Probing Intracellular Potassium Dynamics in Neurons with the Genetically Encoded Sensor lc-LysM GEPII 1.0 in vitro and in vivo

doi:10.21203/rs.3.rs-3715395/v1

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Probing Intracellular Potassium Dynamics in Neurons with the Genetically Encoded Sensor lc-LysM GEPII 1.0 in vitro and in vivo

https://doi.org/10.21203/rs.3.rs-3715395/v1

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published 14 Jun, 2024

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Neuronal activity is accompanied by a net outflow of potassium ions (K⁺) from the intra- to the extracellular space. While extracellular [K⁺] changes during neuronal activity are well characterized, intracellular dynamics have been less well investigated due to lack of respective probes. In the current study we characterized the FRET-based K⁺ biosensor lc-LysM GEPII 1.0 for its capacity to measure intracellular [K⁺] changes in primary cultured neurons and in mouse cortical neurons in vivo. We found that lc-LysM GEPII 1.0 can resolve neuronal [K⁺] decreases in vitro during seizure-like and intense optogenetically evoked activity. [K⁺] changes during single action potentials could not be recorded. We confirmed these findings in vivo by expressing lc-LysM GEPII 1.0 in mouse cortical neurons and performing 2-photon fluorescence lifetime imaging. We observed an increase in the fluorescence lifetime of lc-LysM GEPII 1.0 during periinfarct depolarizations, which indicates a decrease in intracellular neuronal [K⁺]. Our findings suggest that lc-LysM GEPII 1.0 can be used to measure large changes in [K⁺] in neurons in vitro and in vivo but requires optimization to resolve smaller changes as observed during single action potentials.

Potassium ions (K⁺) are the most abundant intracellular cations in most cell types and are involved in an array of vital processes both on the cellular and supracellular level¹. These processes notably include the diffusion or transport of K⁺ across membranes to form an electrochemical gradient. Together with the electrochemical gradients of other ions, it makes up the membrane potential and can be utilized both as energy storage and for intracellular signaling².

Dynamic shifts of intracellular K⁺ concentrations are fundamental during signal transduction by neurons, which transmit information via action potentials. After the initial depolarization phase of an action potential, the neuronal membrane is repolarized by the outflow of K⁺ through voltage-gated K⁺ channels or K⁺ leak channels to allow a new action potential to occur^3,4. Therefore, the dynamics and the regulation of K⁺ ions play a crucial role in neurotransmission, and changes in brain K⁺ homeostasis are known to influence neuronal excitability^5,6 as well as action potential shape^7,8. In addition, restoration of the ion gradients after activity, mainly achieved via the Na⁺/K⁺-ATPase, is one of the primary energy-consuming processes in neurons, making ion homeostasis a key player in brain energetics and its regulation^9,10. Perturbed K⁺ dynamics or mutations of K⁺ channels are directly responsible for many pathological conditions, such as epilepsy^11,12 and migraine^13,14. In contrast, other disorders, such as ischemic stroke, are often exacerbated or accompanied by perturbed K⁺ homeostasis¹⁵.

So far, technical limitations in measuring K⁺ fluxes in neurons have hampered our understanding of neuronal potassium dynamics. While electrode-based potassium measurements are fast and accurate, they are invasive, can measure K⁺ only very locally, and are primarily used to assess extracellular K⁺ dynamics^16,17. Imaging methods such as MRI^18,19 or live cell imaging with K⁺ sensitive dyes²⁰ can potentially overcome these limitations; however, they lack (sub-)cellular resolution or specificity.

The development of genetically encoded fluorescent sensors for K⁺ ions holds great potential for overcoming those limitations, as they can be expressed in a cell-type-specific manner or with subcellular localization. In addition, they allow observation of K⁺ dynamics in single neurons and whole tissues non-invasively and can be measured with high temporal resolution. We and others²¹, recently developed novel FRET-based genetically encoded K⁺ ion indicators (GEPIIs) with high specificity. While these biosensors may help to understand K⁺ dynamics in the brain, they have been mainly tested in non-neuronal cells and have not been characterized neither in vitro nor in vivo for their feasibility to measure K⁺ dynamics in neurons.

Therefore, the aim of the current is to characterize the capacity of GEPIIs to observe K⁺ changes during neurotransmission, both in vitro and in vivo. To achieve this goal, we expressed the potassium sensor lc-LysM GEPII 1.0 in primary cultured neurons or in the cortex of living mice. We then recorded K⁺ signals of individual neurons during either spontaneous or evoked neuronal activity to assess the degree to which lc-LysM GEPII 1.0 can resolve K⁺ dynamics during neuronal activity.

Plasmids and cloning

We used plasmids carrying the genetically encoded potassium sensor lc-LysM GEPII 1.0 with different subcellular targeting sequences as previously described²¹. The sensor was targeted either to the cytosol (cyto), the plasma membrane (SubPM), or the mitochondria (mito). To achieve AAVmediated expression, we subcloned the ORFs encoding the different variants of lc-LysM GEPII 1.0 in AAV backbones under the control of a CAG promoter for ubiquitous expression (pAAV-CAG-GFP, Addgene #37825), a hSyn promoter for neuronal expression (pAAV-hSyn-eGFP, Addgene #50465) or a GFAP promoter for astrocytic expression (pAAV.GFAP.iGABASnFr, Addgene #112172). All three backbones were linearized by cutting out the existing transgenes using BamHI and HindIII restriction sites. These restriction sites were added to cyto-lc-LysM GEPII 1.0 and SubPM-lc-LysM GEPII 1.0 via overhang PCR to insert them directly via subcloning. Since mito-lc-LysM GEPII 1.0 contains a BamHI restriction site, this transgene was inserted using the Gibson Assembly method. After cloning, DNA was transformed into competent DH5α E. coli bacteria, colonies were screened via colony PCR and candidate clones were confirmed by sequencing before amplification and AAV production.

pAAV.Syn.NES-jRCaMP1b.WPRE.SV40 was a gift from Douglas Kim & GENIE Project (Addgene plasmid # 100851; http://n2t.net/addgene:100851; RRID: Addgene_100851). For bacterial expression of jRCaMP1b, the construct was subcloned into EKAR2G_design1_mTFP_wt_Venus_wt vector (Addgene plasmid # 39813) using the restriction sites BamHI and XhoI.

For imaging the concentration of cytosolic calcium ions in vitro, we used pGP-AAV-syn-jGCaMP8m-WPRE, a gift from the GENIE Project (Addgene plasmid #162375)²². For optogenetic stimulation, we used pAAV-Syn-ChrimsonR-tdT, a gift from Edward Boyden (Addgene plasmid #59171)²³.

Protein purification:

Bacterial expression plasmids containing lc-LysM GEPII 1.0 and jRCaMP1b were transformed into Rosetta (DE3) competent cells. For the selection of lc-LysM GEPII 1.0 and jRCaMP1b positive cells, kanamycin and ampicillin were used, respectively. For purification of the biosensors, positive clones were inoculated in 5 mL LB with appropriate antibiotic, and after 8 hours, the cultures were transferred into 250 mL LB plus antibiotic and incubated at 37°C. At an OD value between 0.4 and 0.6 protein expression was induced using IPTG with a final concentration of 0.5 mM and further incubated for 16 hours at 18°C. Purification of the 6x histidine-tagged biosensors was performed using gravity-based Ni-NTA affinity chromatography method as described elsewhere²⁴. Purified proteins were concentrated using Ultracel® Regenerated Cellulose (30kDa MWCO) Amicon tubes and kept at -80°C for further usage. The functionality of the purified biosensors was tested using a SpectraMax i3 Multi-Mode Microplate Reader. Samples were loaded on a 96-well plate with a solid black bottom. The intensiometric calcium biosensors jRCaMP1b were excited at 535/25 nm and emission was collected at 595/35 nm. The FRET-based biosensors lc-LysM GEPII 1.0 were excited with 430/9 nm and emission was collected at 485/9-535/15 nm, respectively.

AAV Production

Adeno-associated viral particles were produced in HEK293T cells grown in DMEM supplemented with 10% FBS and 100 U/mL Penicillin-Streptomycin in a humidified incubator at 37°C/5% CO2. Cells were grown to 70–80% confluency and triple transfected with pHelper, pAAV-DJ (both from Cell Biolabs, Cat: VPK-400-DJ) and a pAAV-ITR-vector carrying the transgene. Triple transfection was achieved using polyethylenimine (PEI) titrated to pH 7.0. Two to three days after transfection, cells were detached using 1/80 of the culture volume of 0.5 M EDTA in PBS pH 7.4 and AAV particles were extracted using the AAVpro® Purification Kit (All serotypes) from Takara Bio Inc. (Cat: #6666). Cells were centrifuged at 1700 g for 10 min at 4°C and the supernatant was discarded. The resulting cell pellet was lysed by vortexing it with 650 µL of AAV Extraction Solution A plus. Subsequently, cell debris was pelleted at 14 000 g for 10 min at 4°C and the supernatant was collected in a new tube. Finally, 65 µL of AAV Extraction Solution B was added, and the viral solution was aliquoted and stored for further use at -80°C. AAV titration was performed by qPCR using the AAVpro® Titration Kit (for RealTime PCR) Ver.2 from Takara (Cat: #6233) according to the manufacturer’s instructions. Titration was performed using primers that annealed in the ITR repeats of the viral backbone (ITR F: GGAACCCCTAGTGATGGAGTT and ITR R: CGGCCTCAGTGAGCGA).

Preparation of cryo-stocks of mixed cortical cultures

Cryopreserved mixed cortical cell culture stocks were prepared from E17 embryos of Sprague Dawley rats. Brains were removed and immediately placed in ice-cold HBSS supplemented with 7 mM HEPES pH 7.4. Cortices were dissected, meninges were removed, and the tissue was cut into small pieces with a scalpel. Then the tissues from all embryos of one litter were digested in HBSS supplemented with 0.5% Trypsin and 10 µg/mL DNAse I for 15 min at 37°C. Digestion was stopped by the addition of MEM supplemented with 10% FBS, the tissue was washed twice with HBSS and subsequently triturated using a glass Pasteur pipette coated with 4% BSA in HBSS. After counting the concentration of cells in the suspension, the cell suspension was diluted with MEM supplemented with 10% FBS to 2 million cells per mL. Finally, DMSO was added to a volume fraction of 10% and the solution was aliquoted. Aliquots were cryopreserved by placing them in a freezing container filled with isopropanol at -80°C over night. The next morning, the aliquots were transferred into a liquid nitrogen tank for longterm storage.

Mixed cortical cell cultures from cryo-stocks

All cell culture reagents were purchased from Gibco. Mixed cortical cultures were plated on 15 mm round glass cover slips coated with PolyDLysine at a density of 100,000 cells per cover slip. A cryopreserved aliquot was thawed at 37°C and diluted in an appropriate amount of culture medium (Neurobasal-A medium, no D-glucose, no sodium pyruvate supplemented with 1x B27, 10 mM glucose, 2 mM GlutaMAX™, 1 mM sodium pyruvate and 100 U/mL Penicillin-Streptomycin). The cell suspension was plated on the coverslips placed in 12-well plates and maintained in a humidified incubator at 37°C/5% CO₂. Half of the culture medium was replaced twice a week and cells were used after 22–24 days. For transgene expression, cells were transduced 3–4 days before the experiment with the appropriate AAV at a MOI of 1000.

Immunostainings

For immunocytochemistry, mixed cortical cultures were washed with PBS and subsequently incubated with 4% paraformaldehyde (PFA)/4% sucrose in PBS for 15 min at room temperature (RT). Fixed cells were washed three times with PBS, incubated with 50 µM NH₄Cl in PBS for 10 min at RT, followed by another three washes with PBS. Fixed cells were then permeabilized using 0.1% TritonX in PBS for 3 min at RT and blocked with blocking buffer (0.2% BSA, 0.2% FCS, 0.02% fish-skin gelatin in PBS) for 1 h at RT. Subsequently, cultures were incubated with primary antibodies against gp-NeuN (Synaptic Systems, 266 004) and mouse-GFAP (Cell Signaling, #3670) diluted in blocking buffer for 2 h at RT. Subsequently, primary antibodies were removed, cells were washed 3 times with PBS for 5 min and then incubated with anti-guinea pig- AlexaFluor® 647 (Jackson, 706-606-148) and anti-mouse AlexaFluor® 594 (Jackson, 715-586-150) diluted in blocking buffer for 1 h at RT. During the last 5 minutes of this step, DAPI (5 µg/mL) was added to stain the nuclei. Finally, coverslips were washed three times with PBS for 5 min each and mounted on microscopy slides. Cells were imaged at a confocal microscope.

For immunohistochemistry, animals virally transduced with AAV-hSyn-lc-LysM GEPII 1.0 were anesthetized with MMF and transcardially perfused with PBS followed by 4% PFA in PBS until the liver was devoid of blood. Brains were extracted, post-fixed in 4% PFA in PBS overnight and then stored in PBS at 4°C until further use. Brains were mounted in 4% agarose and sectioned using a vibratome to create 100 µm thick brain slices. PFA-fixed brain sections were incubated in a blocking and permeabilizing primary antibody buffer solution (1% BSA, 0.1% fish-skin gelatin, 0.1% Triton X-100, 0.05% Tween 20 in PBS) with rabbit anti-NeuN (Abcam, ab177487) antibodies at 1:100 dilution on a rotary shaker at 4°C for 2–3 days. Sections were then washed in PBS three times for 30 minutes and incubated with a secondary antibody buffer mix (0.2% BSA, 0.2% FCS, 0.02% fish-skin gelatin in PBS) containing 1:300 anti-rabbit AlexaFluor® 647 (Jackson, 711-606-152) antibody at 4°C on a rotary shaker for two days. Sections were then washed three times in PBS and during the last washing step, DAPI was added at a concentration of 5 µg/mL for 30 minutes to stain the nuclei. Washing using PBS was repeated three times for 30 minutes prior to mounting the sections on glass coverslips. Brain section overviews were imaged with a confocal microscope using a 10x air objective.

Live cell imaging

For live cell imaging of mixed cortical cultures expressing either GCaMP8m or lc-LysM GEPII 1.0, coverslips with the cultures were transferred in an open imaging chamber and placed on an inverted microscope. The microscope was equipped with a 20x air objective (NA 0.8), an LED light source, an emission image splitter, and a CCD camera. Cells were constantly superfused with aCSF (in mM: 125 NaCl, 2.5 KCl, 1.25 NaH₂PO₄, 26 NaHCO₃, 1 MgCl₂, 1.25 CaCl₂, 2 glucose, 0.5 sodium lactate, 0.05 sodium pyruvate) gassed with 5% CO₂/95% air to maintain a stable pH at room temperature. GCaMP8m was excited with 10 Hz at 469 ± 19 nm and emission was collected with a bandpass filter at 525 ± 25 nm. For FRET imaging of lc-LysM GEPII 1.0, cells were excited at 436 ± 10 nm and emitted light was split at 515 nm on the camera for simultaneous recording of mseCFP and cpV using a dichroic mirror (t515lp, Chroma). Emission of the individual channels was collected using bandpass filters (480 ± 15 nm for mseCFP, 535 ± 15 nm for cpV). Depending on the experiment, cells expressing lc-LysM GEPII 1.0 were excited with frequencies ranging from 0.2 to 10 Hz. Exposure time, LED power, and excitation frequencies were adjusted to minimize bleaching and phototoxicity while still obtaining sufficient signal-to-noise ratio. The remaining bleaching was corrected using a customwritten Python script and the data was represented as FRET ratio cpV/mseCFP.

Optogenetic stimulation

For optogenetic stimulation, we co-transduced cultures with AAV-Syn-ChrimsonR-tdT and either AAV-CAG-lc-LysM GEPII 1.0 or AAV-Syn-GCaMP8. Optogenetic stimulation was achieved by placing a red LED (M617L4, Thorlabs) over the cells during imaging using a custom-made 3Dprinted holder. To maximize stimulation strength, the emitted light was collected using an aspheric condenser lens. A custom-made microcontroller-gated LED driver allowed programmable and precise light pulses for stimulation. Cells were stimulated with trains of 10 ms light pulses at 1 Hz, allowing enough time in between stimulation trains for complete recovery of the signal.

To avoid imaging artifacts from the stimulation light, a 590 nm long pass filter was placed in front of the LED and additional 550 nm short pass filters were added in the emission light path. The intensity of the excitation light for either GCaMP8m or lc-LysM GEPII 1.0 was minimized to avoid cross-stimulation of ChrimsonR. If the minimal LED power led to neuronal stimulation, it was further reduced using a neutral density filter OD 1.3.

Animals

Two to three months old male C57BL/6J mice were used. The animals were group-housed under pathogen-free conditions and bred in the animal housing facility of the Center of Stroke and Dementia Research, with food and water provided ad libitum (21 ± 1°C, at 12/12-hour light/dark cycle). All experiments were carried out in compliance with the ARRIVE guidelines, the National Guidelines for Animal Protection, Germany and with the approval of the regional Animal Care Committee of the Government of Upper Bavaria.

Cranial window implantation and stereotactic virus injection

Before use, surgical tools were sterilized in a glass-bead sterilizer (Fine Science Tools). Mice were anesthetized by an i.p. injection of medetomidine (0.5 mg/kg), midazolam (5 mg/kg), and fentanyl (0.05 mg/kg). Subsequently, mice were placed onto a heating blanket (37°C), and the head was fixed in a stereotactic frame. Eyes were protected from drying by applying eye ointment. The scalp was washed with swabs soaked with 70% ethanol. A flap of skin covering the cranium was excised using small scissors. The periosteum was scraped away with a scalpel. The prospective craniotomy location over the somato-sensory cortex was marked with a biopsy punch (diameter 4 mm). The exposed skull around the area of interest was covered with a thin layer of dental acrylic (iBond Self Etch, Heraeus Kulzer) and hardened with an LED polymerization lamp (Demi Plus, Kerr). A dental drill (Schick Technikmaster C1, Pluradent) was used to thin the skull around the marked area. After applying a drop of sterile phosphate buffered saline on the craniotomy, the detached circular bone flap was removed with forceps. Subsequently, 0.5 µl of a virus suspension (AAV.PhPeb-Syn.NES- lc-LysM GEPII 1.0, Vectorbuilder and AAV.9-Syn.NES-jRCAMP1b, Addgene #100851-AAV9 mixed 1:1 to a final concentration of 1x10¹² vg/ml) was injected via a glass capillary and Nanoliter 2020 Injector (World Precision Instruments) at a speed of 50 nl/min into the somatosensory cortex at the following coordinates: -1.5 mm rostrocaudal, -1.5 mm lateral and − 0.25 mm dorsoventral. A circular coverslip (4 mm diameter) was placed onto the craniotomy and glued to the skull with histoacryl adhesive (Aesculap). The exposed skull was covered with dental acrylic (Tetric Evoflow A1 Fill, Ivoclar Vivadent), and a head-post was attached parallel to the window for head-fixing mice in subsequent imaging sessions. After surgery, mice received a s.c. dose of the analgesic Carprophen (7.5 mg/kg body weight). Anesthesia was antagonized using Atipamezol (2.5 mg/kg), Naloxone (1.2 mg/kg), and Flumazenil (0.5 mg/kg) i.p. Finally, mice were allowed to recover in a 35°C warming chamber until full recovery. In vivo imaging was commenced three to four weeks after surgery.

Mouse cerebral ischemia model

For remote occlusion of the middle cerebral artery (MCAo) in mice²⁵ the left common carotid artery was exposed and the superior thyroid artery branching was cauterized. The common carotid artery was ligated and the external carotid artery (ECA) was fully occluded. A filament with a diameter of 0.21 ± 0.02 mm (6021PK10, Doccol Corporation, Sharon, MA, USA) was inserted through as small incision into the ECA and advanced for 5 mm. A custom-made occlusion filament was inserted into the ECA while the Doccol filament was simultaneously removed. The filament was then further advanced along the internal carotid artery towards the middle cerebral artery (MCA). After placing the under the 2-photon microscope the MCA could be occluded remotely. MCA occlusion (MCAo) caused a cortical infarct which induced periinfarct depolarizations waves (PIDs) within the surrounding tissue.

In vivo two-photon microscopy

In vivo two-photon imaging was performed three weeks after cranial window implantation using a Leica SP8 DIVE 2-photon microscope equipped with a fs-laser, a 25x water immersion objective and a motorized stage. lc-LysM GEPII 1.0 and jRCaMP1b were co-excited at 850 nm and 1045 nm, respectively, and the emission was collected at 450–500 nm and 575–625 nm with non-descanned detectors placed directly behind the objective. Laser power below the objective was kept around 50 mW to minimize phototoxicity. Throughout the imaging session, mice were anesthetized with isoflurane (1% in oxygen, 0.5 l/min) and kept on a heating pad to maintain body temperature at 37°C. XY time-lapse series (7.51 Hz) with 1 µm axial resolution and 128 x 128 pixels per image frame of lc-LysM GEPII 1.0 and jRCaMP1b expressing neurons were subsequently recorded at a depth of 150–200 µm underneath the cortical surface. After 5 min of baseline recordings, periinfarct depolarizations were induced via remote MCAo and recordings were continued for further 10 min. Fluorescence lifetime measurements were subsequently analyzed in LAS X (Leica).

After confirming that we were able to successfully express lc-LysM GEPII 1.0 in a cell type-specific manner and to target different subcellular compartments (see supplementary Fig. 1), we explored the potential for resolving K⁺ changes during neuronal activity by live cell imaging. Ca²⁺ dynamics were measured in parallel experiments using GCaMP8m to monitor neuronal activity. lc-LysM GEPII 1.0 fluorescence was measured in rat primary mixed cortical cultures containing neurons and astrocytes.

lc-LysM GEPII 1.0 resolves [K⁺] decreases in response to tetanic neuronal stimulation in vitro

During neuronal activity, neurons depolarize due to the inflow of sodium (Na⁺) and calcium ions (Ca²⁺) and repolarize due to the outflow of K⁺. Our goal was to investigate if and to what extent we could observe this K⁺ outflow during neuronal activity using lc-LysM GEPII 1.0. Therefore, as a proofofprinciple experiment, we induced neuronal hyperactivity in our mixed cortical cultures by inhibiting inhibitory neurons with Bicuculline, a selective GABA_A receptor antagonist. Ca²⁺ imaging confirmed that Bicuculline induced a strong, epileptic-like neuronal firing pattern. The addition of Bicuculline for 50 seconds led to an immediate increase of the intensiometric signal of GCaMP8m, which peaked within seconds (Fig. 1A and C). After Bicuculline was washed out, the Ca²⁺ signal recovered to baseline. When we applied the same stimulus (50 µM Bicuculline for 50 seconds) to cultures expressing lc-LysM GEPII 1.0, we observed a pronounced decrease of the FRET ratio signal by 12.7 ± 3.4%, indicating a reduction in the intracellular K⁺ concentration (Fig. 1B and D). The FRET ratio signal of the K⁺ biosensor continued to decrease even after removing Bicuculline and reached a minimum after 3.9 ± 0.4 minutes before it slowly returned to baseline. These data show that Bicuculline massively reduces cytosolic K⁺ levels and demonstrates the suitability of the FRET-based biosensor to monitor K⁺ changes in response to neuronal hyperactivity.

Measuring lc-LysM GEPII 1.0 fluorescence during spontaneous neuronal activity in cultured neurons

After confirming that we can resolve cytosolic [K⁺] alterations during intensive neuronal activity associated with large changes in intracellular ion concentrations, we aimed to investigate if we could observe K⁺ changes during spontaneous neuronal activity characterized by mainly single action potentials. Ca²⁺ imaging using GCaMP8m showed that our cultures displayed spontaneous neuronal activity, which was completely abolished in the presence of tetrodotoxin (TTX), a potent inhibitor of voltage-gated sodium channels (Fig. 1E). When measuring lc-LysM GEPII 1.0 fluorescence, the obtained recordings did not display any apparent changes of the FRET ratio signal (Fig. 1F). When inhibiting neuronal activity using TTX, the FRET ratio traces were indistinguishable from the ones before TTX application (Fig. 1F). To assess whether the lc-LysM GEPII 1.0 FRET ratio signals were due to neuronal activity or technical noise, we hypothesized that changes in the FRET ratio signal in response to actual K⁺ fluctuations should be independent of signal intensity, while FRET changes due to noise should correlate with signal intensity. To test this hypothesis, we plotted the standard deviation (SD) of the full calcium traces against the average intensity of GCaMP8m. Indeed, our analysis showed no correlation between SD and signal intensity in spontaneously firing neurons (Fig. 1G, gray symbols) indicating that neuronal activity induced changes in the Ca²⁺ signal disturb the correlation between SD and intensity, while in neurons silenced with TTX these parameters correlated well (Fig. 1G, orange symbols) suggesting that the recorded signal represented background noise. Performing the same analysis for signals derived under the same conditions from neurons expressing lc-LysM GEPII 1.0, we observed no changes between the control condition, i.e., when neurons fired spontaneously (Fig. 1H, gray symbols) and the time when spaontaneous neuronal activity was inhibited by TTX (Fig. 1H, orange symbols). These data indicate that neuronal activity did not contribute to the FRET ratio signal changes of lc-LysM GEPII 1.0., i.e., the recorded signal was background noise. These findings suggest that lc-LysM GEPII 1.0 is not able to measure changes in intracellular [K⁺] elicited by single action potentials or, alternatively, that single action potentials may not result in global reductions of cytosolic [K⁺].

lc-LysM GEPII 1.0 signals following optogenetic stimulation of neurons

Having identified that lc-LysM GEPII 1.0 can resolve K⁺ decreases in response to multiple but not to single action potentials, we wanted to investigate the threshold above which lc-LysM GEPII 1.0 is able to resolve detectable [K⁺] changes in neurons. We combined optogenetic neuronal stimulation with FRET ratio imaging of lc-LysM GEPII 1.0, allowing us to control neuronal activity during live imaging of [K⁺] changes. We chose the red-shifted optogenetic tool ChrimsonR to avoid cross-stimulation with the excitation light during imaging and co-expressed it with either GCaMP8m or lc-LysM GEPII 1.0 (Fig. 2A). We decided to stimulate our mixed cortical cultures using low-frequency stimulation trains with increasing pulses to investigate the minimal activity required for detecting K⁺ changes using lc-LysM GEPII 1.0. We chose 10 ms as the individual pulse length as this was the shortest duration reliably eliciting a Ca²⁺ signal during imaging with GCaMP8m (see supplementary Fig. 2). We stimulated neurons with up to 120 light pulses and allowed the signal to return to baseline before triggering the next stimulation train. Each stimulation triggered an individual peak of [Ca²⁺]_i as evidenced by imaging with GCaMP8m (Fig. 2B, green traces); when measuring potassium using lc-LysM GEPII 1.0 we could observe a decrease of FRET ratio signals (normalized FRET change: -0.63 ± 0.12%) only after a stimulation train of 30 pulses (Fig. 2B, purple traces and Fig. 2C), again confirming that multiple action potentials are necessary to decrease global cytosolic [K⁺] to levels detectable with lc-LysM GEPII 1.0. Increasing the number of pulses per stimulation train to 60 or 120 further reduced FRET ratio signals of the K⁺ biosensor (normalized FRET changes − 1.12 ± 0.37% and − 2.31 ± 0.93%, respectively) linearly (R² = 0.997).

As we expressed lc-LysM GEPII 1.0 in neurons and astrocytes, we could also examine the effect of neuronal stimulations on cytosolic [K⁺] in neighboring astrocytes. However, we did not monitor any significant changes in the FRET ratio signal of lc-LysM GEPII 1.0 in astrocytes, independent of the number of stimulations (Fig. 2D).

lc-LysM GEPII 1.0 allows measuring [K⁺]_i in neurons in vivo

After having characterized lc-LysM GEPII 1.0 in cultured cells, we wanted to investigate if we can also measure neuronal K⁺ dynamics in the brain of living mice by expressing hSyn-lc-LysM GEPII 1.0 and Syn-jRCaMP1b with AAV-based viral vectors in the cerebral cortex (supplementary Fig. 3). Since we observed changes of cytosolic [K⁺] only in response to intense neuronal activity in vitro, we expected a similar situation in vivo. To generate vigorous neuronal activity in vivo, we chose a model resulting in massive depolarizations of all neurons within a given spatial and temporal window. For this purpose, we induced focal cerebral ischemia by remote occlusion of the middle cerebral artery (MCAo; Fig. 3A), a condition well known to cause waves of massive neuronal depolarizations in the vicinity of ischemic tissue, so called periinfarct depolarizations (PIDs) while mice were placed under a 2-photon microscope for simultaneous recordings of jRCaMP1b and lc-LysM GEPII 1.0. Within minutes of MCA occlusion, PIDs were recorded in the mouse cerebral cortex as evidenced by increases in neuronal [Ca²⁺]_i spreading in cortical tissue with the expected velocity of about 2 mm/min (Fig. 3B). As a PID also elicits an increase in cerebral blood flow that may interfere with intensity-based fluorescent imaging approaches, we decided to measure jRCaMP1b and lc-LysM GEPII 1.0 using fluorescence lifetime imaging (FLIM), which is independent of intensity changes and therefore provides a robust readout. PIDs elicited a substantial increase in the fluorescence lifetime of jRCaMP1b by 111 ± 30% (Fig. 3C), crossing the field of view (Fig. 3D). Having now established in vivo FLIM microscopy, we used the same experimental protocol to check if we could measure [K⁺] changes during PIDs using lc-LysM GEPII 1.0. For this purpose, we measured the fluorescence lifetime of the donor mseCFP as a proxy of the FRET efficiency of the sensor. After induction of PIDs by cerebral ischemia, we recorded an increase in the fluorescence lifetime of the donor fluorescence of lc-LysM GEPII 1.0 by 5.9 ± 2.4% (Fig. 3E). When placing regions of interest in different parts of the field of view, we observed the wave-like character of the PID, confirming that the signal increase occurred in response to the PID (Fig. 3F). Since the fluorescence lifetime of lc-LysM GEPII 1.0 is inversely correlated with the K⁺ concentration, an increase in the lifetime corresponds to a decrease of neuronal [K⁺] (supplementary Fig. 4), as is expected during strong neuronal depolarization. As lifetime changes are not affected by intensity or potential volume changes during the PID, confounders are unlikely to account for this effect. When we correlated the normalized fluorescence lifetime values of lc-LysM GEPII 1.0 with those of jRCaMP1b, we could not observe any correlation between the two sensors before PID induction (R = -0.019, p = 0.437 - Fig. 3G). This confirms that we cannot resolve intracellular [K⁺] changes using lc-LysM GEPII 1.0 during spontaneous neuronal activity. However, an analogous correlation between the FLIM values for lc-LysM GEPII 1.0 and jRCaMP1b during the rising phase of the PID revealed a highly significant correlation (R = 0.359, p < 0.001 – Fig. 3G). The range of FLIM values of jRCaMP1b is considerably more extensive during PIDs, indicating that a very intense neuronal activation is required to elicit changes in [K⁺]_i big enough to be detected by lc-LysM GEPII 1.0. These data strongly suggest that we managed for the first time to record neuronal [K⁺] changes in the brain of living mice.

In the current study, we characterized the functionality of the FRET-based genetically encoded K⁺ sensor lc-LysM GEPII 1.0 in neuronal cells with a specific focus on measuring changes of [K⁺]_i during neuronal activation. We provide proof-of-principle evidence that lc-LysM GEPII 1.0 can resolve differences in [K⁺]_i during intense neuronal activity in cultured cells and in vivo. Changes of [K⁺]_i during single action potentials, as observed during spontaneous neuronal activity, could not be detected either in vitro or in vivo. However, we could successfully measure [K⁺]_i changes in the mammalian brain in vivo using the FRET-based K⁺ biosensor if massive neuronal activity was induced.

Brain potassium homeostasis is crucial for proper neuronal function, including controlling neuronal excitability or neurovascular and -metabolic coupling²⁶ and its disruption, e.g., through mutations in K⁺ channels are involved in severe neurological pathologies such as epilepsy¹² or migraine²⁷. Therefore, precise monitoring of K⁺ dynamics in brain tissue would be desirable to understand better K⁺ fluxes and how they are controlled. However, this understanding has been hampered by technical challenges.

The emergence of genetically encoded sensors for potassium^21,28–30 sparked hope that imaging approaches can help answer questions that could not be tackled with other methods^5,31,32. However, applications of genetically encoded K⁺ sensors to record intracellular neuronal potassium dynamics are sparse. One possible reason is that available sensors have a very high affinity for K⁺ and are saturated when expressed in neurons, cells which have a very high intracellular K⁺ concentration of around 140 mM⁹. For the current study, we chose lc-LysM GEPII 1.0, which, at the start of the study, had the highest K_d of all available sensors (i.e., 27 mM in vitro and 60 mM in cells^21,28). In the meantime, a new generation of K⁺ biosensors optimized for intracellular recordings has been published, yet they still need to be investigated in neurons³⁰.

When we induced hyperactivity in our neuronal cultures using Bicuculline or intensely stimulated these cells optogenetically (at least 30 times at 1 Hz), we observed a decrease of the FRET ratio of lc-LysM GEPII 1.0, indicating a reduction in neuronal [K⁺]_i. This is expected, as K⁺ outflow is the main contributor to neuronal repolarization after an action potential and was also corroborated by all so far published studies measuring intracellular K⁺ dynamics in neurons using genetically encoded sensors independent of the stimulation paradigm^28,29,33. Before concluding that lc-LysM GEPII 1.0 indeed measures [K⁺]_i one has to consider that next to changes in [K⁺]_i strong neuronal activity also leads to an intracellular acidification by up to 0.3 pH units^34,35. Since genetically encoded sensors are based on fluorescent proteins prone to pH-induced changes, intracellular pH shifts may result in apparent changes in [K⁺]_i. Wu and colleagues suggested that the decrease of the signal of the K⁺ sensor GINKO2 was, to a large degree, caused by acidification of the neuronal cytoplasm rather than K⁺ changes²⁹. The specific pH stability of the K⁺ indicator used in the current study, i.e., lc-LysM GEPII 1.0, has not been evaluated; however, lc-LysM GEPII 1.0 is derived by only 3 point mutations from GEPII 1.0, which shows only negligible pH sensitivity in the physiological range²¹. Therefore, it is reasonable to conclude that the decrease of the FRET ratio of lc-LysM GEPII 1.0 in response to Bicuculline observed in the current study is indeed caused by a reduction in [K⁺]_i. Hence, we can conclude with a reasonable confidence level that lc-LysM GEPII 1.0 accurately detects [K⁺]_i during neuronal activity.

After ensuring the general functionality of lc-LysM GEPII 1.0 in neurons, we aimed to investigate whether the biosensor can detect changes of [K⁺]_i occurring during single action potentials. Our cultured cortical neurons displayed spontaneous neuronal activity consisting of multiple single action potentials as evidenced by single, sharp increases in [Ca²⁺]_i, which could be silenced with the inhibitor of voltage-gated sodium channels TTX. After characterizing these robust spontaneous [Ca²⁺]_i fluctuations in our cell culture system, we transduced our cells with lc-LysM GEPII 1.0. We could not observe any distinguishable changes in the lc-LysM GEPII 1.0 signals that indicated spontaneous K⁺ dynamics. In addition, the signal was not affected by TTX-induced neuronal silencing and correlated with the expression level of the sensor. These findings suggest that lc-LysM GEPII 1.0 failed to detect [K⁺]_i changes induced by single action potentials. This conclusion is further supported by the observation that controlled optogenetic stimulation of neurons with low frequencies also failed to elicit any change in lc-LysM GEPII 1.0 fluorescence.

These data may be interpreted in three ways: 1) either lc-LysM GEPII 1.0 is not sensitive enough to resolve small changes of [K⁺]_i, or 2) up to a certain number of action potentials [K⁺] does not decrease in the cytoplasm or decreases only in the vicinity of the cell membrane, or 3) both scenarios are valid. To understand which of these scenarios is the most likely, it is essential to know that [K⁺]_i in neurons is around 140 mM while in the extracellular space, it is only 3 mM⁶. In addition, the extracellular space only makes up about 20% of the brain volume³⁶. Hence, a change of a few mM of potassium can lead to a significant relative change of [K⁺]_e, while barely affecting [K⁺]_i. As the membrane potential is determined by the ratio between intra- and extracellular ions, the contribution of potassium to the membrane potential is nearly exclusively governed by [K⁺]_e³⁷. Therefore, most studies investigating K⁺ alterations in response to neuronal activity measured extracellular rather than intracellular changes. For example, strong, tetanic neuronal stimulation or epileptic seizures reliably lead to increases of [K⁺]_e that plateau around 10 to 12 mM^38,39, likely because of efficient potassium clearance via astrocytes^39,40. Cortical spreading depolarizations can further increase extracellular K⁺ levels to around 65 mM^38,41,42. In contrast, strong tetanic stimulation of frog motor neurons led to a decrease of intracellular [K⁺] by not more than 5 mM⁴³, while Raimondo et al. estimated that epileptic seizures lead to a decrease of intracellular potassium concentrations of only 2 mM⁴⁴. The relatively small activity-dependent changes in intracellular potassium are even more relevant when considering spontaneous neuronal activity. A single action potential leads to an increase of extracellular potassium by only 0.2–0.8 mM^45,46. Hence, changes in [K⁺]_i are most likely much smaller.

As the EC₅₀ of lc-LysM GEPII 1.0 (27 mM – 60 mM) is far from the intracellular K⁺ concentration in neurons (~ 140 mM), the sensor will either be close to saturation or fully saturated when expressed in the intracellular space. Indeed, lc-LysM GEPII 1.0 is saturated at 150 mM potassium and a drop to 100 mM leads to a change of the dynamic range by only about 10%²¹. Therefore, small changes of intracellular K⁺ are unlikely to elicit a change of the FRET ratio of lc-LysM GEPII 1.0 strong enough to be resolved. We conclude that to resolve intracellular K⁺ changes in response to spontaneous neuronal activity, lc-LysM GEPII 1.0 needs to be improved towards lower affinity for K+.

Combining live cell imaging with optogenetic stimulation can be used to control [K⁺] changes tightly and therefore allows comparing the performance of potential candidate variants. This might help facilitating the development of improved potassium sensors focusing on resolving K⁺ changes during neuronal activity. The inability to measure [K⁺]_i following single action potentials can be attributed to the anticipated little changes in [K⁺]_i in such scenarios and the very low EC₅₀ of lc-LysM GEPII 1.0 for neuronal measurements.

Despite these shortcomings of lc-LysM GEPII 1.0 for the measurement [K⁺]_i in neurons, we were able to express lc-LysM GEPII 1.0 in cortical neurons at a sufficiently high level to be visualized by in vivo imaging and to perform the first measurements of [K⁺]_i in the living mouse brain. Like in cultured cells, we did not detect any changes of [K⁺]_i during spontaneous neuronal activity. However, we observed increased fluorescence lifetime (FLIM) of lc-LysM GEPII 1.0 during massive neuronal depolarizations triggered by cerebral ischemia. Other groups already used imaging approaches to measure [K⁺] in response to cortical depolarization waves in vivo, however, only in the extracellular space. Transient extracellular K⁺ increases were recorded after injection of the K⁺-sensitive dye APG-2 in the cisterna magna⁴⁷ or topical application of GINKO2 on the open cortex in mice²⁹. Wu and colleagues tried to measure [K⁺]_i dynamics in neurons and astrocytes of drosophila in response to induced neuronal activity; however, they reported that their data obtained with the single fluorophore sensor GINKO2, which displays a strong pH dependency, is quite likely confounded by activity-induced acidification artifacts²⁹. Also, cortical depolarization waves cause intracellular acidification in neurons^48,49; however, as discussed above, GEPIIs are considerably more pH stable than GINKO2. Therefore, we do not expect that pH affected our measurements considerably. Future measures with improved GEPIIs in terms of K_D and pH stability and a pH-sensor such as pHRed⁵⁰ could help disentangle the actual potassium signal from a potential pH artifact⁵¹.

Our data show that intracellular K⁺ imaging in neurons is, in principle, possible in vivo. However, the low K_D of lc-LysM GEPII 1.0 and the high technical requirements of such measurements (in vivo FLIM) currently limit the practical use of this approach. Hence, optimizing lc-LysM GEPII 1.0 with respect to affinity, dynamic range, and fluorescence intensity is critical to allowing the measurement of intracellular K⁺ to become a standard tool in neuroscience research. We believe such a task is worth pursuing because reliably detecting intracellular K⁺ dynamics in vivo will help investigating neuronal phenomena and other critical neurobiological processes such as astrocytic K⁺ clearance and spatial K⁺ buffering^31,32.

In conclusion, we provide a first functional characterization of lc-LysM GEPII 1.0 in cortical neurons both in vitro and in vivo. We show that lc-LysM GEPII 1.0 can resolve [K⁺] changes in response to intense neuronal activity, however, fails to detect K⁺ dynamics in response to mild activity or single action potentials. Future optimization of lc-LysM GEPII 1.0, especially with respect to an increased K_D, is needed for allowing precise K⁺ measurements in physiologic settings.

Data availability

The data generated during the study are available from the corresponding author upon reasonable request.

Acknowledgements

The current study was mainly funded by a joint grant provided by the Deutsche Forschungsgemeinschaft (DFG, German Research Foundation) and the Austrian Science Fund (FWF) to NP (PL249/15-1) and RM (I 3716). Further funding was received by the DFG under Germany’s Excellence Strategy within the framework of the Munich Cluster for Systems Neurology (EXC 2145SyNergy – ID 390857198; NP and AL).

Author Contributions

B.G., S.B., H.B., R.M. and N.P. conceived the study and contributed to the study design. B.G., G.M.C., C.R., J.S., G.L., A.G.Z. and S.F. contributed to the acquisition and analysis of the data. B.G., E.E., A.L., R.M., S.F. and N.P. contributed to the interpretation of the data. B.G., S.F. and N.P. drafted the manuscript. All authors read and approved the final manuscript.

Competing Interests

The authors declare no competing interests.

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Probing Intracellular Potassium Dynamics in Neurons with the Genetically Encoded Sensor lc-LysM GEPII 1.0 in vitro and in vivo

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