Fluorescence Microscopy and Fluorescent Markers Notes

A. Fluorescence

1. What is Fluorescence?

Fluorescence is the property of certain atoms and molecules to absorb light of a specific wavelength and, after a brief interval (the fluorescence lifetime), re-emit light at longer wavelengths.

It requires an external energy source and is a result of light absorption, involving the emission of electromagnetic radiation (light).
This differs from chemoluminescence, where a chemical reaction creates the excited state. Chemoluminescence involves a chemical reaction that produces light without heat. Bioluminescence is a type of chemoluminescence found in living organisms.

2. Absorption and Emission Spectra

A fluorescent molecule is called a fluorophore. Fluorophores are designed with chemical structures that allow them to undergo fluorescence.
Each fluorophore has a characteristic absorption spectrum. The absorption spectrum indicates the wavelengths at which the fluorophore absorbs the most light.
The ability of a molecule to absorb light at a certain wavelength is described by the molar extinction coefficient $e$ . $Abs = ecl$ where $c$ is concentration and $l$ is pathlength. This equation is based on the Beer-Lambert Law, which relates the attenuation of light to the properties of the material through which the light is traveling. The law is commonly applied in spectrophotometry to measure the concentration of a substance.
A fluorescent molecule also has a characteristic emission spectrum, which is always shifted towards the red (longer wavelength, lower energy) compared to the excitation spectrum. This shift is the Stokes shift. The Stokes shift is due to energy loss during the excited state lifetime.
The Quantum yield of a fluorophore is the ratio of emitted photons per absorbed photon. A high quantum yield indicates a bright fluorophore.
The Brightness of a fluorophore is the product of the extinction coefficient and the quantum yield. Brighter fluorophores are easier to detect and provide better signal in microscopy.
The difference in wavelength between the absorption maximum and the emission maximum is the Stokes shift, which is critical in fluorescence microscopy as it allows separation of excitation light and emission light. Without the Stokes shift, the emitted light would be indistinguishable from the excitation light.

3. Fluorescent Molecules

Also called fluorophores or fluorochromes, they typically have highly delocalized electrons that can absorb energy from photons in the UV to visible light range. The delocalized electrons allow the molecule to absorb light energy efficiently, leading to fluorescence.
The fluorescence spectrum is affected by the size of the conjugated system (larger systems require less energy to excite, thus longer wavelength). Fluorophores with extended conjugated systems tend to emit at longer wavelengths.
Alexa Fluor dyes are a commonly used modern class of fluorophores. Alexa Fluor dyes are known for their brightness and photostability.
Each fluorophore has its characteristic emission and excitation spectra, and it is important to know the spectral properties of the fluorophore being used. Proper selection of fluorophores ensures optimal excitation and detection.

4. How to Observe Fluorescence in a Microscope

An Excitation filter selects the correct wavelengths to excite the fluorophore. The excitation filter ensures that only the desired wavelengths of light reach the sample.
A Barrier filter allows observation of only a selected range of wavelengths from the fluorophore. It is necessary to block the excitation wavelengths, as the observer or detector would otherwise be blinded by reflected excitation light. The barrier filter ensures that only the emitted light from the fluorophore is observed.

B. The Epifluorescence Microscope

1. Overview

Typical design of an epifluorescence microscope includes observation tubes, eyepiece, filter turret, digital camera systems, epi-fluorescence lamphouse, UV shield, objective, stage, condenser, field diaphragm, base, microscope control circuit board, filters, collector lens and tungsten-Halogen lamphouse. The arrangement of these components allows for efficient excitation and detection of fluorescence signals.

2. The Filter Cube

Optical filters make it possible to excite and detect specific fluorophores. These filters are crucial for selective excitation and detection of fluorophores.
The filter cube contains an exciter filter, a dichroic mirror (beamsplitter), and an emission filter (barrier filter). The components of the filter cube are optimized for specific fluorophores.
Properties of the excitation and emission filters and dichroic mirror must match the excitation and emission spectra of the fluorophore. Matching the filter properties to the fluorophore spectra maximizes signal and minimizes background.
Filter sets (excitation filter, emission filter, and dichroic beamsplitter) maximize image contrast and maintain image quality by ensuring high throughput (wide bandwidth & high transmittance) and low cross-talk. High throughput ensures efficient light transmission, while low cross-talk prevents unwanted signals from other fluorophores.

3. Light Sources for Excitation

Mercury lamp: Under high pressure and has a limited life time. Historically very common. Mercury lamps provide a broad spectrum of light, suitable for exciting many fluorophores.
Xenon arc lamp: Weak in the UV region. Xenon arc lamps also provide a broad spectrum but are less intense in the UV region.
Metal-halide lamps: Often mercury-based with long life-times. Currently very common. Metal-halide lamps offer a good balance of intensity and lifespan.
LEDs (light-emitting diodes): Increasingly more common. Each LED emits a specific wavelength. LEDs are energy-efficient and can be precisely tuned to specific excitation wavelengths.
Lasers: Discussed in relation to confocal laser scanning microscopy. Lasers provide highly focused and intense light, ideal for confocal microscopy.
The spectrum of a high-pressure mercury arc lamp and a xenon arc lamp differ significantly across wavelengths. This difference affects the choice of light source for specific fluorophores.

4. Light Path and Summary

The light path in an epifluorescence microscope involves:

Light from the arc lamp passing through a heat filter (removing IR). The heat filter protects the sample from excessive heat.
The light then goes through the exciter filter. The exciter filter selects the wavelengths that will excite the fluorophore.
The light reaches the dichroic mirror. The dichroic mirror reflects the excitation light towards the objective and transmits the emission light to the detector.

5. Comments on Image Acquisition

Images are typically acquired by digital cameras in wide-field fluorescence microscopy. Digital cameras provide high-resolution images with quantitative data.
Generation of digital images involves digital sampling (dividing the image into pixels) and pixel quantization (assigning an intensity value to each pixel). Digital sampling and pixel quantization convert the analog signal from the camera into a digital format.
Example: A cooled CCD camera is monochromatic (greyscale), has a high pixel count (e.g., $1300 \times 1030$ ), and is cooled to reduce noise (e.g., to $-30$ °C). Cooling reduces thermal noise, improving image quality.
Digital color images are generated by using three separate channels, each representing the primary colors: Red, Green, and Blue (RGB images). Each channel contains grayscale values for the respective color. Pseudocoloring of multichannel images is often used in fluorescence microscopy, where the sample is stained with several fluorophores with different spectra. Each fluorophore is visualized separately with a specific filter set and a black-and-white image for each filter set. Each image is assigned a color, and these channels are combined to create the final RGB color image. Pseudocoloring allows for easy visualization and differentiation of multiple targets in a sample.

C. More About Fluorescent Molecules

1. Fluorescence Lifetime and Ways to Leave the Excited State

Fluorescence lifetime is the average time a molecule stays in the excited state. The fluorescence lifetime is an intrinsic property of the fluorophore and can be used for various applications.
The molecule can leave the excited state in several ways, not only by emitting a photon including:-
- Non-radiative relaxation to the ground state. Non-radiative relaxation involves the conversion of excitation energy into heat.
- Quenching by surrounding molecules. Quenching reduces the fluorescence intensity by transferring energy to other molecules.
- Intersystem crossing to a triplet state (non-fluorescent and reactive, often reacting with oxygen, leading to ROS formation and photobleaching). The triplet state is a long-lived excited state that can lead to photobleaching and the formation of reactive oxygen species (ROS).
- Förster resonance energy transfer (FRET), which leads to excitation of another acceptor molecule. FRET is a distance-dependent transfer of energy between two fluorophores.

2. Photobleaching

In the excited state, a molecule may undergo chemical reactions that destroy the fluorophore. Molecules that enter a long-lived triplet state have a higher likelihood of reacting with oxygen and thus undergo faster photobleaching. Photobleaching is accelerated by the presence of oxygen and high light intensity.
Photobleaching is irreversible, involves permanent loss of fluorescence. Once a fluorophore is photobleached, it can no longer emit light.
Photostability is an important feature of a fluorophore, describing the extent to which a fluorophore is photobleached when being constantly excited. Highly photostable fluorophores are preferred for long-term imaging.
Removing oxygen from the sample can reduce photobleaching; this is how some antifading agents work. Antifading agents protect fluorophores from photobleaching, allowing for longer observation times.

D. Fluorescent Dyes and Their Uses in Microscopy, Part 1

1. Overview: What Can Fluorescent Dyes Be Used For?

Organelle Probes: Fluorescent molecules that bind to specific cell components or accumulate in specific compartments (e.g., dyes binding to nucleic acids, mitochondria, Golgi, ER). Organelle probes are used to visualize specific cellular structures.
Fluorophores coupled to other molecules and proteins: Allow for targeting of specific structures (e.g., Phalloidins binding the actin cytoskeleton, Lectins binding specific sugars and Antibodies for Immunofluorescence microscopy). These conjugates are used to target specific molecules or structures within the cell.
Environmental Probes: Change spectral properties in response to environmental conditions to monitor specific conditions inside the cell (e.g., probes sensitive to $Ca^{2+}$ concentration, pH, membrane potential). Environmental probes provide information about the cellular environment.
Labeling of specific protein tags in vivo: SNAP-tags, FlAsH-tags, HALO-tags. These tags allow for specific and efficient labeling of proteins in living cells.

2. Staining of Specific Cell Components – Organelle Probes

Examples of fluorescent stains that are used to stain specific compartments or parts of cells, including:

DNA Stains: DAPI, 7-aminoactinomycin D, Propidium iodide, Acridine orange, Hoechst 33342. These stains are used to visualize DNA in cells.
Mitochondria: Mitotracker Green FM. Mitotracker Green FM is used to label mitochondria in living cells.
Lysosomes: Lysotracker probes. Lysotracker probes are used to label lysosomes.
ER, Golgi: DiOC6. DiOC6 is used to stain the endoplasmic reticulum (ER) and Golgi apparatus.
Membranes: FM4-64. FM4-64 is used to label cell membranes.

3. Conjugating Fluorophores to Other Molecules

Example a: Staining actin cytoskeleton with labelled phallotoxins like phallicidin conjugated to a fluorophore. Phallotoxins bind to actin filaments, allowing for visualization of the actin cytoskeleton.
Example b: Monitoring cell wall synthesis in bacteria: Vancomycin labeled with a fluorophore labels peptidoglycan synthesis sites, as does fluorescently labelled D-amino acids incorporate into peptidoglycan. These labels are used to study cell wall synthesis in bacteria.

4. Immunofluorescence Microscopy

Involves coupling fluorophores to antibodies using the protocol:
1. Fix and permeabilize cells with formaldehyde, and attach to microscope slide. Fixation preserves cell structure, and permeabilization allows antibodies to enter cells.
2. Incubate with primary antibody. The primary antibody binds to the target protein.
3. Wash to remove unbound antibody. Washing removes excess primary antibody.
4. Incubate with secondary antibody conjugated to fluorophore. The secondary antibody binds to the primary antibody, allowing for visualization of the target protein.
5. Wash. Washing removes excess secondary antibody.
6. (if needed, stain other structures or components in cells). Additional staining can be used to visualize other cellular structures.
7. Mount under coverslip and view in microscope. Mounting protects the sample and provides a clear image.
The technique involves using a primary antibody that binds specifically to the target protein, and a secondary antibody conjugated to a fluorophore (e.g., FITC, Cy3, Alexa) that binds to the primary antibody. The secondary antibody amplifies the signal, making the target protein easier to visualize.

5. Indicators of Intracellular Calcium (Ca2+) are Examples of Environmental Probes

Fura-2 is an example that, upon binding of $Ca^{2+}$ , causes a shift in the absorption (excitation) spectrum and it requires microinjection into cells. Acetomethyl-esters of fura-2 are cell-permeant and, once inside the cell, are cleaved to cell-impermeant products by intracellular esterases. Fura-2 is commonly used to measure intracellular calcium concentrations.
Using fura-2 to indicate intracellular free $[Ca^{2+}]$ involves capturing two images with excitation at different wavelengths and then generating a new image by calculating the ratio of the two, pixel by pixel, which is colour-coded using a colour look-up table (LUT). Ratiometric imaging reduces artifacts caused by uneven dye loading or photobleaching.
Fluo-4 is another example used to visualize $[Ca^{2+}]$ oscillations in living cells using confocal microscopy. Fluo-4 is a bright, calcium-sensitive dye suitable for live-cell imaging.

6. Live/Dead Staining

A range of dye combinations can be used to distinguish living from dead cells. The principle is that some dyes can and others cannot penetrate normal intact cytoplasmic membranes. A typical example, used in bacteria, involves two DNA-staining dyes:

SYTO 9 can penetrate and stain DNA in living cells (green). SYTO 9 is a cell-permeant dye that stains DNA in all cells.
Propidium iodide can only penetrate and stain DNA in dead cells with permeabilized membranes (orange). Propidium iodide is a cell-impermeant dye that only stains DNA in cells with damaged membranes.

E. Fluorescent Proteins (FPs)

1. The Green Fluorescent Protein (GFP)

Isolated from the jellyfish Aequorea victoria. GFP revolutionized cell biology by allowing researchers to visualize proteins in living cells.
The protein Aequorin in this organism is chemoluminescent, emitting a blue light in solution.
In the living organism, excited Aequorin transfers its energy to GFP without emitting blue light (FRET), and the excited GFP then emits photons of green light. This is an example of bioluminescence resonance energy transfer (BRET).
Aequorea victoria jellyfish emits green light, not blue.
GFP exhibits fluorescence, while Aequorin exhibits chemoluminescence.

2. Mutations That Improve GFP:

UV excitation. Changing Ser65 to Thr (S65T) abolishes the 370 nm excitation peak and results in only anionic fluorophore, also speeds up formation of the fluorophore 4-fold and gives increased brightness. This mutation is found in the popular EGFP variant. The S65T mutation is a key improvement in GFP variants.
Temperature sensitivity. A. victoria lives in cold water, and wild-type GFP does not fold well at high temperatures like 37°C. Many mutations improve folding, e.g., F64L. The F64L mutation improves the folding of GFP at mammalian body temperatures.
Slow formation of fluorophore (takes ~2h in wt GFP). Also improved by mutations. Faster fluorophore formation allows for quicker visualization of tagged proteins.
EGFP (enhanced GFP) (GFP with F64L and S65T) 488/507. EGFP is a widely used variant of GFP with improved brightness and folding.

3. Engineering the Fluorescent Protein Palette

Specific mutations shift the excitation and emission spectra:-
- Y66H (histidine instead of tyrosine in fluorophore) gives blue (EBFP). Replacing tyrosine with histidine shifts the emission spectrum to blue.
- Y66W (tryptophan instead of tyrosine in fluorophore) gives cyan (ECFP). Replacing tyrosine with tryptophan shifts the emission spectrum to cyan.
- Placing another tyrosine (T203Y) stacked on the fluorophore (and changing S65G) gives yellow fluorescence (EYFP). Adding another tyrosine and changing S65G shifts the emission spectrum to yellow.
New fluorescent proteins from corals and other marine organisms, GFP-like proteins have been found in a range of other marine organisms, including FPs that emit light in the red region of the spectrum.
- DsRed from Discosoma striata (a coral).

Native DsRed is tetrameric. Genetic engineering of DsRed and other proteins have yielded a range of useful proteins that together with the GFP-derivatives cover most of the visible part of the spectrum.

4. Examples of What FPs Can Be Used For

FPs as reporters of gene expression. By genetic engineering, placing the gfp gene under control of the promoter for the b-tubulin gene in the nematode C. elegans FPs can be used to monitor gene expression in real-time.
Lifestyle choices in the bacterium Bacillus subtilis. Promoter of a general stress response gene fused to gene for GFP. FPs can be used to study bacterial stress responses.
Monitoring individual cells, detection of bacteria that produce GFP inside mammalian cellAlso virus particles carrying a GFP tagged protein in their capsid can be detected. FPs can be used to track individual cells and viruses.
Fusion of target protein to GFP. This allows for visualization of the target protein in living cells.

5. Limitations and Problems with FP Fusions

Problems can arise, as some FPs have a weak tendency to dimerize. When such an FP is fused to a protein that forms multimers (like the ClpP protease), there are many surfaces for interaction between FPs on such multimers, leading to increased affinity and formation of clusters, hence producing clustering artefacts. Dimerization can lead to mislocalization or altered function of the tagged protein.

6. Photoactivatable and Photoswitchable FPs

FP varieties that can be triggered to change spectral properties by illumination with specific wavelengths. These varieties of FPs have become very important for new types of microscopy, including super-resolution methods. Photoactivatable and photoswitchable FPs are essential for super-resolution microscopy techniques.

7. SplitGFP

A way to monitor protein-protein interactions i.e. a fluorescent protein (FP) is expressed in two parts (e.g. one is only beta strand 11). The two parts do not have enough affinity for each other to reconstitute a functional FP, so there is no fluorescence.
The two parts may be fused to two different proteins (A and B). If protein A and protein B interact, they bring FP(11) close to FP(1-10), which allows reconstitution of a functional FP. Thus. interaction of A and B results in fluorescence. SplitGFP is a powerful tool for studying protein interactions in vivo.

8. Genetically Encoded Sensors: e.g., Calcium-Ion Sensors

By genetically engineering, Calmodulin and a calmodulin-binding peptide has been added to an FP. Binding of $Ca^{2+}$ to calmodulin induces large conformational change. This is propagated to affect the environment of the fluorophore in the FP, leading to large change in fluorescence intensity. Genetically encoded calcium sensors allow for real-time monitoring of calcium dynamics in cells.

F. Labelling Proteins with Fluorophores In Vivo

Systems have been developed by which an organic fluorophore can be specifically attached to protein tags in cells. These can work in living cells and allow for live cell imaging. These systems offer advantages over FP tagging.

Some advantages over FP tagging include:

Organic fluorophores give stronger and more stable signals. Organic fluorophores are often brighter and more photostable than FPs.
There is flexibility in which fluorophores can be used. This allows for a wider range of applications.
Some tags are small and may interfere less with protein function than FPs. Smaller tags are less likely to disrupt protein function.

1. SNAP-Tag

Can be genetically fused to a protein of interest and a benzylguanine-derivative with a conjugated fluorophore is added to the cells (there are cell-permeant derivatives). The SNAP-tag will break the bond between the benzyl group and the guanine, and will become covalently linked to the benzyl group with its conjugated fluorophore and as there are several possible fluorophores to choose from, one tagged protein can be labelled with different colours in different experiments. SNAP-tags allow for versatile labeling of proteins in living cells.

2. HALO-Tag

Based on a haloalkane dehalogenase enzyme from the bacterium Rhodococcus rhodochrous.Halo-tag is based on a haloalkane dehalogenase enzyme from the bacterium Rhodococcus rhodochrous. • A genetically modified version of this enzyme is used in which the substrate becomes covalently attached to the enzyme and cannot be released.., which is used as a genetically engineered tag to label specific proteins and by adding synthetic substrates, they become covalently linked to the HALOtag. HALO-tags provide a stable and specific method for protein labeling.

3. FlAsH-Tag

A 15 amino acid peptide containing the tetracysteine motif (CCPGCC) is added to the protein (by genetic engineering), and this tagged protein is produced by the cells. A fluorophore with two arsenic groups (biarsenical fluorophore) is added to the cells and is cell-permeant. In the reducing environment in the cell, the biarsenical fluorophore reacts with and gets bound to the tetracysteine motif, converting the fluorophore to the fluorescent form. FlAsH-tags are useful for labeling proteins with small, cell-permeant fluorophores.

G. Quantum Dots

Nanometer-sized crystals of purified semiconductors. Quantum dots exhibit unique optical properties due to their size and composition.
The fluorescence emission wavelength is dependent on the particle size (i.e. size of the nanoparticle) but independent of the excitation wavelength. This allows for tuning of the emission wavelength by controlling the size of the quantum dot.
There is no problem with photobleaching. Quantum dots are highly resistant to photobleaching, making them suitable for long-term imaging.
It is not easy to functionalize for use with biological samples and living cells. Functionalizing quantum dots for biological applications can be challenging.

H. Some Microscopy Methods That Depend on Specific Properties of Fluorophores

1. Photobleaching-Based Methods to Study Protein Dynamics:

Fluorescence recovery after photobleaching (FRAP). FRAP is used to measure the mobility of proteins in living cells.
In FLIP, a region of interest is constantly photobleached, bleaching all molecules that pass through this region and simultaneously, the fluorescence signal is measured in another part of the cell to monitor to what extent photobleached molecules are spreading throughout the cell.

FRAP results are interpreted by plotting recovery of intensity after bleaching to determine the mobile and immobile fractions.

2. Förster Resonance Energy Transfer (FRET)

Called ”Fluorescence Resonance Energy Transfer”. In the excited state, a fluorophore (the donor) transfers energy to another fluorophore (the acceptor) without emitting a photon (non-radiative transfer). The acceptor then becomes excited at ground state.

Requirements for FRET to occur:

The donor and acceptor must be very close (less than approx 10 nm). Efficiency depends inversely on the 6th power of the distance. FRET is highly sensitive to the distance between the donor and acceptor.
The emission spectrum of the donor should overlap with the excitation spectrum of the acceptor (the energy of the excited donor has to match the energy needed to excite the acceptor). Spectral overlap is essential for efficient energy transfer.
The orientation of the molecules with respect to each other matters (the transfer occurs via a long-range dipole-dipole coupling). The orientation of the donor and acceptor affects the efficiency of energy transfer.

FRET allows monitoring of distance between molecules. If FRET is detected between two molecules, they are so close that likely interact with each other.

3. Fluorescence Lifetime Imaging (FLIM)

Fluorescence lifetime can be regarded as the average time the fluorophore spends in the excited state before returning to the ground state. It is measured as the time it takes for the fluorescence signal to decay to 1/e of the peak intensity. FLIM-FRET is measuring FRET with FLIM, which affect the fluorescence life-time i.e. If FRET occurs, the average fluorescence life-time becomes shorter and FLIM can be a reliable way to measure FRET.

4. Single-Molecule Localization Microscopy (SMLM)

If knowing that the signal that is detected as an Airy disc comes from one single fluorophore molecule, you can estimate the likely localization of this molecule in the middle of the Airy disc pattern.

Different forms of SMLM

PALM: Photoactivated Localisation Microscopy by by photoactivatable FPs, which can be switched from a dark state to a fluorescent state by light of certain wavelength. PALM relies on photoactivatable FPs to achieve super-resolution.
STORM (Stochastic Optical Reconstruction Microscopy): • In the most common variety