Cell Biology: Origin and Evolution of Cells PDF

Scanning electron micrograph of E. coli, grown in culture and adhered to a slide. Under optimal culture conditions, E. coli divides every 20 min....

Scanning electron micrograph of E. coli, grown in culture and adhered to a slide. Under optimal culture conditions, E. coli divides every 20 min. Rapid growth medium: glucose, salts, and various organic compounds, such as amino acids, vitamins, and nucleic acid precursors. Slow growing medium: salts, a nitrogen source (such as ammonia), and a carbon and energy E. coli colonies growing on the source (such as glucose). surface of an agar medium. 41  It is a multicellular organism in the shape of an elongated tube that thins at the ends. Simple multicellular organism: 959 somatic cells, and 1000 to 2000 germ cells. It can be easily reproduced and subjected to genetic manipulation in the laboratory. The whole body is covered by a fine outer cuticle. Cells are organized into fairly simple organs and systems. It has a digestive system made up of a stoma (or mouth), pharynx, and intestine. It also has sexual organs (gonads) and a rudimentary nervous system. Caenorhabditis elegans 51  First multicellular organism to have its genome sequenced (1998). The C. elegans genome is approximately 100 million (108) base pairs long and consists of six chromosomes and one mitochondrial genome. The genome contains an estimated 19,000 protein-coding genes. 36% of the genes of C. elegans have human homologues. 53  Danio rerio The zebrafish (Danio rerio) is a small and active fish and very common in aquariums. Its natural habitat is more or less calm waters, sometimes stagnant, of central Asia, particularly of the region of the Ganges in India. Danio rerio Adult individuals are usually between 3 and 5 cm long and 1 cm (Zebrafisk) wide depending on environmental conditions. It has an elongated shape with a dorsal fin. On the sides it has between 5 and 9 bluish bands that overlap the background color, which is golden in males and silver in females. This striped look has earned it the popular name of zebrafish. The ventral area is whitish and pink. 62 72 Heydari Z, Moeinvaziri F, Agarwal T, et al. Organoids: a novel modality in disease modeling [published online ahead of print, 2021 Aug 9]. Biodes Manuf. 2021;1-28. 73 Summary of generated human organoid disease models INDEX 1.1. Origin and evolution of cells 1.2. Cells as experimental models 1.3. Cell biology instruments 77 1.3. Cell Biology Instruments (a) Optical microscopy Electron microscopy Super-Resolution Microscopy 78 Light microscope Transmission electron microscope 79  Magnification: the factor by which an image appears to be enlarged. o Light microscope: 1000 X o Electron microscope: 150.000-200.000 X Resolving power: ability of the objective lens to distinguish as separate and different two points very close to each other. o Human eye: 0,1 mm. o Light microscope: 0,2 mm. o Transmission electron microscope: 1-2 nm (10-20 ángstrom (Å)) 80 82  Frog egg (naked eye) 83  Epithelial tissue (Optical microscopy) 84  Ribosome Transcription machinery (electron microscopy) Polypeptide chain mRNA molecule 85  Conventional light microscopy The conventional optical microscope is made up of three systems: Mechanical: foot (base), column (arm), stage (sample holder) and tube (head). Optical: Objectives and eyepieces Lighting: spotlight, condenser and diaphragm. Magnification and resolution Magnification: About 1000 times Most cells can be seen Resolution: 0.2 mm It does not allow to observe small details of the cellular structure    = wavelength Resolution = NA NA = Numerical Aperture  visible light = 0,5 mm NA = n * sen a n = refractive index of the medium (1,0 for air. Max. 1,4 for immersion oil) a = the maximal half-angle of the cone of light that can enter    Resolution the lens limit = = 0,22 mm amax = 90 ͦ 1,4 sen amax = 1 Types of optical microscopy Light field microscope Phase contrast microscope Differential interference-contrast microscope Fluorescence microscope Confocal microscope Multiphoton microscope  Light field microscope The light passes directly through the cell. Ability to distinguish different cell parts depends on the contrast obtained from the absorption of visible light by cellular components. Fixation: Stabilize and preserve cell structures (alcohol, acetic acid, formaldehyde...) Staining: Highlight Contrast Light field micrograph of stained tissue. Unsuitable techniques for observing Section of a benign kidney tumor. living cells.  Phase contrast microscope Allows us to view live samples without the need for staining techniques. It manipulates the light in such a way that it is possible to increase the contrast of the observed sample, allowing to observe structures that are invisible through a conventional microscope. The phase contrast microscope works by increasing the contrast of the phases of the waves reaching the objective. Small phase differences turn into intensity changes. Phase: measurement of the wave's position relative to a reference point. Yeast observed in the phase contrast microscope  Differential Interference Contrast (DIC) Nomarski interference contrast (NIC) or Nomarski microscopy Similar to phase contrast, DIC microscopy is a contrast- enhancing technique. DIC uses polarized light and prisms to convert phase delays into intensity changes (contrast). It is useful for rendering contrast in transparent samples and gives brilliant pseudo-3D relief shading images. DIC is used to image live, unstained biological samples. High sensitivity to thin cellular material. Although DIC images look very appealing, the pseudo-3D effect might be misleading in some cases, making it seem that the cells have structures that they do not have. As an example, areas inside a living cell with a different refractive index, like vacuoles and chromatin, appear as DIC microscopy of Sacharomyces bumps, which is actually an optical impression. cerevisiae.  Light field microscope (A) Phase contrast microscope (B) Differential interference- contrast microscope (C) Microscopic observation of living cells. Photomicrographs of human buccal cells.  Fluorescence Microscopy Fluorescence microscopy is based on the property of fluorescent substances to absorb light from a certain , and emit light from another . To see structures that have been selectively labeled with fluorescent substances (fluorochromes) and for the study of the intracellular distribution of molecules in living or fixed cells. Almost any celular component can be “stained” and thereby specifically imaged. - Nucleic acids. Specific genes or RNA transcripts can be detected by hybridization with complementary probes labeled with fluorescent markers. - Proteins. They can be detected using labeled antibodies that specifically recognize those proteins that we want to study, or by fusing with fluorescent proteins (such as GFP).  Widefield Fluorescence Microscopy The light passes through the excitation filter to select the light from a  (eg. blue) that excites the dye. Then a dichroic mirror deflects the excited light towards the sample. The light emitted by the sample (e.g green) passes through the mirror and a barrier filter that selects the  emitted by the fluorescent dye. Fluorescent micrograph of a newt lung cell in which the DNA is stained blue and the microtubules in the cytoplasm green. Fluorescent dyes can be used to stain specific cell components, such as acridine orange, which selectively stains the nucleus. https://www.thermofisher.com/es/es/home/life-science/cell-analysis/cell-structure.html A culture of BPAE (Bovine Pulmonary Artery Endothelial Cells) cells was stained with MitoTracker Red CMXRos, vividly labeling the intracellular mitochondrial network. The specimen was also labeled for filamentous actin and DNA with Alexa Fluor 488 (green emission) conjugated to phalloidin and DAPI (blue emission), respectively.  Green fluorescent protein (GFP) It was isolated from the Aequorea victoria jellyfish. Revolutionary breakthrough in fluorescence microscopy. GFP can be fused to any protein of interest by recombinant DNA technology. To visualize proteins inside living cells. It does not require fixation or staining, unlike the use of antibodies. Cover of the February 11, 1994 issue of the journal Science (from Chalfie M, et al., Green fluorescent protein as a marker for gene expression, Science, 1994. 263. p. 802). It was the beginning of the GFP revolution. The cover image showed green-glowing sensory neurons in C. elegans. FRAP: fluorescence recovery after photobleaching Method to follow the movement of GFP-labeled proteins in living cells. A region of interest in a cell that expresses a GFP- tagged protein is bleached by exposure to high intensity light. Fluorescence recovers over time due to the movement of unbleached GFP-labeled molecules into the bleached region. The rate of fluorescence recovery therefore provides a measure of the rate of protein movement within the cell. FRET: fluorescence resonance energy transfer The two proteins bind to different fluorescent markers, such as two GFP variants (A, B) that absorb and emit light at different , so that the light emitted by the first excites the second. The interaction between two proteins can be detected by illuminating the cell with a light of  that excites the first GFP variant and analyzing the  of the emitted light. If there is interaction, the light emitted by A will excite B by spatial closeness, resulting in the emission of light at a  typical of B.  Confocal Microscopy It enables the acquisition of images of increased detail by analyzing the fluorescence of a single plane of the sample. Structures within thicker objects can be conveniently visualized. Instead of illuminating the whole sample at once, laser light is focused onto a defined spot at a specific depth within the sample. This leads to the emission of fluorescent light at exactly this point. A pinhole inside the optical pathway cuts off signals that are out of focus, thus allowing only the fluorescence signals from the illuminated spot to enter the light detector. By scanning the specimen in a raster pattern, images of one single optical plane are created. 3D objects can be visualized by scanning several optical planes and stacking them using a suitable microscopy deconvolution software (z-stack). It is also possible to analyze multicolor immunofluorescence stainings using state-of-the-art confocal microscopes that include several lasers and emission/excitation filters.  Multiphoton Microscopy The sample is illuminated with a light of one  such that the excitation of the fluorescent dye requires the simultaneous absorption of two or more photons, which only happens at the point of the sample where the laser is focused. Thus, fluorescence is only emitted from the focus plane of the light.  Electron microscopy Due to the limited resolution of the light microscope, the analysis of the details of the cell structure has necessitated this much more powerful technique. Developed in 1930. First applied to biological samples in 1940-1950 by Albert Claude, Keith Porter, and George Palade. The electron microscope uses an accelerated electron beam as a 'illumination' source. The  of electrons (up to 0.004 nm) is much lower than that of visible light, which allows greater resolution, allowing the observation of cellular organelles. The theoretical maximum resolution is 0.002 nm, but it has not been achieved, as it also depends on the numerical aperture of the objective lens, whose aperture angle is limited to 0.5 degrees (AN = 0.01). Resolution limit = 0.2 nm. In biological samples, the lack of contrast limits the resolution to 1-2 nm.  Transmission electron microscopy (TEM) It consists of a cathode ray tube (electron streams in vacuum tubes), approx. 1 m long. At the top is the cathode, which emits a beam of electrons, through a potential difference, towards a perforated anode that is lower. The beam passes through the anode and is directed to the basal part of the column. On its way, the beam is first modified by an electromagnetic coil that acts as a condenser, focusing the electrons towards the sample. The beam then passes through the sample and is then modified by a coil that acts as an objective and another that serves as a projection system. In order to conduct the electrons, a system of This image is obtained on a fluorescent screen. pumps that generate vacuum is necessary, so it is not possible to carry out in vivo studies.  In order for the electron beam to pass through the sample, it must be very thin (10-100 nm). The contrast in TEM is achieved by differential absorption of heavy metals in the form of salts, which bind to biological structures. Typical contrast substances: lead citrate, osmium tetroxide, and uranium acetate. Structures that have a higher affinity for these substances partially impede the passage of electrons (the electrons collide with the heavy metal ions and are reflected, so they do not contribute to the final image), so it is seen on the screen as a dark area (electrodense areas). The areas that allow a greater passage of electrons are seen as light areas (electrolucent areas).  Scanning electron microscopy (SEM) It is used to view tissue and cell surfaces in 3D at high magnification and resolution. The samples are placed at the base of the microscope. They have been covered with a heavy metal, so the electrons do not pass through them, but rather they will be swept by the electrons in the three directions of space, giving a three-dimensional image. Electrons emitted from the sample surface are collected by a sensor and projected onto a television monitor. The display aspect is metallic and three- dimensional. Pollen grains Blood cells More information: https://ibidi.com/content/cat egory/127-microscopy- techniques 1.3. Cell Biology Instruments (b) Specimen preparation Flow citometry Subcellular separation Growth of animal cells in culture Virus BLOQUE 1: INTRODUCCIÓN. UNIDAD DIDÁCTICA 1 111 Specimen preparation Histological processing consists of four steps: a. Fixation b. Tissue embedding c. Sectioning d. Staining FIXATION To stop the degradation processes that occur after cell death (autolysis). It preserves the structures and chemical components of the cells and gives the sample mechanical consistency (hardness), allowing subsequent manipulations. a) Physical: cryofixation. The tissue is exposed to temperatures below -70ºC, decreasing enzymatic lysis activity and immobilizing cell structures. It is a reversible method, which stops working when the temperature increases. it can cause the formation of ice crystals that damage cell structure. Cryoprotective substances, such as sucrose or glycerol, prevent the formation of these crystals. Very fast fixation method, very useful in the processing of urgent biopsies. b) Chemicals: uses chemical substances that interact with cellular macromolecules, stabilizing them.  TISSUE EMBEDDING Objective: to remove the water from the tissues and replace it with a medium that solidifies, allowing subsequent cutting. It provides structural support to the piece, which makes it possible to obtain fine cuts to allow the passage of light or electrons.. dehydration: using increasing concentrations of ethanol solidification: paraffin is used for optical microscopy and resins for EM. Paraffin: mixture of waxes Resins: they are harder than composed of long-chain paraffin, so they allow finer hydrocarbons with a melting cuts. They are liquid point of 50ºC. The sample is substances in monomeric immersed in liquid paraffin form, and polymerize after (contained in a mold) that treatment with heat or UV infiltrates the intra- and light, giving rise to solid extracellular spaces of the polymers of great hardness. tissue. SECTIONING Histological sections are made in a microtome (between 3 and 5 mm, OM) or ultramicrotome (between 10-100 mm, EM). Paraffin sections are collected on glass slides and TEM sections on copper or nickel grids. The sections of the freeze-fixed samples are thicker (6-8 mm minimum), and are made in a cryostat, which maintains the samples at a temperature of -20 ° C during the section. These samples do not require the dehydration step.  There are samples that do not require fixing or sectioning steps, but the visualization is done directly. For example, cells in liquid medium (blood smear) or samples of various biological fluids, such as sputum or bronchioalveolar lavage. Extensions are made directly on the slide. STAINING In OM, dyes with different affinity for cell organelles are used, depending on their chemical composition. Examples of basic dyes: are thionin, safranin, toluidine blue, methylene blue, hematoxylin. They have a craving for acidic substances in tissue such as DNA, thus revealing the cell nucleus. Examples of acid dyes are acid fuchsin, quick green, orange G, or eosin. They have a craving for basic substances, especially protein structures located in the cell cytoplasm and also for the collagen of the extracellular matrix. One of the most commonly used stains in histology is hematoxylin-eosin. A basic dye (hematoxylin) and another acid (eosin) are used to stain the acidic and basic structures of the cell differently. In EM, the contrast is performed by means of high molecular weight molecules that enhance the dispersion capacity of the sample (uranium acetate and lead citrate).  Sample preparation for light microscopy: https://www.youtube.com/watch?v=4DJm4NLECQs Sample preparation for electron microscopy: https://www.youtube.com/watch?v=Ad5VGbA-_vk IMMUNOHISTOCHEMICAL TECHNIQUES Purpose: to identify the presence of a specific protein in a tissue section by using specific antibodies. Immunohistochemistry is essential in pathological anatomy laboratories, due to its ability to help in the diagnosis of diseases. Example: identification of viral proteins, overexpression of certain oncogenes, etc.  Immunohistochemical techniques can be divided into two large groups: Direct: The antibody that recognizes the antigen of interest (primary antibody) is conjugated to a substance that may be detectable under a microscope (an enzyme, a fluorochrome, or a gold particle). Indirect: the primary antibody is not conjugated, but a secondary antibody is used, which recognizes the primary one, and which in turn is conjugated with a substance visible under the microscope. Therefore, the signal will come from the secondary antibody. Indirect methods are the most widely used, since the antibodies used can recognize the Fc fraction of a given species, and use the same antibody for many primary antibodies, regardless of the antigen they recognize.  Flow cytometry Flow cytometry is a technique for analyzing the number, size and complexity of a cell suspension. It is also used to quantify the number of cells in certain cell populations specifically labeled with fluorescent substances and to isolate cell populations. Especially useful for the study of normal and pathological populations of blood cells and bone marrow. The cytometer has the following components: ✓ Fluidic sample transport system. ✓ Optical laser illumination system. ✓ Electronic detector, which converts light into digital signal. The cells pass in a very fine capillary so that they pass one by one.  As the cells pass, the beam hits them and a beam distortion is produced, which is what is registered by the detector. This system quantifies the number of events (cells) that are impacted by the laser, but it is also capable of analyzing the shape and internal complexity of the cell.. Visible light scatter is measured in two different directions, the forward direction (Forward Scatter or FSC) which can indicate the relative size of the cell and at 90° (Side Scatter or SSC) which indicates the internal complexity or granularity of the cell. COMPLEXITY SIZE  neutrophil neutrophils monocytes lymphocyte monocyte lymphocytes Size 125 Subcellular separation To study the chemical composition and function of certain organelles and other subcellular particles, it is convenient to separate them from the rest of the cytoplasmic and nuclear components. The first thing to do is to break the plasma membrane, which will make it possible to obtain a suspension with all the subcellular components. There are various methods of breaking down cells. Most used physical methods: osmotic shock, ultrasound, mechanical grinding. These procedures break down most cell membranes (plasma membrane and endoplasmic reticulum membranes) Enzymatic methods, usually lysozyme is used. When cells have a wall (plant or bacterial cells). DIFFERENTIAL CENTRIFUGATION A typical homogenate or lysate is made up of a suspension of organelles, cellular components and macromolecules that can be separated by centrifugation. An ultracentrifuge processes the samples at high speed (100,000 rpm) to produce forces around 500,000 times greater than gravity. Cellular components settle to the bottom of the centrifuge tube forming a precipitate (sedimentation) to a degree that depends on size and density: larger and heavier structures settle more quickly. Centrifuges are equipment that generate this type of force and have a rotor (where the samples are placed) that rotates at very high speeds. DENSITY GRADIENT CENTRIFUGATION It allows to obtain a higher degree of purification. The organelles are separated by sedimentation as a function of the gradient of a dense substance, such as sucrose or cesium chloride. Speed separations: Particles of different sizes settle down the gradient at different scales, moving as bands. The separated particles can be obtained in individual fractions  Equilibrium separations: The gradient is very concentrated. Subcellular components separate regardless of their size and shape, according to their density. The particle does not move when its density is equal to that of the medium (equilibrium position).  Centrifugation also allows cells to be separated, for which gradients such as Ficoll, Percol or Nicodenz are used. The separation of Peripheral Blood Mononuclear Cells (PBMC) may be required for certain applications including flow cytometry. Cell cultures The study of cells in a living organism is very difficult. An alternative is to isolate and maintain the cells in the laboratory under conditions that allow their survival and growth, a procedure that is known by the term cell culture. Although the process is technically much more difficult than the culture of bacteria and yeast, a wide variety of animal and plant cells can be cultured and manipulated in vitro. Cultured cells have several advantages over intact organisms for research: the experimental conditions can be better controlled, in many cases a single cell can easily grow into a colony of many identical cells. The resulting strain of cells, which is genetically homogeneous, is called a clone. ANIMAL CULTURES They are started by dispersing a part of tissue in a suspension of its cellular components. The cells are added to a culture dish containing a nutrient medium. Primary cultures: initial cultures of cells established from a culture. The cells grow to cover the surface of the culture dish. They can then be removed from the plate and replenished at low density to form secondary cultures. This process can be repeated many times, although most normal cells cannot grow indefinitely in culture.E. g.: human fibroblasts tolerate 50-100 passages.  In contrast, cells that are derived from tumors often proliferate indefinitely in culture and are called immortal cells. Continuous and uniform source of cells that can be manipulated and cultivated indefinitely. In humans it was obtained for the first time in 1951: HeLa cells, obtained from a biopsy of Henrietta Lacks' cervical tumor. Embryonic stem cells. They are established from early embryos. Ability to differentiate into all cell types of the adult organism. Developmental gene function studies. HeLa cells Dr. George Gey, in his cancer research, spent 30 years trying to grow cancer cells in the laboratory. On February 1, 1951, Henrietta Lacks was taken to John Hopkins Hospital. She had a cervical tumor. Cells from part of her tumor were retained in the hospital's cancer unit as Gey had discovered that they could be grown in the laboratory indefinitely. It was what he had sought for so many years. For the first time anything could be tested in living human cells. One of the first uses was the development of the vaccine against the polio virus. At present they continue to be widely used in human cell and molecular biology.  One of the most used tools in the field of cell biology. Cells grown in vitro come from a normal or tumor organ or tissue. These cells are kept in culture media of known composition and under controlled oxygen and temperature conditions (usually 37°C, 5% CO2 and 95% O2). The necessary culture medium is complex (salts, glucose, aa and vitamins). It usually also includes serum, a source of polypeptide growth factors necessary for cell division. Identification of specific growth factors for specific cell types eliminates serum necessity. VIRUSES Viruses need a host in which to carry out their life cycle. To grow viruses in the laboratory, it is essential to grow an animal, plant, fungal or bacterial cell culture, depending on what infects the virus. The host cells will enable the virus to increase its numbers. They provide simple systems for studying the functions of cells. Studies with viruses have revealed fundamental aspects of cell biology: molecular genetic mechanisms, RNA genetic potential, discovery of oncogenes...