Fluorescence & Advanced Microscopy Techniques

Microscopy Techniques: Fluorescence

Bright Field vs. Dark Field

• Bright Field Microscopy: Utilizes white light and staining to visualize samples.

  • Contrast is achieved through the absorption of light by the sample. Structures that absorb more light appear darker, while those that absorb less appear brighter.

• Dark Field Microscopy: A contrast-enhancing technique that uses light diffracted by the sample to create a bright image on a dark background.

  • Employs diffraction to visualize structures based on their ability to diffract light. Instead of directly illuminating the sample, light is directed at an angle, so only light scattered by the sample is collected by the objective lens.

Phase Contrast Microscopy

• Utilizes white light.

• Specimens are unstained, making it useful for observing live cells.

• Contrast is generated via refraction. It exploits differences in refractive index within the sample to produce contrast. Light passing through denser regions of the sample is slowed down and experiences a phase shift, which is then converted into amplitude (brightness) variations.

Differential Interference Contrast (DIC) Microscopy

• Employs polarized light and refraction to generate contrast.

• Relies on the birefringent properties of substances like collagen. DIC microscopy uses polarized light beams that pass through the sample and recombine. Differences in the refractive index and thickness of the sample alter the path of the light, creating an interference pattern that enhances contrast, giving a 3D appearance.

Fluorescence Microscopy

• Uses specific wavelengths of colored light to excite fluorescent molecules.

• Involves labeling with fluorophores. These are molecules that emit light of a specific wavelength when excited by light of another wavelength.

• Contrast is achieved through the Stokes shift: incoming light is absorbed and then emitted at a longer wavelength. The difference in wavelength between the excitation and emission light is known as the Stokes shift.

Fluorescence Microscopy

• A primary technique in life science research due to high contrast and molecular specificity.

• Enables labeling of cells and tissue structures using molecularly specific probes, allowing researchers to target and visualize specific components within a sample.

Confocal Microscopy

• A laser scanning system that uses point illumination and spatial filtering to eliminate out-of-focus light, resulting in sharper images.

Light as a Particle

• Photons: Packets of energy. The energy of a photon is inversely proportional to its wavelength ($E = \frac{hc}{\lambda}$, where $E$ is energy, $h$ is Planck's constant, $c$ is the speed of light, and $\lambda$ is wavelength).

• Fluorophores absorb energy and emit light at a different, usually longer, wavelength. This process is fundamental to fluorescence.

• Thermal relaxation accounts for energy loss: Shorter wavelengths have more energy, longer wavelengths less. When a fluorophore absorbs light, it enters an excited state, and some energy is lost due to molecular vibrations (thermal relaxation) before emitting light.

Fluorescence Process

• Light is absorbed, exciting electrons in the fluorophore to a higher energy state. The fluorophore's electrons jump to a higher energy level when it absorbs light of a specific wavelength.

• Thermal relaxation leads to energy loss. Some of the absorbed energy is dissipated as heat through molecular vibrations.

• Fluorescence occurs when electrons return to their ground state, emitting light of a longer wavelength. As the electron returns to its normal energy level (ground state), it emits light with less energy (longer wavelength) than the absorbed light.

Fluorophores

• Used to label proteins or molecules of interest, allowing their visualization under a fluorescence microscope.

• Common examples: Alexa Fluor dyes, known for their brightness and photostability.

• Often used in multicolor experiments (e.g., red, green, blue), enabling the simultaneous visualization of multiple targets within the same sample.

• Alexa dyes are named according to their absorption peak (e.g., Alexa Fluor 488 absorbs maximally at 488 nm). The number in the name indicates the wavelength at which the dye absorbs the most light.

Excitation and Emission Spectra

• Excitation (or absorption) spectra: Shows how much light is absorbed by a molecule at different wavelengths. It plots the absorption efficiency of a fluorophore across a range of wavelengths.

• Emission spectra: Shows the wavelengths of light emitted by the molecule after excitation. It plots the intensity of emitted light across a range of wavelengths.

• Emission is proportional to excitation but less efficient if the excitation wavelength is off-peak. The closer the excitation wavelength is to the fluorophore's absorption peak, the more light will be emitted.

Quantum Yield

• Indicates the efficiency of a fluorophore: how many photons are emitted for each photon absorbed. A higher quantum yield means less energy is lost to non-radiative processes.

• Higher quantum yield indicates a better fluorophore. Fluorophores with high quantum yields are brighter and more sensitive.

Wide Field Epifluorescence Microscopy

• Uses a broad spectrum white light source to excite fluorophores in the sample.

• Lacks a condenser. In epifluorescence, the objective lens acts as both the condenser and the objective.

• Features a dichroic mirror within a filter block to filter excitation light and collect emission light. The dichroic mirror reflects specific wavelengths of light (excitation light) and allows other wavelengths (emission light) to pass through.

• Filters are essential to remove excitation light, allowing the weaker fluorescence signal to be observed. These filters ensure that only the emitted light reaches the detector, improving the signal-to-noise ratio.

Inverted Microscope

• An inverted configuration places the objective below the sample, allowing for easier imaging of samples in dishes or well plates.

• Blue excitation light from below excites the sample, causing it to fluoresce. The excitation light is directed through the objective lens onto the sample.

• The dichroic mirror reflects blue light but allows red (fluorescence) to pass through. This separation of excitation and emission light is crucial for fluorescence microscopy.

• An emission filter further cleans up the fluorescence signal; modern dichroics can handle multiple wavelengths but still benefit from emission filters. Emission filters block any remaining excitation light and select specific emission wavelengths.

Wide Field Microscopy

• Images are typically captured using a camera, such as a CCD (charge-coupled device) or CMOS (complementary metal-oxide-semiconductor) camera.

Advantages of Wide Field Microscopes

• Highly flexible and can be used for various applications.

• Good for live imaging due to speed. Wide-field microscopes can capture images quickly, making them suitable for observing dynamic processes in live cells.

• Utilizes molecularly specific probes, advantageous over generic stains as you know exactly what is being labeled. Fluorescent probes allow researchers to target specific molecules or structures within a cell or tissue.

Cellular Labeling

• Targets can include whole cells, organelles (e.g., T tubules, mitochondria), proteins, or lipids. Fluorescent labels can be designed to bind to specific cellular components.

• Functional imaging is possible (e.g., calcium imaging). Fluorescent indicators can be used to monitor changes in ion concentrations or other physiological parameters within cells.

Labeling Methods

• Direct Imaging: Live or fixed cells; molecules bind directly to the target; tags such as green fluorescent protein (GFP). Direct labeling involves attaching a fluorescent molecule directly to the target of interest.

• Immunolabeling: Primarily on fixed samples. Antibodies are used to recognize and bind to specific targets, and then fluorescently labeled secondary antibodies are used to visualize the primary antibody.

• Genetic Tags: Live or fixed samples (e.g., green fluorescent protein). Genetic tags involve inserting a gene encoding a fluorescent protein into the genome of a cell or organism, causing the protein of interest to be fluorescently labeled.

Cell Structure

• Specific probes can reveal the function of different organelles and structures within cells, aiding in disease treatment. By visualizing specific cellular components, researchers can gain insights into their roles in health and disease.

• Super-resolution microscopes can track molecules like tubulin (part of microtubules/cytoskeleton). These advanced microscopes can resolve structures at a higher resolution than traditional light microscopes.

Direct Labeling Methods

• Histological Stains: Applied to tissue sections (e.g., hematoxylin and eosin). These stains provide contrast and highlight different tissue components.

• Ion Indicators: Fluorophores bind to calcium to measure calcium release. These indicators change their fluorescence properties when they bind to specific ions, allowing researchers to monitor ion dynamics.

• Organelle-Specific Labels: Target specific organelles, allowing researchers to visualize their structure and function.

Live Cell Imaging Considerations

• Labels must be non-toxic to avoid disrupting normal cellular processes.

• Getting labels into cells can be challenging due to lipid membranes. The plasma membrane of cells is a barrier to many molecules.

• AM Esters: Allow labels to cross the membrane; esterases within the cell cleave off hydrophobic esters, trapping the fluorophore inside. AM esters are hydrophobic and can diffuse across the cell membrane. Once inside, esterases remove the ester groups, converting the molecule into a charged form that cannot cross the membrane.

• Microinjection: Dye is injected directly into the cell using a fine needle.

• Electroporation: Electric current makes membranes leaky, allowing dye entry. A brief electrical pulse creates temporary pores in the cell membrane, allowing molecules to enter.

Resources for Direct Labeling Dyes

• Thermo Fisher site: Provides information on direct labeling dyes for different organelles. This is a valuable resource for researchers looking for specific dyes.

Examples of Fluorescent Labels

• ER Tracker: Labels the endoplasmic reticulum (protein synthesis).

• MITO Tracker: Labels mitochondria.

• Molecular Probes (now Thermo Fisher) originally developed these fluorescent labels.

Ion Indicators

• Can also detect pH and membrane potential. These indicators change their fluorescence properties in response to changes in pH or membrane potential.

• Change fluorescence upon binding to ions or changes in pH. The change in fluorescence can be used to quantify the concentration of ions or the pH.

• Example: Confocal imaging of cells with changes in fluorescence intensity over time, visualized using pseudo lookup tables. Pseudo lookup tables are color scales that allow researchers to visualize changes in fluorescence intensity.

Indirect Methods: Immunolabeling

• Uses antibodies for detection. Antibodies are proteins that specifically bind to other molecules (antigens).

• A primary antibody recognizes the target, and a secondary antibody recognizes the primary antibody.

  • Secondary antibody has a fluorophore or chromogen, allowing visualization of the primary antibody.

Single-Color Indirect Immunolabeling

• The primary antibody binds to the protein of interest.

• The secondary antibody, raised in a different species, binds to the primary antibody. This allows for signal amplification.

Amplification and Flexibility in Immunolabeling

• Multiple secondary antibodies can bind to one primary, amplifying the signal, making it easier to detect.

• A bulk standard secondary antibody can be used with various primary antibodies as long as it recognizes the species where the primary was raised (e.g., rabbit). This simplifies the labeling process.

Antibodies

• Rabbit antibodies are typically polyclonal. Polyclonal antibodies are produced by multiple B cells and recognize different epitopes on the same antigen.

• Mouse antibodies tend to be monoclonal. Monoclonal antibodies are produced by a single B cell and recognize a single epitope on the antigen.

Antibody Detection Methods

• Original Method: Horseradish peroxidase (HRP) attached to the secondary antibody, using DAB as a chromogen (produces a brown color). This method is based on an enzymatic reaction that produces a visible precipitate.

  • Still used, but less common due to the development of more sensitive and versatile methods.

• Fluorescent Detection: Traditional dyes like Alexa Fluor, FITC, or quantum dots. Fluorescent dyes emit light when excited by specific wavelengths, allowing for sensitive detection.

  • Fluorescent detection is more common now due to its higher sensitivity and the availability of a wide range of fluorescent dyes.

Electron Microscopy (EM) Labeling

• Uses nanogold attached to antibodies for high-resolution visualization; gold is electron dense and appears as dark spots in EM images.

Comparison of Labeling Methods

• DAB chromogen: Brown color, less specific, and can have high background.

• Confocal image: High contrast, very specific, and allows for optical sectioning.

• EM: Best resolution, but lower contrast; gold beads are visible as negative images.

Fluorophore Resources

• Thermo Fisher site contains information on fluorophores and secondary antibodies.

• Alexa Fluor dyes cover the visible light spectrum, providing options for various multicolor experiments.

Designing Multicolor Experiments

• Choose dyes with well-separated emission spectra to minimize signal overlap.

Genetic Labeling: Fluorescent Proteins

• Proteins that are auto-fluorescent, meaning they produce their own light without the need for external dyes.

• Derived largely from jellyfish (Green Fluorescent Protein or GFP) and coral (DsRed).

• The gene sequences are cloned and attached to the gene of a target protein. When the target protein is expressed, it is fused to the fluorescent protein.

• Allows specific targeting and tracking of proteins of interest; when the protein is made, it fluoresces, allowing its location and movement to be tracked.

Origins of Fluorescent Proteins

• Aequorea victoria (jellyfish): Source of GFP, which emits green light.

• Discosoma species (coral): Source of red fluorescence proteins, which emit red light.

Applications of Fluorescent Proteins

• Finding a protein's location within a cell.

• Tracing a protein's movement over time.

• Cell lineage tracing, following the development of cells and tissues.

• pH indicators, using fluorescent proteins that change their emission properties in response to pH changes.

• FRET (Fluorescence Resonance Energy Transfer), a technique to study protein-protein interactions.

• Whole animal studies, allowing the visualization of gene expression and protein localization in living organisms.

Transgenic Animals

• Global expression of GFP can be achieved in animals like mice, allowing for the visualization of cells and tissues throughout the body.

Mutants of GFP

• Mutations in GFP's structure have led to different colors (mutants), expanding the range of fluorescent proteins available.

Nobel Prize

• Awarded for the discovery of GFP and its application in biology, recognizing its transformative impact on biological research.

Bleaching

• Light can damage fluorophores, causing a loss of fluorescence over time, reducing image quality.

Anti-Fade Compounds

• Added to imaging buffers to prevent bleaching, extending the time during which images can be acquired.

• Bleach-resistant fluorophores have also been developed, providing a more robust alternative.

Emission Spectra Overlap

• Avoid fluorophore pairings with overlapping absorption spectra if possible to minimize bleed-through.

• Common pairing: Alexa Fluor 488 and 594, which have well-separated emission spectra.

• Alternatively can use far red dyes like Alexa Fluor 633, which have minimal overlap with other commonly used dyes.

Confocal Microscopy

• In light microscopy, images typically have a thickness of about three microns, which can cause out-of-focus light, making structures appear fuzzy.

• Confocal microscopy allows optical sectioning, where only light from a thin plane within the sample is collected, eliminating out-of-focus light.

• Optical sections of around 600-800 nanometers are achievable, allowing the view of a really thin slice. This improves image resolution and clarity.

• Physically, sections much below three microns are difficult to cut, making optical sectioning a valuable alternative.

• Full illumination in epifluorescence leads to increased photo bleaching because the entire sample is exposed to light.

History of Confocal

• Marvin Minsky patented conceptually in 1960s, but commercial instruments didn't appear until the 1980s due to technological limitations.

Comparison of Widefield and Confocal

• Cardiac Myocyte Labeled for Actin: Labeled for actin, widefield is fuzzy, while confocal is sharper due to the removal of out-of-focus light.

Resolution

• Confocal improves axial (z) and also improves lateral resolution, resulting in clearer and more detailed images.

Key Component: Pinhole

• Confocal systems use a pinhole to eliminate out-of-focus light, set to one airy unit, which is the optimal pinhole size for maximum resolution.

Kinds of Confocal

• Laser Scanning Confocal Microscopy: Uses a single laser beam to scan the sample point by point.

• Spinning Disk Confocal Microscopy (Multiple Pinhole): Uses multiple pinholes on a spinning disk to rapidly scan the sample.

• Multifocal Microscopy (No Pinhole): Uses multiple focal points to image the sample, but does not use a pinhole to reject out-of-focus light.

Convocal Like system components

Elements

• Laser light source, which provides a monochromatic and coherent light beam.

• Objectives, which focus the laser light onto the sample and collect the emitted light.

• Filters, which select specific wavelengths of light for excitation and emission.

• Detectors: Photomultiplier (laser scanning), Camera (nikon).

• Scanning Mirrors, which steer the laser beam across the sample.

• Scanning Prisms, which can also be used to steer the laser beam.

• Pinhole, which blocks out-of-focus light.

How does Convocal Work?

• Laser produces diffraction limited spot and is scanned across the sample using two mirrors (x and y).

Diagram

• Illuminating a small point of the overall volume of a specimen to reduce the exposed area to laser light, minimizing photo bleaching.

• Light emitted and separated by a dichroic mirror.

• In focus light is allowed, and it is recorded by a photomultiplier. The computer generates the image, which is not viewable by the naked eye directly.

Point Spread Function

• This impacts what resolution can be achieved, specifically the airey disk and what size of the area unit is achieved by the brain.

Two Photon Excitation

Resolution

• The advantage is you don't have above and below excitation. In order to measure point spread function you need point source below it's resolution limit. So just buy fluorescent beads.

Spinning Disk Confocal

• Rather than use a single laser point, it uses lots, which are blurred out onto the spinning disk.

Pinhole Array

• This has another spinning disk, which is the equivalent of the pinhole.

• In spinning disk, the laser source blurs onto a spinning disk, which is then sent through pinholes. The detection then uses a camera pixel.

Scientific Cameras

• These do not come as RGB like a regular camera. Instead, they capture intensity values for each pixel.

Benefits

• Good for what you want to measure.

• Fluorescence simulation of using an epifluorescence, opening the pinhole. Sample has a bright sample.

• Can close the pinhole to improve resolution for a bright sample but open if it is a full sample.

Using Confocal

• For reflection, it is used in laser and the sample is used like a mirror, reflecting it back. It still uses pinhole.

• Can be used as a transmission; however, this is not a convocal mode. The benefit is you overlay it with whatever laser and the cell is fluorescing with for contacts and structure.

• Confocal also has optical sectioning, is 3D reconstructable, very good resolution, has all sorts of wavelengths (1-4), is high sensitivity, manipuable digital image extraction, and advanced imaging techniques.

Point Spread Axial Function

• Axial shape is most important, the shape in z as it shows the shape is worse, or not as concise in the z plane.

Stereoscopic Images

• The images you can use after acquiring an image, creating a 3D representation.

Seed Stack

• If you're imaging a sphere, take optical sections at regular intervals to capture the entire volume.

Rendering Methods

• Allows the user, the computer to slice and see where in the molecule it looks best on the inside, providing detailed structural information.

Multi Laboring Tips

• Benefit with convocal is you can use the spectra to elect what you want, so can mitigate bleeding through by electing wavelength with special defection and sequential scanning.

• Spectral mixing also works in close processing.

Channels

• If you're using a few, separated florescence is better; however, if you use more, the options open up and provide different benefits.

Heart Example

• To understand it, understanding kalcium is important for arrhythmia. Isolated cardiomyocytes and you've got a calcium dye, which fluoresces when it binds to calcium, you CAN then get calcium to spread. The multi scale problems can be contributed to at the self cellular level.

• Collagen can function in reflection mode because it shows the fibrills

Benefits of Confocal Use in Sciences

• There is labels, protein trapping is easier, organelle identification, live cell, supercellular with Mitochondria identification, molecular mobility measuring, FRAC

  • FRAC is fluorescence recovery after photo bleaching

Limitations

• The resolution is limited to the 300 Naometter

• It is expensive. To use it and be specific you have to have expensive tags. The laser is limited by what can be used. Also, don't use too must, burn the specimen.

Two Photon Imaging

Lasers

• In two-photon imaging, lasers emit photons with approximately twice the wavelength required for single-photon excitation of a fluorophore (e.g., around 800-1000 nm for a fluorophore that usually absorbs at 488 nm). Because energy is inversely proportional to wavelength, each of these photons has about half the energy needed for excitation. The fluorophore can be excited only when two of these photons arrive nearly simultaneously.

• Two-photon excitation requires high peak power, so it typically uses pulsed lasers. These lasers emit very short bursts of light with very high intensity, increasing the likelihood of two photons arriving at the fluorophore at the same time.

Concentration

• Two-photon excitation depends on the square of the laser intensity; therefore, it occurs only at the focal point where the photon concentration is high enough for there to be a good probability of two photons arriving simultaneously at the fluorophore.

Resolution Function

• Two-photon excitation provides an intrinsic improvement in the point spread function (PSF) and improved axial resolution, particularly for live cell imaging and deep tissue imaging in animals.

Two Photon Imaging Facts

• Two-photon excitation uses longer wavelengths of light, it penetrates deeper into scattering samples. This makes it particularly useful for imaging in thick tissues and live animals. The longer wavelengths used in two-photon imaging are less prone to scattering by molecules in the sample, allowing for deeper penetration.

Two and One Photon Comparison and Benefits

• Better high axial resolution

• No pinhole is needed because excitation is confined to the focal point.

• Good at imaging scattering samples and working with live samples because of reduced photobleaching and deeper penetration.

• One-photon confocal microscopy removes out-of-focus light using a pinhole, while two-photon microscopy achieves optical sectioning through the physics of two-photon excitation.

References

Websites

• Excellent resources are available from microscope manufacturers such as Nikon. Their websites often provide detailed information and interactive tutorials on microscopy techniques.

Advanced Information

• TIRF (Total Internal Reflection Fluorescence) Microscopy: Utilizes the principle of total internal reflection to selectively excite fluorophores in a very thin region (approximately 100 nm) close to the coverslip. This is achieved by directing the excitation light at an angle such that it undergoes total internal reflection at the interface between the coverslip and the aqueous sample. This technique is particularly useful for studying events at the cell membrane.

• FRAP (Fluorescence Recovery After Photobleaching): Used to study protein dynamics and mobility within living cells. A small region of the sample is photobleached using a high-intensity laser beam, and the recovery of fluorescence in the bleached area is monitored over time. The rate of fluorescence recovery reflects the mobility of the fluorescent molecules.

• FRET (Fluorescence Resonance Energy Transfer): A technique used to study protein-protein interactions and molecular proximity. When two fluorophores are in close proximity (typically 1-10 nm), excitation of one fluorophore (the donor) can result in energy transfer to the other fluorophore (the acceptor), causing the acceptor to emit light. The efficiency of energy transfer depends on the distance between the fluorophores, providing information about their proximity.

• FLIM (Fluorescence Lifetime Imaging Microscopy): Measures the time a fluorophore spends in its excited state before returning to the ground state. This lifetime is sensitive to the microenvironment of the fluorophore, including pH, ion concentration, and binding to other molecules. FLIM can provide additional contrast and information beyond that obtained from conventional fluorescence intensity imaging.

TIRF Details

Evanescent Field

• In TIRF microscopy, the excitation light generates an evanescent field, a very thin electromagnetic field that extends approximately 100 nm from the coverslip into the sample. Only fluorophores within this evanescent field are excited, providing highly selective imaging of structures near the coverslip.

• TIRF does not replace confocal microscopy but is useful for specific applications such as visualizing filament growth and cell adhesion events at the cell membrane.

FRAP Details

• FRAP is used to measure the turnover rate of proteins within a specific cellular compartment. After photobleaching, the rate at which fluorescence recovers in the bleached area is determined by the diffusion and binding kinetics of the fluorescently labeled proteins. This provides information about the dynamic behavior of proteins within the cell.

FRET- Fluorescence Resonance Energy Transfer Details

• FRET can provide nanometer-scale information about the distance between two labeled molecules. If the molecules are close enough to interact physically, FRET will occur; if they are farther apart, FRET will not occur. FRET is often used to study protein-protein interactions, conformational changes in proteins, and the activity of signaling pathways.

• In a FRET experiment, the emission spectrum of the donor fluorophore should overlap with the excitation spectrum of the acceptor fluorophore for efficient energy transfer. When the donor is excited, energy is transferred to the acceptor, causing the acceptor to emit light at its characteristic wavelength.

Time-Correlated Single Photon Counting (TCSPC) and Fluorescence Lifetime Imaging (FLIM)

Time-Correlated Single Photon Counting (TCSPC)

• A technique often used in FLIM to precisely measure the arrival times of individual photons emitted by the sample after excitation with a pulsed laser. By analyzing the distribution of photon arrival times, the fluorescence lifetime of the fluorophore can be determined.

Benefits of FLIM

• Improves spatial resolution by providing additional contrast based on fluorescence lifetime.

• Allows monitoring of changes in parameters such as pH and ion concentration, as these can affect fluorescence lifetime.

• Fluorescence lifetime is independent of fluorophore concentration and excitation light intensity, making FLIM a robust technique for quantitative imaging.

Images of Lifetimes

• Fluorophores can show similar spectral emission properties but have distinct differences in their fluorescence lifetimes. FLIM can differentiate between these fluorophores based on their lifetimes, even if their emission spectra overlap.

FLIM-FRET

• FLIM can be combined with FRET to provide additional information about molecular interactions. In FLIM-FRET, the lifetime of the donor fluorophore is measured in the presence and absence of the acceptor fluorophore. A decrease in the donor lifetime indicates that FRET is occurring, providing evidence of a close interaction between the donor and acceptor molecules. This can be visualized using specialized software that maps fluorescence lifetimes in each pixel of the image.

• Example: can be done with caged cyclic AMP to see distinct differences in signaling events.

New Microscopy Techniques

Light Sheet Microscopy

• Light sheet microscopy uses a thin sheet of light to illuminate a single plane of the sample, reducing out-of-focus light and minimizing photobleaching. The sample is typically illuminated from the side, and the emitted light is collected by an objective lens oriented perpendicular to the light sheet. This technique is particularly well-suited for imaging large, transparent samples such as developing embryos and organoids.

Key Features of Light Sheet Microscopy

• Uses a thin sheet of light to optically section