Microscopy for Cell Biology: How to Visualize Cells

Light Microscopy Basics

Light microscopy uses visible light and glass lenses to magnify specimens. The compound light microscope, the workhorse of biology laboratories, passes light through a thin specimen and through a series of lenses (objective and ocular) to produce a magnified image. The theoretical resolution limit of light microscopy is approximately 200 nanometers (0.2 micrometers), determined by the wavelength of visible light and the numerical aperture of the lens system. This means that structures separated by less than 200 nanometers appear as a single blurred object. In practical terms, light microscopy can resolve individual cells, nuclei, mitochondria, and other large organelles, but cannot resolve individual ribosomes, membrane proteins, or the fine details of chromatin structure.

Brightfield microscopy is the simplest form of light microscopy. Light passes directly through the specimen, and contrast is generated by the absorption of light by the sample. Most cells and tissues are nearly transparent and produce very little contrast in brightfield without staining. Histological stains such as hematoxylin and eosin (H&E) are used routinely in medical pathology: hematoxylin stains nucleic acids blue-purple, while eosin stains cytoplasmic proteins pink. Other common stains include the Gram stain for classifying bacteria, the Wright-Giemsa stain for blood cells, and periodic acid-Schiff (PAS) for carbohydrates and glycogen.

Phase Contrast and Differential Interference Contrast

Phase contrast microscopy, invented by Frits Zernike in the 1930s (earning him the Nobel Prize in Physics in 1953), converts differences in the refractive index of cellular components into visible contrast without the need for staining. This makes it invaluable for observing living cells, which cannot be stained without being killed or significantly altered. Phase contrast works by separating the light that passes through the specimen into two beams and recombining them so that differences in optical path length appear as differences in brightness. Under phase contrast, cell nuclei, nucleoli, mitochondria, and other organelles appear as dark or bright structures against a gray background.

Differential interference contrast (DIC, also called Nomarski microscopy) uses polarized light and prism optics to generate a pseudo-three-dimensional image with excellent contrast and no halo artifacts that can occur in phase contrast. DIC produces images with a characteristic shadow-cast appearance that makes cell edges and internal structures stand out clearly. Both phase contrast and DIC are essential for routine examination of living cell cultures, monitoring cell health and morphology, and observing dynamic processes such as cell division, migration, and phagocytosis in real time.

How to Prepare and Image Cells

Fluorescence Microscopy

Fluorescence microscopy has revolutionized cell biology by enabling researchers to visualize specific molecules within cells with extraordinary specificity. The technique exploits fluorescent molecules (fluorophores) that absorb light at one wavelength (excitation) and emit light at a longer wavelength (emission). By using appropriate filter sets that separate excitation light from emission light, the microscope produces images in which only the fluorescently labeled structures are visible against a dark background. Multiple fluorophores with different excitation and emission spectra can be used simultaneously to visualize two, three, or even four different targets in the same cell, revealing their spatial relationships.

Green fluorescent protein (GFP), originally isolated from the jellyfish Aequorea victoria and developed as a research tool by Osamu Shimomura, Martin Chalfie, and Roger Tsien (who shared the 2008 Nobel Prize in Chemistry for this work), transformed cell biology by enabling researchers to tag any protein with a fluorescent marker in living cells. By fusing the GFP gene to the gene encoding a protein of interest, researchers can track that protein location, movement, and dynamics in real time without killing the cell or adding external reagents. An expanding palette of fluorescent proteins in colors from blue to far-red now allows simultaneous imaging of multiple proteins in the same living cell.

Confocal and Super-Resolution Microscopy

Confocal laser scanning microscopy overcomes a major limitation of conventional fluorescence microscopy: the blurring caused by out-of-focus fluorescence from above and below the focal plane. A confocal microscope uses a pinhole aperture to reject out-of-focus light, collecting fluorescence only from a thin optical section (typically 0.5 to 1.5 micrometers thick) at the focal plane. By acquiring a series of optical sections at different depths through the specimen (a z-stack), the confocal microscope can reconstruct a three-dimensional image of the cell or tissue with clarity that is impossible with a conventional fluorescence microscope.

Super-resolution microscopy techniques break the 200-nanometer diffraction limit of conventional light microscopy, achieving resolutions of 20 to 100 nanometers. Structured illumination microscopy (SIM) improves resolution to roughly 100 nanometers by illuminating the sample with patterned light and computationally extracting high-frequency information. Stimulated emission depletion (STED) microscopy uses a depletion laser to narrow the effective fluorescence spot to roughly 30 to 50 nanometers. Single-molecule localization methods such as PALM (photoactivated localization microscopy) and STORM (stochastic optical reconstruction microscopy) achieve resolutions as fine as 20 nanometers by precisely determining the positions of individual fluorescent molecules over thousands of image frames. Eric Betzig, Stefan Hell, and William Moerner shared the 2014 Nobel Prize in Chemistry for developing these super-resolution techniques.

Electron Microscopy

Electron microscopy uses beams of electrons rather than visible light to image specimens, achieving resolutions far beyond the capability of any light microscope. The wavelength of electrons is roughly 100,000 times shorter than that of visible light, enabling theoretical resolutions below 0.1 nanometers, though practical resolution for biological specimens is typically 1 to 2 nanometers for transmission electron microscopy (TEM) and 5 to 10 nanometers for scanning electron microscopy (SEM).

Transmission electron microscopy passes electrons through an ultrathin section of the specimen (typically 50 to 100 nanometers thick), producing a two-dimensional image that reveals internal ultrastructure at extraordinary detail. TEM has been essential for characterizing the fine structure of organelles, the arrangement of cytoskeletal filaments, the architecture of the nuclear pore complex, and the structure of viruses within infected cells. Scanning electron microscopy scans a focused electron beam across the surface of a specimen coated with a thin layer of metal, producing a three-dimensional image of surface topography. SEM images of cells reveal surface features such as microvilli, cilia, filopodia, and the texture of the extracellular matrix with striking clarity.

Cryo-electron microscopy (cryo-EM), which images specimens flash-frozen in vitreous ice without chemical fixation or staining, has become one of the most powerful tools in structural biology. Single-particle cryo-EM can determine the three-dimensional structure of protein complexes at near-atomic resolution (2 to 4 angstroms), and cryo-electron tomography can produce three-dimensional reconstructions of cellular structures in their native context. Jacques Dubochet, Joachim Frank, and Richard Henderson received the 2017 Nobel Prize in Chemistry for developing cryo-EM techniques.