A bioorthogonal small-molecule-switch system for controlling protein function in live cells

Article abstract of DOI:10.1002/anie.201403463

Chemically induced dimerization (CID) has proven to be a powerful tool for modulating protein interactions. However, the traditional dimerizer rapamycin has limitations in certain in vivo applications because of its slow reversibility and its affinity for

Full text of DOI:10.1002/anie.201403463

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Chemie
DOI: 10.1002/anie.201403463
Reversible Dimerization
A Bioorthogonal Small-Molecule-Switch System for Controlling
Protein Function in Live Cells**
Peng Liu, Abram Calderon, Georgios Konstantinidis, Jian Hou, Stephanie Voss, Xi Chen, Fu Li,
Soumya Banerjee, Jan-Erik Hoffmann, Christiane Theiss, Leif Dehmelt, and Yao-Wen Wu*
Abstract: Chemically induced dimerization (CID) has proven
to be a powerful tool for modulating protein interactions.
However, the traditional dimerizer rapamycin has limitations
in certain in vivo applications because of its slow reversibility
and its affinity for endogenous proteins. Described herein is
a bioorthogonal system for rapidly reversible CID. A novel
dimerizer with synthetic ligand of FKBP’ (SLF’) linked to
trimethoprim (TMP). The SLF’ moiety binds to the F36V
mutant of FK506-binding protein (FKBP) and the TMP
moiety binds to E. coli dihydrofolate reductase (eDHFR).
SLF’-TMP-induced heterodimerization of FKBP(F36V) and
eDHFR with a dissociation constant of 0.12 mm. Addition of
TMP alone was sufficient to rapidly disrupt this heterodime-
rization. Two examples are presented to demonstrate that this
system is an invaluable tool, which can be widely used to
rapidly and reversibly control protein function in vivo.
(FKBP) and the FKBP-rapamycin-binding (FRB) domain of
the mammalian target of rapamycin (mTOR),[3] rapamycin-
based CID has been widely used to control a variety of
protein functions including gene transcription, signal trans-
duction, post-translational protein modification, and protein
degradation.[1a,4]
However, because rapamycin also binds to endogenous
proteins, it can produce off-target effects. For example,
heterodimerization of FKBP and mTOR by rapamycin
inhibits the kinase activity of mTOR, thus leading to
undesirable biological activities including immunosuppres-
sion and induction of autophagy.[5] And, FKBP-binding
ligands such as rapamycin and FK506 have been shown to
interfere with the cellular functions of FKBP proteins, such as
regulation of intracellular calcium release.[6] Moreover, since
FKBPs are ubiquitous and abundant in mammals, they could
sequester rapamycin and attenuate its efficacy as a dimerizer.
To eliminate the off-target effects of rapamycin on the native
mTOR protein, extensive work has been done to design
improved rapamycin analogues (rapalogues) which only bind
to a mutant of the FRB domain.[7] However, these rapalogues
are not truly bioorthogonal, as they still interact with
endogenous FKBP. Moreover, chemical modifications based
on the already complex rapamycin molecule are highly
restricted. Therefore, it is not surprising that only limited
progress has been achieved in the development of selective
rapalogues. Similarly, the “bump and hole” strategy was used
to engineer a synthetic homodimerizer (SLF) which is only
recognized by the F36V mutant of FKBP.[8] More recently,
bioorthogonal CID systems which use plant hormones, such
as abscisic acid (ABA) and a gibberellin analogue (GA3-AM),
as chemical dimerizers have been introduced.[9] These devel-
opments have substantially expanded the toolkit of CID
systems.[2a,10]
Another challenge in the development of CID systems is
the reversible control of dimerization. This type of control is
necessary to dissect the complexity of many biological
processes, such as signal transduction, which are often
regulated in a reversible manner (e.g., phosphorylation and
dephosphorylation, activation and inactivation of molecular
switches, and reversible protein–protein interactions). Yet
most CID systems are essentially irreversible. For the
rapamycin CID system, this irreversibility is due to the high
affinity of rapamycin for its binding partners and the
extremely slow dissociation of the dimerization complex.[11]
Attempts to disrupt rapamycin-induced dimerization using
medium exchange or competition with FK506 have not been
very successful.[12] And, although the heterodimerization of
the tobacco 14-3-3 protein and the C-terminal domain of
M
ethods to perturb and control the activity or localization
of proteins in cells are enormously useful to probe a variety of
biological processes.[1] Chemically induced dimerization
(CID) systems have been used to bring two proteins of
interest into close proximity, and can result in modulation of
their function and perturbation of associated cellular pro-
cesses.[2] Since the discovery that the natural product rapa-
mycin induces heterodimerization of FK506-binding protein
[*] Dr. P. Liu,[+] Dr. G. Konstantinidis,[+] Dipl. S. Voss, Dr. X. Chen, F. Li,
C. Theiss, Dr. Y. Wu
Chemical Genomics Centre of the Max Planck Society
Otto-Hahn-Str. 15, 44227 Dortmund (Germany)
and
Abteilung Physikalische Biochemie, Max-Planck-Institut fꢀr mole-
kulare Physiologie, Otto-Hahn-Str. 11, 44227, Dortmund (Germany)
E-mail: yaowen.wu@mpi-dortmund.mpg.de
Homepage: http://www.cgc.mpg.de/index.php/research-groups/
rg-dr-yaowen-wu/research
A. Calderon,[+] Dr. J. Hou,[+] S. Banerjee, Dr. J. Hoffmann,
Dr. L. Dehmelt
Abteilung Systemische Zellbiologie, Max-Planck-Institut fꢀr mole-
kulare Physiologie (Germany)
A. Calderon,[+] S. Banerjee, Dr. J. Hoffmann, Dr. L. Dehmelt
Chemische Biologie, Technische Universitꢁt Dortmund
Otto-Hahn-Str. 6, 44227 Dortmund (Germany)
[+] These authors contributed equally to this work.
[**] This work was supported in part by DFG grants (no.: SPP 1623 and
SFB 642) to Y.W.W., the BMBF (grant no. 0315258) to L.D. and A.C.
and the MERCUR (grant no. Pr-2010-0022) to L.D. and A.C. We
thank James C. Hu for the kind gift of the E. coli. DHFR plasmid and
Gary Bokoch for the kind gift of the Rac1Q61L plasmid.
Supporting information for this article is available on the WWW
under http://dx.doi.org/10.1002/anie.201403463.
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tobacco H+-ATPase (PMA), induced by the natural product
fusicoccin, was reversed by medium exchange, both the
dimerization and the dissociation proceeded at relatively slow
rates.[13] Activation and deactivation of signal transmission in
cells has been shown by the reversible homodimerization of
FKBP induced by the addition of FK1012 (a homodimeric
variant of FK506) with subsequent treatment with the
competitor FK506.[2c] Similarly, FK506-induced heterodime-
rization of FKBP and calcineurin was blocked by rapamycin,
which competes for binding to FKBP.[14] However, once again,
the small molecules used in these studies bind to endogenous
proteins, thus leading to undesirable biological activities.
A recent study that employed the sequential application
of two orthogonal CID systems (the rapamycin and gibber-
ellin systems) was used to turn Rac-/PI3K-dependent mem-
brane ruffling on and off.[12b] However, because each of the
CID systems used in this study is irreversible, multiple rounds
of recruitment and dissociation are not possible with this
approach. Moreover, in addition to the off-target effects of
rapamycin, this approach is also subject to the side effects of
gibberellin, side effects which have been shown to induce cell
acidification.
Therefore, despite significant advances in the develop-
ment of CID systems, there remains a high demand for
reversible CID systems with fast reaction rates, minimal
disruption of endogenous functions, and low cytotoxicity.[2b,15]
Herein, we report a novel bioorthogonal and reversible CID
system for modulating protein function in living cells. To our
knowledge, the present work is the first to establish a bio-
orthogonal small-molecule-switch system for controlling
protein function in cells.
mammalian form of DHFR (Scheme 1).[16a] The other moiety
consists of synthetic ligand of FKBP’ (SLF’), which binds with
high affinity (subnanomolar) to the F36V mutant of FKBP
(hereafter referred to as FKBP’) and displays a 1000-fold
selectivity for this mutant over the wild-type FKBP.[8] The
TMP molecule alone is used as a competitor to dissociate the
SLF’-TMP-induced dimerization. Importantly, the selectivity
of TMP and SLF’ conjugates has been proven before in
cellular labeling studies from other groups and our group,
thus showing that those components also do not significantly
interact with other cellular components.[16b–d] Therefore, we
reasoned that SLF’-TMP should display minimal off-target
effects and should not interfere with native biological systems.
The crystal structures of the eDHFR:TMP and
FKBP’:SLFꢀ complexes provide a good structural basis for
the selection of crosslinking sites on each of the moieties, as
well as the selection of an appropriate linker length.[8,16a,17]
The 4’-O of the trimethoxybenzyl group of TMP and 3-O of
the phenyl group of SLFꢀ (Scheme 1) were chosen as cross-
linking sites since previous studies have shown that modifi-
cations at these positions do not significantly disrupt the
binding of the individual moieties to their cognate pro-
teins.[8,18] SLF’ was connected to TMP with a triethyleneglycol
(TEG) linker (Scheme 1). The synthesis of SLF’ and the SLF’-
TMP conjugate is described in the Supporting Information.
To test the ability of SLF’-TMP to induce heterodimeri-
zation, recombinant eDHFR and FKBP’ proteins were mixed
together in either the presence or the absence of an equimolar
amount of the dimerizer, and subsequently subjected to size-
exclusion chromatography (Figure 1A). A higher molecular
weight species was eluted in the presence of SLF’-TMP, thus
indicating formation of the heterodimer.
The dimerizer contains two moieties, each of which binds
to a separate bioorthogonal protein module. The trimetho-
prim (TMP) moiety has a high affinity (KI = 1 nm) for E. coli
dihydrofolate reductase (eDHFR), an 18 kDa monomeric
protein and a substantially lower affinity (KI = 4–8 mm) for the
Importantly, neither the SLF’-TMP dimerizer nor the
TMP competitor showed signs of cytotoxicity toward COS-7
cells up to a concentration of 50 mm. This is in contrast to that
of rapamycin, which displays significant cytotoxicity at 50 mm.
Scheme 1. Schematic presentation of reversible chemically induced dimerization and translocation between two subcellular sites using
a combination of SLF’-TMP and rapamycin systems. MT=mitochondria, PM=plasma membrane, POI=protein of interest.
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profile of ECFP-eDHFR and Citrine-FKBP’ with SLF’-TMP
was fitted to a binding equation, thus giving a dissociation
constant of (0.12 Æ 0.04) mm.
To test the ability of SLF’-TMP to induce heterodimeri-
zation in cells, we expressed EGFP with two copies of FKBP’
(EGFP-2 ꢁ FKBP’) and TagBFP with two copies of eDHFR
and a plasma-membrane-targeting sequence from K-Ras
(TagBFP-2 ꢁ eDHFR-CAAX) in Neuro-2a cells. Total inter-
nal reflection fluorescence microscopy (TIRF-M) was used to
observe the recruitment of EGFP-2 ꢁ FKBP’ to the plasma
membrane. About 10 mm SLF’-TMP rapidly induced dimeri-
zation in cells as observed by measuring the kinetics of EGFP-
2 ꢁ FKBP’ translocation. The half-life for maximal recruit-
ment (t1/2) was calculated to be (44 Æ 4.3) s, which is similar to
that of rapamycin-induced dimerization in cells.[12b] To con-
firm the chemically induced dimerization in cells using FRET,
we expressed mCherry-2 ꢁ FKBP’ and Citrine-2 ꢁ eDHFR-
CAAX in HeLa cells. If FRET occurs, energy transfer from
the donor molecule to the acceptor molecule will cause
a decrease in the lifetime of the donor, and can be measured
by fluorescence lifetime imaging microscopy (FLIM). Since
FLIM-based FRET measurements are insensitive to the
concentrations of the fluorophores, artifacts caused by
changes in concentration and emission intensity can be
avoided. Therefore, FLIM is a very useful technique for
monitoring protein interactions in cells.[19] Addition of 1 mm
SLF’-TMP to cells expressing Citrine-2 ꢁ eDHFR-CAAX and
mCherry-2 ꢁ FKBP’ led to a time-dependent decrease of the
fluorescence lifetime of Citrine, a decrease which was not
observed in control cells expressing Citrine-2 ꢁ eDHFR-
CAAX and mCherry (Figure 2). In those experiments,
a lower concentration of SLF’-TMP was used (1 mm). There-
fore, the association kintetics were slower compared to the
experiments shown in Figure 1B. These results suggest that
SLF’-TMP-induced heterodimerization of eDHFR and
FKBP’ also occurs in living cells. Subsequent addition of
Figure 1. SLF’-TMP induces heterodimerization of eDHFR and FKBP’
in vitro and in cells. A) Size-exclusion chromatograms of a mixture of
100 mm eDHFR and 100 mm FKBP’ in the absence (solid line) or the
presence of 100 mm SLF’-TMP (dashed line). B) Measurement of EGFP
fluorescence intensity by TIRF was used to monitor the plasma
membrane recruitment dynamics of EGFP-2ꢂFKBP’ constructs after
addition of 10 mm SLF’-TMP. Neuro-2a cells were co-transfected with
TagBFP-2ꢂeDHFR-CAAX and EGFP-2ꢂFKBP’.
This data demonstrates that the
SLF’-TMP dimerizer and the TMP
competitor are orthogonal to bio-
logical systems (see Figure S3 in
the Supporting Information).
To further confirm the effi-
ciency of SLF’-TMP-induced het-
erodimerization, we made ECFP-
eDHFR and Citrine-FKBP’ fusion
proteins. Fluorescence resonance
energy transfer (FRET) between
ECFP and Citrine was used to
assess the extent of heterodimeri-
zation. Addition of the dimer
inducer led to a dose-dependent
increase of FRET as judged by
a significant decrease of the ECFP
fluorescence intensity at l =
Figure 2. FLIM measurements of the reversible heterodimerization of eDHFR and FKBP’ in live cells.
A) Lifetime images demonstrating the reversible heterodimerization of Citrine-2ꢂeDHFR-CAAX and
mCherry-2ꢂFKBP’ in HeLa cells induced by the addition of 1 mm SLF’-TMP followed by 10 mm TMP.
B) Measurement of the average lifetime of Citrine in HeLa cells coexpressing Citrine-2ꢂeDHFR-CAAX
and mCherry-2ꢂFKBP’ (solid circles) and in HeLa cells coexpressing Citrine-2ꢂeDHFR-CAAX and
475 nm and a concomitant increase
of the fluorescence intensity at l =
527 nm upon excitation at l =
370 nm (see Figure S1 in the Sup-
porting Information). A titration mCherry (open squares) upon addition of 1 mm SLF’-TMP followed by 10 mm TMP.
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10 mm of the competitor (TMP) led to the recovery of the
fluorescence lifetime (Figure 2), thus showing that the
induced protein–protein interaction was fully reversed
(Figure 3, and see Figure S2 in the Supporting Information).
When the sequential application of the dimerizer and
competitor is combined with medium exchange, multiple
rounds of protein dimerization and dissociation can be
induced. FLIM measurements were used to detect switches
between the dimerization on and off states (Figure 3B; see
Figure S2). The rapid translocation of mCherry-2 ꢁ FKBP’
between the cytosol and the plasma membrane was also
monitored by confocal microscopy (Figure 3A).
Because rapamycin can also bind to FKBP’, it is possible
to combine the SLF’-TMP system (ST system) and the
rapamycin system to induce double relocation of the FKBP’
protein in cells. In light of this idea, we made a construct
containing mCherry with two copies of the FRB domain and
a mitochondrial localization sequence from the Listeria
monocytogenes protein ActA at its C terminus (mCherry-
2 ꢁ FRB-ActA). For COS-7 cells coexpressing TagBFP-2 ꢁ
eDHFR-CAAX, Citrine-2 ꢁ FKBP’, and mCherry-2 ꢁ FRB-
ActA, confocal microscopy revealed that these three proteins
can be found at the plasma membrane, in the cytosol and the
nucleus, and at mitochondria, respectively (Figure 4). Addi-
tion of SLF’-TMP to the cells induced translocation of
Citrine-2 ꢁ FKBP’ to the plasma membrane. And, subsequent
addition of TMP together with rapamycin led to translocation
of Citrine-2 ꢁ FKBP’ to the mitochondria, because of the
disruption of the ST-dimerization and the concomitant
formation of the rapamycin dimerization (Figure 4 and
Scheme 1).
We next tested whether the ST system could be used to
reversibly induce a biological process in cells. The signaling
activity of Rac1, a small Rho GTPase, has been shown to lead
to the formation of thin actin-based cell protrusions called
lamellipodia.[19] The post-translational prenylation of wild-
type Rac1 enables it to bind to the plasma membrane, where
it is often activated by membrane-bound guanine nucleotide
exchange factors (GEFs).[20] Once it is active, Rac1 recruits
downstream effectors to the plasma membrane, which
promote lamellipodial actin assembly.[21,22] Therefore, to
control Rac1 signaling activity in cells, we generated a con-
stitutively active Rac1 mutant which is largely cytosolic
because of the removal of its membrane anchor (Rac1[Q61L;
C189S; D190-192], hereafter referred to as Rac1Q61LD
CAAX) and we fused this mutant to the C terminus of
EGFP-2 ꢁ FKBP’. We expressed this cytosolic fusion protein
together with the plasma membrane anchored TagBFP-2 ꢁ
eDHFR-CAAX in Neuro-2a cells, a cell line which forms
extensive Rac1-dependent lamellipodia.[23] TIRF-M was used
to observe plasma membrane translocation of the
Rac1Q61LDCAAX. These cells were also co-transfected
with mCherry to facilitate detection of changes in the cell
adhesion area independent of fluorescence intensity changes
(Figure 5A,C). The reversible plasma membrane transloca-
tion of Rac1Q61LDCAAX (Figure 5B) was closely corre-
lated with a reversible increase in cell adhesion area (Fig-
ure 5C). This correlation shows that SLF’-TMP-mediated
plasma membrane targeting of Rac1Q61LDCAAX can be
Figure 3. Confocal microscopy and FLIM measurements for multiple
rounds of chemically induced protein dimerization and dissociation.
A) Confocal and lifetime images demonstrating multiple rounds of
dimerization and dissociation of EGFP-2ꢂeDHFR-CAAX and mCherry-
2ꢂFKBP’ in COS-7 cells in response to sequential treatment with 1 mm
SLF’-TMP and 1 mm TMP. Scale bar: 15 mm. B) Measurement of the
oscillating average lifetime of Citrine in HeLa cells coexpressing
Citrine-2ꢂeDHFR-CAAX and mCherry-2ꢂFKBP’ or control mCherry
produced by switching dimerization on with 2 mm SLF’-TMP and off
with 2 mm TMP (n=3 cells).
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Figure 4. Double relocation of proteins in live cells. A) Confocal microscopy images showing the translocation of Citrine-2ꢂFKBP’ between two
intracellular organelles. COS-7 cells coexpressing TagBFP-2ꢂeDHFR-CAAX, Citrine-2ꢂFKBP’ and mCherry-2ꢂFRB-ActA were treated with 1 mm
SLF’-TMP to induce plasma membrane targeting (arrow) and subsequently with 0.25 mm TMP together with 0.25 mm rapamycin to re-route Citrine-
2ꢂFKBP’ to the mitochondria (arrow head). Scale bar: 10 mm. B) Enlarged views of parts of the plasma membrane and mitrochondria.
used to acutely switch on Rac1 signaling activity and
associated cellular processes, such as the induction of cell
protrusions, in living cells. Furthermore, this signaling activity
can also be acutely switched off by addition of the TMP
competitor. Interestingly, the rapid reversal of Rac1 signaling
activity caused the cell adhesion area to shrink to a value that
was similar to that observed before addition of the dimerizer.
The ST system described here offers several advantages
over existing CID systems for the control of protein function
in cells. First, the dimerizer SLF’-TMP and the competitor
TMP molecules selectively bind to exogenously expressed
protein modules. Therefore, they are bioorthogonal, thus
exhibiting minimal interference with endogenous cellular
systems. Second, the dimerization induced by SLF’-TMP can
be rapidly disrupted by the addition of the competitor TMP,
thus making this a reversible CID system. Third, this
approach can be used to induce multiple rounds of dimeriza-
tion and dissociation. Finally, the features of the ST system
make it possible to carry out the temporally controlled,
double relocation of a protein of interest to two distinct
subcellular locations when combined with the rapamycin
system. Although this strategy is subject to the limitations of
the rapamycin system (e.g., irreversibility and off-target
effects), it nevertheless offers advantages over the recently
reported approach.[12b] It only requires three protein modules,
while the previous one requires four protein modules.
Furthermore, with the approach presented here, the initial
translocation step could be performed in a reversible way
before the irreversible rapamycin-based translocation to
a final distinct site.
In conclusion, we have developed a bioorthogonal and
reversible system of chemically induced protein dimerization.
We demonstrated that SLF’-TMP rapidly induces the hetero-
dimerization of eDHFR and FKBP’ in live cells and that this
dimerization is rapidly disrupted by addition of the compet-
itor TMP. We also demonstrated that this CID system could
be used to reversibly target a constitutively active Rac1
mutant to the plasma membrane in live cells, which led to the
rapid and reversible formation of lamellipodia. We believe
that this system possesses many useful features which will
make it an invaluable tool for controlling protein function
in vivo.
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Figure 5. Reversible induction of lamellipodia formation in cells. Neuro-2a cells were co-transfected with mCherry, TagBFP-2ꢂeDHFR-CAAX and
either EGFP-2ꢂFKBP’ (control) or EGFP-2ꢂFKBP’-Rac1Q61LDCAAX. A) TIRF-M images of mCherry before and after addition of either 10 mm SLF’-
TMP or TMP. Green arrows point to regions of reversible lamellipodium formation. B) Measurement of EGFP fluorescence intensity in the TIRF
field was used to monitor the reversible plasma membrane recruitment dynamics of EGFP-2ꢂFKBP’ constructs after addition of 10 mm SLF’-TMP
and TMP. Within 30 sec after SLF’-TMP addition, a significant increase in fluorescence intensity of EGFP-2ꢂFKBP’ or EGFP-2ꢂFKBP’-
Rac1Q61LDCAAX was observed (n=3 cells; *: p<0.05; ***: p<0.001; Student’s t-test). C) Morphometric analysis of cell adhesion area
corresponding to the recruitment kinetics shown in (B). D) Representation of average EGFP fluorescence intensity and average cell area
measurements from panel (C) during the time periods before addition of SLF’-TMP, with addition of SLF’-TMP, and with addition of TMP (n=3
cells; *: p<0.05; **: p<0.01; ***: p<0.001; One-way ANOVA with Tukey’s multiple comparison test. All significant changes are marked with
brackets).
Received: March 19, 2014
Revised: June 23, 2014
Published online: July 25, 2014
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