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Cell_Biology_Alberts. cells, so as to follow all of their Regulatory systems often employ network motifs that generate useful behaviors: changes in real time? a rapid negative feedback loop dampens the response to input signals; a delayed negative feedback loop creates a biochemical oscillator; positive feedback yields a models to accurately describe the systems that generate transient signal pulses or respond only to sustained inputs. enormous complexity of cellular The dynamic behavior of these network motifs can be dissected in detail with deter-networks and to predict undiscovered ministic and stochastic mathematical modeling. components and mechanisms? Which statements are true? explain why or why not. Discuss the following problems.


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Cell_Biology_Alberts. Which statements are true? explain why or why not. Discuss the following problems. 8–1 Because a monoclonal antibody recognizes a spe-8–7 A common step in the isolation of cells from a cific antigenic site (epitope), it binds only to the specific sample of animal tissue is to treat the tissue with trypsin, protein against which it was made. collagenase, and EDTA. Why is such a treatment nece ssary, and what does each component accomplish? And 8–2 Given the inexorable march of technology, it why does this treatment not kill the cells?


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Cell_Biology_Alberts. seems inevitable that the sensitivity of detection of molecules will ultimately be pushed beyond the yoctomole 8–8 Tropomyosin, at 93 kd, sediments at 2.6S, whereas level (10–24 mole). the 65-kd protein, hemoglobin, sediments at 4.3S. (The sedimentation coefficient S is a linear measure of the rate 8–3 If each cycle of PCR doubles the amount of DNA of sedimentation.) These two proteins are drawn to scale synthesized in the previous cycle, then 10 cycles will give a in Figure Q8–1. How is it that the bigger protein sediments 103-fold amplification, 20 cycles will give a 106-fold amplimore slowly than the smaller one? Can you think of an fication, and 30 cycles will give a 109-fold amplification. analogy from everyday experience that might help you 8–4 To judge the biological importance of an interac-with this problem? tion between protein A and protein B, we need to know quantitative details about their concentrations, affinities, and kinetic behaviors.


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Cell_Biology_Alberts. 8–5 The rate of change in the concentration of any molecular species X is given by the balance between its rate of appearance and its rate of disappearance. 8–6 After a sudden increase in transcription, a protein with a slow rate of degradation will reach a new steady state level more quickly than a protein with a rapid rate of degradation. tropomyosinhemoglobinFigure Q8–1 Scale models of tropomyosin and hemoglobin (Problem 8–8). 8–9 Hybridoma technology allows one to generate monoclonal antibodies to virtually any protein. Why is it, then, that genetically tagging proteins with epitopes is such a commonly used technique, especially since an epitope tag has the potential to interfere with the function of the protein?


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Cell_Biology_Alberts. 8–10 How many copies of a protein need to be present in a cell in order for it to be visible as a band on an SDS gel? Assume that you can load 100 μg of cell extract onto a gel and that you can detect 10 ng in a single band by silver staining the gel. The concentration of protein in cells is about 200 mg/mL, and a typical mammalian cell has a volume of about 1000 μm3 and a typical bacterium a volume of about 1 μm3. Given these parameters, calculate the number of copies of a 120-kd protein that would need to be present in a mammalian cell and in a bacterium in order to give a detectable band on a gel. You might try an order-of-magnitude guess before you make the calculations.


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Cell_Biology_Alberts. 8–11 You have isolated the proteins from two adjacent spots after two-dimensional polyacrylamide-gel electrophoresis and digested them with trypsin. When the masses of the peptides were measured by MALDI-TOF mass spectrometry, the peptides from the two proteins were found to be identical except for one (Figure Q8–2). For this peptide, the mass-to-charge (m/z) values differed by 80, a value that does not correspond to a difference in amino acid sequence. (For example, glutamic acid instead of valine at one position would give an m/z difference of around 30.) Can you suggest a possible difference between the two peptides that might account for the observed m/z difference? Masses of peptides measured by MALDI-TOF mass spectrometry (Problem 8–11). Only the numbered peaks differ between the two protein samples.


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Cell_Biology_Alberts. Masses of peptides measured by MALDI-TOF mass spectrometry (Problem 8–11). Only the numbered peaks differ between the two protein samples. 8–12 You want to amplify the DNA between the two stretches of sequence shown in Figure Q8–3. Of the listed primers, choose the pair that will allow you to amplify the DNA by PCR. 8–13 In the very first round of PCR using genomic DNA, the DNA primers prime synthesis that terminates only when the cycle ends (or when a random end of DNA is encountered). Yet, by the end of 20 to 30 cycles—a typical amplification—the only visible product is defined precisely by the ends of the DNA primers. In what cycle is a double-stranded fragment of the correct size first generated? DNA to be amplifed Figure Q8–3 DNA to be amplified and potential PCR primers (Problem 8–12). 8–14 Explain the difference between a gain-of-function mutation and a dominant-negative mutation. Why are both these types of mutation usually dominant?


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Cell_Biology_Alberts. 8–14 Explain the difference between a gain-of-function mutation and a dominant-negative mutation. Why are both these types of mutation usually dominant? 8–15 Discuss the following statement: “We would have no idea today of the importance of insulin as a regulatory hormone if its absence were not associated with the human disease diabetes. It is the dramatic consequences of its absence that focused early efforts on the identification of insulin and the study of its normal role in physiology.” 8–16 You have just gotten back the results from an RNA-seq analysis of mRNAs from liver. You had anticipated counting the number of reads of each mRNA to determine the relative abundance of different mRNAs. But you are puzzled because many of the mRNAs have given you results like those shown in Figure Q8–4. How is it that different parts of an mRNA can be represented at different levels?


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Cell_Biology_Alberts. 8–17 Examine the network motifs in Figure Q8–5. Decide which ones are negative feedback loops and which are positive. Explain your reasoning. 8–18 Imagine that a random perturbation positions a bistable system precisely at the boundary between two stable states (at the orange dot in Figure Q8–6). How would the system respond? Figure Q8–4 RNA-seq reads for a liver mRNA (Problem 8–16). The exon structure of the mRNA is indicated, with protein-coding segments indicated in light blue and untranslated regions in dark blue. The numbers of sequencing reads are indicated by the heights of the vertical lines above the mRNA. Figure Q8–5 Network motifs composed of transcription activators and repressors (Problem 8–17).


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Cell_Biology_Alberts. 8–19 Detailed analysis of the regulatory region of the Lac operon has revealed surprising complexity. Instead of a single binding site for the Lac repressor, as might be expected, there are three sites termed operators: O1, O2, and O3, arrayed along the DNA as shown in Figure Q8–7. To probe the functions of these three sites, you make a series of constructs in which various combinations of operator sites are present. You examine their ability to repress expression of β-galactosidase, using either tetrameric (wild type) or dimeric (mutant) forms of the Lac repressor. The dimeric form of the repressor can bind to a single operator (with the same affinity as the tetramer) with each monomer binding to half the site. The tetramer, the form normally expressed in cells, can bind to two sites simultaneously. When you measure repression of β-galactosidase expression, you find the results shown in Figure Q8–7, with higher numbers indicating more effective repression. concentration of X


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Cell_Biology_Alberts. concentration of X Figure Q8–6 Perturbations of a bistable system (Problem 8–18). As shown by the green lines, after perturbation 1 the system returns to its original stable state (green dot at left), and after perturbation 2, the system moves to the other stable state (green dot at right). Perturbation 3 moves the system to the precise boundary between the two stable states (orange dot). A. Which single operator site is the most important for repression? How can you tell? b. Do combinations of operator sites (Figure Q8–7, constructs 1, 2, 3, and 5) substantially increase repression by the dimeric repressor? Do combinations of operator sites substantially increase repression by the tetrameric repressor? If the two repressors behave differently, offer an explanation for the difference. C. The wild-type repressor binds O3 very weakly when it is by itself on a segment of DNA. However, if O1 is included on the same segment of DNA, the repressor binds O3 quite well. How can that be?


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Cell_Biology_Alberts. C. The wild-type repressor binds O3 very weakly when it is by itself on a segment of DNA. However, if O1 is included on the same segment of DNA, the repressor binds O3 quite well. How can that be? Figure Q8–7 Repression of β-galactosidase by promoter regions that contain different combinations of Lac repressor binding sites (Problem 8–19). The base-pair (bp) separation of the three operator sites is shown. Numbers at right refer to the level of repression, with higher numbers indicating more effective repression by dimeric (2-mer) or tetrameric (4-mer) repressors. (From S. Oehler et al., EMBO J. 9:973–979, 1990. With permission from John Wiley and Sons.) Ausubel FM, Brent R, Kingston RE et al. (eds) (2002) Short Protocols in Molecular Biology, 5th ed. New York: Wiley. Brown TA (2007) Genomes 3. New York: Garland Science Publishing. Spector DL, Goldman RD & Leinwand LA (eds) (1998) Cells: A Laboratory Manual. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press.


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Cell_Biology_Alberts. Spector DL, Goldman RD & Leinwand LA (eds) (1998) Cells: A Laboratory Manual. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press. Watson JD & Berry A (2008) DNA: The Secret of Life. New York: Alfred A Knopf. Watson JD, Myers RM & Caudy AA (2007) Recombinant DNA: Genes and Genomes – A Short Course, 3rd ed. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press. Ham RG (1965) Clonal growth of mammalian cells in a chemically defined, synthetic medium. Proc. Natl Acad. Sci. USA 53, 288–293. Harlow E & Lane D (1999) Using Antibodies: A Laboratory Manual. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press. Herzenberg LA, Sweet RG & Herzenberg LA (1976) Fluorescence-activated cell sorting. Sci. Am. 234, 108–116. Milstein C (1980) Monoclonal antibodies. Sci. Am. 243, 66–74. de Duve C & Beaufay H (1981) A short history of tissue fractionation. J. Cell Biol. 91, 293s–299s.


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Cell_Biology_Alberts. Milstein C (1980) Monoclonal antibodies. Sci. Am. 243, 66–74. de Duve C & Beaufay H (1981) A short history of tissue fractionation. J. Cell Biol. 91, 293s–299s. Laemmli UK (1970) Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227, 680–685. Scopes RK (1994) Protein Purification: Principles and Practice, 3rd ed. New York: Springer-Verlag. Simpson RJ, Adams PD & Golemis EA (2008) Basic Methods in Protein Purification and Analysis: A Laboratory Manual. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press. Wood DW (2014) New trends and affinity tag designs for recombinant protein purification. Curr. Opin. Struct. Biol. 26, 54–61. Choudhary C & Mann M (2010) Decoding signalling networks by mass spectrometry-based proteomics. Nat. Rev. Mol. Cell Biol. 11, 427–439. Domon B & Aebersold R (2006) Mass spectrometry and protein analysis. Science 312, 212–217.


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Cell_Biology_Alberts. Domon B & Aebersold R (2006) Mass spectrometry and protein analysis. Science 312, 212–217. Goodrich JA & Kugel JF (2007) Binding and Kinetics for Molecular Biologists. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press. Kendrew JC (1961) The three-dimensional structure of a protein molecule. Sci. Am. 205, 96–111. Knight ZA & Shokat KM (2007) Chemical genetics: where genetics and pharmacology meet. Cell 128, 425–430. O’Farrell PH (1975) High resolution two-dimensional electrophoresis of proteins. J. Biol. Chem. 250, 4007–4021. Pollard TD (2010) A guide to simple and informative binding assays. Mol. Biol. Cell 21, 4061–4067. Wüthrich K (1989) Protein structure determination in solution by nuclear magnetic resonance spectroscopy. Science 243, 45–50. Cohen S, Chang A, Boyer H & Helling R (1973) Construction of biologically functional bacterial plasmids in vitro. Proc. Natl Acad. Sci. USA 70, 3240–3244.


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Cell_Biology_Alberts. Cohen S, Chang A, Boyer H & Helling R (1973) Construction of biologically functional bacterial plasmids in vitro. Proc. Natl Acad. Sci. USA 70, 3240–3244. Green MR & Sambrook J (2012) Molecular Cloning: A Laboratory Manual, 4th ed. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press. International Human Genome Sequencing Consortium (2001) Initial sequencing and analysis of the human genome. Nature 409, 860–921. Jackson D, Symons R & Berg P (1972) Biochemical method for inserting new genetic information into DNA of Simian Virus 40: circular SV40 DNA molecules containing lambda phage genes and the galactose operon of Escherichia coli. Proc. Natl Acad. Sci. USA 69, 2904–2909. Kosuri S & Church GM (2014) Large-scale de novo DNA synthesis: technologies and applications. Nat. Methods 11, 499–507. Maniatis T, Hardison RC, Lacy E et al. (1978) The isolation of structural genes from libraries of eucaryotic DNA. Cell 15, 687–701.


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Cell_Biology_Alberts. Maniatis T, Hardison RC, Lacy E et al. (1978) The isolation of structural genes from libraries of eucaryotic DNA. Cell 15, 687–701. Mullis KB (1990) The unusual origin of the polymerase chain reaction. Sci. Am. 262, 56–61. Nathans D & Smith HO (1975) Restriction endonucleases in the analysis and restructuring of DNA molecules. Annu. Rev. Biochem. 44, 273–293. Saiki RK, Gelfand DH, Stoffel S et al. (1988) Primer-directed enzymatic amplification of DNA with a thermostable DNA polymerase. Science 239, 487–491. Sanger F, Nicklen S & Coulson AR (1977) DNA sequencing with chain-terminating inhibitors. Proc. Natl Acad. Sci. USA 74, 5463–5467. Shendure J & Lieberman Aiden E (2012) The expanding scope of DNA sequencing. Nat. Biotechnol. 30, 1084–1094. Botstein D, White RL, Skolnick M & Davis RW (1980) Construction of a genetic linkage map in man using restriction fragment length polymorphisms. Am. J. Hum. Genet. 32, 314–331.


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Cell_Biology_Alberts. Botstein D, White RL, Skolnick M & Davis RW (1980) Construction of a genetic linkage map in man using restriction fragment length polymorphisms. Am. J. Hum. Genet. 32, 314–331. DeRisi JL, Iyer VR & Brown PO (1997) Exploring the metabolic and genetic control of gene expression on a genomic scale. Science 278, 680–686. Esvelt KM, Mali P, Braff JL et al. (2013) Orthogonal Cas9 proteins for RNA-guided gene regulation and editing. Nat. Methods 10, 1116–1121. Fellmann C & Lowe SW (2014) Stable RNA interference rules for silencing. Nat. Cell Biol. 16, 10–18. Mello CC & Conte D (2004) Revealing the world of RNA interference. Nature 431, 338–342. Nüsslein-Volhard C & Wieschaus E (1980) Mutations affecting segment number and polarity in Drosophila. Nature 287, 795–801. Palmiter RD & Brinster RL (1985) Transgenic mice. Cell 41, 343–345. Weigel D & Glazebrook J (2002) Arabidopsis: A Laboratory Manual. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press.


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Cell_Biology_Alberts. Palmiter RD & Brinster RL (1985) Transgenic mice. Cell 41, 343–345. Weigel D & Glazebrook J (2002) Arabidopsis: A Laboratory Manual. Cold Spring Harbor, NY: Cold Spring Harbor Laboratory Press. Wilson RC & Doudna JA (2013) Molecular mechanisms of RNA interference. Annu. Rev. Biophys. 42, 217–239. Mathematical Analysis of Cell Functions Alon U (2006) An Introduction to Systems Biology: Design Principles of Biological Circuits. Boca Raton, FL: Chapman & Hall/CRC. Alon U (2007) Network motifs: theory and experimental approaches. Nat. Rev. Genet. 8, 450–461. Ferrell JE Jr (2002) Self-perpetuating states in signal transduction: positive feedback, double-negative feedback and bistability. Curr. Opin. Cell Biol. 14, 140–148. Ferrell JE Jr, Tsai TY & Yang Q (2011) Modeling the cell cycle: why do certain circuits oscillate? Cell 144, 874–885. Gunawardena J (2014) Models in biology: ‘accurate descriptions of our pathetic thinking’. BMC Biol. 12, 29.


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Cell_Biology_Alberts. Gunawardena J (2014) Models in biology: ‘accurate descriptions of our pathetic thinking’. BMC Biol. 12, 29. Lewis J (2008) From signals to patterns: space, time, and mathematics in developmental biology. Science 322, 399–403. Mogilner A, Allard J & Wollman R (2012) Cell polarity: quantitative modeling as a tool in cell biology. Science 336, 175–179. Novak B & Tyson JJ (2008) Design principles of biochemical oscillators. Nat. Rev. Mol. Cell Biol. 9, 981–991. Silva-Rocha R & de Lorenzo V (2008) Mining logic gates in prokaryotic transcriptional regulation networks. FEBS Lett. 582, 1237–1244. Tyson JJ, Chen KC & Novak B (2003) Sniffers, buzzers, toggles and blinkers: dynamics of regulatory and signaling pathways in the cell. Curr. Opin. Cell Biol. 15, 221–231.


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Cell_Biology_Alberts. Understanding the structural organization of cells is essential for learning how they function. In this chapter, we briefly describe some of the principal microscopy methods used to study cells. Optical microscopy will be our starting point because cell biology began with the light microscope, and it is still an indispensible tool. The development of methods for the specific labeling and imaging of individual cellular constituents and the reconstruction of their threedimensional architecture has meant that, far from falling into disuse, optical microscopy continues to increase in importance. One advantage of optical microscopy is that light is relatively nondestructive. By tagging specific cell components with fluorescent probes, such as intrinsically fluorescent proteins, we can watch their movement, dynamics, and interactions in living cells. Although conventional optical microscopy is limited in resolution by the wavelength of visible light, new methods cleverly bypass this


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Cell_Biology_Alberts. their movement, dynamics, and interactions in living cells. Although conventional optical microscopy is limited in resolution by the wavelength of visible light, new methods cleverly bypass this limitation and allow the position of even single molecules to be mapped. By using a beam of electrons instead of visible light, electron microscopy can image the interior of cells, and their macromolecular components, at almost atomic resolution, and in three dimensions.


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Cell_Biology_Alberts. This chapter is intended as a companion, rather than an introduction, to the chapters that follow; readers may wish to refer back to it as they encounter applications of microscopy to basic biological problems in the later pages of the book. Looking at CeLLs in the Light MiCrosCope A typical animal cell is 10–20 μm in diameter, which is about onefifth the size of the smallest object that we can normally see with the naked eye. Only after good light microscopes became available in the early part of the nineteenth century did Schleiden and Schwann propose that all plant and animal tissues were aggregates of individual cells. Their proposal in 1838, known as the cell doctrine, marks the formal birth of cell biology.


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Cell_Biology_Alberts. Animal cells are not only tiny, but they are also colorless and translucent. The discovery of their main internal features, therefore, depended on the development, in the late nineteenth century, of a variety of stains that provided sufficient contrast to make those features visible. Similarly, the far more powerful electron microscope introduced in the early 1940s required the development of new techniques for preserving and staining cells before the full complexities of their internal fine structure could begin to emerge. To this day, microscopy often relies as much on techniques for preparing the specimen as on the performance of the microscope itself. In the following discussions, we therefore consider both instruments and specimen preparation, beginning with the light microscope.


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Cell_Biology_Alberts. The images in Figure 9–1 illustrate a stepwise progression from a thumb to a cluster of atoms. Each successive image represents a tenfold increase in magnification. The naked eye can see features in the first two panels, the light microscope allows us to see details corresponding to about the fourth or fifth panel, and the electron microscope takes us to about the seventh or eighth panel. Figure 9–2 shows the sizes of various cellular and subcellular structures and the ranges of size that different types of microscopes can visualize. Looking at CeLLs in the Light MiCrosCope Looking at CeLLs anD MoLeCULes in the eLeCtron MiCrosCope 20 mm 2 mm 0.2 mm 20 µm 2 µm 0.2 µm 20 nm 2 nm 0.2 nm the Light Microscope Can resolve Details 0.2 μm apart


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Cell_Biology_Alberts. Looking at CeLLs anD MoLeCULes in the eLeCtron MiCrosCope 20 mm 2 mm 0.2 mm 20 µm 2 µm 0.2 µm 20 nm 2 nm 0.2 nm the Light Microscope Can resolve Details 0.2 μm apart For well over 100 years, all microscopes were constrained by a fundamental limitation: that a given type of radiation cannot be used to probe structural details much smaller than its own wavelength. A limit to the resolution of a light microscope was therefore set by the wavelength of visible light, which ranges from about 0.4 μm (for violet) to 0.7 μm (for deep red). In practical terms, bacteria and mitochondria, which are about 500 nm (0.5 μm) wide, are generally the smallest objects whose shape we can clearly discern in the light microscope; details smaller than this are obscured by effects resulting from the wavelike nature of light. To understand why this occurs, we must follow the behavior of a beam of light as it passes through the lenses of a microscope (Figure 9–3).


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Cell_Biology_Alberts. Because of its wave nature, light does not follow the idealized straight ray paths that geometrical optics predicts. Instead, light waves travel through an optical system by many slightly different routes, like ripples in water, so that they Figure 9–1 A sense of scale between living cells and atoms. each diagram shows an image magnified by a factor of ten in an imaginary progression from a thumb, through skin cells, to a ribosome, to a cluster of atoms forming part of one of the many protein molecules in our body. atomic details of biological macromolecules, as shown in the last two panels, are usually beyond the power of the electron microscope. While color has been used here in all the panels, it is not a feature of objects much smaller than the wavelength of light, so the last five panels should really be in black and white. limit of limit of super-conventional 0.1 nm(1 A)


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Cell_Biology_Alberts. limit of limit of super-conventional 0.1 nm(1 A) Figure 9–2 Resolving power. sizes of cells and their components are drawn on a logarithmic scale, indicating the range of objects that can be readily resolved by the naked eye and in the light and electron microscopes. note that new superresolution microscopy techniques, discussed in detail later, allow an improvement in resolution by an order of magnitude compared with conventional light microscopy.


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Cell_Biology_Alberts. interfere with one another and cause optical diffraction effects. If two trains of waves reaching the same point by different paths are precisely in phase, with crest matching crest and trough matching trough, they will reinforce each other so as to increase brightness. In contrast, if the trains of waves are out of phase, they will interfere with each other in such a way as to cancel each other partly or entirely (Figure 9–4). The interaction of light with an object changes the phase relationships of the light waves in a way that produces complex interference effects. At high magnification, for example, the shadow of an edge that is evenly illuminated with light of uniform wavelength appears as a set of parallel lines (Figure 9–5), whereas that of a circular spot appears as a set of concentric rings. For the same reason, a single point seen through a microscope appears as a blurred disc, and two point objects close together give overlapping images and may merge into one.


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Cell_Biology_Alberts. the following units of length are commonly employed in microscopy: μm (micrometer) = 10–6 m nm (nanometer) = 10–9 m Å (Ångström unit) = 10–10 m Figure 9–3 A light microscope. (a) Diagram showing the light path in a compound microscope. Light is focused on the specimen by lenses in the condenser. a combination of objective lenses, tube lenses, and eyepiece lenses is arranged to focus an image of the illuminated specimen in the eye. (B) a modern research light microscope. (B, courtesy of Carl Zeiss Microscopy, gmbh.) Although no amount of refinement of the lenses can overcome the diffraction limit imposed by the wavelike nature of light, other ways of cleverly bypassing this limit have emerged, creating socalled superresolution imaging techniques that can even detect the position of single molecules.


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Cell_Biology_Alberts. The limiting separation at which two objects appear distinct—the socalled limit of resolution—depends on both the wavelength of the light and the numerical aperture of the lens system used. The numerical aperture affects the lightgathering ability of the lens and is related both to the angle of the cone of light that can enter it and to the refractive index of the medium the lens is operating in; the wider the microscope opens its eye, so to speak, the more sharply it can see (Figure 9–6). The refractive index is the ratio of the speed of light in a vacuum to the speed of light in a particular transparent medium. For example, for water this is 1.33, meaning that light travels 1.33 times slower in water than in a vacuum. Under the best conditions, with violet light (wavelength = 0.4 μm) and a numerical aperture of 1.4, the basic light microscope can theoretically achieve a limit of resolution of about 0.2 μm, or 200 nm. Some microscope makers at the end of the nineteenth century


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Cell_Biology_Alberts. μm) and a numerical aperture of 1.4, the basic light microscope can theoretically achieve a limit of resolution of about 0.2 μm, or 200 nm. Some microscope makers at the end of the nineteenth century achieved this resolution, but it is routinely matched in contemporary, factoryproduced microscopes. Although it is possible to enlarge an image as much as we want—for example, by projecting it onto a screen—it is not possible, in a conventional light microscope, to resolve two objects in the light microscope that are separated by less than about 0.2 μm; they will appear as a single object. It is important, however, to distinguish between resolution and detection. If a small object, below the resolution limit, itself emits light, then we may still be able to see or detect it. Thus, we can see a single fluorescently labeled microtubule even though it is about ten times thinner than the resolution limit of the light microscope. Diffraction effects, however, will cause it to appear blurred


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Cell_Biology_Alberts. a single fluorescently labeled microtubule even though it is about ten times thinner than the resolution limit of the light microscope. Diffraction effects, however, will cause it to appear blurred and at least 0.2 μm thick (see Figure 9–16). In a similar way, we can see the stars in the night sky, even though their diameters are far below the angular resolution of our unaided eyes: they all appear as similar, slightly blurred points of light, differing only in their color and brightness.


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Cell_Biology_Alberts. photon noise Creates additional Limits to resolution When Light Levels are Low Any image, whether produced by an electron microscope or by an optical microscope, is made by particles—electrons or photons—striking a detector of some sort. But these particles are governed by quantum mechanics, so the numbers reaching the detector are predictable only in a statistical sense. Finite samples, collected by imaging for a limited period of time (that is, by taking a snapshot), will show random variation: successive snapshots of the same scene will not be exactly identical. Moreover, every detection method has some level of background signal or noise, adding to the statistical uncertainty. With bright illumination, corresponding to very large numbers of photons or electrons, the features of the imaged


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Cell_Biology_Alberts. Figure 9–4 Interference between light waves. When two light waves combine in phase, the amplitude of the resultant wave is larger and the brightness is increased. two light waves that are out of phase cancel each other partly and produce a wave whose amplitude, and therefore brightness, is decreased. Figure 9–5 Images of an edge and of a point of light. (a) the interference effects, or fringes, seen at high magnification when light of a specific wavelength passes the edge of a solid object placed between the light source and the observer. (B) the image of a point source of light. Diffraction spreads this out into a complex, circular pattern, whose width depends on the numerical aperture of the optical system: the smaller the aperture, the bigger (more blurred) the diffracted image. two point sources can be just resolved when the center of the image of one lies on the first dark ring in the image of the other: this is used to define the limit of resolution.


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Cell_Biology_Alberts. Figure 9–6 Numerical aperture. the path of light rays passing through a transparent specimen in a microscope illustrates the concept of numerical aperture and its relation to the limit of resolution. the objective lens collects a cone of light rays to create an image the condenser lens focuses a cone of light rays onto each point of the specimen RESOLUTION: the resolving power of the microscope depends on the width of the cone of illumination and therefore on both the condenser and the objective lens. It is calculated using the formula 0.61 ˛ where: ˜ = half the angular width of the cone of rays collected by the objective lens from a typical point in the central region of the specimen (since the maximum width is 180o, sin ˜ has a maximum value of 1) n = the refractive index of the medium (usually air or oil) separating the specimen from the objective and condenser lenses ˛ = the wavelength of light used (for white light a fgure of 0.53 µm is commonly assumed)


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Cell_Biology_Alberts. NUMERICAL APERTURE: n sin ˜ in the aperture, the greater the resolution and the equation above is called the numerical aperture brighter the image (brightness is important in of the lens and is a function of its light-fuorescence microscopy). However, this advancollecting ability. For dry lenses this cannot be tage does necessitate very short working distances more than 1, but for oil-immersion lenses it can and a very small depth of feld. be as high as 1.4. The higher the numerical specimen are accurately determined based on the distribution of these particles at the detector. However, with smaller numbers of particles, the structural details of the specimen are obscured by the statistical fluctuations in the numbers of particles detected in each region, which give the image a speckled appearance and limit its precision. The term noise describes this random variability. Living Cells are seen Clearly in a phase-Contrast or a Differential-interference-Contrast Microscope


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Cell_Biology_Alberts. Living Cells are seen Clearly in a phase-Contrast or a Differential-interference-Contrast Microscope There are many ways in which contrast in a specimen can be generated (Figure 9–7A). While fixing and staining a specimen can generate contrast through color, microscopists have always been challenged by the possibility that some components of the cell may be lost or distorted during specimen preparation. The only certain way to avoid the problem is to examine cells while they are alive, without fixing or freezing. For this purpose, light microscopes with special optical systems are especially useful.


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Cell_Biology_Alberts. In the normal bright-field microscope, light passing through a cell in culture forms the image directly. Another system, dark-field microscopy, exploits the fact that light rays can be scattered in all directions by small objects in their path. If oblique lighting from the condenser is arranged, which does not directly enter the objective, focused but unstained objects in a living cell can scatter the rays, some of which then enter the objective to create a bright image against a black background (Figure 9–7B).


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Cell_Biology_Alberts. When light passes through a living cell, the phase of the light wave is changed according to the cell’s refractive index: a relatively thick or dense part of the cell, such as a nucleus, slows the light passing through it. The phase of the light, consequently, is shifted relative to light that has passed through an adjacent thinner region of the cytoplasm (Figure 9–7C). The phase-contrast microscope and, in a more complex way, the differential-interference-contrast microscope increase these phase differences so that the waves are more nearly out of phase, producing amplitude differences when the sets of waves recombine, thereby creating an image of the cell’s structure. Both types of light microscopy are widely used to visualize living cells (see Movie 17.2). Figure 9–8 compares images of the same cell obtained by four kinds of light microscopy.


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Cell_Biology_Alberts. Figure 9–7 Contrast in light microscopy. (a) the stained portion of the cell will absorb light of some wavelengths, which depends on the stain, but will allow other wavelengths to pass through it. a colored image of the cell is thereby obtained that is visible in the normal bright-field light microscope. (B) in the dark-field microscope, oblique rays of light focused on the specimen do not enter the objective lens, but light that is scattered by components in the living cell can be collected to produce a bright image on a dark background. (C) Light passing through the unstained living cell experiences very little change in amplitude, and the structural details cannot be seen even if the image is highly magnified. the phase of the light, however, is altered by its passage through either thicker or denser parts of the cell, and small phase differences can be made visible by exploiting interference effects using a phase-contrast or a differential-interference-contrast microscope.


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Cell_Biology_Alberts. Phasecontrast, differentialinterferencecontrast, and darkfield microscopy make it possible to watch the movements involved in such processes as mitosis and cell migration. Since many cellular motions are too slow to be seen in real time, it is often helpful to make timelapse movies in which the camera records successive frames separated by a short time delay, so that when the resulting picture series is played at normal speed, events appear greatly speeded up.


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Cell_Biology_Alberts. In recent years, electronic, or digital, imaging systems, and the associated technology of image processing, have had a major impact on light microscopy. Certain practical limitations of microscopes relating to imperfections in the optical system have been largely overcome. Electronic imaging systems have also circumvented two fundamental limitations of the human eye: the eye cannot see well in extremely dim light, and it cannot perceive small differences in light intensity against a bright background. To increase our ability to observe cells in these difficult conditions, we can attach a sensitive digital camera to a microscope. These cameras detect light by means of chargecoupled devices (CCDs), or high sensitivity complementary metaloxide semiconductor (CMOS) sensors, similar to those found in digital cameras. Such image sensors are 10 times more sensitive than the human eye and can detect 100 times more intensity levels. It is therefore possible to observe cells for long periods


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Cell_Biology_Alberts. found in digital cameras. Such image sensors are 10 times more sensitive than the human eye and can detect 100 times more intensity levels. It is therefore possible to observe cells for long periods at very low light levels, thereby avoiding the damaging effects of prolonged bright light (and heat). Such lowlight cameras are especially important for viewing fluorescent molecules in living cells, as explained below.


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Cell_Biology_Alberts. Because images produced by digital cameras are in electronic form, they can be processed in various ways to extract latent information. Such image processing makes it possible to compensate for several optical faults in microscopes. Moreover, by digital image processing, contrast can be greatly enhanced to overcome Figure 9–8 Four types of light microscopy. Four images are shown of the same fibroblast cell in culture. all images can be obtained with most modern microscopes by interchanging optical components. (a) Bright-field microscopy, in which light is transmitted straight through the specimen. (B) phase-contrast microscopy, in which phase alterations of light transmitted through the specimen are translated into brightness changes. (C) Differential-interference-contrast microscopy, which highlights edges where there is a steep change of refractive index. (D) Dark-field microscopy, in which the specimen is lit from the side and only the scattered light is seen.


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Cell_Biology_Alberts. the eye’s limitations in detecting small differences in light intensity, and background irregularities in the optical system can be digitally subtracted. This procedure reveals small transparent objects that were previously impossible to distinguish from the background. intact tissues are Usually Fixed and sectioned Before Microscopy Because most tissue samples are too thick for their individual cells to be examined directly at high resolution, they are often cut into very thin transparent slices, or sections. To preserve the cells within the tissue they must be treated with a fixative. Common fixatives include glutaraldehyde, which forms covalent bonds with the free amino groups of proteins, crosslinking them so they are stabilized and locked into position.


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Cell_Biology_Alberts. Because tissues are generally soft and fragile, even after fixation, they need to be either frozen or embedded in a supporting medium before being sectioned. The usual embedding media are waxes or resins. In liquid form, these media both permeate and surround the fixed tissue; they can then be hardened (by cooling or by polymerization) to form a solid block, which is readily sectioned with a microtome. This is a machine with a sharp blade, usually of steel or glass, which operates like a meatslicer (Figure 9–9). The sections (typically 0.5–10 μm thick) are then laid flat on the surface of a glass microscope slide.


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Cell_Biology_Alberts. There is little in the contents of most cells (which are 70% water by weight) to impede the passage of light rays. Thus, most cells in their natural state, even if fixed and sectioned, are almost invisible in an ordinary light microscope. We have seen that cellular components can be made visible by techniques such as phasecontrast and differentialinterferencecontrast microscopy, but these methods tell us almost nothing about the underlying chemistry. There are three main approaches to working with thin tissue sections that reveal differences in types of molecules that are present. First, and traditionally, sections can be stained with organic dyes that have some specific affinity for particular subcellular components. The dye hematoxylin, for example, has an affinity for negatively charged molecules and therefore reveals the distribution of DNA, RNA, and acidic proteins in a cell (Figure 9–10). The chemical basis for the specificity of many dyes, however, is not known.


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Cell_Biology_Alberts. movement of microtome arm ribbon of ribbon of sections on glass slide, stained and mounted under a glass cover slip Figure 9–9 Making tissue sections. this illustration shows how an embedded tissue is sectioned with a microtome in preparation for examination in the light microscope. Second, sectioned tissues can be used to visualize specific patterns of differential gene expression. In situ hybridization, discussed earlier (see Figure 8–34), reveals the cellular distribution and abundance of specific expressed RNA molecules in sectioned material or in whole mounts of small organisms or organs. This is particularly effective when used in conjunction with fluorescent probes (Figure 9–11). A third and very sensitive approach, generally and widely applicable for localizing proteins of interest, also depends on using fluorescent probes and markers, as we explain next.


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Cell_Biology_Alberts. A third and very sensitive approach, generally and widely applicable for localizing proteins of interest, also depends on using fluorescent probes and markers, as we explain next. Fluorescent molecules absorb light at one wavelength and emit it at another, longer wavelength (Figure 9–12A). If we illuminate such a molecule at its absorbing wavelength and then view it through a filter that allows only light of the emitted wavelength to pass, it will glow against a dark background. Because the background is dark, even a minute amount of the glowing fluorescent dye can be detected. In contrast, the same number of molecules of a nonfluorescent stain, viewed conventionally, would be practically indiscernible because the absorption of light by molecules in the stain would result in only the faintest tinge of color in the light transmitted through that part of the specimen.


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Cell_Biology_Alberts. The fluorescent dyes used for staining cells are visualized with a fluorescence microscope. This microscope is similar to an ordinary light microscope except that the illuminating light, from a very powerful source, is passed through two sets of filters—one to filter the light before it reaches the specimen and one to Figure 9–10 Staining of cell components. (a) this section of cells in the urine-collecting ducts of the kidney was stained with hematoxylin and eosin, two dyes commonly used in histology. each duct is made of closely packed cells (with nuclei stained red) that form a ring. the ring is surrounded by extracellular matrix, stained purple. (B) this section of a young plant root is stained with two dyes, safranin and fast green. the fast green stains the cellulosic cell walls while the safranin stains the lignified xylem cell walls bright red. (a, from p.r. Wheater et al., Functional histology, 2nd ed. London: Churchill Livingstone, 1987; B, courtesy of stephen grace.)


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Figure 9–11 RNA in situ hybridization.

Cell_Biology_Alberts. Figure 9–11 RNA in situ hybridization.


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Cell_Biology_Alberts. as described in Chapter 8 (see Figure 8–62), it is possible to visualize the distribution of different rnas in tissues using in situ hybridization. here, the transcription pattern of five different genes involved in patterning the early fly embryo is revealed in a single embryo. each rna probe has been fluorescently labeled in a different way, some directly and some indirectly; the resulting images are displayed each in a different color (“false-colored”) and combined to give an image where different color combinations represent different sets of genes expressed. the genes whose expression pattern is revealed here are wingless (yellow), engrailed (blue), short gastrulation (red), intermediate neuroblasts defective (green), and muscle specific homeobox (purple). (From D. kosman et al., Science 305:846, 2004. With permission from aaas.) energy of orbital electronin fuorophore emission of photon 3 second barrier flter: cuts out unwanted fuorescent signals, passing the specifc green


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Cell_Biology_Alberts. Science 305:846, 2004. With permission from aaas.) energy of orbital electronin fuorophore emission of photon 3 second barrier flter: cuts out unwanted fuorescent signals, passing the specifc green fuorescein emission between 520 and 560 nm 2 beam-splitting mirror: refects light below 510 nm but transmits light above 510 nm


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Cell_Biology_Alberts. Figure 9–12 Fluorescence and the fluorescence microscope. (a) an orbital electron of a fluorochrome molecule can be raised to an excited state following the absorption of a photon. Fluorescence occurs when the electron returns to its ground state and emits a photon of light at a longer wavelength. too much exposure to light, or too bright a light, can also destroy the fluorochrome molecule, in a process called photobleaching. (B) in the fluorescence microscope, a filter set consists of two barrier filters (1 and 3) and a dichroic (beam-splitting) mirror (2). this example shows the filter set for detection of the fluorescent molecule fluorescein. high-numerical-aperture objective lenses are especially important in this type of microscopy because, for a given magnification, the brightness of the fluorescent image is proportional to the fourth power of the numerical aperture Figure 9–14 Different fluorescent probes can be visualized in the same cell. in this composite micrograph of a


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Cell_Biology_Alberts. of the fluorescent image is proportional to the fourth power of the numerical aperture Figure 9–14 Different fluorescent probes can be visualized in the same cell. in this composite micrograph of a cell in mitosis, three different fluorescent probes have been used to label three different cellular components (Movie 9.1). the spindle microtubules are revealed with a green fluorescent antibody, centromeres with a red fluorescent antibody, and the Dna of the condensed chromosomes with the blue fluorescent dye Dapi. (Courtesy of kevin F. sullivan.) (see Figure 9–13) but, like many organic fluorochromes, they fade fairly rapidly when continuously illuminated. More stable fluorochromes have been developed based on inorganic chemistry. Tiny crystals of semiconductor material, called nanoparticles, or quantum dots, can be excited to fluoresce by a broad spectrum of blue light. Their emitted light has a color that depends on the exact size of the nanocrystal, between 2 and 10 nm in diameter,


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Cell_Biology_Alberts. or quantum dots, can be excited to fluoresce by a broad spectrum of blue light. Their emitted light has a color that depends on the exact size of the nanocrystal, between 2 and 10 nm in diameter, and additionally the fluorescence fades only slowly with time (Figure 9–15). These nanoparticles, when coupled to other probes such as antibodies, are therefore ideal for tracking molecules over time. If introduced into a living cell, in an embryo for example, the progeny of that cell can be followed many days later by their fluorescence, allowing cell lineages to be tracked.


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Cell_Biology_Alberts. Figure 9–15 Fluorescent nanoparticles or quantum dots. (a) Quantum dots are tiny particles of cadmium selenide, a semiconductor, with a coating to make them water-soluble. they can be coupled to protein molecules such as antibodies or streptavidin and, when introduced into a cell, will bind to a target protein of interest. Different-sized quantum dots emit light of different colors—the larger the dot, the longer the wavelength—but they are all excited by the same blue light. Quantum dots can keep shining for weeks, unlike most fluorescent organic dyes. (B) in this cell, microtubules are labeled (green) with an organic fluorescent dye (alexa 488), while a nuclear protein is stained (red) with quantum dots bound to streptavidin. on continuous exposure to strong blue light, the fluorescent dye fades quickly while the quantum dots continue to shine. (C) in this cell, the labeling pattern is reversed; a nuclear protein is labeled (green) with an organic fluorescent dye (alexa 488), while


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Cell_Biology_Alberts. dye fades quickly while the quantum dots continue to shine. (C) in this cell, the labeling pattern is reversed; a nuclear protein is labeled (green) with an organic fluorescent dye (alexa 488), while microtubules are labeled (red) with quantum dots. again, the quantum dots far outlast the fluorescent dye. (B and C, from L. Medintz et al., Nat. Mater. 4:435–446, 2005. With permission from Macmillan publishers Ltd.)


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Cell_Biology_Alberts. Later in the chapter, additional fluorescence microscopy methods will be discussed that can be used to monitor changes in the concentration and location of specific molecules inside living cells. antibodies Can Be Used to Detect specific Molecules


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Cell_Biology_Alberts. antibodies Can Be Used to Detect specific Molecules Antibodies are proteins produced by the vertebrate immune system as a defense against infection (discussed in Chapter 24). They are unique among proteins in that they are made in billions of different forms, each with a different binding site that recognizes a specific target molecule (or antigen). The precise antigen specificity of antibodies makes them powerful tools for the cell biologist. When labeled with fluorescent dyes, antibodies are invaluable for locating specific molecules in cells by fluorescence microscopy (Figure 9–16); labeled with electrondense particles such as colloidal gold spheres, they are used for similar purposes in the electron microscope (discussed below). The antibodies employed in microscopy are commonly either purified from antiserum so as to remove all nonspecific antibodies, or they are specific monoclonal antibodies that only recognize the target molecule.


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Cell_Biology_Alberts. When we use antibodies as probes to detect and assay specific molecules in cells, we frequently use chemical methods to amplify the fluorescent signal they produce. For example, although a marker molecule such as a fluorescent dye can be linked directly to an antibody—the primary antibody—a stronger signal is achieved by using an unlabeled primary antibody and then detecting it with a group of labeled secondary antibodies that bind to it (Figure 9–17). This process is called indirect immunocytochemistry. Some amplification methods use an enzyme as a marker molecule attached to the secondary antibody. The enzyme alkaline phosphatase, for example, in the presence of appropriate chemicals, produces inorganic phosphate that in turn primary antibody: secondary antibodies: Figure 9–16 Immunofluorescence.


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Cell_Biology_Alberts. Figure 9–16 Immunofluorescence. a transmission electron micrograph of the periphery of a cultured epithelial cell showing the distribution of microtubules and other filaments. (B) the same area stained with fluorescent antibodies against tubulin, the protein that assembles to form microtubules, using the technique of indirect immunocytochemistry (see Figure 9–17). Red arrows indicate individual microtubules that are readily recognizable in both images. note that, because of diffraction effects, the microtubules in the light microscope appear 0.2 μm wide rather than their true width of 0.025 μm. (From M. osborn, r. Webster and k. Weber, J. Cell Biol. 77:r27–r34, 1978. With permission from the rockefeller University press.) Figure 9–17 Indirect immunocytochemistry.


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Cell_Biology_Alberts. Weber, J. Cell Biol. 77:r27–r34, 1978. With permission from the rockefeller University press.) Figure 9–17 Indirect immunocytochemistry. this detection method is very sensitive because many molecules of the secondary antibody recognize each primary antibody. the secondary antibody is covalently coupled to a marker molecule that makes it readily detectable. Commonly used marker molecules include fluorescent probes (for fluorescence microscopy), the enzyme horseradish peroxidase (for either conventional light microscopy or electron microscopy), colloidal gold spheres (for electron microscopy), and the enzymes alkaline phosphatase or peroxidase (for biochemical detection).


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Cell_Biology_Alberts. leads to the local formation of a colored precipitate. This reveals the location of the secondary antibody and hence the location of the antibody–antigen complex. Since each enzyme molecule acts catalytically to generate many thousands of molecules of product, even tiny amounts of antigen can be detected. Although the enzyme amplification makes enzymelinked methods sensitive, diffusion of the colored precipitate away from the enzyme limits the spatial resolution of this method for microscopy, and fluorescent labels are usually used for the most sensitive and precise optical localization. imaging of Complex three-Dimensional objects is possible with the optical Microscope


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Cell_Biology_Alberts. imaging of Complex three-Dimensional objects is possible with the optical Microscope For ordinary light microscopy, as we have seen, a tissue has to be sliced into thin sections to be examined; the thinner the section, the crisper the image. Since information about the third dimension is lost upon sectioning, how, then, can we get a picture of the threedimensional architecture of a cell or tissue, and how can we view the microscopic structure of a specimen that, for one reason or another, cannot first be sliced into sections? Although an optical microscope is focused on a particular focal plane within a threedimensional specimen, all the other parts of the specimen, above and below the plane of focus, are also illuminated and the light originating from these regions contributes to the image as “outoffocus” blur. This can make it very hard to interpret the image in detail and can lead to fine image structure being obscured by the outoffocus light.


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Cell_Biology_Alberts. Two distinct but complementary approaches solve this problem: one is computational, the other optical. These threedimensional microscopic imaging methods make it possible to focus on a chosen plane in a thick specimen while rejecting the light that comes from outoffocus regions above and below that plane. Thus one sees a crisp, thin optical section. From a series of such optical sections taken at different depths and stored in a computer, a threedimensional image can be reconstructed. The methods do for the microscopist what the computed tomography (CT) scanner does (by different means) for the radiologist investigating a human body: both machines give detailed sectional views of the interior of an intact structure.


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Cell_Biology_Alberts. The computational approach is often called image deconvolution. To understand how it works, remember that the wavelike nature of light means that the microscope lens system produces a small blurred disc as the image of a point light source (see Figure 9–5), with increased blurring if the point source lies above or below the focal plane. This blurred image of a point source is called the point spread function (see Figure 9–36). An image of a complex object can then be thought of as being built up by replacing each point of the specimen by a corresponding blurred disc, resulting in an image that is blurred overall. For deconvolution, we first obtain a series of (blurred) images, usually with a cooled CCD camera or more recently a CMOS camera, focusing the microscope in turn on a series of focal planes—in effect, a (blurred) threedimensional image. Digital processing of the stack of digital images then removes as much of the blur as possible. In essence, the computer program uses the


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Cell_Biology_Alberts. focal planes—in effect, a (blurred) threedimensional image. Digital processing of the stack of digital images then removes as much of the blur as possible. In essence, the computer program uses the measured point spread function of a point source of light from that microscope to determine what the effect of the blurring would have been on the image, and then applies an equivalent “deblurring” (deconvolution), turning the blurred threedimensional image into a series of clean optical sections, albeit still constrained by the diffraction limit. Figure 9–18 shows an example.


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Cell_Biology_Alberts. the Confocal Microscope produces optical sections by excluding out-of-Focus Light The confocal microscope achieves a result similar to that of deconvolution, but does so by manipulating the light before it is measured; it is an analog technique rather than a digital one. The optical details of the confocal microscope are complex, but the basic idea is simple, as illustrated in Figure 9–19, and the results are Figure 9–18 Image deconvolution. a light micrograph of the large polytene chromosomes from Drosophila, stained with a fluorescent Dna-binding dye. the same field of view after image deconvolution clearly reveals the banding pattern on the chromosomes. each band is about 0.25 μm thick, approaching the diffraction limit of the light microscope. (Courtesy of the John sedat Laboratory.)


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Cell_Biology_Alberts. Figure 9–19 The confocal fluorescence microscope. (a) this simplified diagram shows that the basic arrangement of optical components is similar to that of the in Figure 9–12, except that a laser is used to illuminate a small pinhole whose image is focused at a single point in the three-dimensional (3-D) specimen. (B) emitted fluorescence from this focal point in the specimen is focused at a second (confocal) pinhole. (C) emitted light from elsewhere in the specimen is not focused at the pinhole and therefore does not contribute to the final image. By scanning the beam of light across the specimen, a very sharp two-dimensional image of the exact plane of focus is built up that is not significantly point of focus fuorescent specimen is illuminated with a focused point of light from a pinhole far superior to those obtained by conventional light microscopy (Figure 9–20A and B).


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Cell_Biology_Alberts. The confocal microscope is generally used with fluorescence optics (see Figure 9–12), but instead of illuminating the whole specimen at once, in the usual way, the optical system at any instant focuses a spot of light onto a single point at a specific depth in the specimen. This requires a source of pinpoint illumination that is usually supplied by a laser whose light has been passed through a pinhole. The fluorescence emitted from the illuminated material is collected at a suitable light detector and used to generate an image. A pinhole aperture is placed in front of the detector, at a position that is confocal with the illuminating pinhole—that is, precisely where the rays emitted from the illuminated point in the specimen come to a focus. Thus, the light from this point in the specimen converges on this aperture and enters the detector.


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Cell_Biology_Alberts. By contrast, the light from regions out of the plane of focus of the spotlight is also out of focus at the pinhole aperture and is therefore largely excluded from the detector (see Figure 9–19). To build up a twodimensional image, data from each point in the plane of focus are collected sequentially by scanning across the field from left to right in a regular pattern of pixels and are displayed on a computer screen. Although not shown in Figure 9–19, the scanning is usually done by deflecting the beam with an oscillating mirror placed between the dichroic mirror and the objective lens in such a way that the illuminating spotlight and the degraded by light from other regions of the specimen. Figure 9–20 Confocal fluorescence microscopy produces clear optical sections and three-dimensional data sets. the first two micrographs are of the same intact gastrula-stage Drosophila embryo, which has been stained with a fluorescent probe for actin filaments.


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Cell_Biology_Alberts. (a) the conventional, unprocessed image is blurred by the presence of fluorescent structures above and below the plane of focus. (B) in the confocal image, this out-of-focus information is removed, resulting in a crisp optical section of the cells in the embryo. (C) a three-dimensional reconstruction of an object can be assembled from a stack of such optical sections. in this case, the complex branching structure of the mitochondrial compartment in a single live yeast cell is shown. (a and B, courtesy of richard Warn and peter shaw; C, courtesy of stefan hell.)


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Cell_Biology_Alberts. Figure 9–21 Multiphoton imaging. infrared laser light causes less damage to living cells than visible light and can also penetrate further, allowing microscopists to peer deeper into living tissues. the two-photon effect, in which a fluorochrome can be excited by two coincident infrared photons instead of a single high-energy photon, allows us to see nearly 0.5 mm inside the cortex of a live mouse brain. a dye, whose fluorescence changes with the calcium concentration, reveals active synapses (yellow) on the dendritic spines (red) that change as a function of time; in this case, there is a day between each image. (Courtesy of thomas oertner and karel svoboda.) confocal pinhole at the detector remain strictly in register. Variations in design now allow the rapid collection of data at video rates.


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Cell_Biology_Alberts. The confocal microscope has been used to resolve the structure of numerous complex threedimensional objects (Figure 9–20C) including the networks of cytoskeletal fibers in the cytoplasm and the arrangements of chromosomes and genes in the nucleus. The relative merits of deconvolution methods and confocal microscopy for threedimensional optical microscopy depend on the specimen being imaged. Confocal microscopes tend to be better for thicker specimens with high levels of outoffocus light. They are also generally easier to use than deconvolution systems and the final optical sections can be seen quickly. In contrast, the cooled CCD or CMOS cameras used for deconvolution systems are extremely efficient at collecting small amounts of light, and they can be used to make detailed threedimensional images from specimens that are too weakly stained or too easily damaged by the bright light used for confocal microscopy.


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Cell_Biology_Alberts. Both methods, however, have another drawback; neither is good at coping with very thick specimens. Deconvolution methods quickly become ineffective any deeper than about 40 μm into a specimen, while confocal microscopes can only obtain images up to a depth of about 150 μm. Special microscopes can now take advantage of the way in which fluorescent molecules are excited, to probe even deeper into a specimen. Fluorescent molecules are usually excited by a single highenergy photon, of shorter wavelength than the emitted light, but they can in addition be excited by the absorption of two (or more) photons of lower energy, as long as they both arrive within a femtosecond or so of each other. The use of this longerwavelength excitation has some important advantages. In addition to reducing background noise, red or nearinfrared light can penetrate deeper into a specimen. Multiphoton microscopes, constructed to take advantage of this two-photon effect, can obtain sharp images, sometimes even


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Cell_Biology_Alberts. noise, red or nearinfrared light can penetrate deeper into a specimen. Multiphoton microscopes, constructed to take advantage of this two-photon effect, can obtain sharp images, sometimes even at a depth of 250 μm within a specimen. This is particularly valuable for studies of living tissues, notably in imaging the dynamic activity of synapses and neurons just below the surface of living brains (Figure 9–21).


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Cell_Biology_Alberts. Even the most stable cell structures must be assembled, disassembled, and reorganized during the cell’s life cycle. Other structures, often enormous on the molecular scale, rapidly change, move, and reorganize themselves as the cell conducts its internal affairs and responds to its environment. Complex, highly organized pieces of molecular machinery move components around the cell, controlling traffic into and out of the nucleus, from one organelle to another, and into and out of the cell itself.


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Cell_Biology_Alberts. Various techniques have been developed to visualize the specific components involved in such dynamic phenomena. Many of these methods use fluorescent proteins, and they require a tradeoff between structural preservation and efficient labeling. All of the fluorescent molecules discussed so far are made outside the cell and then artificially introduced into it. But use of genes coding for protein molecules that are themselves inherently fluorescent also enables the creation of organisms and cell lines that make their own visible tags and labels, without the introduction of foreign molecules. These cellular exhibitionists display their inner workings in glowing fluorescent color.


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Cell_Biology_Alberts. Foremost among the fluorescent proteins used for these purposes by cell biologists is the green fluorescent protein (GFP), isolated from the jellyfish Aequorea victoria. This protein is encoded by a single gene, which can be cloned and introduced into cells of other species. The freshly translated protein is not fluorescent, but within an hour or so (less for some alleles of the gene, more for others) it undergoes a selfcatalyzed posttranslational modification to generate an efficient fluorochrome, shielded within the interior of a barrellike protein, which will now fluoresce when illuminated appropriately with blue light (Figure 9–22). Extensive sitedirected mutagenesis performed on the original gene sequence has resulted in multiple variants that can be used effectively in organisms ranging from animals and plants to fungi and microbes. The fluorescence efficiency has also been improved, and variants have been generated with altered absorption and emission spectra from the


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Cell_Biology_Alberts. ranging from animals and plants to fungi and microbes. The fluorescence efficiency has also been improved, and variants have been generated with altered absorption and emission spectra from the blue–green, like blue fluorescent protein or BFP, to the far visible red. Other, related fluorescent proteins have since been discovered (for example, in corals) that also extend the range into the red region of the spectrum, like red fluorescent protein or RFP.


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Cell_Biology_Alberts. One of the simplest uses of GFP is as a reporter molecule, a fluorescent probe to monitor gene expression. A transgenic organism can be made with the GFPcoding sequence placed under the transcriptional control of the promoter belonging to a gene of interest, giving a directly visible readout of the gene’s expression pattern in the living organism (Figure 9–23). In another application, a peptide location signal can be added to the GFP to direct it to a particular cell compartment, such as the endoplasmic reticulum or a mitochondrion, lighting up these organelles so they can be observed in the living state (see Figure 12–31).


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Cell_Biology_Alberts. The GFP DNA coding sequence can also be inserted at the beginning or end of the gene for another protein, yielding a chimeric product consisting of that protein with a GFP domain attached. In many cases, this GFP fusion protein behaves in the same way as the original protein, directly revealing its location and activities by means of its genetically encoded fluorescence (Figure 9–24). It is often possible to prove that the GFP fusion protein is functionally equivalent to the untagged protein, for example by using it to rescue a mutant lacking that protein. GFP tagging is the clearest and most unequivocal way of showing the distribution and dynamics of a protein in a living organism (Figure 9–25 and see Movie 16.8).


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Cell_Biology_Alberts. Fluorescent proteins are now exploited not just to see where in a cell a particular protein is located, but also to uncover its kinetic properties and to find out whether it might interact with other molecules. We now describe three techniques in which fluorescent proteins are used in this way. First, interactions between one protein and another can be monitored by fluorescence resonance energy transfer, also called Förster resonance energy


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Cell_Biology_Alberts. First, interactions between one protein and another can be monitored by fluorescence resonance energy transfer, also called Förster resonance energy Figure 9–23 Green fluorescent protein (GFP) as a reporter. For this experiment, carried out in the fruit fly, the gFp gene was joined (using recombinant Dna techniques) to a fly promoter that is active only in a specialized set of neurons. this image of a live fly embryo was captured by a fluorescence microscope and shows approximately 20 neurons, each with long projections (axons and dendrites) that communicate with other (nonfluorescent) cells. these neurons are located just under the surface of the animal and allow it to sense its immediate environment. (From W.B. grueber et al., Curr. Biol. 13:618–626, 2003. With permission from elsevier.)


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Cell_Biology_Alberts. Figure 9–22 Green fluorescent protein (GFP). the structure of gFp, shown here schematically, highlights the eleven β strands that form the staves of a barrel. Buried within the barrel is the active chromophore (dark green) that is formed post-translationally from the protruding side chains of three amino acid residues. (From M. ormö et al., Science 273:1392–1395, 1996. With permission from aaas.)


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Cell_Biology_Alberts. Figure 9–24 GFP-tagged proteins. this living cell from a tobacco plant is expressing high levels of green fluorescent protein, fused to a protein that is targeted to mitochondria, which accordingly appear green. the mitochondria are seen to cluster around the chloroplasts, whose chlorophyll autofluorescence marks them out in red. (Courtesy of olivier grandjean.) transfer, both abbreviated FRET. In this technique, two molecules of interest are each labeled with a different fluorochrome, chosen so that the emission spectrum of one fluorochrome, the donor, overlaps with the absorption spectrum of the other, the acceptor. If the two proteins bind so as to bring their fluorochromes into very close proximity (closer than about 5 nm), one fluorochrome, when excited, can transfer energy from the absorbed light directly (by resonance, nonradiatively) to the other. Thus, when the complex is illuminated at the excitation wavelength of the first fluorochrome, fluorescent light is produced at the


Cell_Biology_Alberts

Cell_Biology_Alberts. absorbed light directly (by resonance, nonradiatively) to the other. Thus, when the complex is illuminated at the excitation wavelength of the first fluorochrome, fluorescent light is produced at the emission wavelength of the second. This method can be used with two different spectral variants of GFP as fluorochromes to monitor processes such as the interaction of signaling molecules with their receptors, or proteins in macromolecular complexes at specific locations inside living cells (Figure 9–26). The FRET can be measured by quantifying the reduction of the donor fluorescence in the presence of the acceptor.


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Cell_Biology_Alberts. A second example of a fluorescencetagging technique that allows detailed observations of proteins within cells involves synthesizing an inactive form of the fluorescent molecule of interest, introducing it into the cell, and then activating it suddenly at a chosen site in the cell by focusing a spot of light on it. This process is referred to as photoactivation. Many inactive photosensitive precursors of this type, often called caged molecules, have been made based on a variety of fluorescent molecules. A microscope can be used to focus a strong pulse of light from a laser on any tiny region of the cell, so that the experimenter can control exactly where and when the fluorescent molecule is photoactivated. The technique allows us to follow complex and rapid intracellular processes, such as the actions of signaling molecules or the movements of cytoskeletal proteins.


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Cell_Biology_Alberts. When a photoactivatable fluorescent tag is attached to a purified protein, it is important that the modified protein remain biologically active: labeling with a caged fluorescent dye adds a bulky group to the surface of a protein, which can easily change the protein’s properties. A satisfactory labeling protocol is usually found by trial and error. Once a biologically active labeled protein has been produced, it needs to be introduced into the living cell where its behavior can be followed. Tubulin labeled with caged fluorescein, for example, can be injected into a dividing cell, where it is incorporated into microtubules of the mitotic spindle. When a small region of the spindle is illuminated with a laser, the labeled tubulin


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Cell_Biology_Alberts. Figure 9–25 Dynamics of GFP tagging. this sequence of micrographs shows a set of three-dimensional images of a living nucleus taken over the course of 135 minutes. tobacco cells have been stably transformed with gFp fused to a spliceosomal protein that is concentrated in small nuclear bodies called Cajal bodies (see Figure 6–46). the fluorescent Cajal bodies, easily visible in a living cell with confocal microscopy, are dynamic structures that move around within the nucleus. (Courtesy of kurt Boudonck, Liam Dolan, and peter shaw.) becomes fluorescent, so that its movement along the spindle microtubules can be readily followed (Figure 9–27).


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Cell_Biology_Alberts. A further development in photoactivation is the discovery that the genes encoding GFP and related fluorescent proteins can be engineered to produce protein variants, usually with one or more amino acid changes, that fluoresce only weakly under normal excitation conditions, but can be induced to fluoresce either more strongly or with a color shift (for example, from green to red) by activating them with a strong pulse of light at a different wavelength. In principle, the microscopist can then follow the local in vivo behavior of any protein that can be expressed as a fusion with one of these GFP variants. These genetically encoded, photoactivatable fluorescent proteins allow the lifetime and behavior of any protein to be studied independently of other newly synthesized proteins (Figure 9–28).


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Cell_Biology_Alberts. A third way to exploit GFP fused to a protein of interest is known as fluorescence recovery after photobleaching (FRAP). Here, one uses a strong focused beam of light from a laser to extinguish the GFP fluorescence in a specified region of the cell, after which one can analyze the way in which remaining unbleached fluorescent protein molecules move into the bleached area as a function of time.


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Cell_Biology_Alberts. Figure 9–26 Fluorescence resonance energy transfer (FRET). to determine whether (and when) two proteins interact inside a cell, the proteins are first produced as fusion proteins attached to different color variants of green fluorescent protein (gFp). (a) in this example, protein X is coupled to a blue fluorescent protein, which is excited by violet light (370–440 nm) and emits blue light (440–480 nm); protein Y is coupled to a green fluorescent protein, which is excited by blue light (440–480 nm) and emits green light (510 nm). (B) if protein X and Y do not interact, illuminating the sample with violet light yields fluorescence from the blue fluorescent protein only. (C) When protein X and protein Y interact, the resonance transfer of energy, Fret, can now occur. illuminating the sample with violet light excites the blue fluorescent protein, which transfers its energy to the green fluorescent protein, resulting in an emission of green light. the fluorochromes must be quite close


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Cell_Biology_Alberts. with violet light excites the blue fluorescent protein, which transfers its energy to the green fluorescent protein, resulting in an emission of green light. the fluorochromes must be quite close together—within about 1–5 nm of one another—for Fret to occur. Because not every molecule of protein X and protein Y is bound at all times, some blue light may still be detected. But as the two proteins begin to interact, emission from the donor blue fluorescent protein falls as the emission from the acceptor gFp rises.


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Cell_Biology_Alberts. Figure 9–27 Determining microtubule flux in the mitotic spindle with caged fluorescein linked to tubulin. a metaphase spindle formed in vitro from an extract of Xenopus eggs has incorporated three fluorescent markers: rhodamine-labeled tubulin (red) to mark all the microtubules, a blue Dna-binding dye that labels the chromosomes, and caged-fluorescein-labeled tubulin, which is also incorporated into all the microtubules but is invisible because it is nonfluorescent until activated by ultraviolet (UV) light.


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Cell_Biology_Alberts. a beam of UV light activates, or “uncages,” the caged-fluorescein-labeled tubulin locally, mainly just to the left side of the metaphase plate. over the next few minutes—after 1.5 minutes in (C) and after 2.5 minutes in (D)—the uncagedfluorescein–tubulin signal moves toward the left spindle pole, indicating that tubulin is continuously moving poleward even though the spindle (visualized by the red rhodamine-labeled tubulin fluorescence) remains largely unchanged. (From k.e. sawin and t.J. Mitchison, J. Cell Biol. 112:941–954, 1991. With permission from the rockefeller University press.) 546 Chapter 9: Visualizing Cells This technique is usually carried out with a confocal microscope and, like photoactivation, can deliver valuable quantitative data about a protein’s kinetic parameters, such as diffusion coefficients, active transport rates, or binding and dissociation rates from other proteins (Figure 9–29).


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Cell_Biology_Alberts. One way to study the chemistry of a single living cell is to insert the tip of a fine, glass, ionsensitive microelectrode directly into the cell interior through the plasma membrane. This technique is used to measure the intracellular concentrations of common inorganic ions, such as H+, Na+, K+, Cl–, and Ca2+. However, ionsensitive microelectrodes reveal the ion concentration only at one point in a cell, and for an ion present at a very low concentration, such as Ca2+, their responses are slow and somewhat erratic. Thus, these microelectrodes are not ideally suited to record the rapid and transient changes in the concentration of cytosolic Ca2+ that have an important role in allowing cells to respond to extracellular signals. Such changes can be analyzed with ion-sensitive indicators, whose light emission reflects the local concentration of the ion. Some of these indicators are luminescent (emitting light spontaneously), while others are fluorescent (emitting light on exposure to


Cell_Biology_Alberts

Cell_Biology_Alberts. whose light emission reflects the local concentration of the ion. Some of these indicators are luminescent (emitting light spontaneously), while others are fluorescent (emitting light on exposure to light).


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Cell_Biology_Alberts. Figure 9–28 Photoactivation. photoactivation is the light-induced activation of an inert molecule to an active state. in this experiment, shown schematically in (a), a photoactivatable variant of gFp is expressed in a cultured animal cell. (B) Before activation (time 0 sec), little or no gFp fluorescence is detected in the selected region (red circle) when excited by blue light at 488 nm. after activation of the gFp with an ultraviolet laser pulse at 413 nm, it rapidly fluoresces brightly in the selected region (green). the movement of gFp, as it diffuses out of this region, can be measured. since only the photoactivated proteins are fluorescent within the cell, the trafficking, turnover, and degradative pathways of proteins can be monitored. (B, from J. Lippincott-schwartz and g.h. patterson, Science 300:87–91, 2003.)


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Cell_Biology_Alberts. Figure 9–29 Fluorescence recovery after photobleaching (FRAP). a strong focused pulse of laser light will extinguish, or bleach, the fluorescence of gFp. By selectively photobleaching a set of fluorescently tagged protein molecules within a defined region of a cell, the microscopist can monitor recovery over time, as the remaining fluorescent molecules move into the bleached region (see Movie 10.6). (a) the experiment shown uses monkey cells in culture that express galactosyltransferase, an enzyme that constantly recycles between the golgi apparatus and the endoplasmic reticulum (er). the golgi apparatus in one of the two cells is selectively photobleached, while the production of new fluorescent protein is blocked by treating the cells with cycloheximide. the recovery, resulting from fluorescent enzyme molecules moving from the er to the golgi, can then be followed over a period of time. (B) schematic diagram of the experiment shown in (a). (a, from J. Lippincott-schwartz,


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Cell_Biology_Alberts. from fluorescent enzyme molecules moving from the er to the golgi, can then be followed over a period of time. (B) schematic diagram of the experiment shown in (a). (a, from J. Lippincott-schwartz, Histochem. Cell Biol. 116:97–107, 2001. With permission from springer-Verlag.) only molecules in the evanescent feld fuoresce


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Cell_Biology_Alberts. Figure 9–32 TIRF microscopy allows the detection of single fluorescent molecules. (a) tirF microscopy uses excitatory laser light to illuminate the cover-slip surface at the critical angle at which all the light is reflected by the glass–water interface. some electromagnetic energy extends a short distance across the interface as an evanescent wave that excites just those molecules that are attached to the cover slip or are very close to its surface. (B) tirF microscopy is used here to image individual myosin–gFp molecules (green dots) attached to nonfluorescent actin filaments (C), which are invisible but stuck to the surface of the cover slip. (Courtesy of Dmitry Cherny and Clive r. Bagshaw.) particular molecule of interest. This problem can be solved by the use of a special optical technique called total internal reflection fluorescence (TIRF) microscopy. In a TIRF microscope, laser light shines onto the coverslip surface at the precise critical angle at which total internal


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Cell_Biology_Alberts. technique called total internal reflection fluorescence (TIRF) microscopy. In a TIRF microscope, laser light shines onto the coverslip surface at the precise critical angle at which total internal reflection occurs (Figure 9–32A). Because of total internal reflection, the light does not enter the sample, and the majority of fluorescent molecules are not, therefore, illuminated. However, electromagnetic energy does extend, as an evanescent field, for a very short distance beyond the surface of the cover slip and into the specimen, allowing just those molecules in the layer closest to the surface to become excited. When these molecules fluoresce, their emitted light is no longer competing with outoffocus light from the overlying molecules, and can now be detected. TIRF has allowed several dramatic experiments, for instance imaging of single motor proteins moving along microtubules or single actin filaments forming and branching. At present, the technique is restricted to a thin layer


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Cell_Biology_Alberts. dramatic experiments, for instance imaging of single motor proteins moving along microtubules or single actin filaments forming and branching. At present, the technique is restricted to a thin layer within only 100–200 nm of the cell surface (Figure 9–32B and C).


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Cell_Biology_Alberts. individual Molecules Can Be touched, imaged, and Moved Using atomic Force Microscopy


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Cell_Biology_Alberts. While TIRF allows single molecules to be visualized under certain conditions, it is strictly a passive observation method. In order to probe molecular function, it is ultimately useful to be able to manipulate individual molecules themselves, and atomic force microscopy (AFM) provides a method to do just that. In an AFM device, an extremely small and sharply pointed tip, often of silicon or silicon nitride, is made using nanofabrication methods similar to those used in the semiconductor industry. The tip of the AFM probe is attached to a springy cantilever arm mounted on a highly precise positioning system that allows it to be moved over very small distances. In addition to this precise movement capability, the AFM device is able to collect information about a variety of forces that it encounters—including electrostatic, van der Waals, and mechanical forces—which are felt by its tip as it moves close to or touches the surface (Figure 9–33A). When AFM was first developed, it was


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Cell_Biology_Alberts. it encounters—including electrostatic, van der Waals, and mechanical forces—which are felt by its tip as it moves close to or touches the surface (Figure 9–33A). When AFM was first developed, it was intended as an imaging technology to measure molecularscale (B) (C) 1000 800 600 400 200 0 0 100 200 300 extension (nm) laser photodiode (detector) detector and feedback electronics springy cantilever arm AFM tip stretching molecule attached to substrate e.g., mica (A) 28 nm force (pN)


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Cell_Biology_Alberts. Figure 9–33 Single molecules can be imaged and manipulated by atomic force microscopy. (a) schematic diagram of the key components of an atomic force microscope (aFM), showing the force-sensing tip attached to one end of a single protein molecule, as in the experiment described in (D). (B) and (C) an aFM in imaging mode created these images of a single heteroduplex Dna molecule with a Muts protein dimer (larger white regions) bound near its center, at the point of a mismatched base pair. Muts is the first protein that binds to Dna when the mismatch repair process is initiated (see Figure 5–19). the smaller white dots are single streptavidin molecules, used to label the two ends of each Dna molecule. (D) titin is an enormous protein molecule that provides muscle with its passive elasticity (see Figure 16–34). the extensibility of this protein can be directly tested using a short, artificially produced protein that contains eight repeated immunoglobulin (ig) domains from one region of


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Cell_Biology_Alberts. (see Figure 16–34). the extensibility of this protein can be directly tested using a short, artificially produced protein that contains eight repeated immunoglobulin (ig) domains from one region of the titin protein. in this experiment, the tip of the aFM is used to pick up, and progressively stretch, a single molecule until it eventually ruptures. as force is applied, each ig domain suddenly begins to unfold, and the force needed in each case (about 200 pn) can be recorded. the region of the force–extension curve shaded green records the sequential unfolding event for each of the eight protein domains. (B and C, from Y. Jiang and p.e. Marszalek, EMBO J. 30:2881–2893, 2011. reprinted with permission of John Wiley & sons; D, adapted from W.a. Linke et al., J. Struct. Biol. 137:194–205, 2002. With permission from elsevier.) features on a surface. When used in this mode, the probe is scanned over the surface, moving up and down as necessary to maintain a constant interaction force with


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Cell_Biology_Alberts. With permission from elsevier.) features on a surface. When used in this mode, the probe is scanned over the surface, moving up and down as necessary to maintain a constant interaction force with the surface, thus revealing any objects such as proteins or other molecules that might be present on the otherwise flat surface (Figure 9–33B and C). AFM is not limited to simply imaging surfaces, however, and can also be used to pick up and move single molecules that adsorb strongly to the tip. Using this technology, the mechanical properties of individual protein molecules can be measured in detail. For example, AFM has been used to unfold a single protein molecule in order to measure the energetics of domain folding (Figure 9–33 D).


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Cell_Biology_Alberts. The variations on light microscopy we have described so far are all constrained by the classic diffraction limit to resolution described earlier; that is, to about 200 nm (see Figure 9–6). Yet many cellular structures—from nuclear pores to nucleosomes and clathrincoated pits—are much smaller than this and so are unresolvable by conventional light microscopy. Several approaches, however, are now available that bypass the limit imposed by the diffraction of light, and successfully allow objects as small as 20 nm to be imaged and clearly resolved: a remarkable, orderofmagnitude improvement.


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Cell_Biology_Alberts. The first of these socalled superresolution approaches, structured illumination microscopy (SIM), is a fluorescence imaging method with a resolution of about 100 nm, or twice the resolution of conventional brightfield and confocal microscopy. SIM overcomes the diffraction limit by using a grated or structured pattern of light to illuminate the sample. The microscope’s physical setup and operation is quite complex, but the general principle can be thought of as similar to creating a moiré pattern, an interference pattern created by overlaying two grids with different angles or mesh sizes (Figure 9–34). In a similar way to creating a moiré pattern, the illuminating grid and the sample features combine into an interference pattern, from which the original highresolution contributions to the image of features beyond the classical resolution limit can be calculated. Illumination by a grid means that the parts of the sample in the dark stripes of the grid are not illuminated and therefore


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Cell_Biology_Alberts. image of features beyond the classical resolution limit can be calculated. Illumination by a grid means that the parts of the sample in the dark stripes of the grid are not illuminated and therefore not imaged, so the imaging is repeated several times (usually three) after translating the grid through a fraction of the grid spacing between each image. As the interference effect is strongest for image components close to the direction of the grid bars, the whole process is repeated with the grid pattern rotated through a series of angles to obtain an equivalent enhancement in all directions. Finally, mathematically combining all these separate images by computer creates an enhanced superresolution image. SIM is versatile because it can be used with any fluorescent dye or protein, and combining SIM images captured at consecutive focal planes can create threedimensional data sets (Figure 9–35).


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Cell_Biology_Alberts. Figure 9–34 Structured illumination microscopy. the principle, illustrated here, is to illuminate a sample with patterned light and measure the moiré pattern. shown are (a) the pattern from an unknown structure and (B) a known pattern. (C) When these are combined, the resulting moiré pattern contains more information than is easily seen in (a), the original pattern. if the known pattern (B) has higher spatial frequencies, then better resolution will result. however, because the spatial patterns that can be created optically are also diffraction-limited, siM can only improve the resolution by about a factor of two. (From B.o. Leung and k.C. Chou, Appl. Spectrosc. 65:967–980, 2011.)


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Cell_Biology_Alberts. Figure 9–35 Structured illumination microscopy can be used to create three-dimensional data. these three-dimensional projections of the meiotic chromosomes at pachytene in a maize cell show the paired lateral elements of the synaptonemal complexes. (a) the chromosome set has been stained with a fluorescent antibody to cohesin and is viewed here by conventional fluorescence microscopy. Because the distance between the two lateral elements is about 200 nm, the diffraction limit, the two lateral elements that make up each complex are not resolved. (B) in the three-dimensional siM image, the improved resolution enables each lateral element, about 100 nm across, to be clearly resolved, and the two chromosomes can clearly be seen to coil around each other. (C) Because the complete three-dimensional data set for the whole nucleus is available, the path of each separate pair of chromosomes can be traced and artificially assigned a different color. (Courtesy of C.J. rachel Wang, peter Carlton


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Cell_Biology_Alberts. data set for the whole nucleus is available, the path of each separate pair of chromosomes can be traced and artificially assigned a different color. (Courtesy of C.J. rachel Wang, peter Carlton and Zacheus Cande.) approximates a gaussian distribution, whose width at half-maximum height under ideal conditions is about 200 nm. (C) two point sources that are about 200 nm apart


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Cell_Biology_Alberts. To get around the diffraction limit, the other two superresolutiontechniques exploit aspects of the point spread function, a property of the optical system mentioned earlier. The point spread function is the distribution of light intensity within the threedimensional, blurred image that is formed when a single point source of light is brought to a focus with a lens. Instead of being identical to the point source, the image has an intensity distribution that is approximately described by a Gaussian distribution, which in turn determines the resolution of the lens system (Figure 9–36). Two points that are closer than the width at halfmaximum height of this distribution will become hard to resolve because their images overlap too much (see Figure 9–36C).


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Cell_Biology_Alberts. In fluorescence microscopy, the excitation light is focused to a spot on the specimen by the objective lens, which then captures the photons emitted by any fluorescent molecule that the beam has raised from a ground state to an excited state. Because the excitation spot is blurred according to the point spread function, fluorescent molecules that are closer than about 200 nm will be imaged as a single blurred spot. One approach to increasing the resolution is to switch all the fluorescent molecules at the periphery of the blurry excitation spot back to their ground state, or to a state where they no longer fluoresce in the normal way, leaving only those at the very center to be recorded. This can be done in practice by adding a second, very bright laser beam that wraps around the excitation beam like a torus. The wavelength and intensity of this second beam are adjusted so as to switch the fluorescent molecules off everywhere except at the very center of the point spread function, a


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Cell_Biology_Alberts. beam like a torus. The wavelength and intensity of this second beam are adjusted so as to switch the fluorescent molecules off everywhere except at the very center of the point spread function, a region that can be as small as 20 nm across (Figure 9–37). The fluorescent probes used must be in a special class that is photoswitchable: their emission can be reversibly switched on and off with lights of different wavelengths. As the specimen is scanned with this arrangement of lasers, fluorescent molecules are switched on and off, and the small point spread function at each location is recorded. The diffraction limit is breached because the technique ensures that similar but very closely spaced molecules are in one of two different states, either fluorescing or dark. This approach is called STED (stimulated emission depletion microscopy) and various microscopes using versions of the general method are now in wide use. Resolutions of 20 nm have been achieved in biological specimens, and


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Cell_Biology_Alberts. STED (stimulated emission depletion microscopy) and various microscopes using versions of the general method are now in wide use. Resolutions of 20 nm have been achieved in biological specimens, and even higher resolution attained with nonbiological specimens (see Figure 9–37).


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Cell_Biology_Alberts. If a single fluorescent molecule is imaged, it appears as a circular blurry disc, but if sufficient photons have contributed to this image, the precise mathematical center of the disclike image can be determined very accurately, often to within a few nanometers. But the problem with a specimen that contains a large number