Semisynthetic fluorescent pH sensors for imaging exocytosis and endocytosis

The GFP-based superecliptic pHluorin (SEP) enables detection of exocytosis and endocytosis, but its performance has not been duplicated in red fluorescent protein scaffolds. Here we describe “semisynthetic” pH-sensitive protein conjugates with organic fluorophores, carbofluorescein, and Virginia Orange that match the properties of SEP. Conjugation to genetically encoded self-labeling tags or antibodies allows visualization of both exocytosis and endocytosis, constituting new bright sensors for these key steps of synaptic transmission.

S ynaptic transmission is mediated by the rapid fusion of synaptic vesicles (SVs) with the plasma membrane. Precise monitoring of exocytosis is important for elucidating fundamental mechanisms of cell-cell communication, and investigating the underlying causes of neurological disorders 1 . The lumen of synaptic vesicles are typically acidified (pH 5.6) by the action of vesicle-resident V-ATPases, which creates the driving force for neurotransmitter uptake. Upon fusion with the plasma membrane, the contents of the vesicle rapidly equilibrate with the extracellular environment (pH 7.4). This large change in pH allows for the visualization of exocytosis using a pH-sensitive variant of green fluorescent protein (GFP) that is expressed as a fusion with a vesicular membrane protein 2 . This "superecliptic pHluorin" (SEP) exhibits ideal properties for detecting the change in pH upon vesicle fusion, with near-ideal pK a , cooperative protonation, and low background fluorescence in the protonated state, making it an excellent tool for monitoring exocytosis in living cells 2 .
A useful extension of this technology has been the creation of pH sensors based on red fluorescent proteins (RFPs) such as mOrange2 3 , pHTomato 4 , pHoran4 5 , and pHuji 5 . Longer excitation wavelengths are less phototoxic, elicit lower levels of autofluorescence, facilitate multicolor imaging experiments, and allow concomitant use of optogenetics. Nevertheless, it has proven difficult to engineer red-shifted pHluorins that match the optimal pK a , cooperativity, and dynamic range of SEP, perhaps due to inherent limitations in RFP scaffolds. More generally, these techniques rely on overexpression of reporter proteins in SVs and the effect of overexpression is a confounding factor in interpreting experimental results 6 .
To circumvent the problems with genetically encoded red fluorophores, we develop a "semisynthetic" sensor platform using red-shifted chemical fluorophores derived from fluoresceincarbofluorescein (CFl) and Virginia Orange (VO)-with pH sensitivities similar to SEP. We have adapted these fluorophores to serve as probes for exo-and endocytosis in two ways. First, we synthesize benzylguanine derivatives, which bind specifically to SNAP-tag ligands 7 , to label exogenous vesicular proteins fused with the SNAP-tag. Second, we label an antibody targeted to the extracellular epitope of a vesicular protein, synaptotagmin1, which allow the imaging of the exo-/endocytosis cycle of an endogenous protein.

Results
In vitro characterization of CFl and VO as pH sensors. Given the limitations of pH-sensitive RFPs and the potential problems with overexpression of sensor proteins, we pursued an alternative strategy: creation of semisynthetic pH indicators using organic pH-sensitive dyes attached to either expressed self-labeling tags such as the SNAP-tag 7 or antibodies that recognize native vesicular proteins (Fig. 1a). To match the performance of SEP, we required a pH-sensitive organic dye that can undergo a cooperative transition from a bright, fluorescent form at neutral pH to a nonfluorescent form at low pH. Unfortunately, the majority of pH-sensitive dyes do not meet these requirements. The    archetypical small-molecule pH sensor is fluorescein (Fl, 1, Fig. 1b), which transitions between a highly fluorescent dianion (1 2− ) and a less fluorescent monoanion (1 − ) with a relatively low pK a value of 6.3 ( Fig. 1c) 8 . Other unsuitable synthetic pH probes include the ratiometric seminapthorhodofluor (SNARF) dyes 9 that exhibit high background, as well as cyanine and rhodaminebased pH sensors that show the opposite pH sensitivity profile 10,11 . We recently synthesized new derivatives of fluorescein (1) where the xanthene oxygen was replaced with a gem-dimethylcarbon moiety. This work resulted in "carbofluorescein" (CFl, 2, Fig. 1b) 12 , and the difluorinated derivative "Virginia Orange" (VO, 3, Fig. 1b) 13 . We discovered that this oxygen→carbon substitution elicited significant changes in photophysical and chemical properties of the fluorescein scaffold. Fl (1) exhibits λ ex / λ em = 491 nm/510 nm at high pH, whereas CFl (2) and VO (3) are red-shifted with λ ex /λ em = 544 nm/567 nm and 555 nm/581 nm, respectively. In addition to this bathochromic shift, the pH sensitivity of the dyes was markedly different. Fluorescein exhibits strong visible absorption at both pH 5.6 (vesicle pH) where the monoanion 1 − dominates, and pH 7.4 (extracellular pH) where the dianion form 1 2− is prevalent-this can be observed by eye (Fig. 1c, d). In contrast, CFl (2) undergoes a cooperative transition between a highly colored dianion species (2 2− ) and a colorless lactone form (2 lactone ; Fig. 1c). This is also evident visually as a solution of CFl (2) is colorless at pH 5.6, but shows robust visible absorption at pH 7.4 (Fig. 1d). Fluorescence-based titrations (Fig. 1e) gave pK a values of 6.3 and Hill coefficient (η H ) value of 0.97 for fluorescein (1), consistent with previous reports 8 . CFl (2) and VO (3) displayed pK a values of 7.5 and 6.7, and η H values of 1.6 and 1.5, respectively. This cooperative transition likely stems from the altered lactone-quinoid equilibrium observed in the carbon-containing analogs of fluorescein and rhodamine dyes 12 . These pH sensitive red fluorophores have extinction coefficients and quantum yields similar to the pH sensitive red fluorescent proteins pHoran4 and pHuji 5 (Supplementary Table 1). Moreover, we tested the resistance of these fluorophores to photobleaching ( Supplementary Fig. 1a). While CFl showed similar photobleaching rate as Fl (τ = 5.33 s and τ = 5.35 s), VO was two to three times more photostable (τ = 14.54 s). Therefore, the longer absorption and emission wavelengths, higher pK a , resistance to photobleaching, and the cooperative colorless→colored transition upon increasing pH make both CFl and VO attractive scaffolds for building indicators to monitor synaptic vesicle fusion events.
To allow for specific labeling of expressed proteins, we prepared the SNAP-tag ligands attached to CFl (4) or VO (5) (Fig. 1f, Supplementary Fig. 1b). We tested the effects of protein conjugation on the properties of the dye by labeling SNAP-tag protein in vitro with CFl-SNAP-tag ligand 4 (Fig. 1g). We observed a shift in pK a to 7.3, and a decreased Hill coefficient (η H = 1.2; Fig. 1h). The active site of the SNAP-tag enzyme is flanked with two Lys, one Arg, and one His (PDB structure 3KZZ, DOI: 10.2210/pdb3kzz/pdb). The resulting Coulombic interaction between these positively charged amino acid residues and the CFl label most likely explains the decrease in pK a upon conjugation 8 . This polar surface might also stabilize the open form of the dye, resulting in the decreased cooperativity of the colored-colorless transition. Despite this lower pK a value and Hill coefficient, the fluorescence of the SNAP-tag-CFl conjugate is still completely suppressed at pH 5.6 (Fig. 1i). Detection of single exocytosis events in PC12 cells. Next, we tested these SNAP-tag-based probes in living cells. Building on existing SEP-based constructs 1, 2 , we designed several SNAP-tag fusion proteins: (i) SNAP-tag inserted within an intra-luminal loop of the vesicular acetylcholine transporter VAChT (VAChT-SNAP), and glutamate transporter VGluT1 (VGluT1-SNAP), and (ii) SNAP-tag protein attached to the luminal C-terminal side of the vesicle protein VAMP2 (VAMP2-SNAP). We first expressed VAChT-SNAP and VAMP2-SNAP in neuroendocrine PC12 cells. VAChT is targeted to small synaptic-like vesicles (SSLV) while VAMP2 is found in both SSLV and large dense core vesicles 14,15 . We found that the propensity of the CFl and VO fluorophores to adopt the neutral lactone form (Fig. 1c, d) allows for efficient intracellular labeling (Fig. 2a) with SNAP-tag ligands 4 or 5 without the use of other masking groups (e.g., acetate esters), which are typically required for fluorescein-based compounds. To monitor exocytosis, we depolarized cells with stimulation buffer containing high [K + ] and imaged single small vesicles as they fused with the plasma membrane using total internal reflection fluorescence (TIRF) microscopy. Cells expressing VAChT constructs displayed events at high frequency. Events detected in cells co-expressing VAChT-SEP and VAChT-SNAP (labeled with CFl ligand 4) showed comparable fold increases in fluorescence at exocytosis (2.19 ± 0.07 vs. 2.40 ± 0.12, mean ± s.e.m.) with similar decay kinetics in both the green and red channels (Fig. 2b, c). We also compared VAChT-SEP to VAChT-SNAP-VO (Fig. 2d, e) and VAChT-pHuji (Fig. 2f, g). Like the semisynthetic indicator from CFl ligand 4, the VAChT-SNAP-VO derived from compound 5 also showed comparable performance to the SEP sensor (2.65 ± 0.10 vs. 2.60 ± 0.16-fold increase; Fig. 2d, e). However, in PC12 cells the RFP-based VAChT-pHuji sensor showed lower relative performance when compared with VAChT-SEP under the same conditions (2.01 ± 0.05 vs. 1.32 ± 0.02-fold increase; Fig. 2f, g) making events harder to detect with pHuji than with the other pH-sensitive proteins. We also observed individual fusion events using VAMP2-SEP or VAMP2-SNAP-CFl ( Supplementary Fig. 2), albeit at low frequency, perhaps due to poor incorporation of this construct in PC12 cells.
Monitoring exocytosis and recycling of synaptic vesicles. We then tested these sensors in living neurons, focusing first on VAMP2-based constructs, which have been used extensively to follow SV exocytosis in neurons 1 . We co-transfected hippocampal neurons with VAMP2-SEP and either VAMP2-pHuji or VAMP2-SNAP incubated with CFl ligand 4 or Virginia Orange ligand 5. For all the sensors, we observed a robust increase in fluorescence following electrical stimulation in fields covered with transfected axons, signaling SV exocytosis. The relative increase in fluorescence upon SV exocytosis was slightly higher for the SEP channel relative to the red-shifted fluorescent indicators, VAMP2-SNAP-CFl (Fig. 3a, b), VAMP2-SNAP-VO (Fig. 3c, d), and VAMP2-pHuji (Fig. 3e, f), which behaved similarly. The kinetics of decay, which tracks endocytosis and re-acidification of the vesicle, were similar for all four labels (Fig. 3b, d, f). We then tested whether the added CFl or VO could label the whole SV population efficiently. To do so, we normalized the fluorescence increase induced by stimulation with an application of a buffer containing 50 mM NH 4 Cl, which quickly increases the intravesicular pH to extracellular pH 2 . We found that the proportion of SVs undergoing exocytosis estimated with this method was similar for all three red fluorescent probes (SNAP-CFl, SNAP-VO, and pHuji) and similar to SEP (Supplementary Fig. 3a-f). We also tested whether the semisynthetic pH sensor system could be used in multicolor imaging experiments with GFP-based indicators. We co-transfected neurons with GCaMP6f 16 and VAMP2-SNAP, which we labeled with CFl ligand 4. This allowed simultaneous imaging of both calcium ion transients and vesicle fusion in the same cell ( Supplementary Fig. 3g-j). Finally, we tested the ability to monitor exocytosis by labeling another SV protein, VGluT1, which has been used previously with SEP to monitor SV exocytosis 17 . The three probes (VGluT1-SNAP-CFl, VGluT1-SNAP-VO, and VGluT1-pHuji) were able to report SV exocytosis ( Supplementary Fig. 3k-m).
Imaging of endogenous protein exo-/endocytosis cycling. One interesting feature of small organic fluorophores is their ability to label not only genetically encoded domains, such as SNAP-tag, but also ligands and antibodies and hence endogenous proteins. To enable imaging of endogenous vesicular proteins, we labeled a monoclonal antibody that recognizes a luminal epitope of synaptotagmin 1 (Syt1), a SV protein, with VOAc 2 -NHS ester (6, Supplementary Fig. 1b) followed by mild deprotection of the acetate esters using hydroxylamine (Fig. 4a). This antibody has previously been used to detect endogenous Syt1 present on the plasma membrane after exocytosis in active synapses 18 . To mark vesicular Syt1, we incubated neurons with this antibody-VO conjugate (10 nM) for 3 h in stimulation buffer, followed by extensive washing to remove the unbound antibodies (Fig. 4b).
The antibody labeling was done in neurons transfected with Syt1-SEP or VAMP2-SEP to compare the performance of this labeling technique with the overexpressed, genetically encoded GFP-based pH sensors. Electrical stimulation evoked a robust increase in fluorescence in axons transfected with Syt1-SEP (Fig. 4c, d) or VAMP2-SEP (Fig. 4e), and in axons of untransfected neurons without detectable SEP. We found that the decay kinetics after stimulation was faster for the VO-antibody conjugate (8.8 ± 0.5 s) than the Syt1-SEP (12.2 ± 0.9 s, n = 41; paired t-test p < 0.0001, Fig. 4d) and VAMP2-SEP (Fig. 4f), suggesting a difference in overexpressed vs. endogenous protein behavior after SV exocytosis 19 . Remarkably, the fluorescence transients were substantially higher in untransfected than in transfected neurons (Fig. 4d, e), perhaps stemming from steric hindrance of the overexpressed SEP proteins or through quenching of the two fluorophores.

Discussion
We have developed new "semisynthetic" pH-sensitive proteins that allow for the imaging of synaptic vesicle fusion events in living cells. This sensor system combines the highly tunable properties of small-molecule fluorophores with the specificity of self-labeling tags or antibodies. The SNAP-tag-based system constitutes the first genetically encoded long-wavelength pH sensor with similar or better performance than SEP in different cell types. The antibody-based pH sensor allows for imaging of vesicle fusion events without the need for overexpression of sensor proteins. Addition of other self-labeling or epitope tags by genome editing methods could allow cell-and protein-specific labeling without the need for overexpression 20 . Future improvements of both the protein and the dye within this semi-synthetic scaffold should further enable imaging of this key biological process in increasingly complex systems.

Methods
General organic synthesis methods. Commercial reagents were obtained from reputable suppliers and used as received. All solvents were purchased in septumsealed bottles stored under an inert atmosphere. All reactions were sealed with septa through which a nitrogen atmosphere was introduced unless otherwise noted. Reactions were conducted in round-bottomed flasks or septum-capped crimp-top vials containing Teflon-coated magnetic stir bars. Heating of reactions was accomplished with a silicon oil bath or an aluminum reaction block on top of a stirring hotplate equipped with an electronic contact thermometer to maintain the indicated temperatures.
Reactions were monitored by thin layer chromatography (TLC) on precoated TLC glass plates (silica gel 60 F 254 , 250 µm thickness) or by LC/MS (Phenomenex Kinetex 2.1 mm × 30 mm 2.6 μm C18 column; 5 μL injection; 5-98% MeCN/H 2 O, linear gradient, with constant 0.1% v/v HCO 2 H additive; 6 min run; 0.5 mL/min flow; ESI; positive ion mode). TLC chromatograms were visualized by UV illumination or developed with anisaldehyde, ceric ammonium molybdate, or KMnO 4 stain. Reaction products were purified by flash chromatography on an automated purification system using pre-packed silica gel columns or by preparative HPLC (Phenomenex Gemini-NX 30 × 150 mm 5 μm C18 column). Analytical HPLC analysis was performed with an Agilent Eclipse XDB 4.6 × 150 mm 5 μm C18 column under the indicated conditions. High-resolution mass spectrometry was performed by the Mass Spectrometry Center in the Department of Medicinal Chemistry at the University of Washington and the High Resolution Mass Spectrometry Facility at the University of Iowa.
NMR spectra were recorded on a 400 MHz spectrometer. 1 H and 13 C chemical shifts (δ) were referenced to TMS or residual solvent peaks, and 19 F chemical shifts (δ) were referenced to CFCl 3 . Data for 1 H NMR spectra are reported as follows: chemical shift (δ ppm), multiplicity (s = singlet, d = doublet, t = triplet, q = quartet, dd = doublet of doublets, m = multiplet), coupling constant (Hz), integration. Data for 13 C NMR spectra are reported by chemical shift (δ ppm) with hydrogen multiplicity (C, CH, CH 2 , CH 3 ) information obtained from DEPT spectra. Original spectra for all reported 1 H and 13 C NMR data are given in Supplementary Figs. 4-26. Since the amount of compound 5 was not enough for 13 C NMR, only LC/MS analysis was performed (Supplementary Fig. 27).
Live cell imaging and analysis of PC12 cells. Cells were imaged in buffer containing 130 mM NaCl, 2.8 mM KCl, 5 mM CaCl 2 , 1 mM MgCl 2 , 10 mM HEPES and 10 mM glucose. pH was adjusted to 7.4 with 1 N NaOH. The stimulation buffer contained 50 mM NaCl, 105 mM KCl, 5 mM CaCl 2 , 1 mM MgCl 2 , 10 mM HEPES and 1 mM NaH2PO 4 . pH was adjusted to 7.4 with 5 M KOH. Experiments were carried out at 25°C using TIRF microscopy 14,23 . Cells were imaged on an inverted fluorescent microscope (IX-81, Olympus), equipped with a ×100, 1.45 NA objective (Olympus). Lasers (488 and 561 nm) (Melles Griot) were combined and passed through a LF405/488/561/635 dichroic mirror. The laser was controlled with an acousto-optic tunable filter (Andor). Emitted light was separated using a 565 DCXR dichroic mirror on the image splitter (Photometrics), and projected through 525Q/50 and 605Q/55 filters onto the chip of an EM-CCD camera. Image acquisition was done using the Andor IQ2 software. Images were acquired sequentially with alternate 488 and 561 nm excitation at 100 ms exposure. The red and green images were aligned post acquisition using projective image transformation 14,23 . Before experiments, 100 nm yellow-green fluorescent beads (Invitrogen) were imaged in the green and red channels, and superimposed by mapping bead positions.
Image analysis was performed using Metamorph (Molecular Devices) and custom scripts on MATLAB (Mathworks). The co-ordinates of the brightest pixel in the first frame of each fusion event in the green channel was identified by eye, and time was normalized to 0 s. A circular ROI of 6 pixels (~990 nm) diameter and a square of 21 pixels (~3.5 µm) were drawn around the fusion co-ordinates. The average minimum pixel intensity in the surrounding square from five frames before fusion was subtracted from the intensity in the circular ROI, and the values were normalized to the frame before fusion in the green and red channels.
Experiments were performed on an inverted microscope (IX83, Olympus) equipped with an Apochromat N oil ×100 objective (NA 1.49). Images were acquired with an electron multiplying charge coupled device camera (QuantEM:512SC; Roper Scientific) controlled by MetaVue7.1 (Roper Scientific). Samples were illuminated by a 473-nm laser (Cobolt) for green imaging, as well as by a coaligned 561-nm laser (Cobolt) for red imaging. Emitted fluorescence was detected after passing filters (Chroma Technology Corp.): 595/50 nm for pHuji, CFl and VO imaging, and 525/50 nm for SEP/GFP imaging. Simultaneous dual color imaging was achieved using a DualView beam splitter (Roper Scientific). To correct for x/y distortions between the two channels, images of fluorescently labeled beads (Tetraspeck, 0.2 µm; Invitrogen) were taken before each experiment and used to align the two channels 5 . Time lapse images were acquired at 1 or 2 Hz with integration times from 50 to 150 ms.
Image analysis was performed with custom macros in Igor Pro (Wavemetrics) using an automated detection algorithm 22 . The image from the time series showing maximum response during stimulation was subjected to an "à trous" wavelet transformation. All identified masks and calculated time courses were visually inspected for correspondence to individual functional boutons. The intensity values were normalized to the 10 frames before stimulation in the green and red channels. Photobleaching in the red channels was corrected using an exponential decay fit applied on the non-responsive boutons. All data are represented as mean ± s.e.m. of the specified number of replicates in text.