Fluorescence Microscopy: Comparison
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Fluorescence microscopy has become a critical tool for researchers to understand biological processes at the cellular level. Micrographs from fixed and live-cell imaging procedures feature in a plethora of scientific articles for the field of cell biology.

  • luorescence microscopy
  • microscopy techniques
  • imaging agents
  • cellular imaging

1. Introduction

Since the inception of fluorescence microscopes in the early 1900s, their use as a research tool for observing discrete subcellular structures and processes has grown immensely [1]. Initially, auto-fluorescent specimens were visualised with fledgling fluorescence microscopes, until the introduction of fluorescent stains in the 1930s, which enabled non-fluorescent specimens to be visualised [2]. The use of fluorescently-labelled antibodies in the 1940s [3] enabled precision visualisation of target structures in cells, and with the Nobel prize winning discovery of green fluorescent proteins (GFP) in the 1960s and their subsequent development as a genetic tag [4], the field has evolved to permit targeted live cell imaging. The discovery of novel fluorescent markers has driven the development of fluorescence-competent microscope technologies, further enabling researchers to discover and understand the intricate dynamics of subcellular biology and their detailed mechanics at ever-higher spatial and temporal resolutions.

2. Fluorescence Microscope Hardware Systems

2.1. Multiphoton and Other Advanced Microscopy Techniques

Modifying the laser excitation source of a confocal microscope with an ultra-fast pulsed laser enables two- or three-photon microscopy [5]. Ultra-fast lasers allow deeper penetration of samples using longer excitation wavelengths, and optical sections are created through confinement of the two (or multi) photon effect to a single focal plane. This technique is suited to imaging large tissue samples, where penetration depths of 100 μm or more are needed to investigate macro-scale biological processes. Multiphoton microscopes are also needed for intravital imaging, where both gentle illumination and imaging depth are critical for imaging live whole organisms. However, multiphoton microscopes are limited by the added expense of purchasing an additional ultra-fast laser, which can add significantly to the cost of ownership and operation. Laser scanning microscopes have been adapted to observe dynamic events in cellular function and exploit the properties of imaging probes, such as fluorescence lifetimes, and their interactions within and between cells. The three most common technologies employed are fluorescence lifetime imaging microscopy (FLIM) [6], fluorescence resonance energy transfer (FRET) [7], and fluorescence recovery after photobleaching (FRAP) [6]. FLIM requires the observation of intensity changes in a fluorophore over a course of time, which has been useful for quantifying decay rates of cell metabolites [8]. FRET measures energy transfer from one fluorophore to another using a single excitation, which allows for a variety of applications, such as calcium imaging [9] and protein–protein interactions/transfers [10]. Finally, FRAP measures the recovery in signal of a molecular probe after photobleaching, which enables the observation of diffusion kinetics in both cells and tissue samples [11]. Another recent technique, used to image larger tissue samples quickly and efficiently, is light-sheet fluorescence microscopy (LSFM) [12]. Light-sheet microscope operation differs from confocal microscopy, as samples are excited using a sheet of light from the side, as opposed to a focussed beam of light from the top or bottom of a sample. Emitted light is then detected perpendicular to the excitation sheet, unlike a confocal microscope, which detects the emitted signal in the opposite direction to the excitation beam. The thickness of the sheet of light determines the thickness of the optical section captured, and the sample can be rotated and moved through this plane to produce a three-dimensional tomographic image. Light-sheet fluorescence microscopy enables imaging of larger, multicellular samples, such as organoids and whole organisms (e.g., insects, plants, and animals [13][14][15]) that may not be adequately captured in a timely manner using single or multi-photon microscopy, although this does require tissue-clearing methodology to create optically transparent samples [16][17][18].

2.2. Super Resolution Microscopy (SRM); Going beyond the Limits of Light

Improvements in laser scanning microscopes have enhanced imaging resolutions beyond Abbe’s limit (Equation (1)), the physical limitation due to the physics of diffraction, for example, a 200 nm resolution limit in air when using an excitation wavelength of 400 nm [19]; however, compromises such as hardware cost, phototoxicity, and low capture speed, remain limitations. The simplest technique to improve resolution using a confocal microscope is to restrict pinhole size, perform a z-stack, which involves combining multiple images captured at sequential focal planes, and perform post-processing of images using software deconvolution. However, acquiring a suitable z-stack of images involves repeated laser excitation, which significantly increases both cellular phototoxicity and the time to capture the complete micrograph [20].
(1)
Equation (1) is Abbe’s formula for the resolution limit, where d is the minimum distance that can be differentiated between two points, λ is the wavelength of light, n is the refractive index of the medium the light is traversing, θ is the angle at which the light is converging, and NA is the numerical aperture [19]. Another approach to improve resolution is stimulated emission depletion (STED) [21], which makes use of two lasers; one for excitation, and the other shaped into a ring or doughnut, used as a de-excitation or depletion spot to limit the size of the emitted fluorescent spot. STED typically improves resolution to 100 nm in all three axes, but this has compromises with phototoxicity and a lack of compatibility with conventional fluorophores and involves an increase in system cost. A unique hardware addition by ZEISS to improve image resolution involves the use of an AiryScan detector in their confocal microscopes [22]. The AiryScan uses an array of detectors laid out in a honeycomb pattern, which retains the light collection sensitivity of a conventional confocal microscope but enables an increase in resolution of approximately 1.4-fold without modifications to sample preparation. The compromise for this enhanced imaging technique is currently the increased hardware cost, and acquisition and processing time; the latter of which improves with continuous computational advances [22][23].

3. Biological Models for Fluorescence Imaging

3.1. Monolayer Cell Culture

There are over 4000 well-characterised, readily available cell lines serving as models for human disease and development, which can be studied by researchers using a variety of imaging applications. Cell culture systems provide several advantages as models of more complex biological systems, including their potential for high-throughput screening, reproducibility, cost-effectiveness, reduced ethical considerations, well-documented protocols, and their ability to be easily manipulated. For example, cells can be transiently or stably transfected to introduce a gene of interest, enabling visualisation of a given protein using fluorescent protein tags. Cell lines offer an isolated monoculture of a single cell type, or co-culture of multiple cell types, which typically takes the form of a thin adherent monolayer or suspension culture [24][25]. For adherent cells, a uniform monolayer permits improved light penetration for imaging and consistent staining/immunolabelling, without the need to permeate dyes and antibodies into deeper layers of a sample. This reduces sample-sample variability, providing more reproducible imaging results. Moreover, cultured cells exhibit less endogenous fluorescence, which can interfere with label detection, compared to processed tissues and organs [26], and do not require time-consuming tissue processing for imaging. A microscope equipped with an incubation system enables real-time visualisation of live cells [27] and time-lapse imaging [16], with minimal interference to normal cellular function due to environmental disturbance. Microscopy using cell line models enables the exploration of subcellular processes that are implicated in disease pathogenesis, as well as cellular responses to therapeutics [28][29][30]. Whole cell imaging of two-dimensional (2D) cultures allows the examination of cell morphology [31] and intercellular communication networks, mediated by structures such as filopodia or cytonemes [32], tunnelling nanotubes [33], and cellular bridges [34]. Live cell imaging of cell cultures also enables the study of dynamic cell behaviour, such as extracellular vesicle formation, cell motility and migration in wound healing [35], and cancer metastasis [29]. Fluorescence microscopy of cell culture is also widely used to image subcellular components, including organelles and molecules, to provide greater insight into their structure, function, and subcellular localisation. Recent examples of the impressive details of cellular structures visualised by super-resolution microscopy (SRM) include nuclear pore complex organisation, membrane-associated periodic skeleton in neurons and synaptic structures [36]. Multispectral imaging, using confocal and lattice light-microscopy, revealed an intricate spatial-temporal organelle interactome in live cells [37], illustrating the usefulness of novel microscopy techniques to better understand complex cell biology.

3.2. 3D Cell Cultures

The need for animal-free disease models that avoid the limitations of traditional cell culture monolayers, yet better mimic human in vivo environments and recapitulate the complex interactions between different cell types, has resulted in advances in 3D cell culture models. These can represent many tissues, including brain [38], breast [39], and prostate tissue [40], utilising scaffold or scaffold-free techniques to induce or cultivate their formation or maintenance. There are multiple types of 3D cultures, ranging from spheroids of cell lines to organoids derived from patient tissue (reviewed by Caleb and Yong [41]). Indeed, patient-derived organoids are utilised to offer personalised treatments and novel therapeutic discoveries (e.g., [42][43]). Generated from cell lines cultures, spheroids or organoids may preserve the cell phenotypes observed in tissues, which result from interactions and responses to the microenvironment, such as detecting changes to necrotic tissue due to nutrient starvation in the centre of spheroids, mimicking rapidly growing non-angiogenic tumour tissue [44]. An example of the differences between 2D and 3D culture was demonstrated in cardiac cells; cells grown in 2D exhibited a large network of microfilaments and microtubules, whilst cells in a 3D environment were smaller in size, had many junctions between cells, and exhibited increased alpha actinin cytoskeletal protein [45]. Thus, 3D cell cultures may provide significant new insights into more complex cell biology, as well as disease biomarkers and therapeutics to improve translation.

3.3. Tissue Sections

A truer representation of in vivo biology is of course to take it directly from the source: tissue. The major advantage of using tissue sections compared to cell line models is that the complex interactions between a cell with its microenvironment are preserved, including the presence of the supporting extracellular matrix [46], the influence of stroma and immune cells [47], and the maintenance of cell polarity in the hierarchical architecture of the tissue [48]. The ability to collect a snapshot of in vivo biology and preserve this for future study is invaluable, and biobanks with archived tissue exist for a multitude of diseases.
Although imaging of both 3D cell culture and tissue sections are widespread in investigating cell biology, to date, most biological research relies on standard cell culture and ex vivo models that cannot fully recapitulate physiological conditions, sometimes leading to artefacts and inaccurate results [49][50]. Imaging of biological processes in the context of a living organism opens new avenues in biomedical research, allowing visualisation of cellular and molecular associations in real time in their natural environment. Intravital or in vivo fluorescent microscopy has become a quintessential tool in the direct visualisation of the biological processes in living animals, with significant applications in studying alterations in tissue morphology and function [51][52], redox dynamics [53], cell proliferation and differentiation [54][55], cell migratory behaviour [56][57], tumour microenvironment [58][59] reviewed in [60], intracellular ionic activity [61][62], and host–pathogen interactions [63].

3.4. Intravital Imaging

Intravital imaging requires access to a high-resolution confocal, multiphoton, and/or light-sheet microscope, and the development of suitable animal models. Transgenic animals modified to express fluorescently tagged proteins provide a true in vivo environment for evaluating various cellular processes, with constitutive replenishment of fluorescently labelled proteins enabling long-term tracing in a cell- and tissue-specific manner. This technique mainly makes use of small organisms, such as fruit flies [64][65][66], zebrafish [53][67][68], and mice [69][70], due to their short generation times, established genetic lines, and relatively low cost. Subcutaneous and orthotopic xenograft models used in intravital studies can accurately provide insights into tumour heterogeneity and responses to drug treatments [71][72]. With the use of intravital microscopy, it has also become possible to visualise the inter-individual variability at a microscopic level in response to drug treatment. An advantage of intravital imaging is the continual monitoring of physiological changes over days and weeks, which is especially important in developmental cells [55], stem cell [73][74], and tumour biology [75].

4. Imaging Agents Used for Fluorescence Microscopy

4.1. Fluorescent Proteins

The discovery of GFP in 1962 [76], and its subsequent cloning 30 years later [4], has led to FPs becoming an integral tool for cell biologists to monitor cellular processes using fluorescence microscopy. Since that time, a variety of FPs have been synthesised, which traverse the visible spectrum and have been well covered in some excellent reviews [77][78]. Fluorescent proteins are large molecules (typically 25–30 kDa) that exhibit bright fluorescence and excellent bioavailability [77].
Typically, FPs are unable to image acidic environments [79][80]; however, examples do exist where this property has been exploited. For example, Trejo and co-workers used a pH-sensitive GFP (pHluorin-mKate2) to monitor starvation-induced autophagy in LC3B transgenic mice, where the FP was non-fluorescent in acidic compartments, but was emissive in neutral or basic environments [81].
A major disadvantage of FPs is their high background fluorescence, which can make the processing of images difficult. To circumvent this issue, FPs have been designed where a non-fluorescent protein becomes emissive upon activation by a trigger [82].
Simultaneous expression of several FP markers allows multicolour labelling of cellular process, compartments, or proteins of interest to be investigated in parallel within a cell. The “Brainbow” strategy relies on a handful of spectroscopically distinct FPs to provide information of different cellular types and environments, by emitting a broad palette of detectable hues [83]. Hematopoietic stem cells, which give rise to B and T cells, have recently been imaged by transplanting a range of fluorescently labelled cells with differing emission profiles, to provide information on hematopoietic cell lineage in Rag1−/− mice using confocal microscopy [84]. Similarly, live pluripotent stem cells were imaged by confocal microscopy to track their eventual derivatisation using the Brainbow technique (Figure 1) [85]. This approach has recently been supplemented with a secondary near-infra red FP (mCardinal), whose expression is driven independently of the Brainbow cassette [86]. This novel combinatorial system opens the door to further expand the Brainbow and related multi-colour techniques of FPs.
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Figure 1. Live human-induced pluripotent stem cells after 12 h with transfected with eGFP, mOrange2 and mKate2. Scale bar = 20 µm. Reproduced with permission [85]. Copyright 2020 Elsevier.
Despite the considerable effort to construct new FPs with different optical parameters, the field is still largely limited by spectral overlap between their broad fluorescence profiles [87][88]. This characteristic of fluorophores is taken advantage of in applications such as FRET. A recent example of a FP-based FRET system was reported by Shen et al., who created three independent potassium ion (K+) sensors to measure both intra and extracellular K+ [89]; a critical electrolyte for cell function [90]. This was achieved in live HeLa cells using confocal microscopy. The overall design of each system (Figure 2) relied on a “turn-on” effect when two potassium binding proteins encapsulated K+, which brought the FRET donor and acceptor FPs close enough to elicit a FRET response.
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Figure 2. Schematic of the KIRIN1-GR FRET system where emission at a higher wavelength is detected after potassium binding. Image adapted with permission [89]. Copyright 2019 Springer Nature.

4.2. Graphene Quantum Dots

Graphene quantum dots (GQDs) are zero-dimensional fluorescent nanomaterials that have been exploited for a variety of imaging and sensing applications. This category of imaging technology is the most recent of the classes reviewed here; however, their potential for greater use in fluorescence microscopy is significant with a surge of recent examples within the literature. We have focussed on GQDs rather than other varieties of inorganic QDs, which often suffer from solubility and toxicity issues owing to their reliance on incorporated heavy metals such as cadmium, mercury, and lead [91][92]. Conversely, GQDs are water soluble materials which have excellent toxicity profiles [93][94][95]. The photophysical properties of GQDs are amenable to fluorescence microscopy with higher molar absorptivity coefficients, improved photostability and longer-lived emission compared to organic fluorophores [92]. A major advantage of using GQDs is how easily they can be synthesised using either a ‘top-down’ or ‘bottom-up’ approach [96][97][98]. Of these two strategies, the ‘bottom-up’ approach has become the favoured route, given starting material affordability and the simplicity of the method, which does not require specialist chemistry or cellular biology training, unlike other imaging strategies reviewed here.
A drawback of using GQDs is that by virtue of their synthesis, a heterogenous and inseparable mixture of chemical structures is formed, which inherently makes characterisation difficult when using traditional analytical techniques employed for small molecules, such as NMR spectroscopy and mass spectrometry [99]. As a result, techniques such as IR, XPS, and UV-visible spectrophotometry are relied on to characterise the bulk material. This structural ambiguity means that functionalisation of the GQD surface is also more difficult when compared to organic fluorophores, as the exact number (and sometimes type) of reactive handles are unknown and reaction monitoring is almost impossible.
Defining subcellular structures is a common goal in fluorescence microscopy and tools that allow this level of specificity are much sought after. Although this domain is mostly owned by organic fluorophores, GQDs have been utilised to track subcellular components too. For example, GQDs were functionalised with ethylenediamine groups (via an acid chloride route) to generate a probe which selectively stained the nucleolus in live HepG2 cells, as evidenced with co-staining experiments with the commercially available SYTO RNA-Select nucleolus stain [100].

4.3. Metal Ion Complexes

Metal ion complexes have been successfully used in cellular imaging applications such as subcellular compartment staining and visualisation of cellular processes. A variety of sensitising pathways have been utilised for these types of metal complexes, such as metal-to-ligand charge transfer (MLCT) and ligand-to-ligand charge transfer (LLCT), which have been reviewed previously [101]. The major advantage of using metal ion complexes as imaging agents are their long-lived emission profiles, which facilitate the use of time-gated fluorescence microscopy experiments, such as FLIM, and enables the visualisation of cellular events, which is otherwise not possible due to endogenous autofluorescence within cells [102]. Moreover, metal ion complexes offer highly tuneable excitation and emission profiles, which span the entire visible and near-IR spectrum. They also offer the largest Stokes shifts of the four classes covered in this review (typically greater than 5000 cm−1), which obviates self-quenching issues encountered by most organic fluorophores [103].

4.4. Organic Fluorophores

Organic fluorophores enjoy a privileged place as the most prominent class of compounds for fluorescence imaging. Owing to their ready accessibility, small size, wide variety, and excellent emissive properties, organic fluorophores are frequently used for imaging cellular components, visualising cellular processes and tagging larger molecules (including antibodies and drugs) for insights into their cellular activity. The advantages and applications of organic fluorophores have been thoroughly covered in many excellent revstudiews [104][105][106][107][108], and their commercial availability renders their use as “plug and play” for cell biologists.

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