Photoactivatable BODIPYs Designed to Monitor the Dynamics of Supramolecular Nanocarriers

Article abstract of DOI:10.1021/ja5125308

Self-assembling nanoparticles of amphiphilic polymers can transport hydrophobic molecules across hydrophilic media and, as a result, can be valuable delivery vehicles for a diversity of biomedical applications. Strategies to monitor their dynamics noninva

Full text of DOI:10.1021/ja5125308

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Article
Photoactivatable BODIPYs Designed to Monitor
the Dynamics of Supramolecular Nanocarriers
Yang Zhang, Subramani Swaminathan, Sicheng Tang, Jaume Garcia,
Marcia Boulina, Burjor Captain, James D. Baker, and Françisco M. Raymo
J. Am. Chem. Soc., Just Accepted Manuscript • DOI: 10.1021/ja5125308 • Publication Date (Web): 20 Mar 2015
Downloaded from http://pubs.acs.org on March 23, 2015
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Journal of the American Chemical Society
Photoactivatable BODIPYs Designed to Monitor
the Dynamics of Supramolecular Nanocarriers
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Yang Zhang,† Subramani Swaminathan,† Sicheng Tang,† Jaume Garcia-
Amorós,† Marcia Boulina,‡ Burjor Captain,† James D. Baker§ and
†
Françisco M. Raymo*,
Laboratory for Molecular Photonics, Department of Chemistry, University of Miami, 1301 Memorial
Drive, Coral Gables, FL 33146-0431, Department of Biology, University of Miami, 1301 Memorial
Drive, Coral Gables, FL 33146-0431 and Analytical Imaging Core Facility, Diabetes Research
Institute, University of Miami, 1450 NW 10th Avenue, Miami, FL 33136
E-Mail: fraymo@miami.edu
Abstract: Self-assembling nanoparticles of amphiphilic polymers can transport hydrophobic molecules
across hydrophilic media and, as a result, can be valuable delivery vehicles for a diversity of biomedical
applications. Strategies to monitor their dynamics noninvasively and in real time are, therefore, essential
to investigate their translocation within soft matrices and, possibly, rationalize the mechanisms
responsible for their diffusion in biological media. In this context, we designed molecular guests with
photoactivatable fluorescence for these supramolecular hosts and demonstrated that the activation of the
fluorescent cargo, under optical control, permits the tracking of the nanocarrier translocation across
hydrogel matrices with the sequential acquisition of fluorescence images. In addition, the mild
illumination conditions sufficient to implement these operating principles permit fluorescence activation
†
‡
§
Department of Chemistry
Diabetes Research Institute
Department of Biology
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within developing Drosophila Melanogaster embryos and enable the monitoring of the loaded
nanocarriers for long periods of time with no cytotoxic effects and no noticeable influence on
embryogenesis. These photoresponsive compounds combine a borondipyrromethene (BODIPY)
chromophore and a photocleavable oxazine within their covalent skeleton. Under illumination at an
appropriate activation wavelength, the oxazine ring cleaves irreversibly to bring the adjacent BODIPY
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fragment in conjugation with an indole heterocycle.
This structural transformation shifts
bathochromically the BODIPY absorption and permits the selective excitation of the photochemical
product with concomitant fluorescence. In fact, these operating principles allow the photoactivation of
BODIPY fluorescence with large brightness and infinite contrast. Thus, our innovative structural design
translates into activatable fluorophores with excellent photochemical and photophysical properties as
well as provides access to a general mechanism for the real-time tracking of supramolecular nanocarriers
in hydrophilic matrices.
Introduction
Amphiphilic polymers incorporate hydrophilic and hydrophobic segments within their macromolecular
backbone.1–7 In aqueous environments, multiple copies of these macromolecular constructs assemble
spontaneously into single particles with nanoscaled dimensions. A subtle balance of enthalpic and
entropic terms brings the hydrophobic segments of distinct polymers together in order to minimize their
direct exposure to water. The concomitant solvation of the hydrophilic counterparts ensures the effective
dispersion of the nanoparticles and prevents their further association into large aggregates. The overall
result is the spontaneous formation of supramolecular hosts with hydrophilic surface and hydrophobic
core, where molecular guests can be encapsulated. Indeed, such nanostructured constructs can capture
multiple hydrophobic molecules in their interior and transport them across hydrophilic media.
Furthermore, the structural design of these nanocarriers can be engineered to allow their internalization
in living cells and the transfer of their cargo from the extracellular to the intracellular space. As a result,
these fascinating supramolecular assemblies can become viable delivery vehicles for a diversity of
biomedical applications.8–26 In this context, the development of strategies to monitor their translocation
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across soft matrices in real time, without significant perturbations of the sample, would be particularly
valuable. If available, such methods would offer the opportunity to investigate and optimize the transport
properties of the nanocarriers as well as facilitate the rationalization of the mechanisms responsible for
their cellular internalization. Specifically, the noninvasive character of fluorescence measurements,27
together with the ability of these supramolecular containers to preserve the photophysical properties of
encapsulated chromophores,28 would be especially convenient to design appropriate mechanisms to track
the dynamics of these particular nanoparticles.
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Fluorescence recovery after photobleaching (FRAP) is a valuable experimental protocol to monitor the
diffusion of fluorescent species within a diversity of samples in real time.29 It is based on the illumination
of a geometrically-defined portion of the sample at the excitation wavelength (Ex) of the chromophores
with sufficiently high intensity and for an appropriately long time to bleach the emissive species and turn
fluorescence off in the irradiated area. The subsequent and sequential acquisition of images, under
illumination of the entire sample at Ex with moderate intensity, then reveals the gradual recovery of
fluorescence in the bleached area, if the many intact fluorophores present in the rest of the sample can
slowly diffuse into this particular region. Under these conditions, the temporal dependence of the
recovering emission intensity can provide quantitative information on the diffusion process. The
bleaching step necessary for the local fluorescence depletion, however, limits significantly the
implementation of FRAP. Indeed, the relatively high illumination intensities required for the
photoinduced degradation of the emissive species can have deleterious consequences on the sample under
analysis. In addition, fairly long irradiation times are often required to ensure the complete bleaching of
the fluorophores, preventing the investigation of diffusion processes that occur on relatively short
timescales. Furthermore, significant amounts of molecular oxygen are also necessary to ensure the
photodegradation of most fluorophores, restricting the application of FRAP to oxygenated samples.
The limitations inherent to the operating principles of FRAP can be overcome with the use of
photoactivatable fluorophores.30–35 These compounds require irradiation at a given activation
wavelength (Ac), prior to illumination at Ex, in order to produce fluorescence. Generally, the absorption
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of photons at Ac induces a photochemical reaction designed to enable the subsequent absorption at Ex
and the population of the excited state responsible for fluorescence. Alternatively, the photochemical
transformation can be engineered to suppress a quenching pathway that would, otherwise, deactivate
nonradiatively the excited state responsible for emission. In both instances, a concatenation of
photochemical and photophysical processes ensures the activation of fluorescence exclusively after
illumination at both wavelengths. Under these conditions, the irradiation of a portion of the sample,
labeled with photoactivatable fluorophores, at Ac turns fluorescence on exclusively in the illuminated
area. The subsequent and sequential acquisition of images, under irradiation at Ex of the entire specimen,
then reveals the gradual decrease in emission intensity within the activated area, if the fluorescent species
can slowly diffuse out of this region. Once again, the temporal evolution of the emission intensity can
then provide quantitative information on the diffusion process. In contrast to the bleaching step of FRAP,
however, photoactivation generally occurs efficiently even after short irradiation pulses at moderate
illumination intensity and does not require any molecular oxygen. As a result of these obvious
advantages, photoactivatable fluorophores are becoming invaluable tools to monitor dynamic events in
a diversity of biological samples and synthetic materials.36–41 Additionally, the spatiotemporal control
of fluorescence, possible with these molecules, offers also the opportunity to overcome the limitations
that diffraction imposes on the spatial resolution of conventional images and permits the visualization of
structural features at the nanometer level.42–49 In principle, the entrapment of such photoresponsive
probes within the hydrophobic interior of self-assembling nanoparticles of amphiphilic polymers would
enable the tracking of the resulting supramolecular constructs, and perhaps even their localization with
subdiffraction resolution, on the basis of fluorescence photoactivation.
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The borondipyrromethene (BODIPY) chromophore is a versatile building block for the construction of
fluorescent probes.50–55 It can be assembled conveniently from appropriate pyrrole precursors and
modified in any position along the perimeter of its heterocyclic platform with established synthetic
procedures. Furthermore, BODIPY derivatives generally have large molar absorption coefficients and
large fluorescence quantum yields. In spite of their synthetic accessibility as well as of their excellent
photophysical properties, mechanisms to photoactivate BODIPY fluorescence remain limited only to a
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few representative examples.56,57 In these systems, a photocleavable quencher is attached covalently to
the BODIPY platform in order to suppress fluorescence. Specifically, excitation of the BODIPY
chromophore at Ex results in the transfer of one electron to or from the quencher, instead of the radiative
deactivation of the former. Illumination at Ac, however, cleaves the covalent linkage between quencher
and fluorophore. The physical separation of the two prevents electron transfer upon excitation of the
latter and ensures fluorescence instead. Nonetheless, these photoactivation mechanisms translate into
minimal changes in the absorption spectrum of the BODIPY chromophore. As a result, absorption at Ex
occurs before and after activation and, unless the quenching mechanism is designed to be extremely
efficient, noticeable background fluorescence is detected. Additionally, the lack of any significant
change in absorption with activation prevents the possibility to bleach selectively the activated state and,
hence, to implement schemes for subdiffraction imaging.58–64 Indeed, the exclusive bleaching of the
emissive species, after their localization at the single-molecule level, is an essential step in the sequential
reconstruction of images with subdiffraction resolution on the basis of fluorescence activation. Thus, the
identification of alternative operating principles to activate the emission of BODIPY chromophores with
concomitant changes in absorption would be particularly valuable to develop photoactivatable
fluorophores with optimal photochemical and photophysical properties to monitor diffusion as well as to
acquire superresolution images. In particular, the attachment of a BODIPY chromophore to a
photoswitchable auxochrome, instead of a photocleavable quencher, appears to be a viable strategy to
regulate the absorption characteristics of the emissive component under optical control. Such structural
design should provide the opportunity to photoinduce a sufficient shift in the BODIPY absorption to
permit the selective excitation of the photochemical product in the presence of the corresponding
reactant. As a result, infinite fluorescence contrast and exclusive product bleaching can in principle be
achieved on the basis of this mechanism for the photoactivation of BODIPY emission.
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In search of strategies to photoactivate fluorescence, we developed a family of molecular dyads
integrating fluorescent and photochromic components within the same covalent skeleton.65 Upon
illumination at Ac, the heterocyclic core of the photochrome opens to extend the electronic conjugation
of the fluorophore. This structural transformation shifts bathochromically the main absorption band of
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the fluorophore and permits the exclusive excitation of the photogenerated state at Ex. The
photochemical reaction responsible for fluorescence activation, however, is thermally reversible. The
fluorescent state survives only for a few microseconds before reverting to the initial species. As a result,
dynamic processes occurring on timescales that are longer than the lifetime of the activated species
cannot be monitored with these photoswitchable systems. In order to overcome this limitation, we
envisaged the possibility of modifying the structural design of the photochromic component of our dyads
to enable the irreversible, rather than reversible, activation of their fluorescence. In this article, we report
the implementation of these operating principles as well as demonstrate that they can be employed to
activate irreversibly the fluorescence of BODIPY chromophores and monitor the translocation of
supramolecular nanocarriers in soft matrices.
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Results and Discussion
Design, Synthesis and Structural Characterization: The photochromic component of our switchable
molecular dyads fuses 2H,3H-indole and 2H,4H-benzo[1,3]oxazine heterocycles within its molecular
skeleton (a in Figure 1).66 The latter fragment incorporates a nitro group in the para-position, relative to
the oxygen atom, and is responsible for absorbing the activating photons at Ac. Upon excitation, the
[C–O] bond at the junction of the two heterocyclic fragments cleaves to open the oxazine ring and
generate a zwitterionic isomer (b in Figure 1). The photogenerated species then reverts back
spontaneously to the original state with first-order kinetics. In addition to the cleaving [C–O] bond, also
a [C–N] bond between the nitrogen atom and the adjacent benzyl group holds the oxazine ring together.
The relocation of the nitro group from the meta- to the ortho-position, relative to the benzyl group, is
expected to allow the irreversible photochemical opening of the oxazine heterocycle. Indeed, ortho-
nitrobenzyl derivatives are common photocleavable protecting groups that convert irreversibly into the
corresponding ortho-nitrosobenzaldehydes.67 On the basis of these considerations, we designed
compound 1 and its BODIPY conjugates 2 and 3 (Figure 2). The [C–N] bond of their ortho-nitrobenzyl
group is expected to cleave, upon illumination at an appropriate Ac, and generate corresponding 3H-
indoles 4–6, after the elimination of ortho-nitrosobenzaldehyde 7.
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Compound 1 was synthesized in three steps from 2-methyl-3-methoxynitrobenzene (Figure 3).
Specifically, bromination of the methyl group to generate 8, cleavage of the methoxy group to produce
9 and condensation with 4 afford the target oxazine in an overall yield of 8%. The structural identities
of the intermediates and final compound were confirmed by electrospray ionization mass spectrometry
(ESIMS), 1H and 13C nuclear magnetic resonance (NMR) spectroscopies. In addition, single crystals of
1, suitable for X-ray diffraction analysis (Table S1), could be obtained from an acetonitrile solution of
the compound. The resulting structure (Figure 4) clearly reveals the oxazine core of this molecule
together with the nitro substituent in the ortho-position, relative to the benzyl group, necessary to induce
the designed photochemical transformation.
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The covalent attachment of BODIPY chromophores to the methyl group on the chiral center of 1 can be
performed in a single additional synthetic step (Figure 3). In particular, condensation of 1 with 10 or 11
generates 2 or 3 in yields of 15 or 17% respectively. Once again, the structural identities of the resulting
1
13
BODIPY–oxazine dyads were confirmed by ESIMS as well as H and C NMR spectroscopies. In
1
addition, the H NMR spectra of both compounds reveal an AX system for the pair of protons on the
ethyne bridge, separating the two functional components of each dyad, with coupling constants of 16 and
17 Hz respectively. These relatively large values demonstrate that the associated [C=C] bond adopts a
trans configuration in both compounds.
Photochemical and Photophysical Properties: Illumination of an acetonitrile solution of 1 at a Ac of
350 nm results in the cleavage of the ortho-nitrobenzyl group with the formation of 4 and 7 (Figure 2),
in agreement with the logic behind the design of this particular oxazine. The progress of the
photochemical transformation can be monitored conveniently by high-performance liquid
chromatography (HPLC) and absorption spectroscopy. In particular, chromatograms (Figure 5),
recorded at regular intervals of time during photolysis, reveal the gradual decrease of a peak with a
retention time of 13 min and the concomitant growth of a peak with a retention time of 4 min. These
peaks correspond to 1 and 4 respectively and the temporal dependence of their intensities indicates the
activation quantum yield for the transformation of the reactant into the product to be 0.043 (A in Table
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1). Similarly, spectra (Figure 6), recorded during photolysis, show a decrease in absorbance for the bands
of 1 at 236 and 277 nm together with the concomitant growth of the characteristic absorption of an ortho-
nitrosobenzaldehyde chromophore at ca. 400 nm, consistently with the photoinduced formation of 7.
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The absorption spectrum (a in Figure 7) of model compound 12 shows a band at 521 nm (Ab in Table
1) for its BODIPY chromophore. Covalent attachment of this chromophoric unit to the photocleavable
oxazine has a negligible influence on its absorption in the case of 2 (b in Figure 7), while it shifts the
band to 548 nm for 3 (d in Figure 7). Presumably, steric hindrance prevents the ethyne bridge from
adopting a co-planar arrangement with the adjacent chromophoric fragment in the ground state of 2. Its
geometry, however, appears to relax significantly in the excited state and, in fact, the Stokes shift of 2 is
25 nm, while those of 12 and 3 are only 13 and 14 nm respectively. Specifically, the BODIPY emission
appears at 534, 555 and 562 nm for 12, 2 and 3 respectively (Em in Table 1 and Figure S1). Nonetheless,
the fluorescence quantum yields of 2 and 3 are only 0.06 and 0.07 respectively (F in Table 1), while that
of 12 is 0.77. Thus, the proximal oxazine component tends to encourage the nonradiative deactivation
of the BODIPY fluorophore in both dyads. Presumably, electron transfer either from the nitrogen atom
of the 2H,3H-indole heterocycle to the excited BODIPY or from the excited chromophore to the nitro
group on the adjacent phenoxy fragment is responsible for this behavior. Indeed, the redox potentials of
1 and 12 suggest that both electron-transfer processes are exergonic with free-energy changes of –0.18
and –0.30 eV respectively.68
In analogy to the photochemical behavior of 1, illumination of 2 and 3 at a Ac of 350 nm cleaves their
ortho-nitrobenzyl group to generate 5 and 6 respectively together with 7. Consistently, the absorption
spectra (a and b in Figure 8) of acetonitrile solutions of the two dyads reveal a decrease in the absorbance
of their BODIPY fragments at 520 and 548 nm respectively during photolysis. Furthermore, a band for
the ortho-nitrosobenzaldehyde chromophore of 7 develops between 300 and 400 nm in both instances
and an absorption for the BODIPY fragments of 5 and 6 appears at 534 and 588 nm respectively. In fact,
the latter absorptions are identical to those observed for the isolated compounds (c and e in Figure 7).
Thus, the photoinduced transformation of 2 into 5 and of 3 into 6 shifts bathochromically the BODIPY
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absorption by 14 and 40 nm respectively. Such a shift offers the opportunity to excite exclusively the
BODIPY chromophore of the photochemical product by illuminating the sample at a Ex positioned
within the developing absorption. Nonetheless, excitation of 5 produces only a weak emission at 620
nm with a fluorescence quantum yield lower than 0.01 (Table 1 and Figure S1). By contrast, 6 emits at
602 nm with a fluorescence quantum yield of 0.50. This particular value in conjunction with a molar
absorption coefficient of 134 mM–1 cm–1 ( in Table 1) at Ab corresponds to a brightness of 67 mM–1
cm–1 ( × F in Table 1). Such a combination of photophysical properties translates into the activation
of intense fluorescence with the photochemical conversion of 3 into 6. In fact, emission spectra (c in
Figure 8), recorded with a Ex of 580 nm at regular intervals of time during photolysis at a Ac of 350
nm, show the appearance and growth of the characteristic band of the photochemical product. Therefore,
this particular choice of chromophoric building blocks offers the opportunity to activate the emission of
a bright fluorophore on the basis of the interplay between activating and exciting sources in conjunction
with a mechanism that enables the selective excitation of the product, satisfying all the requirements
behind the rationale for the design of these molecular switches.
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The chromophoric fragments of 3 are relatively hydrophobic and, as a result, this dyad is essentially
insoluble in aqueous solutions. However, it dissolves readily in phosphate buffer saline (PBS, pH = 7.4)
under the assistance of 13 (Figure 9). This particular macromolecular construct has a number average
molecular weight (Mn) of 30 kDa and incorporates hydrophilic oligo(ethylene glycol) and hydrophobic
decyl side chains in a ratio of 3:1 along a common poly(methacrylate) backbone.69 In aqueous
environment at concentrations greater than 5.7 g mL–1, multiple copies of this polymer assemble
spontaneously into particles with an average hydrodynamic diameter of 31 nm.70 The resulting
supramolecular hosts can capture molecular guests in their hydrophobic interior.65d,f,g,j,71 Indeed, the
absorption spectrum of nanoparticles of 13, loaded with 3, reveals the characteristic BODIPY absorption
at 550 nm (a in Figure 9). Upon illumination at a Ac of 350 nm, 3 switches to 6 within the interior of
the nanoparticles and the characteristic absorption band of the product appears at 596 nm (b in Figure 9).
Irradiation at a Ex positioned within this band excites 6 selectively with the concomitant appearance of
an emission at 607 nm in the corresponding spectrum (c in Figure 9). Thus, the switchable dyad retains
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its photochemical and photophysical properties essentially unaltered, despite encapsulation within the
supramolecular containers.
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Fluorescence Imaging: The emission bands (c and d in Figure S1) of 3 and 6 are sufficiently resolved
to permit the monitoring of the photochemical conversion of one compound into the other with two
independent detection channels. Specifically, illumination of a rectangular portion of a poly(butyl
methacrylate) (PBMA) film, doped with the dyad, at a Ac of 405 nm converts 3 into 6 exclusively within
the irradiated area. The subsequent acquisition of an image (a in Figure 10) with a Ex of 514 nm, where
3 absorbs, and a detection window of 530–650 nm, where 3 emits, shows significant fluorescence only
outside the activated area. By contrast, an image (b in Figure 10) of the very same sample collected with
a Ex of 594 nm, where 6 absorbs, and a detection window of 610–700 nm, where 6 emits, shows
significant fluorescence exclusively within the activated area. Furthermore, exactly the same fluorescent
square can be observed in images (Figure S4) collected with these excitation and detection conditions
several minutes after activation. These results confirm the photoinduced conversion of 3 into 6 and
demonstrate that the photochemical product cannot diffuse within the rigid polymer matrix.
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The fluorescence activation associated with the transformation of 3 into 6 can be exploited to monitor
the translocation of nanoparticles of 13 across soft matrices. In particular, these supramolecular
nanocarriers can be loaded with 3 and dispersed in an alginate hydrogel.72–75 An image of the resulting
sample (a in Figure 11), recorded with a Ex of 594 nm and a detection window of 610–700 nm, does not
reveal any significant fluorescence. Illumination of a rectangular area at the center of the imaging field
at Ac of 405 nm converts 3 into 6 and causes the appearance of intense emission in this particular portion
of the sample (b in Figure 11). The nanocarriers with activated cargo, however, can gradually diffuse
across the soft hydrogel matrix.76 As a result, an image (c in Figure 11), recorded 630 s after activation,
clearly reveals fluorescence outside the activated area because of the translocation of the emissive species
across the sample. In fact, the sequential acquisition of images after activation offers the opportunity to
monitor this dynamic process in real time (Video S1). The very same process can also be tracked with
conventional FRAP schemes (Figure S5 and Video S2) and nanoparticles of 13 containing 12. However,
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the illumination time required to bleach 12 is significantly longer than that sufficient to activate 3, under
otherwise identical conditions. Specifically, illumination at 405 nm for only 10 s converts a sufficient
amount of 3 into 6 to permit the detection of fluorescence against a dark background, while it has
negligible influence on the images recorded with 12 (a and b in Figure S5). The latter compound requires
an illumination time of, at least, 60 s to enable the effective suppression of fluorescence and the
subsequent tracking of the nanoparticle diffusion (a–c in Figure S5).
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The relatively mild illumination conditions sufficient to convert 3 into 6, in conjunction with the
photophysical properties of both compounds, are particularly attractive to probe dynamics within living
organisms. In particular, the fate of nanoparticles of 13, containing 3 and microinjected within a
Drosophila Melanogaster embryo during cellularization,77 can be monitored conveniently on the basis
of the photoactivation of their fluorescent cargo. Overlaid fluorescence and transmittance images (a in
Figure 12) of a labeled embryo, positioned next to an unlabeled counterpart, clearly reveal the intense
emission of the fluorescent nanocarriers exclusively in the former. In this particular experiment, the
imaging field was illuminated at a Ex of 514 nm, where 3 absorbs, and fluorescence was detected within
530–650 nm, where 3 emits. Irradiation of a relatively small region (arrow in b of Figure 12) within the
labeled embryo at 405 nm converts 3 into 6 and causes a pronounced decrease in emission, detected
under the same conditions, exclusively in the activated area. Instead, the corresponding images (c and d
in Figure 12) recorded with a Ex of 594 nm, where 6 absorbs, and a detection window of 610–700 nm,
where 6 emits, show the appearance of intense fluorescence in the activated area of the labeled embryo,
in agreement with the photoinduced generation of this particular compound. Furthermore, the subsequent
acquisition of frames over the course of 2250 s does not reveal any spatial redistribution of the activated
nanoparticles (Video S3). The final image (Figure S7), acquired after this relatively long period of time,
is essentially identical to that recorded immediately after activation (d in Figure 12). These observations
demonstrate that the fluorescent nanocarriers cannot diffuse within the developing embryos, in contrast
to their ability to translocate within the microporous alginate hydrogel (a–c in Figure 11 and Video S1).
Presumably, the nanoparticles remain immobilized within the many hydrophobic domains present in the
growing embryo.77 Additionally, comparison of the labeled and unlabeled embryos over this period of
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time does not show any significant difference in their development (Video S3). Both undergo
gastrulation and continue to grow even after photoactivation, indicating that the illumination conditions
and photochemical products do not have any noticeable cytotoxic effects and influence on the subsequent
steps of embryogenesis.
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Conclusions
A photocleavable 2H,4H-benzo[1,3]oxazine can be assembled in three synthetic steps from a commercial
precursor and condensed to formylated BODIPYs in a further synthetic step. Illumination of the resulting
BODIPY–oxazine dyads at an appropriate Ac cleaves the oxazine ring irreversibly and brings the
corresponding BODIPY chromophore in conjugation with a 3H-indole heterocycle. This structural
transformation shifts bathochromically the BODIPY absorption by up to 40 nm and allows the selective
excitation of the photochemical product at an appropriate Ex with concomitant emission. The molar
absorption coefficients of the photogenerated species are in excess of 105 M–1 cm–1 with fluorescence
quantum yields approaching 0.5. Furthermore, the hydrophobic character of these photoresponsive
molecules encourages their encapsulation within the interior of self-assembling nanoparticles of
amphiphilic polymers. In aqueous environments, the resulting supramolecular hosts retain their
molecular guests and preserve their photochemical and photophysical properties. Indeed, the
fluorescence of the entrapped BODIPY–oxazine dyads can be, once again, activated irreversibly under
the influence of illuminating sources operating at Ac and Ex. In turn, the translocation of supramolecular
nanocarriers with activated cargo across an alginate hydrogel matrix can be probed in real time with the
sequential acquisition of fluorescence images. Additionally, the illumination conditions required to
activate the fluorescence of these photoresponsive probes are significantly milder than those needed to
bleach a conventional fluorophore and implement a FRAP scheme. In fact, the fluorescent cargo of
nanocarriers, microinjected within developing Drosophila Melanogaster embryos, can be photoactivated
and then monitored for prolonged periods of time without cytotoxic effects and any noticeable influence
on embryogenesis. Thus, this particular structural design to photoactivate BODIPY fluorescence with
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large brightness and infinite contrast can evolve into a general experimental protocol to track the
dynamics of supramolecular nanocarriers and their cargo across soft matrices in real time.
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Experimental Section
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Materials and Methods: Chemicals were purchased from commercial sources and used as received.
CH2Cl2 and MeCN were distilled over CaH2. THF was distilled over Na and benzophenone. H2O (18.2
M cm) was purified with a Barnstead International NANOpure DIamond Analytical system. Decyl
methacrylate,
poly(ethylene
glycol)
methyl
ether
methacrylate,
1,4-bis(2-bromo-2-
methylpropionato)benzene, 10 and 12 were prepared according to literature procedures.56,65e,71a,78
ESIMS was performed with a Bruker micrOTO-Q II spectrometer. NMR spectra were recorded with a
Bruker Avance 400 spectrometer. HPLC traces were acquired with a Varian ProStar system, equipped
with a ProStar 330 photodiode array detector (250 nm), using an Agilent Microsorb 100-5 BDS column
(4.6  250 mm) and a MeCN/H2O (60:40, v/v) eluent (1.0 mL min–1). GPC analyses were performed
with the same chromatographic apparatus, using a Phenogel 5-μm 10E4A column (7.8 × 300 mm) and a
THF eluent (1.0 mL min–1). Monodisperse polystyrene standards (2,700–200,000) were employed to
determine the Mn and PDI of the polymers from the GPC traces. DLS measurements were performed
with a Malvern ZEN1600 apparatus. Absorption spectra were recorded with a Varian Cary 100 Bio
spectrometer in quartz cells with a path length of 1.0 cm. Emission spectra were recorded with a Varian
Cary Eclipse spectrometer in aerated solutions. The values of F were determined against cresyl violet
and rhodamine 6G standards, following a literature protocol.27 Those of A were determined with a
potassium ferrioxalate actinometer, according to an established procedure.79 Solutions were irradiated
at 350 nm (4.88 mW cm–2) with a Luzchem Research LZC-4V photoreactor. Voltammograms were
recorded with a CH Instruments 610A electrochemical analyzer in MeCN under Ar, using a three-
electrode cell. The reference was a Ag/Ag+ electrode (1 mM AgNO3 in MeCN). The counter and
working were a platinum wire and a glassy-carbon electrode respectively. The supporting electrolyte
was Bu4NPF6 (0.1 M). The scan rate and pulse amplitude were 50 mV s–1 and 50 mV respectively.
Fluorescence images were recorded with a Leica SP5 confocal laser-scanning microscope.
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1. A solution of 4 (549 mg, 3.4 mmol) and 9 (800 mg, 3.4 mmol) in MeCN (10 mL) was heated for 12
hours under reflux and Ar. After cooling down to ambient temperature, the solvent was distilled off
under reduced pressure. The residue was dissolved in CH2Cl2 (50 mL) and washed with H2O (20 mL).
The organic layer was dried over Na2SO4 and the solvent was distilled off under reduced pressure. The
residue was purified by column chromatography [SiO2, hexanes/AcOEt (95:5, v/v)] to give 1 (150 mg,
14%) as a yellow solid. ESIMS: m/z = 311.1401 [M + H]+ (m/z calcd. for C18H19N2O3 = 311.1390); 1H
NMR (CD3CN): δ = 1.17 (3H, s), 1.53 (3H, s), 1.60 (3H, s), 4.86 (1H, d, 16 Hz), 5.04 (1H, d, 16 Hz),
6.66 (1H, d, 8 Hz), 6.79–6.83 (1H, m), 6.98–7.05 (2H, m), 7.17 (1H, d, 8 Hz ), 7.28 (1H, t, 8 Hz), 7.67
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(1H, d, 8 Hz); C NMR (CDCl3): δ = 16.1, 18.8, 25.8, 38.9, 47.8, 100.6, 108.3, 116.0, 117.1, 120.3,
122.4, 123.9, 127.6, 137.9, 146.8, 147.7, 154.7.
2. A solution of 1 (100 mg, 0.3 mmol), 10 (113 mg, 0.3 mmol) and trifluoroacetic acid (TFA, 0.1 mL,
1.3 mmol) in EtOH (10 mL) was heated for 24 hours under reflux. After cooling down to ambient
temperature, the solvent was distilled off under reduced pressure and the residue was dissolved in CH2Cl2
(5 mL). The addition of hexane (50 mL) caused the precipitation of a solid, which was filtered off,
dissolved in CH2Cl2 (20 mL) and washed with H2O (20 mL). The organic phase was dried over Na2SO4
and the solvent was distilled off under reduced pressure. The residue was purified by column
chromatography [SiO2, hexanes/EtOAc (90:10, v/v)] to give 2 (30 mg, 15%) as a red solid. ESIMS: m/z
= 645.2837 [M + H]+ (m/z calcd. for C38H36BF2N4O3 = 645.2849); 1H NMR (CD3CN): δ = 1.18 (3H, s),
1.29 (3H, s), 1.38 (3H, s), 1.51 (3H, s), 2.50 (6H, s), 4.77 (1H, d, 18 Hz), 5.01 (1H, d, 18 Hz), 5.98 (1H,
d, 16 Hz), 6.14 (1H, s), 6.65 (1H, 16 Hz), 6.71 (1H, d, 8 Hz), 6.83 (1H, t, 7 Hz), 7.07 (1H, t, 7 Hz), 7.11–
7.22 (2H, m), 7.28 (1H, t, 8 Hz), 7.35 (2H, bs), 7.43–7.61 (3H, m), 7.65 (1H, d, 8Hz); 13C NMR (CDCl3):
δ = 157.0, 155.0, 153.4, 147.4, 146.8, 144.2, 142.0, 138.5, 138.0, 134.9, 129.3, 129.2, 129.1, 128.0,
127.6, 127.5, 127.2, 125.5, 123.4, 122.3, 120.4, 117.3, 117.1, 108.7, 101.9, 49.5, 39.7, 34.7, 31.6, 26.5,
25.3, 22.7, 18.3, 14.7, 14.5, 14.1, 13.7, 12.8.
3. A solution of 1 (97 mg, 0.3 mmol), 11 (100 mg, 0.3 mmol) and TFA (0.1 mL, 1.3 mmol) in EtOH (10
mL) was heated for 24 hours under reflux. After cooling down to ambient temperature, the solvent was
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distilled off under reduced pressure. The residue was dissolved in CH2Cl2 (30 mL) and washed with a
saturated aqueous solution of NaHCO3 (2  20 mL). The organic layer was dried over Na2SO4 and the
solvent was distilled off under reduced pressure. The residue was purified by column chromatography
[SiO2, hexanes/AcOEt (95:5  90:10, v/v) to give 3 (30 mg, 17%) as a dark violet solid. ESIMS: m/z =
687.3338 [M + H]+ (m/z calcd. for C41H42BF2N4O3 = 687.3318); 1H NMR (CD3CN): δ = 0.97 (6H, q, 8
Hz), 1.24 (3H, s), 1.28 (3H, s), 1.31 (3H, s), 1.59 (3H, s), 2.35 (2H, q, 8 Hz), 2.43–2.52 (5H, m), 4.88
(1H, d, 19 Hz), 5.08 (1H, d, 19 Hz), 6.50 (1H, d, 17 Hz), 6.77 (1H, d, 8 Hz), 6.87 (1H, td, 1 and 8 Hz),
7.10 (1H, td, 1 and 8 Hz), 7.15–7.26 (2H, m), 7.26–7.32 (1H, m), 7.31–7.34 (3H, m), 7.51–7.57 (3H, m),
7.69 (1H, dd, 1 and 8 Hz); 13C NMR (CD3CN): δ = 158.3, 154.6, 147.7, 147.0, 145.2, 141.4, 140.9, 138.1,
137.9, 135.1, 135.0, 132.7, 132.5, 131.0, 129.8, 129.2, 129.1, 128.2, 127.9, 127.6, 126.9, 123.4, 123.2,
122.4, 122.3, 120.5, 118.0, 109.0, 102.1, 49.8, 39.5, 29.4, 25.6, 17.7, 17.5, 16.5, 13.6, 12.2, 11.2, 10.6.
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5. A solution of 4 (159 mg, 1.0 mmol), 10 (352 mg, 1.0 mmol) and TFA (0.1 mL, 1.3 mmol) in EtOH
(20 mL) was heated for 24 hours under reflux. After cooling down to ambient temperature, the solvent
was distilled off under reduced pressure and the residue was dissolved in CH2Cl2 (5 mL). The addition
of hexane (50 mL) caused the precipitation of a solid, which was filtered off, dissolved in CH2Cl2 (20
mL) and washed with H2O (20 mL). The organic phase was dried over Na2SO4 and the solvent was
distilled off under reduced pressure. The residue was purified by column chromatography [SiO2,
hexanes/EtOAc (90:10, v/v)] to give 5 (104 mg, 21%) as a black solid. ESIMS: m/z = 494.2601 [M +
H]+ (calcd. for C31H31BF2N3 = 494.2579); 1H NMR (CD3CN) δ = 1.39 (6H, s), 1.42 (3H, s), 1.55 (3H,
s), 2.55 (3H, s), 2.73 (3H, s), 6.20 (1H, s), 6.68 (1H, d, 17 Hz), 7.23 (1H, t, 11 Hz), 7.32 (1H, t, 8 Hz),
7.36–7.46 (3H, m), 7.51 (1H, d, 8 Hz), 7.56–7.62 (3H, m), 7.64 (1H, d, 17 Hz); 13C NMR (CDCl3): δ =
183.7, 157.7, 154.6, 154.0, 146.3, 144.8, 142.1, 139.5, 134.9, 132.4, 130.9, 129.4, 129.3, 129.2, 128.0,
127.8, 125.3, 122.3, 121.1, 120.5, 120.4, 52.5, 31.6, 24.0, 22.7, 14.8, 14.6, 14.2, 14.1, 13.0.
6. A solution of 4 (50 mg, 0.3 mmol), 11 (100 mg, 0.3 mmol) and TFA (0.1 mL, 1.3 mmol) in MeCN
(20 mL) was heated for 24 hours under reflux. After cooling down to ambient temperature, the solvent
was distilled off under reduced pressure. The residue was dissolved in CH2Cl2 (20 mL) and washed with
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a saturated aqueous solution of NaHCO3 (2  20 mL). The organic layer was dried over Na2SO4 and the
solvent was distilled off under reduced pressure. The residue was purified by column chromatography
[SiO2, hexanes/AcOEt (80:20  30:70, v/v) to give 6 (50 mg, 36%) as a dark purple solid. ESIMS: m/z
= 536.3050 [M + H]+ (m/z calcd. for C34H37BF2N3 = 536.3049); 1H NMR (CD3CN): δ = 1.04 (3H, t, 8
Hz), 1.16 (3H, 8 Hz), 1.31 (3H, s), 1.37 (3H, s), 1.78 (6H, s) 2.37 (2H, q, 8 Hz), 2.58–2.80 (5H, m), 7.29–
7.36 (2H, m), 7.36–7.48 (3H, m), 7.51–7.62 (4H, m), 7.85 (1H, d, 6 Hz), 8.50 (1H, d, 17 Hz); 13C NMR
(CDCl3): δ = 183.8, 182.7, 162.2, 152.2, 144.4, 143.1, 140.4, 139.9, 137.3, 136.4, 134.7, 129.5, 129.5,
129.0, 128.0, 127.9, 124.9, 121.7, 119.4, 118.0, 100.0, 52.1, 25.3, 18.4, 17.2, 14.1, 13.9, 12.2, 10.9.
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8. A solution of 2-methyl-3-methoxynitrobenzene (1.0 g, 6.0 mmol), azobis(i-butyronitrile) (AIBN, 98
mg, 0.1 mmol) and N-bromosuccinimide (NBS, 1.3 g, 7.3 mmol) in CCl4 (15 mL) was heated for 14
hours under reflux and Ar. After cooling down to ambient temperature, the reaction mixture was diluted
with CH2Cl2 (30 mL) and was washed with H2O (20 mL). The organic layer was dried over Na2SO4 and
the solvent was distilled off under reduced pressure to give 8 (1.2 g, 82%) as a white solid. ESIMS: m/z
= 245.9760 [M + H]+ (m/z calcd. for C8H9BrNO3 = 245.9760); 1H NMR (CDCl3): δ = 4.00 (3H, s), 4.84
(2H, s), 7.18 (1H, bs), 7.45 (1H, d, 8Hz), 7.55 (1H, d, 8 Hz); 13C NMR (CDCl3): δ = 22.0, 57.4, 115.6,
117.1, 121.6, 130.0, 149.8, 158.5.
9. A solution of BBr3 in CH2Cl2 (1 M, 12.7 mL) was added dropwise to a solution of 8 (1.2 g, 4.9 mmol)
in CH2Cl2 (10 mL) maintained at –78 °C under inert atmosphere. The reaction mixture was allowed to
warm up to ambient temperature, stirred for a further 12 hours, poured into H2O (10 mL) and extracted
with CH2Cl2 (30 mL). The organic layer was dried over Na2SO4 and the solvent was distilled off under
reduced pressure. The residue was dissolved in AcOEt (10 mL) and filtered through a silica plug, which
was washed thoroughly with AcOEt. The solvent was distilled off under reduced pressure to give 9 (800
mg, 71%) as a brown solid. ESIMS: m/z = 231.9604 [M + H]+ (m/z calcd. for C7H7BrNO3 = 231.9604);
1H NMR (CD3CN): δ = 4.78 (2H, s), 7.23 (1H, d, 8 Hz), 7.39 (1H, t, 8 Hz), 7.49 (1H, d, 8 Hz); 13C NMR
(CD3OD): δ 21.7, 115.6, 119.3, 120.0, 129.7, 150.1, 157.2.
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11. A degassed solution of 2,3-dichloro-5,6-dicyano-1,4-benzoquinone (DDQ, 474 mg, 2.1 mmol) in
tetrahydrofuran (10 mL) was added dropwise to a solution of 12 (200 mg, 0.5 mmol) in tetrahydrofuran
(15 mL) and H2O (0.15 mL) maintained at 0 °C under Ar. The mixture was allowed to warm up to
ambient temperature, stirred for a further 12 hours under these conditions, diluted with AcOEt (30 mL)
and washed with H2O (2  20 mL). The organic layer was dried over Na2SO4 and the solvent was distilled
off under reduced pressure. The residue was purified by column chromatography [SiO2, hexanes/AcOEt
(95:5  80:20, v/v) to give 11 (96 mg, 47%) as a brown solid. ESIMS: m/z = 417.1920 [M + Na]+ (m/z
calcd. for C23H25BF2N2ONa = 417.1926); 1H NMR (CDCl3,): δ = 1.03 (6H, t, 8 Hz), 1.28 (3H, s), 1.37
(3H, s), 2.36 (2H, q, 8 Hz), 2.65 (3H, s), 2.70 (2H, q, 8 Hz), 7.30 (2H, m), 7.54 (3H, m), 10.41 (1H, s);
13C NMR (CDCl3):  = 10.5, 12.3, 13.7, 14.1, 14.4, 17.1, 17.6, 127.8, 129.5, 129.5, 132.2, 134.52, 134.6,
135.7, 137.7, 139.8, 142.1, 143.9, 166.1, 186.
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13. A mixture of decyl methacrylate (1.70 g, 75 mmol), poly(ethylene glycol) methyl ether methacrylate
(5.17 g, 2.5 mmol), 1,4-bis(2-bromo-2-methylpropionato)benzene (8 mg, 0.02 mmol), FeCl3 ·6H2O (3

mg, 0.01 mmol), Ph3P (3 mg, 0.01 mmol), AIBN (8.2 mg, 0.05 mmol) in degassed THF (30 mL) was
heated at 75 °C for 48 hours in a sealed tube. After cooling down to ambient temperature, the solvent
was distilled off under reduced pressure and the residue was purified by column chromatography [SiO2,
CH3Cl/MeOH (95:5, v/v)] to give 13 (513 mg, 8%) as a white solid. GPC: Mn = 30 kDa, PDI = 1.56; 1H
NMR (CDCl3,): δ = 0.81–1.16 (25H, m), 1.20–1.45 (48H, m), 3.38 (3H, s), 3.54–3.78 (146H, m), 3.86–
4.01 (6H, m), 4.04–4.20 (2H, m).
Crystallographic Analysis. Single crystals of 1 were obtained after maintaining an acetonitrile solution
of the compound at ambient temperature in a sealed tube. The data crystal was glued onto the end of a
thin glass fiber. X-Ray intensity data were measured with a Bruker SMART APEX2 CCD-based
diffractometer, using Mo K radiation ( = 0.71073 Å).80 The raw data frames were integrated with the
SAINT+ program by using a narrow-frame integration algorithm. Corrections for Lorentz and
polarization effects were also applied with SAINT+. An empirical absorption correction based on the
multiple measurement of equivalent reflections was applied using the program SADABS. The structure
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was solved by a combination of direct methods and difference Fourier syntheses and refined by full-
matrix least-squares on F2 with the SHELXTL software package.81 Hydrogen atoms were placed in
geometrically idealized positions and included as standard riding atoms during the least-squares
refinements. Crystal data, data collection parameters and results of the analyses are listed in Table S1.
Compound 1 crystallized in the monoclinic crystal system. The systematic absences in the intensity data
identified the unique space group P21/n.
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Polymer Nanoparticles. Doped polymer nanoparticles were prepared by combining CHCl3 solutions of
3 (0.1 mg mL–1, 50 μL) and 13 (2.5 mg mL–1, 200 μL) and heating the resulting mixture at 40 °C in an
open vial. After the evaporation of the solvent, the residue was purged with air, dispersed in PBS (1 mL,
pH =7.4), sonicated for 5 min and flashed through a syringe filter with a pore size of 0.2 m. The filtrate
was used for the imaging and spectroscopic measurements without further purification.
Polymer Films. Doped polymer films were prepared by spin coating CH2Cl2 solutions of PBMA (MW =
337 × 103, 20 mg mL–1) and 3 (50 g mL–1) with a Chemat Technologies KW-4A spin coater at 1200
rpm for 60 s on glass coverslips.
Alginate Hydrogels. Doped alginate hydrogels were prepared by mixing a solution of sodium alginate
(20 mg mL–1) in H2O (200 L) with a solution of nanoparticles of 13 (500 μg mL–1), loaded with either
3 or 12 (5 μg mL–1), in H2O (200 L) and then adding a drop of the mixture to a solution of CaCl2 (0.24
M) in H2O (1 mL). The resulting doped alginate bead was rinsed with H2O (1 mL) and transferred on a
glass slide.
Drosophila Melanogaster Embryos. Drosophila Melanogaster embryos were prepared for the imaging
experiments adapting a literature protocol.82 Specifically, embryos were collected during cellularization,
aligned in pairs on glass slides and maintained in a desiccator, together with CaCl2, at ambient
temperature for 8 minutes. The slides were covered with a mixture (7:1, v/v) of Series 700 (Sigma
H8898) and Series 27 (Sigma H8773) halocarbon oils and transferred individually on the stage of a Leica
DMIL LED inverted microscope. A solution of nanoparticles of 13 (5 mg mL–1), containing 3 (50 g
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mL–1), in Dulbecco's PBS (pH = 7.2–7.6) was microinjected in only one embryo of each pair of embryos
with a MN151 Narishige micromanipulator. After 1 hour, the slides were mounted on the stage of a
Leica SP5 confocal laser-scanning microscope and imaged for 40 minutes.
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Acknowledgment. J. G.-A. is grateful for a Beatriu de Pinós postdoctoral grant from the Generalitat de
Catalunya (Spain, 2011 BP-A-00270). F. M. R. acknowledges the National Science Foundation
(CAREER Award CHE-0237578, CHE-0749840 and CHE-1049860) for financial support.
Supporting Information Available. Crystallographic data for 1; emission spectra of 2, 3, 5, 6 and 12;
differential-pulse voltammograms of 1 and 12; dependence of the emission intensity of 12 on the
concentration of 13; fluorescence images of 3 in PBMA; fluorescence images of nanoparticles of 13,
doped with 3 or 12, in alginate hydrogels; fluorescence images of nanoparticles of 13, doped with 3, in
Drosophila Melanogaster embryos. This material is available free of charge via the Internet at
http://pubs.acs.org.
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(1) (Kavarnos, G. J. Fundamentals of Photoinduced Electron Transfer; VCH: New York, 1993).
The oxidation potential (EOx) of the donor and the reduction potential (ERed) of the acceptor in both
instances were determined from the differential pulse voltammograms (Figure S2) of 1 and 12.
The energy change (E00) for the S0  S1 transition of the BODIPY chromophore was estimated
to be +2.35 eV from the absorption and emission spectra of 12. The donor–acceptor distance (d)
was approximated to 0.5 nm in both instances.
e2
G° = e EOx – e ERed – E00 –
(1)
4  0 r d
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69 Macromolecule 13 was synthesized from the corresponding methacrylate monomers (ref. 71a),
adapting a literature protocol (ref. 78) for atom-transfer radical polymerization. Gel permeation
chromatography (GPC) and 1H NMR spectroscopy were employed to determine Mn and the ratio
between the hydrophobic and hydrophilic chains respectively.
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70 The critical concentration of 13, required to encourage the assembly of nanoparticles, was
estimated from the dependence of the emission intensity of 12 on the polymer concentration (Figure
S3), adapting a literature protocol (ref. 71b). Dynamic light scattering (DLS) measurements were
employed to determine the average hydrodynamic diameter of the nanoparticles.
71 (a) Yildiz, I.; Impellizzeri, S.; Deniz, E.; McCaughan, B.; Callan, J. F.; Raymo, F. M. J. Am. Chem.
Soc. 2011, 133, 871–879. (b) Swaminathan, S.; Fowley, C.; McCaughan, B.; Cusido, J.; Callan, J.
F.; Raymo, F. M. J. Am. Chem. Soc. 2014, 136, 7907–7913.
72 Solutes of various dimensions and shapes, including relatively large biomolecules, are known to
diffuse through alginate hydrogels (ref. 73–75). These materials hold large amounts of water
within intricate networks of polyanionic alginate chains bridged by divalent calcium cations.
73 (a) Quignard, F.; Valentinw, R.; Di Renzo, F. New J. Chem. 2008, 32, 1300–1310. (b) Quignard,
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76 In principle, the photoactivatable guests can leak out of their supramolecular hosts (Jiwpanich, S.;
Ryu, J.-H.; Bickerton, S.; Thayumanavan, S. J. Am. Chem. Soc. 2010, 132, 10683–10685) and
diffuse across the sample. The hydrophobic character of 3 and 6, however, translates into aqueous
solubilities in the subpicomolar regime. At such low concentration levels, the fluorescence of any
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activated guest, which might be present outside the nanocarriers, is undetectable under the
experimental conditions employed to acquire the images in Figure 11. As a result, the observed
intense emission can only be associated with the large fraction of fluorophores encapsulated within
the supramolecular containers.
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77 Campos-Ortega, J. A.; Hartenstein, V. The Embryonic Development of Drosophila Melanogaster;
Springer: Berlin, 1997.
78 Zhu, G.; Zhang, L.; Zhang, Z.; Zhu, J.; Tu, Y.; Cheng, Z.; Zhu, X. Macromolecules 2011, 44, 3233–
3239.
79 Scaiano, J. C. Handbook of Organic Photochemistry; CRC Press: Florida, 1989.
80 Apex2 Version 2.2-0 and SAINT+ Version 7.46A; Bruker Analytical X-ray System, Inc., Madison,
Wisconsin, USA, 2007.
81 (a) Sheldrick, G. M. SHELXTL Version 6.1; Bruker Analytical X-ray Systems, Inc., Madison,
Wisconsin, USA, 2000. (b) Sheldrick, G. M. Acta Cryst. 2008, A64, 112–122.
82 Spradling, A. C.; Rubin, G. M. Science 1982, 218, 341–347.
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Table 1. Photochemical and photophysical parameters of 1–3, 5, 6 and 12 in MeCN at 25 °C.
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9

Ab [a]
 [b]
(mM–1 cm–1)

Em [a]
F [c]
 × F [d]
(mM–1 cm–1)
A [c]
(nm)
333
520
548
534
588
521
(nm)
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1
2
2
—
—
—
5
0.043
0.002
0.002
—
84
555
562
620
602
534
0.06
0.07
<0.01
0.50
0.77
3
111
88
8
5
<1
67
66
6
134
86
—
12
—
[a] Wavelengths at the absorption (Ab) and emission (Em) maxima. [b] Molar absorption coefficient at Ab. [c] Fluorescence (F) and activation
(A) quantum yields. [d] Brightness at Ab
.
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Figure 1. Reversible opening of an oxazine ring (a) to form a zwitterionic isomer (b) under illumination
at an activating wavelength (Ac).
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Figure 2. Irreversible opening of the oxazine ring of 1–3 to form 4–6 and 7 under illumination at an
activating wavelength (Ac).
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