"if a picture is worth a thousand words, then a movie may be worth a million words." - Anonymous
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 dynamics1. Live-cell imaging has offered countless insights into biological marvels. It is the study of living cell/s with time-lapse microscopy. It helped scientists to achieve a better understanding of biological function through the study of cellular dynamics.
Live‐cell microscopy usually involves conciliation between obtaining image quality and maintaining healthy cells. Continual advances in imaging techniques and the design of fluorescent probes improve the power of this approach, ensuring that live‐cell imaging will continue to be an important tool in biology. To begin with, the milestone historical events starting from the first attempt in 1907 by Julius Ries, the article also explain the different types of Microscopy in live-cell imaging.
According to Daniel Gottschling in his book “Epigenetics” (2007) "based on static images of chromatin and the refractory nature of silent chromatin, I was convinced that once established, a heterochromatic state was solid as granite." It is indeed easy to forget that chromatin - exhaustively analyzed biochemically, histologically detailed for over a century that appears in such characteristic form in electron and light microscopy - is always moving and undergoing constant change”2.
"What is true in a cell at one moment in time may not be true at another moment under different conditions."
The first attempt to Live-cell imaging
The first time-lapse films of embryonic development and cultured cells were made in France in 1907 by Julius Ries (Fig. 1). The films were made of the fertilization and development of the sea urchin egg as a technique of teaching medical students cell theory. Julius Ries assumed that students would never believe that all cells came from other cells and that organisms were made up of cells unless they had a moving, living proof. The entire development process took about 14 hours. Ries converted the 14 hours long time-consuming film into two minutes on the screen. As per Ries's opinion, Drawings and fixed sections were inadequate, exactly because they "differed from the living in their motionlessness"3.
Figure 1 | Stills from Fertilization and Development of the Sea Urchin Egg by Julius Ries.
Commercialization of Microcinematography
After the introduction of microcinematography, researchers quickly realized that the technique is a more convenient option to demonstrate known phenomena. They were now able to see things that were not visible in any other way, as the movement of many cells was so slow as to be beneath the threshold of human perception. The first purpose-built micro-cinematographic devices became commercially available in Europe in 1914. Time-lapse microcinematography has been used to study cellular behavior in vitro ever since the early work of Warren Lewis, demonstrating pinocytosis as a basic cellular phenomenon and the cellular movement studies of Michael Abercrombie that followed4.
The practice was revived with the development of phase contrast microscopy coupled with a film camera and later with video technology in 1930. Phase-contrast allowed the observers into the cell: intracellular organelles and their previously untraceable movements could now be seen in much greater detail.
Criticism makes it a more powerful tool
The visualization efforts in microcinematography were clearly directed at making structures more elucidated, e.g. to observe the behavior of chromatin or mitochondria over time and their reaction to injury or pharmacological agents. However, dynamic modes of imaging were in their turn critiqued as inaccurate and non-scientific, precisely because of their qualitative nature. In the 1940s, Nobelist Peter Medawar wrote in his memoir, scolding film-making cell biologists for having been "delighted, distracted, and beguiled by the sheer beauty" of cells on film, and as a result, having missed the opportunity to use cytological methods to - as he put it - "solve biological problems"5.
Introduction of mathematical logic into Biology
New methods and technologies are introduced for many disparate reasons, but one of them is the craving to make biology a more exact science, to create laws or statistical tools such as are found in other sciences. For instance, J.H. Woodger, attempts to construct theoretical biology in the late 1930s by the introduction of mathematical logics into biology. This was an attempt to assign logical tools to biological knowledge to produce self-explanatory theories. This he pursued because "an intense interest in, and intimate first-hand acquaintance with organisms, indispensable as it is, will not alone lead biology to the goal of exact science"6. Although Woodger's notation system did not persist. His drive to formalize qualitative observation was prominent in the early years of mathematical and theoretical biology.
Labeling of cell molecules
In the 1950s, labeling of molecules with radioisotopes and then visualizing their passage through cells, tissues, and ecosystems was introduced. It was a complete new form of perception because it permitted temporal visualization of the molecular objects enumerated and fixed by x-ray crystallography, the ultracentrifuge, and the electron microscope.
With continuous technical innovation in the addition of fluorescent probes into living cells and the microscopy (to visualize their appearance and movement), the escalation of live cell imaging in the past decades has been spectacular. Roger Tsien, recognized along with Martin Chalfie and Osamu Shimomura with the Nobel prize in chemistry for founding work in fluorescent labeling, stated a feeling that precisely echoed the spirit of microcinematographers's critiques of histology a century ago: "genome sequences alone lack spatial and temporal information and are therefore as dynamic and informative as census lists or telephone directories"7.
The tension between the still and the moving image has been, and will no doubt remain, a highly productive force in the generation of new scientific knowledge.
LIVE‐CELL IMAGING-TECHNIQUES AND MICROSCOPY
Factors that matter most
The sensitivity of the detector (signal‐to‐noise), specimen viability, and the speed required for image acquisition are the main three factors that matter most when choosing an optical microscopy system for live-cell imaging.
Currently, there is no all‐purpose live‐cell imaging system that is suitable for all possible examinations. Researchers must therefore confront this by determining the optimal parameters while reducing cell damage or death. They also need to understand the pros and cons of different microscopes. Most cellular processes occur in three dimensions over time. Therefore, cells need to be imaged in four dimensions to obtain a complete picture. Most epi‐illumination microscopes and confocal systems acquire data series in four dimensions.
Widefield microscopy (WFM)
A microscope technique where the entire sample is exposed to light is known as 'widefield' imaging. This is the simplest, least expensive, and oldest imaging modality used for live-cell imaging. The advantage of the WFM is that it requires the lowest photon dose, particularly for transmitted light imaging, i.e., phase or differential interference contrast (DIC). While fluorescence has largely replaced transmitted light imaging, these techniques are still very useful in answering biological questions. Widefield microscopes are brilliant for generating 2D images of samples as the entire field can be captured at once (Fig. 3). With fast temporal resolution, the amount and localization of specific fluorescent molecules can be seen. Processes like neural signaling in live cells can be measured in real-time. The low photon dose of this modality allows multi-dimensional data to be collected while still maintaining happy and healthy cells.
Alternatively, developments in label-free image analysis techniques have enabled long-term cell culture imaging. These non-invasive techniques can be used to process videos obtained using bright-field microscopes. Such methods require sophisticated algorithms, but are not hampered by phototoxicity effects inherent to fluorescent microscopy.
Looking for an automated live cell microscope? The CytoSMART Omni allows for multi-well imaging for hours up to weeks at a time.
Confocal laser scanning microscopy (CLSM)
CLSM is the mainstay of the imaging lab; however, it is less than ideal for the cell's health and normal functioning. It syndicates high-resolution optical imaging with depth selectivity which permits to do optical sectioning. With the help of CLSM, visual sections of minute structures that would be difficult to physically section (e.g. embryos) and construct 3-D structures from the acquired images can be viewed. In CLSM, a laser beam is passed through a light source aperture which is then focused by an objective lens into a small area of the sample, and an image is built up pixel-by-pixel by collecting the emitted photons from the fluorophores in the sample.
Two-photon or multi-photon microscopy (MPM)
Multiphoton microscopy (MPM) can be used for imaging of living, intact biological tissues on length scales from the molecular level through the whole organism. Since the fluorescence is only generated within the imaging region, there is no need for descanning the image (Fig. 2). For thick specimens, such as in vivo imaging or where UV excitation is required (uncaging, excitation, etc.), MPM is an excellent and often only choice. This is especially true for live-organism imaging. Additionally, with MPM, it is possible to image label-free samples such as collagen fibers or other non-centrosymmetric samples with second harmonic generated microscopy.
Figure 2 | Distribution of fluorescein fluorescence (green) as viewed in the x-z plane, on inverted microscope, focused with 20 X objective lens 0.75NA. (left) Wide-field microscopy (WFM) mercury source and filter cube excitation. (middle) Confocal scanning microscopy (CLSM), 488 nm laser excitation, and (right) two-photon (2P) excitation using femtosecond pulses of 850-nm light.
Spinning disk confocal microscopy (SDCM)
There are some implementations of the confocal principle where a pinhole rejects the out-of-focus light but maintains the ability to generate wide-field images, thereby overcoming the speed limitations and phototoxicity of CLSM. These include slit scanning and pinhole multiplexing methods, including swept-field and spinning disk confocal. The most commonly used is SDCM. This technique uses a pair of rotating disks with thousands of pinholes in a spiral. The Yokogawa scanner adds micro-lenses to the second disk, which increases the light efficiency of the system 8. Because of the high scan speed (up to 360 frames per second) and the parallel collection of images with a high-QE CCD camera, there is a dramatic (10–15-fold) decreased photo-bleaching and phototoxicity compared with point scanning9. While it does not have as low a photon dose as WFM, it does offer the advantage of confocality at a reasonable photon dose, thereby making it well suited to high magnification (due to single pinhole size) live-cell imaging, especially in 4D.
Light-sheet fluorescence microscopy (LSFM)
In this technique the specimen is illuminated with a thin sheet of light from the side of the specimen, thereby illuminating a single XY plane with fluorescence detection occurring in the direction perpendicular to the excitation. Speed is greatly increased relative to point-scanning methods, as well as the entire field of view being captured in parallel on high QE, a low noise CCD camera. The excitation is confined to the focal plane without the use of a pinhole, thereby increasing the light throughput and reducing the photon dose. Additionally, the spatial resolution is lower than other techniques; therefore most studies have been of whole embryos or tissue slices rather than at the single-cell level. LSFM has allowed remarkable observations of vertebrate embryonic development due to its speed and sensitivity as well as having the ability to image over long time intervals10.
At the bleeding edge
In the past two decades or so, some advanced techniques have been introduced in live-cell imaging, such as fluorescence resonance energy transfer (FRET), fluorescence correlation spectroscopy (FCS), fluorescence recovery after photobleaching (FRAP), fluorescence-lifetime imaging (FLIM), and others. Nanoscopy will allow new areas of research at detection levels previously unreachable and frankly unthinkable a decade ago.
Improvements have not only been made to improve spatial resolution. Automation of microscopes and in image analysis opens possibilities for high-throughput microscopic methods (see the CytoSMART Omni for automated live cell microscopy). Parallel analysis of samples in full multi-well plates is now a possibility and can replace labor intensive microscopy work in screening experiments.