Theoretical and experimental investigations of large stokes shift fluorophores based on a quinoline scaffold

Article abstract of DOI:10.3390/molecules25112488

A series of novel styrylquinolines with the benzylidene imine moiety were synthesized and spectroscopically characterized for their applicability in cellular staining. The spectroscopic study revealed absorption in the ultraviolet–visible region (360–380

Full text of DOI:10.3390/molecules25112488

Article
Theoretical and Experimental Investigations of Large
Stokes Shift Fluorophores Based on a
Quinoline Scaffold
Barbara Czaplińska 1, Katarzyna Malarz 2,3, Anna Mrozek-Wilczkiewicz 2,3, Aneta Slodek 4,
Mateusz Korzec 4 and Robert Musiol 1,*
1
Institute of Chemistry, Faculty of Science and Technology, University of Silesia in Katowice,
75 Pułku Piechoty 1A, 41-500 Chorzów, Poland; czaplinska.basia@gmail.com
2
A. Chełkowski Institute of Physics, Faculty of Science and Technology, University of Silesia in Katowice,
75 Pułku Piechoty 1, 41-500 Chorzów, Poland; katarzyna.malarz@us.edu.pl (K.M.);
anna.mrozek-wilczkiewicz@us.edu.pl (A.M.-W.)
3
Silesian Center for Education and Interdisciplinary Research, University of Silesia in Katowice,
75 Pułku Piechoty 1A, 41-500 Chorzów, Poland
4
Institute of Chemistry, Faculty of Science and Technology, University of Silesia in Katowice, Szkolna 9,
40-007 Katowice, Poland; aneta.slodek@us.edu.pl (A.S.); mateusz.korzec@us.edu.pl (M.K.)
* Correspondence: robert.musiol@us.edu.pl
Academic Editor: Pascale Moreau
Received: 7 May 2020; Accepted: 21 May 2020; Published: 27 May 2020
Abstract: A series of novel styrylquinolines with the benzylidene imine moiety were synthesized
and spectroscopically characterized for their applicability in cellular staining. The spectroscopic
study revealed absorption in the ultraviolet–visible region (360–380 nm) and emission that covered
the blue-green range of the light (above 500 nm). The fluorescence quantum yields were also
determined, which amounted to 0.079 in the best-case scenario. The structural features that are
behind these values are also discussed. An analysis of the spectroscopic properties and the
theoretical calculations indicated the charge-transfer character of an emission, which was
additionally evaluated using the Lippert–Mataga equation. Changes in geometry in the ground and
excited states, which had a significant influence on the emission process, are also discussed.
Additionally, the capability of the newly synthesized compounds for cellular staining was also
investigated. These small molecules could effectively penetrate through the cellular membrane.
Analyses of the images that were obtained with several of the tested styrylquinolines indicated their
accumulation in organelles such as the mitochondria and the endoplasmic reticulum.
Keywords: green fluorophores; quinoline; large Stokes shift; intramolecular charge transfer; cell
imaging; DFT calculations
1. Introduction
Quinoline is an attractive scaffold for designing new fluorescent agents due to its small
molecular size; the presence of nitrogen, which creates good coordination properties; its ability to
form hydrogen bonds, and a fair synthetic availability [1]. At the same time, quinoline itself is also
known as a fluorophore unit, which further increases its potential. It is also widely used as a core
molecular fragment in a broad range of drugs and biologically active substances [2–4]. For this reason,
it has been claimed to be a privileged structure [5]. Our research group is interested in synthesizing
and investigating quinoline derivatives [6–8]. In recent years, we obtained a series of new
styrylquinolines with anticancer and antifungal properties [9,10]. Besides active compounds, some of
Molecules 2020, 25, 2488; doi:10.3390/molecules25112488
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the newly synthesized structures also turned out to be good fluorophores [11], which is in agreement
with many reports concerning the application of quinoline’s structure in biological imaging. Due to
their planar and rigid aromatic structure, quinolines interact with proteins and nucleic acids.
Therefore, they were used for DNA and RNA imaging with good results (Figure 1A) [12–15]. Both
activities (therapeutic and diagnostic) of a styrylquinoline derivative were also exploited by Staderini
et al. to stain the Aβ amyloid plaques in Alzheimer’s disease (Figure 1B) [16]. Highly fluorescent
quinolines can also detect both gram-positive and gram-negative bacteria (Figure 1C) [17]. Quinoline
derivatives, especially 8-hydroxy substituted are particularly good chelators for metal ions, and
therefore, this type of probes have dominated this field of chemosensing [18]. Among them are the
Zn2+ sensors, which have very good properties (Figure 1D–I) [19–22]. Indeed, the first and most
commonly known fluorescent zinc ion indicator was 8-hydroxyquinoline, which was used in 1968,
for detection in human plasma [23].
Figure 1. Examples of quinoline-based fluorophores and sensors. (A) carbocyanine dyes for DNA
and RNA; (B) styrylquinoline for amyloid plaques; (C) quinoline dyes for bacterial application; (D)
bisquinoline dye for metal detection; (E) and (F) dipicolylamine derivatives of quinoline as
chemosensing dyes; (G) 8-aminoquinoline dye for physiological fluorescent labeling of Zn2+; (H) and
(I) fluorescein based probes for Zn2+ detection.
Another appealing feature of the quinoline-related compounds is their tendency to have an
intramolecular charge transfer characteristic (ICT) [24,25]. A plausible reason for this is the electron-
accepting nature of the moiety because of a well-defined dipole moment. Compounds with ICT
exhibit different properties in the ground and excited states, which is caused by an electronic
redistribution after photoexcitation. This transient alternation leads to large changes in dipole
moments and to the relaxation of the structure. The energy difference between the Frank–Condon
state and the ICT state is the reason for the large Stokes shift, which is preferred in most fluorescence
methods [26].
In biological applications, particularly fluorescent microscopy, the large stokes shift which is
usually over 80 nm are particularly useful for the reduction of chromatic aberrance and increase of

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resolution [27]. Stimulated emission depletion (STED) microscopy is a technique that utilizes res-
shifted light pulse for reducing the excitation of the molecules peripheral to the excitation spot [28].
Consequently, their excitation is reduced, which allows us to overcome the diffraction barrier in high
resolution images. An excellent review on large stokes shift dyes in such STED microscopy was
published recently [29].
Another application of large stokes shift dyes is the Fӧrster resonance energy transfer (FRET)
microscopy, a technique based on pair dyes that can undergo direct excitation of an acceptor by the
adjacent donor molecules [30]. An effective dye pair needs overlapping in the emission and excitation
spectra of donor and acceptor molecule, respectively, which inevitably leads to cross-excitation of the
acceptor upon the donor, resulting in false-positive results. Large stokes shift dyes are helpful to
overcome this problem, increasing the threshold of detection. Moreover, compounds of this type have
attracted much attention because of their possible applications in photoelectronic devices, as non-
linear materials, emitters, or in organic light-emitting diodes (OLED) [31–33]. Since the ICT state
emission is usually strongly dependent on environmental parameters like the solvent polarity or
viscosity, push–pull compounds are also widely used as chemical sensors [34,35]. Based on our
experience with the styrylquinoline compounds, we decided to substitute this scaffold with a Schiff
base moiety (Figure 2).
Figure 2. The synthetic pathway of compounds 3a–f. Reagents and conditions: a) Aromatic
aldehyde/Ac2O, b) SnCl2/EtOH, and c) aromatic aldehyde/EtOH.
Schiff bases are frequently reported for their antifungal, antibacterial, antiproliferative, anti-
inflammatory, antimalarial, and antiviral activity [36–38]. They are also used as metal ions or pH
sensors [39,40]. On the other hand, Schiff bases are relatively rarely exploited for their fluorescent
potential. However, the unique properties of the imine bond, namely its polarized electron density
and propensity to form a tautomeric structure, make it useful for designing compounds with ICT
characteristics. Thus, in the present report, we provide the chemical, physicochemical, and optical
profiles of the newly obtained compounds, together with the theoretical data and biological assays.
2. Results and Discussion
2.1. Spectroscopic Properties
2.1.1. Absorption Properties
The absorption and fluorescence of all newly synthesized compounds were investigated using
UV–VIS spectroscopy. When selecting the solvents for the experiments, various properties were
taken into account, such as polarity and the ability to create hydrogen bonds, since these might

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influence fluorescence. The results that were obtained in this study are presented in Table 1. The
obtained compounds exhibited a good excitation/emission profile. All absorption spectra consisted
of two bands and the maxima of the more red-shifted ones were over 360 nm, which is desirable in
cell imaging. Wavelengths shorter than 350 nm carry enough energy to destroy the complex protein
structure, which interferes with the observation process and, therefore, are unsuitable for long-term
experiments [41]. Absorption wavelengths above 360 nm enable avoidance of any autofluorescence
from naturally occurring molecules whose excitation profile embrace higher-energy wavelengths,
such as 270 nm (tyrosine) [42], 280 nm (tryptophan) [43], or 340 nm (NAD(P)H) [44]. When the
absorption spectra (Figure 3) were analyzed, a relationship between a structure and the band shift
were observed. The biggest red shift was recorded for compound 3a, which had a strong electron-
donating group in its structure (N(CH3)2). Therefore, attaching an electron-donating N(CH3)2
substituent in 3a led to a markedly enhanced extinction coefficient of the low energy band, similar to
the compound 3c bearing the OH group (Table 1). Then, the spectra maxima shifted toward the longer
wavelengths, together with an increase of the donating power of the substituent that was attached to
a phenyl moiety (3a > 3c > 3b > 3f > 3e). Apparently, compound 3d did not fit into a relationship that
could be explained on a structural basis. The phenyl vinyl substituent constituted a larger conjugated
aromatic system that diluted the partitive effects and changed the overall dipole of a molecule.
Table 1. Fluorescent and absorption properties of the Schiff bases 3a–f in ethanol.
Absorption
λmax/nm (ε/103 M−1 cm−1)
388 (51.1)
Emission
λmax/nm
Stokes Shift
nm/cm−1
Compound
φ
3a
3b
3c
3d
3e
3f
519
512
514
512
514
514
0.041
131/6505
144/7642
144/7571
140/7350
152/8169
148/7867
336 (33.3)
368 (44.4)
0.054
0.052
0.034
0.058
0.079
276 (46.8)
370 (52.3)
272 (139.6)
372 (46.1)
274 (131.3)
362 (24.3)
270 (35.6)
366 (33.3)
272 (23.4)
Figure 3. UV–Vis absorption spectra of the Schiff bases 3a–f in ethanol (1.25 × 10−5 M).

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2.1.2. Fluorescence Properties
The fluorescence spectra were recorded in various solvents as well, which enabled the charge
transfer character of the emission to be revealed. All fluorescence spectra are presented in Figure 4
and the corresponding data are listed in Table 2. As one can see, the compounds exhibited the best
fluorescent properties in ethanol. The emission peaks occurred around 514 nm in ethanol for all 3a–f
compounds, and were intense and strongly red-shifted compared to those in the chloroform and
acetonitrile and were comparable to those in DMSO (except for 3a). On the other hand, in DMSO and
acetonitrile, the intensity of the emission drastically decreased in most cases, while in the chloroform,
an average intensity and blue-shifted spectra could be observed.
Table 2. Solvatochromic properties of the compounds 3a–f; S–Stokes shift.
Chloroform
λabs λem
Acetonitrile
λabs λem
DMSO
λabs Λem
Ethanol
Λabs Λem
S
S
S
S
Compound
nm
nm
nm/cm−1
nm
nm
nm/cm−1
nm
nm
nm/cm−1
nm
nm
nm/cm−1
3a
3b
3c
3d
3e
3f
386
370
368
374
362
370
467
450
454
458
456
459
81/4493
80/4804
86/5147
84/4904
94/5694
89/5240
387
370
367
372
364
368
513
500
502
500
501
500
126/6346
130/7027
135/7327
128/6881
137/7512
132/7174
400
374
374
382
370
374
557
511
510
512
516
507
157/7046
137/7168
136/7130
130/6646
146/7647
133/7014
388
368
370
372
362
366
519
512
514
512
514
514
131/6505
144/7642
144/7571
140/7350
152/8169
148/7867
These data suggest that the emission spectra are strongly dependent on the solvent polarity,
which probably arises from a big difference in the dipole moment between the ground and excited
states [45]. At the same time, it is easily noticeable that the changes had a specific tendency, which is
called positive solvatochromism—an increase of the solvent polarity induces the red shift of an
emission band. The intramolecular charge transfer phenomenon is another reason for the very large
Stokes shift that was observed in the synthesized compounds. The changes of the dipole moment that
occurred in the excited state caused the relaxation of the solvent around an excited molecule, to obtain
a state of minimum energy [46].
The smallest Stokes shift of 131 nm (6505 cm−1) was observed for 3a and the largest one was 152
nm (8169 cm−1) for 3e, for the measurements that were performed in ethanol. The Stokes shift is critical
to the sensitivity of the fluorescence method because it enables a good detection of photons against a
low background as well as the separation from an excitation source. Unfortunately, such large Stokes
shifts are usually associated with small fluorescence quantum yields. These were also measured for
all compounds, as presented in Table 1. The smallest quantum yield was observed for 3d and
amounted to 0.034. Whereas, for 3f, the value was 0.079, which was the highest fluorescence efficiency
that was observed. Nevertheless, all of these features together made these compounds suitable for in
vitro biological imaging.

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Figure 4. Fluorescence spectra of compounds 3a–f in the various solvents (1.25 × 10−5 M).
2.1.3. Influence of Water on the Spectroscopic Properties
Additionally, we decided to conduct an in-depth investigation of the spectroscopic properties of
the dyes in a water environment, as is typical in analytical conditions that include the cell interior.
For this purpose, because of the low water solubility of our compounds, we carried out a series of
measurements in ethanol, with an increasing amount of water. The sets of the relationships that were
obtained through these experiments are presented in Table 3.

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Table 3. Spectroscopic properties of 3a–f depending on the amount of water in a solution. S—BioTek
Stokes shift.
Ethanol + Water
(80%/20%)
Ethanol + Water
(50%/50%)
Ethanol + Water
(40%/60%)
λabs λem
Ethanol
λabs λem
Compound
S
Λabs Λem
S
λabs Λem
S
S
nm
nm
nm/cm−1
nm
nm
nm/cm−1
nm
nm
nm/cm−1
nm
nm
nm/cm−1
3a
3b
3c
3d
3e
3f
388
368
366
372
362
366
519
512
514
512
514
514
131/6505
144/7642
146/7867
140/7350
152/8169
148/7867
388
368
366
372
360
366
527
524
524
524
525
522
139/6797
156/8090
158/8238
152/7797
165/8730
156/8165
388
368
366
372
362
366
534
530
530
532
529
530
146/7046
162/8305
164/8454
160/8084
167/8720
164/8454
354
366
366
366
360
362
540
534
534
536
533
534
186/9730
168/8595
168/8595
170/8665
173/9016
172/8897
According to the results, water had almost no influence on the absorption spectra. The
absorbance maxima of each compound did not shift, compared to their maxima in ethanol. An
exception was compound 3a whose absorption maximum was significantly blue-shifted in the
40%/60% mixture, which could be an effect of the higher protonation of the dimethylamine group in
the highly protic solvent [47]. Fluorescence, on the other hand, is strongly dependent on the
environmental factors, because an increased amount of water shifts the emission to the longer
wavelengths. At the same time, the Stokes shifts also had higher values in the aqueous solutions, e.g.,
172 nm vs. 148 nm in the case of compound 3f, which was in accordance with the increase of
fluorescence intensity. In light of these results, we decided to further study this phenomenon.
Moreover, a recently published article that investigated the hydrolysis and aggregation processes of
the Schiff bases was brought to our attention, which raised more doubts about the changes that occur
in water. Due to the possibility that the compounds would degrade, we used the HPLC method for
the samples that contained 25% water and 75% ethanol, and measured the composition of the
mixtures after 5, 15, 30, and 60 min. The results of this study are presented in Figure 5.
Figure 5. Plot presenting the decomposition rate of compounds 3a–f in a water environment.
For some compounds, the decomposition was quite rapid. For compounds 3b and 3f, only 30–
40% of the product remained after one hour. On the other hand, compounds 3d and 3e appeared to
be rather resistant to hydrolysis. Interestingly, these observations correlated with the structural
features that possibly affect the dipole moment of a molecule. The methoxy- (3b) and ethoxy-
substituted (3f) compounds decomposed quickly, while the more lipophilic, less polarized ones were
stable. The styryl- (3d) or 2-fluorophenyl- (3e) groups did not have an increased electron density on
the imine bond, which had a positive effect on stability. It should be noted here that this type of
decomposition does not always disturb the applications in water-based environments, e.g., biological

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systems. Recently, we published a report on the application of a pH-sensitive Schiff base, as a cellular
probe for staining lysosomes [48].
2.1.4. Estimating the Dipole Moment (the Lippert–Mataga Equation)
In order to explain the large Stokes shifts in relation to changes in the structure in the excited
states, we applied the Lippert–Mataga Equation (1) [49,50]. Estimating the difference between the
excited- and ground-state dipole moments in this equation (µ = µe−µg) exploited the relationship
between a Stokes shift and the solvent polarity. This method is widely used in photophysical studies
of the push–pull molecules to elucidate the internal charge transfer (ICT) characteristics [24,25,51,52].
The equation used was as follows:
ꢍ
ꢌ
ꢇꢈꢉ ꢅꢉ
ꢊ
ꢋ
(1)
(2)
∆ꢀ = ∆ꢁꢂꢃꢄꢅꢂꢆ
=
∆ꢑ + ꢒꢓꢔꢕꢖ.
ꢐ
ꢎꢄꢏ
ꢍ
ꢗꢅꢘ
∆ꢑ = ꢇꢗꢙꢘ
ꢚ ꢅꢘ
−
ꢍ
ꢇꢚ ꢙꢘ
where Δν is the Stokes shift in a given solvent (cm−1), h is the Planck’s constant, c is the speed of light
in a vacuum, and a is the Onsager cavity radius. The Onsager cavity radii and the ground-state dipole
moments (μg) were obtained from the quantum chemical calculation (Gaussian09, DFT/B3LYP/6-
311+G(d)) [41,53]. The Δf is the orientation polarizability of the solvent, which measures both the
electron mobility and dipole moment of a solvent molecule. Δf was calculated using Equation (2),
where ε is the dielectric constant of the solvent and
n is the optical refractive index of the
solvent. Figure 6 presents the Lippert–Mataga plots, which provide the slopes to the further
calculations of equation:
2
ℎꢠꢡꢢ
ꢛꢜꢝꢓꢞꢟ =
ꢈ∆ꢣꢂꢤꢌꢇꢥ
(3)
whose values are presented in Table 4, together with the results that were obtained from the equation.
The calculated values of Δμeg were 16.7 D, 18.4 D, 16.3 D, 17.8 D, 16.6 D, and 15.8 D, respectively, for
compounds 3a–f. Such big differences between the dipole moments indicate the internal charge
transfer (ICT) as being a process that is involved in acquiring the excited state.
Table 4. The Onsager cavity radii, slopes, and changes of the dipole moments of compounds 3a–f.
Slope
(cm−1)
Δμeg
μg
μe
Compound Onsager Cavity Radius (Å)
(Debye) (Debye) (Debye)
3a
3b
3c
3d
3e
3f
5.94
6.00
5.48
5.66
5.63
6.27
13,438 ± 5578
16,925 ± 3660
13,903 ± 3722
14,614 ± 2235
14,304 ± 3404
15,202 ± 2888
16.7
18.4
16.3
17.8
16.6
15.8
5.6
3.8
3.1
2.3
3.2
3.5
22.3
22.2
19.4
20.1
19.3
19.3

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Figure 6. Plots of the Stokes shifts as a function of the solvent polarity parameter Δf (ε, n2) for
compounds 3a–f.
2.2. Theoretical Calculations
Electronic Structure and Transition Energies
To gain a more precise insight into the spectroscopic properties of the newly synthesized
compounds, theoretical calculations were performed. The Time-Dependent Density Functional
Theory with the CAM-B3LYP functional and the 6-311+G(d, p) basis set (TD-DFT//CAM-B3LYP//6-
311+G (d, p)) appeared to be the best method for this task. Additionally, the PCM model was used to
evaluate the solvent effects. The results are presented in Table 5, which includes the characteristics
(oscillator strength-f, transition energy–λ/nm together with the main orbital configurations) of the
main electronic transitions that were responsible for the band maxima. Furthermore, Figure 7
presents a visualization of the border orbitals HOMO and LUMO, which were involved in the main
electronic transitions. The applied method gave satisfactory results compared to the experimental
data. The high values of the oscillator strengths resulted from the method that was used for the
calculation. According to the theoretical data, the experimental absorption maximum of the first band
was mainly assigned to the S0→S1 transition. This transition employed the HOMOs and LUMOs,
whose shapes indicated a π→π* type of transition. In the case of compound 3a, the electron density
of the highest occupied orbital was located on the phenyl ring with the N(CH3)3 group, and the lowest
unoccupied orbital embraced the quinoline structure. There was a distinct charge transfer between
the donor and the acceptor units. A similar property could be observed in the case of the rest of the
compounds. However, the charge transfer process was no longer distinguishable. The absorption
spectra of all compounds consisted of two bands. The second bands were assigned to the higher

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transitions, such as S0→S5 or S0→S6, and used the HOMO orbitals with a lower energy (like HOMO−1,
HOMO−3, and HOMO−6) and the LUMO orbitals with a higher energy (such as LUMO+1 and
LUMO+2, and LUMO+3). Quantum calculations were also performed for the spin-allowed singlet→
singlet (Sn→S0) transitions, in order to determine the theoretical values of the fluorescence emission
energy. The results are presented in Table 6. The obtained values of the emission maxima correctly
reflected the experimental data.
Table 5. Electronic transition data obtained by TD–DFT//B3LYP//6-311+G (d), using a polarizable
continuum model (PCM) (solvent–ethanol) for compounds 3a–f, at the DFT-optimized geometry.
Electronic
λ
Molecular
Percentage
Experimental
Compound
3a
f
Transition (nm)
Orbital
Contribution (%)
λ (nm)
388
S0→S1
S0→S6
S0→S1
367
274
356
2.2134
0.0800
1.8851
HOMO→LUMO
HOMO−1→LUMO+4
HOMO→LUMO
HOMO−1→LUMO
HOMO−1→LUMO+1
HOMO→LUMO
HOMO−1→LUMO
HOMO−3→LUMO+1
HOMO→LUMO
HOMO−2→LUMO
HOMO−6→LUMO
HOMO→LUMO
HOMO→LUMO+2
HOMO−1→LUMO
HOMO→LUMO
HOMO→LUMO+2
HOMO→LUMO+3
57
55
86
23
18
88
23
18
85
15
14
88
21
12
87
21
11
282
368
3b
3c
3d
3e
3f
S0→S5
S0→S1
S0→S5
S0→S1
S0→S5
S0→S2
S0→S5
S0→S1
S0→S6
271
355
267
364
280
354
265
358
265
0.2781
1.8087
0.3937
2.1305
0.2265
1.6569
0.4493
1.7470
0.2758
276
370
272
372
274
362
270
366
272
Table 6. Emission data obtained for 3a–f by TD–DFT//B3LYP//6-311+G (d), using a PCM model
(ethanol).
Electronic
Transition
S0→S1
Λ
(nm)
532
502
501
540
-
Molecular
Orbital
Percentage
Experimental
Compound
f
Contribution (%)
λ (nm)
519
3a
3b
3c
3d
3e
3f
2.5873
2.2812
2.2154
2.4128
-
HOMO→LUMO
HOMO→LUMO
HOMO→LUMO
HOMO→LUMO
-
91
93
93
91
-
S0→S1
512
S0→S1
514
S0→S1
512
S0→S1
514
S0→S1
522
1.9072
HOMO→LUMO
91
514
Due to the structural relaxation, there was a decrease in the dihedral angle between the
styrylquinoline plane and the phenyl ring (C38–N28–C27–C24, red color in Figure 8). In other words,
the molecule became more planar. As a result of this change, the energy difference between the
HOMO and LUMO also decreased and the energy of the emission was lower than the energy of the
excitation, which resulted in a larger Stokes shift (Figure 9). These relationships were in agreement
with the Jablonski diagram. The changes in the dihedral angles are presented in Table 7 and are
depicted in Figure 8.

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HOMO
LUMO
3a
E = −6.68 eV
E = −1.24 eV
3b
E = −7.09 eV
E = −1.29 eV
3c
3d
3e
E = −7.10 eV
E = −7.10 eV
E = −7.19 eV
E = −1.28 eV
E = −1.41 eV
E = −1.34 eV

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3f
E = −6.55 eV
E = −1.85 eV
Figure 7. Visualization of the molecular orbitals that were involved in the main electronic transitions.
Figure 8. The geometry of compound 3a in the ground and excited states, which was obtained by
TD–DFT//B3LYP//6-311+G (d).
Table 7. Comparison of a compound’s geometry in the ground and excited states.
Dihedral Angle (C38-N28-C27-C24)/[°]
Compound
Ground State S0
Excited State S1
Difference
38.0
3a
3b
3c
3d
3e
3f
42.5
42.5
42.9
41.7
-
4.5
15.9
17.0
13.3
-
26.6
25.9
28.4
-
42.7
16.0
26.7

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Figure 9. Geometry relaxation of compound 3a after photoexcitation. The more planar structure was
the reason for the decrease in the energy gap between the HOMO and LUMO orbitals. CC—
configuration change and IC—internal conversion.
2.3. Biological Activity and Cellular Imaging
The combination of large Stokes shifts, absorption over 350 nm, and a good fluorescent profile
predestine the investigated compounds to be molecular probes for biological staining. For such
applications, various physicochemical properties such as lipophilicity, protein binding, ion chelation,
or halochromism could be exploited, in order to achieve specific staining effects. However, inactivity
toward the biological systems is of special importance. We performed cytotoxicity tests on the human
colon cancer cell line (HCT 116) and the normal human fibroblast cell line (NHDF), to evaluate the
potential applications in this appealing field. The results are shown in Table 8. In general, all tested
compounds had no prolonged cytotoxicity against the HCT 116 and NHDF cells, in concentrations
that are useful for staining.
Incubation with synthesized compounds 3a–f for 72 h did not affect the viability of the cells. This
fact, together with their good spectroscopic properties, make them potentially useful as fluorescent
dyes for biological specimens. However, with regards to the possibility of the decomposition of
compounds in an aqueous environment, only two derivatives (3e,d) were selected for the cell imaging
experiments. Additionally, for the co-localization experiments, the incubation time was reduced to 1
h, in order to minimize the risk of decomposition. The site of the accumulation of the selected
compounds was examined in the HCT 116 cells, using distinct specific-organelle markers for
mitochondria (MitoTracker Orange), lysosomes (LysoTracker® Red DND-99), and the endoplasmic
reticulum (ER-Tracker™ Red BODIPY® TR Glibenclamide). Apparently, these compounds tend to
accumulate in the membranous structures, and their associated organelles accumulate mainly in the
mitochondria and endoplasmic reticulum (ER) (Figure 10). Moreover, there is a subtle but evident
relationship between the structure of a dye and the apparent site of accumulation. According to the
scientific literature, it has not yet been determined why the fluorescent probes for the ER are selective.
However, compounds with an affinity to the mitochondria and ER, including compounds 3d and 3e
have a few common features, which might be crucial for their behavior in cells. Namely, these dyes
usually have a medium-size aromatic system (20–30 atoms) and a moderate (or higher) lipophilicity
(3d—logP = 7.1, 3e—logP = 5.3). Furthermore, imines might be partially protonated in a cell, which
might increase their affinity to the negatively charged mitochondrial membrane. The cationic
character of a compound was also a typical feature for the ER dyes [54].
Table 8. Antiproliferative activity on the HCT 116 cell line.
Cytotoxicity–IC50 [µM]
Compound
HCT 116
>25
NHDF
>25
3a
3b
3c
3d
3e
3f
>25
>25
>25
>25
>25
>25
>25
>25
>25
>25

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Figure 10. Fluorescence images of the HCT 116 cells that were incubated with 3d (I) and 3e (II) (both
at a concentration of 25 µM), for 1 h at 37 °C. The images show a co-localization with specific-organelle
trackers—the mitochondria, lysosomes, and endoplasmic reticulum. The first panel represents the
fluorescence of the 3d and 3e compounds alone, the middle panel—the specific dyes, and the last—
the merged. Scale bars indicate 50 µm. The absorption profile of the tested compounds enabled their

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excitation with a DAPI wavelength and the large Stokes shift helped to minimize the self-quenching
effect and also boosted the signal-to-noise ratio, which in turn enabled us to obtain images of a good
resolution.
3. Experimental Method
3.1. Materials and Basic Measurements
All reagents were purchased from Sigma-Aldrich. The TLC experiments were performed on
aluminum-backed silica gel 40 F254 plates (Merck, Darmstadt, Germany). The plates were
illuminated under UV (254 nm and 365 nm). The melting points were determined on an Optimelt
MPA-100 apparatus (SRS, Stanford, CA, USA). The structures of the newly obtained compounds
were confirmed by NMR spectroscopy. All NMR spectra were recorded in deuterated DMSO-d6 as
the solvent on a Bruker AM-series (Bruker BioSpin Corp., Karlsruhe, Germany). The working
frequency for each compound was known. Chemical shifts were reported in ppm (δ), against the
internal standard Si(CH3)4. The mass spectra were performed on a WATERS LCT Premier XE system
(high-resolution mass spectrometer with a Time Of Flight (TOF) analyzer).
3.2. Synthesis and Characterization
3.2.1. Synthesis of the Starting Material
Step 1: 2-[(E)-2-(4-nitrophenyl)ethenyl]quinoline (1): The synthesis was performed according to the
procedure described in [10]. The quinaldine (10 mmol) and 4-nitrobenzaldehyde (10 mmol) were
mixed in acetic anhydride and heated for 20 h at 130 °C. The excess solvent was evaporated in vacuo
and the obtained solid was purified using column chromatography, with dichloromethane as the
eluent. The product was obtained as a bright yellow solid with a yield of 73%, and a melting point of
1
167 °C. H NMR (400 MHz, DMSO) δ 8.41 (d, J = 8.6 Hz, 1H), 8.27 (d, J = 8.6 Hz, 2H), 8.06–7.99 (m,
4H), 7.97 (d, J = 6.4 Hz, 1H), 7.94 (d, J = 8.6 Hz, 1H) 7.78 (t, J = 7.8 Hz, 1H), 7.73 (d, J = 16.4 Hz, 1H),
7.60 (t, J = 7.4 Hz, 1H). 13C NMR (126 MHz, DMSO) δ 155.27, 148.11, 147.36, 143.49, 137.24, 133.63,
132.22, 130.52, 129.30, 128.67, 128.35, 127.77, 127.17, 124.53, 120.82.
Step 2: 4-[(E)-2-(quinolin-2-yl)ethenyl]aniline (2): In a round-bottom flask, compound 1 was placed in
ethanol, together with anhydrous SnCl2, at a molar ratio of 1:5. The mixture was heated for 2 h at a
temperature of 70 °C, under nitrogen. Then, the mixture was cooled, placed in a flask with ice, and
neutralized with a 5% solution of sodium bicarbonate (NaHCO3). Next, the extraction with ethyl
acetate was performed, the organic layer was washed with brine and dried over anhydrous sodium
sulfate (Na2SO4). The product was obtained as a red solid with a yield of 61% and a melting point of
174 °C. 1H NMR (400 MHz, DMSO) δ 8.26 (d, J = 8.6 Hz, 1H), 7.93 (d, J = 8.4 Hz, 1H), 7.89 (d, J = 8.0
Hz, 1H) 7.77 (d, J = 8.6 Hz, 1H), 7.73–7.69 (m, 1H), 7.67 (d, J = 16.1, 1H), 7.50 (t, J = 7.4 Hz, 1H), 7.42 (d,
J = 8.3 Hz, 2H), 7.13 (d, J = 16.2 Hz, 1H), 6.61 (d, J = 8.3 Hz, 2H), 5.54 (s, 2H). 13C NMR (126 MHz,
DMSO) δ 157.03, 150.39, 148.22, 136.54, 135.54, 130.07, 129.21, 128.81, 128.18, 127.10, 125.95, 124.22,
123.28, 120.06, 114.33. 13C NMR (126 MHz, DMSO) δ 157.03, 150.39, 148.22, 136.54, 135.54, 130.07,
129.21, 128.81, 128.18, 127.10, 125.95, 124.22, 123.28, 120.06, 114.33.
3.2.2. Representative Procedure for Compounds 3a–f
Compound 2 and the corresponding aldehyde were placed in a round-bottom flask, at a ratio of
1:1. Then, 5 mL of ethanol and 3 drops of acetic acid were added. The mixture was exposed to
microwave irradiation for 20 min in conditions of 50 W and 83 °C. The obtained compounds were
filtered and washed with ethanol. The general route for the synthesis is presented in Figure 2.
(N-{[4-(Dimethylamino)phenyl]methylidene}-4-[(E)-2-(quinolin-2-yl)ethenyl]aniline (3a): The product was
1
obtained as a yellow solid with an yield of 62% and a melting point of 202 °C. H NMR (500 MHz,
DMSO) δ 8.49 (s, 1H), 8.35 (d, J = 8.6 Hz, 1H), 7.99 (d, J = 8.5 Hz, 1H), 7.95 (d, J = 7.3 Hz, 1H), 7.87 (d, J
= 9.00 Hz, 1H), 7.86 (d, J = 16.5 Hz, 1H), 7.79–7.73 (m, 5H), 7.56 (t, J = 7.4 Hz 1H), 7.46 (d, J = 16.3 Hz,

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1H), 7.27 (d, J = 8.4 Hz, 2H), 6.80 (d, J = 9.0 Hz, 2H), 3.03 (s, 6H). 13C NMR (126 MHz, DMSO) δ 160.31,
156.29, 153.07, 153.10, 148.24, 136.84, 134.33, 133.63, 130.88, 130.21, 129.11, 128.69, 128.22, 128.14,
127.47, 126.49, 124.40, 121.94, 120.39, 112.01. ESI–MS calcd. for C26H24N3 378.1970 [M+H]+ found
378.1973.
N-[(3,4-Dimethoxyphenyl)methylidene]-4-[(E)-2-(quinolin-2-yl)ethenyl]aniline (3b): The product was
1
obtained as a yellow solid with an yield of 70% and a melting point of 172 °C. H NMR (500 MHz,
DMSO) δ 8.59 (s, 1H), 8.36 (d, J = 8.6 Hz, 1H), 8.00 (d, J = 8.4 Hz, 1H), 7.95 (d, J = 7.5 Hz, 1H), 7.87 (d, J
= 9.0 Hz, 1H), 7.86 (d, J = 16.5 Hz, 1H), 7.80 (d, J = 8.4 Hz, 2H), 7.76 (t, J = 7.7 Hz, 1H), 7.58–7.55 (m,
2H), 7.49 (d, J = 16.5 Hz, 1H), 7.48 (d, J = 8.5 Hz, 1H), 7.32 (d, J = 8.4 Hz, 2H), 7.12 (d, J = 8.4 Hz, 1H),
3.85 (s, 6H). 13C NMR (126 MHz, DMSO) δ 160.66, 156.20, 152.43, 152.37, 149.51, 148.16, 136.9, 134.22,
134.14, 130.30, 129.50, 129.10, 128.77, 128.46, 128.28, 127.46, 126.59, 124.74, 122.09, 120.45, 111.79,
109.88. ESI–MS calcd. for C26H23N2O2 395.1760 [M+H]+ found 395.1763.
4-(N-{4-[(E)-2-(quinolin-2-yl)ethenyl]phenyl}carboximidoyl)phenol (3c): The product was obtained as a
yellow solid with an yield of 67% and a melting point of 248 °C. 1H NMR (500 MHz, DMSO) δ 10.17
(s, 1H), 8.54 (s, 1H), 8.36 (d, J = 8.6 Hz, 1H), 8.00 (d, J = 8.4 Hz, 1H), 7.95 (d, J = 7.9 Hz, 1H), 7.88 (d, J =
8.5 Hz, 1H), 7.86 (d, J = 16.5 Hz, 1H), 7.82–7.73 (m, 5H), 7.56 (t, J = 7.5 Hz, 1H), 7.47 (d, J = 16.3 Hz, 1H),
7.29 (d, J = 8.3 Hz, 2H), 6.90 (d, J = 8.5 Hz, 2H). 13C NMR (126 MHz, DMSO) δ 161.26, 160.54, 156.22,
152.62, 148.16, 136.92, 134.20, 134.02, 131.30, 130.30, 129.09, 128.74, 128.33, 128.27, 127.97, 127.45,
126.57, 122.03, 120.43, 116.17. ESI–MS calcd. for C24H19N2O 351.1497 [M+H]+ found 351.1500.
N-[(2E)-3-phenylprop-2-en-1-ylidene]-4-[(E)-2-(quinolin-2-yl)ethenyl]aniline (3d): The product was
1
obtained as a yellow solid with an yield of 68% and a melting point of 195 °C. H NMR (400 MHz,
DMSO) δ 8.49 (d, J = 8.9 Hz, 1H), 8.36 (d, J = 8.6 Hz, 1H), 8.00 (d, J = 8.4 Hz, 1H), 7.95 (d, J = 8.0 Hz,
1H), 7.89 (d, J = 8.8, 1H), 7.87 (d, J = 16.8 Hz, 1H), 7.81–7.70 (m, 5H), 7.51 (s, 1H), 7.49, (d, J = 16.4 Hz,
1H), 7.45–7.40 (m, 4H), 7.30 (d, J = 8.3 Hz, 2H), 7.20 (dd, J = 16.0, 8.9 Hz, 1H).13C NMR (126 MHz,
CD2Cl2) δ 161.47, 155.89, 152.07, 148.34, 144.13, 136.21, 135.69, 134.55, 133.53, 129.63, 129.60, 129.08,
128.90, 128.66, 128.43, 128.16, 127.53, 127.36, 126.04, 121.49, 119.60. ESI–MS calcd. for C26H21N2
361.1705 [M+H]+ found 361.1707.
N-[(2-fluorophenyl)methylidene]-4-[(E)-2-(quinolin-2-yl)ethenyl]aniline (3e): The product was obtained as
a yellow solid with an yield of 68% and a melting point of 124 °C. 1H NMR (400 MHz, DMSO) δ 8.85
(s, 1H), 8.37 (d, J = 8.6 Hz, 1H), 8.14 (t, J = 7.1 Hz, 1H), 8.00 (d, J = 8.4 Hz, 1H), 7.96 (d, J = 8.0 Hz, 1H),
7.89 (d, J = 8.8, 1H), 7.87 (d, J = 16.8 Hz, 1H), 7.83 (d, J = 8.3 Hz, 2H), 7.76 (t, J = 7.6 Hz, 1H), 7.67–7.54
(m, 2H), 7.51 (d, J = 16.3 Hz, 1H), 7.43–7.33 (m, 4H). 13C NMR (126 MHz, DMSO) δ 163.65, 161.64,
156.11, 153.80, 151.80, 148.14, 136.96, 135.09, 134.22 (d, J = 8.4 Hz), 133.99, 130.32, 129.01 (d, J = 23.5
Hz), 128.80, 128.31 (d, J = 8.5 Hz), 127.49, 126.64, 125.42, 123.88 (d, J = 8.8 Hz), 122.26, 120.46, 116.69
(d, J = 20.6 Hz). ESI–MS calcd. for C24H18FN2 353.1448 [M+H]+ found 353.1452.
N-[(2-ethoxyphenyl)methylidene]-4-[(E)-2-(quinolin-2-yl)ethenyl]aniline (3f): The product was obtained as
a yellow solid with an yield of 55% and a melting point of 167 °C. 1H NMR (500 MHz, DMSO) δ 8.91
(s, 1H), 8.36 (d, J = 8.6 Hz, 1H), 8.04 (dd, J = 7.7, 1.8 Hz, 1H), 8.00 (d, J = 8.5 Hz, 1H), 7.96 (d, J = 8.1 Hz,
1H), 7.89 (d, J = 8.5 Hz, 1H), 7.87 (d, J = 16.5 Hz, 1H), 7.80 (d, J = 8.4 Hz, 2H), 7.78–7.74 (m, 1H), 7.57
(dt, J = 8.0, 6.9, 1.1 Hz, 1H), 7.54–7.51 (m, 1H), 7.49 (d, J = 16.4 Hz, 1H), 7.29 (d, J = 8.4 Hz, 2H), 7.17 (d,
J = 8.2 Hz, 1H), 7.07 (t, J = 7.5 Hz, 1H), 4.18 (q, J = 7.0 Hz, 2H), 1.40 (t, J = 7.0 Hz, 3H). 13C NMR (126
MHz, DMSO) δ 159.23, 156.18, 156.14, 152.85, 148.20, 136.91, 134.51, 134.12, 133.75, 130.26, 129.12,
128.80, 128.65, 128.25, 127.49, 127.43, 126.58, 124.55, 121.96, 121.12, 120.44, 113.54, 64.52, 14.76. ESI–MS
calcd. for C26H23N2O 379.1810 [M+H]+ found 379.1806.
3.3. Biological Properties
3.3.1. Cell Culture
The human colon carcinoma cell line HCT 116 was obtained from ATCC. The cells were grown
as monolayer cultures in 75 cm2 flasks (Nunc) in Dulbecco’s modified Eagle’s medium with antibiotic

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gentamicin (200 µL/100 mL medium). DMEM was supplemented with 12% heat-inactivated fetal
bovine serum (Sigma). The cells were cultured under standard conditions, at 37 °C in a humidified
atmosphere at 5% CO2.
3.3.2. Cytotoxicity Studies
The cells were seeded in 96-wells plates (Nunc) at a density of 5 × 103 cells/well and incubated
at 37 °C for 24 h. The assay was performed, following a 72 h incubation with varying concentrations
of the compounds (3a–f) that were being tested. Then, 20 µL of the CellTiter 96®AQueous One
Solution-MTS (Promega) was added to each well (with 100 µL DMEM without phenol red) and
incubated for 1 h at 37 °C. The optical densities of the samples were analyzed at 490 nm, using a
multi-plate reader Synergy 4 (BioTek, BioTek Instruments, Winooski, VT, USA). The results were
calculated as the inhibitory concentration (IC50) values using the GraphPad Prism 7, and are
expressed as a percentage of the control. Each individual compound was tested in triplicates in a
single experiment, with each experiment being repeated three times.
3.3.3. Cellular Staining
The cells were seeded onto glass slides at a density of 3 × 105 cells/slide and were incubated at
37 °C for 24 h. Then, the medium was removed and the solutions of 3a–f, at a concentration of 25 µM,
were added. The cells were then further incubated for 1 h and 2 h, under standard conditions—at 37
°C in a humidified atmosphere at 5% CO2. After incubation, the cells were washed three times with
PBS and fixed with 3.7% paraformaldehyde, for 10 min. The cellular staining with compounds being
tested was observed using a Nikon Eclipse Ni-U microscope, equipped with a Nikon Digital DS-Fi1-
U3 camera and the corresponding software (Nikon, Tokyo, Japan).
3.3.4. Subcellular Localization
The cells were seeded onto glass slides at a density of 3 × 105 cells/slide and incubated at 37 °C
for 24 h. Then, the medium was removed and the solutions of 3a–f, at a concentration of 25 µM, were
added. The cells were then further incubated for 1 h and 2 h. After incubation, the cells were rinsed
with PBS (pH 7.2) and the staining procedures were performed according to the protocols of the
providers. In brief, a serum-free medium that contained MitoTracker® Orange (100 nM, 30 min
incubation, Molecular Probes), ER-Tracker™ Red BODIPY® TR Glibenclamide (1 µM, 30 min
incubation, Molecular Probes), and LysoTracker® Red DND-99 (500 nM, 1 h incubation, Molecular
Probes) or Hoechst 33342 (6.5 µM, 15 min incubation, Molecular Probes) were added. After staining
with organelle-specific trackers, the cells were washed three times with PBS and fixed with 3.7%
paraformaldehyde for 10 min. The subcellular localization was observed using a Nikon Eclipse Ni-U
microscope equipped with a Nikon Digital DS-Fi1-U3 camera and the corresponding software. The
analysis and processing of the images were performed using an Image J 1.41 (Wayne Rasband,
National Institutes of Health, Bethesda, MD, USA).
3.4. Spectroscopic Studies
The absorption and fluorescence spectra were measured at room temperature in a 10 mm quartz
cell, using a U-2900 spectrophotometer (Hitachi) and an F-7000 spectrofluorometer (Hitachi),
respectively. Due to the low solubility of the compounds in the applied solvents, the solutions that
were used in the test of the solvatochromic behavior contained 1% of DMSO, to improve the
solubility. The quantum yields of fluorescence were determined using the absolute method at room
temperature, using an integrating sphere with a solvent as a blank on an FLS-980 spectrophotometer.
The compounds were excited at the wavelength that corresponded with the absorption wavelength
of the compounds.

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3.5. Theoretical Calculations
The geometrical parameters of an investigated compound and its electronic properties in the
ground and excited states were calculated within the DFT and TD–DFT approximations, using the
Gaussian 09 software package [53] with the CAM-B3LYP exchange-correlation functional [55–57] and
the 6-311+G (d, p) basis set. The effects of the solvent were evaluated using the polarizable continuum
model (PCM) [58,59] in which a cavity was created via a series of overlapping spheres [60] with the
standard dielectric constants (ε) of 24.55 for ethanol. The lowest 50 singlet–singlet vertical electronic
excitations based on the B3LYP optimized geometries and the optimization of the first excited states
were computed using the time-dependent density functional theory (TD–DFT) formalism [61]. The
visualization of the molecular orbitals and analysis of the energy levels were performed using the
Chemissian software.
4. Conclusions
A series of styrylquinoline dyes with a Schiff base fragment was designed and synthesized, and
their physicochemical parameters and spectroscopic features were determined. All compounds had
a high Stokes shift and strong solvatochromism. Along with the calculations that were carried out
accordingly, these suggested an intramolecular charge-transfer mechanism of fluorescence. All
obtained compounds appeared to be non-toxic in a broad range of concentrations, during the in vitro
experiments. Styrylquinolines maintained their fluorescent properties in biological systems, thus
allowing for staining applications to use them. Their selective accumulation in the cellular organelles
was also investigated, which indicated a preference for structures such as the mitochondria and the
endoplasmic reticulum.
Author Contributions: Conceptualization R.M.; chemical synthesis B.C., M.K.; physical investigation B.C., A.S.;
biological assays K.M., A.M.-W., Funding acquisition K.M., A.M.-W.; original draft preparation B.C., K.M.
A.M.W.; text review and editing R.M., project administration R.M. All authors have read and agreed to the
published version of the manuscript.
Funding: The authors wish to thank the National Center for Science grants: 2014/13/D/NZ7/00322 (A.M.W.)
and 2016/23/N/NZ7/00351 (K.M.).
Acknowledgments: The Gaussian 09 calculations were performed in the Wroclaw Centre for Networking and
Supercomputing, WCSS, Wrocław, Poland: http://www.wcss.wroc.pl, under calculation Grant No. 18.
Conflicts of Interest: The authors declare no conflict of interest.
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