Site-specific fluorogenic protein labelling agent for bioconjugation

Article abstract of DOI:10.3390/biom10030369

Many clinically relevant therapeutic agents are formed from the conjugation of small molecules to biomolecules through conjugating linkers. In this study, two novel conjugating linkers were prepared, comprising a central coumarin core, functionalized with a dimaleimide moiety at one end and a terminal alkyne at the other. In our first design, we developed a protein labelling method that site-specifically introduces an alkyne functional group to a dicysteine target peptide tag that was genetically fused to a protein of interest. This method allows for the subsequent attachment of azide-functionalized cargo in the facile synthesis of novel protein-cargo conjugates. However, the fluorogenic aspect of the reaction between the linker and the target peptide was less than we desired. To address this shortcoming, a second linker reagent was prepared. This new design also allowed for the site-specific introduction of an alkyne functional group onto the target peptide, but in a highly fluorogenic and rapid manner. The site-specific addition of an alkyne group to a protein of interest was thus monitored in situ by fluorescence increase, prior to the attachment of azide-functionalized cargo. Finally, we also demonstrated that the cargo can also be attached first, in an azide/alkyne cycloaddition reaction, prior to fluorogenic conjugation with the target peptide-fused protein.

Full text of DOI:10.3390/biom10030369

Article
Site‐Specific Fluorogenic Protein Labelling Agent for
Bioconjugation
Kelvin K. Tsao, Ann C. Lee, Karl É. Racine and Jeffrey W. Keillor *
Department of Chemistry and Biomolecular Sciences, University of Ottawa, Ottawa, ON K1N 6N5, Canada;
ktsao072@uottawa.ca (K.K.T.); alee189@uottawa.ca (A.C.L.); kraci097@uottawa.ca (K.É.R.)
* Correspondence: jkeillor@uottawa.ca; Tel.: +01‐613‐562‐5800 (ext. 6314)
Received: 28 January 2020; Accepted: 25 February 2020; Published: 28 February 2020
Abstract: Many clinically relevant therapeutic agents are formed from the conjugation of small
molecules to biomolecules through conjugating linkers. In this study, two novel conjugating linkers
were prepared, comprising a central coumarin core, functionalized with a dimaleimide moiety at
one end and a terminal alkyne at the other. In our first design, we developed a protein labelling
method that site‐specifically introduces an alkyne functional group to a dicysteine target peptide
tag that was genetically fused to a protein of interest. This method allows for the subsequent
attachment of azide‐functionalized cargo in the facile synthesis of novel protein‐cargo conjugates.
However, the fluorogenic aspect of the reaction between the linker and the target peptide was less
than we desired. To address this shortcoming, a second linker reagent was prepared. This new
design also allowed for the site‐specific introduction of an alkyne functional group onto the target
peptide, but in a highly fluorogenic and rapid manner. The site‐specific addition of an alkyne group
to a protein of interest was thus monitored in situ by fluorescence increase, prior to the attachment
of azide‐functionalized cargo. Finally, we also demonstrated that the cargo can also be attached first,
in an azide/alkyne cycloaddition reaction, prior to fluorogenic conjugation with the target peptide‐
fused protein.
Keywords: protein labelling; heterobifunctional; fluorogenic; CuAAC
1. Introduction
A number of clinically relevant therapeutic agents have been formed from the covalent
attachment of synthetic polymers, drugs, and imaging probes to proteins or antibodies [1,2]. These
protein‐polymer or antibody‐drug conjugates are formed through a number of linking strategies that
connect the biomolecule to cargo. Appending cargo to these biomolecules relies on highly efficient
and bioorthogonal “click” type reactions such as Michael addition, copper‐catalyzed azide‐alkyne
cycloaddition (CuAAC), strained‐promoted azide‐cycloalkyne cycloaddition (SPAAC) and the
Staudinger ligation [3–5]. The link between the protein and the cargo can be non‐site specific, as in
the case of modifying natural amino acids, or highly site specific, as is the case with labelling
genetically fused tags or introducing unnatural amino acids [6,7]. Both the reactions used and the
degree of site‐specificity of the conjugation must be considered and optimized in order to effectively
build new protein conjugates.
Bioconjugates are most commonly synthesized by appending cargo to a protein or antibody
through a linker that provides a site of attachment for the intended cargo. Examples of relevant cargo
include polyethylene glycol [8–11], to improve pharmacokinetic properties, or a small molecule drug
[12,13], for targeted delivery. Traditionally, the conjugation is monitored discontinuously (or ex situ),
using methods such as sodium dodecyl sulfate‐polyacrylamide gel electrophoresis (SDS PAGE), high
throughput liquid chromatography (HPLC), or mass spectrometry. More recently developed linkers
Biomolecules 2020, 10, 369; doi:10.3390/biom10030369
www.mdpi.com/journal/biomolecules

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allow for the continuous (or in situ) monitoring of the bioconjugation event by fluorescence, due to
their fluorogenic nature. Many of these linkers have been reviewed by Liu and coworkers, but three
significant methods stand out [14]. Most recently, Liu reported a fluorogenic linker that allowed the
attachment of cargo through CuAAC on one end of the linker and reaction with a target protein’s
native Cys thiol group through a Michael addition on the other [15]. O’Reilly et al. presented the
fluorogenic conjugation of PEG onto salmon calcitonin (sCT) through an N‐
propargyldibromomaleimide linker [16]. Lastly, Cornelissen and coworkers developed a technique
that conjugated PEG cargo to azide functionalized bovine serum albumin (BSA) through a
fluorogenic 3‐azidocoumarin linker [17]. However, drawbacks of the above linkers include
dependence on the reactivity of an intrinsic Cys residue, reversible conjugation or the requirement of
specific techniques to introduce unnatural amino acids into the protein. Thus, there remains a need
for complementary methods that will expand the scope of biomolecules that can be easily and
efficiently conjugated.
The modular design of novel protein‐cargo conjugates requires a linker that can be easily
integrated into any protein. Furthermore, the conjugation method should also be site specific, so as
not to disrupt the native function of the protein itself. Lastly, it would be advantageous for the linker
to be fluorogenic such that the conjugation can be monitored in situ. Thus, we propose a fluorogenic
linker that reacts site‐specifically with a small peptide tag genetically fused to either terminus of a
target protein. This approach eliminates the necessity of incorporating unnatural amino acid residues
into a protein of interest, while also providing a specific site of modification that can be easily
appended onto any protein with minimal disruption of that protein’s function.
In previous work, we have developed a fluorogenic specific protein labelling method based on
the conjugation between a pro‐fluorescent probe and a small α‐helical peptide genetically fused to
the protein of interest [18,19]. The fluorogenic probes (referred to as FlARe probes) consist of a
fluorophore covalently attached to two maleimide groups that quench the fluorophore’s emission
through a photoinduced electron transfer (PeT) mechanism [20,21]. These bismaleimide probes have
also been shown to react highly efficiently with the two activated cysteine thiols, located 10 Å apart,
on a rationally designed α‐helical target peptide (dC10*) in a Fluorogenic Addition Reaction (FlARe)
[22,23]. When both maleimides react with the chemically tuned thiols, the PeT quenching is abolished,
and the intrinsic fluorescence of the FlARe probe is restored.
Herein, we report a method for the site‐specific and fluorogenic introduction of an alkyne
functional group onto a specific protein. This method uses a heterobifunctional pro‐fluorescent linker
that takes advantage of the FlARe protein labelling technology (Scheme 1). The proposed linker
consists of a coumarin fluorophore core, functionalized on one end with a dimaleimide moiety. These
maleimide functional groups quench the coumarin’s fluorescence and allow the linker to react with
the dC10* tag that is genetically fused to a protein of interest. This coumarin FlARe probe is
additionally functionalized with a terminal alkyne on the other end, such that it modifies a specific
protein bearing the dC10* tag by site‐specifically introducing an alkyne group onto the exposed
terminal target peptide sequence. Since the reaction between the linker and the dC10* tag is
fluorogenic, the initial conjugation can be monitored in situ by fluorescence increase. After the initial
labelling reaction is complete, the modified protein bearing the alkyne functional group can then be
conjugated to a broad array of azide‐functionalized cargo through CuAAC. Overall, this novel linker
allows for a facile and effective method for the in vitro synthesis of fluorescent protein‐conjugates
that is easily amenable to engineered proteins of therapeutic significance.

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Scheme 1. A protein‐cargo conjugate is formed by first labelling a protein with a heterobifunctional
fluorogenic coumarin linker (A). Azide‐functionalized cargo (B) is then attached to the protein
through the alkyne moiety on the fluorogenic probe. Thus, fluorogenic and site‐specific
bioconjugation is achieved.
2. Materials and Methods
2.1. Expression and Purification of Test Protein
The plasmid coding for the tagged test protein maltose binding protein (MBP‐dC10*) was
designed and created in a previously published study in which the reactivity of the original dC10 tag
was improved through the introduction of positively charged residues around the cysteines [24]. The
expression and purification of MBP‐dC10* followed the protocol from the same study and consisted
of transforming the expression plasmid into the BL21‐Gold (DE3) strain of E. coli cells [24]. Expression
of MBP‐dC10* was induced with IPTG (0.3 mM) and purification was performed an amylose resin
column and elution with 10 mM maltose. According to the Bradford Assay, protein yields generally
ranged around 12 mg from 500 mL of expression culture.
2.2. Determination of Fluorescence Enhancement Ratios
Emission spectra and fluorescence intensity measurements were recorded at 25 °C with a
Synergy H4 Hybrid Multi‐Mode Microplate Reader with excitation and emission monochromators
set at 9‐nm bandpass. The initial excitation and emission spectra were recorded for a solution of 10
μM labelling reagent (compounds 1, 6, 10, or 15) in 50 mM HEPES (I = 100 mM with NaCl, pH 7.4)
with 10% DMSO and 50 μM TCEP. A solution of 10 μM labelling reagent in 50 mM HEPES (I = 100
mM with NaCl, pH 7.4) with 10% DMSO and 50 μM TCEP with the addition of 10 μM MBP‐dC10*
was allowed to react for 4 h (in the case of 1 or 15) or 24 h (in the case of 6 or 10). After the reaction
was complete, the final excitation and emission spectra were recorded. The ratio of fluorescence
intensity at the maximum emission between the initial and final spectra gave the fluorescence
enhancement (FE).
2.3. Kinetic Studies
Kinetic experiments were carried out at 25 °C using a Synergy H4 Hybrid Multi‐Mode
Microplate Reader with excitation and emission monochromators set at a 20‐nm bandpass. Solutions
containing 10 μM MBP‐dC10* in 50 mM HEPES buffer (I = 100 mM, pH 7.4) with 10% DMSO and 50
μM TCEP were prepared in a 96‐well plate. Labelling reagent was added immediately before
recording to a final concentration of 10 μM. Samples were excited at 435 nm for 10 and 15 or 415 nm
for 1 and fluorescence intensity was followed at 485 nm for 10 and 15 or 460 nm for 1 as a function of
time. All time‐dependent fluorescence curves were fitted to a second order rate equation to obtain
the second order rate constant (k ) for the reaction between the label and MBP‐dC10*.
2
2.4. Bioconjugation of Rhodamine B Azide to MBP‐dC10*‐15 or 6 Adduct
The protocol for performing the CuAAC between rhodamine B azide (RhB‐N ) and MBP‐dC10*
3
labelled with 6 or 15 was adapted from procedures developed by Nasheri, et al. and Yang, et al.
[25,26]. An aliquot of 5 μM 15 was incubated with 2.5 μM MBP‐dC10* and 50 μM TCEP in HEPES

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buffer (50 mM HEPES (I = 100 mM NaCl), pH 7.4) at 25 °C and a total volume of 200 μL for 1 h. The
reaction was monitored using a Synergy H4 Hybrid Multi‐Mode Microplate Reader with excitation
(435 nm) and emission (485 nm) monochromators set at a 20‐nm bandpass. After complete reaction,
the mixture was transferred to a 500‐μL 10‐kDa MWCO Amicon filter and buffer exchanged into
fresh HEPES buffer three times. After the third time, the solution was adjusted to 100 μL. A 500‐μL
click mixture (1.0 mM CuSO4, 100 μM TBTA, 1.0 mM TCEP‐HCl, 50 μM RhB‐N3, PBS pH 7.4, 0.25%
DMSO) was prepared and 100 μL of this click mixture was added to the buffer exchanged solution
containing the MBP‐dC10*‐15 adduct. The click reaction was allowed to progress for 2 h in the dark
with gentle shaking at 25 °C. After this, 500 μL of cold acetone (chilled to −80 °C) was added and the
protein was precipitated overnight at ‐80 °C. The protein was pelleted by spinning at 14,000 rpm for
15 min at 4 °C before removing the supernatant and drying off any excess acetone under a gentle
stream of air. After, the pellet was washed with 100 μL MeOH chilled to −80 °C before pelleting again
at 14,000 rpm for 15 min at 4 °C. The precipitate was then taken up in 35 μL SDS PAGE loading buffer
(50 mM TCEP) and shaken for 15 min at room temperature. The samples were spun at 14,000 rpm for
5 min and then boiled at 90 °C for 5 min before 10 μL was loaded onto a 10%‐acrylamide SDS‐PAGE
gel and run for 1.5 h at 120 V.
In the case of 6, the reaction between MBP‐dC10* and 6 was allowed to react for 24 h and the
reaction was not monitored.
2.5. Bioconjugation of Rh‐B‐N3‐15 Adduct to MBP‐dC10*
The protocol for performing the CuAAC between Rhodamine B azide (RhB‐N3) and 15 was
adapted from procedures developed by Nasheri, et al. and Yang, et al. [25,26]. A 500‐μL click mixture
(1.0 mM CuSO4, 100 μM TBTA, 1.0 mM TCEP‐HCl, 50 μM RhB‐N3, PBS pH 7.4, 0.25% DMSO) was
first prepared and 90 μL of this click mixture was incubated with 10 μL of a 200 μM 15 stock (final
concentration 20 μM). The click reaction was allowed to progress in the dark at r.t. for 2 h with gentle
shaking. After, 100 μL of a 10‐μM MBP‐dC10* solution was added to the mixture and the entire
solution was allowed to react for 1 h at r.t. Next, 500 μL of cold acetone (chilled to −80 °C) was added
and the protein was precipitated overnight at −80 °C. The protein was pelleted by spinning at 14,000
rpm for 15 min at 4 °C before removing the supernatant and drying off any excess acetone under a
gentle stream of air. The pellet was washed with 100 μL MeOH chilled to −80 °C before pelleting
again at 14,000 rpm for 15 min at 4 °C. The precipitate was taken up in 35 μL SDS PAGE loading
buffer (50 mM TCEP) and sonicated for 3 minutes prior to being shaken for 15 min at room
temperature. The samples were spun at 14,000 rpm for 5 min and then boiled at 90 °C for 5 min before
5 μL was loaded onto an 10% acrylamide SDS‐PAGE gel and run for 1.5 h at 120 V.
2.6. Synthesis
All reagents and solvents for reactions were used as received unless otherwise stated.
Dichloromethane, dimethylformamide, methanol and acetonitrile were dried using a solvent
purification system from LC Technology Solutions Inc (Salisbury, USA).
Reactions were monitored by thin layer chromatography (TLC) using E. Merck silica gel 60F254
precoated aluminum plates. Components were visualized by illumination with a short‐wavelength
ultraviolet light or long‐wavelength visible light. Flash column chromatography was performed on
®
ZEOCHEM silica gel 60 (ECO 40–63 μm) using ethyl acetate/n‐hexane or methanol/dichloromethane
as eluting solvents.
Nuclear magnetic resonance (NMR) spectra were recorded in deuterated DMSO or deuterated
chloroform at ambient temperature unless otherwise stated. The experiments were performed either
1
on a Bruker Avance 400 Fourier Transform Spectrometer operating at 400 MHz for H and at 100
13
1
MHz for C or on a Bruker Avance 600 Fourier Transform Spectrometer operating at 600 MHz for H
13
and at 150 MHz for C. EI‐MS spectra were recorded on a Kratos Concept mass spectrometer for both
low resolution and high‐resolution mass spectra. ESI‐MS spectra were recorded on a Waters
Micromass Q‐TOF mass spectrometer. Melting points were measured on an EZ‐Melt automated

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melting point apparatus and are uncorrected. Ultraviolet absorbance spectra and fluorescence
spectroscopic studies were performed on a Synergy H4 Hybrid Multi‐Mode Microplate Reader.
Compound 5: Compound 5 was synthesized according to the procedures laid out by Chen and
coworkers [19].
Compounds 2 and 3 were synthesized according to the procedures laid out by Abdizadeh and
coworkers [27].
Compound 2: To a solution of 4‐hydroxysalicylic acid (1.38 g, 10.0 mmol) and diethylmalonate
(1.0 eq, 1.53 mL, 10.0 mmol) in EtOH (5 mL) was added piperidine (0.02 eq, 19.8 μL, 0.2 mmol) and
acetic acid (0.02 eq, 11.4 μL, 0.2 mmol). The solution was stirred at room temperature for 20 min and
then heated to 78 °C and stirred for 3 h. The reaction was cooled to room temperature and the product
1
was crystallized as an orange solid in 81% yield. H NMR (400 MHz, DMSO) δ 11.04 (s, 1H), 8.73–8.53
(m, 1H), 7.71 (d, J = 8.6 Hz, 1H), 6.80 (dd, J = 8.6, 2.3 Hz, 1H), 6.75–6.47 (m, 1H), 4.22 (q, J = 7.1 Hz, 2H),
13
1.25 (t, J = 7.1 Hz, 3H). C NMR (100 MHz, DMSO) δ 164.5, 163.4, 157.6, 156.9, 149.9, 132.6, 114.5,
+
112.6, 110.9, 102.3, 61.3, 14.6. HRMS (ESI): calcd for C12H10O5Na ([MNa] ): 257.0426, found: 257.0414.
Compound 3: Compound 2 (50 mg, 0.159 mmol) was dissolved in acetonitrile (3.0 mL), to which
a solution of NaOH (10.0 eq, 63.6 mg, 1.59 mmol) dissolved in water (2.0 mL) was added dropwise.
The solution was stirred at room temperature for 3.5 h after which it was diluted with 10 mL water
and then acidified with 10 mL 1 M HCl. The aqueous solution was extracted with EtOAc (3 × 30 mL)
and the combined organic layers were dried with MgSO4. After removing the solvent on the rotary
1
evaporator, compound 3 was obtained as a yellow solid in quantitative yield. H NMR (400 MHz,
13
DMSO) δ 12.00 (bs, 1H), 8.64 (s, 1H), 7.73 (d, J = 8.4 Hz, 1H), 6.83 (d, J = 8.4 Hz, 1H), 6.72 (s, 1H).
C
NMR (100 MHz, DMSO) δ 164.32 (d, J = 14.1 Hz), 157.75, 156.99, 148.96, 131.91, 114.17, 112.69, 110.53,
13
101.83. C NMR (100 MHz, DMSO) δ 164.3, 157.8, 157.0, 149.0, 131.9, 114.2, 112.7, 110.5, 101.8. HRMS
+
(ESI): calcd for C10H6O5Na ([MNa] ): 229.0113, found: 229.0128.
Compound 1: To a solution of compound 3 (26.0 mg, 0.130 mmol) in anhydrous DMF (2.5 mL)
was added HBTU (1.1 eq, 52.0 mg, 0.138 mmol). The mixture was stirred at room temperature for 10
min before 5 (1.5 eq, 59.0 mg, 0.188 mmol) was added. The mixture was stirred for an additional 20
min before trimethylamine (3.0 eq, 50 μL, 0.380 mmol) was added. It was then stirred overnight
followed by diluting the mixture in EtOAc (100 mL) and washing with 1 M HCl (2 × 100 mL). The
organic layer was dried with MgSO4 before the solvent was removed using a rotary evaporator. The
crude was purified by flash chromatography using a gradient elution system of 0%–5% MeOH in
1
DCM to obtain compound 1 as a pale‐yellow solid in 7% yield. H NMR (600 MHz, DMSO) δ 11.18
(bs, 1H), 10.86 (s, 1H), 8.83 (s, 1H), 7.86 (d, J = 8.7 Hz, 1H), 7.78 (d, J = 1.7 Hz, 2H), 7.12 (t, J = 1.8 Hz,
1H), 6.92 (dd, J = 8.6, 2.2 Hz, 1H), 6.85 (d, J = 2.0 Hz, 1H), 6.83 (q, J = 1.8 Hz, 2H), 2.09 (d, J = 1.7 Hz,
13
6H). C NMR (150 MHz, DMSO) δ 170.3, 169.4, 164.1, 161.2, 160.7, 156.5, 149.6, 148.4, 145.9, 138.7,
132.6, 132.3, 127.6, 120.0, 116.8, 114.6, 114.1, 111.2, 102.0, 10.9. HRMS (ESI): calcd for C26H17N3O8Na
+
([MNa] ):522.0913, found: 522.0929. mp: decomp. at 267.7‐271.1 °C.
Compound 7: The synthesis of compound 7 was done according to Jiang, et al. [28]. A Finkelstein
reaction was performed to synthesize 7 in which 6‐chloro‐1‐hexyne (8.30 mmol, 1.00 mL) was added
to dry acetone (100 mL) along with NaI (58.1 mmol, 8.71 g, 7 eq). The solution was brought to reflux
and stirred overnight after which the acetone was removed using a rotary evaporator. The residue
was then partitioned between hexanes (100 mL) and water (100 mL) and the organic layer was
washed with water (100 mL) and brine (100 mL). The organic layer was then dried with MgSO4 and
the solvent was removed using a rotary evaporator to obtain compound 7 as a transparent liquid in
1
94% yield (7.78 mmol, 1.62 g). H NMR (400 MHz, CDCl3) δ ppm 3.20 (t, J = 6.91 Hz, 2 H), 2.22 (td, J =
6.98, 2.60 Hz, 3 H), 1.95 (m, 3 H), 1.63 (tt, J = 7.30 Hz, 3 H).
Compound 8: Compound 8 (1.00 mmol, 234 mg) was dissolved in acetonitrile after which
compound 7 was added. K2CO3 (1.2 mmol, 165 mg, 1.2 eq) and a catalytic amount of 18‐crown‐6 (0.1
mmol, 26.4 mg) was then added to the reaction mixture, which was brought to reflux and stirred
overnight. The acetonitrile was removed using a rotary evaporator and the residue was partitioned
between EtOAc (20 mL) and water (20 mL). The aqueous layer was extracted with EtOAc (3 × 30 mL)
and the combined organic layers were dried with MgSO4 and the solvent was removed. The residue

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was purified by flash chromatography using a gradient elution system of 10%–30% EtOAc in
1
hexanes. Compound 8 was obtained as a white solid in 44% yield (0.437 mmol, 138 mg). H NMR (400
MHz, CDCl3) δ ppm 8.50 (s, 1 H), 7.49 (d, J = 8.72 Hz, 1 H), 6.88 (dd, J = 8.67, 2.20 Hz, 1 H), 6.80 (d, J =
2.06 Hz, 1 H), 4.40 (q, J = 7.15 Hz, 2 H), 4.08 (t, J = 6.22 Hz, 2 H), 2.30 (td, J = 6.91, 2.55 Hz, 2 H), 1.97
13
(m, 3 H), 1.74 (tt, J = 7.30 Hz, 2 H), 1.40 (t, J = 7.10 Hz, 3 H). C NMR (100 MHz, CDCl3) δ ppm 164.6,
163.5, 157.6, 157.2, 149.0, 130.7, 114.0, 114.0, 111.6, 100.8, 83.7, 68.9, 68.3, 61.7, 27.9, 24.9, 18.1, 14.3.
+
HRMS (ESI): calcd for C18H18O5Na ([MNa] ): 337.1052, found: 337.1074. mp: 92.6–93.6 °C.
Compound 9: To a solution of 8 (0.437 mmol, 138 mg) dissolved in acetonitrile, NaOH (4.93
mmol, 197 mg, 10 eq) was added as a 2 M solution in water (2.5 mL). The reaction was stirred for 3.5
h after which it was partitioned between water (10 mL), 1 M HCl (10 mL) and EtOAc (30 mL). The
aqueous phase was extracted with EtOAc (3 × 30 mL) and the combined organic phases were dried
with MgSO4. The remaining EtOAc was removed using a rotary evaporator to obtain compound 9 as
1
a white solid in 97% yield (0.425 mmol, 122 mg). H NMR (400 MHz, DMSO) δ ppm 12.97 (br. s., 1 H),
8.71 (s, 1 H), 7.82 (d, J = 8.72 Hz, 1 H), 7.01 (m, 2 H), 4.14 (t, J = 6.22 Hz, 2 H), 2.79 (t, J = 2.55 Hz, 1 H),
13
2.24 (td, J = 6.69, 2.20 Hz, 2 H), 1.83 (tt, J = 8.50, 7.70 Hz, 2 H), 1.61 (tt, J = 7.30 Hz, 2 H). C NMR (100
MHz, DMSO) δ ppm 164.1, 163.9, 157.2, 156.8, 148.9, 131.5, 113.7, 113.5, 111.5, 100.6, 84.1, 71.4, 68.0,
+
27.4, 24.4, 17.3. HRMS (ESI): calcd for C16H14O5Na ([MNa] ): 309.0739, found: 309.0727. mp: 170.0–
170.9 °C.
Compound 6: Compound 9 (0.349 mmol, 100 mg) was dissolved in SOCl2 (6 mL). The solution
was brought to reflux and stirred for 6 h before removing excess SOCl2 using the rotary evaporator.
The residue was dissolved in pyridine (5 mL) to which compound 5 (0.349 mmol, 149 mg, 1.0 eq)
dissolved in pyridine (5 mL) was added and the mixture was left to stir overnight. The solution was
then taken up in EtOAc (20 mL) and then washed with 1 M HCl (3 × 20 mL), 5% CuSO4 (20 mL) and
brine (20 mL). The organic layer was dried with MgSO4 and the solvent was removed on a rotary
evaporator to obtain the crude residue, which was purified by flash chromatography using a 5:4:1
hexanes/DCM/EtOAc mixture. Compound 6 was obtained as a pale‐yellow solid in 12% yield (0.0431
1
mmol, 25 mg). H NMR (400 MHz, CDCl3) δ ppm 10.96 (s, 1 H), 8.92 (s, 1 H), 7.85 (d, J = 1.86 Hz, 2 H),
7.62 (d, J = 8.72 Hz, 1 H), 7.28 (t, J = 1.86 Hz, 1 H), 6.96 (dd, J = 8.72, 2.35 Hz, 1 H), 6.89 (d, J = 2.25 Hz,
1 H), 6.50 (q, J = 1.67 Hz, 2 H), 4.11 (t, J = 6.27 Hz, 2 H), 2.31 (td, J = 6.96, 2.65 Hz, 2 H), 2.18 (d, J = 1.86
13
Hz, 6 H), 1.99 (m, 3 H), 1.75 (m, 2 H). C NMR (100 MHz, CDCl3) δ ppm 170.1, 168.9, 164.7, 162.1,
160.1, 156.9, 149.3, 145.9, 138.9, 132.7, 131.2, 127.6, 118.4, 116.4, 114.7, 114.3, 112.4, 100.8, 83.7, 69.0, 68.4,
+
27.9, 24.8, 18.1, 11.2. HRMS (ESI): calcd for C32H25N3O8Na ([MNa] ): 602.1539, found: 602.1550. mp:
decomp. at 192.9–194.2 °C.
Compound 12: To a solution of 3‐aminophenol (1 g, 9.2 mmol, 2.0 eq) in THF (20 mL) was added
4‐ethynylbenzaldehyde. The solution was stirred for 1.5 h at r.t. prior to adding STAB (1.36 g, 6.44
mmol, 1.4 eq). After stirring for 3 h at r.t., the reaction was quenched with sat. NaHCO3 (60 mL) and
extracted with ether (3 × 20 mL). Compound 12 was purified using flash chromatography with a 2:8
1
EtOAc/hexanes eluent and obtained as a transparent viscous oil in 75% yield. H NMR (400 MHz,
CDCl3) δ ppm 7.51–7.39 (m, 2H), 7.31 (d, J = 8.4 Hz, 2H), 7.01 (t, J = 8.0 Hz, 1H), 6.20 (dddd, J = 10.4,
13
8.0, 2.3, 0.8 Hz, 2H), 6.08 (t, J = 2.3 Hz, 1H), 4.32 (s, 2H), 3.06 (s, 1H). C NMR (100 MHz, CDCl3) δ
156.8, 149.6, 140.4, 132.6, 130.4, 127.4, 121.1, 106.2, 104.9, 99.8, 83.6, 48.1. HRMS (EI): calcd for
C15H13NO: 223.09971, found: 223.09831.
Compound 13: Compound 12 (460 mg, 2.06 mmol) was dissolved in MeOH and cooled to 0 °C.
Once cooled, acetaldehyde (4.0 eq, 0.461 mL, 8.24 mmol) was added and the solution was stirred for
2 h. After, the solution was removed from ice and then NaBH4 (6.0 eq., 468 mg, 12.4 mmol) was added
and the mixture stirred at room temperature overnight. Purification by flash chromatography by
gradient elution (0%–30% EtOAc in hexane) yielded compound 13 in 50% yield as a transparent
1
viscous oil. H NMR (400 MHz, CDCl3) δ ppm 7.43 (d, J = 8.23 Hz, 2 H), 7.18 (d, J = 7.84 Hz, 2 H), 7.02
(t, J = 8.28 Hz, 1 H), 6.25 (m, 1 H), 6.15 (m, 2 H), 4.62 (s, 1 H), 4.48 (s, 2 H), 3.43 (q, J = 7.09 Hz, 2 H),
13
3.04 (s, 1 H), 1.19 (t, J = 7.05 Hz, 3 H). C NMR (100 MHz, CDCl3) δppm 156.7, 149.9, 140.1, 132.4,
+
130.2, 126.5, 120.5, 105.1, 103.3, 99.2, 83.6, 53.8, 45.4, 12.2. HRMS (ESI): calcd for C17H18NO ([MH] ):
252.1388, found: 252.1362.

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Compound 11: The synthesis of compound 4 was adapted from a procedure developed by
Takakura, et al. [29]. POCl3 (0.464 mL, 4.77 mmol, 4.0 eq) was added dropwise to DMF (1.06 mL, 13.7
mmol, 1.5 eq.) over ice and stirred for 5 min. Compound 13 was then added and the solution was
stirred at r.t. for 1 h. Afterwards, the solution was heated to 70 °C and stirred for 1 h before cooling
the solution to r.t. Hydrolysis was performed by adding H2O (10 mL) to the reaction and stirring for
30 min. The solution was then neutralized with NaOH and extracted with EtOAc (3 × 50 mL).
Compound 11 was purified by flash chromatography using a 2:8 EtOAc/hexanes elution system and
1
was obtained as a white solid in 44% yield. H NMR (400 MHz, CDCl3) δ ppm 11.57 (s, 1H), 9.57–9.18
(m, 1H), 7.45 (d, J = 8.3 Hz, 2H), 7.26 (t, J = 4.4 Hz, 2H), 7.13 (d, J = 8.5 Hz, 2H), 6.26 (dd, J = 8.9, 2.4 Hz,
1H), 6.12 (d, J = 2.4 Hz, 1H), 4.59 (s, 2H), 3.52 (q, J = 7.1 Hz, 2H), 3.06 (s, 1H), 1.26 (t, J = 7.1 Hz, 3H).
13
C NMR (100 MHz, CDCl3) δ 192.4, 164.3, 154.9, 138.0, 135.5, 132.6, 126.2, 121.2, 112.1, 104.8, 97.5,
83.3, 53.4, 45.8, 12.2. HRMS (EI): calcd for C18H17NO2: 279.12593, found: 279.12498. mp: 97.1–97.8 °C.
Compound 14: Compound 11 (184 mg, 0.658 mmol) and Meldrum’s acid (1.0 eq, 95 mg, 0.658
mmol) was dissolved in 99% EtOH (5 mL) to which piperidine (0.02 eq, 1.3 μL 0.0130 mmol) and
AcOH (0.02 eq, 0.75 μL, 0.0130 mmol) was added. The solution was stirred at r.t. for 20 min before it
was heated to 78 °C and stirred for 3 h. After, the mixture was cooled to r.t. and then to 0 °C and
allowed to crystallize overnight. The product was filtered and collected as a yellow‐green solid in
1
48% yield. H NMR (400 MHz, CDCl3) δ ppm 12.30 (s, 1 H), 8.67 (s, 1 H), 7.46 (t, J = 8.52 Hz, 3 H), 7.13
(d, J = 8.33 Hz, 2 H), 6.72 (dd, J = 9.01, 2.45 Hz, 1 H), 6.57 (d, J = 2.25 Hz, 1 H), 4.67 (s, 2 H), 3.62 (q, J =
13
7.15 Hz, 2 H), 3.08 (s, 1 H), 1.32 (t, J = 7.10 Hz, 3 H). C NMR (100 MHz, CDCl3) δ ppm 165.3, 164.1,
157.8, 154.3, 150.4, 136.8, 132.8, 132.0, 126.1, 121.7, 111.3, 109.2, 106.8, 97.7, 83.0, 53.9, 46.4, 12.1. HRMS
+
(ESI): calcd for C21H17NO4Na ([MNa] ): 370.1055, found: 370.1065. mp: 207.3–208.8 °C.
Compound 10 and 15: POCl3 (3.0 eq, 40.4 μL, 432 μmol) was added to a solution of compound
5 (50 mg, 144 μmol) dissolved in DCE (1.4 mL) after which the solution was heated to 84 °C and
stirred for 4 h to form the acyl chloride. The solvent was removed by rotary evaporation and the
residue was dissolved in acetonitrile (1.4 mL). A separate solution containing 5 or 4 (to make 10 or 15
respectively) (1.2 eq., 49 mg, 173 μmol) and DIPEA (2.0 eq, 50 μL, 288 μmol) in acetonitrile (1.4 mL)
was then added to the solution of the acyl chloride and the combined reaction mixture was stirred at
room temperature overnight. The solvent was removed by rotary evaporation and the crude was
purified by flash chromatography using a gradient elution system of 0%–50% EtOAc in hexanes
elution system to afford the product.
1
Compound 10: yellow solid in 28% yield. H NMR (600 MHz, CDCl3) δ ppm 11.00 (s, 1H), 8.77
(s, 1H), 7.83 (d, J = 1.9 Hz, 2H), 7.51–7.41 (m, 3H), 7.23 (t, J = 1.8 Hz, 1H), 7.14 (d, J = 8.3 Hz, 2H), 6.67
(dd, J = 8.9, 2.5 Hz, 1H), 6.56 (d, J = 2.3 Hz, 1H), 6.48 (q, J = 1.7 Hz, 2H), 4.64 (s, 2H), 3.59 (q, J = 7.1 Hz,
13
2H), 3.07 (s, 1H), 2.17 (d, J = 1.8 Hz, 6H), 1.30 (t, J = 7.1 Hz, 3H). C NMR (150 MHz, CDCl3) δ 170.2,
169.1, 162.9, 161.2, 157.7, 153.6, 149.0, 146.0, 139.4, 137.5, 132.9, 132.7, 131.6, 127.7, 126.3, 121.6, 118.3,
116.5, 110.8, 109.4, 97.7, 83.3, 77.7, 54.0, 46.4, 12.3, 11.3. HRMS (ESI): C37H28N4O7 calcd for: 663.1856,
+
found: 663.1855 ([MNa] ). mp: decomp. at 330.0–331.2 °C.
1
Compound 15: yellow solid in 37% yield. H NMR (600 MHz, DMSO) δ ppm 10.93 (s, 1H), 8.74
(s, 1H), 7.79 (d, J = 1.8 Hz, 2H), 7.74 (d, J = 9.1 Hz, 1H), 7.46 (d, J = 8.3 Hz, 2H), 7.23 (d, J = 8.4 Hz, 2H),
7.21 (s, 4H), 7.11 (t, J = 1.8 Hz, 1H), 6.86 (d, J = 11.2 Hz, 1H), 6.70 (s, 1H), 4.78 (s, 2H), 4.16 (s, 1H), 3.64
13
(q, J = 7.0 Hz, 2H), 1.19 (t, J = 7.1 Hz, 3H). C NMR (150 MHz, DMSO) δ 169.7, 162.0, 161.0, 157.1,
153.4, 148.3, 138.8, 134.8, 132.4, 132.0, 131.9, 127.7, 126.8, 126.0, 120.4, 120.2, 117.1, 110.9, 109.8, 108.4,
+
96.8, 83.3, 80.8, 52.8, 45.7, 29.8, 12.2. HRMS (ESI): calcd for C35H24N4O7Na ([MNa] ): 635.1543, found:
635.1537. mp: decomp. at 184.8–184.9 °C.
3. Results and Discussion
3.1. Initial Design
We initially envisioned attaching a dimaleimide quencher moiety at the C3 position of a
coumarin fluorophore through an amide bond, while appending a hexyne tail at the C7 position
through an ether linkage. Thus, we first synthesized 1 (Scheme 2) to confirm that the coumarin FlARe

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probe without the alkyne tail could be quenched by the dimaleimide quencher. Fluorogen 1 also
allowed us to subsequently determine the photophysical effects of adding the terminal alkyne tail.
Scheme 2. Synthetic route for the preparation of fluorogen 1.
First the coumarin core was formed via a Knoevenagel condensation between 4‐formylresorcinol
and diethylmalonate, providing ester 2 in good yield (Scheme 2). Carboxylic acid 3 was then obtained
in excellent yield after hydrolysis of 2. We originally attempted to couple acid 3 with aniline 4;
however, the instability of the unsubstituted maleimide groups of aniline 4 made this synthetic
strategy difficult and impractical. Therefore, we switched to aniline 5, whose methyl‐substituted
maleimide groups are more stable. Aniline 5 was coupled to carboxylic acid 3 using HBTU to obtain
the FlARe probe 1 [30]. The low yield of the final coupling reaction is probably due to the low
nucleophilicity of substituted aniline 5.
Probe 1 was then reacted with an equimolar concentration of a representative target protein–
namely, maltose binding protein genetically fused to the dC10* target peptide (MBP‐dC10*), in order
to validate the activation of the probe by FlARe (Figure 1) [22]. The reaction of 1 with MBP‐dC10*
was observed to be fluorogenic with a fluorescence enhancement of 12.7. This is considerably lower
than the FE manifested by a previous FlARe probe comprised of a 7‐(diethyl)aminocoumarin instead
of the 7‐hydroxylcoumarin [19]. Notably, the overall fluorescence intensities of both the activated and
quenched form of 1 are over 10‐fold higher than those of the previously reported YC5 [19]. Regarding
reaction kinetics, the majority of 10 μM 1 reacts with 10 μM dC10* tag within 40 minutes, while the
reaction was essentially complete after 2 hours (Figure 1B). From the time course of this reaction, the
−1 −1
second order rate constant was calculated as k2 = 245 ± 8 M s . This rate constant is similar to that
measured for our previous 7‐(diethyl)aminocoumarin FlARe probe,[19] suggesting that the change
of substituent at the C7 position on the coumarin did not influence reactivity. Encouraged by these
results, we proceeded to append the terminal hexyne tail through an ether linkage at the C7 hydroxyl
group, in the synthesis of compound 6 (Scheme 3).

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Figure 1. (A) Excitation (dotted line) and emission (solid line) spectra of 1 before (grey line) and after
reacting with MBP‐dC10* (black line). (B) Kinetic time course for the fluorogenic reaction between 10
μM of 1 with 10 μM of MBP‐dC10* (black), relative to the stability of 1 in the absence of MBP‐dC10*
(grey). Rate constant k
= 245 ± 8 M−1s−1.
2
O
O
6, Et3N,
NaI, 56 oC,o.n.
100 oC, o.n.
O
Cl
I
Acetone
94%
DMF
44%
O
O
7
8
1) SOCl2, 75 oC, 6 h
O
O
O
O
N
N
MeCN
97%
5
2) , Pyridine, r.t., o.n.
OH
O
O
2 M NaOH, r.t.,
3.5 h
12%
O
O
N
H
O
O
O
O
9
6
Scheme 3. Synthetic route for the preparation of alkyne 6.
In this approach, we attached the hexyne group before attaching the dimaleimide quenching
intermediate 5 (Scheme 3). Thus, 7 was first synthesized from 6‐chlorohexyne before using 2 to
displace the iodide in 7, obtaining 8 in decent yield. As performed in the synthesis of 1, ester 8 was
then hydrolyzed to provide 9 in excellent yield, which was then converted to the acid chloride using
SOCl . The acid chloride was then reacted with aniline 5 in pyridine to attach the quenching
2
dimaleimide moiety and form 6. This coupling of aniline 5 also gave a low yield, although slightly
better than the HBTU‐mediated reaction (Scheme 2).
As before, this novel FlARe linker was incubated with an equimolar concentration of MBP‐dC10*
(10 μM). However, no fluorescence increase was observed, even after 24 h of incubation (data not
shown). Since 6 only differs from 1 by the presence of the terminal hexyne tail, this functional group
was suspected of disrupting the coumarin’s intrinsic fluorescence. Density Functional Theory (DFT)
calculations of 6 showed that neither the LUMO nor the HOMO of the alkyne group were of
appropriate energy to quench coumarin fluorescence through PeT (Figure S1). Thus, it is unlikely
that the coumarin fluorescence of 6 was being quenched by this specific mechanism, as opposed to
what has been proposed to be the case for other alkyne‐functionalized coumarins [31].
Rather, different studies have shown that the coumarin fluorophore’s emission is very sensitive
to the functional group’s rigidity at the C position [32]. Indeed, this sensitivity is exploited in enzyme
7

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activity probes in which a carbohydrate linked to umbelliferone at the C
7
position is enzymatically
cleaved to release a highly fluorescent 7‐hydroxylcoumarin fluorophore [33,34]. Therefore, in the case
of 6, the terminal hexyne tail at the C position offers a means of rotational relaxation that could
7
severely diminish the probe’s fluorescence. It is striking to note that the 7‐propargyl coumarin probe
developed by Liu, bearing a slightly shorter alkyl chain is indeed fluorescent [15]. Thus, extending
the alkyl chain beyond three carbons on 7‐alkoxycoumarins apparently has a detrimental effect on
fluorescence.
Despite the lack of fluorogenic activation by FlARe, we tested whether 6 could successfully
introduce an alkyne group site‐specifically to a target protein, albeit in a non‐fluorogenic manner.
Specifically, we tested the utility of 6 to couple RhB‐N
reaction. We first reacted MBP‐dC10* with 6 for 24 h to ensure complete labelling. Afterwards, we
performed the CuAAC with RhB‐N and evaluated the bioconjugation by fluorescent SDS‐PAGE
(Figure 2). On the gel, we observed a protein band that is red fluorescent but not blue fluorescent
(Figure 2B, lane 3). This was indicative of formation of the MBP‐dC10*‐RhB‐N conjugate using
3
to MBP‐dC10* through a CuAAC conjugating
3
3
compound 6 as the linker. The lack of any blue fluorescence is consistent with the non‐fluorogenic
nature of 6 that was previously observed (Figure 2C). The protein bands in lane 5 of Figure 2 did not
fluoresce at all, confirming that without first introducing the alkyne handle using 6, RhB‐N
3
could
not be conjugated to MBP‐dC10*. In conclusion, although compound 6 was not observed to be
fluorogenic as intended, it still introduced an alkyne group into MBP‐dC10*, to which RhB‐N could
3
then be conjugated through cycloaddition.
Figure 2. SDS PAGE gel of protein conjugates obtained after first reacting 6 with MBP‐dC10* by the
FlARe reaction followed by the conjugation of rhodamine B azide (RhB‐N ) to MBP‐dC10* through 6
3
using CuAAC. A) Coomassie stained gel showing MBP‐dC10* bands. B) Unstained gel excited with
Green Epi Illumination and imaged using a 605/50 filter. C) Unstained gel excited with Blue Epi
Illumination and imaged with a 530/28 filter. Lanes: 1) Pre‐stained protein standards 2) MBP‐dC10*
3) MBP‐dC10* reacted with 6 and then conjugated to RhB‐N 4) MBP‐dC10* reacted with 6 5) MBP‐
3
dC10* incubated with RhB‐N under CuAAC conditions.
3
3.2. Improved Design
As previously discussed, the non‐fluorogenic nature of 6 made it difficult to ascertain how
rapidly and efficiently the alkyne group was introduced. To address this issue, we designed a second
fluorogenic bioconjugating linker where the alkyne group was attached to the coumarin core at the
C position as a 4‐ethynylbenzylamine substituent. We hypothesized that this functional group
7
would present less rotational flexibility and maintain the coumarin’s intrinsic fluorescence. These
modifications are comprised in the design of the fluorogenic linker 10 shown in Scheme 4.

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Scheme 4. Synthetic route for the preparation of fluorogenic alkynes 10 and 15.
Prior to the formation of 10, the salicylaldehyde 11 was prepared (Scheme 4). First, the terminal
alkyne was attached by reacting 3‐aminophenol with p‐ethynylbenzaldehyde in a reductive
amination using sodium triacetoxyborohydride (STAB). This afforded the secondary amine 12 in
good yield, which was then capped with acetaldehyde through a second reductive amination using
STAB to yield the tertiary amine 13 in moderate yield. A Vilsmeier–Haack reaction was performed
to obtain the functionalized salicylaldehyde 11 in moderate yield, which was then followed by a
Knoevenagel condensation with Meldrum’s acid to obtain coumarin 14. Finally, the
di(methylmaleimide) aniline intermediate 5 was attached to 14 by first converting the latter to its acid
chloride with POCl3 and then forming the amide bond to afford 10 in good yield.
Reacting equimolar amounts of 10 with MBP‐dC10* at 10 μM resulted in a 37‐fold enhancement
in fluorescence after complete reaction with the tag (Figures 3A and S2A). Not only did this indicate
successful fluorescence activation by FlARe, it is also a significant improvement from the FE observed
with probe 1. However, nearly 10 h were required for the complete reaction of 10 with MBP‐dC10* at
−1 −1
10 μM, providing a rate constant of k2 = 8.4 ± 0.7 M s , which is over an order of magnitude lower
than the rate constant measured for 1 (Figure S2B). This suggests that the modified alkyne tail of 6
may hinder the labelling reaction. On a practical level, this means that making micromolar
concentrations of a protein conjugate would require at least two days to complete, calling for a
quicker alternative. As mentioned before, unsubstituted maleimides react more quickly than
methylmaleimides [30]. Thus, the di(methylmaleimide) aniline moiety was replaced with
dimaleimide aniline in the design of probe 15 (Scheme 4).

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Figure 3. (A) Excitation (dotted line) and emission (solid line) spectra of 10 μM 10 (grey lines) and 15
(blue lines) before (light) and after (dark) reacting with 10 μM MBP‐dC10*. (B) Kinetic time course for
the fluorogenic reaction between 10 μM of 10 (grey lines) and 15 (blue lines) with 10 μM of MBP‐
dC10* (dark lines) (black), relative to the stability of 10 and 15 in the absence of MBP‐dC10* (light
lines).
Using the same synthetic strategy outlined in Scheme 4, aniline 4 was coupled to 14 through the
acyl chloride afforded by POCl to obtain 15 in decent yield. Upon reacting with equimolar amounts
3
of MBP‐dC10* at 10 μM, the fluorescence intensity of 15 was found to increase 10‐fold. The final MBP‐
dC10*‐15 adduct displayed a quantum yield of Φ = 0.093, which is 57 times higher than the quantum
yield of the unreacted form of 15 (Figure S3). This FE was three‐fold lower compared to 10 but still
comparable to the FE observed with 1 (Figure 3A). The rate constant of the reaction between the target
−1 −1
peptide and 15 was calculated to be k = 180 ± 10 M s , with completion at the 10‐μM concentrations
2
achieved after only 30 minutes (Figure 3B). Thus, switching to the unsubstituted maleimide
accelerated the labelling reaction by over an order of magnitude, making it comparable to the reaction
between 1 and MBP‐dC10*.
Furthermore, the total time required to prepare a protein conjugate was reduced to a one‐day
process. Encouraged by both the fluorogenic nature of 15 and its rapid labelling reaction, the utility
of 15 for bioconjugation was assessed in a manner similar to 6. Thus, 10 μM of 15 was incubated with
10 μM MBP‐dC10* for 1 h, after which RhB‐N was then conjugated to MBP‐dC10* by CuAAC
3
through the alkyne handle introduced by 15. SDS‐PAGE analysis showed a protein band that was
both red and blue fluorescent (Figure 4B and 4C, lane 4), indicating that MBP‐dC10* was conjugated
to the red fluorescent rhodamine B through the blue fluorescent 15 coumarin linker. Interestingly, the
blue fluorescence of this protein band is considerably dimmer than MBP‐dC10* labelled only with 15
(Figure 4C, lane 2). We suspect this is because of a FRET interaction between the coumarin
fluorophore of 15 and the rhodamine B group of the model ‘cargo’. This FRET interaction has been
identified previously in the literature, and a comparison of the coumarin emission spectrum of the
adduct of 15 and the excitation spectrum of rhodamine B (see Figure S4) reveals the significant
spectral overlap required for FRET [20,35,36]. Furthermore, molecular modelling of the triazole
adduct formed after CuAAC coupling suggests the fluorophores would be separated by less than 30
Å, allowing for efficient resonance energy transfer. The protein band in lane 5 is neither red nor blue
fluorescent, confirming that the linker 15 is required to successfully conjugate RhB‐N to MBP‐dC10*
3
(Figure 4B, lane 5). Therefore, 15 successfully introduced an exposed alkyne group in a site‐specific
and fluorogenic manner, which allowed for the attachment of the azide functionalized cargo, RhB‐
N3.

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Figure 4. SDS PAGE gel of protein conjugates obtained after first reacting 10 μM 15 with 10 μM MBP‐
dC10*, followed by the conjugation of rhodamine B azide (RhB‐N3) to MBP‐dC10* through 15 using
CuAAC. A) Coomassie stained gel showing MBP‐dC10* bands. B) Unstained gel imaged using a
605/50 filter. C) Unstained gel excited with Blue Epi Illumination and imaged with a 530/28 filter.
Lanes: 1) MBP‐dC10* 2) MBP‐dC10* reacted with 15 3) pre‐stained protein molecular weight markers
4) MBP‐dC10* reacted with 15 and then conjugated to RhB‐N3 5) MBP‐dC10* incubated with RhB‐N3
under CuAAC conditions.
In addition to its rapid kinetics, 15 was also noted to be resistant to hydrolysis over the course
of 3 h under the assay conditions. This prompted us to perform the CuAAC between 15 and RhB‐N3
prior to conjugation of the triazole product to MBP‐dC10*, as we hypothesized that the maleimides
would remain intact after the CuAAC. Thus, after performing the CuAAC between 15 and RhB‐N3,
we added MBP‐dC10* in equimolar concentrations to 15 and allowed the 15‐RhB triazole product to
react with the dC10* tag for 1 h. As before, the bioconjugation was analysed by SDS‐PAGE (Figure
5). The presence of a red and blue fluorescent protein band (Figure 5B and C, lane 2) indicates the
successful formation of the MBP‐dC10*‐rhodamine conjugate. Like the previous experiment, in the
lane representing the conjugation of the RhB‐N3 cargo to MBP‐dC10* after labelling the protein with
15, the blue fluorescence of this band is dimmer than that of MBP‐dC10* labelled only with 15, and
no red fluorescence is observed from the band in lane 4. This reflects both the FRET interaction
occurring between the fluorophores and the importance of 15 for successful bioconjugation.
However, it is also noteworthy that the intensity of the fluorescence of all the protein bands
shown in Figure 5 appears to be lower than those from the previous experiment (shown in Figure 4).
The similarity of the density of the Coomassie stained protein bands between the gels (Figure 4A vs.
Figure 5A) suggests this is not due to a difference in gel loading. We hypothesize that the labelling
reaction of the 15‐RhB triazole adduct with MBP‐dC10* (shown in Figure 5) was less efficient than
the reaction of 15 with MBP‐dC10* (shown in Figure 4), leading to a lower proportion of protein being
labelled after 1 hour of incubation, and a less intense signal being observed. It is conceivable that the
steric bulk of the adduct pre‐formed between rhodamine B and fluorogen 15 decreases the efficiency
of the subsequent reaction with MBP‐dC10*, slowing the labelling reaction in a manner similar to
what was observed on comparing 1 and 10. Therefore, although we have demonstrated that it is
possible to perform the FlARe protein‐labelling reaction either before or after the CuAAC reaction, it
appears to be more efficient to label the protein first and then attach the cargo to the labelled protein
through the alkyne group.

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Figure 5. SDS PAGE analysis of protein conjugates obtained after first reacting 15 with rhodamine B
azide (RhB‐N3) in the CuAAC reaction, followed by conjugation to MBP‐dC10* by the FlARe reaction.
A) Coomassie stained gel showing MBP‐dC10* bands. B) Unstained gel excited with Green Epi
Illumination and imaged using a 605/50 filter. C) Unstained gel excited with Blue Epi Illumination
and imaged with a 530/28 filter. Lanes: 1) Pre‐stained protein standards 2) MBP‐dC10* reacted with
the 15‐RhB‐N3 product of the CuAAC. 3) MBP‐dC10* reacted with 15 subjected to a mock CuAAC
without RhB‐N3. 4) MBP‐dC10* incubated with RhB‐N3 under CuAAC conditions. 5) MBP‐dC10*.
4. Conclusions
In summary, we have developed a new bioconjugation method that first introduces an alkyne
group onto a target protein in a site‐specific, rapid and fluorogenic manner. This allows for the
subsequent site‐specific attachment of a broad array of azide‐functionalized cargo. Our initially
designed bifunctionalized conjugating agent (6) proved to be non‐fluorescent, presumably due to its
long, flexible hexyne tail. Despite being non‐fluorogenic, 6 was still capable of introducing an alkyne
group to the test protein MBP‐dC10*, onto which the model cargo RhB‐N3 could then be attached. In
our second design, the long alkyne tail of 6 was replaced with a shorter and more rigid alkynyl amine
substituent, while the di(methylmaleimide) aniline was replaced with dimaleimide aniline, resulting
in the heterobifunctional fluorogenic linker 15. These modifications resulted in a linker whose
fluorescence increased 10‐fold upon reacting with the dC10* target peptide (Φ = 0.093), allowing the
introduction of the exposed alkyne group to be monitored in situ by fluorescence increase.
Subsequently, model cargo RhB‐N3 could be attached to MBP‐dC10* through the site‐specifically
installed alkyne handle presented by 15. Furthermore, we demonstrated that azide‐functionalized
cargo can also be incorporated into 15 by CuAAC prior to reacting with the target protein. Overall,
our method allows the site‐specific introduction of an exposed alkyne group into a protein of interest
genetically fused to a dC10* tag, representing an approach that is easily amenable for the synthesis
of new, therapeutically important protein‐cargo conjugates. Additionally, the fluorogenic nature of
the bioconjugating linker allows the initial protein modification step to be easily monitored in situ by
fluorescence. Therefore, our linker provides a facile, modular and adaptable method for the in vitro
synthesis of new protein‐cargo conjugates in the future.
Supplementary Materials: The following are available online at www.mdpi.com/xxx/s1, Figure S1, Figure S2,
Figure S3, Figure S4, Figure S5, synthetic protocols for RhB‐N3, compound 4 and its intermediates, 1H‐NMR and
13C‐NMR spectra.
Author Contributions: Conceptualization, K.K.T. and J.W.K.; methodology, K.K.T., A.C.L. and K.É.R.;
validation, K.K.T., and J.W.K.; formal analysis, K.K.T.; investigation, K.K.T, A.C.L. and K.É.R.; resources, J.W.K.;
data curation, K.K.T.; writing—original draft preparation, K.K.T.; writing—review and editing, J.W.K.;

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visualization, J.W.K.; supervision, K.K.T and J.W.K.; project administration, K.K.T and J.W.K.; funding
acquisition, J.W.K. All authors have read and agreed to the published version of the manuscript.
Funding: This research was funded by the Natural Sciences and Engineering Research Council of Canada,
Discovery Grant number 05893. The APC was funded by the same grant.
Acknowledgments: The authors would like to thank our departmental colleague Dr. John Pezacki and his
research group for their assistance with the CuAAC bioconjugation.
Conflicts of Interest: The authors declare no conflict of interest. Furthermore, the funders had no role in the
design of the study; in the collection, analyses, or interpretation of data; in the writing of the manuscript, or in
the decision to publish the results.
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