Live-cell imaging

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A live-cell microscope. Live-cell microscopes are generally inverted. To keep cells alive during observation, the microscopes are commonly enclosed in a micro cell incubator (the transparent box). Olympus FluoView FV1000 Confocal Microscope - NCMIR.jpg
A live-cell microscope. Live-cell microscopes are generally inverted. To keep cells alive during observation, the microscopes are commonly enclosed in a micro cell incubator (the transparent box).

Live-cell imaging is the study of living cells using time-lapse microscopy. It is used by scientists to obtain a better understanding of biological function through the study of cellular dynamics. [1] Live-cell imaging was pioneered in the first decade of the 21st century. One of the first time-lapse microcinematographic films of cells ever made was made by Julius Ries, showing the fertilization and development of the sea urchin egg. [2] Since then, several microscopy methods have been developed to study living cells in greater detail with less effort. A newer type of imaging using quantum dots have been used, as they are shown to be more stable. [3] The development of holotomographic microscopy has disregarded phototoxicity and other staining-derived disadvantages by implementing digital staining based on cells’ refractive index. [4] [5]

Contents

Overview

Biological systems exist as a complex interplay of countless cellular components interacting across four dimensions to produce the phenomenon called life. While it is common to reduce living organisms to non-living samples to accommodate traditional static imaging tools, the further the sample deviates from the native conditions, the more likely the delicate processes in question will exhibit perturbations. [6] The onerous task of capturing the true physiological identity of living tissue, therefore, requires high-resolution visualization across both space and time within the parent organism. [7] The technological advances of live-cell imaging, designed to provide spatiotemporal images of subcellular events in real time, serves an important role for corroborating the biological relevance of physiological changes observed during experimentation. Due to their contiguous relationship with physiological conditions, live-cell assays are considered the standard for probing complex and dynamic cellular events. [8] As dynamic processes such as migration, cell development, and intracellular trafficking increasingly become the focus of biological research, techniques capable of capturing 3-dimensional data in real time for cellular networks ( in situ ) and entire organisms ( in vivo ) will become indispensable tools in understanding biological systems. The general acceptance of live-cell imaging has led to a rapid expansion in the number of practitioners and established a need for increased spatial and temporal resolution without compromising the health of the cell. [9]

Types of microscopy used

Phase contrast

Phase contrast microscopy time-lapse video of dividing rattle grasshopper spermatocytes. This historic film, which popularized phase contrast microscopy, was made in the early 1940s by Kurt Michel of the Carl Zeiss company. [10]

Before the introduction of the phase-contrast microscope, it was difficult to observe living cells. As living cells are translucent, they must be stained to be visible in a traditional light microscope. Unfortunately, the process of staining cells generally kills them. With the invention of the phase-contrast microscopy it became possible to observe unstained living cells in detail. After its introduction in the 1940s, live-cell imaging rapidly became popular using phase-contrast microscopy. [11] The phase-contrast microscope was popularized through a series of time-lapse movies (see video), recorded using a photographic film camera. [12] Its inventor, Frits Zernike, was awarded the Nobel Prize in 1953. [13] Other later phase-contrast techniques used to observe unstained cells are Hoffman modulation and differential interference contrast microscopy.

Fluorescent

Fluorescent microscopy time-lapse video of a dividing purple sea urchin embryo. [14]

Phase-contrast microscopy does not have the capacity to observe specific proteins or other organic chemical compounds which form the complex machinery of a cell. Synthetic and organic fluorescent stains have therefore been developed to label such compounds, making them observable by fluorescent microscopy (see video). [15] Fluorescent stains are, however, phototoxic, invasive and bleach when observed. This limits their use when observing living cells over extended periods of time. Non-invasive phase-contrast techniques are therefore often used as a vital complement to fluorescent microscopy in live-cell imaging applications. [16] [17] Deep learning-assisted fluorescence microscopy methods, however, help to reduced light burden and phototoxicity and allow even repeated high resolution live imaging. [18]

Quantitative phase contrast

Quantitative phase contrast microscopy video of a dividing breast cancer cells. [19]

As a result of the rapid increase in pixel density of digital image sensors, quantitative phase-contrast microscopy has emerged as an alternative microscopy method for live-cell imaging. [20] [21] Quantitative phase-contrast microscopy has an advantage over fluorescent and phase-contrast microscopy in that it is both non-invasive and quantitative in its nature.

Due to the narrow focal depth of conventional microscopy, live-cell imaging is to a large extent currently limited to observing cells on a single plane. Most implementations of quantitative phase-contrast microscopy allow creating and focusing images at different focal planes from a single exposure. This opens up the future possibility of 3-dimensional live-cell imaging by means of fluorescence techniques. [22] Quantitative phase-contrast microscopy with rotational scanning allow 3D time-lapse images of living cells to be acquired at high resolution. [23] [24] [4]

Holotomography

Holotomography (HT) is a laser technique to measure three-dimensional refractive index (RI) tomogram of a microscopic sample such as biological cells and tissues. Because the RI can serve as an intrinsic imaging contrast for transparent or phase objects, measurements of RI tomograms can provide label-free quantitative imaging of microscopic phase objects. In order to measure 3D RI tomogram of samples, HT employs the principle of holographic imaging and inverse scattering. Typically, multiple 2D holographic images of a sample are measured at various illumination angles, employing the principle of interferometric imaging. Then, a 3D RI tomogram of the sample is reconstructed from these multiple 2D holographic images by inversely solving light scattering in the sample.

The principle of HT is very similar to X-ray computed tomography (CT), or CT scan. CT scan measures multiple 2D X-ray images of a human body at various illumination angles, and a 3D tomogram (X-ray absorptivity) is then retrieved using the inverse scattering theory. Both the X-ray CT and laser HT shares the same governing equation – Helmholtz equation, the wave equation for a monochromatic wavelength. HT is also known as optical diffraction tomography.

The combination of holography and rotational scanning allows long-term, label-free, live-cell recordings.

Non-invasive optical nanoscopy can achieve such a lateral resolution by using a quasi-2π-holographic detection scheme and complex deconvolution. The spatial frequencies of the imaged cell do not make any sense to the human eye. But these scattered frequencies are converted into a hologram and synthesize a bandpass, which has a resolution double the one normally available. Holograms are recorded from different illumination directions on the sample plane and observe sub-wavelength tomographic variations of the specimen. Nanoscale apertures serve to calibrate the tomographic reconstruction and to characterize the imaging system by means of the coherent transfer function. This gives rise to realistic inverse filtering and guarantees true complex field reconstruction. [24]

In conclusion, the 2 terminologies of (i) optical resolution (the real one) and (ii) sampling resolution (the one on the screen) are separated for 3D holotomographic microscopy.

Instrumentation and optics

Live-cell imaging represents a careful compromise between acquiring the highest-resolution image and keeping the cells alive for as long as possible. [25] As a result, live-cell microscopists face a unique set of challenges that are often overlooked when working with fixed specimens. Moreover, live-cell imaging often employs special optical system and detector specifications. For example, ideally the microscopes used in live-cell imaging would have high signal-to-noise ratios, fast image acquisition rates to capture time-lapse video of extracellular events, and maintaining the long-term viability of the cells. [26] However, optimizing even a single facet of image acquisition can be resource-intensive and should be considered on a case-by-case basis.

Lens designs

A) Upright lens configuration. B) Inverted lens configuration. Lens Configuration2.png
A) Upright lens configuration. B) Inverted lens configuration.

Low-magnification "dry"

In cases where extra space between the objective and the specimen is required to work with the sample, a dry lens can be used, potentially requiring additional adjustments of the correction collar, which changes the location of the lens in the objective, to account for differences in imaging chambers. Special objective lenses are designed with correction collars that correct for spherical aberrations while accounting for the cover-slip thickness. In high-numerical-aperture (NA) dry objective lenses, the correction collar adjustment ring changes the position of a movable lens group to account for differences in the way the outside of the lens focuses light relative to the center. Although lens aberrations are inherent in all lens designs, they become more problematic in dry lenses, where resolution retention is key. [27]

Oil-immersion high-NA

Oil immersion is a technique that can increase image resolution by immersing the lens and the specimen in oil with a high refractive index. Since light bends when it passes between media with different refractive indexes, by placing oil with the same refractive index as glass between the lens and the slide, two transitions between refractive indices can be avoided. [28] However, for most applications it is recommended that oil immersion be used with fixed (dead) specimens because live cells require an aqueous environment, and the mixing of oil and water can cause severe spherical aberrations. For some applications silicone oil can be used to produce more accurate image reconstructions. Silicone oil is an attractive medium because it has a refractive index that is close to that of living cells, allowing to produce high-resolution images while minimizing spherical aberrations. [27]

Water-immersion

Live-cell imaging requires a sample in an aqueous environment that is often 50 to 200 micrometers away from the cover glass. Therefore, water-immersion lenses can help achieve a higher resolving power due to the fact that both the environment and the cells themselves will be close to the refractive index of water. Water-immersion lenses are designed to be compatible with the refractive index of water and usually have a corrective collar that allows adjustment of the objective. Additionally, because of the higher refractive index of water, water-immersion lenses have a high numerical aperture and can produce images superior to oil-immersion lens when resolving planes deeper than 0 µm. [27]

Dipping

Another solution for live-cell imaging is the dipping lens. These lenses are a subset of water-immersion lenses that do not require a cover slip and can be dipped directly into the aqueous environment of the sample. One of the main advantages of the dipping lens is that it has a long effective working distance. [29] Since a cover slip is not required, this type of lens can approach the surface of the specimen, and as a result, the resolution is limited by the restraints imposed by spherical aberration rather than the physical limitations of the cover slip. Although dipping lenses can be very useful, they are not ideal for all experiments, since the act of "dipping" the lens can disturb the cells in the sample. Additionally, since the incubation chamber must be open to the lens, changes in the sample environment due to evaporation must be closely monitored. [27]

Phototoxicity and photobleaching

Today, most live imaging techniques rely on either high-illumination regimes or fluorescent labelling, both inducing phototoxicity and compromising the ability to keep cells unperturbed and alive over time. Since our knowledge of biology is driven by observation, it is key to minimize the perturbations induced by the imaging technique.

The rise of confocal microscopy is closely correlated with accessibility of high-power lasers, which are able to achieve high intensities of light excitation. However, the high-power output can damage sensitive fluorophores, so the lasers usually run significantly below their full power output. [30] Overexposure to light can result in photodamage due to photobleaching or phototoxicity. The effects of photobleaching can significantly reduce the quality of fluorescent images, and in recent years there has been a significant demand for longer-lasting commercial fluorophores. One solution, the Alexa Fluor series, show little to no fading even at high laser intensities. [31]

Under physiological conditions, many cells and tissue types are exposed to only low levels of light. [32] As a result, it is important to minimize the exposure of live cells to high doses of ultraviolet (UV), infrared (IR), or fluorescence exciting wavelengths of light, which can damage DNA, raise cellular temperatures, and cause photobleaching respectively. [33] High-energy photons absorbed by the fluorophores and the sample are emitted at longer wavelengths proportional to the Stokes shift. [34] However, cellular organelles can be damaged when the photon energy produces chemical and molecular changes rather than being re-emitted. [35] It is believed that the primary culprit in the light-induced toxicity experienced by live cells is a result of free radicals produced by the excitation of fluorescent molecules. [32] These free radicals are highly reactive and cause the destruction of cellular components, which can result in non-physiological behavior.

One method of minimizing photo-damage is to lower the oxygen concentration in the sample to avoid the formation of reactive oxygen species. [36] However, this method is not always possible in live-cell imaging and may require additional intervention. Another method for reducing the effects of free radicals in the sample is the use of antifade reagents. Unfortunately, most commercial antifade reagents cannot be used in live-cell imaging because of their toxicity. [37] Instead, natural free-radical scavengers such as vitamin C or vitamin E can be used without substantially altering physiological behavior on shorter time scales. [38] Phototoxicity-free live-cell imaging has recently been developed and commercialised. Holotomographic microscopy avoids phototoxicity thanks to its low-power laser (laser class 1: 0.2 mW/mm2). [4] [5] [39]

See also

Related Research Articles

<span class="mw-page-title-main">Microscopy</span> Viewing of objects which are too small to be seen with the naked eye

Microscopy is the technical field of using microscopes to view objects and areas of objects that cannot be seen with the naked eye. There are three well-known branches of microscopy: optical, electron, and scanning probe microscopy, along with the emerging field of X-ray microscopy.

<span class="mw-page-title-main">Microscope</span> Scientific instrument

A microscope is a laboratory instrument used to examine objects that are too small to be seen by the naked eye. Microscopy is the science of investigating small objects and structures using a microscope. Microscopic means being invisible to the eye unless aided by a microscope.

<span class="mw-page-title-main">Optical microscope</span> Microscope that uses visible light

The optical microscope, also referred to as a light microscope, is a type of microscope that commonly uses visible light and a system of lenses to generate magnified images of small objects. Optical microscopes are the oldest design of microscope and were possibly invented in their present compound form in the 17th century. Basic optical microscopes can be very simple, although many complex designs aim to improve resolution and sample contrast.

<span class="mw-page-title-main">X-ray microscope</span> Type of microscope that uses X-rays

An X-ray microscope uses electromagnetic radiation in the X-ray band to produce magnified images of objects. Since X-rays penetrate most objects, there is no need to specially prepare them for X-ray microscopy observations.

A total internal reflection fluorescence microscope (TIRFM) is a type of microscope with which a thin region of a specimen, usually less than 200 nanometers can be observed.

<span class="mw-page-title-main">Fluorescence microscope</span> Optical microscope that uses fluorescence and phosphorescence

A fluorescence microscope is an optical microscope that uses fluorescence instead of, or in addition to, scattering, reflection, and attenuation or absorption, to study the properties of organic or inorganic substances. "Fluorescence microscope" refers to any microscope that uses fluorescence to generate an image, whether it is a simple set up like an epifluorescence microscope or a more complicated design such as a confocal microscope, which uses optical sectioning to get better resolution of the fluorescence image.

<span class="mw-page-title-main">Confocal microscopy</span> Optical imaging technique

Confocal microscopy, most frequently confocal laser scanning microscopy (CLSM) or laser scanning confocal microscopy (LSCM), is an optical imaging technique for increasing optical resolution and contrast of a micrograph by means of using a spatial pinhole to block out-of-focus light in image formation. Capturing multiple two-dimensional images at different depths in a sample enables the reconstruction of three-dimensional structures within an object. This technique is used extensively in the scientific and industrial communities and typical applications are in life sciences, semiconductor inspection and materials science.

<span class="mw-page-title-main">Two-photon excitation microscopy</span>

Two-photon excitation microscopy is a fluorescence imaging technique that is particularly well-suited to image scattering living tissue of up to about one millimeter in thickness. Unlike traditional fluorescence microscopy, where the excitation wavelength is shorter than the emission wavelength, two-photon excitation requires simultaneous excitation by two photons with longer wavelength than the emitted light. The laser is focused onto a specific location in the tissue and scanned across the sample to sequentially produce the image. Due to the non-linearity of two-photon excitation, mainly fluorophores in the micrometer-sized focus of the laser beam are excited, which results in the spatial resolution of the image. This contrasts with confocal microscopy, where the spatial resolution is produced by the interaction of excitation focus and the confined detection with a pinhole.

<span class="mw-page-title-main">Autofluorescence</span>

Autofluorescence is the natural emission of light by biological structures such as mitochondria and lysosomes when they have absorbed light, and is used to distinguish the light originating from artificially added fluorescent markers (fluorophores).

In fluorescence microscopy, colocalization refers to observation of the spatial overlap between two different fluorescent labels, each having a separate emission wavelength, to see if the different "targets" are located in the same area of the cell or very near to one another. The definition can be split into two different phenomena, co-occurrence, which refers to the presence of two fluorophores in the same pixel, and correlation, a much more significant statistical relationship between the fluorophores indicative of a biological interaction. This technique is important to many cell biological and physiological studies during the demonstration of a relationship between pairs of bio-molecules.

<span class="mw-page-title-main">STED microscopy</span>

Stimulated emission depletion (STED) microscopy is one of the techniques that make up super-resolution microscopy. It creates super-resolution images by the selective deactivation of fluorophores, minimizing the area of illumination at the focal point, and thus enhancing the achievable resolution for a given system. It was developed by Stefan W. Hell and Jan Wichmann in 1994, and was first experimentally demonstrated by Hell and Thomas Klar in 1999. Hell was awarded the Nobel Prize in Chemistry in 2014 for its development. In 1986, V.A. Okhonin had patented the STED idea. This patent was unknown to Hell and Wichmann in 1994.

High-content screening (HCS), also known as high-content analysis (HCA) or cellomics, is a method that is used in biological research and drug discovery to identify substances such as small molecules, peptides, or RNAi that alter the phenotype of a cell in a desired manner. Hence high content screening is a type of phenotypic screen conducted in cells involving the analysis of whole cells or components of cells with simultaneous readout of several parameters. HCS is related to high-throughput screening (HTS), in which thousands of compounds are tested in parallel for their activity in one or more biological assays, but involves assays of more complex cellular phenotypes as outputs. Phenotypic changes may include increases or decreases in the production of cellular products such as proteins and/or changes in the morphology of the cell. Hence HCA typically involves automated microscopy and image analysis. Unlike high-content analysis, high-content screening implies a level of throughput which is why the term "screening" differentiates HCS from HCA, which may be high in content but low in throughput.

<span class="mw-page-title-main">Optical sectioning</span> Imaging of focal planes within a thick sample

Optical sectioning is the process by which a suitably designed microscope can produce clear images of focal planes deep within a thick sample. This is used to reduce the need for thin sectioning using instruments such as the microtome. Many different techniques for optical sectioning are used and several microscopy techniques are specifically designed to improve the quality of optical sectioning.

Super-resolution microscopy is a series of techniques in optical microscopy that allow such images to have resolutions higher than those imposed by the diffraction limit, which is due to the diffraction of light. Super-resolution imaging techniques rely on the near-field or on the far-field. Among techniques that rely on the latter are those that improve the resolution only modestly beyond the diffraction-limit, such as confocal microscopy with closed pinhole or aided by computational methods such as deconvolution or detector-based pixel reassignment, the 4Pi microscope, and structured-illumination microscopy technologies such as SIM and SMI.

<span class="mw-page-title-main">Multifocal plane microscopy</span>

Multifocal plane microscopy (MUM) or multiplane microscopy or multifocus microscopy is a form of light microscopy that allows the tracking of the 3D dynamics in live cells at high temporal and spatial resolution by simultaneously imaging different focal planes within the specimen. In this methodology, the light collected from the sample by an infinity-corrected objective lens is split into two paths. In each path the split light is focused onto a detector which is placed at a specific calibrated distance from the tube lens. In this way, each detector images a distinct plane within the sample. The first developed MUM setup was capable of imaging two distinct planes within the sample. However, the setup can be modified to image more than two planes by further splitting the light in each light path and focusing it onto detectors placed at specific calibrated distances. It has later been improved for imaging up to four distinct planes. To image a greater number of focal planes, simpler techniques based on image splitting optics have been developed. One example is by using a customized image splitting prism, which is capable of capturing up to 8 focal planes using only two cameras. Better yet, standard off-the-shelf partial beamsplitters can be used to construct a so-called z-splitter prism that allows simultaneous imaging of 9 individual focal planes using a single camera. Another technique called multifocus microscopy (MFM) uses diffractive Fourier optics to image up to 25 focal planes.

Photo-activated localization microscopy and stochastic optical reconstruction microscopy (STORM) are widefield fluorescence microscopy imaging methods that allow obtaining images with a resolution beyond the diffraction limit. The methods were proposed in 2006 in the wake of a general emergence of optical super-resolution microscopy methods, and were featured as Methods of the Year for 2008 by the Nature Methods journal. The development of PALM as a targeted biophysical imaging method was largely prompted by the discovery of new species and the engineering of mutants of fluorescent proteins displaying a controllable photochromism, such as photo-activatible GFP. However, the concomitant development of STORM, sharing the same fundamental principle, originally made use of paired cyanine dyes. One molecule of the pair, when excited near its absorption maximum, serves to reactivate the other molecule to the fluorescent state.

<span class="mw-page-title-main">Light sheet fluorescence microscopy</span> Fluorescence microscopy technique

Light sheet fluorescence microscopy (LSFM) is a fluorescence microscopy technique with an intermediate-to-high optical resolution, but good optical sectioning capabilities and high speed. In contrast to epifluorescence microscopy only a thin slice of the sample is illuminated perpendicularly to the direction of observation. For illumination, a laser light-sheet is used, i.e. a laser beam which is focused only in one direction. A second method uses a circular beam scanned in one direction to create the lightsheet. As only the actually observed section is illuminated, this method reduces the photodamage and stress induced on a living sample. Also the good optical sectioning capability reduces the background signal and thus creates images with higher contrast, comparable to confocal microscopy. Because light sheet fluorescence microscopy scans samples by using a plane of light instead of a point, it can acquire images at speeds 100 to 1,000 times faster than those offered by point-scanning methods.

Lattice light-sheet microscopy is a modified version of light sheet fluorescence microscopy that increases image acquisition speed while decreasing damage to cells caused by phototoxicity. This is achieved by using a structured light sheet to excite fluorescence in successive planes of a specimen, generating a time series of 3D images which can provide information about dynamic biological processes.

Holotomography (HT) is a laser technique to measure the three-dimensional refractive index (RI) tomogram of a microscopic sample such as biological cells and tissues. Because the RI can serve as an intrinsic imaging contrast for transparent or phase objects, measurements of RI tomograms can provide label-free quantitative imaging of microscopic phase objects. In order to measure 3-D RI tomogram of samples, HT employs the principle of holographic imaging and inverse scattering. Typically, multiple 2D holographic images of a sample are measured at various illumination angles, employing the principle of interferometric imaging. Then, a 3D RI tomogram of the sample is reconstructed from these multiple 2D holographic images by inversely solving light scattering in the sample.

Super-resolution dipole orientation mapping (SDOM) is a form of fluorescence polarization microscopy (FPM) that achieved super resolution through polarization demodulation. It was first described by Karl Zhanghao and others in 2016. Fluorescence polarization (FP) is related to the dipole orientation of chromophores, making fluorescence polarization microscopy possible to reveal structures and functions of tagged cellular organelles and biological macromolecules. In addition to fluorescence intensity, wavelength, and lifetime, the fourth dimension of fluorescence—polarization—can also provide intensity modulation without the restriction to specific fluorophores; its investigation in super-resolution microscopy is still in its infancy.

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