Prof. Francolini Flashcards
synthetic calcium indicators
In general, calcium indicators allow us to understand the amount of free calcium or bound calcium to specific probe, within the cell. It is possible because in free or bound state, the excitation and the emission spectra might change. Knowing the dissociation constant (Kd) of the sensor, it is possible to evaluate the amount of free or bound calcium. In particular, synthetic calcium indicators are the result of artificial manipulation (they were developed from ’60 and ’70) from similar molecule structures. Synthetic calcium indicators are fluorescent molecules as sensors able to bind calcium. They are very sensitive and versatile. These indicators came from the hybridization of highly calcium-selective chelators, such as EGTA (that has several carboxyl groups able to bind calcium, but it is not fluorescent), conjugated with a chromophore to obtain fluorescent molecules. Some examples are:
• BAPTA → it is designed to detect calcium fluctuations. It is the same of EGTA, but two benzene rings were added to give fluorescence to the molecule. At 0-calcium amount, BAPTA is very efficiently excited at low wavelength. Calcium concentration causes the carboxy-groups conformational modifications, transferred then to the fluorophore and this can alter the absorption in BAPTA. If the Ca concentration increases, the absorption of photons of BAPTA decreases and it does not emit fluorescence anymore. It is incompatible with life because it is detected with a very short wavelength.
• QUIN-2 → this indicator absorbs at longer wavelength and so it is compatible with life. it is excited by UV-light (339 nm) and it is the first dye used in biological experiments. This molecule is not very bright and needs to be used at high intracellular concentration to overcome cellular autofluorescence.
Other example of new generation calcium indicators are FURA-2, INDO-1, FLUO-3 and indicators with intermediate or low affinity (intermediate or low Kd). Low affinity is useful in the case of measurement of superfast events because the indicator that I use has to be low and rapid reverse.
The next two examples are ratiometric dyes, it means that they are indicators that can collect photons both from bound and free dye, during the excitation or emission event. Ratiometric dyes are compounds which have a shift in their spectra on binding ions or remaining alone. Excitation or emission spectra change as a function of free Ca2+ concentration. The ratio between fluorescence intensities of two different wavelengths estimate the real intracellular calcium concentration. This mechanism allows the normalization of several events such as the uneven loading of the dyes, the shift in the focal plane during acquisition, the photobleaching and the different optical path. Two examples are:
• FURA-2 → it has two excitation peaks: at 340 nm the peak is high, when the concentration of free calcium is high; the other peak is at 380 nm, where the concentration of calcium is low because it is bound.
• INDO-1 → it has two emission peaks, the one at 400 nm that is high when calcium is bound, and the one peak at 475 nm when the calcium is free. The ratio between the two fluorescence intensities is useful to evaluate the real calcium concentration within the cell.
One single dye, allows me to calculate the real calcium concentration within the cell.
And then there is FLUO-3, that is a non-ratiometric dye because it has only one peak of excitation and emission, not double and if it is used in combination with another non-ratiometric dye, such as RHOD or FURA-RED (that have the opposite behavior of FLUO-3), we can obtain ratiometric information about the concentration of free calcium within the cell. This means that one has the emission peak when the calcium is free in the cell, while the other has the emission peak when the calcium is bound. Using non-ratiometric dyes you can avoid UV light, they can be excited with
visible light and give the same result, if they are used in combination. You can use basic confocal microscope, can reduce autofluorescence and cellular damage.
Synthetic molecules are generally hydrophilic and so they do not pass spontaneously the cellular membrane, so the main methods to load synthetic indicators into the cell are:
• Microinjection → we use a glass capillary under high pressure to “shot” the molecules within the cells or the tissue. This can be quite limited because the result of my experiment depends on how many cells I successfully microinjected, (we cannot microinject 200 cells because it takes time and cells are going to suffer).
• Electroporation of the cells with the dye. You favour the entry of a polar dye within the cell creating transient pores in the membrane. It is based on a short pulse that allows the entrance of some molecules within the cells.
• Use of hydrophilic dye forms, through the creation of an esterified dye using AM (acetoxymethyl). For example, one AM dye is Calcein, which spontaneous passes the membrane. An ester occupied the carboxyl group and it is not able to bind the calcium ions. The esterified dye, once intracellular, is converted in a hydrophilic polar and calcium sensitive form in this way the dye can not go out from the cell anymore and it is able to bind calcium ions. Using this method you have to pay attention about the compartmentation of the dye, in other word if the same amount of dye is well distributed in different compartmentations; moreover you have to consider the loss of fluorescence during the process and the incomplete hydrolysis of AM ester and so you can have different calcium sensitive dye within the different compartments. This method is also toxic for the cells.
These methods can be applicated in:
• cell samples through microinjection, electroporation, single cell loadings with sharp electrode and whole-cell patch clamp. All they allow electrophysiological recordings because the membrane is broken and the cell is loaded with dye solution.
• tissue models through Acute network loading for instance in the brain. Many neurons are labelled simultaneously by acetoxymethyl ester (AM) loading (I inject the esterified dyes in a given brain area), so I can perform a local tissue perfusion by injected esters (observing dye behavior) or I can loading of the dye in small vesicles (dextran-conjugated dye, a small polymer that is transported along axons and dendrites to the cell stroma, so I will have the loading of my neuron by applying a dextran-conjugated dye on neurites), and by bulk electroporation (I can electroporate a tissue section to upload the dye in order to treat a population of neuron with the selected dye). All these methods can be useful in brain dynamic studies.
The amount of photons collected from any given point of the sample will depend on both the amount of intracellular calcium and on the amount of the indicator, but this problem is solved thanks to the ratiometric dyes.
methods to evaluate the ligand-receptor complex
• DETERMINATION OF LIGAND-BINDING USING PHYSICAL SEPARATION OF BOUND AND FREE LIGAND. This configuration is based on the size of the molecule (PM). You have a population of free ligand and bound ligand: because of their difference in size, it is possible to separate them and then determinate the total fluorescence of the two population because they go in different fractions.
• DETERMINATION OF LIGAND BINDING BASED ON CHANGES IN FLUORESCENCE EMISSION SPECTRUM OR INTENSITY OF A BULK SOLUTION CONTAINING RECEPTORS AND LIGANDS. The emission wavelength of the free ligand is different from the emission wavelength of the bound one. It is possible to measure the concentration of the receptor.
• FCS (fluorescent correlation spectroscopy). It is possible to determinate the binding between ligand and receptor by means of the detection of the changes in ligand mobility. Free ligand moves faster than a bound ligand, so we can monitor the diffusion coefficient of the bound ligand with respect to the diffusion coefficient of free ligand by using the small detection volume that is given us by the two photons confocal excitation. The fast diffusion molecule will pass very rapidly in the small volume, the slow diffusion molecule instead will pass very slowly. So, we’ll be able to count the number of molecules that is moving fast and the number of molecules that is moving slowly, so we can evaluate the amount of free ligand and bound ligand and correlate that in function of time. The rational of this technique is that rapid fluctuations in fluorescence emission of ligand in a small illuminated volume reflect the diffusion time (mobility) of the labeled entity. One complication of using Fluorescence Correlation Spectroscopy is that multiple species of ligand with different characteristic diffusion times are often detected for a given ligand bound to a particular type of receptor. This confounding effect may reflect different states of homomeric or heteromeric protein–protein association or different subcellular localization of the particular receptor that for example can be embedded into a membrane. In these cases, the diffusion is dramatically changed.
• FLOW CYTROMETRY: both the ligand and the receptor have a fluorescent tag or singly fluorescent. Illumination of a small volume containing receptors expressed on cells or bound to beads allows determination of the amount of bound fluorescent ligand. Free or bound ligand will go in different channels. Data is presented as an histogram, showing the percentage of cells or beads exhibiting a given fluorescence emission intensity.
• FRET (fluorescent resonance energy transfer): it is based on energy transfer between 2 different molecules. Both the ligand and the receptor are tagged with a photon emitting molecule, one is a donor molecule and the other is an acceptor (both fluorescent): when the ligand interacts with the receptor, the energy from the donor will be absorbed by the acceptor and so exciting the donor we have the emission of the acceptor. Donor and acceptor represent what is called “a FRET pair”. In this case the binding of the ligand to the receptor leads to the transfer of the energy by resonance (there is no emission of photons) from the donor to the acceptor that can be detected via a decrease on donor emission or an increase in acceptor emission. If the ligand and receptor are interacting, if I excite the donor, I will see emission in the acceptor. This phenomenon is called sensitized emission.
• BRET (bioluminescent resonance energy transfer): the donor is a bioluminescent molecule. The advantage is that we don’t need to excite the donor because the emission of photons is triggered by a chemical reaction, for instance after a stimulus like the changing of the environment (ex. Renilla Luciferase). BRET is based on non-radiative energy transfer: there is no emission of photons, it is resonance between two molecules. Donor and acceptor molecules must be in close proximity (1-10 nm, 10-100 A) and it is a true measure of interaction. The absorption spectrum of the acceptor and the emission spectrum of the donor must overlap and the more they overlap the more efficient will be the transfer of energy from the donor to the acceptor. So, one protein is linked to donor and the second protein is linked to the acceptor: if the two proteins don’t interact, I can see only photons released by the donor, if the two proteins interact with each other, the donor releases
energy that is transfer to the acceptor by resonance and I can see photons released by the acceptor (NB: I see also photons from the donor, it is never “all or nothing”).
Optical voltage sensors
Optical voltage sensors, called also VSFPs, are fluorescent protein made by voltage segments of endogenous protein, that usually derives from Ciona organism, which has genes that encodes for membrane phosphatases whose activity is dependent on membrane voltage. So, the researchers fused the sensing domain of the phosphatases with a FRET pair, in this way changes in the voltage of the membrane detected by these sensors, are transferred in a conformation change of FRET pair and so in a change in FRET ratio and final fluorescence. These sensors are useful to monitor membrane potential by mean of FRET. VSFPs have a phosphatase domain composed by 4 transmembrane domains. It is possible to use ratiometric measurement of FRET because it is an intramolecular FRET, where the donor and the acceptor are fused in the same construct. If we apply current, we have a membrane depolarization, the voltage increases and the fluorescence emission of the acceptor increases, while the fluorescent emission of the donor decreases and so the FRET ratio increases. We can say that when the membrane is depolarized, the FRET ratio increases. In the opposite situation, obviously, FRET ratio decreases, and we will measure less fluorescence emission. Sensing domain of protein that come from Ciona organism are called Ci-VSD, but there are some examples of optical voltage sensors that do not derived from Ciona organism and are:
• Gg-VSD→ the fluorescent protein is fused to C terminal, while N terminal is the sensing domain. The donor is mCitrine (a variant of YFP) and the acceptor is mKate and they are not fused in the same construct. In physiological condition, Citrin is excited and the emission of the acceptor is very low, meaning that in this condition FRET is very low or absent. Upon membrane depolarization, donor and acceptor are close together because of conformational change within the voltage sensing domain and so FRET ratio increases (ratiometric measurement).
• MacQ Rhodopsin → where the sensing domain is a fraction of Rhodopsin (receptor for photons). When the rhodopsin is not protonated, it is not fluorescent. It is able to collect photons from the excitated Citrine. The Citrine is the donor of the FRET pair, we can excite it and in normal condition I see fluorescence in Citrine and not in rhodopsin, because if it is not protonated it is not able to absorbe photons from the citrine. Upon depolarization of the membrane, the rhodopsin is protonated and so it is able to absorb photons from the citrine. In this case, the donor (citrine) is not fluorescent anymore, because the energy is transfer to the acceptor (rhodopsin), that actually is not able to emit photons. So, in this case, the read out will be reduction in the fluorescence of the donor.
• ArcLight → its sensing domain derived from Ciona, but it has a mutated pHluorin (in position 277) that is no longer sensitive to pH, but it becomes sensitive to voltage. In physiological condition, if the pHluorine is excited it emits photons, but upon membrane depolarization the sensing domain of Ciona undergoes some conformational changes and after excitation it does not emit so many photons anymore.
• ASAP1 → this fluorescent protein is able to do circular permutation. The protein is fused to the sensing domain of chicken phosphate and in physiological condition it is fluorescent. Upon membrane depolarization because of conformational changes of sensing domain, the protein is no longer able to emit photons.
Which are the photoperturbation methods and how they can be used in solving cell pharmacology issues?
The photoperturbation methods are: • PHOTOBLEACHING: with this technique, a small region of interest (ROI) is bleached and it is possible to measure the recovery of the fluorescence. We can say that a fluorescent protein is mobile, if the molecule is able to move from the unbleached area to the bleached one. There is more than one technique for the photobleaching: o FRAP (fluorescent recovery after photobleaching) is used to monitor the movement of the molecules, to quantify it and to analyze photobleaching. This is possible by measuring the diffusion coefficient of the molecules and the mobile fraction. These two parameters determine the recovery of the sample. We can obtain the halftime of equilibrium (T ½) that represents half of the time needed for the recovery of ROI and gives us the diffusion coefficient. Then, we can obtain the ration between the mobile fraction and the immobile one. ESEMPIO cardiomyocytes (cancein (non-cell permanent hydrophobic dye) passes among the cells) and Td-tomatoTM17 fused with cytochrome b5. o FLIP (fluorescence loss in photobleaching), that consist in repetitive bleaches of the ROI. If you measure the fluorescence outside the bleached ROI, this will be decreased in time because we are depleting the cell of all the fluorescent molecules. o iFRAP (inverse FRAP): in this case it is bleached the region outside the ROI and a decrease in fluorescence will be measured in the sample. • PHOTOACTIVATION: non-fluorescent molecules within a ROI are activated in fluorescent through illumination at a given wavelength. Then, if the molecules can diffuse, you will measure a reduction of fluorescence intensity within the ROI over time. It has been carried on an experiment in the post-Golgi with PA-GFP to measure the trafficking of transferrin receptor. • PHOTOCONVERSION: it means that initially the cell is, for example, green due to the GFP and then the ROI is photoconverted using irradiation at a given wavelength which induces a translation of its fluorescence spectrum towards longer wavelength and the ROI becomes red. Then, it is possible to observe that the ROI loses the red fluorescence due to the diffusion outside and the general fluorescence become orange due to the mix of green GFP and red. A photoconvertible molecule is Kaede. The aim of all these techniques is to alter the steady-state fluorescence distribution in a specimen photoperturbing fluorescence in selected regions. These methods allow us to study molecular mobility and from this point of view they are useful in solving cell pharmacology issues because allow us to understand, for example, if a particular drug, has achieved the right compartment of the cell and if it is a useful therapy. Fluorescent proteins are used in vivo imaging (the idea is to express the fluorescent protein under the control of a cell specific promoter and then excite it with long and penetrate wavelength and observe it from outside) and in medical imaging, to monitor and study tumor growth, metastasis and angiogenesis. We can target with a fluorescent protein the tumor cells and monitor the dimension of it before and after a treatment with anti-cancer drugs to understand whether these drugs are useful to reduce the tumor size. We can monitor it because the fluorescence is proportional to the tumor size. Another example of use of PA-GFP to measure the mobility of a nuclear protein: you want to know whether NFkB is migrating from the cytoplasm to the nucleus after drug administration in cell. NFkB is a transcriptional factor that promotes the transcription of some specific gene that will encode for some specific protein. In this case, NFkB is transfected with PA-GFP into the cells. Initially, it is not possible to see the fluorescence because the fluorescent molecule is not activated yet. You select a ROI and excite it with a specific wavelength and so you active PA-GFP. Now you give the drug in order to see in time whether the factor migrates from the cytoplasm to the nucleus to active some specific genes.
FRAP: rationale, and possible application in the analysis of diffusion of fluorescent moieties and their mobile fraction;
FRAP means Fluorescence Recovery After Photobleaching. Performing this method, the sample is stained with fluorescent molecules and the fluorescence of a specific region of interest (ROI) is selectively bleached once (generally through high intensity excitation).
Then the recovery of the fluorescence within the bleached the area is monitored. It depends on the movement of fluorescent molecules form the unbleached area toward the bleached one.
Therefore, based on the recovery of fluorescence within the bleached area we can obtain some parameters:
• Halftime of equilibrium (t1/2): half of the time needed for the recovery of the fluorescence within the bleached area. This gives us an indication of the diffusion coefficient (Deff) of our molecule that is the movement of molecules by diffusion.
• Ratio between the mobile fraction (Mf) and the immobile fraction of the molecules within the sample.
Example 1: neonatal rat cardiomyocytes are loaded with an exogenous cell permeant fluorescent dye (Calcein AM) (FRAP is not tighly dependent on the use of fluorescent protein. We can do FRAP with any fluorescent molecule). Inside the cytoplasm, Calcein AM is converted into a non-cell permeant dye and so it can diffuse from cell to cell only through gap junctions. Images are acquired before bleaching the ROI (Prebleach) and immediately after bleaching (Postbleach) to monitor recovery of fluorescence in the bleached area thanks to the diffusion of Calcein AM among adiacent cells.
Example 2: cells are transfected with a construct made of TdTomato-TM17 (Tandem dimer Tomato) fused with the transmembrane domain of cytochrome B5. The ER is targeted with fluorescent protein. A region of one cell is bleached and then the recovery of fluorescence in time is observed.
Molecules within the lipid bilayer of the ER membranes are very motile, so the recovery of fluorescence is very rapid and the mobile fraction is close to 100%. In the field of observation, one cell is studied, and one cell is always used to monitor or normalize for bleaching due to imaging (control).
In these two examples, the mobile fraction if high because the cytoskeleton allows the movement of the molecule within the cells. If we use drugs (such as a mix of latrunculin, cytochalasin, jasplakinolide, blebbistatin and Y27632) that inhibit actin dynamicity (polymerization and depolymerization) and the movement of the motor protein acting on the actin filament, we will observe that no recovery after photobleaching occurs in this cells after FRAP, because the cytoskeleton is completely fixed and there is mobile fraction.
how to test Biosensors
Intermolecular versus intramolecular energy transfer: rationale and use in the study of ligand/receptor interaction.
Experimental design using BRET to study GPCR interaction with downstream effectors, activation and desensitization.
What are the receptor types?
- Membrane receptors:
- plasma membrane
- endosome
- lysosome
- endoplasmic reticulum (ER)
- Intracellular receptor
What are the drug types?
- Lipophilic => cross the plasma membrane => reach intracellular compartments, easily
- Hydrophilic => interact with surface receptors
What is Fluorescent Lifetime?
The average time the molecule stays in its excited state before emitting a photon and going back in the resting state
Where do you find GFP?
Aequorin protein
Aequorea victoria jellyfish
first calcium sensonr => shines with movement
Why we can not use wild type GFP?
Temperature.
Aequorea victoria lives in the water => cold
Protein folding temperature is not suitable for human.
Thus, we use popular humanized version of GFP= enhanced GFP (EGFP)
What do you do if you want to know specific amounts of two proteins in numbers (in quantitative way)?
1) You can tag another protein that you know its concentration.
The quantum yields of different proteins are very different from each other, even when they have the exact same number of molecules.
So, you can collect and meausre the number of photons by quantitative fluorescence microscopy.
2) You can use Western Blot (WB) by antibodies that are against tags, not the proteins itself. Meaning, you will use antibodies that will bind to your tags like YFP and mGrape.
The antibody that is directed against YFP can also recognize other derivatives of GFP, but not mGrape. Same way, the antibody that is directed against mGrape can recognize other derivatives of RFP or mFruits, but not GFP
What are the types of functional fluorophore variants?
- Photoactivable Fluorescent Protein
- Photoconvertible Fluorescent Protein
- Photoswitchable Fluorescent Protein
