Elucidation of the Mechanisms for the Underlying Depolarization and Reversibility by Photoactive Molecule
Tomohiro Numataa Ryosuke Fukudab Mitsuru Hiranoc Kazuma Yamaguchic,d Kaori Sato-Numataa,e Hiroshi Imahorif,g Tatsuya Murakamib,g
aDepartment of Physiology, School of Medicine, Fukuoka University, Fukuoka, Japan, bDepartment of Biotechnology, Graduate School of Engineering, Toyama Prefectural University, Toyama, Japan, cDepartment of Synthetic Chemistry and Biological Chemistry, Graduate School of Engineering, Kyoto University, Kyoto, Japan, dHiroshima Regional Taxation Bureau, Hiroshima, Japan, eJapan Society for the Promotion of Science, Tokyo, Japan, fDepartment of Molecular Engineering, Graduate School of Engineering, Kyoto University, Kyoto, Japan, gInstitute for Integrated Cell-Material Sciences (iCeMS), Kyoto University Institute for Advanced Study (KUIAS), Kyoto University, Sakyo-ku, Kyoto, Japan
Key Words Photo-induced
depolarization • Membrane capacitance • Potassium channel • Patch-clamp • Photo-induced
charge-separation molecule Abstract Background/Aims: Light-induced
control of the cell membrane potential has enabled important advances in the
study of biological processes involving the nervous system and muscle activity.
The use of these light-induced modifications is expected in various medical
applications, including the control of physiological responses and the recovery
of lost functions by regulating nerve activity. In particular,
charge-separating linkage molecules (Charge-Separation (CS) molecules) can
depolarize cells by photoexcitation without genetic processing. However, the
molecular mechanisms underlying cell membrane depolarization are unknown and
have hindered its application. Here, we show that CS molecules localized in the
cell membrane of PC12 cells using a high-density lipoprotein (HDL)-based drug
carrier can excite the cells through a novel membrane current regulation
mechanism by light irradiation. Methods: Membrane
potential, channel activity, and membrane capacitance were measured by patch
clamp method in rat adrenal gland pheochromocytoma (PC12) cells and KV-overexpressing
PC12 cells. CS molecules localized in the cell membrane of PC12 cells using
HDL-based drug carrier. The localization of CS molecule was measured by a confocal
microscopy. The mRNA expression was tested by RT-PCR. Results: Current
clamp measurements revealed that the photo-activated CS molecule causes a sharp
depolarization of about 15 mV. Furthermore, it was shown by voltage clamp measurement
that this mechanism inactivates the voltage-dependent potassium current and simultaneously
generates photo-activated CS molecule induced (PACS) current owing to the loss
of the cell membrane capacitance. This activity continues the depolarization of
the target cell, but is reversible via a regenerative mechanism such as
endocytosis and exocytosis because the cell membrane is intact. Conclusion:
Thus, the mechanism of photo-induced depolarization concludes that
photo-activated TC1 causes depolarization by generating PACS current in
parallel with the suppression of the K+ current. Moreover, the
depolarization slowly restores by internalization of TC1 from the membrane and
insertion of new lipids into the cell membrane, resulting in the restoration of
KV to
normal activity and eliminating PACS currents, without cell damage. These
results suggest the possibility of medical application that can safely control
membrane excitation. Introduction Optogenetic therapies that
enable local excitement have high potential as therapeutic strategies for
neurological diseases, such as Alzheimer's, because they enable precise
spatiotemporal control of cells [1]. Most of these technologies require prior
gene manufacturing, such as the gene transfection of light-sensitive proteins
or modification of target molecules with light-sensitive compounds, and their
use is limited [2]. Using these applications adds technical complexity and
risk. In contrast, quantum dots, gold nanomaterials, magneto-electric
nanomaterials, piezoelectric nanomaterials, and caged compound have been shown
to cause membrane excitation in cells simply by treating the cells with
nanomaterials before stimulation [3-7]. Therefore, it is attractive as a tool
for basic research and clinical applications related to various membrane
excitations. Despite increasing evidence that
photosensitive compounds can be used to control cell function, the underlying
mechanisms are inadequate. Photostimulation mediated by photosensitive
compounds is thought to generate heat or reactive oxygen species, affect ion
channel gene expression and gating, and excite cells by increasing membrane
conductance [3, 8-10]. In part, this is because most previous experiments with
light-induced charged molecules did not directly assay target cells through
electrophysiology, but rather a downstream analysis of their effects (e.g.,
imaging, biochemistry). Furthermore, studies on cells excited by light-induced
molecular stimulation only change the state of the cells, and many studies
still have the problem of reversibility and safety [11]. Thus, solving this
problem will further advance research areas for cells and biological
applications. Here, the mechanism of
photostimulation-induced depolarization by the photo-induced charge-separation
(CS) molecular probe TC1, a ferrocene–zinc porphyrin–fullerene linked triad [12],
in mammalian neural PC12 (rat pheochromocytoma) cells was directly investigated
via electrophysiology. It was expected that the photoexcitation of TC1 would
result in photo-induced electron transfer from the zinc porphyrin excited
singlet state to the fullerene followed by a second electron transfer from the
ferrocene to the zinc porphyrin radical cation, generating the ferrocenium
cation-zinc porphyrin–fullerene radical anion pair in PC12 cell membrane [12]. We
then attempted to perform membrane current measurements for an extended time,
which showed that repolarization was possible by removing TC1 from the cell
membrane via membrane transport. This finding has important implications for
the photostimulation application of light-induced CS probes in the nervous
system and other organs, and reports the effect of the cell membrane quality on
membrane ion transport. Materials
and Methods Cell culture and
cDNA expression Rat adrenal
gland pheochromocytoma (PC12) cells and KV-overexpressing PC12 cells
were cultured in Dulbecco’s modified Eagle’s minimal essential medium (DMEM)
with high glucose (Merck Millipore, Darmstadt, Germany) supplemented with 5% fetal
bovine serum (FBS), 5% horse serum (HS), 30 U/mL penicillin and 30 μg/mL
streptomycin (Nacalai Tesque, Inc., Kyoto, Japan) under 5% CO2 and
95% air at 37°C. PC12 cells were used for the following experiments with this
culturing condition unless stated otherwise. PC12 cells for overexpression were
plated on culture dishes for 24 hours. Then, the PC12 cells were transfected
with KV1.6-IRES2-AcGFP1, KV2.1-IRES2-AcGFP1, KV3.4-IRES2-AcGFP1,
or KV4.2-IRES2-AcGFP1. Lipofectamine 2000 (Invitrogen, Carlsbad, CA,
USA) was used as the transfection reagent in accordance with the manufacturer’s
instructions. Electrophysiological measurements were performed at 36–72 h
after transfection. Rat KV1.6, KV2.1, KV3.4,
and KV4.2 (GenBank accession No. NM_023954, NM_013186, NM_001122776,
and NM_031730.2, respectively) were cloned from rat whole brain Marathon-Ready
cDNA (BD Biosciences, San Jose, CA, USA) using a PCR-based approach, and
subcloned into the expression vector pIRES2-AcGFP1 (Clontech, Mountain View,
CA, USA). CS molecule and
drug delivery system Materials. The following
materials were used as per the manufacturer’s instructions. 1-Palmitoyl-2-oleoyl-glycero-3-phosphocholine
(POPC: NOF Corporation, NY, USA), sodium cholate, dimethyl sulfoxide, (16:0)
Liss Rhod PE (Avanti Polar Lipids), urea (Nacalai Tesque, Inc.), Spectra/Por
Dialysis Membrane (Spectrum Laboratories, Inc., CA, USA), potassium bromide,
Phospholipid C-test (FUJIFILM Wako Pure Chemical Corporation, Osaka, Japan),
EMD Millipore Amicon Ultra-15 Centrifugal Filter Units (Fisher Scientific, MA,
USA), DC Protein Assay Kit (Bio-Rad Laboratories, CA, USA), PD-10 desalting
column (GE Healthcare, CA, USA). The donor–acceptor (D-A) linked molecule, TC1
was prepared according to the literature [12]. The TC1 molecule was designed to
reduce the aggregation by increasing the bulkiness around the porphyrin moiety
of the ferrocene–zinc porphyrin–fullerene (Fc–ZnP–C60) triad [13]. The
photo-induced charge-separation (CS) yields of TC1 are reported to be 50% in
DMSO/H2O mixture (1/99, v/v) and 18% in liposome in DMSO/H2O
(1/99, v/v) [12]. Preparation of
genetically engineered HDL and loading of TC1 in HDL. The HDL mutant
used in this study was prepared in accordance with a method reported previously
with a minor modification [14]. The required amount of POPC and (16:0) Liss
Rhod PE at a molar ratio of 99:1 was solubilized in phosphate buffered saline (PBS)
containing 30 mg/mL sodium cholate (SC) at a molar ratio of lipid: SC = 1:3.9.
The protein component of the mutant, which was an apoA-I protein with the
N-terminal 44 amino acids deleted and a TAT (transactivator of transcription)
peptide fused at the C-terminus, was solubilized in PBS containing 4 M urea and
mixed with the above lipid/SC mixture at a protein:lipid molar ratio of 1:300.
The mixture was incubated overnight at 4°C and then dialyzed against PBS at 4°C
with a Spectra/Por Dialysis Membrane (MWCO = 50 kDa). The dialyzed dispersion
was centrifuged at 20,000 × g at 4°C for 30 min to remove any debris.
The obtained HDL sample was purified by density gradient ultracentrifugation in
accordance with the method by Suda et al. [15] with minor modifications.
Briefly, the density of the HDL sample (3 mL) was adjusted to 1.31 g/mL using
potassium bromide. A four-step gradient solution of potassium bromide (9.0 mL,
1.21 g/mL; 11.4 mL, 1.063 g/mL; 9.9 mL, 1.019 g/mL; 3.6 mL, 1.006 g/mL) was
prepared in a polyallomer tube (Beckman Coulter, Inc., Tokyo, Japan).
Ultracentrifugation was carried out at 16°C in a HITACHI CP80NX using a 70 Ti
rotor (Beckman Coulter, Inc.) at 286,000 × g. The sample was collected
from a fraction in the 1.019 g/mL density range and was dialyzed against PBS at
4°C overnight. The dialysate was centrifuged at 20,000 × g at 4°C for 15
min. The supernatant was collected and concentrated with an Amicon Ultra
Centrifugal Device (MWCO = 50 kDa) by centrifugation at 5,000 × g at 4°C
until the total sample volume was ~3 mL. Incorporation of TC1 was conducted by
following our previous methods [13]. Briefly, the HDL mutant (91 µg protein/mL)
in PBS was mixed with TC1 in dimethyl sulfoxide (0.4 mM) at a volume ratio of
9:1, and then the mixture was incubated at room temperature for 1 h. The
TC1-HDL mutant complex was purified with a PD-10 desalting column equilibrated
with 0.9% NaCl. Characterization
of TC1-HDL mutant complex. The protein and POPC concentrations in the HDL
mutant were determined by the Lowry method using the DC Protein Assay Kit and a
phospholipid-specific enzymatic assay using the Phospholipid C-test Wako
(FUJIFILM Wako Pure Chemical Corporation), respectively. The concentration of the
CS molecules or (16:0) Liss Rhod PE was determined spectroscopically with a
V-630 spectrophotometer (JASCO Corporation, Tokyo, Japan, 300–700 nm) or a
Fluoromax 4 spectrophotometer (HORIBA, Ltd., Kyoto, Japan, 575–640 nm). The size
distribution and zeta potential measurements were performed with a Nanotrac
UPA-EX250 particle size analyzer (MicrotracBEL Corp., Osaka, Japan) in PBS and
a Zetasizer Nano Z (Malvern Panalytical, Malvern, UK) in 20 mM Tris·HCl (pH
7.4) containing 10% PBS, respectively. Confocal
microscopy PC12 cells were
adhered to a poly-L-lysine (PLL) coated cover glass. After 24 h, the medium was
removed, and the cells were washed once with Tyrode solution ((in mM) 140 NaCl,
5 KCl, 2 CaCl2, 1 MgCl2, 10 HEPES, and 10 D-glucose (pH
adjusted to 7.4 with NaOH, and osmolality adjusted to 300 mOsmol/kg H2O
with D-mannitol) and resuspended with Tyrode solution. Then, an aqueous
solution of the (16:0) Liss Rhodamine PE-labeled TC1-cell-penetrating (cpHDL)
was added to the cells to adjust the concentration to a final value of 0.5
μM based on the compound. The cells were incubated for 3 min and washed with
Tyrode solution once and resuspended in Tyrode solution with or without
inhibitors. The cells incubated at room temperature (22-25°C) for 0 and 60 min
after 2-min illumination (525−550 nm, input power 2 mW cm−2,
see in Patch Clamp Measurements), the cells were fixed with 4%
paraformaldehyde (Nacalai Tesque, Inc.), and then mounted in Fluoromount-G
mounting medium (Southern Biotechnology, Birmingham, AL, USA). Fluorescence
images were acquired with a confocal laser-scanning microscope (Zeiss LSM710,
Carl Zeiss Microscopy GmbH, Jena, Germany), which was equipped with a 40× oil
objective lens. Rhodamine signals were acquired and line analyses were made
with ZEN software (Carl Zeiss). Eight lines were drawn from the center with a
40° angle. The rhodamine signal for each line was considered positive when both
points where the line crossed the edge of the cell showed a signal intensity
above the half-maximum intensity. The relative rhodamine signal per cell was
defined as the ratio of the positive lines. Patch Clamp
Measurements Whole-cell patch
recordings for the current clamp and voltage clamp were recorded using a nystatin-perforated
patch technique on PC12 cells at room temperature (22–25°C) with an Axopatch
200B (Molecular Devices, Axon Instruments, Sunnyvale, CA, USA) patch-clamp
amplifier. The patch electrodes prepared from borosilicate glass capillaries
had a resistance of 4-5 MΩ. For perforated whole-cell recordings, low
access resistance measurements were achieved by creating a blunt-tip electrode
on the pipette that increased the total surface area of the membrane piece
drawn into the pipette [16]. In nystatin-perforated whole cell recordings,
series resistance (<15 MΩ) was compensated (to 70–80%) to minimize
voltage errors. In current clamp recordings, currents were clamped to zero by
the fast current clamp mode of the Axopatch 200B. Current signals were filtered
at 5 kHz with a four-pole Bessel filter and digitized at 20 kHz. pCLAMP
software (version 10.5: Molecular Devices, Axon Instruments) was used for
command pulse control, data acquisition, and analysis. For membrane capacitance
recordings were measured using the established conventional patch-clamp voltage
application method [17]. Briefly, the membrane test mode of Clampex software
(Molecular Devices, Axon Instruments) was used to obtain the values calculated
from the capacitance component. The nystatin perforated patch was measured
according to procedures based on previous reports [18, 19]. Briefly, the
pipette tip was dipped into a normal pipette solution for about 10 seconds,
then backfilled with the solution containing nystatin. Then, within a few
minutes, we achieved a high input seal resistance by setting up the electrode
holder, approaching the cell, and forming a giga-seal on the cell. The nystatin
solution was dissolved by placing 200 mg of nystatin in 1 ma of
dimethylsulfoxide (DMSO: FUJIFILM Wako Pure Chemical Corporation), vortex for 1
min, and then sonication for 10 min. From this stock solution, 1 μL was
added to 1 mL pipette solution, vortexed again for 1 min, and sonicated for 10
min. The final solution was filtered through a 0.20 µm filter (Millipore
Corporation, Japan) to obtain a pipette solution. This solution was used within
the day. For conventional and nystatin-perforated whole-cell recordings, the Na+-based
bath solution contained (in mM) 140 NaCl, 5 KCl, 2 CaCl2, 1 MgCl2,
10 HEPES, and 10 D-glucose (pH adjusted to 7.4 with NaOH, and osmolality
adjusted to 320 mOsmol/kg H2O with D-mannitol). The pipette solution
contained (in mM) 55 K2SO4, 20 KCl, 5 MgCl2,
0.2 EGTA, and 5 HEPES (pH adjusted to 7.4 with KOH, and osmolality adjusted to
300 mOsmol/kg H2O with D-mannitol). An Ag-AgCl pellet-3M KCl-agar
bridge was used for reference electrode. The hν-induced Vm in
Fig. 1 was calculated using the following equation: hν-induced Vm
(mV) = VCtl−Vhν, where VCtl
and Vhν are the membrane potential values observed
before and by the end of the illumination. Iafter is a stable value
of I after 30 minutes from illumination. I1 and I2 show a
whole-cell current at +100 mV, and at –100 mV, respectively. For Fig. 2, 3, 4,
5, and 6, ramp pulses were applied every 5 s from –100 mV to +100 mV from a
holding potential of −60 mV at a speed of 4 mV/ms. For Fig 6b, the
recovery of membrane potential (%) was calculated using the following equation:
recovery of membrane potential (%) = ((Vpeak-Vafter)/(Vpeak-Vbefore))
× 100, where Vbefore, Vpeak are the peaks values before
and after illumination of Vm, respectively. Vafter is a
stable value of Vm after 30 minutes from illumination. The recovery
of K+ current (%) in Fig. 6d was calculated using the following
equation: recovery of K+ current (%) = ((Ipeak1-|Ipeak2|)-(Iafter1-|Iafter2|))/((Ipeak1-|Ipeak2|)-(Ibefore1-|Ibefore2|)))
× 100, where Ibefore, Ipeak are the peaks values before
and after illumination of I, respectively. Iafter is a stable value
of I after 30 minutes from illumination. The recovery of PACS current (%) in
Fig. 6e was calculated using the following equation: recovery of PACS current
(%) = (|Ipeak2|-|Iafter2|))/(|Ipeak2|-|Ibefore2|)
× 100, where Ibefore, Ipeak are the peaks values before
and after illumination of I, respectively. The light used in the experiment was
from a mercury lamp house (C-HGFIE: Nikon, Tokyo, Japan) attached to an
inverted microscope Ti (Nikon) that excited cells through an objective lens
through TRITC and ND filters (Nikon). The wavelength and intensity of the
excitation light were measured with a spectrophotometer C-7000 (SEKONIC, Tokyo,
Japan) and confirmed to be 525 to 550 nm and 2 mW cm-2,
respectively. RNA isolation
and RT-PCR Total cellular
RNA was extracted from PC12 cells using ISOGENE (Nippon Gene, Tokyo, Japan) in accordance
with the protocol supplied by the manufacturer. Five hundred nanograms of total
RNA were reverse-transcribed into the first-strand cDNA by use of the RNA LA
PCR kit (AMV) Ver1.1 (Takara, Shiga, Japan) at a final volume of 20 μL.
Expression levels of KV 1–12 mRNA in PC12 cells were determined by
RT-PCR. Gene-specific primers used for PCR were designed with Primer3 software
(http://bioinfo.ut.ee/primer3/) and NCBI BLAST (www.ncbi.nlm.nih.gov/blast/) to
identify complementary sequences in the rat genome. The primers used for PCR
amplification and the predicted lengths of the PCR products are summarized in
Supplementary Table S1 (for all supplementary material see
www.cellphysiolbiochem.com). PCR was conducted with a GeneAmp PCR system 9700
(Applied BioSystems, Foster City, CA, USA) using LA Taq polymerase with GC
buffer (Takara) for 32 cycles under the following conditions: initial denaturation
was 3 min at 95°C, then 30 sec at 95°C, followed by a 30-sec annealing step at
63°C and 60-sec elongation at 72°C, and a final elongation of 7 min at 72°C.
Each RT-PCR experiment was independently repeated twice to test the amplification
reproducibility. The specificity of the amplicons was checked by sequencing the
PCR products to confirm that its sequence corresponded to the target gene. Detection of
photo-induced cell damage Detection of
membrane damage was performed using the Cytotoxicity LDH Assay Kit-WST (Dojindo
Laboratories, Kumamoto, Japan) in accordance with the manufacturer's protocol.
Nuclear cell viability analysis was performed by double staining using acridine
orange (AO) and propidium iodide (PI) assays. For the AO/PI reagents, a Cell
Viability Kit (Logos Biosystems, Korea) was added to each sample. Briefly,
reagent-loaded cells were left at room temperature for 5 minutes in accordance
with the manufacturer's protocol, and then the cell sample images were obtained
from a Countess II-FL automated cell counter (Thermo Fisher Scientific,
Waltham, MA, USA). AO-positive cells were counted as live cells, and
PI-positive cells were counted as necrotic cells. In these assays, TC1-treated
cells were light-stimulated with Tyrode's solution for 5 minutes and then
stimulated for 1 hour at room temperature under the same conditions as in the
electrophysiology experiments. Each assay was performed three times and the
results were analyzed statistically. Drugs The K+
channel blockers and their suppliers were: tetraethylammonium (TEA: Sigma, St.
Louis, MO, USA), 4-Aminopyridine (4-AP: Sigma), and XE-991 (10,10-bis(4-pyridinylmethyl)-9(10H)
-anthracenone dihydrochloride: alomone labs, Jerusalem, Israel). The cation and
anion channel blockers and their suppliers were: SKF96365 (1{β-[3-(4-methoxyphenyl)propoxyl]-4-methoxyphenethyl}-1H-imidazole
hydrochloride: Sigma) and DIDS (4,4'-diisothiocyanostilbene-2,2'-disulfonic
acid: Sigma). The blockers in Fig. 6, 7 and their suppliers were: brefeldin A
(BFA), genistein (GEN), and phenylarsine oxide (PAO) were purchased from TCI (Tokyo,
Japan). The TEA and 4-AP were dissolved in the experimental solution before
use. Concentrated stock solutions of SKF96365 (10 mM), DIDS (100 mM), XE-991
(10 mM), BFA (50 mM), GEN (200 mM), and PAO (5 mM) were prepared in DMSO and
stored at −20°C until required. The final concentration of DMSO was
always kept below 0.1%, a concentration that did not interfere with the
measurements. Statistical
analyses All data are
expressed as means ± SEM. We accumulated the data for each condition from at
least three independent experiments. We evaluated statistical significance with
Student’s t-test for comparisons between two mean values. The data in Fig. 1e
were compared using one-way analyses of variance (ANOVA), followed by Tukey’s
multiple comparisons test. In all cases, a value of P < 0.05 was considered
significant. Results TC1 loaded in a
lipoprotein-based drug carrier causes efficient light-induced depolarization by
decreasing K+ current and increasing PACS current Using an appropriate drug
delivery system, light-induced CS molecules that efficiently localized to cell
membranes caused large membrane depolarization [13]. The TC1 molecule, one of a
series of previously developed CS molecules, showed only a very small
depolarization (~6.3 mV) in cellular applications because it did not perform the
appropriate drug delivery system strategy [12]. This greatly delayed its
technological development for biological applications. Achieving more efficient
depolarizing systems is a prerequisite for the development of biological tools
and medical applications. Therefore, we adopted a previously developed discoidal
high-density lipoprotein (HDL)-based drug carrier [14] that could be
efficiently localized to the cell membrane. A system for encapsulating TC1 was
designed to evaluate the light-induced depolarization in PC12 cells using
nystatin-perforated whole-cell recordings. As shown in Fig. 1, the cells
treated with the TC1 molecule incorporated in cell-penetrating high-density lipoprotein (TC1-cpHDL) caused
depolarization gradually after light irradiation, and reached a stable
depolarization of 16 mV within 5 minutes after stimulation. In contrast, TC1
cells without cpHDL reached a stable depolarization of 6 mV with 10 min after
stimulation. Cells without TC1 or cpHDL did not show a significant change in the
membrane potential upon light irradiation. These results indicate that we
developed a more efficient depolarizing system of TC1 that was 159% larger and
twice as fast when using cpHDL in comparison with TC1 without the drug delivery
system. This increase in efficiency is consistent with the current (Fig 7a,
control) and previous results [12, 13] using cpHDL, which is due to the uniform
distribution and increased number of CS molecules (i.e. from 1.6 × 106
to 2-3 × 107) inserted into the cell membrane. Also, TC1-cpHDL cells
induced depolarization by illumination, and reached a specific range of
depolarization, indicating that this recording was performed stably. Under the
experimental conditions based on the above results, we then investigated the
underlying membrane current that causes membrane potential changes. Because rapid changes in the cell
membrane potential often need to be driven by ion-flux, the effect of TC1 on the
cell membrane currents in PC12 cells under nystatin-perforated patch clamps was
next examined by voltage clamp. As shown in Fig. 2a and 2b, the current-voltage
relationship induced by the ramp pulse in TC1-treated cells was found to show current
properties with an outward rectification and a reversal potential of −75 to
−80 mV (Fig. 2b, inset x). Next, photostimulation of the cells caused the
whole-cell current to slowly increase at −100 mV, while simultaneously
decreasing at 100 mV. Later, both changes reached a plateau within 5 minutes.
(Fig. 2a–d). After the current reached the plateau, a reversal potential near 0
mV and a linear I-V curve was observed (Fig. 2b, inset y). Since
the photostimulated CS molecule, TC1, generated a membrane current, we call
this current the photo-activated CS molecule-induced (PACS) current. The biophysical properties of
the current shown in the cells before photostimulation are typical because they
showed a value close to the ideal K+ equilibrium potential, which
was −82 mV from the experimental solution conditions. Furthermore, the
channel activity increased with depolarization. These biophysical features suggested
that this is a voltage-gated K+ channel. A decrease in the outward
current and a shift in the reversal potential to around 0 mV were observed after
photostimulation, suggesting that the TC1 activity mainly suppressed the voltage-dependent
K+ currents. To investigate the PACS current characteristics
with a linear rectification increased by light stimulation, we tried to inhibit
the ion channel activity by drugs. Photostimulation-induced whole-cell currents
with linear I-V relationships were not affected by the
application of sufficient amounts of 100 µM DIDS, a broad anion channel
inhibitor [20], or 10 µM SKF96365 a broad cation channel inhibitor [21] (Fig. 3a–d). Electrophysiological membrane
models generally assume that cell membranes are insulators and capacitors.
However, because the membrane is not a perfect insulator, electricity may flow
because the membrane capacity and permittivity are affected by physical changes
such as the lipid content and the generation of conductive resistance. To
examine the cell membrane properties, the membrane capacitance was measured using
the established conventional patch clamp voltage application method [17]. As
shown in Fig. 3e, photostimulation through TC1 gradually reduced the membrane
capacitance. After an 8-minute stimulation time, the membrane capacitance was
reduced by 24.7% in comparison with the control without TC1 (Fig. 3f). These
results suggest that the photostimulated TC1 altered the membrane properties
independently of ion transport, causing PACS currents. TC1 suppresses intrinsic and
extrinsic voltage-gated K+ channels in PC12 cells Because the data shown in Fig. 2
suggested that light-induced TC1 suppressed endogenously expressed
voltage-gated K+ (KV) channels in PC12 cells, we
investigated the molecular and functional basis of the KV channels. PC12
cells are frequently used as a neuronal cell line, and the functional molecular
expression of KV1–3 has been previously reported [22]. However, no
studies have comprehensively investigated KV expression. Robust amplification of PCR
products of the expected size (see Supplementary Table S1) from the reverse
transcribed RNA was obtained with specific primers for KV1.1, KV1.2,
KV1.3, KV1.4, KV1.5, KV1.6, KV2.1,
KV2.2, KV3.1, KV3.3, KV3.4, KV4.2,
KV4.3, KV7.2, KV7.3, KV10.1, KV11.1,
KV11.2, and KV12.2. Sequencing analysis of KV
channels expressed endogenously in PC12 cells showed that almost all expressed KV
channel families were completely identical to the corresponding sequence of
each rat voltage-gated K+ channel (Fig. 4a). Next, we attempted to perform
pharmacological studies using the well-known K+ channel blockers TEA
and 4-AP, for the molecular classification of KV channels and the KV7-specific
channel blocker XE-991. The voltage-gated K+ channel current in the PC12
cells was inhibited by 50% with 5 mM TEA and 65% with 2 mM 4-AP. The current
inhibition by XE-991 at 10 µM was minimal (<10%) and is summarized in Fig.
4b and 4c. KV1, KV4, and KV7 were selected as
candidates in consideration of the gene expression, the fact that the K+
currents were activated near the resting membrane potential of PC12 cells
(<0 mV, Fig. 4b, control), and because the molecular identification was
based on low-potential (LVA)-type KV channels. Furthermore,
considering the pharmacological characterizations of TEA, 4-AP, and XE-991 [23,
24], KV4 was found to be a current mainly suppressed by TC1. To
investigate the TC1 target discrimination more clearly, an experimental PC12
cell system overexpressing rat KV4.2, KV1.6, KV2.1,
and KV3.4 channels that showed relatively higher expression in a
was constructed. As shown in Fig. 5a, almost all large outward currents
observed with the control currents in KV1.6, KV2.1, KV3.4
or KV4.2 expressing cells were unexpectedly suppressed by photo-induced
TC1 stimulation. All cells notably showed a PACS current of ~2 nA at −100
mV, similar to that seen in Fig. 2. Therefore, in the analysis of this
experiment, the absolute value of the current obtained at −100 mV was
subtracted from the current value obtained at +100 mV to obtain a purer K+-derived
current. These results imply that TC1
suppresses all KV channel types, thus making changes to the cell
membrane that may commonly affect membrane transport proteins. The TC1-induced depolarizing
effect of light stimulation is reversible via membrane recycling Previous work showed that
photoreversibility studies on the effects of light-induced CS probes on
cell-membrane depolarization remained a challenge [13]. In particular, the lack
of CS molecule light-off reactions led to problems with the potential of safely
using CS molecules for biological applications. To overcome these problems, we
tracked the membrane potential and current through an incredibly long record of
sustained depolarization, independent of the TC1 molecule off-response in PC12
cells. As shown in the control of Fig.
6a, the light-induced TC1 activity caused a depolarization of approximately 15 mV
(as in Fig. 1), followed by a sustained depolarization even after the
irradiation was stopped. However, after 30–40 minutes of sustained
depolarization, they surprisingly repolarized to the basal membrane potential
(Fig. 6a, control). In addition, the membrane current underlying the membrane
potential change was examined by evaluating the recovery rates of the K+
current and PACS current to the initial recorded values (Fig. 6c–e, control). One plausible mechanism of the cell
membrane function recovery involves the recycling of membranes using endocytic
and exocytotic pathways [25]. To further investigate this possibility, we
examined the membrane potential and membrane current in the presence of
inhibitors that target the membrane transport pathway. Administration of brefeldin A
(BFA), an inhibitor of the exocytosis pathway from the Golgi apparatus to the
cell surface, had no effect on restoring the membrane potential in comparison
with the controls (Fig. 6a and 6b: BFA). However, genistein (GEN), an inhibitor
that blocks caveolae-mediated endocytosis, had a small effect on restoring the membrane
potential. The clathrin-mediated endocytosis inhibitor phenylarsine oxide (PAO)
also showed greater inhibition (Fig. 6a and 6b). As shown in Fig. 6c–e, the
effect of membrane transport pathway inhibitors on the recovery of the K+
current did not change with BFA treatment. However, the GEN and PAO treatments
were significantly weakened. The effect of the PACS current on the recovery
rate was reduced for all three treatments. The above results suggest that the
endocytic pathway is important for the K+ current restoration.
Furthermore, both exocytosis and endocytosis pathways are plausibly involved in
the restoration of membrane lipids. Finally, confocal microscopy was
used to determine whether the membrane potential recovery upon depolarization from
light-induced TC1 activity was dependent on changes in the TC1 localization. As
shown in Fig. 7a, control, treatment of TC1-encapsulated rhodamine-labeled cpHDL
was localized to the PC12 cell membrane at 0 minutes, which is consistent with
a report that previously investigated the cell membrane localization of CS-encapsulated
cpHDL [13]. We then investigated the cellular localization of TC1 using the
same membrane transport pathway inhibitors as with the experiment shown in Fig.
6. After 4 minutes of light stimulation on the TC1-treated cells, the rhodamine
fluorescence observation disappeared 60 minutes later from the cell membrane in
the control sample. The same was observed for the BFA-treated cells. However,
the cells treated with GEN and PAO showed that the rhodamine fluorescence
partially remained in the cell membrane. In a series of these experiments, LDH (lactate
dehydrogenase) release experiments and PI/Hoechst staining showed that there
was almost no damage to the cell membrane (Fig. 7c and 7d). In summary, the TC1-induced
depolarization from changes in the membrane property was repolarized before
light stimulation by TC1 internalization and membrane function regeneration of
intact PC12 cells via the membrane-trafficking system. Discussion Although the potential utility
of photosensitive molecules, including photo-induced CS probes, has been
demonstrated in recent cell biology studies, a poor understanding of the
underlying mechanisms has slowed the development of valuable scientific and
clinical applications. This study revealed an important and unexpected
mechanism that causes the light-induced depolarization of CS molecules. This
reversibility from depolarization has been shown to cause repolarization in
correlation with cell membrane recycling (Fig. 8). Controlling the membrane
potential by photochemical switching is well known as an attractive way to
control cellular and biological functions, as demonstrated in optogenetics and
caged compounds. In particular, the control of membrane excitability using a
photosensitive compound does not involve genetic manipulation, and may lead to
attractive applications for disease treatment through realistic neural activity
control. However, the control of a light-induced membrane potential, using
photosensitive compounds, has not been fully elucidated for depolarization and
its reversibility. The rate of change in the light-induced membrane potential
and membrane current was measured directly by the patch-clamp method in both
the neural cell line and overexpression cell system treated with TC1. These
depolarizing currents are basically irreversible, but exhibit reversibility in
the form of repolarization upon membrane recycling. Here, for the first time, we
demonstrate that the mechanism of light-induced depolarization of TC1 results
from KV channel activity suppression related to the resting membrane
potential (Fig. 5) and the occurrence of PACS current with the loss of cell
membrane capacitance (Fig. 2, 3). In addition, direct real-time recording of
the current revealed that repolarization from depolarization was caused by TC1 internalization
through endocytosis, and insertion of new membrane material through exocytosis
(Fig. 6, 7). Notably, the membrane disk-shaped nanocarrier cpHDL enabled
efficient light-induced depolarization by localizing the triad to the outer
surface of the intact cell membrane (Fig. 1, 7). These results revealed a mechanism
by which the light-induced TC1 activity, localized precisely at the cell
membrane, caused a decrease in the plasma membrane resistance and subsequently sustained
depolarization by KV suppression and generation of PACS current. In
addition, the intact cells restored KV and PACS current through
membrane recycling and achieved repolarization. We have previously observed
that CS molecules, which do not have the same reactive oxygen species and
cytotoxicity as TC1, have neural firing after photo-induced depolarization in
cultured rat hippocampal neurons [26]. These findings suggest that the
application of TC1 in combination with cpHDL can be used more efficiently and
safely in biological applications. Biological membranes play
multiple physiological roles, such as physical barriers for ions and solutes,
regulators of membrane protein function, and mediators of signal transduction.
In particular, ion channel activity is moderately affected by substances that
directly and indirectly affect cell membrane-protein interactions (polyunsaturated
fatty acids such as arachidonic acid and DHA), and amphiphiles such as Triton X
and capsaicin [27, 28]. Our results indicate that the endogenous and exogenous
expression of voltage-gated K+ channel activity is completely abolished,
regardless of the molecular species. Thus, TC1 does not bind directly to the
channel and is unlikely to stimulate specific membrane signaling. Indeed, the
results in Fig. 6b, 6d, and 6e show that the K+ channel function was
not impaired for ~1 hour after 4 min of TC1 stimulation. However, the K+
channel activity was restored indirectly by regeneration of cell membranes
using the cytosis pathway. This is consistent with previous reports that such
membrane lipids may affect the K+ channel activity [29, 30]. Our
results showed that voltage-dependent modulation of K+ channel
activity was not modulated, but was inactivated because it did not respond to
voltage pulses after light-induced TC1 activation. Thus, the observations seen
in the ion channel inactivation from the lipid quality changes are consistent
with those of the gramicidin channel inactivation caused by oxide damage [31]. Our
results showed that TC1 had no effect on membrane pore formation and recycling
mechanism (Fig. 7), but had a significant effect on ion transport mechanism. An
investigation of the properly tuned control of ion channel activity requires
further study of the interaction between membrane lipids and ion channels. Previous research on cell
control using light-induced nanomaterials overlooked the possibility of
membrane current generation from changes in the membrane quality [3]. Therefore,
we considered the possibility of PACS currents through the membrane. The most probable cause of a
decreased membrane resistance is the ion channel activity. However, our
observations showed no effect from the broad cation and anion channel inhibitor
application (Fig. 3a–d), so the PACS current is unlikely to be the current
through the ion channel. Second, CS molecules have been reported to generate
very large voltages (~106 V/cm; 500 mV/5 nm) upon photostimulation [32].
Extremely high voltages on the membrane can reduce membrane resistance by
forming pores in the membrane, as seen in electroporation technology [33]. Our
results, as shown in Fig. 7c and 7d, show that very small molecules such as LDH
(MW: 144 kDa) and PI (MW: 668.4 g/mol) do not pass through the membrane after
TC1 stimulation (Fig. 7c and 7d). Thus, TC1 activity is unlikely to cause
physical damage. Furthermore, TC1 has not been detected to generate reactive
oxygen species [12], and is therefore unlikely to be PACS current from membrane
damage. Third, the possibility of changing the cell membrane properties can be
understood by applying the idea of a dielectric breakdown, which is the basis
of physics. The leakage current magnitude of a capacitor that has caused a
dielectric breakdown is proportional to the damaged membrane capacitance (i.e.,
Ileak ∞ Cd = ε / td;
Ileak, leakage current; Cd, damaged
capacitance; ε, dielectric constant; td, damaged membrane
thickness). In our results, a 24.7% capacity loss was observed from the TC1
activity induced by photostimulation, which may be accompanied by PACS current
(Fig. 3e). As described above, because the TC1 activity does not damage the
cells, it is plausible that the PACS current occurs when the relative
dielectric constant increases. Indeed, the dielectric constant of the damage was
around 2 [34], but the previously reported dielectric constant of CS molecules
of the same type as TC1 was ~4 in amphiphilic solvents [26]. Therefore, it is
likely that TC1 activated in the cell membrane increased the cell membrane
dielectric constant and lowered the membrane resistance. Recently, an interdisciplinary
study of physics and biology by Bezanilla and colleagues elucidated the
mechanism by which a local temperature rise with near-infrared lasers caused
instantaneous changes in the capacitance and leakage current [35]. The most
recently, a research group by Lanzani and Benfenati has developed a method in
which millisecond pulses of visible light induce transient hyperpolarization
followed by a delayed depolarization that triggers action potential firing in
neurons [36]. The administration of a photosensitive azobenzene compound to
cells can be stimulated for 7 days without directly affecting the ion channels
and local temperature. Although Near-infrared treatment and light-sensitive
azobenzene treatment are very effective when applied to short-term neuronal
cell excitement, our technique is that 5 minutes light irradiation of TC1-cpHDL
can maintain cell excitement for 1 hour. It also has the advantage with regard
to phototoxicity owing to drug metabolism. On the contrary, it is not suitable
for treatment limited to short-term nerve stimulation. Therefore, it has
potential medical applications in the regulation of autonomic excitability and
endocrine cell secretion controlled by persistent excitability [37-40]. It is
also important to note that the differentiating targets that can be stimulated
in a cell-specific manner by improving cpHDL drug delivery system are
candidates for advanced alternative tissue excitation techniques. The integrity of cell membranes
and membrane proteins depends on the endocytic and exocytotic pathways [41].
Although the membrane transport mechanisms of ion channels (including K+
channels) have been demonstrated mainly through imaging and biochemical
techniques, the direct real-time evaluation was inadequate [42]. Our results
show that the assessment of ion channel activity during membrane recycling can
be directly performed electrophysiologically in one hour in real time, which is
the first study of this kind to our knowledge. Our CS-cpHDL also caused changes
in membrane function on the target cells with light-induced timing. Therefore, the
use of TC1 can be expected as a response tool for membrane research, such as
autophagy and interaction analysis between membrane transport proteins and
lipids. Because the use of cpHDL
improved the degree and rate of depolarization, improved cpHDL will certainly
result in greater efficiency. Therefore, the use of recently developed cpHDL [43]
that can achieve the efficient and appropriate targeted transport of CS
molecules (such as cell membrane delivery efficiency) may contribute to the
development of more efficient light-induced depolarization systems. For
example, the absorption spectrum of TC1 solubilized with liposomes was reported
to be similar to that in H2O containing 10% DMSO, and their
extinction coefficient at the excitation wavelength was also comparable [12].
Under this condition, the charge-separation quantum yield was decreased by
>50% in the presence of liposomes, which could be accounted for by intermolecular
electron transfer and self-quenching caused by unfavorable aggregation. In the
same report, the number of TC1 molecules per cell was determined to be 1.9 × 106
upon its direct addition to cell culture media, which was one-tenth of 2–3 × 107
molecules/cell for the delivery by cpHDL [13]. In order to improve the charge
separation yield, bioapplicable TC1s need to be improved by strategies that
solve the problems of intermolecular aggregation and number of molecules in the
cell membrane. Experiments investigating the
effect of TC1 on current with a voltage-clamp showed that most K+
currents were suppressed during illumination of TC1-treated cells (Fig. 2).
However, these results were lower than the expected K+
channel-dependent depolarization values in response to the same TC1 treatment
in current-clamp mode (Fig. 1). This value of depolarization suggests the
involvement of Na+-K+ pumps [44] and Cl-
channels [45, 46] that form the resting membrane potential, and the effects of
TC1 on the activity of these targets require further study. Further research is needed to
better characterize the depolarization and repolarization mechanisms of TC1
photostimulation, and to explore how that knowledge could enhance the use of
this technology in vivo. For example, it would be interesting to model
the effect of TC1 on intramolecular lipid dipole moments at the molecular level
and the interaction of lipids experienced by voltage-gated K+
channels. It may also help to understand the membrane properties of various cell
types expressed in vivo and analyze whether they mediate downstream
physiological effects. Herein, we demonstrated that TC1 photostimulation
affects the membrane potential via ion transport by altering cell membrane properties,
including cell membrane resistance. Our discovery supports the application of
this unique optical technology as a tool in physics, biology, pharmacology and
medicine. Conclusion The ability to non-invasively
alter cell membrane excitability, via unique light-controlled depolarization
applications, has attractive therapeutic translation potential. The
light-induced regulation of membrane potentials with drug delivery systems is
fascinating as a strategy for promoting research on the biological processes of
excitable cells, and for the spatiotemporal control of the nervous system and
muscle activity. Our findings concluded that depolarization of the CS molecule
simultaneously inactivates the voltage-dependent potassium currents and
produces PACS currents. This activity continues to depolarize target cells, but
is reversible via a regeneration mechanism because the cell membrane is intact.
Unraveling the underlying mechanism of cellular photoexcitation highlights the
generality of membrane excitability regulation and potential medical
applications to autonomic nerves and secretory organ that exert physiological
function by continuous excitation. Acknowledgements We thank Mr. Masahiro Ohara
(National Institute for Physiological Science) for helpful discussion. We thank
Rosalie Tran, PhD, from Edanz Group
(https://en-author-services.edanzgroup.com/) for editing a draft of this
manuscript. Author contributions T.N. conducted all experiments
and analysis. R.F. and T.M. conducted the CS molecule and drug delivery system
experiments. H.M. and Y.K. performed the PCR experiments. K.S-N. assisted in
the culture design, imaging analysis, and discussion of the data. H.I. and T.M.
discussed and commented on the draft. T.N. conceived and designed the work, and
wrote the manuscript. Funding Sources This work was supported in part
by Grants in-Aid for Scientific Research (KAKENHI) from the Japan Society for
the Promotion of Science and the Ministry of Education, Culture, Sports,
Science (No. 15K08197 and 18K06864), and Central Research Institute of Fukuoka
University (No. 177009). Disclosure
Statement The authors have no conflicts of
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