A Click Cage: Organelle‐Specific Uncaging of Lipid Messengers

Abstract Lipid messengers exert their function on short time scales at distinct subcellular locations, yet most experimental approaches for perturbing their levels trigger cell‐wide concentration changes. Herein, we report on a coumarin‐based photocaging group that can be modified with organelle‐targeting moieties by click chemistry and thus enables photorelease of lipid messengers in distinct organelles. We show that caged arachidonic acid and sphingosine derivatives can be selectively delivered to mitochondria, the ER, lysosomes, and the plasma membrane. By comparing the cellular calcium transients induced by localized uncaging of arachidonic acid and sphingosine, we show that the precise intracellular localization of the released second messenger is crucial for the signaling outcome. Ultimately, we anticipate that this new class of caged compounds will greatly facilitate the study of cellular processes on the organelle level.

(E) pH dependency of coumarin fluorescence. Fluorescence spectra of compounds 6, 12, 17 and 19 were measured in PBS buffer with pH 7.4, 7, 6 and 5. Spectra are normalized to the emission observed at the physiological pH 7.4. Changes of fluorescence intensity with changing pH appear to be minor in the physiological pH range. Figure S2. Normalized HPLC elution profiles of compound 14 and compound 21 without UV illumination (red) and after 3 min illumination (blue). Elution profiles of the respective standards (sphingosine (15) or arachidonate (5)) are shown in grey. UV cleavage and HPLC analysis were carried out with diluted samples to minimize substance loss. Figure S3. Bleed-through controls for representative caged lipids. Compounds were loaded as described below (section "Loading procedures of caged compounds and organelle markers"). Left panels: Cells loaded with the caged lipid and the respective organelle marker were illuminated solely with the 405 nm laser while sampling both coumarin (blue, between 425-475 nm) and organelle tracker fluorescence (green, between 500-550 nm). Note that signal is detected only in the blue channel and no bleed-through occurs. Middle panels: The same cells were subsequently illuminated solely with the 488 nm laser while sampling both coumarin (blue, between 425-475 nm) and organelle tracker fluorescence (green, between 500-550 nm). Note that signal is only in the green channel. Right panels: The same cells were subsequently illuminated with both lasers while sampling both coumarin (blue, between 425-475 nm) and organelle tracker fluorescence (green, between 500-550 nm). Note that signal is now detected in both channels. All images were acquired using the same magnification, the scale bar indicates 20 µm. Identical settings were used for each caged lipid / organelle marker pair. Fluorescence intensities tended to be slightly lower in the last set of measurements due to photobleaching.  Figure S5. Bar graphs show Pearson correlation coefficients (PCC) of non-thresholded images and Manders correlation coefficients above the autothreshold for both channels, in every case derived from a minimum of four images. Only images that could be thresholded using a standardized automated routine were included for the determination of Manders overlap coefficients to avoid human bias. Error bars indicate standard deviation.  -table) show the subcellular localization of organelle-targeted caged lipids, right panels (red lookup-table) show the respective organelle markers. Figure S6. For absorbance, the detection range was set to 200-500 nm, the spectral resolution to 2 nm and the averaging time to 0.1 s. The path length of the cuvette was 0.1989 cm (d), baseline correction was carried out by subtraction of the background signal of an ethanol sample. Extinction coefficients were determined by plotting the maxima of a serial dilution, using a linear fit to determine the slope (m) and calculated as following:

= * 1000
Fluorescence spectra were recorded with 5-nm resolution. Excitation wavelength for all coumarin derivatives was 380 nm and emission collection at 400-600 nm, with a step size of 2 nm, and 40 µs integration. All spectra were recorded in ethanol at an analyte concentration of 62.5 µM and are background corrected for ethanol. Quantum Yields (QYs) where calculated as following: where QYref is the QY of the reference compound (coumarin 1, QY=0.73) [1] , η is the refractive index of the solvent (ethanol for all samples and the reference), A is the absorbance at the excitation wavelength, I is the integrated fluorescence intensity. Spectra were recorded using a concentration range from 0.065 mM to 0.5 mM. To observe the effect of pH on the fluorescence of the caged compounds, spectra of four representative compounds (the un-clicked derivatives 6 and 17 and the lysosomal targeted derivatives 12 and 19) were acquired at room temperature at 50 µM concentration in PBS buffer adjusted with 3% HCl to the respective pH value 7.4, 7, 6 and 5. The spectra as well as the data are shown in Figure S1.

Determination of photocleavage efficiency in vitro
Photocleavage efficiency after exposure of caged compounds to UV light was analyzed by high performance liquid chromatography (HPLC), using a Knauer Azura system (AZURA P 2.1L pump, ASM 2.1L assistant, DAD 6.1 and a Sedere Sedex 85 ELSD detector) with a reverse phase Knauer Eurosphere II 100-5 C4 column. Newly synthesized compounds 6, 11-14 and 17-21 were compared to previously reported caged arachidonate and sphingosine derivatives 22 and 23, which are equipped with the 7-(diethylamino)-4-(hydroxymethyl)-coumarin group. UV irradiation was carried out using a 1000 W Mercury-Xenon lamp equipped with a 335 nm high pass filter. Irradiation samples were prepared by diluting 20 µl of 10 mM stock solutions of the respective caged compounds in DMSO with 180 µl MeOH in a 250 µl transparent Eppendorf tube. Samples were placed in a stand 20 cm away from the light source. For each compound a sample was illuminated for 1, 3 and 5 min, respectively. Samples were analyzed by semi-preparative reversed phase-HPLC directly after illumination (conditions below). Elution conditions were chosen to enable analysis of the photo-liberated parent molecules and the caged precursors during the same HPLC run (Table S1). Eluting compounds were detected by light scattering using a Sedere Sedex 85 ELSD detector and retention times (Rt) of the free parent molecules sphingosine (15) and arachidonic acid (5) and their caged counterparts were determined. During the analysis of the 5 min UV irradiation sample set, peaks matching the retention times of arachidonate and sphingosine were collected and analyzed by high resolution mass spectrometry to confirm the identity of the photo-liberated molecules. Results of the photocleavage analysis are summarized in table S2. Elution profiles are exemplary shown for arachidonate, sphingosine, and compounds 14 and 21 before and after 3 min UV irradiation in Figure S2. Caged lipids were stored as 10 mM DMSO stock solutions, whereas organelle markers were stored as 1 mM DMSO stock solutions. Caged lipid loading solutions for cellular uptake were generated by appropriate dilution with imaging buffer. HeLa cells were grown to 95% confluence in 8-well Lab-Tek dishes. The growth medium was removed prior to compound loading, and the cells were washed with imaging buffer (1 x 200 µl per well). Subsequently, the cells were treated with the respective caged lipid loading solutions (200 µl per well) for the time indicated in Table S1. The loading solution was removed, the cells washed with imaging buffer (1 x 200 µl per well) and subsequently treated with a solution of the respective organelle marker in imaging buffer. The organelle marker concentrations for each case are given in Table S3. The subcellular distributions of caged lipid probes and organelle markers were assessed using dual color confocal fluorescence microscopy on an Olympus FV1000 setup (see previous section for details). The two channels were acquired in sequential mode (line wise) to avoid bleed-through. Caged lipids were stored as 10 mM DMSO stock solutions. Caged lipid loading solutions for cellular uptake were generated by appropriate dilution with imaging buffer. HeLa cells were grown to 50% confluence in 8-well Lab-Tek dishes. The growth medium was removed prior to compound loading, and the cells were washed with imaging buffer (1 x 200 µl per well). Subsequently, the cells were treated with the respective caged lipid loading solutions (200 µl per well) and incubated for 10 min at 37 °C. The loading solution was removed, the cells washed twice with imaging buffer, either kept at 37 °C for additional 20 min (for assessment of compound uptake) or incubated with Fluo-4-AM (5 µM Fluo-4-AM in imaging buffer) at 37 °C for 20 min. Prior to imaging, the cells were washed once more with imaging buffer. The optimized loading conditions chosen for calcium imaging are displayed in Table S4, comparable loading was achieved for compounds 11-13 and 18-20, whereas incorporation of compounds 14 and 21 was found to be either slightly (21) or significantly (14) less efficient (compare Figure S6).

Colocalization analysis
The colocalization plugin Coloc2 for the ImageJ/Fiji software was used to generate 2D histograms and to determine Pearson correlation coefficients and Manders overlap coefficients. Noncorrected and non-thresholded images of caged lipid probes and respective organelle markers were loaded into the plugin. Mean Pearson correlation coefficients for each caged lipid/organelle marker pair were determined from the entire dataset. Thresholding was carried out automatically using the "automatic-biscetion" settings. Images were not included for the calculation of mean average thresholded Manders overlap coefficients, if the automatic threshold determination failed to reduce bias introduced by manually chosen threshold values.
Confocal time lapse live cell microscopy and uncaging assay For calcium imaging experiments (Figures 2, S7), cells loaded with caged compounds and Fluo-4-AM (as described above) were imaged in eight-well microscopy dishes (80826, Ibidi) using a Zeiss LSM800 confocal microscope with a Plan-Apochromat 63x/1.40 Oil objective at ambient temperature in imaging buffer containing 20 mM HEPES, 115 mM NaCl, 1.8 mM CaCl2, 1.2 mM MgCl2, 1.2 mM K2HPO4, 10 mM glucose. Coumarin dyes were excited using a 405 nm laser (at 1.0% laser power) and emission was collected between 400 nm and 590 nm. The calcium dye Fluo-4 was excited using a 488 nm laser (at 1.2%) and emitted light was collected between 400 and 590 nm. Coumarin and Fluo-4 fluorescence was acquired sequentially in line-mode. Timelapse images were acquired using the Zen Blue software and the Experiment Designer plug-in set at one frame per second. Uncaging was performed after acquiring a 10 frames baseline, using widefield-illumination of the whole field-of-view by the 385 nm LED of the Colibri 7 module set to 30 % power for 1.5 s. After uncaging, time-lapse images of Coumarin and Fluo-4 fluorescence were acquired for 60 more frames. In the end, baseline and post-uncaging frames were stitched using the Timestitch function in the Experiment Designer plug-in and the resulting time-lapses were exported as .czi files.

Data analysis of calcium time lapses
The acquired time lapse series were analyzed with the Fiji software using the FluoQ macro [2] set to the following parameters: Background subtraction method: Do not remove background Noise reduction / smoothing method: None Threshold method: Interactively with ImageJ's built-in threshold window ROI segmentation: Semi-automatically with binary mask modification Calculate amplitude changes: Using maximum observed amplitude change The maximum amplitude values were calculated by subjecting the raw traces to a central moving average. The maximal amplitude x �������� ��� of these smoothed traces was used to calculate the amplitude change in percent%∆ � ��� according to the following formula: The resulting intensity series/amplitude values represent mean values of whole cells and were further analyzed using R. Figures 2 and S7 were prepared using the ggplot2 package [3] .