Photocontrol of ion permeation in lipid vesicles with (bola)amphiphilic spirooxazines

Article abstract of DOI:10.1039/c5ob01993a

Three (bola)amphiphilic spirooxazines have been synthesized and their photochromism has been characterized. The large biphotochromic structure of 2 significantly affects its conformational flexibility and the rate constants for thermal ring closure are particularly dependent on the lipid phase state. Two comprehensive ion permeation studies were performed to examine the effect of spirooxazine inclusion and isomerization on membrane permeability. In all cases, the open-ring isomers of these spirooxazines are more disruptive in bilayer membranes than their closed-ring isomers. Further, the rate of ion permeation and net release are highly dependent on the lipid bilayer phase state and the relative position of the photochromic moiety in the bilayer membrane. Moreover, the difference in potassium ion permeability under UV and visible irradiation is more pronounced than previously reported photoresponsive membrane disruptors with reversible photocontrols.

Full text of DOI:10.1039/c5ob01993a

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Page 1 of 34
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Photocontrol of Ion Permeation in Lipid Vesicles with (Bola)amphiphilic
Spirooxazines
Yamuna S. Kandasamy, Jianxin Cai, John G. Ottaviano, Kelti A. Smith, Ashley N. Williams, Jarod
Moore, Kristen M. Louis, Lindsay Selzler, Alisha Beler, Tobechi Okwuonu and R. Scott Murphy*
Department of Chemistry and Biochemistry, Research and Innovation Centre, University of Regina,
3737 Wascana Parkway, Regina, SK, S4S 0A2, Canada
scott.murphy.uregina@gmail.com
Abstract
Three (bola)amphiphilic spirooxazines have been synthesized and their photochromism has been
characterized. The large biphotochromic structure of 2 significantly affects its conformational
flexibility and the rate constants for thermal ring closure are particularly dependent on the lipid
phase state. Two comprehensive ion permeation studies were performed to examine the effect of
spirooxazine inclusion and isomerization on membrane permeability. In all cases, the openꢀring
isomers of these spirooxazines are more disruptive in bilayer membranes than their closedꢀring
isomers. Further, the rate of ion permeation and net release are highly dependent on the lipid
bilayer phase state and the relative position of the photochromic moiety in the bilayer membrane.
Moreover, the difference in potassium ion permeability under UV and visible irradiation is more
pronounced than previously reported photoresponsive membrane disruptors with reversible
photocontrols.
1

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Introduction
Biocompatible lipid vesicles that integrate photochromic molecules have shown potential in
1
ꢀ10
biomedical applications such as drug delivery and ion transport.
These photoresponsive systems
provide a high level of spatial and temporal control for onꢀdemand dosing. Further, photocontrols
based on the reversible photoisomerization of photochromic molecules could be used to regulate the
1
1, 12
delivered dosage. To date, amphiphilic photochromic molecules comprised of azobenzene
,
1
3
14
spiropyran , and dithienylethene have been used to photocontrol ion permeation in lipid vesicles
with varying degrees of success. For example, when an amphiphilic spiropyran was included into
1
3
phosphatidylcholine (PC) vesicles the rate of potassium ion permeation increased. Upon exposure
to ultraviolet (UV) light the rate of ion permeation decreased, as the closedꢀring spiro isomer was
converted to the openꢀring merocyanine isomer. However, the difference in permeability between
these two isomeric states was within the experimental error and the observed perturbation order was
not anticipated, as Hurst and coworkers admittedly desired a photochromic system that would
promote ion leakage with UV irradiation.
1
5
Relatively few spirooxazine amphiphiles have been synthesized and characterized in lipid
1
6, 17
vesicles.
In addition, their photocontrol of ion permeability in lipid vesicles has never been
examined, even though spirooxazines are well known for their greater resistance to
1
8, 19
photodegradation than the structurally related spiropyrans.
Notably, spiropyrans specifically
2
0ꢀ22
derived from the prevalent 6ꢀnitroꢀBIPS core have been shown to have poor hydrolytic stability.
Surprisingly, the preparation of a bolaamphiphilic photochromic dimer has never been reported.
Albeit, Fujiwara and coworkers have recently examined the photochromic reactivity of
23, 24
nonamphiphilic spirooxazine dimers in organic solution
, and their photocontrol of guest binding
2
5
while tethered to a polymer. Although the functionality achieved by these systems is slowly
2
6
improving , biphotochromic systems have never been examined in lipid vesicles.
2

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1
7
In an earlier report, we examined the thermal isomerization of 1 in lipid vesicles (Chart 1).
1
6
Consistent with similar spirooxazines, the rate constant for thermal ring closure (k ) increased and
T
the wavelength of maximum absorption (λmax) of the openꢀring isomer underwent a small
bathochromic shift with increasing solvent polarity. Upon inclusion of 1 in
dipalmitoylphosphatidylcholine (DPPC) vesicles, the λmax for the openꢀring isomer underwent a
hypsochromic shift relative to toluene, which suggested that the photochromic moiety of 1 is most
likely located in the less polar aliphatic region of the bilayer membrane. Building on this previous
work, we report on the synthesis of the first bolaamphiphilic biphotochromic dimer 2 and a new
analogous amphiphilic spirooxazine 3. Further, we have characterized their thermal isomerization
in organic solution and lipid vesicles of varying lamellar phase, and have examined their reversible
photocontrol of ion permeability (i.e., protons and potassium ions) through a bilayer membrane.
3

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Chart 1. Structures of the (bola)amphiphilic spirooxazines photoisomers for 1–3.
Results and discussion
Synthesis of 1–3. We have previously reported on the procedures for the synthesis of the
1
7
amphiphilic spirooxazine 1. Similar procedures were used to prepare the analogous hexyl
derivative, 3. In general, the synthesis of the bolaamphiphilic spirooxazine dimer 2 involves the
coupling of two moles of a substituted phenylboronic acid with a diiodospirooxazine dimer. We
had originally prepared a substituted alkylboronic acid to couple with the diiodospirooxazine dimer;
however, these attempts were unsuccessful due to the limited solubility of this alkylboronic acid.
Also, lower reactivity is frequently observed in SuzukiꢀMiyaura reactions of alkylboronic acids
2
7
when compared with arylboronic acids. Consequently, our synthetic methodology began with the
4

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inclusion of a cationic alkyl ammonium substituent on phenylboronic acid (Scheme 1). Initially, the
Sonogashira reaction was used to couple 1ꢀbromoꢀ4ꢀiodobenzene with pentadecꢀ14ꢀynꢀ1ꢀol to give
4
in moderate yield. The internal alkyne was then reduced by catalytic hydrogenation using
hydrogen gas over platinum(IV) oxide to produce 5 in high yield. Notably, platinum(IV) oxide was
used instead of the more common palladium on carbon because the former displays good
chemoselectivity as debromination was not observed. tertꢀButyldimethylsilyl chloride (TBSCl) and
imidazole were then used to protect the terminal hydroxyl group of 5 to give 6 in good yield. The
conversion of 6 to its corresponding boronic acid and subsequent protection using pinacol gave the
boronate ester 7 in moderate yield. Deprotection of the hydroxyl group with hydrochloric acid in
methanol produced 8 in high yield. Next, the primary alcohol was converted to the alkyl bromide
using Nꢀbromosuccinimide and triphenylphosphine to give 9 in moderate yield. The nucleophilic
substitution of the alkyl bromide was achieved in high yield to produce the cationic quaternary
ammonium substituent in 10 using triethylamine. This was followed by a convenient twoꢀstep
2
8
procedure for the deprotection of the boronate ester. The boronate ester was first converted to the
potassium trifluoroborate salt 11 in good yield using potassium hydrogen fluoride. Treatment of the
trifuoroborate salt with trimethylsilyl chloride afforded the corresponding boronic acid, which was
used without further purification in the final coupling reaction with the diiodospirooxazine dimer 13.
This photochromic dimer was synthesized using phenanthrenequinone and 2,3,3ꢀ
trimethylindolenine as starting materials for the preparation of the oxime 12 and the bisindoline unit,
respectively. The iodination of phenanthrenequinone via an electrophilic aromatic substitution
reaction produced 2ꢀiodoꢀ9,10ꢀphenanthrenedione in low yield due to the formation of a side
product, 2,6ꢀdiiodophenanthrenequinone, and the presence of unreacted starting material. The
conversion of the diketone to the oxime 12 was considerably more successful. The precursor to the
bisindoline unit was a bisindoleninium bromide salt. This salt was prepared from successive
5

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nucleophilic substitution reactions of 1,4ꢀdibromobutane with excess 2,3,3ꢀtrimethylindolenine.
Subsequently, the bisindoleninium bromide salt was converted to the bisindoline unit with base in
moderate yield. Thereafter, 13 was formed from the electrocyclic reaction of the bisindoline unit
and 12. The SuzukiꢀMiyaura reaction was then used to couple two moles of the substituted
phenylboronic acid with 13 to produce the final bolaamphiphilic spirooxazine dimer 2.
Scheme 1. Synthesis of 2.
Photochromism of 1–3. Absorption in the visible region was observed for all spirooxazines prior
to irradiation with UV light. The thermochromic behavior of analogous phenanthrylꢀbased
29ꢀ32
33
spirooxazines has been well established in homogeneous solution
, micelles , and lipid
1
6, 17
vesicles
. Further, this thermal equilibrium has been shown to favour the formation of the openꢀ
3
3
ring isomer as the polarity of the microenvironment increases. Upon UV irradiation, spirooxazine
6

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absorption increased further in the visible region, which represents the formation of the openꢀring
isomer. In addition, the photoisomerization of 1–3 was reversible in organic solution and lipid
vesicles. Unfortunately, we were unable to differentiate between the openꢀring–closedꢀring and the
openꢀring–openꢀring isomers of 2 as the two photochromic units are not electronically conjugated.
Nonetheless, following UV irradiation, thermal equilibration led to a decrease in absorbance at the
λmax in the visible region giving back the absorbance spectrum prior to UV irradiation. The rate
constants for thermal ring closure (k ) were determined from exponential fits of the absorbance
T
decays at λmax. We measured the k for 1–3 in acetonitrile and toluene to examine the effect of
T
solvent polarity and molecular structure on the thermal ring closure (Table 1). The k for 1 and 3
T
were both five fold larger in acetonitrile than toluene. Although 2 was not soluble in toluene, the
precursor 13 was examined in both solvents. Similarly, k for 13 was 18 fold larger in acetonitrile
T
1
6, 17, 29, 30, 34
than toluene. Consistent with reports on analogous phenanthrylꢀbased spirooxazines
,
these results suggest that k for 1–3 increase with increasing solvent polarity. Furthermore, k for 1
T
T
and 3 in acetonitrile and toluene were within experimental error of each other. Therefore, the
difference in the alkyl chain length has a minor effect on thermal ring closure in organic solution.
However, these rate constants in acetonitrile were two fold larger than 2. A similar reduction in kT
3
4
has been reported for a biphotochromic system when compared with its corresponding monomer.
This suggests that the dimeric structure of 2, specifically the linked indoline moieties, does
influence the rate of ring closure through a steric effect.
The k for 1–3 were also measured in two different lipid systems, namely DPPC and 1,2ꢀ
T
dioleoylꢀsnꢀglyceroꢀ3ꢀphosphocholine (DOPC), to examine the effect of the lipid bilayer phase state
on the rate of thermal ring closure. In addition, the activation energies for ring closure of 1–3 in
lipid vesicles were determined from their Arrhenius behavior. The phase state of these lipid
systems is dependent on temperature. At ambient temperature, DPPC vesicles are in the gel phase,
7

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3
5
whereas DOPC vesicles are in the fluid phase. The k for 1 and 3 were two fold larger in DOPC
T
vesicles than DPPC vesicles. This trend was anticipated given that local viscosity of DPPC bilayers
in their gel phase is higher and more restrictive to isomerization than DOPC bilayers in their fluid
phase. In addition, k is at least 50% larger for 1 when compared with 3 in both lipid systems.
T
Table 1 Thermal ring closure rate constants (k ) for spirooxazines in organic solution and lipid
T
a
vesicles at a mole ratio of 1:20 (i.e., spirooxazine/lipid)
ꢀ
1 ꢀ1
k (10 s )
T
SpOx
Acetonitrile
Toluene
DPPC
DOPC
1
2
4.16 ± 0.12 (590)
2.14 ± 0.12 (600)
0.78 ± 0.02 (591)
N/A
0.83 ± 0.01 (589)
1.91 ± 0.06 (584)
b
b
0.079 ± 0.002 (610) 0.48 ± 0.03 (606)
b
b
0.011 ± 0.002
0.10 ± 0.01
3
4.45 ± 0.48 (595)
3.14 ± 0.16 (600)
0.81 ± 0.18 (598)
0.17 ± 0.01 (597)
0.56 ± 0.07 (608)
N/A
1.03 ± 0.02 (606)
N/A
13
a
The error is the standard deviation for the mean taken from a minimum of three independent
measurements, which were recorded at, or extrapolated to 7 °C. The absorption maximum of the
b
openꢀring isomer is shown in parentheses and the error is ± 1 nm. The rate constants were
obtained from biexponential fits.
This latter result suggests that the relative positioning of the photochromic moiety of 1 and 3 in the
lipid bilayer is different owing to differences in alkyl chain length. A similar trend has been
3
6
previously observed for spiropyrans with assorted alkyl chain lengths in lipid vesicles.
Specifically, Wohl and coworkers examined the free volume distribution in a DPPC lipid bilayer.
In this work, the largest available free volume was detected near the bilayer midplane, while the
smallest free volume was found in the highly ordered region of the hydrocarbon chains. Since the
3
7
polarity in these regions of the bilayer is similar , the viscosity of the microenvironment will have a
dominant effect on k . Consequently, we postulate that the thermal ring closure of 1 is less
T
8

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hindered as it is most likely positioned closer to the less dense bilayer midplane, thereby enhancing
rates. Not surprisingly, k for 2 was significantly smaller when compared with the monomer
T
systems. Interestingly, kinetic decays with biexponential behavior were observed for 2 in DPPC
and DOPC vesicles (Figure 1). Timeꢀresolved studies on phenanthrylꢀbased spirooxazines have
previously attributed this biexponential character to the cisꢀtrans isomerization of the openꢀring
1
7, 30
isomer prior to ring closure.
Our ability to observe this shorterꢀlived component in lipid
vesicles highlights the reduced conformational flexibility of the dimer system due to the higher
viscosity within the bilayer membrane when compared with organic solution. Analogous to the
monomer systems, the longerꢀlived component of the thermal decay for 2 was nine fold larger in
DOPC vesicles than DPPC vesicles. Compared with the monomer systems, k for 2 was 75 and 50
T
fold smaller than 1 and 3 in DPPC vesicles, and 20 and 10 fold smaller in DOPC vesicles,
respectively. Also, the activation energies for the thermal ring closure of 1, 2, and 3 were 97, 119
ꢀ1
ꢀ1
and 104 kJ mol in DPPC vesicles, and 87, 74 and 84 kJ mol in DOPC vesicles, respectively, over
a temperature range of 2–21 °C (Figures S1–S6, ESI). Thus, the activation energies are higher in
the more viscous gel phase of DPPC vesicles than the fluid phase of DOPC vesicles. This
correlation with lipid phase state is consistent with a similar examination of spiropyrans in DPPC
3
6
vesicles. Again, these results demonstrate the increased rigidity of a DPPC bilayer when
compared with a DOPC bilayer, and the effect of a more viscous microenvironment on the thermal
ring closure of these (bola)amphiphilic spirooxazines. Overall, the isomerization kinetics show that
the larger molecular structure of 2 greatly affects the conformational mobility necessary for ring
closure when compared with the monomer systems.
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Figure 1. Normalized decay kinetics for the openꢀring isomer of 2 in DOPC vesicles at a mole ratio
of 1:20 (i.e., spirooxazine/lipid) monitored at 606 nm following irradiation with UV light, and
recorded at various temperatures. The inset is a series of absorption spectra recorded in the visible
region at 12 °C following a 20 s irradiation with UV light, as the openꢀring isomer reverts back to
the closedꢀring isomer.
1
4, 38, 39
Proton permeation studies in lipid vesicles. A fluorescence assay
was used to assess the
photocontrol of proton permeability for lipid vesicles incorporating spirooxazines 1–3. In this
assay, the photoresponsive lipid vesicles entrap a pHꢀsensitive, fluorescent probe, 8ꢀhydroxypyreneꢀ
1
,3,6ꢀtrisulfonic acid (HPTS). Then a transmembrane pH gradient is formed by adding base to the
external buffer. The collapse of this pH gradient is monitored over time from changes in the
fluorescence of HPTS. Subsequently, the fluorescence data is converted to a normalized extent of
permeation (N) from which the rate constant for proton permeation (k ) is determined (Figure S7,
H+
ESI). In general, the pH gradient is expected to collapse more rapidly if the incorporation of
spirooxazines as either photoisomer enhances membrane permeability. The permeation of protons
4
0
across bilayer membranes has been described by a sequential twoꢀstep mechanism. This biphasic
10

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route proceeds from a transient–pore phase to a solubility–diffusion phase. During the transient–
pore phase, which is initiated by the addition of base, a steep rise in N is often observed. This
increase in proton permeation is due to the formation of transient hydrated pores within the bilayer
produced by thermal fluctuations as the vesicles adjust to both the electrical potential and osmotic
4
1
gradients formed by the rapid change in pH. Once both gradients are closer to equilibrium a
relatively slower rate of proton permeation is observed. This second phase of proton permeation is
consistent with the solubility–diffusion model whereby hydrated protons partition into the inner
leaflet of the bilayer, diffuse through the hydrophobic core, and partition out of the outer leaflet into
4
0
the external buffer. The rate constants presented in this study were determined from this second
phase of proton permeation because permeation during this phase best represents the effect of
spirooxazine inclusion and isomerization on membrane permeability.
We have recently reported on the photocontrol of proton permeation in lipid vesicles including
thermally irreversible, amphiphilic dithienylethene monomers using the same fluorescence assay
1
4
described here. In that report, we examined pure lipid vesicles to provide a comparison with
vesicles incorporating dithienylethenes. This was necessary given the moderate batchꢀtoꢀbatch
1
4, 38
variability normally observed in lipid vesicle studies.
Correspondingly, we reꢀexamined the kH+
1
4
and N for DOPC and DPPC control vesicles. Consistent with our earlier study , the rate constants
and N were four fold higher in DOPC than DPPC (Table 2), confirming that DOPC vesicles are more
permeable to protons than DPPC vesicles. In addition, the changes in k and N following UV
H+
irradiation were also similar. Overall, the proton leakage observed in pure lipid vesicles occurs
primarily during the transient–pore phase following the formation of the transmembrane pH gradient.
Nonetheless, comparisons with these control samples will ensure that changes in membrane
permeability are related to the inclusion and isomerization of 1–3 in lipid vesicles.
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The k and N were initially examined for amphiphilic spirooxazines 1 and 3 in DOPC and
H+
DPPC vesicles to assess the effect of the position of the photochromic moiety on membrane
permeability. Varying the alkyl chain length has been shown to be an effective approach for
3
6
controlling the embedding depth of the photochromic moiety in bilayer membranes. Prior to UV
irradiation, k for 1 in DOPC vesicles at a mole ratio of 1:30 1/DOPC was two fold higher than the
H+
control vesicles, whereas, k for 3 in DOPC vesicles was similar to the control vesicles (Table 2).
H+
However, N for 1 and 3 were both two fold higher than the control vesicles prior to the addition of
detergent, which might suggest that 1 and 3 are equally disruptive overall in DOPC vesicles. Yet, N
immediately following the addition of base is lower for 1 when compared with 3 (Figure 2). These
results suggest that the presence of 3 with a shorter hexyl chain enhances proton permeation during
the transient–pore phase but provides no significant increase during the solubility–diffusion phase
when compared with 1. Conversely, the inclusion of the 1 with a longer dodecyl chain appears to
provide moderate stability during the transient–pore phase yet the rate of proton permeation is two
fold higher than 3 during the solubility–diffusion phase. Overall, the inclusion of amphiphilic
spirooxazine monomers does disrupt bilayer packing in DOPC vesicles. Although, the extent of the
disruption is dependent on a combination of factors: the relative position of the photochromic moiety
in the bilayer membrane and the phase during which proton permeation occurs.
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Table 2 Proton permeation rate constants and normalized extent of proton permeation determined
a
for 1–3 in DOPC and DPPC vesicles at a mole ratio of 1:30 spirooxazine/lipid
ꢀ
4 ꢀ1
DOPC
k (10 s )
N
H+
UV
No
Yes
No
Yes
Control
1.8 ± 0.5 (57)
4.3 ± 0.4 (6)
8.9 ± 0.6 (6)
1.8 ± 0.1 (6)
1.4 ± 0.6 (52)
4.1± 0.2 (6)
8.7 ± 0.2 (6)
1.7 ± 0.4 (6)
0.31 ± 0.07 (47)
0.57 ± 0.02 (6)
0.89 ± 0.01 (7)
0.57 ± 0.03 (6)
N
0.37 ± 0.06 (45)
0.62 ± 0.05 (6)
0.90 ± 0.01 (6)
0.61 ± 0.03 (6)
1
2
3
ꢀ
4 ꢀ1
DPPC
UV
k (10 s )
H+
No
Yes
No
Yes
Control
1
0.42 ± 0.24 (61)
1.1 ± 0.3 (6)
1.0 ± 0.3 (6)
1.7 ± 0.6 (6)
0.31 ± 0.27 (52)
0.99 ± 0.50 (6)
0.83 ± 0.35 (6)
1.4 ± 0.6 (6)
0.07 ± 0.05 (54)
0.65 ± 0.05 (6)
0.53 ± 0.11 (6)
0.22 ± 0.06 (6)
0.20 ± 0.07 (49)
0.71 ± 0.07 (6)
0.62 ± 0.09 (6)
0.40 ± 0.09 (6)
2
3
a
The error is the standard deviation of the mean taken from n independent measurements. The value
of n is shown in parentheses.
Following UV irradiation no significant change in k or N was observed for 1 or 3 when
H+
compared to the nonirradiated vesicles. This was somewhat anticipated given that thermal reversion
to the closedꢀring isomers at room temperature is most likely complete prior to the formation of the
transmembrane gradient. Consequently, this fluorescence assay is not ideal for assessing the
photocontrol of membrane permeability for thermally reversible photochromic molecules in lipid
vesicles. Nevertheless, this assay does provide a sensitive method for evaluating the relative effect of
spirooxazine inclusion on the membrane permeability of lipid vesicles. As a result, we have also
examined these systems using a potassium ion permeation assay to better assess the effect of
spirooxazine isomerization on membrane permeability (vide infra).
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Figure 2. Normalized extent of proton permeation as a function of time for 1–3 in DOPC (a) and
DPPC (b) vesicles at a mole ratio of 1:30 (i.e., spirooxazine/lipid) prior to UV irradiation. For clarity
error bars are not shown, and only the control is shown before the base pulse and after the addition of
detergent.
In DPPC vesicles, k for 1 and 3 at a mole ratio of 1:30 were at least three fold higher than
H+
the control vesicles (Table 2). In addition, N for 1 and 3 were nine and three fold higher when
compared with the control vesicles, respectively (Figure 2). Given the kH+ for 1 and 3 are within
experimental error of one another, the large difference in N suggests that the inclusion of a
spirooxazine with a longer alkyl chain significantly enhances proton permeation during the transient–
pore phase for lipid vesicles in the rigid gel phase. Thus, 1 is more disruptive in DPPC vesicles
because the photochromic moiety is presumably more deeply embedded in the bilayer membrane.
Also, the relative change in k and N for both monomers is greater in DPPC vesicles than those
H+
observed in DOPC vesicles when compared to their respective control vesicles. A similar trend with
1
4
the phase state of the lipid vesicles has also been observed for amphiphilic dithienylethenes. Like 1
and 3 in DOPC vesicles, there was no significant change in k in DPPC vesicles following UV
H+
irradiation. However, there was a two fold increase in N for 3 but no significant change was observed
for 1. This suggests that the photoisomerization of the hexyl chain analogue disrupts the bilayer
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membrane to a greater extent than the dodecyl analogue in DPPC. Yet, one must be cautious
considering N for the DPPC control vesicles also increases with irradiation. Interestingly, at lower
concentrations of 3 in DPPC vesicles (i.e., 1:40 3/DPPC; Table S1 and Figure S2, ESI) a four fold
increase in N was observed following irradiation, whereas at higher concentrations (i.e., 1:20) no
significant change was seen. Given this concentration dependence on N, we suggest that these
changes are more of a reflection of the higher stability of nonirradiated DPPC vesicles, and those
incorporating low concentrations of spirooxazines, than solely due to the photoisomerization of 3.
That is, the inclusion of spirooxazines at higher concentrations is the primary contributor to a
decrease in vesicle stability and an increase in N. To support this proposal further, we determined N
for both 1 and the dimer 2 in DPPC at 1:50 (Tables S2 and S3, and Figures S9 and S11, respectively,
ESI) and found that both were two fold higher following irradiation. Yet at 1:30, both spirooxazines
had similar N. Overall, the inclusion of the dodecyl derivative in DPPC vesicles appears to be more
disruptive than the hexyl derivative in both nonirradiated lipid systems at 1:30 because k is two fold
H+
larger in DOPC and N is three fold larger in DPPC. As a result, these studies provide additional
support to the hypothesis that the relative position of the photochromic moiety in the bilayer
membrane is important to vesicle permeability. Interestingly, the effect of chain length on membrane
permeability is more pronounced for these spirooxazines than the series of dithienylethenes we
1
4
recently examined. This particular comparison suggests that vesicle stability is more sensitive to the
relative location of a spirooxazine than a dithienylethene, and the former may be a more effective
membrane disruptor.
The bolaamphiphilic spirooxazine 2 was also examined using the fluorescence assay to
determine if the inclusion of a larger dimer is a more disruptive to a bilayer membrane than a
monomer like 1. Ideally, we envisage 2 spanning a bilayer membrane by positioning charged
tethers within the hydrophilic headgroup region of the inner and outer leaflets. However, it is
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possible that the dimer may adopt a Uꢀshaped or folded conformation in lipid vesicles where both
charged tethers reside in a single leaflet. Nonetheless, the inclusion of 2 should cause greater
disruption than a monomer that resides in only one leaflet. The k for 2 in DOPC vesicles at a mole
H+
ratio of 1:30 was five fold higher than the control vesicles, and three and five fold higher than 1 and
3
in DOPC vesicles, respectively (Table 2). Also, N for 2 in DOPC vesicles was three fold higher
than the control vesicles, and 50% higher than both 1 and 3 in DOPC vesicles. We postulate that
the higher k and N values are correlated with the increased molecular size of 2 when compared
H+
with the monomer systems. In DPPC vesicles, 2 also exhibited a large increase in proton
permeation but it was similar to that observed for the monomeric spirooxazines. Following UV
irradiation, and similar to 1 and 3, changes in k and N for 2 in both DOPC and DPPC vesicles
H+
were within experimental error of the nonirradiated samples. Thus, photoisomerization and the
subsequent thermal isomerization back to the closedꢀring isomer of 2 prior to the addition of base
does not have a significant effect on the permeability or the overall integrity of these lipid vesicles.
Notably, k is nine fold higher in DOPC vesicles than in DPPC vesicles, which demonstrates the
H+
importance of the lipid bilayer phase state on membrane permeability. In fact, the curvature
observed for the plot of 2 in DOPC vesicles following the addition of base is more pronounced than
the other systems, which qualitatively indicates that the rate of proton permeation is fast (Figure 2).
Interestingly, the molecular size of these spirooxazines is positively correlated with k in DOPC
H+
vesicles. This suggests that a structure–activity relationship does exist within this series of
photochromic molecules. By contrast, this relationship is not observed in DPPC vesicles, as kH+
was similar for all three molecules. As a result, this lack of selectivity implies that the inclusion of
spirooxazines in gel phase lipid vesicles exhibits an ‘allꢀorꢀnothing’ activity, whereas in fluid phase
lipid vesicles spirooxazine structure is important to activity. Like the spirooxazine comparisons
presented here, k for 2 was at least three fold higher than the dithienylethene monomers in DOPC
H+
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vesicles, which further supports our hypothesis that molecular size is correlated with membrane
permeability in fluid phase vesicles. In addition, similar to our observations with spirooxazines,
there was no correlation between k and the molecular size of the dithienylethenes in DPPC
H+
vesicles.
To measure the effect of spirooxazine concentration on vesicle permeability, we examined
1
–3 in lipid vesicles at various mole ratios. As expected, the spirooxazine concentration in DOPC
and DPPC vesicles do affect both k and N. However, the magnitude of these changes was
H+
dependent on the spirooxazine–lipid system studied. Increasing the mole ratio of 2 from 1:30 to
1
:20 in both DOPC and DPPC resulted in unstable lipid vesicles and complete collapse of the pH
gradient as shown from N values approaching unity (Table S3, and Figures S10 and S11, ESI).
Decreasing the mole ratio to 1:50 significantly reduced proton leakage three fold in DPPC vesicles,
but had a relatively small effect in DOPC vesicles. However, both lipid vesicles were two fold
more permeable than their respective controls. In DOPC vesicles, the k increased 50% from 1:50
H+
to 1:30, whereas in DPPC vesicles the k was constant over the same concentration range. Again,
H+
this highlights the effect of the lipid bilayer phase state on the observed behavior. In DPPC
vesicles, N is dependent on concentration, whereas k is not. Conversely, in DOPC vesicles k is
H+
H+
dependent on concentration, but N is moderately dependent. These trends suggest that the lipid
bilayer phase state affects the predominant mechanism that governs permeation through the bilayer
membrane. That is, the inclusion of 2 primarily affects proton permeation during the solubility–
diffusion phase in fluid phase vesicles, whereas in gel phase vesicles permeation is largely altered
during the transient–pore phase. Similar to 2, the k were reasonably correlated to the
H+
concentration of 3 in DOPC vesicles, although the changes were small by comparison. Unlike 2,
DPPC and DOPC vesicle stability was compromised only when 3 was incorporated at a higher mole
ratio of 1:10 (Table S1, and Figures S8 and S12, ESI). This suggests that the inclusion of the
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amphiphilic monomer is relatively less disruptive in lipid vesicles than the larger bolaamphiphilic
dimer. Interestingly, the highest mole ratio at which lipid vesicles incorporating 1 were not
compromised was dependent on the lipid bilayer phase state. In DPPC, vesicle stability was
compromised when 1 was included at a mole ratio of 1:20 but stable in DOPC vesicles at a mole
ratio of 1:10, although they were very permeable (Table S2, and Figures S9 and 3, ESI). All
together, the permeability of DOPC vesicles incorporating these (bola)amphiphilic spirooxazines is
found to increase in the order of 3 ≤ 1 < 2 when one considers both k and N collectively. Yet for
H+
DPPC vesicles, permeability increases in the order of 3 < 1 ≈ 2. Once again, these differences in
relative permeability indicate that the lipid bilayer phase state and spirooxazine structure are
important to vesicle integrity prior to UV irradiation.
Figure 3. Normalized extent of proton permeation as a function of time for various mole ratios of 1
in DOPC vesicles (i.e., 1 /DOPC) prior to UV irradiation (a) and after 3 min of UV irradiation (b).
Each plot is an average of a minimum of six independent measurements. For clarity error bars are
not shown, and only the control is shown before the base pulse and after the addition of detergent.
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Potassium ion permeation studies in lipid vesicles. As discussed earlier, the fluorescence assay is
not ideal for evaluating the reversible photocontrol of membrane permeability for thermally
reversible photochromic molecules in lipid vesicles. In practice, the fluorescence assay is sensitive
to background, scattered light such that the measurement must be stopped while the irradiation
source is illuminated. In order to monitor changes in permeability uninterrupted with continuous
irradiation, an assay that is not sensitive to background light is required. As a result, we have also
examined spirooxazines in lipid vesicles using a potassium ion permeation assay to better assess the
effect of spirooxazine isomerization on membrane permeability. This assay was used to determine
the rate constant for potassium ion permeation (k ) and the percentage of potassium ion release (%
K+
release). Initially, the kK+ and % release were determined for DOPC and DPPC control vesicles
prior to, and following irradiation with UV and visible light. Consistent with fluorescence assay,
DOPC vesicles are more permeable to potassium ions than DPPC vesicles. In all cases, both the kK+
and % release were at least two fold higher in DOPC than DPPC vesicles (Table 3, and Figures 4
and S13, ESI). Following UV irradiation a 50% increase in k and % release was also observed for
K+
DOPC vesicles, whereas for DPPC vesicles the changes were within experimental error of the
nonirradiated samples. The changes observed for DOPC vesicles may stem from the susceptibility
of the unsaturated double bonds in the DOPC lipid tails to photooxidation, which can increase
4
2ꢀ45
membrane permeability.
By contrast, visible irradiation had no significant effect on k for
K+
either lipid system. In general, the efflux detected from the control samples is largely due to the
high concentration gradient and occurs primarily via the solubilityꢀdiffusion mechanism. Still,
comparisons with these control samples will ensure that changes in membrane permeability are
related to the inclusion and isomerization of 1–3 in lipid vesicles.
Prior to irradiation, the k for both 1 and 3 in DPPC vesicles at a mole ratio of 1:20 were
K+
three fold higher than the control vesicles, whereas in DOPC vesicles, they were 50% higher. This
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Table 3 Potassium ion permeation rate constants and normalized percentage of potassium ion
release determined for 1–3 in DOPC and DPPC vesicles at a mole ratio of 1:20 spirooxazine/lipid
a
ꢀ8
ꢀ1
DOPC
k (10 M s )
% release
No
K+
b
Light
No
UV
Visible
UV and Visible
21 ± 2
Control
1.4 ± 0.1
2.1 ± 0.1
2.7 ± 0.1
2.0 ± 0.1
2.1 ± 0.1
4.7 ± 1.1
3.9 ± 0.5
4.0 ± 0.2
1.9 ± 0.1
0.3 ± 0.2
1.4 ± 0.3
0.9 ± 0.2
15 ± 1
29 ± 4
33 ± 1
25 ± 3
% release
No
1
49 ± 1
2
54 ± 3
3
41 ± 2
ꢀ8
ꢀ1
DPPC
Light
Control
1
k (10 M s )
K+
b
No
UV
Visible
UV and Visible
7 ± 1
0.7 ± 0.1
1.8 ± 0.1
1.6 ± 0.1
1.9 ± 0.1
0.8 ± 0.1
2.8 ± 0.4
4.0 ± 0.5
2.6 ± 0.3
0.6 ± 0.1
2.7 ± 0.4
2.8 ± 0.2
2.4 ± 0.2
6 ± 1
21 ± 1
18 ± 3
24 ± 1
39 ± 3
2
60 ± 5
3
34 ± 1
a
The error is the standard deviation of the mean taken from three independent measurements.
The percentage of potassium ion release was determined after 3.5 cycles of UV and visible light
b
irradiation.
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Figure 4. Normalized percentage of potassium ion release as a function of time for 1–3 in DOPC
a) and DPPC (b) vesicles at a mole ratio of 1:20 (i.e., spirooxazine/lipid) with alternating periods of
(
continuous UV and visible irradiation. Each plot is an average of three independent measurements.
For clarity error bars are not shown.
similarity in k for 1 and 3 suggests that the alkyl chain length has little effect on the permeation of
K+
potassium ions in both lipid systems. The % release for 1 and 3 were also similar at two and four
fold higher in DOPC and DPPC vesicles than the control vesicles, respectively. The results of this
assay are consistent with the fluorescence assay in that the inclusion of amphiphilic spirooxazine
monomers does disrupt bilayer packing in DOPC and DPPC vesicles and increases their
permeability. However, the two fold difference observed in k between 1 and 3 in DOPC vesicles
H+
was not evident in this assay. This variance highlights the increased sensitivity of the fluorescence
assay to changes in molecular structure given proton permeation can also occur by the Grotthuss
mechanism (i.e., hopping along hydrogenꢀbonded chains of water), which is not available to
41
potassium ions. Upon UV irradiation, k for both 1 and 3 in DOPC vesicles increased two fold.
K+
In DPPC vesicles, a small increase was also observed. These results suggest that
photoisomerization of spirooxazines to their more polar ringꢀopened form leads to greater
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disruption of the local bilayer structure, which allows for increased water penetration and enhanced
permeability. Interestingly, the k for both 1 and 3 in DOPC vesicles were significantly lower
K+
upon irradiation with visible light to below control levels. Specifically, 1 and 3 were fourteen and
fourteen fold lower than those observed following UV irradiation, respectively. The large changes
observed in these rate constants suggest that isomerization to the less polar closedꢀring isomers
restores the normal bilayer membrane structure, significantly reducing permeation rates. In
addition, the rate constants were lower than the nonirradiated samples, which suggest that an
equilibrium mixture of both isomers exists in DOPC vesicles prior to irradiation. This thermal
equilibrium is a consequence of the thermochromic behavior of these spirooxazines. By
comparison, the k for both 1 and 3 in DPPC vesicles were relatively unchanged upon irradiation
K+
with visible light. The % release for these systems was also measured after several cycles of
alternating UV and visible irradiation. In DPPC vesicles, the net change in % release for 1 and 3
was nineteen and nine fold higher than the control vesicles, respectively. In DOPC vesicles, both
spirooxazines were two fold higher than the controls. Similar to the fluorescence assay, we also
accredit the relative magnitude of these changes to the higher stability of gel phase DPPC vesicles,
which are three fold less permeable than the fluid phase DOPC vesicles. Once again, the dodecyl
derivative appears to be a more disruptive than the hexyl derivative in both lipid systems because
kK+ upon UV and visible irradiation underwent the largest change in DOPC vesicles and the net
change in % release was largest in DPPC vesicles. These studies are consistent with the
fluorescence assay analysis and further support our hypothesis that the relative position of the
photochromic moiety in the bilayer membrane is important to vesicle permeability. Intriguingly,
the photocontrol of potassium ion permeation from lipid vesicles incorporating a spiropyran with a
1
3
single alkyl chain has been reported previously. Unlike 1 and 3, this spiropyran did not contain a
terminally charged ammonium group. As a result, the perturbation order for controlling membrane
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permeability was reversed when compared with our systems. That is, the closedꢀring isomers
caused greater disruption of the local bilayer structure than the openꢀring isomers. This opposing
result suggests that the inclusion of a terminally charged ‘tether’ improves the organization of the
spirooxazines within the bilayer membrane, enhancing membrane stability. Consequently, this
comparison clearly shows that the perturbation order can be conveniently controlled through the
insertion or deletion of this structural feature. In addition, the differences in potassium ion
permeability under UV and visible irradiation are considerably more pronounced for these
spirooxazines than the spiropyran derivative previously reported.
Similarly, 2 was examined to determine if the isomerization of a biphotochromic molecule is
a more disruptive to a bilayer membrane than 1 or 3. Prior to UV irradiation, k for 2 in DOPC
K+
vesicles at a mole ratio of 1:20 was two fold higher than the control vesicles, and 30% higher than 1
and 3 (Table 3). In DPPC vesicles, k for 2 was similar to 1 and 3. The % release for 2 was also
K+
two and three fold higher in DOPC and DPPC vesicles, respectively, than the control vesicles, and
similar to both 1 and 3. Again, this highlights the lower sensitivity of this assay to changes in
molecular structure, as the large differences in k between 2 and the monomers in DOPC vesicles
H+
were not apparent. Nonetheless, this assay is beneficial in examining the effect of
photoisomerization on membrane permeability. Upon UV irradiation, the k for 2 in DOPC
K+
vesicles increased 50%, whereas a two fold increase was observed in DPPC vesicles. By
comparison, these rate constants were 50% higher than the monomers in DPPC vesicles, but similar
in DOPC vesicles. In general, these increases also suggest that photoisomerization of spirooxazine
dimer to its more polar ringꢀopened form leads to greater disruption of the local bilayer structure
when compared with the nonirradiated vesicles. Like the monomer systems, upon visible
irradiation a three fold decrease in k was observed for 2 in DOPC vesicles, whereas a small
K+
reduction was found in DPPC vesicles. The net change in % release for 2 following irradiation was
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three and 44 fold higher in DOPC and DPPC vesicles than the control vesicles, respectively.
Interestingly, the molecular size of these spirooxazines is also positively correlated with net change
in % release, particularly in DPPC vesicles. As a result, the photoinduced changes in ion
permeation from lipid vesicles incorporating these (bola)amphiphilic spirooxazines is found to
increase in the order of 3 < 1 < 2. As shown earlier, the rate of thermal ring closure is highly
dependent on the lipid bilayer phase state. Consequently, the variation in kK+ prior to, and upon UV
and visible irradiation for these spirooxazines has a cumulative effect, which impacts the net change
in % release. In DPPC vesicles, a trend with molecular size clearly emerges, whereas in DOPC
vesicles this particular trend is less discernable. Overall, these results demonstrate that changes in
membrane permeability can be reversibly photocontrolled with (bola)amphiphilic spirooxazines.
We are currently examining the permeation of encapsulated molecules from lipid vesicles
incorporating these spirooxazines. A preliminary study on the permeation of HPTS from DPPC
vesicles incorporating 1 at a mole ratio of 1:40 shows that for each independent experiment a small
net change in % release is observed upon UV irradiation when compared to nonirradiated vesicles
(Figure 5). However, the average % release values are within experimental error of each other. As
a result, further study is required to assess the activity of these spirooxazines as membrane
disruptors for the delivery of small molecules from lipid vesicles. Moreover, we are currently
exploring mixed lipid systems that produce coexisting liquidꢀdisordered and liquidꢀordered phase
states.
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Figure 5. Normalized percentage of HPTS release as a function of time for 1 in DPPC vesicles at a
mole ratio of 1:40 (i.e., spirooxazine/lipid) with and without periods of UV irradiation. The error is
the standard deviation for the mean taken from five independent measurements.
Conclusions
In summary, we have synthesized (bola)amphiphilic spirooxazines and have examined their
photochromism in organic solution and lipid vesicles. Solvent polarity and the lipid bilayer phase
state have a large effect on the isomerization kinetics. Notably, the thermal ring closure rate
constant for 2 was significantly lower in DPPC bilayers as the higher local viscosity of the gel phase
and the large dimeric structure severely reduces the conformational flexibility required for
spirooxazine isomerization. In addition, two ion permeation assays were used to assess the effect of
spirooxazine inclusion and isomerization on the membrane permeability of lipid vesicles. In all
cases, photoisomerization of the incorporated spirooxazines to their openꢀring form disrupts the
bilayer membrane structure and enhances permeability. However, ion permeation is clearly
sensitive to the lipid bilayer phase state and the relative position of the photochromic moiety in the
bilayer membrane. Overall, we have established that membrane permeability can be reversibly
photocontrolled with (bola)amphiphilic spirooxazines.
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Experimental
Supporting data tables and figures, and synthetic procedures, including NMR and highꢀresolution
mass spectra, are described in the electronic supplementary information (ESI).
1
13
Instrumentation. H and C NMR spectra were recorded at 300.18 and 75.48 MHz, respectively,
1
3
on a Varian Mercury plus spectrometer. Chemical shifts are referenced to solvent signals.
C
1
NMR spectra were recorded with H decoupling. Highꢀresolution mass spectral analyses were
carried out by the University of Calgary on a VG 70SE mass spectrometer and the University of
Saskatchewan on a API Qstar XL mass spectrometer or JEOL AccuTOFꢀGCv 4G mass
spectrometer, using various ionization techniques (e.g., chemical ionization, electron ionization,
electrospray ionization, or field desorption). Steadyꢀstate absorption spectra and kinetics were
obtained at constant temperature (e.g., 21.0 ± 0.1 °C) on a Cary 300 Bio UV–Vis spectrophotometer
equipped with a dual cell Peltier circulator accessory. Absorption spectra were recorded at a scan
ꢀ1
rate, step size, and integration time of 300 nm min , 0.5 nm, and 0.1 s, respectively. Kinetic
measurements were recorded at the λmax for the openꢀring isomer following UV irradiation and with
an integration time of 0.1 s. The effective rate constants for thermal equilibration (k ) were
T
determined from exponential fits of the absorbance decays at λmax. In particular, k = k + k , where
T
1
ꢀ1
k and k are the thermal rate constants for the ringꢀopening and ringꢀclosing reactions,
1
ꢀ1
respectively. Due to the detection limits of our instrumentation, we were unable to determine k1
i.e., k >> k ). It has been shown previously for spirooxazines containing phenanthryl moieties that
(
1
ꢀ1
1
6
k ≈ k . Thus to a good approximation, we will specifically refer to k as the rate constant for
ꢀ1
T
T
thermal ring closure. Steadyꢀstate fluorescence spectra were obtained at constant temperature (i.e.,
1.0 ± 0.1 °C) with a PTI QuantaMaster spectrofluorometer, and the excitation and emission slits
were set such that the bandwidths were 2 nm. Excitation ratio experiments using HPTS were
2
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performed with excitation wavelengths of 403 and 460 nm and an emission wavelength of 510 nm.
The integration time was 2 s. The emission experiments using HPTS were performed with an
excitation wavelength of 413 and an emission wavelength range of 450ꢀ700 nm. The integration
+
time was 0.25 s. Potassium ion permeation was monitored using a MIꢀ442 K ion microelectrode
and a MIꢀ402 Ag/AgCl reference microelectrode from Microelectrodes, which were connected to a
Thermo Scientific Orion Star A325 meter. The electrode potentials were recorded once per minute.
For the proton assay, all samples were irradiated with a 300 W xenon light source from Luzchem,
and measured in a quartz Suprasil fluorescence cell (10 mm × 10 mm) from Hellma. For ultraviolet
irradiations, the xenon light source was filtered with a bandpass filter (Hoya Uꢀ340, λ = 340 ± 42
c
nm). For visible irradiations, a bandpass filter (Thorlabs FB530ꢀ10, λ = 530 ± 5 nm) was used.
c
For the potassium ion assay, all samples were measured in a quartz test tube (14 mm OD, 12 mm
ID, 100 mm long) from Luzchem, which was positioned in a waterꢀfilled and waterꢀjacketed beaker
at constant temperature (i.e., 21.0 ± 0.1 °C). For ultraviolet irradiations, a 6 W UVA (λ = 365 nm)
c
UltraLum TLC lamp was used. For visible irradiations, a 300 W Osram EXR halogen lamp of a
Kodak Ektagraphic III E Plus slide projector fitted with a longpass filter (Schott GGꢀ495, λ = 495
c
±
6 nm) was used. For the fluorescence assay, all samples were measured in a quartz fluorescence
cell at constant temperature (i.e., 20.0 ± 0.1 °C), but irradiated in Pyrex test tubes. The tubes were
loaded onto a rotating sample rack that was placed into a 2 L Pyrex beaker containing a cupric
sulfate solution (100 mM) at constant temperature (20 ± 2 °C), which was positioned in a Rayonet
RPRꢀ100 photochemical reactor equipped with eight 8 W UVA (λ = 350 nm) lamps from
c
Luzchem. For the kinetic studies, all samples were measured in a quartz fluorescence cell and
irradiated with the 6 W UVA lamp for 20 s. The decay in absorbance was monitored at λMC until a
constant baseline was obtained.
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Materials. All reactants (99+%, SigmaꢀAldrich), deuterated solvents (99.9 atom % D, Sigmaꢀ
Aldrich), palladium catalysts (99.9+% Pd, Strem Chemicals), Triton Xꢀ100 (scintillation grade,
Eastman Kodak Company), buffer salts (99+%, SigmaꢀAldrich and Alfa Aesar), 1,2ꢀdioleoylꢀsnꢀ
glyceroꢀ3ꢀphosphocholine (DOPC; >99%, Avanti Polar Lipids), and 1,2ꢀdipalmitoylꢀsnꢀglyceroꢀ3ꢀ
phosphocholine (DPPC; >99%, Avanti Polar Lipids) were purchased and used as received. Copper
(
I) iodide, dimethyl formamide, 1,4ꢀdioxane, tetrahydrofuran, and triethylamine were purified based
4
6
on literature procedures. Flash column chromatography was performed on silica gel (230ꢀ400
mesh, 60 Å, Sillicycle), neutral aluminum oxide (activated, 150 mesh, 58 Å, SigmaꢀAldrich), and
basic aluminum oxide (activated, 60 mesh, 58 Å, Alfa Aesar). 8ꢀHydroxypyreneꢀ1,3,6ꢀtrisulfonic
acid trisodium salt (HPTS; dye content ca. 75%, SigmaꢀAldrich) was recrystallized from ethanol
five times. Deionized water was obtained from a MilliꢀQ Gradient A10 water system (Millipore
Corp.). All aqueous solutions were prepared in phosphate buffer (10 mM Na PO , 100 mM NaCl,
3
4
pH adjusted to 6.4 using H PO ). The organic solvents used in the absorption and kinetic studies
3
4
were spectrophotometric grade.
Procedure for lipid vesicle preparation. The preparation of lipid vesicles has been previously
4
7
described in detail. The same procedures were used to prepare lipid vesicles for the proton and
potassium ion permeability assays; however, higher lipid concentrations were used for the latter.
The concentrations or volumes used for the proton and potassium ion assays are indicated in
parentheses, respectively. Briefly, lipid stock solutions (2 or 10 mg/mL) of DOPC and DPPC were
prepared in chloroform. Separate stock solutions of 1–3 in chloroform (1 mM), 8ꢀhydroxypyreneꢀ
1
,3,6ꢀtrisulfonic acid trisodium salt (HPTS; 10 ꢀM) in phosphate buffer, and potassium chloride
0.20 M) in phosphate buffer were prepared. As an example, to prepare DDPC vesicles containing
at a mole ratio of 1:20 (2/DPPC), a stock solution of 2 (127 ꢀL or 254 ꢀL) and DPPC (1 mL or
(
2
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4
00 ꢀL) were added to a 10 mL round bottom flask. The chloroform was evaporated under reduced
pressure to deposit a lipid film. The lipid film was dried under reduced pressure for at least 3 h and
then hydrated with HPTS or potassium chloride (1 mL) at 50.0 ± 0.1 °C for a minimum of 3 h. The
lipid suspension was left to stand overnight at 4 °C. The hydrated lipid suspension underwent five
freeze and thaw cycles by placing the sample in a dry ice/acetone bath, followed by a warm water
bath and vortexed for 30 s. The sample was extruded 21 times at constant temperature (50.0 ± 0.5
°
C) using 0.1 ꢀm polycarbonate membrane filters. The sample was loaded onto a buffered PD
MidiTrap Gꢀ25 desalting column and eluted with 1.5 mL of buffer to remove extravesicular HPTS
or potassium chloride. The first 5 drops of eluent were discarded to give a purified lipid vesicle
sample (1.3 mg/mL or 2.7 mg/mL).
Proton permeability assay. The protocol for the HPTS fluorescence assay has been previously
1
4
described. Briefly, 2 mL of phosphate buffer and 100 ꢀL of the lipid vesicle sample were
transferred to a fluorescence cell and stirred. For nonirradiated samples, 50 ꢀL of 0.5 M NaOH was
added following a 150 s equilibration period. After 700 s, the vesicles were lysed with 50 ꢀL of a
5
% Triton Xꢀ100 solution. For irradiated samples, the base was added following a 70 s
equilibration period, which was preceded by 3 min of UV irradiation, and an initial 80 s
equilibration period. All data collected during the 150 s equilibration period and ca. 30 s after the
addition of base was not used when fitting the data (i.e., ca. 180 s was t ). The relative emission
0
intensity E /E (I) was calculated and then the extent of ion permeation (N ) was calculated using
4
03 460
t
Eq. 1,
(It − I
)
)
0
N =
(1)
t
(
I∞ − I0
29