In this chapter we describe the technical details of using these probes to monitor inositol lipids in live cells. Most of this methodology requires basic equipments, which are now widely available.
3.1. Expression of the fusion proteins in mammalian cells
The easiest method for delivery of the fluorescent riporter into the cell is by transfection with a mammalian expression plasmid. Alternative methods used for delivering the construct are microinjection of cRNA or the bacterially expressed and purified fusion proteins. However, these latter methods require more preparations and a microinjection apparatus and will not be discussed in this chapter. For most microscopy application 5 × 105 cells in 2 ml culture medium are plated onto 25 mm glass coverslips (PG Science, # 60–4884–25). The coverslips are usually precoated with Poly-lysine (Sigma, #P-8920) by preparing a 1:50 or 1:100 dilution of poly-lysine in sterile deionized water (some cells require the higher concentrations, but others do not tolerate it well) and adding 1.0 ml of the diluted poly-lysine for 1 h to the coverslips that had been rinsed with 98% ethanol and air dried. After 1 h treatment, the poly-lysine is removed and the coverslips are air-dried and are ready fro plating.
Cells are grown for 1–2 days in the appropriate medium containing 10% FBS and antibiotics before transfection. The preferred transfection method varies between cell-to-cell and even among laboratories. For COS-7 and HEK293 cells we routinely use Lipofectamine 2000 (Invitrogen) or Fugene (Roche) and follow the manufacturer’s recommended protocols. The time of transfection can also depend on the application. However, we do not recommend keeping the cells longer than 24 h after transfection because of the potential toxicity of the lipid-binding fusion protein due to interference with the functions of the lipids that they bind. An important control that we always recommend doing is to use a mutant version of the lipid-binding domain that does not bind lipids. Many constructs also localize to the nucleus, but this localization is not dependent on lipid binding based on the fact that the mutant versions also show nuclear localization. It is also recommended that one simultaneously transfect cells with the GFP protein alone without the lipid-binding domain. These two controls help to track phenotypic changes and potential cellular toxicity associated with overexpression of the lipid-binding domain, as well as serve to serve as controls for monitoring localization that depends on lipid binding
3.2. Confirming the integrity of the expressed protein using a phosphorimager
For this, we use cells seeded in 12 well plates (2 × 105 cells/2ml) and transfected the next day with plasmid DNA. After 24 h, cells are washed with 2 ml of PBS and dissolved in 75 μl of SDS-PAGE loading Laemmli buffer. Cells are sonicated to disrupt the DNA but are not boiled. A 25 μl aliquot is loaded on a small (10 cm) SDS-PAGE gel at an acrylamide concentration that will resolve proteins in the size range of interest. After running, the gel is removed from the casing and is subjected to scanning in its wet-form in a phosphorimager using the blue laser line. It is our experience that the electrophoretic mobility of the EGFP molecule (or the fusion protein) can be very different from their calculated mass. This is due to altered migration of the non-denatured protein due to conformational effects. For accurate size analysis, it is recommended that samples are boiled and, after SDS-Page, detected with an anti-GFP antibody using standard protocols.
3.3. Analysis of transfected cells in the microscope. Fixed or live cells?
Once the plasmids are completed and the fusion protein is found to be intact, transfected cells can be observed under the microscope. Cells can be analyzed either live or fixed, and there are pros and cons for both cases. Fluorescence can persist in fixed cells under proper fixation conditions for a period of time. Fixed cells can be also processed for immunostaining with an anti-GFP antibody that is often more sensitive than the GFP fluorescence. Immunostaining can also help determine co-localization of the GFP construct with markers for which antibodies are available. Also, fixed cells can be stored and studied when convenient. However, one disadvantage of using fixed cells is that changes cannot be followed as they happen in real time in stimulated cells. Moreover, fixation and permeabilization procedures may distort cellular morphology. For example, we find that vesicular structures shrink during fixation, and long canaliculi can turn into small vesicles. The use of live cells definitely is the most reliable way of assessing undistorted morphology, but it is also the most time-demanding and least efficient in terms of data collection. For live cell imaging, it is best to use an inverted microscope. New water-immersion objectives make it possible to look at cells in upright microscopes, but it requires to that the bulky and difficult-to-clean objective be immersed in the culture medium.
3.3.1. Fixing and immunostaining of cells expressing GFP fusion proteins
Cells are rinsed with 2 ml PBS before the addition of 2 ml 2% paraformaldehyde (PFA) solution for 10 min at room temperature. This solution should be freshly made by dissolving electron microscopy (EM) grade PFA in phosphate-buffered saline (PBS) followed by heating in a chemical hood to 60 °C until fully dissolved with loose cover to allow gas to escape. After cooling to room temperature the pH is adjusted to 7.4 with NaOH. After the PFA incubation cells are washed three times with 2 ml of PBS for 10 min each. At this point the cells can be examined under the microscope for the GFP fluorescence or can be further processed for immunostaining. For this the cells are incubated for 10 min at room temperature with 2 ml of Blocking Solution (10% solution of FBS in PBS) to block nonspecific antibody binding. This is followed by addition of the primary antibody diluted appropriately in Antibody Diluent (10% FBS and 0.2% Saponin in PBS). The cells are incubated with the primary antibody for 1 hour at room temperature. (100 μl of diluted antibody should be sufficient for a 25 mm circular or 22×22 mm square coverslip, if it is inverted on a glass slide and incubated in a humidified petri dish). After the 1st antibody incubation, cells are washed three times with 2 ml of Blocking Solution for 5 min each before adding the fluorescent secondary antibody also diluted in Antibody Diluent. Incubations continue for 1 hour at room temperature protected from light. Finally the cells are washed three times with 2 ml of PBS and air dried until the coverslips are only damp. The coverslips are then mounted with the cells down on a glass slide using Aqua Poly/Mount and sealed on the slide with clear nail polish to prevent drying.
3.4. Examination of the transfected cells in the microscope
With the rapid advances in microscope technology there are different ways the cells can be studied depending on the application and the purpose of the experiment. However, it is always advised that cells are first examined in a wide-field fluorescence microscope. In fact, it is more difficult to get a general impression of how cells expressing the construct look in specialized microscopes, such as a confocal microscope or TIRF, than in a fluorescence microscope. Single cells, especially COS cells, show enormous variability in their shape, size, and general appearance, and often the level of expression changes their appearance. This morphological variability of cells is the largest problem when one would like to determine the effects of the expressed proteins on cellular morphology. Conventional fluorescence microscopy is a significantly more efficient way to browse through many cells and notice trends in cellular morphology. Moreover, many cells are flat in culture (especially COS cells), so there is a limited benefit in analyzing the cells in a confocal microscope. Confocal microscopy can be saved for recording cells and changes in fluorescence distribution once the conditions have been optimized with a fluorescence microscope. An additional advantage of viewing the cells with a fluorescence microscope is that autofluorescence often can be distinguished from the GFP signal because its color is different from the color of GFP. It is important to remember that confocal microscopes measure light intensity, but no colors; the “color” that is given is artificial. Therefore, in each case, the autofluorescence has to be determined so that the fluorescence signal can be reliably used. For this, observation of a set of untransfected cells is a very useful control.
3.4.1. What kind of cells should be selected for analysis?
Looking into the fluorescence microscope, there is a wide range of cells with varying fluorescence intensities (). Depending on the quality of the microscope and the intensity of the light source, sometimes only the cells with the highest expression levels are visible and these are the cells one would like avoid. It is always best to study cells in which the fluorescent signal is as low as possible but is still clearly distinguishable from the autofluorescence. High expression levels of lipid probes exert profound effects on the cells and also increase the background of signal in the cytosol as the lipid binding sites may become saturated. The best way to be certain that even the lowest level of expression is visible in the microscope is to find the autofluorescence of untransfected cells. There are no general rules for this as the sensitivities of confocal microscopes or of the cameras attached to conventional fluorescence microscopes vary from manufacturer to manufacturer. However, the following practices are recommended: 1. Observe the transfected cells (and if necessary, the control untransfected cells) with the fluorescence setting of the confocal microscope. (a separate untransfected cell control is not always necessary, because not all cells will express the fusion proteins). The untransfected cells in the population can usually be distinguished by their autofluorescence, allowing easy identification of transfected cells from the same sample. In COS-7 cells the autofluorescence (in the GFP/FITC setting) appear as small vesicular puncta most densely arranged around the nucleus probably originating from the peroxysomes). 2. Choose healthy looking cells that express protein levels just enough so that the signal can be clearly resolved from the background autofluorescence. Analysis of cells that show signs of toxic effects induced by the expression of the fusion protein should be avoided. For example, cells round up or contain large intracellular vesicles when they express high levels of the PLCd1PH-GFP construct. 3. A good quality image should be obtained with no more than 3% to 5% of the maximum laser power (assuming ~ 30 mW lasers provided with many confocal microscopes) to avoid photobleaching and damaging the cells. Higher % power may be required in the red laser as some red lasers (543 nm) have lower overall power and the red detectors are usually less sensitive.
Figure 2 Selection of cells expressing moderate levels of PLCδ1PH-GFP. Panels a-c show the same imaging field with increasing gain of HEK293 cells transfected with PLCδ1PH-GFP for 24 h. Note that the cells we would select for imaging (shown enlarged (more ...) 3.4.2. Time-lapse analysis of live cells
The most satisfying and exciting aspect of inositol lipid imaging is working with live cells. However, this also is the most time consuming and technically demanding. First of all, live cell imaging requires keeping the cells at the proper temperature. Some cellular processes, such as hormone-induced PLC activation are robust enough to be recorded at 22–25 °C as shown by many studies on Ca2+
signaling. However, trafficking steps such as receptor endocytosis is very inefficient below 30 °C. Therefore, it is desirable to run experiments at a higher temperature. An important point to remember is that even when the medium in the observation chamber is kept at the desired temperature by a heated stage, the objective in an inverted microscope acts as a heat sink keeping cells just within the observation field at a temperature of the objective. Because lasers require proper cooling and, therefore, the microscopy rooms are usually kept on the colder side, this means that even on a heated stage the recorded cells are examined at room temperature. Therefore we recommend the use of objective heaters (available from Bioptechs http://www.bioptechs.com
). However, the heater collar does not fit all objectives, and heating may be damaging to the objective if it is warmed very fast from a cold temperature. Alternative methods of maintaining the proper temperature include perfusing the cells with a high flow of warm medium and using a hair dryer to keep the objective at the proper temperature during live cell imaging. There are complete incubator enclosures available from various companies that can keep both the temperature and CO2
concentration of cells on the stage at the desired levels. Unfortunately, they make manipulations of the cells often difficult.
We use the metal Atto chambers from Molecular Probes (now part of Invitrogen) as holders in a heated stage and an objective heater. After securing the coverslips with the cells in the metal chamber we add 1 ml of prewarmed (37°C) Modified Krebs-Ringer solution (NaCl, 120 mM; KCl, 3.7 mM; Na2HPO4, 1.2 mM; CaCl2, 1.2 mM; MgSO4, 0.7 mM; Glucose, 10 mM; Na-Hepes, pH 7.4, 20 mM; Bovine serum albumin (BSA) 0.1%). Ideally, one should include cells transfected with lipid-binding fusion protein construct, nontransfected control cells, and cells transfected with a control construct that does not bind lipids.
Most current softwares allow recording of time-lapsed images. We usually record at every 5 s to 20 s (depending on the speed of the response), and with a scanning speed of 1.6 μs/pixel, to capture the cellular response with proper image resolution. Remember that the higher the resolution and the more frequent the scans are the more photobleaching and phototoxicity of the cells occur. Addition of stimuli or inhibitors is another technical challenge. The ideal way to stimulate cells is with a constant perfusion system with small dead-volumes and a valve-system that can change the composition of the medium. However, our experience is that with many of the lipophilic compounds (ionomycin, thapsigargin, rapamycin etc.) it is extremely difficult to wash the system with the valves and plastic tubes clean. Washing the Atto chambers with their plastic oring is already a challenge after using these compounds. This minor detail is often overlooked producing all kind of hard to explain artifacts.
3.4.3. Quantification of the data
The most demanding part of the analysis of time-lapsed sequences is the quantification of data. When the fluorescence redistribution is obvious a series of pictures or movies might be sufficient to describe what is happening. However, when determining a dose-response effect or comparing the relative effectiveness of two stimuli, or investigating the efficacy or potency of an inhibitor, it is necessary to quantify the changes. The most common way of this analysis is to create a line-intensity histogram through a selected line spanning the image (). The highest intensities should not saturate the detector, which requires a fine optimization of the dynamic range before recording, especially with 8-bit systems that only have 256 levels of intensity. Saturation is less of a problem with the 12-bit systems that have 4096 levels of intensity. A simple way of quantifying the translocation of a fluorescent protein from the membrane to the cytosol is to monitor the cytosolic pixel intensity of selected cells ( shows an example). A more accurate but also more demanding way of quantification of the extent of membrane association of fluorescent proteins is to use fluorescence resonance energy transfer (FRET) or total internal reflection fluorescence (TIRF) microscopy. The former is able to detect the radiationless energy transfer between two appropriate fluorophore pairs when they are within an optimal distance (see below), while the latter only detects fluorescence coming from the plane of the membrane. The variability of the cell population and the requirement for analysis of a large number of cells to obtain reliable quantitative estimates of the fluorescence changes remain the most laborious part of obtaining reliable, reproducible results.
Figure 3 Quantification methods to assess changes in plasma membrane localization of PLCδ1PH-GFP. (A) Changes in membrane localization after angiotensin II (AngII) stimulation as assessed by confocal imaging of a HEK293 cell shown on the left. The right (more ...) 188.8.131.52. FRET measurements
It is not easy to make quantitative assessment of the membrane localization of fluorescent probes. In fact, some “localization” is a reflection of change in membrane volume or change of membrane shape as opposed to real change in recruitment. Therefore, several attempts were made to utilize the FRET principle to obtain a signal based upon the binding of an inositide binding domain to membranes. The simplest method we used was to co-express the CFP- and YFP-tagged versions of the same PH domain (e.g. PLCδ1-PH). CFP and YFP are the most widely used pairs of fluorophores that when come within FRET distance (<8 nm) will show energy transfer from CFP to YFP. This will cause a decrease in CFP emission (475 nm) and increased YFP emission (525 nm) when only CFP excitation (430 nm) is applied. The efficiency of the energy transfer can be numerically calculated after making all necessary corrections (such as bleed through of the CFP and YFP signals into the other pair’s emission channels). However, if the two wavelengths show opposite changes, the simple fluorescence ratio of 525/475 can be used (). When the two fluorophores are bound to the lipids at the membrane via the attached PH domains there is an efficient energy transfer. However, upon PLC activation the molecules leave the membrane and the FRET signal decreases (34
). This method is quite sensitive and even small PLC activation can be detected and quantified. It can also be used in individual cells or in cell suspensions (35
). A disadvantage of this method is that at low expression levels (which would be desirable to minimize the ill effects of the presence of the probes) the FRET efficiency is not very high and even at high probe concentration there is low FRET signal if the density of the lipids is below a certain level.
In contrast to the above examples using two separate molecules that show intermolecular FRET, a few attempts have been made to generate inositol lipid probes based on intramolecular FRET. In this case, both fluorophores are attached to the same inositide-recognizing domain. Lipid binding then induces an alteration in the distance (or more likely the dipole orientation) of the fluorophores and, hence, a change in the FRET signal. A probe based on the Grp1-PH domain was targeted to different membranes for PtdIns(3
) and a similar principle was utilized to generate FRET probes for monitoring InsP3
changes in the cytoplasm (37
). The challenge in this type of molecular engineering is to make sure that the conformational change upon lipid binding is sufficiently big to change the FRET signal. Construction of a useful probe requires lots of experimentation with the domains themselves as well as with the linkers to connect the fluorophores. In a recent study, the AktPH domain was used to detect PtdIns(3
changes using a unique molecular design. The conformational change between the lipid-bound and unbound stages was achieved by inserting a negatively charged “pseudoligand” in the probe that binds to the PH domain (presumably to the lipid binding site) when lipids are not present. Binding of the appropriate lipids abolishes this intramolecular interaction amplifying the conformational change and generating a larger change in the FRET signal (39
). These single molecule FRET probes do not require coexpression, their readout does not depend on their expression level (once above reliable detection limits) or on lipid density in the membrane. It is expected that more efforts will be made to generate similar probes for the detection of lipid production in the various cellular compartments.
More detailed technical and theoretical background on FRET measurements either with sensitized emission or with fluorescence lifetime imaging microscopy (FLIM), including corrections for bleed-through and uneven illumination, can be found in very comprehensive publications elsewhere (40
184.108.40.206. Quantification of membrane association with TIRF analysis
TIRF (total internal reflection fluorescence) analysis has also been used to monitor plasma membrane association of inositol lipid binding domains (43
). This technique detects fluorescence originating only from the thin membrane area of the cell attached to the coverslips making it a suitable method of detecting membrane-associated fluorescence (). One caveat of this method is that anything that changes the footprint of the cell would cause a change in fluorescence intensity unrelated to the actual amount of fluorescent molecules at the membrane. For this reason, for most accurate results it is desirable to use a fluorescent membrane marker and relate all fluorescence change to this reference signal.