G2 (10); the complexed structures
˚
were refined at 1.7, 1.7 and 2.2 A resolution, and Rfree
values of 0.22, 0.24 and 0.25, respectively. Statistics
from data processing and structure refinement are
summarized in SI Table S1. The location of the active
site tunnel and its occupation by ClMUF–G2 (7) and
ClPhUF–G2 (10) are shown in Fig. 4. The protein
structures are very similar (pairwise RMSD < 0.5) to
previous H. jecorina Cel6A structures, including the
N- and O–glycosylation sites on the surface. The flexi-
ble 177–183 loop near the active centre adopts the
closed conformation, as previously observed with the
D221A and Y169F mutants (PDB codes 1HGY and
1QJW), except in protein molecules A and C of the
MUF–G2 complex where loop residues 179–181 had
insufficient electron density to be resolved.
B
The MUF–G2 complexed crystal contained four
protein molecules per asymmetric unit. Molecules B
and D displayed practically identical binding of one
MUF–G2 molecule at the catalytic centre, with the cel-
lobioside moiety occupying subsites ꢁ2/ꢁ1 and the
MUF group in subsite +1, and without further ligand
electron density in subsites +2 and +3 (Fig. 5B). In the
other two protein molecules A and C, the MUF moi-
ety was found in subsite +3 and the cellobioside in
+1/+2. They also had electron density for another cel-
lobiose moiety in subsites ꢁ1 and ꢁ2, but without
either additional density from MUF at subsite +1 or
from an alpha-axial hydroxyl group on the D-glucoside
moiety occupying subsite ꢁ1. This implies that either
the ꢁ1 and +1 D-glucoses are connected, or that the
electron density is the result of overlapping binding of
cellobiose and MUF–G2 at partial occupancies. All
four subsequent D-glucoses and MUF could be con-
nected by glycosidic bonds without violation of bond
length and angle restraints. Thus we chose to include
Fig. 4. Secondary structure representation of H. jecorina Cel6A
D221A (grey) with bound ClPhUF–G2 (10, blue), and ligand from
the superimposed ClMUF–G2 (7, orange) complex (A chains from
both structures). (A) Tunnel exit view. (B) Rotated 90ꢀ. The flexible
177–183 loop (red) is closed in these structures, but is partly
disordered in chains A and C of the MUF–G2 complex.
and
refine
4–methylumbelliferyl-b–cellotetraoside
(MUF–G4) in protein molecules A and C in the final
model (Fig. 5A).
subsites +1 to +3 (ClMUF group in +3) which is not a
catalytically productive binding position (Fig. 5D).
The cellobioside moieties bound similarly, but there
was a slight deviation in the binding of the ClMUF
The other two crystals had two protein molecules in
the asymmetric unit. With the ClMUF–G2 complex,
both A and B chains showed the substrate occupying
190
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M. Wu et al.
Fluorogenic substrates for GH6
2
ety in subsite ꢁ1 adopted the previously observed SO
A
skew conformation, somewhat tilted towards a B3,O
boat, in both ClPhUF–G2 complexed protein mole-
cules (Fig. 5D), and in the MUF–G2 complexes A and
C where MUF–G4 was modelled (occupying subsites
ꢁ2 to +3 with MUF in +3; Fig. 5A). The 6–hydroxyl
group in ꢁ1 was not positioned above the D-glucose
ring as in the wild-type H. jecorina Cel6A complex
with Glc2–S–Glc2–OMe (1QK2), but had taken on a
more relaxed position pointing away from the D-glu-
cose ring, as observed with the Y169F mutant [31]
(1QJW) However, in the other two protein molecules
(B, D) of the MUF–G2 complex, with the substrate
occupying subsites ꢁ2 to +1 (MUF group in +1), the
B
C
D
4
D-glucoside in ꢁ1 adopted the C1 chair conformation
(Fig. 5B).
Comparison of MUF–G2 and ClPhUF–G2 binding
at the catalytic centre
A superposition of the observed binding modes of
MUF–G2 (2) (B chain) and ClPhUF–G2 (10) (A
chain) within the H. jecorina Cel6A crystal structures
is shown in Fig. 6. The catalytic acid is missing in
these structures due to the D221A mutation; instead
the D221 side chain was taken from the structure of
mutant D175A [23] (1HGW, B molecule), which is the
only H. jecorina Cel6A structure where the catalytic
acid is in position for protonation of the glycosidic
bond [23].
The binding mode of MUF–G2 (2) at the catalytic
centre provides clues to why it is poorly hydrolysed by
the enzyme. Although it occupies the correct ꢁ2 to +1
subsites, the D-glucoside moiety at ꢁ1 is in the unpro-
Fig. 5. Electron density for bound ligands in the crystal structures.
(A) MUF–G2 complex, chain C. (B) MUF–G2 complex, chain B. (C)
ClMUF–G2 complex, chain A. (D) ClPhUF–G2 complex, chain A.
The maximum likelihood/rA weighted 2Fobs ꢁ Fcalc electron-
4
ꢁ ꢀꢁ3
ductive ground state C1 chair conformation. The ano-
density maps were contoured at 0.45 e ꢂA
.
meric carbon, or more precisely its anti-orbital to the
glycosidic bond, is poorly accessible for nucleophilic
attack. Furthermore, the dihedral angle φ2 of the het-
erosidic anomeric bond remains in a minus-synclinal
(φ2 = ꢁ88°) position, thus the stabilizing exo-anomeric
effect is still present. The heteroglycosidic oxygen is
˚
unit, up to 1.6 A for atoms at the outer edge, most
likely because of interference by a neighbouring pro-
tein molecule (symmetry-related chain C) near the
ClMUF in chain B, which is not present in chain A.
In chain B there was also density for a single D-glucose
in subsite ꢁ2, but not in chain A. In the ClPhUF–G2
complexed structure, however, the productive pose of
the substrate was found in both protein molecules,
with almost identical binding of the cellobioside in
subsites ꢁ2/ꢁ1, the 6–chloro umbelliferyl moiety in
subsite +1, and the 4–phenyl substitution occupying
subsite +2, all as predicted by the docking (Fig. 5C).
All the D-glucose moieties found in subsites ꢁ2, +1
˚
also rather distant from the catalytic acid ( ꢀ 3.9 A)
and is not pointing a free electron pair towards it
(Fig. 6A). The orientation of the MUF group versus
subsite +1, as reflected by the dihedral angle w2 (C1′–
O7umb–C7umb–C6umb) = +21°, comes as a surprise. It is
flipped ‘upside down’ compared with the productive
pose in the docking of MUF–G2 (2) at the active site
(w2 = ꢁ108°; Table 1 and SI Fig. S2). The 4–methyl
group is not pointing towards the +2 subsite, but
instead interferes with and prevents closure of the
active-centre loop, since it overlaps with the position
of Ala178 Cb in the closed loop conformation.
4
and +2 did refine in the stable C1 chair conformation,
in similar positions and with same interactions with
the protein as described in previous H. jecorina Cel6A
structures [31] (e.g. 1QK2, 1QJW). The D-glucose moi-
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191

Fluorogenic substrates for GH6
M. Wu et al.
+1, because the chlorine atom would then clash with
C1 and/or O5 of the D-glucoside in subsite ꢁ1. Neither
A
4
can the D-glucoside at ꢁ1 relax to a C1 chair without
O2 of the ClPhUF moiety overlapping with Ala178
from any known conformation of this enzyme (regard-
less if the active-centre loop is closed or open).
Conversely, the D-glucoside in subsite ꢁ1 of the
MUF–G2 complex (B and D chains) cannot adopt the
2SO skew conformation with the current ‘upside down’
orientation of MUF because that would place O2 of
the MUF moiety in close contact with His266 NE2.
However, as shown by the docking, there is no hin-
drance for MUF–G2 (2) to bind in a flipped orienta-
tion similar to ClPhUF–G2 (10) that allows the 2SO
skew conformation at subsite ꢁ1. It should be noted,
however, that subsite +1 is too narrow to allow an um-
belliferyl group to flip once inside. Thus a MUF–G2
molecule bound in the nonproductive ‘upside down’
orientation would first need to dissociate from the
active site and rebind in the productive pose in order
to be hydrolysed.
B
C
Productive versus nonproductive binding
positions
Fig. 6. Comparison of MUF–G2 (2, pink) and ClPhUF–G2 (10, blue)
binding at the catalytic centre of H. jecorina Cel6A D221A, with
D221 reincorporated as positioned in chain B of PDB-entry 1HGW.
(A) The glycosidic oxygen of ClPhUF–G2 (10) is closer to the
catalytic acid D221, and its dihedral angle /2 is in the minus-
anticlinal position, which relieves exo-anomeric stabilization. (B)
Rotated view showing the proposed catalytic base D175 and the
two water molecules (red spheres) involved in the Grotthus
mechanism. Note that the umbelliferyl units are flipped ‘upside
down’ as reflected by the respective w2 dihedral angles. (C) The
D–glucoside unit in subsite ꢁ1 adopts a 4C1 chair with MUF–G2 (2)
and a 2SO skew with ClPhUF–G2 (10). The ligand from chain B of
the MUF–G2 complex was superimposed with chain A of the
ClPhUF–G2 structure.
The observation of two binding positions for MUF–
G2 (2), with the MUF moiety either in subsite +1 or
+3, correlates with differences in crystal packing. In
the B and D chains of the asymmetric unit, subsite +4
is blocked and +3 is partly obstructed by crystal con-
tacts (Leu230–Pro233 of chains C and A, respectively).
Apparently this has a negative impact on MUF bind-
ing at subsite +3 and shifts the binding preference of
MUF–G2 (2) in favour of the ꢁ2 to +1 position. In
protein molecules A and C, subsite +3 is fully accessi-
ble and harbours the MUF moiety, suggesting that
MUF–G2 (2) prefers to bind nonproductively at the
+1 to +3 position when the enzyme is in solution.
A similar correlation does not occur with the
ClMUF–G2 and ClPhUF–G2 complexes. Although
subsites +3 and +4 of chain A are obstructed in the
same way, while they are open in chain B, ClMUF–G2
(7) binds at the nonproductive +1 to +3 position in
both chains. By contrast, ClPhUF–G2 (10) shows the
productive pose at subsites ꢁ2 to +2 in both. Appar-
ently the 4–phenyl substitution shifts the binding pref-
erence, most likely due to its affinity at subsite +2,
because there is no steric hindrance for the ClPhUF
moiety to bind in subsites +3 to +4 in chain B. Obvi-
ously ClMUF–G2 (7) must also be able to bind pro-
ductively at ꢁ2 to +1 and is actually hydrolysed faster
than ClPhUF–G2 (10), despite the preference for non-
productive binding (Table 3).
By contrast, the ClPhUF–G2 complex shows a simi-
lar orientation (w2 = ꢁ137°) of the umbelliferyl moiety
within subsite +1 as predicted by the docking
(w2 = ꢁ115°), with the 4–phenyl group occupying sub-
site +2 (Figs 6B and SI S2). This complex shows all
the requirements for the substrate to be in a catalyti-
cally productive pose: (a) the D-glucoside moiety at
subsite ꢁ1 has adopted the pre-TS 2SO skew confor-
mation; (b) the dihedral angle φ2 of the heterosidic
anomeric bond is now in minus-anticlinal position
(φ2 = ꢁ105°) which relieves the stabilizing exo-anomeric
effect; and (c) the glycosidic oxygen is closer to the
˚
catalytic acid ( ꢀ 3.0 A) and now points a free electron
pair towards it.
Because of the C6 substitution, ClPhUF–G2 (10)
cannot bind in the ‘upside down’ orientation observed
for MUF–G2 (2) within the confined space of subsite
192
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M. Wu et al.
Fluorogenic substrates for GH6
These structures illustrate how the presence of multi-
ple subsites may allow for alternate binding positions
along the active site, which complicates enzyme kinet-
ics and ligand-binding studies. Nonproductive sub-
strate-binding modes that overlap and compete with
productive binding will lower the apparent kcat and
KM values by the same factor (1/[1 + nonproductive
Kd/productive KM]) depending on the relative affinities.
Weaker productive than nonproductive binding of
ClMUF–G2 (2) in H. jecorina Cel6A implies that the
‘true’ intrinsic kcat and KM values must be at least
twice as high as the measured apparent values
(Table 4). Apparent KM values are thus not necessarily
direct measures of binding strength for substrate at the
catalytic centre. Differences observed between enzymes
and/or substrates may as well be due to differences in
nonproductive binding at other positions along the
active site.
preferentially adopted. Modifications of the umbellife-
ryl motif by introduction of a chlorine at C6 and/or
substitution of the 4–methyl group with 4–phenyl or
4-trifluoromethyl, as predicted by substrate dockings,
restrict the conformational freedom in the confined
space of the active site and effectively turn the umbel-
liferyl group ‘upside down’, with the glycosidic bond
better positioned for protonation (Fig. 6).
The modified umbelliferyl b–cellobioside substrates
are indeed hydrolysed faster than MUF–G2 (Table 3).
The largest rate increase with GH6 enzymes, ꢀ 150
times, is obtained with ClF3MUF–G2 (8). But this
compound is only soluble up to ꢀ 0.1 mM concentra-
tion. Furthermore, the 4–phenyl substituted fluoro-
phores PhUF (5) and ClPhUF (6) show much weaker
fluorescence. Therefore, ClMUF–G2 (7) was consid-
ered most useful for enzyme kinetics measurements.
All the fluorophores exhibit stronger fluorescence at
pH 10.2 than at pH 5.0 [ ꢀ 12–fold for ClMUF (3);
Table 2], and thus the sensitivity of activity assays can
be significantly enhanced by alkali addition prior to
fluorescence measurements.
Comparison of fluorogenic substrate and
oligosaccharide binding at the catalytic centre
The configuration at the ꢁ1/+1 glycosidic bond in these
and comparable crystal structures with GH6 enzymes
is listed in Table 1. We note that with D-glucose or
thio-D-glucose at subsite +1, the φ2 angle is rotated
around ꢁ20° further away from the minus-synclinal
position compared with the ClPhUF–G2 complex.
Perhaps this is necessary for complete removal of exo-
anomeric stabilization and/or optimal alignment of a
free electron pair with the catalytic acid, which would
then explain why these fluorogenic substrates are still
hydrolysed slowly compared with oligosaccharide sub-
strates. We propose that further development of artifi-
cial substrates should strive at optimizing the fit of the
aglycone at subsite +1 such that the φ2 angle will be
restrained within the range observed when D-glucoside
is attached at subsite +1 (Table 1).
The rate of fluorophore release from the modified
substrates is still two to three orders of magnitude
lower with the tested GH6 enzymes than with H. jeco-
rina Cel7B, pointing at a potential for additional
improvements. Based on the comparison with relevant
GH6 structures with D–glucoside or thio-D–glucoside
connected at subsite +1, we speculate that further rate
enhancement may be achieved if the φ2 angle of the
scissile bond is rotated further away from the minus-
synclinal position than observed in the ClPhUF–G2
complex. However, further studies are needed to inves-
tigate this.
Materials and methods
Docking method
Protein molecule A of the H. jecorina Cel6A crystal structure
Conclusions
1QJW,
containing
methyl
4-S-b–cellobiosyl-4-thio-
b–cellobioside (Glc)2–S–(Glc)2–OMe within its ꢁ2 to +2 sub-
sites, was selected for docking experiments with AutoDock-
Vina [43] using AutoDockTools [48] for preparations of
enzyme and ligands. The calcium, water and noncovalent
ligands were removed, and the Y169F mutation was rena-
tived. The original ligand was successfully redocked as the first
This work demonstrates the successful use of substrate
docking at the active site in combination with theory
on mechanistic requirements on bond geometry for
catalysis, as guides for rational modification of the
substrate towards better complementarity with the cat-
alytic machinery of the enzyme. Our results indicate
that poor hydrolysis of MUF–G2 (2) by GH6 enzymes
is due to preferential nonproductive binding at the
active site. At least with H. jecorina Cel6A, MUF–G2
(2) prefers to bind out-of-register with the catalytic
centre (in subsites +1 to +3), and when bound in-regis-
ter at the catalytic centre a nonproductive pose is
pose found by Vina (calculated affinity ꢁ11.3 kcalꢂmolꢁ1
)
when its four anomeric bonds were set as unrotatable in their
original dihedral angles, resp. φ1 (O5″–C1″–O4′–C4′) = ꢁ72°,
φ2 (O5′–C1′–S4–C4) = ꢁ140°, φ3 (O5–C1–O4*–C4*) = ꢁ75°
and φ4 (O5*–C1*–O4Me–CMe) = ꢁ69°. The umbelliferyl–
cellobioside ligands, with the central D-glucoside in a 2SO skew
FEBS Journal 280 (2013) 184–198 ª 2012 The Authors Journal compilation ª 2012 FEBS
193

Fluorogenic substrates for GH6
M. Wu et al.
conformation, were drawn in 3D with AVOGADRO (http://
avogadro.openmolecules.net/) and were minimized with the
MMFF94 force field [49]. Dockings were with the anomeric
bond between both D-glucoside moieties always locked in the
synclinal dihedral angle as obtained after MMFF94 minimiza-
tion (φ1 (O5″–C1″–O4′–C4′) = ꢁ71°), whereas the heterosidic
anomeric bond was kept as rotatable. Vina box settings
This was done at room temperature, both at pH 5.0 and at
pH 10.2 after addition of sodium–glycine buffer.
Monitoring hydrolysis of fluorogenic substrates
on a UV table
Hydrolysis reactions were set up in 96-well plates at pH 5.0
or 5.5 and 40 °C, with 0.1 mM ClF3MUF–G2 (8) due to
limited solubility, and 0.5 mM of the other fluorogenic
cellobioside substrates, alone or with the following enzymes
and concentrations: H. jecorina Cel6A, 2 lM; Cel7A, 1 lM;
Cel7B, 0.1 lM (pH 5.0); T. fusca Cel6A, 2 lM; Cel9A,
1 lM; Cel9B, 0.1 lM (pH 5.5). The fluorescence was moni-
tored on a UV table (UV Transilluminator Biometra TI 1,
Biometra GmbH, Goettingen, Germany) and photographed
with a Kodak EDAS 290 camera (Thermo Fisher Scientific).
The picture in Fig. 3 was taken after 1 h of incubation. Free
fluorophore was included as reference at 10 and 20 lM
concentration for MUF (1), ClMUF (3) and ClF3MUF
(4) or at 10, 20 100 and 500 lM concentration for PhUF
(5) and ClPhUF (6), at pH 5.0.
˚
were: size x, y and z = 20 A centred at x = 28.5, y = ꢁ36.3
and z = 35.9; docking exhaustiveness was set at 64. Only
those poses were evaluated where the cellobioside moiety
occupied subsites ꢁ2 to ꢁ1 near identical to that in the
original ligand.
Compound synthesis
MUF–G2 (2) was synthesized as described previously [50].
Details of synthesis, analysis and compound data for the
four modified umbelliferones, respectively 6–chloro-4–
methyl- (ClMUF, 3), 6–chloro-4-trifluoromethyl- (ClF3MU
F, 4), 4–phenyl- (PhUF, 5) and 6–chloro-4–phenyl- (ClP
hUF, 6) and for the corresponding b–cellobiosides ClMUF
–G2 (7), ClF3MUF–G2 (8), PhUF–G2 (9) and ClPhUF–
G2 (10) are provided in the Supporting Information (SI).
Determination of hydrolysis rates and enzyme
kinetics parameters
Enzymes and general experimental conditions
Hydrolysis rates were measured at pH 5.0 (pH 5.5 for
T. fusca Cel6A) and 37 °C, with 0.1 mM ClF3MUF–G2 (8)
and 0.5 mM of the other cellobioside substrates, and with
H. jecorina Cel6A (1 lM), Cel7B (0.01 lM), Cel5A (1 lM),
Cel45A (2 lM) or T. fusca Cel6A (1 lM). The fluorescence
was recorded at regular intervals for 10–20 min in a micro-
plate reader (Infinite M200, TECAN, with MAGELLAN 6.55
software) at their respective Ex and Em wavelengths. The
hydrolysis rates ([P]/t/[E]) were derived from the slopes
using standard curves for the respective free fluorophores.
Enzyme kinetics parameters (kcat, KM and kcat/KM) of
H. jecorina Cel6A (1 lM; pH 5.0) and T. fusca Cel6A
(1 lM; pH 5.5) were determined using ClMUF–G2 (7) as
substrate, at concentrations ranging from < 1/5 KM to > 5
KM. After 20 min at 37 °C for H. jecorina Cel6A and
50 °C for T. fusca Cel6A the reaction was stopped by addi-
tion of 60 lL 1 M sodium glycine buffer, pH 10.2, and fluo-
rescence intensity was read (Ex 360 nm; Em 450 nm; Cary
Eclipse). The substrate consumption was < 10% in all
cases. After subtraction of background fluorescence, kcat
and KM were extracted by fitting the Michaelis–Menten
equation to the experimental data by nonlinear regression
using MMFIT of the SIMFIT package (http://www.simfit.man-
chester.ac.uk/) [55]. The efficiency constant, kcat/KM was
calculated using RFFIT of the SIMFIT package by fitting the
equation below.
Wild-type enzymes from H. jecorina for hydrolysis experi-
ments, prepared as described from culture filtrate of strain
QM9414 (ATCC 26921; formerly T. reesei) were Cel5A
and Cel7B [51], Cel6A [52], Cel7A [14] and Cel45A [53].
Enzymes Cel6A, Cel9A and Cel9B (formerly E2, E4, E1)
from T. fusca [54] were kind gifts from David B. Wilson
(Cornell University, Ithaca, NY, USA). Construction and
expression of the catalytically deficient mutant D221A of
H. jecorina Cel6A and preparation of the catalytic domain
has been described previously [23]. Unless otherwise stated,
hydrolysis experiments were carried out in 240 lL of 25 mM
sodium acetate buffer, pH 5.0 or 5.5, in black 96-well
microtiter plates with cover (NUNC, Thermo Fisher Scien-
tific, Roskilde, Denmark), and fluorescence was measured
at these conditions or after addition of 60 lL 1 M sodium
glycine buffer, pH 10.2. MUF was from Sigma-Aldrich
(St. Louis, MO, USA). All other chemicals were of analyti-
cal grade.
Spectroscopic property analysis
Fluorescence spectra of 20 lM free fluorophores MUF (1),
ClMUF (3), ClF3MUF (4), PhUF (5), ClPhUF (6) and
0.5 mM ClMUF–G2 (7) were scanned from 300 to 550 nm
wavelength in a plate reader (Cary Eclipse, Varian, Agilent
Technologies, Santa Clara, CA, USA) to determine their
respective Ex (excitation) and Em (emission) wavelength max-
ima, where relative fluorescence intensities were measured.
kcat
½Eꢅ½Sꢅ
v ¼
ꢄ
K
ð1 þ ½Sꢅ=K Þ
m
m
194
FEBS Journal 280 (2013) 184–198 ª 2012 The Authors Journal compilation ª 2012 FEBS

M. Wu et al.
Fluorogenic substrates for GH6
(PyMOL Molecular Graphics System, Version 1.5.0.4
€
Schrodinger, LLC). Atom coordinates and structure factors
HPLC analysis of hydrolysis products
Hydrolysis reactions were set up in 200 lL buffer, pH 5.0, at
37 °C, with 50 lM substrate and 1 lM enzyme (H. jecorina
Cel6A or T. fusca Cel6A). Reactions with ClMUF–G2 (7)
were incubated for 2 and 4 h, and with ClPhUF–G2 (10) for
16 h, aiming for > 10% substrate consumption. Glucose and
cellobiose in the hydrolysates were separated and quantified
by high-performance anion exchange chromatography cou-
pled with pulsed amperometric detection (HPAEC-PAD) on
a Dionex ICS-3000 HPLC system equipped with a ED40
electrode, using a PA-200 column (3 9 250 mm; Dionex,
Thermo Fischer Scientific), 4 min equilibration with 50 mM
NaOH, 20 mM sodium acetate, 5 min isocratic elution and
5 min gradient elution to 200 mM NaOH, 20 mM sodium
acetate at 0.4 mLꢂminꢁ1 flow rate. The ED40 detector was
set to use the carbohydrate quadruple waveform as provided
by the manufacturer. Free fluorophores ClMUF (3) and
ClPhUF (6) were analysed for reference, and standards of
glucose and cellobiose were used for quantification (0.5, 1,
2.5, 5, 10, 20 lM).
have been deposited with the Protein Data Bank [64] with
accession codes 4AX7 (MUF–G2 (2) soak), 4AU0
(ClMUF–G2 (7) soak) and 4AX6 (ClPhUF–G2 (10) soak).
Acknowledgements
This work was supported by the Faculty for Natural
Resources and Agriculture, Swedish University of
Agricultural Sciences; Swedish Farmers Research
Foundation (SLF); Swedish Energy Agency; through
the research program MicroDrivE
– Microbially
Derived Energy, and by Japan Society for the Promo-
tion of Science (JSPS) through fellowship to TI. Dr
Anu Koivula and Dr Laura Ruohonen, VTT Techni-
cal Research Centre of Finland, are gratefully
acknowledged for providing the H. jecorina Cel6A
D221A mutant protein, and Prof. David B. Wilson,
Cornell University, Ithaca, NY, USA, for kind gift of
T. fusca enzymes.
X-Ray crystallography
References
The Cel6A D221A catalytic domain (10 mgꢂmLꢁ1) was
crystallized by the hanging drop vapour diffusion method
[56] with 20% poly(ethylene glycol) 5000 monomethyl ether
(Fluka, Sigma-Aldrich) and 20 mM sodium Mes buffer, pH
6.0, as precipitant solution. Streak and micro-seeding were
used to improve crystal quality. Ligands were introduced
by transferring crystals to soaking drops containing 1 mM
of MUF–G2 (2), ClMUF–G2 (7) or ClPhUF–G2 (10) in
precipitant solution with 15% glycerol as a cryoprotectant.
After 1.5 h of soaking at room temperature, crystals were
flash frozen in liquid nitrogen. X–Ray diffraction data were
collected from single crystals at the European Synchrotron
Radiation Facility (ESRF; Grenoble, France) beamlines
ID14–1 and ID23–1. The datasets were reduced and scaled
using the CCP4 suite of programs [57]. The complex struc-
tures were solved by molecular replacement using MOLREP
(auto-MR), a program in the CCP4 package, and using the
previous structure of H. jecorina Cel6A D221A [23] (PDB
code 1HGY) as search model. A topology file for refine-
ment of substrate structure was obtained by the PRODRG
server (http://davapc1.bioch.dundee.ac.uk/prodrg/) [58].
Water molecules were initially added automatically using
the water picking function in ARP/wARP [59]. The struc-
tures were improved through several cycles of inspection
and rebuilding against electron density using COOT [60] and
structure refinement with REFMAC5 [61]. For cross-valida-
tion, 5% of the reflections were excluded from refinement
for calculation of Rfree. Uppsala Software Factory (http://
xray.bmc.uu.se/usf/) programs MOLEMAN2 [62] and LSQMAN
[63] were used for pdb-file handling and structure compari-
son. Protein structure images were prepared using PYMOL
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Supporting information
Additional supporting information may be found in
the online version of this article at the publisher’s web
site:
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