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A sensitive and specific genetically-encoded potassium ion biosensor for in vivo applications across the tree of life [1]

['Sheng-Yi Wu', 'Department Of Chemistry', 'University Of Alberta', 'Edmonton', 'Alberta', 'Yurong Wen', 'Department Of Biochemistry', 'Center For Microbiome Research Of Med-X Institute', 'The First Affiliated Hospital', 'Xi An Jiaotong University']

Date: 2022-09 

Potassium ion (K + ) plays a critical role as an essential electrolyte in all biological systems. Genetically-encoded fluorescent K + biosensors are promising tools to further improve our understanding of K + -dependent processes under normal and pathological conditions. Here, we report the crystal structure of a previously reported genetically-encoded fluorescent K + biosensor, GINKO1, in the K + -bound state. Using structure-guided optimization and directed evolution, we have engineered an improved K + biosensor, designated GINKO2, with higher sensitivity and specificity. We have demonstrated the utility of GINKO2 for in vivo detection and imaging of K + dynamics in multiple model organisms, including bacteria, plants, and mice.

Funding: This work was supported by grants from the Canadian Institutes of Health Research (CIHR, FS-154310 to REC) and the Natural Sciences and Engineering Research Council of Canada (NSERC, RGPIN 2018-04364 to REC, RGPIN-2020-05514 to KB, and RGPIN-2016-06478 to MJL). SYW was supported by NSERC Canada Graduate Scholarships – Doctoral program, Alberta Innovates Technology Future (AITF) Graduate Scholarship, and the University of Alberta. YW was supported by the Alberta Parkinson Society Fellowship and National Natural Science Foundation of China (No. 31870132 and No. 82072237). This research used resources of the Advanced Photon Source (APS), a U.S. Department of Energy (DOE) Office of Science User Facility operated for the DOE Office of Science by Argonne National Laboratory under Contract No. DE-AC02-06CH11357. X-ray crystallography was performed using Beamline 23IDB at APS. GM/CA@APS has been funded by the National Cancer Institute (ACB-12002) and the National Institute of General Medical Sciences (AGM-12006, P30GM138396). Data were also collected at beamline CMCF-ID at the Canadian Light Source, a national research facility of the University of Saskatchewan, which is supported by the Canada Foundation for Innovation (CFI), the NSERC, the National Research Council (NRC), the CIHR, the Government of Saskatchewan, and the University of Saskatchewan. NBCS and MF work was supported by the European Research Council (Grant No. 803048) and Charles University Primus (Grant No. PRIMUS/19/SCI/09). CCHL, AGD, HH, and MN were supported by the Novo Nordisk Foundation (NNF19OC0058058 to HH and NNF13OC0004258 to MN) and the Lundbeck Foundation (R155-2016-552 to MN and R263-2017-4062 to AGD). BRT was supported by NIH grant R01GM095903. AA and KP were supported by Howard Hughes Medical Institute. ARK and DFE were supported in part by NSF grant 2037828. Two-photon characterization work (MD and RSM) was supported by the NIH BRAIN grant U24 NS109107 (Resource for Multiphoton Characterization of Genetically-Encoded Probes). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

Data Availability: Data availability Plasmids and DNA sequences are available via Addgene (Addgene ID 177116, 177117). The GINKO1 structure is deposited in the Protein Data Bank (PDB ID:7VCM). Seeds are deposited in the NASC Arabidopsis seed repository ( https://arabidopsis.info/ ID: N2111001). Data supporting the findings in this research are included in the supplementary data file. Code availability Root elongation quantification code is available at https://sourceforge.net/projects/lbopsis/ .

This is an open access article, free of all copyright, and may be freely reproduced, distributed, transmitted, modified, built upon, or otherwise used by anyone for any lawful purpose. The work is made available under the Creative Commons CC0 public domain dedication.

(A) Schematic representation of GINKO. In the top panel, the linear DNA representation of GINKO gene shows the ligand recognition domain Kbp (BON in cyan and LysM in yellow) inserted in the split EGFP (green). In the bottom panel, the illustration shows a K + -binding induced conformational change of Kbp leading a change in fluorescence. (B) Cartoon representation of the structure of GINKO1 with the BON (bacterial OsmY and nodulation) domain of Kbp in cyan, the LysM (lysin motif) domain of Kbp in yellow, and the EGFP in green. The chromophore and the K + ion (green) are shown in sphere representation. (C) The K + is coordinated by carbonyl backbone atoms of 6 amino acids. The distance (in Å) of each amino acid backbone oxygen to the K + ion was measured in PyMOL. (D) Structure alignment of the Kbp domain in GINKO1 and the previously reported NMR structure of Kbp (PDB ID: 5FIM). Kbp NMR structure ensemble is shown in ribbon representation. GINKO1 BON domain is in cyan; GINKO1 LysM domain is in yellow. Kbp NMR structure BON domain is in blue, and LysM domain is in orange. (E) Zoom-in view of the binding pocket in the GINKO1 crystal structure and the Kbp NMR structure (PDB ID: 5FIM). Structure coloring is the same as in (D). EGFP, enhanced green fluorescent protein; Kbp, K + -binding protein.

A high-performance genetically-encoded fluorescent biosensor for K + could enable a variety of applications that are currently impractical or impossible by enabling targeted expression and noninvasive in vivo imaging. We have previously reported a prototype intensiometric K + biosensor, designated GINKO1, based on the insertion of K + -binding protein (Kbp) [ 12 ] into enhanced green fluorescent protein (EGFP) ( Fig 1A ) [ 13 ]. Ratiometric genetically-encoded biosensors have also been reported [ 13 , 14 ]. To create a more robust K + biosensor with broader utility, we undertook an effort to further improve the sensitivity and specificity of GINKO1.

The potassium ion (K + ) is one of the most abundant cations across biological systems [ 1 ]. It is involved in a variety of cellular activities in organisms ranging from prokaryotes to multicellular eukaryotes [ 2 – 4 ]. While studies of other biologically important cations, notably calcium ion (Ca 2+ ), have been revolutionized by the availability of high-performance genetically-encoded biosensors [ 5 , 6 ], the development of analogous biosensors for K + has lagged far behind. Canonical methods to monitor K + include K + -sensitive microelectrodes and synthetic dyes. Microelectrodes are considered the gold standard for their sensitivity and selectivity, but they are invasive and not suitable for high-throughput cellular or subcellular K + detection [ 7 ]. Synthetic dye-based approaches allow K + visualization in live cell populations with improved spatiotemporal resolution [ 8 – 11 ]; however, they still require dye loading and washing procedures and lack the targetability to specific cell types or subcellular compartments.

Results and discussion

Structure of GINKO1 To better understand the K+-dependent fluorescence response mechanism of GINKO1 and facilitate further engineering, we determined the crystallographic structure of GINKO1 in the K+-bound state at 1.85 Å (Figs 1B and S1 and S1 Table). Well-diffracting crystals of the unbound state were unattainable. The K+-bound crystal structure revealed the location and coordination geometry of the K+-binding site of Kbp (Fig 1C), which was not apparent from the previously reported NMR structure (Fig 1D and 1E) [12]. Notably, the K+ ion is coordinated via 6 backbone carbonyl oxygen atoms (from amino acids V154, K155, A157, G222, I224, and I227). This coordination via backbone carbonyl oxygen atoms is similar to that observed in the K+ selective filters of KcsA (PDB ID: 1BL8) [15] and TrkH (PDB ID: 4J9U) [16], as well as K+-coordinating compound valinomycin [17]. The distances of coordinating carbonyl oxygens to K+ in GINKO1 range from 2.6 to 3.2 Å with a mean value of 2.8 Å (Fig 1C), similar to those in KscA (2.70 to 3.08 Å, with a mean value of 2.85 Å) [18], and valinomycin (2.74 to 2.85 Å) [17]. One difference is that K+ is coordinated via 8 oxygens from backbone carbonyls in both KcsA and TrkH, and 6 backbone carbonyls in Kbp. In the previous study that described the Kbp NMR solution structure, it was suggested that crystallization of Kbp for X-ray crystallography was challenging [12]. We suspect that fusing Kbp to EGFP constrains the conformational mobility of Kbp, thus increasing the stability of Kbp protein for it to be crystallized as a domain in GINKO1. A similar approach has recently been reported to stabilize small transmembrane proteins for crystallization [19]. The Kbp region of the GINKO1 structure aligns well with the previous Kbp NMR solution structure (Fig 1D and 1E). The BON domain and the LysM domain of Kbp were both well resolved in the GINKO1 structure. The structure further revealed that the K+ binding site is located in the BON domain, close to the interface between the BON and LysM domains (Fig 1D). This is consistent with the previous finding that the BON domain binds K+ and the LysM domain stabilizes the K+-bound BON domain [12].

Monitoring intracellular K+ concentration in bacteria with GINKO2 To determine whether GINKO2 could be used to monitor intracellular K+ in bacteria, we attempted to use it in E. coli to monitor the decreasing intracellular K+ concentration that can be induced by growth in a low-K+ medium (Fig 4A). Real-time detection of intracellular K+ concentration dynamics could allow the relationship between extracellular low-K+ availability, intracellular K+ concentration, and bacterial growth rate, to be established. The excitation ratiometric change of GINKO2 presents a unique solution to monitor K+ concentration changes in proliferating E. coli, in which intensity-based measurements are impeded by the increasing biosensor expression level during cell growth. GINKO2-expressing E. coli grown in a medium with 20 μM K+ exhibited a 58% decrease in excitation ratio R 470/390 (Fig 4B), corresponding to an estimated decrease in intracellular K+ from 103 ± 21 mM to 20 ± 3 mM based on a calibration in E. coli (Fig 4C and 4D). In contrast, cells grown in a medium with 800 μM K+ showed unchanged intracellular K+ concentration at around 80 mM during the same growth period (Fig 4D). An excitation ratiometric pH biosensor pHluorin [24] was used to confirm that the intracellular pH remained stable. This application of GINKO2 demonstrated its practicality for real-time monitoring of intracellular K+ in E. coli. PPT PowerPoint slide

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TIFF original image Download: Fig 4. Monitoring intracellular K+ concentrations with GINKO2 in E. coli grown in K+-depleted media. (A) E. coli are capable of accumulating K+ to a higher concentration than the environment. The free intracellular K+ concentration is around 100 mM when cells are cultured with sufficient K+ in the environment such as in LB. In this work, we aimed to investigate the intracellular K+ concentrations of E. coli growing in K+-depleted media. (B) Excitation ratio (R 470/390 ) of GINKO2 in E. coli cells grown in K+-deficient media. Optical density at 600 nm (OD 600 ) reflects cell density during the growth. Two low K+ concentrations (open circle: 800 μM, solid circle: 20 μM) were used for the experiment: only the medium supplemented with 20 μM K+ induced detectable K+ decrease during the growth. n = 6–8 for E. coli expressing GINKO2 in 20 μM K+; n = 3–8 for E. coli expressing GINKO2 in 800 μM K+; n = 3 for E. coli expressing pHluorin in 20 μM K+; n = 3–6 for E. coli expressing pHluorin in 800 μM K+. (C) A K+ titration calibration curve was obtained with E. coli cells pretreated with 30 nM valinomycin for 5 min. The GINKO2-expressing cells (solid circle and dashed line) and nonexpressing cells (control, empty circle) were both titrated with K+ at OD 600 approximately 0.1. The calibration curve (solid circle and continuous line) was obtained by subtracting the fluorescence readings of control from those of GINKO2-expressing cells. (D) K+ concentrations in (B) were estimated based on the calibration curve in (C). Fig 4A was created with BioRender.com. The underlying data for Fig 4B-4D can be found in S1 Data. https://doi.org/10.1371/journal.pbio.3001772.g004

In vivo imaging of intracellular K+ dynamics in plants with GINKO2 To evaluate the utility of GINKO2 in vivo in plants, we attempted to use GINKO2 to monitor intracellular K+ concentration changes in Arabidopsis thaliana under stress conditions. K+ is an essential nutrient for plants and regulates root growth, drought resistance, and salt tolerance [25,26]. Despite the importance of K+, its detailed spatiotemporal dynamics remain elusive in plants, largely due to the lack of high-performance imaging probes. A. thaliana stably transformed with GINKO2 expressed under the control of the g10-90 constitutive promoter exhibited homogeneous fluorescence in leaf epidermis, hypocotyls, primary root tips, and primary mature roots (Fig 5A). GINKO2 expression did not affect root elongation (S8 Fig) nor the overall plant development. GINKO2 fluorescence was visible in the cytoplasm but absent in vacuoles. Vacuoles are K+ reservoirs with concentrations as high as 200 mM. This significant store of vacuolar K+ is available to be released into the cytoplasm for the regulation of the cytoplasmic K+ concentration [27]. Due to the low vacuolar pH (pH = 5.0 to 5.5) [28], GINKO2 fluorescence would be quenched if it was targeted to vacuoles. Therefore, even if it was localized to the vacuole, GINKO2 is likely to be unsuitable for reporting vacuolar K+ concentration changes. When the seedlings were transferred from the plant standard growing medium (½MS medium) with 10 mM K+, to K+ gradient buffers (0.1, 1, 10, and 20 mM) for 2.5 d, cytosolic GINKO2 fluorescence reported no significant differences in R 488/405 across the concentration range (S9A Fig), suggesting that the vacuolar pools of K+, invisible to GINKO2, might buffer the low K+ in the treatments. PPT PowerPoint slide

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TIFF original image Download: Fig 5. Monitoring K+ efflux in Arabidopsis thaliana with GINKO2 during salt stress. (A) Expression and characterization of GINKO2 in A. thaliana. Representative fluorescence images of g10-90::GINKO2 expressing tissues excited at 405 nm and 488 nm. Scale bar = 50 μm. (B) Effect of increasing concentrations of KCl and 2 μM valinomycin on g10-90::GINKO2 R 488/405 after 6 h of K+ depletion with a 0-mM KCl and 2-μM valinomycin pretreatment. n = 16–21 individual roots. Letters indicate the significantly different statistical groups with P < 0.05 minimum. Statistical analysis was conducted with a nonparametric multiple comparison. (C) Effect of 100 mM NaCl on g10-90::GINKO2 R 488/405 (top panel) in root tips, mature root stele, and epidermis with K+ depleted for 6 h. Treatment was applied at time 0. n = 14 (root tip), 8 (mature root stele and epidermis) individual seedlings. pHGFP expressing roots (bottom panel) were used as controls. n = 9 individual seedlings for root tips, mature root stele, and epidermis. The underlying data for Fig 5B and 5C can be found in S1 Data. https://doi.org/10.1371/journal.pbio.3001772.g005 It has been previously reported that during low K+ treatment, the vacuolar pool of K+ gradually decreases to sustain the cytosolic pool, and only when the vacuolar pool is severely diminished does the cytosolic K+ concentration start to decline [27]. Therefore, we thought to deplete the vacuolar K+ before imaging to reduce its buffering effect by transferring the seedlings onto a medium containing 0 mM K+ and the K+-specific ionophore valinomycin (2 μM). This predepletion of K+ enabled the direct manipulation of the cytosolic K+ concentration using media of different K+ concentrations, allowing GINKO2 to display its full sensing capacity. In permeabilized and K+-depleted seedlings, we observed a significant decrease of the GINKO2 R 488/405 , indicating a lowered cytoplasmic K+ concentration (S9B Fig). GINKO2 excitation ratio R 488/405 correlated well with the medium K+ concentrations in the physiological range of 1 to 100 mM (Fig 5B). We next imaged K+ dynamics in roots under salt (NaCl) stress. The Na+ influx to the roots triggers a K+ efflux to counterbalance the membrane depolarization [29]. NaCl treatment without predepletion of K+ produced an initial increase in the cytoplasmic K+ concentration followed by a decrease after 10 min (S9C Fig). This, again, could be attributed to the vacuoles exporting K+ into the cytoplasm. With K+ predepletion and a treatment of 100 mM NaCl, GINKO2 reported the K+ efflux with substantial decreases in R 488/405 in root tips (35%), mature root stele (19%), and mature root epidermis (13%) (Fig 5C, top panel, S10 Fig and S1 and S2 Movies). While cytosolic pH of plant cells is known to be tightly regulated and well maintained [30], even under an induced salt stress [31], we investigated the possibility that pH changes could be responsible for the observed changes in GINKO2 fluorescence. We used the ratiometric pHGFP, a pH sensor modified from ratiometric pHluorin for plant expression, which exhibits an increase in R 488/405 with a decrease in pH [32,33]. Ratiometric measurement of pHGFP fluorescence suggested intracellular pH remained relatively stable after the NaCl treatment (Fig 5C, lower panel). Specifically, in root tips, pH is transiently lowered (3% increase in pHGFP R 488/405 ) upon the addition of NaCl but quickly returned to the baseline level. In mature root stele, the pH remained unchanged throughout the experiment. These pH control experiments suggested that the observed decline in GINKO2 ratio under salt stress (Fig 5C) resulted from a change of K+ concentration rather than pH. In contrast, in the mature root epidermis, pHGFP reported an overall 5% R 488/405 increase, indicating a slight pH decrease. Accordingly, we were unable to conclude that the observed R 488/405 change (13%) of GINKO2 in the epidermis was solely caused by a decrease in the K+ concentration. Taken together, these results demonstrated that GINKO2 is capable of reporting cytoplasmic K+ dynamics in vivo in the roots of A. thaliana with great sensitivity and have provided insight into the complexity of K+ regulation in plants. With appropriate protocols and controls, GINKO2 represents a substantial step forward for the study of K+ homeostasis in plants with the potential to be applied to a variety of experimental paradigms, including detection and characterization of mutant phenotypes (e.g., mutations in K+ transporters), and characterization of changes in K+ dynamics under stress conditions.

In vivo imaging of extracellular K+ changes in mice with GINKO2 To further explore GINKO2 applications, we tested whether GINKO2 is capable of reporting extracellular K+ changes in vivo during cortical spreading depolarization (CSD) in the mouse brain. CSD is a propagating, self-regenerating wave of neuronal depolarization moving through the cortex and is associated with severe brain dysfunctions such as migraine aura and seizures [34]. On the molecular level, CSD is accompanied by propagating waves of increased extracellular K+ from a baseline of 2.5 to 5 mM to a peak concentration of 30 to 80 mM [35]. As previously reported for Kbp-based K+ biosensor GEPII [36], we have been unable to express and display functional GINKO2 on the extracellular membrane for reasons that remain unclear to us. To circumvent this limitation, we turned to the exogenous application of bacterially expressed GINKO2 as an alternative method to evaluate extracellular K+ concentration dynamics during CSD. Purified GINKO2 protein (6.55 mg/mL in artificial cerebrospinal fluid (aCSF)) was exogenously applied to the extracellular space of deeply anesthetized mice above the somatosensory cortex (Fig 6A). To experimentally elicit CSD, we applied 1 M KCl to a separate frontal craniotomy [35] (Fig 6A), after which multiple waves of GINKO2 fluorescence intensity increase were observed, propagating at a velocity of 2.4 ± 0.8 mm/min (Figs 6B, 6C, 6D and S11A and S3 Movie). The fluorescence intensity increased by 1.0 ± 0.2× (Fig 6E), with a fast rise at 0.29 ± 0.07% s−1 and a significantly slower decay at 0.03 ± 0.01% s−1 (Fig 6F). The duration of the waves (width at half maximum) was 22 ± 6 s (Fig 6G). The fluorescence increases observed with GINKO2 during CSD (Fig 6) correspond well to descriptions of the extracellular K+ concentration dynamics previously reported during CSD [35]. A control experiment using EGFP (2.13 mg/mL in aCSF) was performed to evaluate pH changes under the same treatment (S11 Fig). A 30% fluorescence decrease under the same treatment indicated a possible decrease in pH based on the pH profile of EGFP [37]. A decline in pH, suggested by either the EGFP control or previous reported pH dynamics during CSD (short increase in pH for approximately 5 s, followed by a decrease in pH [38]), should have resulted in a GINKO2 fluorescence change in the opposite direction of the one we observed. This strongly supported that the observed elevation of GINKO2 fluorescence resulted from a substantial extracellular K+ concentration increase during CSD. Overall, these results suggest that GINKO2 is an effective tool for reporting extracellular K+ concentration changes in vivo in the mouse brain during CSD. PPT PowerPoint slide

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TIFF original image Download: Fig 6. Monitoring the CSD-induced elevation of extracellular K+ concentrations in mice. (A) Experimental setup of 2P microscopy in anesthetized mice. CSD was induced using 1 M KCl applied to a separate frontal craniotomy (small circle) of the imaging window (large circle) at a distance of 3 mm. Exogenously expressed GINKO2 protein was purified and externally applied to the imaging site by pipetting. (B) Averaged image of GINKO2 in the somatosensory cortex (−70 μm) obtained using 2P microscopy. E. coli expressed GINKO2 was applied externally by bath application 1 h before imaging. The image depicts the ROIs corresponding to traces in (C). Scale bar: 100 μm. (C) Example of traces from ROIs in the same animal, depicting the first CSD wave. x-axis: 5 s, y-axis: 100% ΔF/F 0 , mean ± SD. (D) Example of a CSD wave showing decay, rise, width, and amplitude. (E) Comparison between ΔF/F of baseline before each CSD and at peak. N = 2, n = 5, paired t test, ***p = 0.0007. (F) Calculated slope coefficient using simple linear regression of the rise and the decay of CSD waves. N = 2, n = 5, paired t test, **p = 0.0024. (G) Average CSD wave duration N = 2, n = 5, mean ± SD. The underlying data for Fig 6D-G can be found in S1 Data. CSD, cortical spreading depolarization; ROI, region of interest; 2P, 2-photon. https://doi.org/10.1371/journal.pbio.3001772.g006

In vivo imaging of K+ dynamics in Drosophila neurons and glial cells with GINKO2 In an attempt to visualize potential K+ changes in vivo in Drosophila, we fused GINKO2 with a red fluorescent pH biosensor, pHuji [39], to monitor both K+ and pH concurrently. We first characterized pHuji-GINKO2 fusion protein in vitro. Decreasing pH reduces the green fluorescence of GINKO2 but does not change the affinity for K+ (S12 Fig). The red fluorescence of pHuji is not sensitive to the K+ concentration. We then produced transgenic flies expressing pHuji-GINKO2 under control of the Gal4-UAS system, either in neurons (elav-Gal4) or in glia (repo-Gal4). Fly brains were stimulated either by rapidly elevating the extracellular K+ concentration or electrically with a glass electrode. In neurons, stimuli led to a decline in GINKO2 fluorescence, while in glia, the same stimuli led to an increase in GINKO2 fluorescence (Fig 7). However, these stimuli also led to similar changes in pHuji fluorescence, indicating substantial pH changes (Fig 7). It is expected that stimulated neuronal activities would likely lead to a K+ efflux, as previously observed by others in several different preparations [40]. However, due to the susceptibility of GINKO2 to pH interference, the GINKO2 fluorescence changes observed in this particular set of experiments cannot be conclusively interpreted as K+ changes in the stimulated neurons or glial cells. PPT PowerPoint slide

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TIFF original image Download: Fig 7. pHuji-GINKO2 responses to K+ or electrical stimulation in the Drosophila brain. (A) Fly heads were encapsulated in a photopolymerizable resin (LCR) delivered by a thin needle with the posterior side of the head on the bottom of the petri dish. The LCR-coated heads were covered by a droplet of saline and cured by blue light at 460 nm. The heads then were transversely sectioned through the joints between the second and third antennal segments [58]. Fly brains expressing pHuji-GINKO2 in (B) neurons and (C) glia were stimulated by adding KCl in the bath to a final concentration of 3.2 mM. The black arrow indicates the time at which KCl was added. Fly brains expressing pHuji-GINKO2 in (D) neurons and (E) glia were stimulated by 500 electrical impulses delivered at 50 Hz, starting at the time indicated by the blue arrow, by a glass microelectrode. The heads were oriented with the eyes at the top of the frame during image acquisition. The samples were excited by alternating between 490 nm and 555 nm, and the ROIs used to plot the graphs are indicated by dashed circles. Scale bars: 100 μm. Fig 7A was created with BioRender.com. The underlying data for Fig 7B-7E can be found in S1 Data. LCR, light cured resin; ROI, region of interest. https://doi.org/10.1371/journal.pbio.3001772.g007

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