Introduction

Fluorescence microscopy is a powerful imaging technique widely used in life sciences and medical research to study cell and tissue morphology¹. By using specific fluorescent stains and dyes, researchers can visualize and differentiate cellular structures with high specificity and contrast.

Traditional brightfield microscopy relies on contrast to distinguish cellular components, but it has limitations in studying dynamic and molecular-specific processes². Fluorescence microscopy overcomes these limitations by allowing visualization of dynamic cellular processes, the study of biomolecular interactions and stoichiometric measurements at the molecular level². The introduction of fluorescently labeled antibodies in the 1940s enabled researchers to achieve molecular-specific imaging of cells and subcellular structures, greatly expanding the scope of biological discovery.

Immunocytochemistry/immunofluorescence - anti-Cytokeratin 18 antibody [SP69] (ab93741); from: Anti-Cytokeratin 18 antibody [SP69] (ab93741) | Abcam

Immunocytochemistry/ Immunofluorescence analysis of HeLa (human cervix adenocarcinoma epithelial cell) cells labeling Cytokeratin 18 with purified ab93741 at 1:100 (2.4 µg/mL). Cells were fixed in 4% paraformaldehyde and permeabilized with 0.1% Triton X-100. Cells were counterstained with Alexa Fluor^® 594 Anti-alpha Tubulin antibody [DM1A] - Microtubule Marker (ab195889). Anti-alpha Tubulin antibody [DM1A] (ab7291) - Microtubule Marker (Alexa Fluor^® 594) 1:200 (2.5 µg/mL). Goat anti-rabbit IgG (Alexa Fluor^® 488, Goat Anti-Rabbit IgG H&L (ab150077) was used as the secondary antibody at 1:1000 (2 µg/mL) dilution. DAPI nuclear counterstain. PBS instead of the primary antibody was used as the secondary antibody-only control.

Fluorescence microscopy techniques are essential for analyzing protein-protein interactions, which are vital for cellular function and homeostasis. Methods such as Förster resonance energy transfer (FRET)³ and bimolecular fluorescence complementation (BiFC)⁴ allow researchers to probe interactions with high sensitivity and spatial resolution.

Fundamentals of fluorescence microscopy

Fluorescence microscopy operates by exciting fluorophores, molecules that can re-emit light, using specific wavelengths of light⁴. Upon excitation, these fluorophores emit light at longer wavelengths, enabling highly specific imaging of biological structures. Common fluorophores include fluorescent dyes, nanostructures, biomolecule analogs, and fluorescent proteins.

Here’s a step-by-step breakdown of the fluorescence process⁴:

1. Excitation of fluorophores: Fluorophores absorb photons from an external light source, typically in the ultraviolet or visible range. This absorption process, known as photoluminescence, elevates the electrons within the molecule to an excited state.

2. Energy dissipation (occurs in picoseconds): Once excited, the electrons undergo rapid relaxation through two main mechanisms:

a. Non-radiative vibrational relaxation: The excess energy is dissipated as heat within the same electronic state without light emission.

b. Internal conversion: The electrons transition between adjacent electronic energy levels, losing energy non-radiatively.

3. Emission of fluorescence (occurs in nanoseconds): The electrons eventually return to the ground state (S₀), releasing energy in the form of visible light. This emitted light, known as fluorescence, is detected using photosensitive devices such as cameras or photomultiplier tubes.

4. The Stokes shift: The difference between the excitation and emission wavelengths is called the Stokes shift⁵. Emission always occurs at a longer wavelength due to the energy lost during non-radiative processes. A larger Stokes shift is advantageous because it reduces signal overlap, minimizes self-absorption, and improves resolution in experiments involving multiple fluorophores^{6, 7}.

The fluorescence mechanism complements techniques like electrophysiology and optogenetics, enabling advancements unattainable by single methods². Its capabilities include multicolor imaging, fluorescence lifetime imaging (FLIM), and FRET for detailed molecular studies.

Key components of a fluorescence microscope

Fluorescence microscopy relies on components like light sources, mirrors, lenses, filters, and detectors, all contributing to image quality and signal accuracy¹. Proper selection of optical elements, such as cover glass thickness and immersion fluid, is essential for optimal performance. Regular maintenance, like alignment of optical elements and cleaning of lenses, is important for accurate and reliable imaging.

Light sources

When light sources are considered, the term “Kohler illumination” comes into play, ensuring the even distribution of light across the field of view of the specimen⁸. Fluorescent sample illumination uses arc lamps like mercury lamps and inert gas lamps like xenon lamps, which emit multiple wavelengths requiring filters. Laser lamps can also be used that provide precise single-wavelength excitation. Fluorophores emit light at longer wavelengths after excitation, captured by photosensitive surfaces.

Filters

The excitation filter blocks unwanted light, allowing only specific wavelengths to excite the fluorochrome, enhancing fluorescence signal specificity⁹. Monochrome astronomy filters can also be used, offering low-energy excitation to reduce photobleaching and specimen damage.

The emission filter blocks excitation wavelengths, allowing only emitted fluorescence to pass, ensuring a high signal-to-noise ratio in images⁹. Advanced composite emission filters can enhance fluorescence signal quality by matching specific fluorescent protein spectra.

The dichroic mirror reflects excitation light toward the specimen while transmitting emitted fluorescence to the detector, ensuring efficient imaging⁹. Some systems eliminate the need for specialized filter cubes, enabling the use of standard laboratory microscopes.

Objective lens and detectors

The objective lens and digital detector system in fluorescence microscopes significantly impact performance and application, with detectors categorized as area or electronic types⁸.

• Area detectors like charge-coupled devices (CCD) and complementary-metal-oxide-semiconductor (CMOS) capture photons simultaneously, suitable for high-speed imaging
• Electronic detectors like photomultiplier tubes (PMTs) scan point-by-point, prioritizing spatial resolution.

Signal regulation in both types depends on factors such as laser intensity, exposure time, and amplification settings. Further, cooled scientific-grade cameras with low readout noise are essential for detecting dim fluorescent signals¹⁰.

Importance of fluorophores, fluorescent dyes, and proteins

Fluorophores differ in brightness and photobleaching rates. Brighter and more photo-stable options maximize the signal. The brightness depends on the extinction coefficient and quantum yield, influenced by the fluorophore environment.

Fluorescence-based dyes are small organic probes offering rapid, selective, and sensitive detection of target analytes¹¹. They are cost-effective, easy to use, and versatile across various analytical conditions. Two widely used fluorescent dyes are Hoechst 33258 and 4ʹ,6-diamidino-2-phenylindole (DAPI), each serving unique purposes in research applications. A few stains and their selective targets are⁸

• Nucleic acids (eg, DAPI)
• Biological membrane lipids (eg, Nile Red, FM dyes)
• Cellular structures like actin filaments (eg, fluorophore-derivatized phallotoxins)

Fluorescent proteins are widely used in microscopy to study protein expression, localization, dynamics, and interactions by genetically fusing them to target proteins¹². Green fluorescent protein (GFP) derived from Aequorea victoria is commonly used. Other alternatives can be red fluorescent protein (RFP) and yellow fluorescent protein (YFP), each providing unique spectral properties.

A few derivatives of GFP, like enhanced GFP (eGFP) and pHluorin-mKate2 (pH-sensitive GFP), have also been documented¹³. Fluorescent nucleoside analogs have been used to study nucleic acid interactions, an example being a highly fluorescent thymine analog, TexT, used to study immune nucleic acid detectors, and toll-like receptors (TLRs)¹⁴.

Other examples include metal-ion complexes (that have the advantages of long-lived emission and larger Stokes shifts, circumventing self-quenching) and zero-dimensional fluorescent nanomaterials called graphene dots. Fluorescence microscopy also uses fluorophores like organic fluorophores (like the Alexa Fluor^®* series) to visualize cellular structures and dynamics, suitable for live-cell imaging and subcellular studies⁸.

Types of fluorescence microscopy

There are several types of fluorescence microscopy, each with its advantages and limitations. Here is a detailed overview of different types of fluorescence microscopes.

Widefield fluorescence microscopy

The widefield fluorescence microscope has become essential for cell biology, offering faster imaging with reduced photobleaching and phototoxic effects⁸. It illuminates the entire field of view. However, it may collect out-of-focus information that compromises resolution. Resolution can be improved through structured illumination or post-acquisition methods like deconvolution, though original images must be compared to avoid artifacts.

Confocal fluorescence microscopy

Fluorescent confocal microscopy uses a laser to scan specimens point-by-point, removing stray light with a pinhole near the detector for precise optical slicing⁸. It enables imaging of thick specimens without tissue sectioning. However, higher laser intensity is often required for this microscopy, which may increase photobleaching and phototoxic effects on cells. Technical advances include white-light and supercontinuum lasers, along with hybrid photomultiplier tube-avalanche photodiode detectors¹⁵.

The accuracy of fluorescence confocal microscopy images of adipose tissue was on par with conventional histology without the frozen-section-related artifacts¹⁶. In another study, the diagnostic power of confocal microscopy was similar to frozen sections and facilitated analyzing structural changes and infiltrates in steatosis, facilitating a new avenue in liver transplant research¹⁷.

Two-photon microscopy

Two-photon, or multiphoton, microscopy uses pulsed near-infrared lasers that enable precise excitation at specific planes without requiring a pinhole⁸. This reduces photobleaching and phototoxicity. This technique allows deeper tissue penetration for long-term imaging of cellular processes but faces resolution challenges at greater depths. This challenge can be tackled by using adaptive optics. Additionally, it supports label-free imaging of biological structures by capturing harmonic generation signals.

Total internal reflection fluorescence (TIRF) microscopy

TIRF microscopy excites fluorophores in a thin axial region (optical region) near the solid-liquid interface using an evanescent wave generated by totally internally reflected light¹⁸. The principle is that the complete internal reflection of excitation light in a transparent solid (like a cover glass) at its interface with liquid generates an electromagnetic field called the evanescent wave at the same frequency as the excitation light.

TIRF provides high-contrast imaging of fluorophores near the plasma membrane with minimal background from the cell interior. It also reduces cellular photodamage and enables rapid exposure time for efficient imaging. TIRF microscopy has been used to study receptor behavior at the plasma membrane in neurons, receptor endocytosis, visualizing the plasmalemmal surface and subplasmalemmal zone, and single-molecule analysis of ion channels¹⁹. A recent development is supercritical illumination microscopy photometric z-localization with enhanced resolution (SIMPLER) to deduce the axial position of single molecules in a TIRF microscope with sub-10 nm axial localization precision²⁰.

Super-resolution fluorescence microscopy

Conventional fluorescence microscopes face resolution limits due to light diffraction, defined by the Abbe and Rayleigh criteria⁸. Resolution is traditionally improved using shorter wavelengths and high-numerical aperture (NA) objectives but remains constrained by diffraction limits. Super-resolution microscopy techniques such as stimulated emission depletion (STED), photoactivated localization microscopy (PALM), stochastic optical reconstruction microscopy (STORM), and structured illumination microscopy (SIM) overcome the diffraction limit of light microscopy, enabling imaging at nanoscale resolutions.

These methods differ in their mechanisms, applications, and trade-offs, making them suitable for distinct biological and materials science research needs. Below is a detailed comparison and explanation of each technique.

Light sheet microscopy

Light sheet microscopy uses perpendicular illumination and detection paths for rapid imaging at cellular and subcellular resolutions⁸. This method selectively illuminates specific specimen planes, significantly reducing fluorophore bleaching and phototoxic effects.

Thus, it is ideal for live-cell imaging. The technique allows sample rotation for multi-angle imaging and often requires unconventional mounting approaches to maintain sample health for long-term studies. However, it generates massive image data, requiring robust computer hardware and software for storage and processing. Research is on to overcome the limitations, such as on-the-fly imaging in real-time, deep learning, and compressed sensing²¹.

Advanced fluorescence imaging techniques

Fluorescence lifetime imaging microscopy (FLIM)

Fluorescence lifetime imaging microscopy is a technique that provides molecular insights by measuring fluorescence decay time, and it has broad research applications²². FLIM captures the temporal aspects of fluorescent decay rather than time-integrated signals, unlike traditional intensity-based methods.

FLIM techniques are categorized into frequency-domain and time-domain methods.

• Frequency-domain FLIM uses modulated light to measure phase shifts and modulation depths, determining fluorescence lifetime by comparison with a reference fluorophore.
• Time-domain FLIM employs pulsed laser excitation and measures fluorescence decay over discrete time channels using ultrafast detectors.

Examples of the use of FLIM include early studies on in vivo imaging of skin, metabolic alterations like redox ratio analysis in organoids and tumors, mitochondrial dynamics, and the time course of apoptosis in zebrafish²³.

Recent advancements focus on improving its speed and three-dimensional imaging capabilities, such as the single-sample image-fusion upsampling based on data-fusion in computational FLIM super-resolution to augment image resolution at high acquisition²⁴.

Förster resonance energy transfer (FRET) imaging

Förster resonance energy transfer is a photophysical process where energy is transferred from a donor fluorophore to an acceptor molecule over distances of 1 to 10 nm²⁵. A combination of quantum mechanics and Coulombic dipole-dipole interactions explains this radiationless transfer. FRET efficiency is commonly used to measure the relative distance between donor and acceptor pairs. FRET can be used to measure the nanoscale distance between two interacting biomolecules, making it highly effective for studying nanoscale interactions in biological systems²⁶. Examples of this application include a DNA-nanotweezer²⁷ with FRET to image mRNA and in vivo pharmacokinetics²⁸ of the organic nanomedicine carriers. FRET has been used for interactions between proteins like G-protein coupled receptor 143 (GPR143) with the melanogenic enzyme tyrosinase (TYR) and conformational rearrangements or changes like the release of calcium in response to ER barrier perturbations²⁹.

Efficient FRET requires three key conditions³⁰:

• There must be an overlap between the donor’s emission spectrum and the acceptor’s excitation spectrum.
• The donor and acceptor must be in close proximity.
• Favorable mutual dipole orientation must be established.

Fluorescence recovery after photobleaching (FRAP)

Fluorescence recovery after photobleaching is a widely used technique for studying biomolecular binding and diffusion kinetics³¹. FRAP has been applied to explore lipid raft properties, cytoplasmic viscosity regulation, and biomolecular dynamics in phase-separated condensates. Its versatility and effectiveness have driven best practice developments, biological insights, and ongoing advancements in biophysics research, such as insights into mRNA³² transport in oocytes, and intermediate filament³³ dynamics.

FRAP involves the following:

• Labeling biomolecules with fluorescence.
• Photobleaching a region with high-intensity light.
• Monitoring fluorescence recovery over time.

The resulting FRAP curve plots fluorescence intensity changes in the bleached region, providing insights into molecular dynamics. A modified approach of FRAP called Line-FRAP was developed to measure photobleaching recovery under a confocal microscope to facilitate diffusion rate measurements in many environments using standard equipment³⁴.

Interpreting fluorescence microscopy images

Interpreting the images formed through the fluorescence microscope is an important factor in deciding the quality of images. There are several methods of analyzing the images, such as:

• Colocalization analysis often involves merging pseudo-colored fluorescence images to visually identify overlapping signals, such as yellow pixels from red and green fluorophores. This qualitative method suggests that both fluorophores are within the same 3D volume.
• Image deconvolution in fluorescence microscopy enhances contrast, making it easier to detect small, dim objects. These algorithms also help reduce out-of-focus fluorescence in digital images. This technique is typically applied after image acquisition for improved clarity.

Image acquisition and processing

Quantitative microscopy measurements are typically made using digital images³⁵. These images are formed when a microscope’s optical image is captured by a detector, such as a CCD camera or PMT, on a grid of pixels. Each pixel represents a defined area and converts detected photons into an intensity value correlated to the photon count.

Fluorescence microscopy pixel intensity reflects the number of fluorophores in a specimen area. Digital images from this technique provide spatial information for calculating distances, areas, and velocities. They also offer intensity data to determine fluorophore concentrations locally.

Challenges in image analysis

Fluorescence microscopy is an excellent tool for quantitative measurements, given that optical sectioning allows quantification with spatial precision. However, meticulous planning and understanding are vital to obtaining meaningful results.

Some of the challenges in image analysis are³⁵:

• Noise creates variations in pixel intensity values, leading to measurement imprecision and uncertainty in digital images. Detecting a signal requires its intensity to be significantly higher than the noise level.
• Measuring fluorescence in deep specimens is challenging due to light scattering and optical aberrations, which reduce signal intensity and are difficult to correct in biological samples. While multi-photon illumination minimizes scattering, optical aberrations and increased photobleaching limit measurement accuracy.
• Uneven illumination affects quantitative measurements by causing variations in intensity for objects with identical fluorophore concentrations across the field of view. Using techniques like scrambling the light source image with a liquid light guide can improve illumination uniformity.
• Bleed-through may occur due to overlapping spectra in the fluorophore’s emission passing through another filter set. Selecting fluorescence filter sets requires balancing optimal excitation and emission collection by minimizing bleed-through and autofluorescence.
• Dim light sources, reduced exposure times, and optical filters in live-cell fluorescent microscopy minimize phototoxicity but result in weak signals.

Applications of fluorescence microscopy

Fluorescence microscopy is widely used in cell biology, molecular biology, and biomedical research to visualize cellular structures and dynamic processes. Its applications include studying protein interactions, tracking live cell activities, and detecting disease biomarkers. For example, FISH is a standard technique in routine diagnosis of genetic aberrations, used in many genetics labs³⁶. The expression status of human epidermal growth factor receptor 2 (HER2) is tested by immunohistochemistry (IHC) and FISH³⁷; HER2 status is more commonly evaluated using FISH and is part of the standardized criteria for diagnosis in the 2018 American Society of Clinical Oncology (ASCO)/College of American Pathologists (CAP) HER2 testing guideline update.

Biological and medical applications

• Fluorescent probes are used to monitor the structure and function of organelles, as well as pH levels and protein transport³⁸. These probes include chemical dyes and genetically encoded protein sensors, each offering unique advantages like brightness and high specificity.
• High-content analysis (HCA) screens use probes to study organelle dysfunction, requiring fluorophores with high photostability, sensitivity, and minimal toxicity.
• Live-cell imaging allows the observation of dynamic processes such as mitochondrial fission, Golgi trafficking, and lysosomal activity³⁹. It enables real-time monitoring of cellular division and DNA replication.
• Fluorescent biosensors are used to measure the ion concentrations, such as calcium. Further, it helps in tracking drug effects on cellular health and metabolism. For example, fluorescent probes like curcumin, Nile Red, and fluorescein have been used in nanocarriers to assess the biodistribution of drug molecules in the eye, with a commercial example being indocyanine green as an FDA-approved near-infrared fluorescence (NIRF) dye to image retinal blood vessels⁴⁰.
• Fluorescence microscopy can be used to visualize the localization and interactions of specific molecules. For example, the interest in O-glycosylation (O-GalNAc-type) type modifications in glycoproteins has received attention given its role in metabolism, microbiome, and disease, leading to research on labeled antibodies to study this process^41, 42. Immunofluorescence (IF) is an antibody-based staining technique that uses immunoglobulins coupled to fluorescent dyes for visualizing proteins in cells or tissues⁹. Selectivity is achieved through specific primary antibodies like the Biotin Anti-Rotavirus NCDV antibody that reacts with rotavirus samples for fluorescent microscopy studies.
• Fluorescence in situ hybridization (FISH) uses fluorescent probes to hybridize with complementary mRNA or DNA sequences in fixed specimens. Multicolor FISH probes enable labeling multiple targets⁸. It is widely used in research to study mRNA expression patterns, subcellular enrichment, locate specific DNA sequences⁴³, diagnosis of genetic diseases, gene mapping, and identification of novel oncogenes or genetic abnormalities associated with disease.

Material science applications

Fluorescent nanoparticles offer superior optical properties, high brightness, and outstanding photostability, making them promising alternatives to dyes and fluorescent proteins. Given the challenge of photobleaching, their small sizes enhance quantum effects, and their functionalized surfaces enable biocompatible, target-specific labeling for advanced microscopy techniques.

Synthetic nanoparticles offer high brightness across the visible spectrum and exceptional photostability, making them superior to traditional probes⁴⁴. Their fluorescent cores and functionalized surfaces enable target-specific and biocompatible labeling in biological microscopy. For example, carbon dots have the advantages of limited photobleaching, stability, small sizes to pass through organelles, non-toxicity, and excitation wavelength-dependent and/or -independent fluorescent emission⁴⁵. They have been used in initial studies for photoimaging, drug delivery studies, environmental monitoring, and detection of pathogens⁴⁵.

Quantum dots (QDs) are fluorescent semiconductor nanocrystals used in biomedical applications like drug delivery and cellular imaging. Their unique feature is that emissions can be tuned with size (broad excitation windows; narrow emission peaks), high brightness, and stability, making them valuable for medical imaging⁴⁶. In addition to in vitro tracking and imaging studies, initial studies have shown the promise of QDs for non-invasive tracking and imaging. Another interesting avenue of research is the potential of graphene quantum dots to detect chloride ions for future cystic fibrosis detection and killing of cancer cells by aluminum phthalocyanine (AlPcS) and CdSe/CdZnS core-shell structure QDs, highlighting their potential⁴⁷.

Advantages and limitations of fluorescence microscopy

Fluorescence microscopy offers high specificity to visualize live cells and multiple targets simultaneously. However, it has limitations such as photobleaching, phototoxicity, and the need for expensive equipment that limits its applications.

Advantages

• Small organic fluorophores and genetically encoded proteins enable selective visualization of nucleic acids, proteins, and other cellular components⁴⁸. They offer high specificity by binding exclusively to target molecules, reducing background noise. For example, a study⁴⁹ used bioorthogonal click chemistry of the strain-promoted inverse electron-demand Diels–Alder cycloaddition between site-specifically incorporated unnatural amino acids and tetrazine dyes to label proteins in living primary mouse neurons. Their high sensitivity allows the detection of even low-abundance molecules, making them essential for applications like rare biomarker identification in cancer research⁵⁰. For example, the blue-yellow mTurquoise2 (donor) and sEYFP (acceptor) pair in FRET provides a Förster radius of 5.9 and ∼2–8 nm sensitivity⁵¹.
• Fluorescence imaging enables real-time monitoring of dynamic cellular processes without disrupting the natural environment³⁹. This is an advantage over the electron microscope, which does not allow live cell visualization.
• Multicolor fluorescence microscopy enables simultaneous visualization of multiple cellular targets using distinct probes with separate excitation and emission spectra⁸. This multicolor imaging allows researchers to study complex biomolecular interactions. For instance, a multicolor fluorescence microscope system under a selective plane illumination microscopy was developed to visualize the localization and dynamics of cellular components functioning in apical growth in living Neurospora crassa hyphae⁵². The application of multicolor fluorescence microscopy has been reported for real-time⁵³ tracking of drug delivery to cancer cells and observing previously “unseen”⁴⁴ structures of neurons.

Limitations

• Fluorophores, including fluorescent proteins, undergo photobleaching when exposed to excitation light for more extended periods⁵⁴. Anti-photobleaching reagents can be used in the mounting medium to reduce photobleaching for certain specimens.
• In digital microscopy, spatial resolution depends on both the microscope and detector, affecting the ability to locate and distinguish close objects precisely⁵⁴. Spatial resolution relies on the image signal-to-noise ratio (SNR), as undetectable objects cannot be resolved. When imaging dynamic specimens, temporal resolution or the image acquisition rate further impacts measurement accuracy.
• Fluorescence microscopes can cost thousands to hundreds of thousands of dollars, making them expensive⁵⁵. As a result, their use is mainly restricted to well-funded institutions, research facilities, and medical laboratories.

Future direction and emerging technologies

Future directions in fluorescence microscopy involve advancements in imaging technologies, such as super-resolution techniques and artificial intelligence (AI)-driven image analysis. These innovations are expanding applications in many fields, like studying neuroscience, cancer, and drug discovery, while enhancing imaging precision and data processing.

Ultrafast widefield systems enhance light collection and spatial resolution for high-throughput imaging. AI is revolutionizing fluorescence microscopy by enhancing image quality and introducing advanced imaging modalities. For example, virtual refocusing helps 3D imaging using a series of 2D images and AI to produce clean outputs from noisy inputs (denoising), often better than traditional methods based on manually defined parameters⁵⁶. In another study, a portable AI fluorescence microscope (πM) with edge computing techniques was developed for real-time targets.

A unified framework categorizes AI-driven methodologies into linear inverse problem-solving and nonlinear prediction. Self-supervised learning techniques further improve data quality, with applications extending to other optical microscopy methods. Quantitative phase imaging enables virtual quantitative fluorescent imaging of live organoids, which are 3D models of studying organs and relevant research⁵⁷.

In cancer research, FLIM⁵⁸ has been used in many applications, such as two-photon⁵⁹ fluorescence lifetime imaging microscopy (2P-FLIM) of metabolites like reduced nicotinamide adenine dinucleotide to distinguish aggressive and non-aggressive cancers and monitoring⁶⁰ histone lysine acetylation with a P300/CBP-associated factor (PCAF) biosensor (HATS) with FLIM.

In pharmacology, FLIM has been used for drug screening, such as assaying⁶¹ the sensitivity of glioma cells derived from patients to temozolomide and identifying⁶² that drugs that could inhibit Drp-1 and simultaneously have antioxidant properties like resveratrol could target diabetic cardiomyopathy better than commercial single-target antioxidant drugs. FLIM has been used in disease modeling, such as studying mitochondrial⁶³ function to assess bioenergetic alterations in hippocampal neuron models of Alzheimer’s disease (AD) and identifying spatial⁶⁴ changes in oxidative phosphorylation in 3D breast spheroids.

The integration of fluorescence microscopy with optogenetics enables precise control and observation of cellular activities, advancing research in neuronal phenotyping and electrophysiology⁶⁵. The synergy allows simultaneous photostimulation and fluorescence imaging. This combination is expected to drive innovations in neuroscience and broader research applications.

FAQs

What are the advantages of using confocal microscopy over widefield microscopy?

Confocal microscopy provides higher resolution and clearer images by eliminating out-of-focus light through optical sectioning. It allows for detailed 3D reconstruction of specimens by capturing multiple focal planes. This technique also enhances contrast and is suitable for thick or complex samples.

How do fluorophores differ from fluorescent proteins in fluorescence microscopy?

Fluorophores are synthetic chemical compounds that emit fluorescence upon excitation, while fluorescent proteins are naturally occurring or engineered proteins with similar properties. Fluorophores are versatile and can be conjugated to various molecules, whereas fluorescent proteins are genetically encoded and expressed directly in living cells. Fluorescent proteins are often preferred for live-cell imaging due to their biocompatibility.

What are the key applications of live cell fluorescence microscopy?

Live cell fluorescence microscopy is crucial for studying dynamic cellular processes in real-time, such as cell division, migration, and intracellular trafficking. It is widely used to track protein localization, interactions, and signal transduction pathways in living cells. Researchers also use it for drug screening and understanding disease mechanisms at the cellular level.

*Alexa Fluor^® is a registered trademark of Molecular Probes, Inc., a Thermo Fisher Scientific Company. Alexa Fluor^® dye conjugates contain(s) technology licensed to Abcam by Life Technologies.

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