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Design of hairpin ribozyme variants with improved activity
for poorly processed substrates
Irene Drude
1,
*, Anne Strahl
2
, Daniel Galla
2
, Oliver Mu
¨
ller
1,
 and Sabine Mu
¨
ller
2
1 Max Planck Institute for Molecular Physiology, Department I, Dortmund, Germany
2 Ernst-Moritz-Arndt Universita
¨
t Greifswald, Institut fu
¨
r Biochemie, Germany
Introduction
In recent years, a number of ribozymes, particularly
the rather small hammerhead and hairpin ribozymes,
have been designed for cleavage of therapeutically rele-
vant targets [1–5]. Cleavage occurs at conserved sites
that first need to be identified on the target; this is
followed by adapting the sequence of the ribozyme
substrate-binding domain to specifically recognize,
bind and cleave the chosen site on the target RNA.

In vitro selection studies have allowed the identification
of hammerhead ribozymes for cleavage of sites with
altered sequences [6,7]. However, in the case of the
hairpin ribozyme, to the best of our knowledge, all
variants that have so far been designed for specific
RNA destruction cleave their substrates within the
consensus sequence 5¢-Y
)2
N
)1
*G
+1
U
+2
Y
+3
B
+4
-3¢.
We have started an effort to design a hairpin ribozyme
Keywords
cleavage; hairpin ribozyme; kinetics; ligation;
RNA
Correspondence
S. Mu
¨
ller, Ernst Moritz Arndt Universita
¨
t
Greifswald, Institut fu

¨
r Biochemie, Felix
Hausdorff Str. 4, 17487 Greifswald,
Germany
Fax: +49 (0) 3834 864471
Tel: +49 (0) 3834 8622842
E-mail:
*Present address
NOXXON Pharma AG, Max-Dohrn-Strasse
8-10, 10589 Berlin, Germany
Present address
University of Applied Sciences Kaiserslau-
tern, Campus Zweibru
¨
cken, Amerikastraße 1,
66428 Zweibru
¨
cken, Germany
(Received 26 August 2010, revised
1 December 2010, accepted 6 December
2010)
doi:10.1111/j.1742-4658.2010.07983.x
Application of ribozymes for knockdown of RNA targets requires the iden-
tification of suitable target sites according to the consensus sequence. For
the hairpin ribozyme, this was originally defined as Y
)2
N
)1
*G
+1

U
+2
Y
+3
B
+4
, with Y = U or C, and B = U, C or G, and C being the
preferred nucleobase at positions )2 and +4. In the context of develop-
ment of ribozymes for destruction of an oncogenic mRNA, we have
designed ribozyme variants that efficiently process RNA substrates at
U
)2
G
)1
*G
+1
U
+2
A
+3
A
+4
sites. Substrates with G
)1
*G
+1
U
+2
A
+3

sites
were previously shown to be processed by the wild-type hairpin ribozyme.
However, our study demonstrates that, in the specific sequence context of
the substrate studied herein, compensatory base changes in the ribozyme
improve activity for cleavage (eight-fold) and ligation (100-fold). In partic-
ular, we show that A
+3
and A
+4
are well tolerated if compensatory muta-
tions are made at positions 6 and 7 of the ribozyme strand. Adenine at
position +4 is neutralized by G
6
fi U, owing to restoration of a Watson–
Crick base pair in helix 1. In this ribozyme–substrate complex, adenine at
position +3 is also tolerated, with a slightly decreased cleavage rate. Addi-
tional substitution of A
7
with uracil doubled the cleavage rate and restored
ligation, which was lost in variants with A
7
,C
7
and G
7
. The ability to
cleave, in conjunction with the inability to ligate RNA, makes these
ribozyme variants particularly suitable candidates for RNA destruction.
Abbreviations
CPG, controlled pore glass; dNTP, deoxynucleoside triphosphate; ds, double strand; EDTA, ethylene diamine tetraacetic acid; lcaa, long chain

amino alkyl; NHS, N-hydroxy succinimidl; NTP, nucleoside triphosphate; PAGE, polyacrylamide gele electrophoresis; RP-HPLC, reversed
phase high performance liquid chromatography.
622 FEBS Journal 278 (2011) 622–633 ª 2010 The Authors Journal compilation ª 2010 FEBS
for cleavage of the CTNNB1 mRNA encoding the
proto-oncoprotein b-catenin. b-Catenin is an effector
of the canonical Wnt signaling pathway, which plays
essential roles in the regulation of cell growth, mobility
and differentiation. High intracellular concentrations
of b-catenin can induce constitutive activation of Wnt
target genes, which has been proposed to be an impor-
tant oncogenic step in cancerogenesis [8,9]. Therefore,
systematic suppression of b-catenin expression by ribo-
zyme-mediated destruction of CTNNB1 mRNA could
be a suitable way to counteract cancer development
and progression.
The hairpin ribozyme is derived from the negative
strand of the tobacco ringspot virus satellite RNA,
and catalyzes the reversible cleavage of a phosphodi-
ester bond through an S
N
2-like mechanism, leading to
characteristic products with 2¢,3¢-cyclic phosphate and
5¢-OH termini [10–12]. The minimal catalytic motif is
characterized by a two-stem structure, each stem con-
sisting of a central loop region flanked by two helices.
For catalysis, the hairpin ribozyme has to undergo
conformational changes that bring the two loops into
close proximity [13–17]. This docking process generates
a complex network of interactions between the bases
in the two loops, with a ribose zipper, hydrogen

bonds, noncanonical base pairs and a Watson–Crick
base pair between G
+1
in loop A and C
25
in loop B as
characteristic elements [18–21]. The consensus sequence
of the hairpin ribozyme, determined by site-directed
mutagenesis [22–27] and in vitro evolution methods
[25–30], defines the helical regions as being highly flexi-
ble in sequence, provided that complementarity is pre-
served. In contrast, base substitutions within the loops
strongly interfere with catalytic activity. Therefore,
suitable RNA substrates were originally supposed to
fulfill the following sequence requirements: reversible
cleavage occurs between the conserved G
+1
and N
)1
within the 5¢-Y
)
N
)1
*G
+1
U
+2
Y
+3
B

+4
-3¢ motif located
in loop A, where N can be any base, B can be cyto-
sine, guanine or uracil (with cytosine being the pre-
ferred base), and Y can be uracil or cytosine (with
cytosine preferred over uracil) (Fig. 1A). In the wild-
type hairpin ribozyme, each of these bases participates
in interactions with partner bases in the ribozyme
strand [12,21]. Therefore, mutations at these positions
could disrupt essential interactions or enforce alterna-
tive ones, with a strong input on catalytic activity.
However, previous results imply that base changes in
the ribozyme domain can compensate for changes in
the conserved bases in the substrate [31], indicating
that the hairpin ribozyme can flexibly respond to base
substitutions in the substrate. The postulated consen-
sus sequence was later refined by Berzal-Herranz and
coworkers [32], who showed that previous studies
failed to evaluate all possible combinations of nucleo-
tides surrounding the cleavage site. On the basis of the
analysis of 64 substrate variants, Pe
´
rez-Ruiz et al. [32]
demonstrated that, in addition to the wild-type
A*GUC substrate, H*GUC (H = A, C or U),
G*GUN, G*GGR (R = A or G), A*GUU and
U*GUA substrates were also sufficiently well cleaved,
although with about five-fold lower activity. When
CTNNB1 mRNA was examined, no target site fully
corresponding to the hairpin ribozyme consensus

sequence YN*GUYB could be identified. Therefore,
we decided to search for a site that keeps at least some
of the required nucleotides intact, and to design hair-
pin ribozyme variants for cleavage at this specific site.
The major criterion for defining a suitable target site
was the presence of a G
+1
immediately at the cleavage
site, because substitution of G
+1
with any of the other
natural RNA bases has been shown to completely
abolish activity [26,28,33,34]. Although substitution of
G
+1
can be compensated for by corresponding substi-
tution of C
25
in loop B, regenerating the interdomain
Watson–Crick base pair, the catalytic activity of the
resulting double mutants was rather low [31]. There-
fore, we decided to retain the essential G
+1
, and chose
a site consisting of U
)2
G
)1
*G
+1

U
+2
A
+3
A
+4
(Fig. 1A), with U
)2
,A
+3
and A
+4
being distinct from
the wild-type hairpin ribozyme substrate. According to
the nomenclature used in the study of Pe
´
rez-Ruiz et al.
[32] mentioned above, the chosen target site corre-
sponds to a G*GUA substrate, which was found to be
cleaved by the hairpin ribozyme with about three-fold
lower activity. In the context of the CTNNB1 substrate
used in this study, cleavage activity was reduced by a
factor of 60 as compared with cleavage of the typical
A*GUC substrate [35] (Table 1). In order to improve
activity to the level of that with the wild-type A*GUC
substrate, we decided to search for hairpin ribozyme
variants with base substitutions in the ribozyme strand
that might restore full activity. Furthermore, the bases
at positions )2 and +4 (not included in the study of
Pe

´
rez-Ruiz et al. [32]) also do not fully correspond to
the consensus sequence (Fig. 1A), and therefore
require additional investigation. In general, there are
two possible ways of adapting the ribozyme sequence
to a specific target sequence. Suitable ribozymes can be
developed by selection of active species from a random
library, or by rational design. For the hairpin ribo-
zyme, a number of crystal structures are available
[20,21,36–40]. Careful inspection of the crystal struc-
tures reveals that nucleobases at positions +3, +4
and )2 of the substrate strand interact only with nu-
cleobases in loop A of the ribozyme strand, without
I. Drude et al. AUG hairpin ribozyme
FEBS Journal 278 (2011) 622–633 ª 2010 The Authors Journal compilation ª 2010 FEBS 623
A C G G A
A
G
G A G -5
′
5′- G G G A G A
U G C C U U
N
G A
A
G C U C
G C
C G
U A
G C

A
G
A
A
A
C
A
C
A
U
U
A
U
A
U
G
G C
A
U G
C G
G
U
U
A U
HP-CTNNB1 N7
WT
6
U= G
7
N= A

A C G G A A
A
U G C C U U
N
G A
A
G C U C
G C
C G
U
G C
A
G
A
A
A
C
A
C
A
U
U
A
U
A
U
G
U G
A U
HP-CTNNB1 N7

–5′
G U G A G -5′
2′,3′cp-
A C G G A A
A
U G C C U U
N
G A
A
GC U C
G C
C G
U
G C
A
G
A
A
A
C
A
C
A
U
U
A
U
A
U
G

3
′
A
U G
A U
HP-CTNNB1 N7
C
C
C
A
A
G
G
A
A
G
G
N
G C
C G
U
G C
A
G
A
A
A
C
A
C

A
U
U
A
U
A
U
G
3
′
A
A U
HP-CTNNB1 N7
C
C
C
A
A
G
G
A
A
G
G
C
U
C
U
U
C

C
U
U
C
G
A
A
A
A
A
A
A
G
G
G
G
G
G
G
G
U
U
U
U
U
U
G
C
C
C

C
C
2′ ,3′cp-GUGAGUCUCUUCCUCG -5′
3
′
U
A
U
U
C
A
G
C
A
G C
A
C G
G
U
U
U
A
U
U
C
A
G
C
G C
A

C G
G
U
U
U
A
U
U
C
A
G
C
A
G C
A
C G
G
U
U
U
A
U
U
C
A
G
C
A
3′- C C C U C U
A

B
C
5′- G G G A G A
3′- C C C U C U
5′- G G G A G A
3
′- C C C U C U
U
A
G
C
U
A
G
C
G
C
G
C
5
′
A
G
C
5′
A
G
C
3′
A

C
C
C
A
A
G
G
A
A
G
G
A
G
C
3′
A
C
C
C
A
A
G
G
A
A
G
G
A
G
C

S-CTNNB1-1
S-CTNNB1-2
S-CTNNB1-3
S-CTNNB1-2
+3
+4
A
–2
6
7
11
U
A
G
N
N N
N
U
G
+3
+4
A
–2
U
Y
C
N
A
A
A

G A
G A
A
A
U
U
G
G
+3
+3
+4
+4
B
C
–2
–2
Y
C
G
G
G
G
N
N
Hairpin ribozyme
consensus sequence
Hairpin ribozyme
wild-type sequence
CTNNB1 target sequence
Fig. 1. Hairpin ribozyme variants for knockdown of CTNNB1 mRNA. (A) Sequences of the wild-type hairpin ribozyme and the consensus

sequence according to [25], and the CTNNB1 target sequence (N = A, C, G, U; B = C, G, U; Y = C, U). Cleavage sites are marked by
arrows. (B) Two-way-junction hairpin ribozymes were used for analysis of the cleavage reaction. In this process, the 20mer S-CTNNB1-1 sub-
strate, corresponding to the CTNNB1 target site, is cleaved into a 15mer S-CTNNB1-2 fragment and a 5mer product, which should rapidly
dissociate from the ribozyme strand. WT refers to a wild-type hairpin ribozyme that was adapted for recognition of the CTNNB1 substrate.
Base changes in comparison with HP-CTNNB1 N7 are shown (boxed area). (C) Hairpin ribozyme constructs for analysis of the ligation reac-
tion. Binding of the 3¢-cleavage product ⁄ ligation substrate, S-CTNNB1-2, and a second ligation substrate, S-CTNNB1-3, leads to the formation
of three-way-junction ribozymes with favored ligation properties. The substrates S-CTNNB1-1 and S-CTNNB1-2 are labeled with the fluores-
cent dye ATTO680 (indicated by the gray dot).
AUG hairpin ribozyme I. Drude et al.
624 FEBS Journal 278 (2011) 622–633 ª 2010 The Authors Journal compilation ª 2010 FEBS
being involved in interdomain interactions. On the
basis of this analysis, it seemed most straightforward
to rationally design compensatory mutations in loop A
of the ribozyme strand, and thus to develop hairpin
ribozyme variants with improved activity for the cho-
sen UG*GUAA substrate.
Results
Design of hairpin ribozymes targeting a CTNNB1
mRNA model substrate at a UG*GUAA site
Literature data show that substitution of wild-type
C
+4
by adenine strongly decreases cleavage activity,
probably because of destabilization of helix 1 as a
result of the emerging G–A mismatch [25]. However,
in vitro selection studies afforded hairpin ribozymes
with uracil instead of cytosine at position +4 in the
substrate strand, and, in addition, adenine instead of
guanine at position 6 of the ribozyme strand [25]. This
result allows for the conclusion that, essentially, a

Watson–Crick base pair is required at this location.
Thus, we replaced G
6
with uracil in the CTNNB1 ribo-
zyme, assuming that the resulting A
+4
–U
6
base pair
would restore activity.
There are no literature data available on compensa-
tory mutations for substitutions at position C
+3
.
On the contrary, it has been shown that the single
substitution C
+3
fi A strongly decreases activity [26].
Careful inspection of crystal structures of the hairpin
ribozyme–substrate complex as four-way-junction
[20,21] and minimal junction-less [36–40] structures,
however, shows that C
+3
forms a noncanonical base
pair with the nucleobase at position 7 in the ribozyme
strand, which naturally is adenine. Therefore, we
investigated whether a single base substitution at posi-
tion 7 in the CTNNB1 ribozyme can compensate for
C
+3

fi A in the substrate. Accordingly, we designed
four hairpin ribozymes carrying any of the four bases
at position 7 (hence dubbed A7, C7, G7 and U7 vari-
ants) and analyzed their cleavage and ligation proper-
ties in comparison with those of a wild-type hairpin
ribozyme that was adapted to recognize CTNNB1
RNA (Fig. 1). To analyze the cleavage efficiencies of
all hairpin ribozyme motifs, we chemically synthesized
a 20mer RNA substrate, S-CTNNB1-1, containing the
target CTNNB1 mRNA sequence and a 3¢-terminal
NH
2
-linker for postsynthetic labeling of the substrate
with ATTO680. This allows for detection and quantifi-
cation of the cleavage event on a DNA sequencer, as
shown previously [41]. The substrate and the ribozyme
form a two-way-junction structure with a 12-bp
helix 1 and a 4-bp helix 2 (Fig. 1B). According to the
model, cleavage and subsequent rapid dissociation of
the 5¢-cleavage product leads to destabilization of
helix 2. Therefore, the reaction pathway for ribozyme-
mediated cleavage of the RNA substrate can be
described as:
R þ S Ð
k
on
k
off
R Á S !
k

cleav
R Á 3
0
P þ 5
0
P
Under these conditions, the time dependence of the
cleavage product concentration typically follows [42]:
½5
0
P¼½5
0
P
1
ð1 ÀðexpÞ
ðk
ðobs;cleavÞ
ÁtÞ
Þ
Hence, the kinetic parameters k
cleav
and K
m
can be
calculated from the observed cleavage rate k
obs,cleav
at
different ribozyme concentrations [R]
o
, using the

following equation [40]:
k
obs;cleav
¼
k
cleav
½R
0
K
m
þ½R
0
with
K
m
¼
k
off
þ k
cleav
k
on
In order to use the same constructs for ligation stud-
ies, we extended the ribozyme by 14 additional nucleo-
tides at the 3¢-end. Thus, upon binding of ligation
Table 1. Kinetic parameters of HP-CTNNB1 N7-mediated cleavage of S-CTNNB1-1 and S-CTNNB1-1U
)2
C under single turnover conditions.
Amplitude values refer to maximal product yield obtained with 50-fold ribozyme excess over substrate. For ribozyme sequences compare
Fig. 1.

Ribozyme
S-CTNNB1-1 substrate S-CTNNB1-1 U
)2
C substrate
k
cleav
(min
)1
) K
m
(nM) Amplitude k
cleav
(min
)1
) K
m
(nM) Amplitude
Wild type 0.007 ± 0.0007 33 ± 3 0.34 ± 0.06 – – –
A7 0.025 ± 0.0007 20 ± 4 0.62 ± 0.02 0.17 ± 0.003 31 ± 3.1 0.6 ± 0.02
C7 0.026 ± 0.006 4.5 ± 2.5 0.64 ± 0.01 0.19 ± 0.003 8.8 ± 1.8 0.58 ± 0.008
G7 (0.005 ± 2) · 10
)5
34 ± 7 0.47 ± 0.02 0.05 ± 0.001 50 ± 6.2 0.33 ± 0.02
U7 0.057 ± 0.001 24 ± 3.6 0.86 ± 0.02 0.75 ± 0.02 29 ± 3.7 0.86 ± 0.02
I. Drude et al. AUG hairpin ribozyme
FEBS Journal 278 (2011) 622–633 ª 2010 The Authors Journal compilation ª 2010 FEBS 625
substrates, a stable ribozyme–substrate complex orga-
nized in a three-way junction results, with both liga-
tion fragments being tightly bound to the ribozyme, by
12 and 15 bp, respectively (Fig. 1C).

Intermolecular cleavage kinetics
Intermolecular single turnover cleavage kinetics were
examined for all hairpin ribozyme variants at 37 °Cin
standard buffer, containing 40 mm Tris (pH 7.5) and
10 mm MgCl
2
. The data are summarized in Fig. 2 and
Table 1. The wild-type hairpin ribozyme cleaves the
CTNNB1 substrate with k
cleav
= 0.007 min
)1
and a
final yield of 34%, showing a 60-fold reduction in the
cleavage rate constant as compared with the cognate
A*GUC substrate ($ 0.42 min
)1
[35]). The U7 variant
showed the best activity among the other four variants,
cleaving about 85% of the CTNNB1 substrate within
100 min. The determined cleavage rate constant of
0.057 min
)1
indicates an eight-fold increase as com-
pared with the wild-type ribozyme, and was only
seven-fold lower than the cleavage rate constant
obtained for wild-type cleavage of the cognate
A*GUC substrate [35]. The A7 and C7 variants
showed similar cleavage properties, with a maximal
product fraction of $ 60% after 4 h. Both ribozymes

catalyzed the cleavage reaction with a k
cleav
of about
0.025 min
)1
, indicating an increase in the cleavage rate
of only three-fold to four-fold as compared with the
wild-type hairpin ribozyme. The G7 variant did not
show any improvement in activity. Only 30% cleavage
product could be detected after 4 h. With a k
cleav
of
0.005 min
)1
, it showed equally low activity as the wild-
type ribozyme for cleavage of the CTNNB1 substrate.
As mentioned above, the CTNNB1 substrate RNA
used has a uracil instead of a cytosine at position –2.
In order to evaluate the sole influence of base substitu-
tions in the ribozyme strand, a modified CTNNB1 sub-
strate was synthesized, carrying the consensus cytosine
at position –2, and the activities of the four variants
for this substrate were tested. C
)2
in the CTNNB1
substrate increased cleavage rate constants about
10-fold in each variant as compared with cleavage of
the U
)2
substrate (Fig. 3; Table 1). The U7 variant

showed a slightly increased cleavage activity (k
cleav
=
0.75 min
)1
) as compared with the activity of the wild
type for the consensus A*GUC substrate [35]. The A7
and C7 variants catalyzed the cleavage reaction with a
k
cleav
of $ 0.2 min
)1
, indicating that the substitutions
at positions +3 (C fi A) and +4 (C fi A) are well
tolerated if adequate compensatory mutations
(G6 fi U; N7 fi A, C or U) are made. The G7 vari-
ant again showed the lowest activity, with k
cleav
=
0.05 min
)1
. All variants catalyzed cleavage of the C
)2
A7
C7
G7
U7
0 100 200 300 400 500 600 700 800
0.00
0.01

0.02
0.03
0.04
0.05
0.06
k
obs
/min
–1
Ribozyme (nM)
0 50 100 150 200 250
0.0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
0.9
A
B
Fraction of cleaved product
Reaction time (min)
Fig. 2. HP-CTNNB1 N7-mediated cleavage of S-CTNNB1-1 under
single turnover conditions. (A) Time course of reactions at 30-fold
excess of ribozyme over substrate. (B) Dependence of k
obs
values

on ribozyme concentration.
A7
C7
G7
U7
0 200 400 600 800
0.0
0.1
0.2
0.3
0.4
0.5
0.6
0.7
0.8
k
obs
(min
–1
)
Ribozyme (nM)
Fig. 3. Dependence of k
obs
values on ribozyme concentration.
Kinetic plot of HP-CTNNB1 N7-mediated cleavage of S-CTNNB1-1
U
)2
C under single turnover conditions.
AUG hairpin ribozyme I. Drude et al.
626 FEBS Journal 278 (2011) 622–633 ª 2010 The Authors Journal compilation ª 2010 FEBS

substrate with a product yield similar to that of the
U
)2
substrate, although with clearly shorter reaction
times.
Intermolecular ligation kinetics
In order to fully characterize the designed hairpin ribo-
zyme variants, we also investigated the ligation behav-
ior. In-trans ligation kinetics were measured in reactions
with ribozyme, 3¢-cleavage product ⁄ ligation substrate,
termed S-CTNNB1-2, and 5¢-ligation substrate,
S-CTNNB1-3 or S-CTNNB1-3 U
)2
C, containing a 2¢,3¢-
cyclic phosphate terminus. Binding of ligation substrates
to the ribozyme resulted in the formation of a stable
ribozyme–substrate complex, forming a three-way-junc-
tion structure (Fig. 1C). Because of the stability of this
complex, ligation should be favored over cleavage,
although cleavage cannot be neglected. Therefore, the
determined ligation rate will reflect an approach to the
equilibrium between cleavage and ligation, provided
that cleavage is much faster than dissociation of the
ribozyme–product complex. It has to be taken into
account that the observed ligation rate will be the sum
of the cleavage and ligation rates. Kinetic parameters
were determined under single turnover conditions, with
increasing concentrations of ribozyme and S-CTNNB1-
3orS-CTNNB1-3 U
)2

C with respect to the 3¢-ligation
substrate.
First, we investigated ribozyme-supported ligation of
S-CTNNB1-2 to S-CTNNB1-3 with uracil at posi-
tion )2. In contrast to cleavage analysis, where all
variants were found to be active, ligation product was
detected only for the wild type and the U7 variant
(Fig. 4), with 17% or 30% yield, respectively (Table 2).
The wild type showed very little ligation activity.
Therefore, ligation was studied only at ribozyme satu-
ration (50-fold excess of ribozyme ⁄ 5¢-ligation substrate
over 3¢-ligation substrate) to determine the correspond-
ing k
obs,lig
(Table 2). For the A7, C7 and G7 variants,
the ligation product levels were too low to be quanti-
fied. Kinetic data for three-way-junction ribozymes
are not available from the literature, but the obtained
k
app
,
lig
value for the U7 variant of 1 min
)1
(Fig. 5B;
Table 2) lies within the range of typical ligation con-
stants for two-way-junction and four-way-junction
hairpin ribozymes [41]. As observed for the wild type
with its cognate substrates [43,44], the U7 variant also
catalyzed ligation about 18 times faster than cleavage.

Therefore, the observed ligation rate constant essen-
tially reflects the ligation step, as the reverse cleavage
2 min
10 min
30 min
1 h
2 h
4 h
6 h
8 h
2 min
10 min
30 min
1 h
2 h
4 h
6 h
8 h
2 min
10 min
30 min
1 h
2 h
4 h
6 h
8 h
2 min
10 min
30 min
1 h

2 h
4 h
6 h
8 h
A7 C7 G7 U7
Fig. 4. Qualitative analysis of HP-CTNNB1 N7-mediated ligation of S-CTNNB1-2 with S-CTNNB1-3. The lower band represents the ATTO680-
labeled ligation substrate, S-CTNNB1-2, and the upper band represents the ATTO680-labeled ligation product.
Table 2. Kinetic parameters of HP-CTNNB1 N7-mediated ligation of S-CTNNB1-2 with S-CTNNB1- 3 and S-CTNNB1-3U
)2
C under single turn-
over conditions. Amplitude values refer to maximal product yield obtained with 50-fold excess of ribozyme and 5¢-ligation substrate over
3¢-ligation substrate. For ribozyme sequences, see Fig. 1. ND, not determined.
Ribozyme
S-CTNNB1-3 substrate S-CTNNB1-3 U
)2
C substrate
k
app,lig
(min
)1
) K
m
(nM) Amplitude k
app,lig
(min
)1
) K
m
(nM) Amplitude
Wild type

a
0.01 ± 0.004 0.17 ± 0.01
A7 ND ND ND 1.3 ± 0.08 105 ± 18 0.29 ± 0.007
C7 ND ND ND 0.68 ± 0.05 123 ± 27 0.32 ± 0.01
G7 ND ND ND 0.26 ± 0.01 61 ± 9 0.18 ± 0.008
U7 1 ± 0.07 380 ± 39 0.34 ± 0.01 1.5 ± 0.08 168 ± 21 0.6 ± 0.03
a
k
obs,lig
at ribozyme saturation.
I. Drude et al. AUG hairpin ribozyme
FEBS Journal 278 (2011) 622–633 ª 2010 The Authors Journal compilation ª 2010 FEBS 627
reaction is negligible. The slightly higher K
m
value
(380 ± 39 nm) may be a result of inactive ribozymes
in the solution [45].
Next, ligation activities of the four variants were
investigated on CTNNB1 substrates with cytosine
instead of uracil at position )2 (S-CTNNB1-3U
)2
C).
As observed for the cleavage reaction, ligation was
also considerably improved by this substitution: rate
constants and product yields were increased for all
four variants (Table 2). All variants showed ligation
activity, with maximal product yields of 65% (U7),
30% (A7 and C7) and 20% (G7) (Fig. 6A). Although
the U7 and A7 variants catalyzed ligation with differ-
ent amplitudes, the rate constants were similar

(1.5 ± 0.08 and 1.3 ± 0.08 min
)1
, respectively). Liga-
tion by the C7 and G7 variants was less efficient, with
k
app
,
lig
values of 0.68 ± 0.05 and 0.26 ± 0.01 min
)1
(Fig. 6B), respectively.
Interestingly, ligation data for the wild type and the
U7 variant were better fitted with a double exponential
than with a single exponential equation, whereas the
other variants showed monophasic kinetics, as expected.
Biphasic kinetics with a fast and slow phase were previ-
ously described for minimal hairpin ribozymes [46], with
the assumption that the slow phase results from inactive
ribozymes that have to undergo structural rearrange-
ment prior to cleavage ⁄ ligation. In investigations of
ribozyme–substrate complexes in native polyacrylamide
gels (data not shown), all ribozyme–substrate complexes
showed a similar band pattern, indicating that there are
no significant differences in global folding. Further
experiments addressing this question might include
0 50 100 150 200 250 300
0.0
0.1
0.2
0.3

0.4
0.5
k
obs
(min
–1
)
Ribozyme (nM)
0 50 100 150 200
250
0.00
0.05
0.10
0.15
0.20
0.25
0.30
0.35
0.40
A
B
Fraction of ligation product
Reaction time (min)
Fig. 5. HP-CTNNB1 U7-mediated ligation of S-CTNNB1-2 with S-
CTNNB1-3 under single turnover conditions. (A) Time course of the
reaction at 30-fold excess of ribozyme and S-CTNNB1-3 over S-
CTNNB1-2. (B) Dependence of k
obs
values on ribozyme concentra-
tion.

0 100 200 300 400 500
0.0
0.2
0.4
0.6
0.8
1.0
1.2
1.4
k
obs
(min
–1
)
Ribozyme (nM)
A7
C7
G7
U7
Fraction of ligation product
Reaction time (min)
0 102030405060
0.0
0.1
0.2
0.3
0.4
0.5
0.6
0.7

A
B
Fig. 6. Ligation of S-CTNNB1-2 with S-CTNNB1-3 U
)2
C catalyzed
by HP-CTNNB1 N7 under single turnover conditions. (A) Time
course of the reaction at 50-fold excess of ribozyme and
S-CTNNB1-3 U
)2
C over S-CTNNB1-2. (B) Dependence of k
obs
val-
ues on ribozyme concentration.
AUG hairpin ribozyme I. Drude et al.
628 FEBS Journal 278 (2011) 622–633 ª 2010 The Authors Journal compilation ª 2010 FEBS
time-resolved folding analysis of individual hairpin
ribozyme variants to look for differences in the folding
kinetics. For the purpose of the study presented here,
the ligation rate constant was assigned to the fast phase
of the reaction, being 0.01 ± 0.004 min
)1
for the wild
type, and 1.0 ± 0.07 min
)1
for ligation of S-CTNNB1-3
by the U7 variant and 1.5 ± 0.08 min
)1
for CTNNB1-3
U
)2

C (Table 2).
Discussion
We have developed hairpin ribozyme variants targeting
UG*GUAA sites on suitable RNA substrates. The
UG*GUAA site was chosen in the context of the
development of a hairpin ribozyme for downregulation
of CTNNB1 mRNA, encoding b-catenin, which is an
essential player in the Wnt signaling pathway, and
which, if present at high cellular concentrations, may
support cancer development and progression [8,9].
We have analyzed the cleavage and ligation properties
of a variety of hairpin ribozymes targeting short model
substrates derived from CTNNB1 mRNA. In particu-
lar, we searched for compensatory mutations in the ri-
bozyme part that are able to counterbalance the effects
of nucleobase substitutions in the substrate, with the
major focus on analysis of C
+3
fi A. The other two
changes were assumed to be less detrimental: U
)2
is
still within the frame of the consensus sequence, and
C
+4
fi A should be easily compensated for by replace-
ment of G
6
with uracil in the ribozyme strand, restor-
ing a Watson–Crick interaction in helix 1 [25].

As known from the crystal structure, C
+3
in the sub-
strate interacts with A
7
in the ribozyme strand. There-
fore, we speculated whether substitution of A
7
would
compensate for C
+3
fi A. Four hairpin ribozymes car-
rying any of the four bases at position 7 have been
prepared (N7 ribozymes) and studied in cleavage and
ligation assays.
A wild-type hairpin ribozyme that recognizes the
CTNNB1 substrate showed 60-fold lower cleavage
activity and 100-fold lower ligation activity than with
the wild-type A*GUC substrate [35,41]. A
+4
in the
substrate is well tolerated if the ribozyme contains a
uracil at position 6, restoring a Watson–Crick base
pair. This result is somewhat surprising, as an A–U
base pair at this position never seemed to have
emerged from in vitro selection experiments [25–30],
such that the nucleobase at position +4 was included
in the consensus sequence as B = C, G or U, but not
A. A substrate with A
+3

in addition to A
+4
was
accepted by all variants, with the U7 variant being the
most active. Interestingly, and in contrast to what we
had expected, C
)2
fi U showed the strongest effect on
activity. The U
)2
substrate was cleaved by all variants.
Activity decreased in the order U7 >
A7 = C7 > G7. The cleavage rate constant of the U7
variant, however, was still reduced 10-fold as com-
pared with the C
)2
substrate, which was cleaved more
rapidly by all four variants, although in the same
activity order: U7 > A7 = C7 > G7. The effect of
U
)2
on ligation was even more pronounced: whereas
ligation of the C
)2
substrate was observed with all
variants, with U7 = A7 > C7 > G7, apart from the
wild-type ribozyme, only the U7 variant could ligate
the U
)2
substrate.

These results expand hairpin ribozyme consensus
rules in different ways. On the basis of previous stud-
ies, it was concluded that the hairpin ribozyme accepts
any base except adenine at position +4 [25].
The results presented here indicate that A
+4
is toler-
ated without loss of activity, if the complementary
base is located at position 6 in the ribozyme strand,
allowing the essential Watson–Crick base pair to be
formed. Furthermore, as previously shown by Ander-
son et al. [26], the C
+3
fi A substitution almost com-
pletely abolished the cleavage activity of the wild-type
hairpin ribozyme. We did not observe such a strong
effect of C
+3
fi A on the ribozymes tested here, in
good agreement with the cleavage of G*GUA sub-
strates reported by Pe
´
rez-Ruiz et al. [32]. Apart from
the different substrate sequence, our A7 variant corre-
sponds to the sequence of the wild-type hairpin ribo-
zyme, with the only difference at position 6 being
uracil instead of guanine (Fig. 1). Apparently, the
G
6
fi U substitution, together with the altered sub-

strate sequence, not only compensates for the replace-
ment of C
+4
with adenine, but also neutralizes the
change from C
+3
to adenine. A possible explanation
for this observation may be found in the spatial situa-
tion around the A
+3
⁄ A
+4
site. In the wild-type hairpin
ribozyme complexed to its A
)1
*G
+1
U
+2
C
+3
C
+4
substrate, the C
+4
–G
6
Watson–Crick base pair stacks
upon a noncanonical base pair formed between C
+3

and A
7
, in which, according to the crystal structure,
the excocyclic amino group of C
+3
donates a proton
to ring nitrogen N1 of A
7
[20,36–40]. In the A7 ribo-
zyme, the Watson–Crick base pair is formed between
A
+4
and U
6
followed by the noncanonical base pair
A
+3
–A
7
. Adenine provides a similar Watson–Crick
edge as cytosine, and the function of the exocyclic
amino group of C
+3
as a hydrogen donor can be
basically retained by adenine. The larger nucleobase
may be tolerated because of the different nature of the
neighboring Watson–Crick base pair, which is A–U
instead of G–C. An A–U base pair is less stable than
G–C, and thus might allow the neighboring A
+3

to
I. Drude et al. AUG hairpin ribozyme
FEBS Journal 278 (2011) 622–633 ª 2010 The Authors Journal compilation ª 2010 FEBS 629
squeeze in the site originally harboring a cytosine. This
interpretation is further supported by the observation
that the U7 variant increased the cleavage rate by
another factor of two, and was the only ribozyme
among the variants with ligation activity. The spatial
situation around the A
+3
⁄ A
+4
site becomes more
relaxed in the U7 variant, because A
+3
now interacts
with the smaller uracil instead of adenine. U
7
still pro-
vides two hydrogen acceptor sites, and thus allows the
noncanonical hydrogen bond with A
+3
to be formed.
Taken together, as compared with the wild-type hair-
pin ribozyme–substrate complex, the situation has
changed from pyrimidine
+4
–purine
6
(Watson–Crick)

and pyrimidine
+3
–purine
7
(noncanonical) to purine
+4
–
pyrimidine
6
(Watson–Crick) and purine
+3
–pyrimidine
7
(noncanonical) in the U7 variant. This may be well
tolerated, owing to the ability of the modified base
pairs to provide the required base-pairing interactions
with similar spatial characteristics.
The most significant effect was observed upon
replacement of C
)2
with uracil. Both cleavage and liga-
tion activities suffered from this substitution, probably
because of the emerging wobble base pair between U
)2
and G
11
closing helix 2. This G–U wobble base pair,
located next to loop A, is presumably less capable of
stabilizing the required loop A conformation than the
regular Watson–Crick base pair that normally occurs

at this site, hampering active site chemistry. The obvi-
ous compensatory mutation of G
11
to adenine in order
to restore the Watson–Crick base pair at this position
was shown to be unable to rescue ribozyme activity
[25]. This is not surprising, as G
11
is involved in forma-
tion of the ribose zipper connecting the loop A domain
with the loop B domain. The destabilization brought
about by the G–U wobble pair influences ligation more
strongly than cleavage (the A7, C7, G7 and U7 variants
were cleavage active, but only the U7 variant showed
ligation activity), because an even more rigid conforma-
tion is required for ligation. This interpretation is given
further support by the recent finding that nucleobase
substitutions that exhibit significant levels of interfer-
ence with tertiary folding and interdomain docking
have relatively large inhibitory effects on ligation rates
while showing little inhibition of cleavage [47].
In conclusion, these results demonstrate that hairpin
ribozymes can be designed for cleavage of sites differing
from the consensus sequence, and thus extend previous
results on hairpin ribozyme specificity [32]. Moreover,
the study shows that, on the basis of careful analysis of
the available structural data, rational design can be a
straightforward and effective strategy for the develop-
ment of catalysts with changed specificity. As compared
with a full in vitro selection experiment, our rational

design study delivered functional ribozymes with less
time, material and costs. Moreover, our results demon-
strate that several changes in the substrate sequence can
be advantageous over just one base substitution, owing
to the cooperative effect of two or more base changes.
The discrepancy between cleavage and ligation activities
observed for the A7, C7 and G7 variants is a useful
property with regard to the use of ribozymes for mRNA
knockdown. Here, only cleavage is required, and liga-
tion activity is undesirable. Altogether, the results of our
study enlarge the window for application of tailor-made
ribozymes in molecular biology and medicine.
Experimental procedures
Substrate preparation
All substrates used for cleavage and ligation analysis were
chemically synthesized on a solid phase as described previ-
ously [48], with the use of phenoxyacetyl-protected phos-
phoamidites (ChemGenes, Wilmington MA, USA) and a
Gene Assembler Special synthesizer (Pharmacia, Freiburg,
Germany). For postsynthetic labeling of the 20mer cleavage
substrate and the 3¢-ligation substrate, 3¢-Amino Modifier
C-3 lcaa CPG (ChemGenes) was used as the solid phase.
Phenoxyacetyl and cyanoethyl protecting groups were
removed with a 1 : 1 mixture of 32% ammonia and 8 m
methylamine in ethanol at 65 °C for 30 min, followed by
lyophilization. Tert-butyldimethylsilyl protecting groups
were removed for 1.5 h at 55 °C in a 3 : 1 mixture of trieth-
ylamine trihydrofluoride and dimethylformamide, and the
reaction was stopped with 25% (v ⁄ v) water. RNA was pre-
cipitated with butanol and purified by PAGE on a 15 %

denaturating polyacrylamide gel. Substrates were obtained
by elution from the gel with 0.3 m sodium acetate (pH 7.0),
followed by ethanol precipitation.
For postsynthetic labeling, 10 nmol of amino-modified
oligonucleotide in 50 lL of 0.2 m sodium bicarbonate
(pH 8.0) was mixed with 100 lg of ATTO680-NHS ester
(ATTOTEC, Siegen, Germany) in 50 lL of dimethylforma-
mide. The reaction was performed for 3 h at room tempera-
ture. After ethanol precipitation, labeled oligonucleotides
were purified by RP-HPLC on a Vario Prep 250-10 Nucleo-
dur 100-5 C18 EC column (Macherey-Nagel, Du
¨
ren,
Germany), with 0.1 m tetraethylammonium acetate (pH 7.5)
and an acetonitrile gradient from 5% to 30%. Product frac-
tions were concentrated and desalted over NAP columns.
Generating RNA fragments with 2¢,3¢-cyclic
phosphate termini
The 16mer 5¢-ligation fragments were obtained by
DNAzyme 8-17-mediated (5¢-AAG AGG ATT CCA GCG
GAT CGA AAC TCA GAG AAG GAG C-3¢; Purimex,
AUG hairpin ribozyme I. Drude et al.
630 FEBS Journal 278 (2011) 622–633 ª 2010 The Authors Journal compilation ª 2010 FEBS
Grebenstein, Germany) cleavage of a chemically synthesized
25mer substrate fragment, S-FR-CTNNB1 (5¢-
GCU CCU
UCU CUG AGU GGU CCU CUU U-3¢), containing the
16mer S-CTNNB1-3 fragment (underlined). Complementary
parts between DNAzyme and substrate are in italics. To
generate the ligation fragments S-CTNNB1-3 and

S-CTNNB1-3U
)2
C with 2¢,3¢-cyclic phosphate termini, RNA
substrate, DNAzyme and Tris (pH 7.5) were mixed to give a
final concentration of 2 lm RNA, 400 nm DNAzyme and
40 mm Tris, heated for 2 min at 95 °C, and incubated for
15 min at 37 °C. After addition of magnesium chloride to a
final concentration of 90 mm, the reaction was performed
for 2 h at 37 °C. RNA was purified on a 10% denaturating
polyacrylamide gel, eluted from the gel with 0.3 m sodium
acetate (pH 7.0) at 4 °C, and precipitated with ethanol.
Ribozyme synthesis
Hairpin ribozymes were transcribed in vitro from a dsDNA
template. The DNA template was obtained from a Klenow
polymerase-mediated fill-in reaction of two synthetic prim-
ers (5¢- CTG TAC TAA TAC GAC TCA CTA TAG GGA
GAT GCC TTN GAA
GCT CAG CTG AGA AAC ACG
AAT C-3¢ and 5¢-GCT CCT TCT CTG GGT AGC TGG
TAA TAT ACC GA A TGC GAA
GAT TCG TGT TTC
TCA GCT GAG C-3¢; biomers.net, Ulm, Germany) over-
lapping at their 3¢-ends by 22 nucleotides (underlined). Both
primers and 10 · KFI buffer (500 mm Tris, pH 7.6,
100 mm MgCl
2
and 500 mm NaCl) were mixed to give final
concentrations of 2 lm each primer and 1 · KFI buffer,
heated for 2 min at 90 °C, and incubated for 15 min at
37 °C. After addition of dNTPs (Fermentas, St Leon-Rot,

Germany) to a final concentration of 500 lm and Klenow
fragment (Fermentas) to a final concentration of
0.05 U lL
)1
, the reaction was performed for 30 min at
37 °C. DNA was purified on a 10% native polyacrylamide
gel, eluted from the gel with 0.3 m sodium acetate (pH 7.0),
and precipitated with ethanol. Transcription was performed
in a reaction mixture with final concentrations of 1 lm
DNA template, 1 · transcription buffer (Fermentas), 2 mm
each NTP (Fermentas) and 0.6 U lL
)1
T7-RNA polymer-
ase (Fermentas) for 3 h at 37 °C. After phenol ⁄ chloroform
extraction, RNA was precipitated with ethanol, purified on
a 10% denaturating polyacrylamide gel, and eluted from
the gel with 0.3 m sodium acetate (pH 7.0). Salt was
removed by ethanol precipitation.
Cleavage kinetics under single turnover
conditions
For kinetic characterization of cleavage events, reactions
were carried out in 40-lL reaction volumes with final con-
centrations of 25 nm substrate, 50–750 nm ribozyme,
40 mm Tris (pH 7.5) and 10 mm MgCl
2
. Substrate and
ribozyme were mixed separately in a 20-lL volume in Tris
and MgCl
2
respectively, denaturated at 90 °C for 2 min,

and incubated for a further 15 min at 37 °C. Reactions
were started by mixing substrate and ribozyme solutions.
At suitable time intervals, aliquots of 1 lL were taken, and
the reaction was immediately stopped by addition of 19 lL
of stop mix (7 m urea and 50 mm Na-EDTA). Samples
were stored on ice before analysis. All reactions were
repeated at least twice. Samples wer e analyzed on a 15% poly-
acrylamide gel with a DNA Sequencer Long ReadIR 4200
(LI-COR Bioscience Bad Homburg, Germany); data were
processed with gene imagir 4.05. The fraction of substrate
cleaved was plotted versus time, and fitted to the single
exponential equation
½3
0
P¼Að1 À e
Àkt
Þ
where [3¢P] is the product concentration, A is the ampli-
tude, k = k
obs,cleave
, and t is the time. Standard deviations
were less than 20% in each case. To determine the enzyme
specific constants, k
cleav
and K
m
, the obtained k
obs,cleav
values were plotted versus ribozyme concentration [R]
0

and
the curve was fitted to the following equation:
k
obs;cleav
¼
k
cleav
½R
0
K
m
þ½R
0
Ligation kinetics under single turnover
conditions
Ligation reactions were carried out in 40-lL reaction vol-
umes with final concentrations of 10 nm 3¢-ligation sub-
strate, 20–500 nm 5¢-ligation substrate, 20–500 nm
ribozyme, 40 mm Tris (pH 7.5) and 10 mm MgCl
2
. First,
ribozyme was denaturated in Tris buffer for 2 min at
90 °C, and this was followed by incubation for 15 min at
37 °C. After addition of MgCl
2
, the solution was incubated
for another 15 min at 37 °C. The 5¢-ligation substrate was
then added, and the solution was incubated again for
15 min at 37 °C. The reaction was started by addition of
the 3¢-ligation substrate. After suitable periods of time,

aliquots of 1.5 lL were taken and immediately added to
8.5 lL of stop mix, and samples were stored on ice until
analysis. Analysis of the ligation reaction was performed on
a DNA sequencer as described for cleavage reactions. The
fraction of ligation product was plotted versus time and fit-
ted to single-exponential or double-exponential equations.
The single-exponential equation was:
½P¼Að1 À e
Àk
obs;lig
Át
Þ
where [P] is the product concentration, A is the amplitude,
and t is the time. The double-exponential equation was:
½P¼A
0
þ A
1
ð1 À e
Àk
1
t
ÞþA
2
ð1 À e
Àk
2
t
Þ
where A

1
and A
2
are the amplitudes of the biphasic time
course and A
0
is the starting signal; k
1
and k
2
represent the
corresponding ligation rates of the fast phase and the slow
I. Drude et al. AUG hairpin ribozyme
FEBS Journal 278 (2011) 622–633 ª 2010 The Authors Journal compilation ª 2010 FEBS 631
phase, respectively, whereby k
1
was assigned to the ligation
rate constant k
obs
,
lig
. The single turnover constants k
app
,
lig
and K
m
were obtained by fitting the k
obs,lig
values to equation

k
obs;lig
¼
k
app;lig
½R
0
K
m
þ½R
0
Acknowledgements
This work was supported by the Ju
¨
rgen Manchot
Foundation. A PhD scholarship of the Konrad-Adena-
uer Foundation to Irene Drude is gratefully acknowl-
edged.
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