Key words

1 Introduction

Second messengers such as Ca2+, IP3, cGMP, and cAMP are critical intermediates that relay signals from membrane-bound receptors to intracellular effectors. Cyclic adenosine monophosphate (cAMP ) plays a key regulatory role in most types of cells; however, the pathways controlled by cAMP may present important differences between organisms. In rare cases, such as the slime mold Dictyostelium [1], cAMP can also convey extracellular signals.

Intracellular concentrations of cAMP change rapidly when it is synthesized from ATP by a family of adenyl cyclases (ACs) or degraded by phosphodiesterases (PDEs). Mammalian ACs are either cytosolic or membrane bound and regulated via (1) phosphorylation by PKA , PKC, and calmodulin-dependent protein kinases (CaMK), (2) second messenger molecules such as Ca2+ and (3) via protein–protein interactions. A prominent mode of activation for mammalian membrane-bound ACs is through interaction with heteromeric G proteins, which in turn are triggered when extracellular signals, such as hormones and neurotransmitters, bind to G protein-coupled membrane receptors (GPCRs). The alpha subunit of the heteromeric G protein (Gα) exchanges GDP for GTP, and as a consequence, dissociates from the G protein complex into Gα and Gβγ. G protein subunits then bind ACs, and depending on the specific subunits and the type of AC, the cAMP producing enzymes are either activated or inhibited [2, 3].

The cAMP concentration is reduced via hydrolysis by members of the PDEs family. Certain PDEs are selective for cAMP , while others can only degrade a closely related second messenger, cGMP (cyclic guanosine monophosphate); the majority however is capable of degrading both cGMP and cAMP . Just as the ACs, PDEs are regulated by the interplay with a variety of cellular signals such as phosphorylation, small signaling molecules, and protein–protein interactions [4]. PDEs have been associated with several diseases, including heart failure, depression, asthma, and inflammation, and many drugs are developed that selectively target PDEs [5]. It is therefore important to keep developing precise real-time measurement techniques for cyclic nucleotide second messengers, that can help elucidate the regulation of their turnover in healthy and diseased cells.

Genetically encoded intramolecular biosensors are powerful tools to study second messenger concentrations with high temporal resolution. Förster resonance energy transfer (FRET) is the non-radiative transfer of energy from a donor fluorophore to an acceptor fluorophore [6] and can directly report protein conformational changes, thus providing a base for a large variety of fluorescent biosensors. Development of these biosensors started around two decades ago, with pioneers such as a calmodulin-based sensor [7] and troponin C-based sensor [8] for Ca2+, cygnet probes for cGMP [9] (with the regulatory domain of PDEs), sensor based on the cGMP-binding domain B from cGMP-dependent protein kinase (GKI) [10], and cAMP sensors based on protein kinase A (PKA) [11] along with the EPAC-based probes [12,13,14].

EPAC1 is a guanine nucleotide exchange factor for Rap1 that is activated by direct binding of cAMP [13]. EPAC1 has an N-terminal DEP (Dishevelled, Egl, Pleckstrin) domain that is essential for membrane localization, a cAMP-binding domain, a REM domain (Ras exchanger motif), and a C-terminal GEF catalytic domain (guanine exchange factor) that regulates the GDP/GTP binding affinity of the Ras-like protein Rap. The EPAC2 protein is identical in domain structure except for a second N-terminal cyclic nucleotide monophosphate (cNMP) binding domain [15].

Several groups reported on EPAC-based cAMP FRET sensors [16]. Nikolaev et al. [14] made a compact FRET sensor by fusing the cyclic nucleotide binding domains of EPAC1 and EPAC2 in-between donor and acceptor fluorophores. The full-length EPAC1 protein was also used [12]. In our design, we deleted the membrane-binding DEP domain (ΔDEP) of EPAC1, and we also introduced point mutations to render EPAC catalytically inactive (CD, T781A, and F782A) so as to prevent unwanted downstream signaling to Rap1 and Rap2 [13, 17]. This proved a good strategy as this configuration yields very robust FRET changes upon cAMP binding. In our original sensor, EPAC (ΔDEP, CD) was sandwiched between CFP and YFP donor and acceptor fluorophores [13]. Since then, we have reported several rounds of optimization [18,19,20].

FRET is extremely sensitive to distance: typically, a fluorescent protein needs to be in the range of 1–10 nm for FRET to occur. This characteristic distance range makes FRET ideal to measure protein–protein interactions as well as conformational changes within proteins in living cells. The most common techniques to read out FRET are fluorescence ratiometry and fluorescence lifetime imaging (FLIM). Ratiometry can be carried out with relatively simple and widely available equipment by recording both donor and acceptor emissions. However, ratiometric measurements are not fully quantitative unless endpoint calibrations are performed or quite elaborate corrections are carried out [21, 22]. FLIM recording, on the other hand, is a much more robust and inherently quantitative technique [23].

FLIM reports on FRET because FRET shortens the donor lifetime. FLIM has the important advantages that lifetimes generally are independent of concentration, bleaching, or excitation fluctuations. However, fluorescence lifetimes of excited fluorophores typically are a few nanoseconds, and FLIM therefore requires complex and dedicated machinery. While being the most quantitative method for the detection of FRET , FLIM is also technically demanding. Time-correlated single photon counting (TCSPC) [24, 25] and frequency domain (FD)-FLIM [26, 27] are the two most common techniques to measure fluorescent lifetimes.

Traditional TCSPC detectors are photon efficient, but have until recently been extremely slow and hence unable to deliver the speed for biological processes occurring at time scales below tens of seconds. Such limitations in speed and experimental complexity were overcome by efforts of several groups, including our laboratory in tight cooperation with Leica Microsystems, and resulted in the development of FLIM instrumentation like the Leica SP8 FALCON [28]. This system is based on a confocal scan head with field-programmable gate array electronics, pulsed laser excitation and fast, spectral single photon counting detectors. FD-FLIM, which is commonly implemented on widefield microscopes, relies on recording a stack of images at different phases and it is inherently fast. Unfortunately, it is also photon inefficient and somewhat prone to producing artifacts when used with living cells, stemming from rapid cellular movements and signal transients. We recently also contributed in methods that overcome these issues by developing, in cooperation with Lambert instruments, a single-image method called siFLIM [29]. In live-cell time-lapse experiments, this approach provides both photon efficiency and speed as well as excellent signal to noise ratio (SNR) along with immunity to lifetime artifacts. Having designed and perfected fast and photon-efficient FLIM techniques for both confocal [28] and wide field microscopy [29] we feel that FLIM should become the method of choice for FRET-based signaling studies as it enables following large numbers of cells in real time, quantitatively, with high data content and minimal photodamage.

Here we describe how to use fast TCSPC FLIM recording to measure changes in cellular cAMP levels. We employ our EPAC-based sensor (EPAC-SH189), a reporter with a truncated version of the cyan fluorescent protein analog mTurquoise2 [30] as donor and a tandem of non-emitting, circular permutated Venus proteins (a yellow fluorescent protein analog) as acceptor [20]. The donor has an exceptionally high quantum yield and brightness, allowing for dim excitation and thus minimizing bleaching and phototoxicity. Unlike most other fluorescent proteins, the decay of excited mTurquoise2 is well fitted with a single exponent, which makes it especially suited for TCSPC FLIM . The tandem dark Venus moiety minimizes emission of the acceptor, enabling collection of mTurquoise2 emission over a wide range without contamination with acceptor signal. This biosensor has an outstanding signal-to-noise ratio and good fluorophore maturation, and it is biochemically inert. Moreover, the EPAC moiety harbors a single point mutation Q270E, which increases its affinity for cAMP . We describe a step-by-step protocol to perform a cAMP recording using the Leica STELLARIS 8 FALCON time domain FLIM confocal setup. The protocol described here should be equally applicable to recordings with other intramolecular FRET sensors.

Note that whereas this protocol focuses on TD-FLIM application to measure cAMP levels in single cells with exceptional quantitative sensitivity, we have previously provided a detailed protocol [31] on the use of FD-FLIM in combination with EPAC-based biosensors and a detailed protocol [32] for cAMP sensing by ratiometric detection of sensitized emission.

2 Materials

All solutions are made with deionized water.

2.1 Stock Solutions

  1. 1.

    1 M NaCl2.

  2. 2.

    1 M NaOH.

  3. 3.

    2.5 M CaCl2.

  4. 4.

    1 M CaCl2.

  5. 5.

    1 M MgCl2.

  6. 6.

    0.1 M KCl.

  7. 7.

    1 M Glucose (see Note 1).

  8. 8.

    1 M HEPES (see Note 1).

2.2 Disposables

  1. 1.

    0.22 μm filters.

  2. 2.

    Attofluor cell chamber (Invitrogen).

  3. 3.

    24 mm Ø, 0.17 mm thick glass coverslips (#1.5).

  4. 4.

    non-pyrogenic polystyrene tubes.

  5. 5.

    6-well cell culture plates.

2.3 Working Solutions

  1. 1.

    2× HEPES buffered saline (HBS-buffer): 280 mM NaCl, 10 mM KCl, 20 mM HEPES, pH = 7.2 at 37 °C. Magnesium (1 mM MgCl2), calcium (1 mM CaCl2), and glucose (10 mM) are added when the dilution to 1× HBS-buffer is made.

  2. 2.

    2× HBS-buffer (for transfection): 280 mM NaCl, 50 mM HEPES, 1.5 mM Na2HPO4, pH 7.2. The optimal pH depends on the cell line that is used (see Note 2).

  3. 3.

    1.5:1 W/V polyethylenimine (PEI) (MW ~ 25.000) ethanol solution. Store in glass at −20 °C (see Note 3).

  4. 4.

    1× phosphate buffered saline (PBS): 137 mM NaCl, 2.7 mM KCl, 10 mM Na2HPO4, and 1.8 mM KH2PO4.

  5. 5.

    DMEM supplemented with 10% FCS and penicillin/streptomycin (pen/strep). Use the appropriate medium with additives if other cell lines are used.

  6. 6.

    0.05% trypsin-EDTA solution.

  7. 7.

    DMEM/F-12 and/or phenol red-free Leibovitz’s L15 is used for imaging (see Note 4).

3 Methods

3.1 Microscope

Confocal microscope equipped with fast time correlated single photon counting detector(s). Here we use Leica DMI8 confocal microscope with STELLARIS 8 FALCON system. For FRET-donor excitation we use the 440 nm line of the white light laser. For FRET-donor emission we use the Power HyD detector. The microscope is operated by Leica LasX software. Images were taken with a 63× oil objective.

3.2 Cell Culture

24 mm Ø, #1.5 glass coverslips are sterilized with 70% alcohol or UV-C and placed in the wells of a 6-well plate. After drying, wells are filled with 2 mL of DMEM with FCS and pen/strep. Cells are trypsinized and resuspended in medium, and approximately 150.000 cells are added to each well (see Note 5). Depending on the cell line, cells are transfected 8–24 h after plating and cultured for another 24–72 h prior to FLIM .

3.3 Transfection

PEI transfection: 2 μL of the PEI solution is mixed with 1 μg of plasmid DNA in 200 μL serum-free medium in a polystyrene tube. Mix gently and incubate for 15–30 min at room temperature. Drop the PEI/medium mixture to the cells, and place the 6-well plate, after gentle swirling, back in the incubator (see Note 6).

Calcium phosphate transfection: Put 86 μL of 2× HBS in a polystyrene tube, and add 2–5 μg of plasmid DNA diluted in H2O (final volume is 194.9 μL). Mix gently and add 5.1 μL of 2.5 M CaCl2 and mix again. Incubate for 20 min at room temperature, and add dropwise to the cells (see Note 7). Place the cells back in a CO2 incubator. Optionally replace the medium after 24 h.

Any other commercial transfection reagent can also be used according to the manufacturer’s protocol.

3.4 Imaging

The cell chamber and imaging medium are prewarmed to 37 °C. A coverslip is taken from the 6-well plate with sharp forceps and carefully mounted in a cell chamber (see Note 8). 1–2 mL HBS+/+ (1× HBS with glucose, CaCl2, and MgCl2) is immediately added to the cell chamber (see Note 9).

Place the loaded cell chamber on the microscope. Use the appropriate immersion liquid on the objective, and focus on the cells. After focusing on the cells, wait up to 5 min to equilibrate the temperature.

3.5 TSCPC FLIM

  1. 1.

    Turn on all the hard- and software, wait for the instrument to initialize. Start up the FLIM software extension.

  2. 2.

    Adjust the software and hardware settings for EPAC sensor imaging: Select the 440 nm excitation laser line, Power HyD detector for donor lifetime detection between 450 and 550 nm, pinhole at the desired value. We used 3 Airy Units in this experiment (see Note 10).

  3. 3.

    Mount the cells on the microscope. Focus by looking through the eyepiece onto the cells or by looking at the digital image in “live” mode.

  4. 4.

    Set up the time-lapse experiment. Measuring the dynamics of changes in cellular cAMP concentration requires a temporal resolution in the order of seconds with minimal photodamage to the cells. In the current example we acquire FLIM images with 5 s frame interval.

  5. 5.

    Select a region with one or more healthy cells (see Note 11). Start the actual experiment by recording a baseline for a few frames before stimulating the cells with a concentrated stock of reagent(s) (see Note 12). Follow the changes in lifetime. Optionally, multiple stimuli can be added. When the plateau is reached after addition of the initial stimulus, wait for another few frames. In Fig. 1, the initial baseline is recorded for approximately 1 min. As the first stimulus, beta2-adrenergic receptor agonist isoproterenol is added at 5 nM final concentration resulting in an increase of intracellular cAMP and a concomitant increase of fluorescence lifetime due to the opening of the FRET sensor. The second stimulus is the same agonist at 25 nM resulting in further, yet not saturating increase in cellular cAMP levels. The final stimulation with 25 μM forskolin directly activates the ACs and increases the intracellular cAMP concentration to the maximum level.

  6. 6.

    Stop the experiment, and make sure the data is properly saved.

  7. 7.

    Photon arrival time histograms from FRET sensor data typically show a multi-exponential curve, indicating the superposition of different FRET states. These FLIM data are well described by a double-exponential fit: a high FRET state with a lifetime of 0.9 ns, and a low/no FRET state with a lifetime of 3.9 ns (see Note 13).

  8. 8.

    The values of lifetime components of the sensor can be plotted in a polar plot (Fig. 1g). The position within the polar plot gives information about several aspects of the sensor such as the fraction of sensor molecules that are in the open and closed conformation, but also on bleaching, and autofluorescence (see Note 13).

  9. 9.

    Photon arrival times registered by the Power HyD detector are fitted with the manufacturer-supplied software (Leica LasX). For the lifetime analysis image binning (e.g., 2x2 pixels) may be applied to increase pixel SNR. For each pixel the photon arrival time histogram is fitted to a double-exponential function with fixed lifetimes for both the high and the low FRET state (see step 7 and Note 13), yielding two images containing the amplitudes of these two components. These images can then be saved as TIF files, reducing the amount of raw data more than 1000-fold (from orders of GB to MB). Further analysis on the fitted lifetime data can be performed with manufacturer-supplied software, or TIF files can be exported for processing with custom scripts in, e.g., Python, Matlab, ImageJ/Fiji (see Note 14).

  10. 10.

    Remove the cell chamber from the microscope and clean the objective with a tissue. Carefully clean the cell chamber, alternatingly with water and 70% alcohol to prevent any residual compound from sticking to the metal and affect future experiments. Prolonged storage of the cell chamber is in 0.5 M NaOH (see Note 15).

  11. 11.

    After all experiments are completed, turn off all equipment.

Fig. 1
figure 1

Time-lapse FLIM imaging of HeLa cells expressing the EPAC-SH189 biosensor . (a) Mean FRET donor fluorescence intensity image with a set of ROIs drawn for single-cell analysis; (b, c, d) intensity weighted lifetime images at t = 50 s, t = 120 s, and t = 360 s; (e) lifetime traces corresponding for the selected ROIs in (a): cells stimulated at 55 s with 5 nM isoproterenol (S1), at 170 s with 25 nM isoproterenol (S2), at 280 s with 25 μM forskolin (S3); (f) lifetime decay for the first 50 s (before stimulation, green) and the last 50 s (after 25 μM forskolin, blue); (g) Polar plot: the population of sensor molecules is shifted toward the open confirmation due to rise in cellular cAMP concentration

Full size image

4 Notes

  1. 1.

    Glucose and HEPES cannot be autoclaved and have to be filtered with a 0.22 μm filter.

  2. 2.

    The optimal pH of the 2× HBS-buffer depends on the cell type used for the experiments. Therefore, a series of different pH buffers is made, ranging from 6.8 to 7.2 in steps of 0.05 and tested on exponentially growing cells (~50% confluent).

  3. 3.

    After 3 months, transfection efficiency drops dramatically. Storage at 4 °C is possible, but 20 °C is recommended to prevent evaporation. It is strongly recommended to use glass containers, since ethanol taken from an Eppendorf tube by itself sometimes affects cells.

  4. 4.

    When imaging for extended times, cell culture medium is preferred over 1× HBS-buffer. Keeping the autofluorescence minimal is important during FLIM recordings. DMEM/F-12 and/or phenol red-free Leibovitz’s L15 is a good option. DMEM/F-12 is buffered with CO2. If no CO2 is available at the microscope, Leibovitz’s L15 is a good choice. If other media are used, riboflavins and phenol red may be a major source of autofluorescence.

  5. 5.

    For a homogenous layer of cells, move the plate twice in south/north direction followed by twice in west/east direction. This will prevent cells from piling up in the middle of the well.

  6. 6.

    After 10 h, the first cells will be fluorescent, and 72 h after transfection, expression is highest. Note that PEI can be toxic to cells, and replacing the medium with fresh medium after 24 h is for some cell types advisable.

  7. 7.

    Do not exceed 20 min of incubation since large precipitates are formed which will decrease transfection efficiency.

  8. 8.

    If the forceps is not sufficiently sharp, a bent needle can be useful for lifting up the coverslip. Prevent leakage by placing the coverslip exactly in the middle of the chamber, and screw the ring tight. However, screwing too tightly can result in breaking the coverslip.

  9. 9.

    Clean the bottom of the coverslip with a paper tissue. Gently press the paper for a second time to the bottom of the coverslip to check if no leakage occurs. If the cell chamber is leaky, either the chamber is not screwed sufficiently tight, the coverslip is not in the middle of the ring or the coverslip is broken.

  10. 10.

    Pinhole (in Airy units, AU) can be opened/closed based on the brightness of the sample and the desired optical sectioning. Increasing the diameter of the pinhole enables collecting more photons at the cost of some image blurring.

  11. 11.

    Make sure the fluorescent cell looks like the non-transfected cells as an indication that they are healthy. Very bright cells may develop a different morphology, e.g., become more rounded, and these are to be avoided. An intermediate bright fluorescent cell is often a good choice for measuring.

  12. 12.

    Usually, a highly concentrated (e.g., 1000×) reagent is used to prevent changes in volume, temperature, and/or osmolarity when adding the stimulus. We mix the reagent by first pipetting a small volume with a yellow tip on a P20. Then we transfer the yellow tip carefully to a P200. Now a small drop of reagent is present in the middle of the yellow tip. To stimulate the cells, we carefully put the tip of the pressed pipette in the medium of the cell chamber and release the plunger. Then we gently mix by pipetting up and expelling 200 μL medium for six more times, while avoiding air bubbles. Do not mix directly on top of the cells as you may wash them away.

  13. 13.

    A polar (phasor) plot can show to what extent the FRET sensor is opened, whether fluorophore bleaching occurs, and the possible contribution of autofluorescence [33, 34]. Perfect mono-exponentially decaying fluorophores are located on the semicircle. Lower lifetimes are located at the right part of the semicircle. FRET sensors usually show a mixture of multiple decays and their phasors form a line within the semicircle. The sensor starts in a closed conformation and opens after increase of cellular cAMP (after addition of isoproterenol and further after addition of forskolin). In case opening of the sensor completely prevents energy transfer, the phasor ends on the semicircle for a mono-exponentially decaying donor. With decreasing cAMP concentration, the fraction of sensor molecules with high FRET increases and the phasor travels to the right and downward in this plot. 100% FRET efficiency is typically not reached with these sensors, but if it would, this would render the molecules invisible because all energy is transferred to the acceptors. The intercepts of the extrapolated line with the semicircle provide us with values for the long (3.9 ns) and the short (0.9 ns) lifetime components characteristic for this particular sensor. These values are used for fitting the photon arrival times from the experimental data to obtain quantitative FRET-donor lifetimes.

  14. 14.

    For calculating the weighted lifetimes for each pixel from the 2-component fit intensity images, users may prepare a simple custom Fiji [35] macro, where the exported intensity images for the short and the long lifetime components are multiplied with their respective values. The sum of these resulting images is then divided by the sum of original (non-lifetime weighted) intensities, yielding a final weighted lifetime image with: τ = 0.9 × I1 + 3.9 × I2/(I1 + I2), in which I1 and I2 denote the intensities of the fast and the slow component, respectively. Transferring the data to ImageJ allows for fast and convenient inspection of the data outside the LAS-X software.

  15. 15.

    Some organic compounds bind to the steel ring and affect the follow-up experiment. Repetitive washing with water and alcohol is sufficient to get rid of most compounds. Other compounds are however not fully removed by washing, and therefore, we place the steel cell chamber in concentrated NaOH for prolonged times when not in use.