Fluorescence microscopy spans from basic epifluorescence/widefield microscopy to advanced techniques like confocal microscopy, FRAP, FLIM, FRET, and TIRF. These cutting-edge technologies enable specimen imaging with exceptional resolution and precision.
Each fluorescence microscopy technique comes with its distinct characteristics, benefits and drawbacks.
In this article we will go into more detail about each microscopy technique, explaining the principle behind each, advantages and disadvantages, and other important information.
Article table of contents:
Widefield, also known as epifluorescence microscopy is the most basic type of fluorescence microscopy where the whole specimen is illuminated. In this technique, both the incident and emitted light waves are transmitted through the same lens.
Figure 1. An image of mouse ear cells. The target protein in this case was b-actin, labeled with green fluorescent protein (green), and its localization was imaged using an epifluorescence microscope. F-actin was labeled with phalloidin (red). Scale bar =1 micrometer. (Roy and Perrin. 2018).
Widefield microscopes have several key features including a white light source, a fluorescence light source, dichoic mirrors, filters and digital cameras.
Widefield microscope features list:
- A fluorescence light source
- A dichroic mirror that reflects the incident waves on the specimen. While transmitting to the detector, the waves emitted from the specimen.
- Filters that block all waves other than the desired ones with a particular wavelength from reaching the specimen, and then allowing only the required emitted wave from reaching the detector.
- Digital camera (with or without a computer).
Gas arc lamps were the primary source of excitation light for widefield microscopy until the light-emitting diodes (LED) were developed and then used as fluorescent light sources.
LEDs offer several advantages over arc lamps, including extended lifespan, improved light output stability, and enhanced reproducibility and quantification.
Nonetheless, the familiarity and availability of the gas arc lamps like mercury-arc lamps and xenon-arc lamps make them a convenient choice for researchers, manufacturers, and users who have already optimized their setups for gas arc lamps.
Mercury arc lamps are also known to produce high-intensity light output, often surpassing the brightness of LEDs.
Despite being powerful light sources for fluorescence microscopy, gas arc lamps have a limited lifespan and need to be replaced frequently.
Widefield microscopes can be either inverted or upright. The distinction affects the type of specimen being studied.
- The specimen is illuminated from above the stage.
- Designed for observing live cells growing on a culture dish.
- The objective lens is below while the condenser is above the sample.
- The specimen is illuminated from below the stage.
- Typically designed for examining samples that are fixed on a slide.
- The objective lens is above and the condenser is below the fixed sample.
- This technique allows you to stain and image nearly any component of living or fixed cells and tissues at the highest magnifications possible.
- Widefield microscopy is frequently employed for real-time observation of specimens allowing users to select fluorescent cells to image. It is critical to be mindful of photobleaching and the observation time, especially if you are imaging your sample after manual observation.
- It is commonly used to analyze immunofluorescence assays.
- In general, the widefield microscopy technique is easy to use and has a high processing speed.
- A drawback is that the light source illuminates the entire specimen or a wide field view. This can be problematic if you want to image a specific component within the cells, such as nuclei, against a dark background of the remainder of the cells.
- Despite being user-friendly, it has limited contrast and resolution due to the out-of-focus fluorescence feature, resulting in blurred images.
Confocal microscopy, a fluorescence optics-based technique, is used to obtain high-resolution and intricate 3D images of cellular and tissue components, including live samples. It is also useful in determining biomolecular interactions and expression patterns using co-localization studies.
Confocal microscopy was invented by Marvin Minsky in 1957 and was introduced to the commercial market in the 1980s. Since then, it has revolutionized cell and molecular biology research.
Just like widefield microscopy, in confocal microscopy, you can employ immunofluorescence to specifically label various components of living or fixed cells and tissue sections, followed by high-resolution visualization.
Figure 3. A
confocal image showing mouse ear cells inside the cochlea. The target protein
fascin-2 was labelled with GFP (green) while F-actin was labelled with
Figure 4. A confocal image showing mouse ear cells inside the cochlea. The target protein in this case was b-actin, labeled with GFP (green), and its localization was imaged using an epifluorescence microscope. F-actin was labeled with phalloidin (red).
A confocal microscope utilizes laser units integrated into the system to serve as the light source for stimulating fluorescent dyes and proteins. Because of this, it is also called confocal laser scanning microscopy (CLSM).
Like widefield microscopy, confocal microscopy utilizes fluorescent optics. However, instead of illuminating the entire specimen, confocal microscopy uses laser light to focus on a precise spot at a specific depth within the sample.
This results in fluorescence emission at that specific point, which helps image your target with high specificity.
A pinhole placed in the optical pathway acts as a filter, eliminating out-of-focus signals and enabling only the fluorescence signals from the illuminated spot to reach the detector.
This technique provides high-resolution, three-dimensional imaging with improved contrast and reduced background noise. This is what helps you get a blur-free image.
- Pinholes: A pinhole acts as a filter, effectively eliminating any out-of-focus signals. This selective filtering enables only the fluorescence signals generated from the illuminated spot to reach the detector, enhancing the accuracy.
- Laser units: Usually 5 different lasers are sufficient to cover the excitation wavelength range of the most commonly used fluorophores like FITC, and Texas red. The two most commonly used laser units are listed below.
- Argon-ion laser: the green wavelengths of the excitation spectrum that excites fluorophores like FITC are covered by this laser.
- Helium-neon laser: the yellow to red of the excitation spectrum that excites Texas red and rhodamine respectively are covered by this laser.
Although gas lasers are commonly used, there are newer light sources such as diode lasers, and fiber lasers that are gaining more popularity.
They generate less heat, enhance stability, uniformity, and they are able to emit wavelengths covering a wider range of the visible spectrum.
- Confocal scan head: equipped with pinholes to remove out-of-focus light, and an array of photomultiplier tubes (PMT) for detecting photons emitted by the specimen. There are typically 3 PMTs that collect green, red, and blue light. However, in the case of collecting reflected or transmitted light, extra PMTs may be utilized.
PMTs are distinct from digital cameras in that they are vacuum tubes with photons entering at one end and an electron-multiplying component present in the tube body. An electrical signal is generated by converting the collected photons prior to the image assembly and display.
Confocal systems enable image capture using both contrast and fluorescence illumination simultaneously. The images are then merged and analyzed by image analysis software built within the computer.
- One notable feature of confocal microscopy is its ability to generate sharp images of the precise plane of focus, without interference from the background or other specimen regions. This allows you to easily visualize structures within even thicker objects. This, in turn, eliminates the blurring effects of fluorescent light from surrounding areas, making it an ideal technique for such applications.
- Additionally, confocal microscopy allows you to analyze 3D structures by stacking multiple images using different optical planes. However, it’s important to note that the penetration depth of the sample can be limited in confocal microscopy.
- One primary benefit of confocal microscopy is that it enables the user to select specifically defined regions of interest, eliminating the requirement of exposing the entire specimen to the fluorescent light source.
- Furthermore, confocal microscopy can generate optical sections through the specimen, which can effectively minimize the out-of-focus fluorescence.
- Confocal microscopes tend to be expensive, making them less accessible for some research labs or institutions.
- Operating and optimizing a confocal microscope requires specialized training and expertise, especially when configuring imaging parameters and optimizing image acquisition settings.
- Although confocal microscopy reduces phototoxicity compared to widefield microscopy, extended exposure to laser light can still cause photodamage and affect cell viability in live cell imaging experiments.
- The scanning mechanism and image acquisition process in confocal microscopy can be time-consuming, especially for large areas or high resolyion imaging, limiting its throughput.
Two things have led to the recent advancements in fluorescence microscopy beyond widefield and confocal setups.
1) Limitations in widefield and confocal setups
2) Advancements in imaging technologies, including the increasing capabilities of computer software for image acquisition and analysis
Different microscopy techniques were subsequently developed in recent years after widefield and confocal microscopy. Advancement in one technique triggered innovation in the other.
Based on these new techniques, novel assays have also been developed.
In terms of software development, image analysis algorithms, machine learning approaches, and data visualization tools have been developed to extract quantitative information from large and complex imaging datasets. This enables researchers to obtain precise measurements and perform quantitative analyses of protein functions and processes in a high-throughput manner.
It is extremely intriguing how resourceful researchers have consistently turned technological limitations into benefits.
For instance, what may be viewed as a drawback or unwanted artifact by one researcher such as photobleaching, was ingeniously employed by another researcher.
Some of the commonly used advanced imaging techniques are discussed below.
FRAP (fluorescence recovery after photobleaching) is a fluorescence microscopy technique that enables you to study how fluorescently-labeled molecules move and behave within living samples.
FRAP is frequently employed to examine
- molecule diffusion within cells
- studying protein-binding interactions
- characterizing biomembrane fluidity
- turnover rates
This enables a deeper understanding of the dynamic behavior of biomolecules in their native cellular environment.
FLIM (fluorescence lifetime imaging microscopy) is an advanced and powerful technique for analyzing and visualizing the spatial distribution of specific cellular components. It provides valuable insights into the localization and dynamics of biomolecules like proteins and nucleic acids within cells.
- It not only provides information about individual molecules but also allows researchers to visualize the dynamic cellular environment surrounding these molecules in living cells.
- Various physiological factors like pH, oxygen concentration, calcium ion levels, molecular interactions and bindings can be measured using FLIM.
- FLIM, in comparison to standard microscopy techniques, offers a greater penetration depth, allowing researchers to analyze and visualize cellular structures and processes in thicker samples.
TIRF (Total internal reflection fluorescence) is an advanced fluorescence microscopy method to visualize cellular events happening at or close to the cell membrane in living cells with a high signal-to-noise ratio.
TIRF is not ideal for visualizing structures that are located deep inside a specimen.
- Capable of investigating various cellular processes such as cell adhesion, signal transduction, endocytosis and exocytosis.
- Quantify the rate at which receptors endocytose in response to ligand binding.
- Can monitor the opening and closing of receptor channels.
- Help determine the function of various proteins like clathrin and adaptor proteins in exocytosis/endocytosis pathways.
FRET (Fluorescence resonance energy transfer) is a robust technique that helps determine the exact location and spatial proximity of molecules labeled with fluorescent probes.
This technique not only provides valuable insights into the colocalization of molecules but also enables analysis of molecules’ associations or interactions within living cells, shedding light on cellular processes and molecular mechanisms in a non-invasive manner.
- FRET helps monitor the intracellular calcium ion signaling in various cellular compartments including endoplasmic reticulum, cytoplasm etc. In addition to detecting ions within cellular compartments, FRET constructs have been employed to study the subsequent events that occur in response to second messenger signaling.
- It enables researchers to study the interactions between receptor and ligand molecules, receptor dimerization, and lipid dynamics between vesicles.
- FRET is also very useful in analyzing the structure, conformation, and hybridization of nucleic acid molecules which helps to map the genome and diagnose genetic mutations.
- STED (stimulated emission depletion)
- STORM (stochastic optical reconstruction microscopy)
- SSIM (saturated structured illumination microscopy)
- PALM (photoactivated localization microscopy)
Super-resolution microscopy is a technique developed to visualize ultra-small structures within living cells or tissue that fail to be resolved by conventional widefield or confocal microscopy.
Super-resolution microscopy pushes beyond the limitations of diffraction, providing impressive 3D resolution. Using this technique, you can gain unprecedented insights into a cell’s intricate structural organization and the behavior of biomolecular assemblies at nanoscale levels.
Some examples of super-resolution microscopy include
Brown. 2007. Fluorescence microscopy-avoiding the pitfalls. J Cell Sci. 120(Pt 10):1703-5. doi: 10.1242/jcs.03433
Chen et al. 1997. Fluorescence energy transfer detection as a homogeneous DNA diagnostic method. PNAS. 94(20): 10756–10761. doi: 10.1073/pnas.94.20.10756
Elliott. 2020. Confocal Microscopy: Principles and Modern Practices. Curr protoc cytom. 92(1). doi: 10.1002/cpcy.68
Fish. 2015. Total Internal Reflection Fluorescence (TIRF) Microscopy. Curr protoc cytom. doi: 10.1002/0471142956.cy1218s50
Ishikawa-Ankerhold. 2012. Advanced Fluorescence Microscopy Techniques—FRAP, FLIP, FLAP, FRET and FLIM. Molecules. 17(4): 4047–4132. doi: 10.3390/molecules17044047
Lippincott-Schwartz et al. 2018. The Development and Enhancement of FRAP as a Key Tool for Investigating Protein Dynamics. Biophys J. 115(7):1146-1155. doi: 10.1016/j.bpj.2018.08.007
Nakabayashi. 2017. Application of Fluorescence Lifetime Imaging (FLIM) to Measure Intracellular Environments in a Single Cell. 1035:121-133. doi: 10.1007/978-3-319-67358-5_8
Roy and Perrin. 2018. The stable actin core of mechanosensory stereocilia features continuous turnover of actin cross-linkers. MBoC. 29(15):1856-1865. doi: 10.1091/mbc.E18-03-0196
Sekar and Periasamy. 2003. Fluorescence resonance energy transfer (FRET) microscopy imaging of live cell protein localizations. J Cell Biol. 160(5): 629–633. doi: 10.1083/jcb.200210140
Yang et al. 2021. Super-resolution Microscopy for Biological Imaging. Adv Exp Med Biol. 3233:23-43. doi: 10.1007/978-981-15-7627-0_2