Can a Single Photograph Reveal Everything About an Animal’s Life? Of course not. The same principle applies to cell biology—a beautiful image of a fixed specimen reveals cellular structure but tells us little about the dynamic processes taking place within the cell. This is why live-cell imaging is becoming increasingly important, allowing researchers to observe cells in their natural, living environment. What are the strengths and limitations of each approach? Let’s take a closer look.

Most of us have probably seen a stuffed fox—or another wild animal—in a natural history museum. Standing up close, we can admire every detail of its fur, claws, whiskers, and perhaps even touch its magnificent tail. We might say, “What a beautiful fox.” But almost immediately another thought follows: “It’s a pity it’s stuffed—you can’t learn anything more about it.” At best, we leave with a photograph, since most of us will never have the opportunity to observe a wild fox from such a close distance.

So where can we learn about its behavior, daily activity patterns, whether it lives alone or with a mate, or if it has cubs? Wildlife biologists typically rely on camera traps—motion-activated cameras that capture images or videos whenever an animal passes nearby. These devices minimize human interference, allowing animals to behave naturally in their environment.

While camera traps may not reveal whether a fox has a small spot in its fur or a chipped tooth—especially since most footage is recorded in black and white at night—they provide something far more valuable. They allow researchers to observe natural behavior, monitor population size, and even assess the overall health of wildlife populations.

But what does this discussion about taxidermy and camera traps have to do with fluorescence confocal microscopy?

Imagine that we want to visualize intracellular structures and determine their localization—for example, protein complexes within the cell nucleus. The standard technique used for this purpose is immunofluorescence, which can be compared to taxidermy. The method involves fixing cells with chemical fixatives (most commonly aldehydes) and permeabilizing their membranes using detergents. The prepared sample is then stained with specific antibodies that recognize target proteins or subcellular markers and imaged using a fluorescence confocal microscope. This provides the spatial resolution needed to observe the proteins of interest within the cell. Like a stuffed fox, however, these cells are no longer alive. They represent only a static snapshot in time and can be imaged without any concern for phototoxicity. Under these conditions, we are free to acquire more densely spaced optical sections along the Z-axis, increase laser power, extend exposure times, and adjust the pinhole diameter as needed. In other words, just as when posing for a photograph with a taxidermy fox, we have complete freedom to optimize every imaging parameter in pursuit of the perfect image. What we cannot obtain, however, is any information about the dynamic biological processes that were taking place inside the cell. We have no way of knowing what happened immediately before immunofluorescence staining or what would have happened to the cell afterward. For example, we cannot determine whether the presence of protein complexes in the nucleus is a consequence of a cell division event that occurred just before fixation. Equally, we cannot tell whether their nuclear localization had already initiated one of the cellular pathways leading to programmed cell death.

In this situation, the logical equivalent of a camera trap would be a method that allows us to observe cells under conditions that preserve their homeostasis—namely, live-cell imaging. As the name suggests, live-cell imaging is performed while the cells are still alive and involves monitoring them in real time. The primary tool is, once again, a fluorescence confocal microscope. This time, however, it is equipped with a live-cell incubation chamber that maintains physiological temperature, CO₂ concentration, and humidity throughout the experiment. Instead of conventional immunofluorescence staining, researchers use specialized fluorescent probes that can enter living cells without disrupting membrane integrity, as well as fusion proteins carrying genetically encoded fluorescent tags, such as GFP (Green Fluorescent Protein). The greatest challenge in live-cell imaging is phototoxicity—light-induced cellular stress that can alter cell behavior or even lead to cell death. Returning to our analogy, phototoxicity is like the constant presence of a person with a camera near a fox’s den. A stressed fox may abandon its den altogether or behave unnaturally. Just as camera traps minimize human disturbance and allow researchers to observe authentic animal behavior, live-cell imaging aims to minimize phototoxicity so that cells can be observed under conditions that are as close as possible to their natural physiological state. Achieving long-term, informative imaging often requires compromising image quality. Laser power and exposure times must be kept to a minimum, while much of the signal amplification is performed digitally during post-processing. Highly sensitive detectors are essential for capturing weak fluorescence signals, and the number of optical sections collected along the Z-axis should be reduced as much as possible. Finally, images must be acquired frequently enough to capture biologically meaningful events, yet infrequently enough to avoid repeatedly exposing the cells to damaging laser illumination. The result of such an experiment is not a collection of beautiful, high-resolution static images of a “stuffed fox,” but rather a time-lapse movie that reveals the cell’s “behavior”—the dynamics of intracellular processes. Live-cell imaging enables researchers to answer questions that fixed-cell imaging simply cannot. For example, how does the localization of a particular protein influence cell behavior? Does it affect cell migration? Is its appearance an early indicator of cell death? Does it influence cell division or other dynamic biological processes? These are the kinds of questions that only living cells can answer.

Does this mean that live-cell imaging inevitably produces flat, blurry, and dim images? Not at all. Most of these challenges can be overcome by carefully optimizing the image acquisition parameters. While laser power is certainly important, detector and scanner settings are equally critical for achieving high-quality images without compromising cell viability. Does this sound time-consuming? In most systems equipped with conventional PMT (Photomultiplier Tube) detectors, it often is. Finding the optimal balance between image quality and phototoxicity can require considerable experience and repeated adjustments. Fortunately, the latest generation of confocal microscopy systems offers solutions that dramatically simplify and accelerate this process, making high-quality live-cell imaging more accessible than ever before.

With conventional PMT (Photomultiplier Tube) detectors, users typically need to optimize several acquisition parameters, including the detector high voltage (HV, or gain), gating, and the timing offset between the laser excitation pulse and the detector gate opening (offset). Finding the right combination of these settings can be both time-consuming and technically demanding. By contrast, SilViR silicon detectors from Evident greatly simplify the workflow. In practice, the detectors are simply activated, and the primary parameter requiring adjustment is the laser power. In addition, SilViR detectors are specifically optimized for single-photon detection, enabling the capture of extremely weak fluorescence signals. As a result, laser power can be reduced to an absolute minimum—typically below 1%—significantly lowering phototoxicity while maintaining excellent image quality.

When it comes to the scanner, there are two options available in standard confocal microscopy: the galvanometric scanner (included as standard) and the resonant scanner (available as an option in hybrid or tandem systems). The two key differences between them are scanning speed and signal-to-noise ratio (SNR). In terms of acquisition speed, the resonant scanner is the clear winner—and by a considerable margin. At an image resolution of 512 × 512 pixels, it can acquire images up to 30 times faster than a galvanometric scanner, making it the preferred choice for capturing highly dynamic processes in living cells. However, this speed comes at a cost. The signal-to-noise ratio (SNR) is substantially lower than with a galvanometric scanner, as each pixel receives approximately 10 to 40 times fewer photons. Consequently, images acquired with a resonant scanner may initially appear noisier. One effective way to improve image quality is through signal accumulation. Rather than generating an image from a single scan, the microscope repeatedly scans the same field of view and sums the pixel intensities from multiple frames or scan lines before displaying the final image. This approach significantly improves SNR while preserving the high acquisition speed of the resonant scanner. For example, on the Evident FV5000 confocal microscope, acquiring a Z-stack consisting of 244 optical sections with 4× accumulation takes only 4.6 minutes using the resonant scanner. Performing the same acquisition with a galvanometric scanner would require approximately 43.5 minutes—while delivering comparable image quality. This dramatic reduction in acquisition time makes resonant scanning particularly advantageous for long-term live-cell imaging, where minimizing experiment duration is essential for preserving cell health and capturing dynamic biological events.

Porównanie jakości obrazu uzyskanego za pomocą skanera galwanometrycznego oraz rezonansowego. Obrazowanie wykonane mikroskopem FV5000 (Evident). Skrawek mózgu myszy. Neurony piramidalne piątej warstwy kory mózgowej znakowane EYFP w linii Thy1‑YFP‑H. Zdjęcie udostepnione dzięki uprzejmości: dr Satoshi Fujimoto i dr Takeshi Imai, Graduate School of Medical Sciences, Uniwersytet Kyushu.”

Another solution specifically designed for live-cell imaging is the spinning disk confocal microscope. In simple terms, a spinning disk is a rapidly rotating disk containing thousands of microscopic pinholes. As the disk spins at several thousand revolutions per minute, it illuminates the specimen at thousands of locations simultaneously, rather than scanning it point by point as in a conventional laser-scanning confocal microscope. As a result, the exposure time for each point within the sample is dramatically reduced, and the amount of light energy delivered to the cells is significantly lower. The outcome is substantially reduced phototoxicity together with much faster image acquisition. These advantages make spinning disk confocal microscopy particularly well suited for studying highly dynamic biological processes, including cell division, intracellular vesicle transport, and even the tracking of individual viral particles during the infection of living cells.

Immunofluorescence. HeLa cells. Blue: nuclei; green: tubulin; red: β-catenin. 60× objective. Image acquired using the Evident IXplore SpinSR spinning disk confocal microscope. Image courtesy of Dr. Natalia Derewońko, Helpi Optics Sp. z o.o.

Does this mean that live-cell imaging will eventually replace fixed-sample imaging in confocal microscopy? Absolutely not. The main reasons are the limitations associated with sample labeling and the technical requirements of live-cell experiments. One of the greatest advantages of immunofluorescence is the vast selection of commercially available antibodies. Researchers can label the majority of proteins in human cells and common model organisms, while a wide range of organelle-specific fluorescent probes enables precise visualization of virtually every major cellular compartment. Live-cell imaging, in contrast, is much more restricted in both labeling strategies and genetic engineering approaches. A good example is the cell membrane. One of the most widely used membrane-associated markers is β-catenin, for which highly specific antibodies are available from virtually every antibody supplier. However, β-catenin itself cannot be labeled directly in living cells using conventional live-cell staining approaches. Although several live-cell membrane dyes are available, each comes with important limitations. Some, such as DiI and DiO, gradually become internalized during prolonged imaging, causing the membrane signal to diminish over time. Others, including WGA (wheat germ agglutinin), may induce cytotoxic effects that compromise cell viability during long-term experiments. As mentioned earlier, fixed cells eliminate concerns about phototoxicity altogether. Consequently, researchers can use as many fluorescent labels as the microscope’s laser lines and detectors permit, producing highly detailed, multicolor images. In live-cell imaging, however, the opposite strategy is recommended. To minimize light exposure and preserve normal cellular physiology, labeling should be kept to the absolute minimum. In practice, this usually means two or, at most, three fluorophores, complemented by transmitted-light imaging to visualize cell boundaries and overall morphology. Genetically encoded fluorescent fusion proteins also present important challenges. The fluorescent tag itself—for example, GFP (~27 kDa)—may interfere with the native behavior of the protein of interest. It can alter protein folding, create steric hindrance that disrupts interactions with binding partners, promote protein aggregation, and ultimately impair normal biological function. These factors must always be carefully considered when designing live-cell imaging experiments.

Another important distinction between these two approaches lies in their instrumentation requirements. Just as observing a taxidermy fox requires nothing more than your eyes, imaging fixed samples can be performed using virtually any well-functioning fluorescence confocal microscope. Live-cell imaging, however, requires additional specialized equipment—the microscopy equivalent of a camera trap. This typically includes a live-cell incubation chamber to maintain physiological conditions throughout the experiment, high-sensitivity detectors capable of capturing weak fluorescence signals while minimizing light exposure, or a spinning disk confocal system for imaging rapid intracellular events with reduced phototoxicity. These specialized components are essential for preserving cell viability while capturing dynamic biological processes with sufficient temporal and spatial resolution.

In summary, just as no one would try to understand a fox’s behavior by studying a museum specimen, no one would go into the forest to set up camera traps simply to learn what a fox looks like. The key is to choose the right method based on the scientific question being asked, as well as the available instrumentation and experimental capabilities. If you’d like to find out which solution is the best fit for your laboratory, feel free to get in touch with us! 😊

AUTHOR: dr Natalia Derewońko, Advanced Microscopy Application Specialist

Dr. Natalia Derewońko is a molecular biologist holding a Ph.D. in Biological Sciences, awarded by the Intercollegiate Faculty of Biotechnology of the University of Gdańsk and the Medical University of Gdańsk (UG/GUMed).
She gained her scientific experience in the Laboratory of Molecular Virology, where she conducted research involving BSL-2+ pathogens. Throughout her academic career, she participated in eight competitive research projects funded by the National Science Centre (NCN) and the National Centre for Research and Development (NCBR), led two research projects, and co-authored six peer-reviewed scientific publications.

In subsequent years, she expanded her expertise at the interface of science and industry, supporting scientific sales, delivering training courses in molecular biology, and advising academic and diagnostic laboratories on selecting optimal research solutions.

Currently, she serves as an Advanced Microscopy Application Specialist at Helpi Optics Sp. z o.o., where she supports researchers in selecting advanced imaging systems and experimental methodologies. Her areas of expertise include confocal fluorescence microscopy, cell culture techniques, and the analysis of proteins and nucleic acids.