Cell Technology 23/24 Lecture Notes PDF

Cell Technology – Sottile 23/24 LECTURE 1 – 4/10 File 1 >CELL SIGNALLING PRINCIPLES Cell phenotype is defined by the expression of genes into mRNA and into protein. Different cell type have different gene expression profile and thus also a different protein expression profile. During differentiat...

Cell Technology – Sottile 23/24 LECTURE 1 – 4/10 File 1 >CELL SIGNALLING PRINCIPLES Cell phenotype is defined by the expression of genes into mRNA and into protein. Different cell type have different gene expression profile and thus also a different protein expression profile. During differentiation cells differentiate and become unique types of cells with unique profiles. Many genes and signalling principles are conserved in different species. Development can be divided in 2 aspects: growth and patterning. Patterning gives us the rules which are coordinated by the cell signalling. Development is controlled and coordinated by signalling between different cell types with different mechanisms such as: Cell proliferation Cell specialization Cell interaction Cell movement In the first fertilized egg (zygote) there are 3 gem layers ->3 main tissue types that give life to all other tissues: mesoderm, endoderm e ectoderm. >GENERATING CELL DIVERSITY: 1. Model: Cell lineage model - diversity from non symmetrical-separation Cells acquire diverse fates while still at the precursor stage, relying primarily on information intrinsic to each cell. From the uncommitted progenitor cell we get 2 daughter cells. The first cleavage produces segregation of determinants. The more division there are the more diverse the cell become. This is because the content inside of the cell is never divided exactly in the middle. Types of content can be mRNA, cytokines, etc that give the cell a gradient that causes different identities/more diversity on the long run.  2. Model: diversity from specific signals from other cells/surrounding When cells divide, they are not physically on the same spot. The surrounding of the cell will give stimuli with factors that can influence the behaviour of the cell. You could also have a single stimulus that forms a gradient capable of inducing differentiation in different cells. 3. Flag model All cells have the same signalling potential. There is a signalling gradient because cells are exposed to different concentration of the signalling molecules. The different levels of signalling trigger different gene expression. Morphogens are one way of generating cell diversity. 4. Direct morphogen gradient Localized production of an inducer diffusing from its source forming a gradient. 5. Indirect morphogen gradient Localized production of an inhibitor diffusing away from its source and blocking the action of a uniformly distributed inducer. So one induces and is uniformly distributed. On one end there is an inhibitor that is distributed in gradient. The result is that you have a resulting gradient in the inducer activity not because it has only once source but because the inhibitor inhibits the inducer 6. Patterning by sequential induction - direct contact of different cell types Inductive interaction can generate many cell types from only a few. There are 2 cells aggregate types in 2 different spaces (A and B). The cells on the border of these 2 aggregates will form an intermediate zone (C. because they act on each other with signals. Now cells in zone C can interact with A and B and influence the neighbouring cells forming more unique compartments (D, E). This creates diversity.  7. Cutting tissue By cutting tissue and removing it from the "normal" ambient of the neighbouring cells one can achieve differentiation. Cutting endoderm from zygote will generate endoderm and then mesoderm. Cutting exoderm from zygote will generate epidermis cells. 8. Spinal cord patterning One can control the spinal cord pattern by Shh/BMP signals. BMP signals come from the dorsal side (roof plate) of the embryo. Shh (sonic hedgehog) signals come from the ventral side (floor plate) of the spinal tube. Depending of the gradient of BMP and Shh the cell receives the cells in the spinal cord will differentiate in the different types of neurons. >DIFFERENT WAYS OF TRANSMITTING SIGNALS 1. Diffusion of molecules or electric signals via pores and receptors 2. Direct contact between 2 cells 3. Gap junction >TYPE OF SIGNALLING: 1. Receptor-mediated signalling - Intracellular signalling cascade An extracellular molecule interacts with a receptor protein and determines an intracellular signal via signalling cascades. The intracellular signal will go to the target proteins which can be enzymes, regulatory proteins or cytoskeletal proteins. The modification are then related to the metabolism, alteration of the gene expression and/or altering the shape of movement of cells. From one binding event we can form a strong signalling event -> signal amplification. - Example: MAPK pathways. In this pathway there are sequential phosphorylation starting from a stimulus which trigger a signalling cascade that leads to cellular responses The binding of one ligand can not only cause an activation but also repress the signalling inducing signal silencing >SIGNALLING PATHWAYS: (STUDY WELL) 1. Wnt pathway - ßcatenin pathway 2 main receptors on cell membrane: LRP and Frizzled ß-catenin is trapped in a complex when Wnt is not there -> the system is off. If Wnt is present in the surrounding is it binds to the receptors and it activated the pathway. Now the ß-catenin is released from the complex and moves near the nucleus where it modifies the transcription of Wnt-responsive genes. It has an ON/OFF system. It determines the body axis in normal development. It is related to the development of the mid-brain and also in haematopoiesis. 2. TGF-ß pathway (BMPs/GDF/…) 2 types of receptors that only bind when the ligand is present. These 2 dimerize and trigger the phosphorylation of one another which triggers the phosphorylation signalling cascade. The cascade is initiated by Smad2 and Co-Smad2. Smad2/3 oligomerize with Smad4 trigger then the transcription of target genes. Important for the left-right asymmetry of the body, formation of the skeleton. BMP tends to rely more on Smad1,5,8 -> in this way although the mechanisms are similar the outcome will be different and the signal of these different pathways will not get lost. BMP is important for neuronal diversity in early development. 3. Hedgehog pathway Sonic hedgehog (Shh), Indian hedgehog (Ihh), desert hedgehog (Dhh). ON/OFF signalling. 2 receptors one patched protein and one smoothened protein which are bound together when there is no ligand. The smoothened protein will be "silenced" by the patched protein. In this case the Ci protein (Gli in vertebrated) is attached to microtubules. This allows PKA and Slimb proteins to cleave Ci into a transcriptional repressor blocking transcription of target genes. When the ligand (hedgehog) binds the conformation changes and releases the smoothened protein which can now release Ci from the microtubules and inactivates the cleavage protein PKA and Slimb. Ci enters the nucleus and acts as a transcriptional activator for target genes. This signalling is important for inducing the floor plate of neural tubes, develop anteroposterior axis and the left-right symmetry.  4. Receptor tyrosine kinase pathway All receptors have in common a intracellular tyrosine kinase component on the intracellular part. A phosphorylation cascade is caused. They are present in many different cells such as the insulin receptor, FGFR1, EGFR, a. PDGF We have dimerized receptors which dimerize when PDGF is present. b. EPH c. FGF The FGF molecules are often bound to something else like glcosaminoglycan chains. So 2 of them bind to each receptor. Receptor then cross- phosphorylate each other and phosphorylate the Ras protein. Then Ras -> Raf -> Mek -> Mapk -> activate genes. This pathway is related with cancer when Ras/Raf are mutated because they initiate the phosphorylation cascades on their own. Mutations in the FGF pathway lead to developmental abnormalities 4. Notch/delta pathway Involved cell-cell contact. It is important in generating specific lineages that have to coordinate in the individual. Cell gaining an advantage deliver strong inhibition to its neighbours. Neighbouring inhibited cells are inhibited from delivering inhibitory signals in return. These cell types are very different from each other but are able to influence each other because one sends a signal saying "I am different, don’t bother me”. If the target cell is expressing the notch receptor on the surface and if another cell is expressing the delta pathway the 2 cells will communicate with each other via these receptors and the notch tail migrates to the nucleus to promote transcription of target genes. It is important in regulation of nervous system, retina development, blood cell development. Notch is important because it keep the progenital neuronal cells in the progenital state and they will keep producing neurons. The cell phenotype is the result of multiple signals received. The resulting responses are integrated at the cellular level and define gene expression profiles. File 2 >CELL PROLIFERATION ANALYSIS Somatic cell division: G0,G1,S,G2,M // prophase, metaphase, anaphase, telophase Cyclin & cyclin-dependent kinases (Cdk) are the proteins involved in the regulation of the cell cycle. The Cdk/Cyclin complex must be phosphorylated to advance cell cycle levels associated with phases of the cycle. Cells which do not go through cell division don't go through the different CDK2 cycle. The content of the cell changes dramatically during cell division. Not only the cytoskeleton goes through many changes but also organelles need to be divided equally between daughter cells. The mitochondria for example are correctly parted between cells. During the S phase there is a genome duplication of both DNA strands. The amount of DNA in the single cell is dependent weather or not the cell is preparing for cell division. A cell that is preparing for mitosis will have double the amount of DNA. Cells reach one point where they are not able to divide anymore, this is because of the telomers. Once the telomers get too short and cannot buff the end of the chromosome anymore the whole genome gets unstable, and a crisis is faced. At this stage some cells can overcome the crisis and continue to proliferate, while other cells. In the cells that overcome this crisis the cells can activate the telomerase and re- synthesise the telomers. Cancer cells can sometimes reactivate the telomerase and thus obtaining the capacity to replicate indefinitely. Quiescence cells are cells that no longer divide but can potentially re-join the cycle and proliferate again. A differentiated cell has lost it's capacity to proliferate. >PHYSIOLOGICAL FACTORS FOR STOPPING PROLIFERATION: Metabolic stimuli/starvation Confluency Chemical compounds (mitomycin C, camptothecin,..) Mechanical stimuli Cellular ageing Transformation/tumorgenesis >CELLL GROWTH MEASUREMENTS 1. Cell proliferation assays a. Single measurement like counting cells with Bürker chambers or automated counting. b. Population Doubling time (PDT): measuring cell growth by calculating the time required for the cell number to double. You give a set amount of time to the cells to duplicate. After this set amount of time you count them and then re-seed them in the same modality. This is repeated many times and plot the numbers. By then dividing the [duration of experimentxlog(2)] / [log(C final) - log(C initial)] you get the population doubling time c. Live imaging: Take image of regular intervals. Quantify how much surface is covered over time with a microscope (2D). Not good if cells detach and/or grow on top of each other, if the single cell surface changes, d. Quantifying protein content in culture wells as a surrogate for cell numbers: using a protein quantification assay (eg. Bradford assay) to measure total protein content. But some cells may naturally produce more proteins so there can be a bias. e. Quantify DNA content in culture wells as a surrogate for cell numbers: Measure DNA content before and after treatment. This is done trough kits which usually use fluorescence to determine the amount of DNA. One of the most popular dyes is one is the propidium Iodide (PI). LECTURE 2 – 06/10 >CELL CYCLE ANALYSIS propidium iodide (PI) is a red-fluorescent nucleic stain that binds to double stranded DNA by intercalating between base pairs without sequence preference. This is used to analyse the different phases of the cell cycle. DNA content per cell depends on the cell cycle phase so the high of the fluorescence depends on how many cells are in a determinate cell cycle phase: more cells are in G2 phase, higher is the fluorescence (for example). You can also identify which stage of the cell cycle the cell is since there are different amounts of DNA in each stage, in the S phase a maximum is reached where each cell has two copied of DNA just before dividing and going back to being one. With flow cytometry we can quantify the DNA content/cell and analyse the cell cycle. In the G0/G1 there is a high peak in cytometry and a smaller peak is seen in G2/M phase. We can thus define the cycle of cell samples treated with different molecules. By comparing the graphs we can quantify the % population in each phase and indicate the proliferation capacity of the sample. We can have two different profiles: one before the treatment and one after the treatment - both show how the proliferation cells capacity changes with two different treatments. >CELL INCORPORATION ASSAYS It is possible to analyse how many nucleotides the DNA polymerases incorporate into new strands of DNA. This is possible by using nucleotide analogues to introduce chemically or radioactively labelled nucleotides into newly synthesized DNA during S phase. The original method uses tritiated thymidine (3H-thymidine) which is incorporated into a new DNA strand during S phase replacing any of the bases. This type of thymidine marks DNA but not RNA (replaced by Uracyl in transcripts).You can also identify which stage of the cell cycle the cell is since there are different amounts of DNA in each stage, in the S phase a maximum is reached where each cell has two copied of DNA just before dividing and going back to being one. Every cell division in the presence of H3-thymidine doubles the amount of labelling. So with this technique and with the knowledge of the cell-cycle we have the amount of 3H-thymidine represents the cells divisions. Protocol: compound added to cell cultures  incubation over several days wash excess 3H - thymidine visualized incorporated 3H- thymidine by autoradiography (not common) or Measure incorporated H3-thymidine using a liquid scintillation counter More 3H- thymidine is incorporated -> more divisions are happening It is possible to used other compounds (non radioactive) such as BrdU (5-bromo-2’-deoxyuridine) which is a thymidine analogue, it is incorporated into newly synthesized DNA and requires DNA denaturation for detection because are used anti-BrdU antibodies for the detection. BrdU labelled proliferating cells can be analysed by flow cytometry. There are other analogues such as IdU (5-iodio- 2’deoxyuridine) or CldU (5-chromosome-2’-deoxyuridine) which are non-radioactive, differently from 3H-thymidine. EdU incorporation assay: Instead of using antibodies to recognise 3H-thymidine, it is possible to use fluorophore labelled azide (N3) which recognise the thymidine. 5-EdU (thymidine) is incorporated in the DNA strand reacts with the N3 trough the click Reaction allowing it's detection in microscopy or flow cytometry. Using EdU is very expensive. This technique allows a direct detection without using Ab. >PROLIFERATION MARKER ANALYSIS We detect markers associated with proliferation. PCNA: During late G1 and S phase there is the proliferation of a cell nuclear antigen called PCNA which is absent in resting G0 cells. To detect this antigen it is possible to use histochemical detection (antibodies). PCNA is not the only antigen but there are for example KI-67 which is a nuclear DNA binding protein expressed in cycling cells (G1, G2, S, M) and MCM-2 which is a DNA replication factor. Another marker is Histone H3 which is a intranuclear antigen present only in mitotic cells. It undergoes phosphorylation during cell mitosis from the prophase to the anaphase. Phosphorylated Histone H3 (pHH3) indicated the % of cells in the M phase. By using an antibody anti-pHH3 we can differentiate between G2 and M phase cells. >DYE DILUTION ASSAYS is based on a specific molecule called CFSE (carboxyfluorescein diacetate, Succinimidyl ester cytoplasmic fluorescent dye). CFSE is loaded in the starting population and the first cells incorporate the dye (CFSE è dato alle prime cellule, viene lavato via l’eccesso e poi si osserva come questo viene dato alle cellule figlie man mano che si procede con le divisioni; ovviamente la fluorescenza diminuisce man mano che si procede con le divisioni perchè il numero di cellule aumenta mentre la quantità di fluorescente rimane costante). When these cells divide, they transmit the fluorescence to the daughter cells causing a procesive dilution for each cell division. The fluorescence level indicates proliferation - there is a progressive dilution of fluorescence over cell division because the amount of initial fluorescence is destined to more and more cells. The fluorescence is measured by flow cytometry. For each round of proliferation the Dye concentration will decrease (low fluorescence=more proliferative cell population / high fluorescence= cell did not divide much so the proliferation has been stopped) Summary of cell proliferation assays: Examples of markers analysis to mark different phases of the cell cycle: File 3 >PROTEIN MARKER DETECTION TECHNIQUES In this part we understand how to phenotype ourselves using protein markers. The purposes of analyse protein markers are: Identify marker profile Sample phenotyping Correlate gene and function To define a cell is important to analyse protein markers and RNA markers (RNA sequence-based recognition-> lesson 4). In this lecture we will see the protein markers like:  Cell surface markers Cytoplasmic/cytoskeleton markers Nuclear markers To recognise the protein markers we will use antibodies. There are different questions that we need to answer: is the target expressed (bulk detection - western blotting or ELISA)? Where is the sample expressed (localised detection - immunostaining )? Which cell expresses the target (single cell detection - flow cytometry or MACS)? >PROTEOMICS APPROACH Bulk sample detection - is the target present or not? There are two techniques used: 1. Protein identification - proteomics. Proteomics is a technology enabling the systematic, wide-scale characterization of the proteins contained in each sample to provide information on identity and amount of protein. Proteomics technology is used to detect protein in different samples, to identify co-expressed proteins, to create a protein fingerprint of the sample. For protein detection: the proteins are extracted from the sample the proteins are digested and separated in peptides there is a mass spectroscopy which is an instrument used to divide and identify all the peptides present in the initial sample the proteins are detected and quantified based on their mass -> have your library Compare results with databases to identify the peptides and proteins Summary: in proteomics technologies proteins are extracted, digested, separated and then analysed in mass spectroscopy. We can analyse down to 10-100 cells in nanodroplets at a time or we can extract protein also from a single cell and make a single-cell mass spectrometry. This type of analysis is important in the case of cancer cells because we know that cancer is a group of different cells, identify protein modifications, elucidate protein-protein interactions and develop protein networks….. 2. Antibody-based technique used to detect and quantify a specific target protein among a mixture of proteins extracted from a sample. western blot (quantitative): The technique involves: o destruction of the simple to extract the sample denaturation of the sample o gel electrophoresis (separation of the proteins based on size) o electroblotting (transferring proteins onto a membrane) o membrane incubation with a primary antibody (used to identify the protein of interest) o incubation with a secondary antibody conjugated to a tag (binding to the primary antibody)  o detection of the secondary antibody tag (revealing the presence of the protein of interest) o if the tag is present, after different washes, it means that the target is present. o colorimetric or chemiluminescent detection using HRP (Horseradish peroxidase) or ALP (Alkaline phosphatase). The second antibody is bound to an enzyme that is able to produce luminescence when the substrate is present. To quantify the protein of interest, the signal must be compared to that of a housekeeping protein, serving as loading control across the samples used - we need something to normalize such as actin. Dot blot method: here the sample is not run on a gel but directly deposited onto a membrane (dot)- There is no protein separation and the membrane is processed for immunodetection as for a Western Blot. In this case: o sample harvest and extraction o sample denaturation o vacuum assisted bottling (transferring proteins onto a membrane by forcing the sample onto the membrane by vacuum) o membrane incubation with the primary antibody (detecting protein of interest) o incubation with the secondary antibody conjugated to a tag (binds to primary antibody) o detection of the secondary antibody tag (same as western blot) to reveal presence of protein of interest A housekeeping protein is used to normalize the signal. In Dot blot there is no gel electrophoresis. ELISA (enzyme - linked immunosorbent assay) which is based on detecting a protein target by immobilizing the antigen to its specific capture antibody. It provides qualitative and quantitative readout for the target protein in the sample. The most known is the Sandwich ELISA. Sandwich assay uses a capture antibody and a detection antibody: antibody - antigen - antibody with signalling tag. Important is the use of protein microarrays which are a large-scale screening of protein targets in samples. Here it is possible to detect protein targets by immobilizing a range of specific antibodies on a slide so it is possible to analyse all the proteins present in a sample at the same time. Protein arrays can be widely adapted to study various types of protein properties such as protein-protein interactions, protein - DNA interactions, protein - peptide interactions. >PROTEIN IMMUNODETECTION APPROACH Localized detection - where is the target expressed? The main technique to use is immunostaining (qualitative) which provides information not only on the presence of a target protein but also on the spatial distribution within the sample. With the bulk detection it is not possible to discriminate between low expression, many cells and high expression, few cells. Immunostaining, on the other hand, can discriminate if there is a low or high expression and in which cells. How it works: sample fixation Blocking (incubation with protein mix to reduce nonspecific binding of the antibody and this reduces background signals)  Sample incubation with primary antibody (recognise target protein in the sample) Incubation with secondary antibody conjugated to a tag (recognises the 1st antibody) Detection of the secondary antibody tag (fluorescence or enzyme-based) There are different detection methods: 1. Direct detection of the primary antibody - primary antibody conjugated to a fluorochrome and directly binds to the target protein 2. Indirect detection of the primary antibody - The primary antibody binds to the protein. The secondary antibody is conjugated to a fluorochrome or to enzyme Horseradish peroxidase HRP (brown colour) or alkaline phosphatase ALP (blue colour) and binds to the Fc of the primary antibody. We use the indirect method because it is more sensitive because of the amplification of the signal - there are three copies of the protein; if we use the direct detection there are three signals while if we use the undercut method we have three primary antibodies and each primary antibody can bind multiple secondary antibodies at the same time and the signal is amplified. It is also possible to increase the amount of enzyme per recognition event (multiple signals molecules are bound to the secondary antibody) and the colour becomes brighter and it means multiplying the number of substrate molecules converted = stronger signal. Important are the controls to confirm that what you are watching is correct and the protocol is correct: Positive control - to ensure the protocol is able to successfully detect the problem. In this case it is needed to run one extra sample containing the protein of interest alongside the experimental sample. It is used to detect false negatives. Negative control - to ensure the protocol does not erroneously produce signal in the absence of the protein. In this case it is needed to run one extra sample containing a sample devoid of the protein of interest and at the same time, run one entry sample containing the protein of interest but no primary antibody. It is important to detect false positives. >MULTIPLEXING is based on detecting several different protein markers in the same sample using different secondary antibodies conjugated to different tags -> co-detection of different fluorochromes. So with multiplexing it is possible to identify different types of proteins in the same sample which are represented by different colours of the fluorochrome. For example it is possible to use HRP and ALP in the same moment. The capturing of the fluorescence can be automated thanks to high content imaging acquires automatically fluorescent images. It can be coupled with multi parameter algorithms to visualize an quantify the different signals such as large-scale datasets. It can analyse multiple wells in the same experiment/multiple labels in a single sample well. (multispectral imaging) Automated fluorescence imagine acquisition Automated image thresholding, segmentation and feature extraction AI/deep learning: Data processing, signal measurements and statistical analysis Key points of the lecture: The protein markers contained in a sample can be analysed through different modalities. Proteomics approaches allow the systematic identification of the whole proteome in a cell sample (not based on target affinity/recognition). Antibody-based approaches allow the detection on target protein markers in samples: Western blot and dot-blot techniques indicate the presence of the target in a sample (quantitative) Immunostaining detects the presence and spatial distribution of the target in a sample (qualitative) Detection can be based on fluorescence or enzymatic-based colour reactions (allows amplification). Appropriate controls are required to avoid false positive and false negative detection. Simultaneous immunodetection for different markers is possible using compatible labels (multiplexing) The process can be scaled-up using high content imaging platforms. LECTURE 3 – 09/10 File 4 >SINGLE-CELL TECHNIQUES FOR PROTEIN DETECTION - WHICH CELL EXPRESSES THE TARGET? 1. Flow cytometry Single cell detection is often made by flow cytometry. It's the ability to characterise the cells one by one in a sample in fluid phase. The machine generates a stream of single cells. The cells pass one at a time through the channel and are hit by a laser from the side. The cell then reflects the light forming types of scatters. The reflected light can be read and detected which can then be transformed into data about the cell. Flow stream passes through a nozzle; this nozzle vibrates and makes it so that there is not a stream of liquid but tiny droplets which should contain each one single cell. The droplets pass trough the laser and reflect the light which is then analysed by detectors. There are different types of detector channels: Forward scatter channel (FSC): collects data on cell size (collects light at 0°) Side scatter channel (SSC): collects data on cell granularity or the content of the cell (takes light at 90°), if the cell is very granular you will get much signal in that channel Fluorescent channel: cannot take physical parameters from the cell but only the fluorescence from the fluorophores, it is dependant on the machinery and which light it can emit Antibody-based labelling Cells labelled with fluorescent antibodies that can be excited at determined light waves. Flow cytometry can use a specific light to excite these fluorescence and make it emit light that can be measured by the machine. Either the Ab is directly labelled with a fluorochrome or there is an indirect coloration with the 1st Ab and the 2nd Ab which is associated to the fluorochrome and recognizes the 1st. Surface marker: o is fully accessible o antibody can be directly incubated with the cell o Compatible with live labelling Intracellular marker: o antigen is not directly accessible because there is a membrane o requires a cell permeabilization (poke holes) to let the antibodies through o Not compatible with live labelling How to analyse the data from flow cytometry? It can be represented in 2 modes: 1. Histogram modality (mono-parametric analysis I analyse 1 parameter at a time (1 fluorescence). On the X-axis is the signal intensity and on the y-axis is the number of cell. The more intense the fluorescence is the more the peak will be on the right. With this technique we can identify sub-population, mixed-population and positive-populations. 2. Dot plot (multiparametric analysis) It gives the overall profile of the population. Every dot represents one droplet that the machine analysed. Each axis shows the intensity of a fluorescence signal. The cells that don't show fluorescence are on the bottom left and are considered "negative". Cells/dots on the right and above show fluorescence. This type of graph is often used when there is more than one parameter (NOT for mono-parametric analysis) because there are machines that have more than one laser -> can do analysis simultaneously. It is better if the chosen colours of the fluorophores are very far from one another so that the detectors can be discriminated easily. There can be analysed up to 20 different parameters (18 fluorophores + FSC/SSC) at a time. Application of flow cytometry: Protein marker detection for phenotyping CD (cluster of differentiation) are markers/surface markers different for different cell types. They are defined by the specific monoclonal antibodies to which they bind. They are used for the immunophenotyping of cell populations, based on established surface profiles. Often used to identify leukocytes. By choosing different CD in combination with each other you can discriminate cell in different populations. Remember that cells can have multiple CD on them. Transfection efficiency/analysis (GFP, dsRed, mCherry,..) Transfect cells with something that make the cell express a (fluorescent) protein. You then see the "positive cells" which are the ones which have actively taken in the trans-gene. If the protein is fluorescent you can then analyse them in the flow cytometry. Fluorescent dye measurement (eg. DNA-binding dye, lipid dyes) Lipids can be detected by specials dyes which can be seen in flow cytometry. Cellular assay based on fluorescent read-out (eg enzymatic assay, calcium assays) Indicators that bind to Ca2+ visible in flow cytometry. Controls for cytometry: They are required to set the limit for "true" signal and avoid false positive/negatives. Unstained controls: determine the level of autofluorescence emitted by the sample. Isotype control: Antibodies similar to the primary antibodies but lacking the specificity to the protein target. They are used to determine the non-specific binding. If this one is also positive we cannot assure that our other result is truly positive Biological samples: known negative and positive samples. Their data can be used to validate my measurements.  2. Cell sorting: FACS (fluorescence activate cell sorting) Separation of cell population based on a set criterion defined by user (eg size, marker fluorescence ,co-expression of markers, dye,..). The stream is separated in single droplets with the nozzle. The droplets pass trough the laser where they are examined. As the droplets go trough the machine will charge the droplets depending on different characteristic it detected. For example i only want green cells -> droplets that are green get positive charged, droplets which are red get a negative charge. The machine then separates/deflects the droplets based on their electric charge. The selection criteria is defined by the user. It has a specificity of 98%. This method is used to purify different subpopulations within a sample to obtain a homogenous subpopulation for further culture, analysis, transplantation etc. The separated sub-population can also be kept in sterility. The machine can separate the cells in tubes or well plates (for clonal analysis). 3. Mass Cytometry (CyTOF) Uses antibodies bound to metal isotopes to label the cell sample. Each Antibody for a specific protein target is labelled with an isotope. It functions similarly to the fluorescence marker but instead of a fluorescence signal you get an isotope signal in the mass cytometry machinery. It provides a measurement based on the time each isotope takes to pass through an electric field towards the detector (the bigger the isotope, the longer the time it takes). It is a destructive type of analysis because the sample have to be destroyed in the process in order to analyse the isotopes. It enables the simultaneous detection of many more labels then with fluorescence because in this technique there are up to 135 channels and there are many more isotopes than possible fluorescence colours. 4. Imaging Flow Cytometry Allows the image capture of the cells as they pass through the laser using a camera. The machine takes a picture of the single cell once it passes through the laser. So on top of the fluorescence channel, FSC and the SSC there is also a camera which takes pictures of every cell. With this method you can analyse many cells very fast and you can also see where the signal is coming from (shows fluorescent label localisation) -> adds information on morphological characteristics and on label location. It is also interesting for cell-cell interaction because the machine can take a picture of these interactions. 5. Magnetic Activated cell sorting (MACS) It is an alternative to the cytometer. The other techniques use all cytometry; this one not. The antibodies are bound to magnetic nanoparticles and thus responsive to magnetic forces. The Ab will recognize the target, then there will be a magnetic separation with a magnetic column (similar to a needle). The "needle" will enter the sample and all the particles that have a magnetic Ab will attach to the column. The rest will be separated. To elute the cells that bound to the column the column is removed from the target and all the magnetic antibodies will remain bound. It is more gentle on the cells so it can be a viable option for sensitive cells/when you don't have many cells (for example: patient tissue)-> you need the cells to survive the sorting process. It is mainly used for mono-parametric experiments and especially when there are large populations. There can be experiments where you sort for more than one parameter but you need to do multiple rounds of selection and in the seps in between you need to release the magnetic particles (cut away/remove the previous magnetic tags). Key points: Flow cytometry is used to identify and analyse different cell populations in a simple (cell suspension) The technique is fast, high throughput and provides a cell-by-cell characterisation of the sample Parameters analysed include size (FSC), granularity (SSC) and fluorescence in different channels Flow cytometry is applicable to live or fixed cells, and allows the analysis of surface and intracellular markers using fluorescently-labelled reagents Flow cytometry can be used to sort distinct cell population, based on one or multiple selection criteria Imaging flow cytometry allows the acquisition of images during flow analysis Cell sorting can also be performed using magnetic-based immune-separation with the MACS technique File 5 >CELL HEALTH ASSAYS Cell health can be affected by intrinsic factors (cellular ageing, cellular control mechanisms, Dna damage, metabolism disruption) and extrinsic factors (pathogens, toxins, cell stress for eg. ROS, chemical stress, loss of physical integrity, mechanical stress). CELL VIABILITY ASSAYS - LIVE/DEAD CELL COUNTS: Are used to measure the proportion of live cells within a population, in many experimental procedures for comparing responses to drugs and for end point analysis or as time-course experiments. 1. Trypan Blue exclusion method Trypan blue dye marks dead cells. It can only enter the membrane of the cell if the cell membrane is compromised so it's dead. Alive cells have an intact membrane and the dye will not enter. Then watch the cells under the microscope and see what % of cells is dead. 2. Nuclear-ID dual dye method Used to discriminate between viable and non-viable cells. Use 2 dyes: blue fluorescent cell-permeable nucleic acid dye that labels all cells and the green fluorescent cell-impermeable nucleic acid dye that labels dead cells. These are fluorescent dye so you can use it in fluorescence microscopy and also the flow cytometer which can also read/count the sample in seconds (automatically also separates if needed with FACS). 3. Esterase activity and membrane integrity Based on using 2 stains based on intracellular esterase activity and plasma membrane integrity. There will be a strong green fluorescence in live cells because their esterase enzyme can modify the substrate (Calcein) and make it be fluorescent. Non-vital cells will not have the enzyme so they will not be green. Then a red nuclear fluorescence molecule is used to determine cells with damages membranes because this dye (eg. propidium iodide) enters only in dying/dead cells with damaged membrane and then bind to their DNA. Alive cells with have a continuous membrane and they will not be red. This type of coloration is visible with a fluorescent microscope and with flow cytometry.  4. ATP assay Principle is that ATP is a marker of viable cells. ATP can be released from live cells by lysis for measurement. If you open all the cell in the sample you will know the total ATP amount and you can do an estimate of the alive cells. Then you can pair the ATP extraction with a luciferin molecule and with the luciferase enzyme. The luciferase will use the ATP to form light from the luciferin. The more ATP you have the more the enzyme can elaborate the luciferin and the more light you will have. After this assay you can not use your sample anymore. CELL VIABILITY ASSAYS - METABOLIC ASSAYS An alive cell will have a metabolism that can be detected via the products of the metabolisms. 1. MTT assay First widely used alternative method. It is based on the measurement of metabolically active cells in culture as an indicator of viability. It is based on the activity of a metabolic enzyme and the MTT dye which is light yellow and cell permeable. MTT is added to the cells and incubated for 4h. The dye enters in the cell and ends up in the mitochondria. In viable cells, the tetrazolium molecule (of MTT) is reduced to formazan (an insoluble purple product) by the mitochondrial succinate dehydrogenase. The product can be quantified by measuring absorbance (at 570nm) which correlates with the amount of viable cells present The limitations are the moderate sensibility, it is not ideal for heterogeneous populations because they can have different levels of ATP, the duration is of 4hrs and the sample may be compromised by the end. 2. Real-time glow assay This method is used to monitor viability in live cultures. A cell-permeant pro-substrate needs to be activated in the live cells to become the active substrate. The luciferase enzyme can produce a bioluminescent signal in the presence of the substrate. In dead cells the pro-substrate remains inactive and it is not usable by the luciferase so there will be no reaction. If the cell is alive the pro-substrate is activated, the luciferase can react with it and there will be a light signal. The advantage of this is that the reagents are not toxic, so the assay can be performed on the same samples several times and it is not a destructive method. The other advantage is that the light emitted is quantifiable so you get a quantitative analysis. Often used to see the effects of radiation. 3. Lactate dehydrogenase (LDH) assay LDH is a cytosolic enzyme present in different cell types. It is released in the medium only if the cell have a damaged plasma membrane. Extracellular LDH can be quantified by an indirect enzymatic reaction. In the first reaction LDH catalyses the conversion of lactate to pyruvate via NAD+ reduction to NADH. In the second reaction resazurin is converted in resurfin by the enzyme Diaphorase by using NADH. So the first reaction uses NADH and the concentration of NADH is dependant on how much LDH there is. Resurfin level is proportional to the amount of LDH released and it emits light that can be measured at 490nm. The more signal you get the more damaged cells you have in your sample. Protocol: Treat it with different vitality compounds->Then collect medium of different conditions and put it in the fresh plate (wells) ->Add reaction components (lactate, diaphorase) leave to incubate for 30min->Add a stop solution-> Measure the absorbance with how much purple light you have LECTURE 4 – 11/10 CELL VIABILITY ASSAYS — APOPTOSIS ASSAYS: Apoptosis is programmed cell death. The main morphological signs for cell death are cell shrinkage, membrane blebbing (protrusiuns in membrane), nuclear fragmentation, chromatin condensation. So if there are more apoptotic cells it means there are more dying cells. Apoptosis can be induced experimentally by withdrawal of serum or growth factors for growth factor-dependent cell lines and with treatments with proteins synthesis inhibitors or DNA topoisomerase I inhibitors (camptothecin). Apoptosis is mediated by the activation of two groups of protein Bcl2 proteins and caspases. 1. Plasma membrane integrity assay Phosphatidylserine (PS) is a phospholipid with a negatively charged head-group and it is an important constituent of the cell membrane. In health cells PS is found only in the inner leaflet of the plasma membrane. During the membrane degradation during apoptosis it causes the exposure of PS in the outer leaflet of the cell membrane. We can target PS residues with Annexin V which is a phospholipid binding protein with a high affinity for PS. A labelled Annexin V is used as a probe to detect PS on the outside of the membrane which stains the apoptotic cells. With Annexin V attached to a fluorescence molecule we can also use flow cytometry. If we add to the Annexin V PI (propidium iodide - vital stain) we can mark dead cells. Only late apoptotic cells will let the PI inside because their membrane is compromised. Normal cells and early stage apoptotic- cells will not be affected because their membrane is not compromised yet. The cells which less PI inside will be stained red. PI is used to see different stages of the apoptosis. If only a normal fluorescence with annexin V and no red from PI -> membrane has PS on the outside so it is doing apoptosis (both early and late stage apoptosis stained), if also red from PI -> late stage of apoptosis. 2. Tunel assay (terminal deoxynucleotidyl transferase dUTP nick end labelling assay) Apoptosis is characterized by fragmentation of genomic DNA into multimers (c.a.200bp). With this technique the DNA ends generated upon DNA fragmentation are enzymatically labelled at the free 3'OH termini. By using modified nucleotides, these fragments can be detected and apoptosis can be analysed. The nucleotide are labelled with fluorophores and only if they get corporated they emit fluorescence. Usually in this technique there is a generic counter-coloration/stain for example with DAPI. The tag is added on a dUTP which will attach on the 3'OH end of the DNA trans break using a terminal deoxynucleotidyl transferase (TdT). The tag is afterwards recognised using an antibody-based recognition. The antibody can either have a fluorescence label or an enzyme and we can identify apoptotic cells. If there is fluorescence we can use the flow cytometer 3. Apoptotic markers - Caspases (cysteine-aspartic proteases) There are different caspases which act at different times in the apoptotic process. The caspases are usually inactive (pro-caspases) and get activated trough cleavage. When active they can cleave a range of substrates and take part in the apoptosis activity. There are different ways to detect caspases and one has also to consider if it is already activated or it is a pro-caspase. There are 3 types of measurements: Antibody-based immunodetection Anti caspase antibody is used for western blot (quantitative), immunostaining (microscope, qualitative) or flow cytometry (quantitative) analysis. It is used to recognise the cleaved (=active) form of caspase-3. The activated caspase has a lower molecular weight then the pro-caspase.  Fluorometric activity assay Caspase substrate becomes fluorescent after cleavage and DNA binding. The fluorogenic substrate is cell-permeant. The DNA is attached to a peptide which keeps it from being fluorescence, once it encounters the active caspases these separate the peptide from the DNA dye. Active caspase-3 and caspase-7 proteins in apoptotic cells cleave the recognition sequence (peptide) in the substrate and free the DNA dye. Once cleaved the DNA dye can enter the nucleus and bind to DNA and emit fluorescence. Bioluminescent activity assay A substrate is given to cells. After cleavage by the caspase, the substrate is able to react with luciferase to produce bioluminescence. In apoptotic cells the substrate is activated upon cleavage by different active caspases. Once cleaved it can be metabolised by luciferase and emit light. The luminescence can be measured quantitative by luminometry. Only apoptotic cells are luminescent and quantifiable. CELL VIABILITY ASSAYS - CELL SENESCENCE: Cells reaching the end of their replicative capacity experience DNA damage, to which they react by entering senescence. Once the cell enters senescence they will not replicate again and die. To know how many cells are senescence there are different test. Definition senescence: irreversible loss of proliferative potential associated with specific morphological and biochemical features. 1. Microscope Just by looking at the cells in the microscope (in vitro) have different morphology. The cells have irregular borders, are very spread out. 2. Expression of senescence-associated ß-galactosidase (SA-ß-gal) The presence of this enzyme can be used as a marker to see how many cells have entered senescence. If you feed a senescence cell which has this enzyme with the substrate you get a blue coloration. The substrate X-Gal is the substrate of the ß-galactosidase. Cells that have not gone into senescence and still proliferate don't have this enzyme so there will be no colour. This technique is used on fixed cells and it is not suitable for real-time imaging of senescent cells. A new substrate is used that if metabolized by the enzyme it will become fluorescent (NIR fluorescence "ON"). This is suitable also on live cells and for real-time imaging of senescent cells. One of these new substrate type is GlycoGREEN. Based on different substrate we can have different coloured fluorescence (red/green/…) that is quantifiable in flow cytometry. CELL VIABILITY ASSAY - RAMAN MICRO-SPECTROSCOPY: It is a technique to detect vibrational modes of chemical bonds. It identifies the chemical composition and creates a chemical fingerprint of the sample. The Raman spectroscopy is based on the Raman effect: for each molecule exposed to laser light a small fraction is scattered with a shift in frequency that can be recorder and is highly specific for each molecule and created a unique fingerprint for the sample. Confocal Raman Microscopy (CRM) is the Raman spectroscopy combined to a confocal microscope. CRM can distinguish cells at different stages of cell cycle and living cells from dead cells. It is a non invasive method to assess cell viability at single-cell level. You can compare the fingerprint of different cells or of the same cell at different stages/treated with different drugs and observe the changes in their scans. In dead cells we usually have more lipids, some cellular compounds are more capsulated, many organelles are degraded so the fingerprints will be different in dead and alive cells. This technique is still very new and the peaks of the graphs are still being analysed to document them in detail. This technique is for now used especially in experiments to see mitochondria. Particles bound to gold nanoparticles can recognise the mitochondria and accumulate in them -> then they are recognisable with CRM. File 6 >GENE EXPRESSION ANALYSIS: RNA MARKERS A. Bulk detection methods - is the transcript expressed and which markers are expressed B. Spatial detection approaches - where is the transcript expressed C. Single-cell techniques - in which cell type is the transcript expressed A.RNA MARKER EXPRESSION ANALYSIS - BULK DETECTION METHODS These methods consider the whole sample and not its single parts. -is target transcript expressed? 1. Northern blot Based on the analysis of RNA material, separated on a gel by electrophoresis and blotted on a membrane. Hybridisation with a probe that is reverse complementary to the target RNA sequence. The probe is labelled with a tag which can be radioactive, fluorescent or biochemical. The detection of the tagged probe indicates the presence of the target RNA in a given sample. The expression level can be semi-quantified if normalised to a housekeeping RNA level. Procedure: Lyse the cell sample to extract the RNA and purify it so you only have the RNA Separate the RNA on an agarose gel (smaller fragments will run faster than big fragments) Transfer the run gel on a membrane Add labelled cDNA probe complementary to the RNA of interest + Hybridize denatured probe with the membrane Wash membrane and expose the film To normalize and quantify the signal you get you need a housekeeping gene like actin (with ImageJ) 2. RT-PCR (reverse transcriptase-polymerase chain reaction) Based on the detection of a target mRNA through the production of a cDNA copy. It requires multiple amplification cycles by Polymerase Chain Reaction. The first step is the reverse transcriptase where the mRNA is reverse transcribed to double strand cDNA. Initially you form a double strand hybrid between RNA and DNA. Via a RNAse the RNA part will be digested while the DNA part will remain. Then you polymerize the complementary DNA strand to the DNA template to get a double strand cDNA. The PCR part will be done by a chain reaction to amplify the cDNA to make multiple copies. The PCR needs primers, taqPolymerase, dNTPs and buff mix. Each amplification cycle is repeated multiple times and each is composed of 3 steps where the temperature is changed constantly. First there is a denaturation of the double strand DNA to separate the strands with high temperatures (>90°C), then there is the annealing where the primers bind to the DNA (≈50°C) the last step is the elongation step (≈72°C) where the taqPolymerase synthesises the new strands starting from the primers. The detection of the amplified fragments can be done by gel electrophoresis or in some cases directly with quantitative real time PRC because the final number of DNA will depends on the initial number used as loading. Usually in "traditional" RT-PCR (no real time) you run a second PCR also with a housekeeping control protein such as actin to have a semi-quantitative analysis.  3. Quantitative RT-PCR (RT-qPCR) Based on analysis of mRNA material, reverse transcribed to cDNA. Hybridisation with 2 primers specific to the target cDNA sequence. Based on multiple amplification cycles using DNA polymerase. What changes is how you detect the amplified fragment. This is done by using a fluorescent tag. Quantitative measurements are made in real time by analysing the fluorescence directly. In the end there will be a graph evaluating the number of copies initially present in the sample. The signal emitted at each cycle determines how many copies there are in each cycle until it will reach a plateau where you will no longer have free primers and dNTPs available to do he reaction. In the graphs there is a threshold of fluorescence signal -> if you need less cycles to reach that threshold it means you had more copies to begin with. This value is called Ct. The lower the Ct (cycle at which the threshold has been reached) the higher the copies in the start of your gene of interest. SYBR Green method Uses SYBR green (binds to minor groove of dsDNA) to produce a fluorescent signal when bound to dsDNA. The fluorescent signal reflects the amount of the DNA in the tube. This is not specific because the amount of SYBER green is related to the amount of dsDNA but it also could not be the target dsDNA (doesn't discriminate between types of genes, only on dsDNA). TaqMan method Uses sequence-specific probes that carry a fluorophore and ist quencher. It binds to the target sequence and it remains non- fluorescent if it remain intact which. Means if the reporter R and the quencher Q are near each other. When the polymerase synthesises a double stranded copy it will displace and remove the probe. This will cause a modification in the probe cleaving it in half. The R and Q will not be close to each other anymore causing the R to emit a fluorescent signal. The fluorescent signal released is proportionate to the number of target copies. This type of mechanism is highly specific because the probe will only attach to your known gene of interest. The other probes even if they attach, they will never emit fluorescent because the R and Q in those cases will always remain together and never emit light. By using multi-well systems we can analyse multiple samples contemporary. LECTURE 5 – 13/10 -which markers are expressed? (bulk detection methods) 1. Marker Survey - Gene Expression Micro Array Based on the analysis of mRNA material. Template RNA is reverse- transcribed with a tag to produce a labelled cDNA sample. The labelled cDNA samples are hybridized onto a probe array (on a microarray slide/panel) where there are present thousands of probes of different genes. Some cDNAs will be complementary to the probe array sequences and will remain attach to the probes even after washing procedures. By detecting the tags of the cDNA we can then detect which ones have attached. We know from the start what type of array probe is attached to each spot of the slide and we also known the corresponding gene of each probe. Via these information we can analyse the colours emitted by the cDNA tags on the whole slide/panel together and known what gene their represent. Protocol: Extract mRNA -> Transcribe RNA in cDNA and attach a probe which can be a fluorescence tag on the end of the sequence or a fluorescent-emitting dNTP -> Hybridize the cDNA to the microarray slide -> Wash the panel to ensure specificity of the target-probe binding -> Detection of the signal by scanning the slide and see in which position you get the signal -> analyze the results Assess if set genes are expressed in the mRNA sample or compare the expression level of set genes of different mRNA samples by using different fluorescent-labeled nucleotides on glass slide arrays. With the second method we can in one experiment see if the both, only one or none of the sample express that gene by looking at the colors in a microarray slide (red would be only one of the two samples, green the other and yellow if both express it because it is a mixture of both colors). With this method you can have million of probes on one slide and it is also used in the clinic for example to compare expression of RNA in different tumor tissues etc.. 2. Nanostring Technology To identify which genes are transcribed in a sample. The probe hybridizes with the target gene in a liquid phase before being immobilized onto a slide. In this type of experiment there is no need to reverse transcribe and amplify the mRNA. The hybridization is done directly to the mRNA. The recognition is based on the complementarity of the probe with the mRNA. The probe can be divided in 2 different sides; on one side there is the fluorescent barcode and the other side there is a capture bead. The capture bead will be complementary to the target. Each probe has a unique tag sequence which is a fluorescent barcode able to mark in a specific way a transcript present in the sample. How can I retrieve and immobilize the recognized mRNA on a solid phase? This is done because of the end of the capture bead where is a little area that will stuck on the solid cartridge. Each immobilized barcode is then read thanks to machines which can decode the barcodes and make a list of all the mRNA (=genes) that are present. The disadvantage of these techniques is that you know what genes you are looking for and tell the company to make the barcodes probes for you. So you will not find new genes.  3. Open marker survey - Rna-seq (illumina sequencing) Identifying in an unbiased way all the transcript presents. With this mechanism you will find new genes because it will sequence all the RNAs available. Based on RNA collection, fragmentation, reverse transcription and sequencing. Sequenced reads are then mapped onto the reference genome to reconstruct transcriptomes. Expression level are quantified and can be compared between samples. This technique is: high throughput it catalogues all transcripts in a given samples used for mRNAs and non-coding small RNAs it is a single base resolution technology because it reads/decode them one single nucleotide at a time It required low amounts of starting materials Is both quantitative and qualitative Procedure (not very important, do not study): RNA processing -> fragmentation into shorter products and then bound to primer Library preparation -> reverse transcribe in cDNA then add adaptors (like primers) to the ends of the cDNA to be able to make a dsDNA Read and map -> map onto the known genome the fragments of the dsDNA (this step is also quantitative because the amount of the fragments similar to the known DNA will be registered so we can trace back how much RNA there was in the sample which is directly correlated to how much it has been transcribed) Bioinformatic analysis -> the machine will give you the absolute number of the fragments but also (if available) the relative number between the ratio of the expression said fragments in 2 different samples B.RNA MARKERS - SPATIAL/LOCALISED DETECTION METHODS Methods to detect where the marker is expressed in a spatial setting. -where is the transcript/target genes expressed? 1. Laser capture microdissection It is applied when working with fixed tissues. First the operator identifies the area of interest in the tissue sample -> if you analyse a tumour you will not choose the part of the tissue where there are healthy cells but where there are more tumour cells. Then the operator cuts that area of interest with the laser of the microscope. Then you can apply all the bulk methods seen before onto that sample. This methods allows you to discriminate the markers of one type of tissue/cell by selecting this sub-population/area from the start but it is not possible to select genes and ask where/how they are distributed within the sample. 2. Target sequence detection - in situ hybridisation (ISH) It is based on the use of a specific probe to hybridise with the sample in order to detect where the target mRNA is expressed within a sample/tissue. It is designed to detect the presence of a sequence of interest within a tissue. The probe is labelled and is often a nucleic acid probe which will recognise and hybridise to the target sequence. Through the label it will be possible to detect and visualize the target. ISH can be done with RNA (RISH) and DNA (DISH). The DNA application are mostly done in genetic diagnostic like with chromosomes and if it is labelled with a fluorochrome it is called FISH. On RNA it is used to determine the expression pattern of a gene in a sample. Protocol: Use a tagged probe specific for the target sequence and add it to the sample (reverse- complementary to the target nucleotide sequence) Probe hybridisation with the sample if the tissue/sample expressed the target Probe recognition thanks to the tag -> usually an Ab it is used to recognize the tag (biochemical tag: enzyme or fluorescent conjugated). The tag can also be fluorescent or radioactive. Probe detection to reveal the presence of the target -> immunostaining The probe is made trough the synthesis of a ribonucleotide starting from a DNA template. The tags attached to the dNTPs can be radioactive, fluorescent and biochemical which can be recognized by Ab. If the DNA probe is labelled with fluorochromes the mechanism of the ISH is called FISH. The detection of the probe depends on how the tag is; so either enzyme-based, coloured, Ab,… On DNA FISH is used to localise specific sequences in the genome, to do karyotypic diagnostics and to do in vivo cell tracking. On RNA it is used to determine the expression pattern of a gene in a sample. This experiment needs a negative control to detect false positives which is a probe the same orientation to the target mRNA because this should never attach to the target mRNA since it is not complementary and thus always give no signal. This technique can tell you: in which cell type/tissue marker expressed, how many cells within the tissue express the gene and if used after having treated the samples it can give indication of the expression of the target in treatment/disease scenarios. The limitations are that it is useful only when you have a small number of genes, if you have 50 genes you would need 50 probes which is not possible and very expensive. It is also not ideal for RNA which are only available in low abundances. 3. Multiplexed target survey - RNAscope It is based on RISH approach using probe pairs and signal amplification to achieve single RNA molecule marking in a tissue or culture sample. We are relying on a pair of probes that needs to bind to the transcript, only if both are bound to the target you'll get a signal. It involved a signal amplification to get higher sensitivity and it allows to multiplex (more targets simultaneously). The transcript needs to be recognised by 2 different probes working in pairs which cover a total of 36-50bp. The probes are complementary and recognise neighbouring parts of the sequence sitting thus next to each other when they recognise their target. The probes reach carry a tail to make them detectable if bound in pair and is very important in the amplification and recognition step; they are Z shaped. So after the probe pairs bind to the target mRNA and position correctly next to each other their tails will be near to each other and will be bound by a preamplifier. The amplifier molecules bind to the preamplifier scaffold and the labelled probes bind to the amplifier branches and detect the target. So the probe (Z) is not labelled but it gets labelled only after 2 of them have bound the target. Procedure: Permeabilize -> the sample are fixed and pre-treated with a RNAscope Pre-treatment Kit to unmask target RNA and permeabilize cells (TLDR: poke holes in the cells) Hybridize -> add the Z probes and let them hybridise with the target Amplify/Labelling -> assembling of the preamplifier and amplifier with the labelled probes which can be fluorescent or enzyme tagged; the amplification step is because each binding event of the labelled probe to the amplifier will cause the emission of the signal (the preamplifier/amplifier are slightly different for each labelled tag so that only that labelled tag will attach) Visualize -> with a microscope, if you use fluorescent you can use different colour Quantify -> manual counting or computer This method is highly specific because both Z need to bind the target and it is sensitive because the signal is amplified and thus even lowly-expressed transcript can be seen. It is not good if you don't know the sequence of your mRNA. LECTURE 6 – 16/10 File 7 4. Multiplexed target survey - Spatial Molecular Imaging (1st generation) Transcriptome analysis with both high resolution and the ability to visualize the expression distribution in the sample. Prepare a tissue section and make the targets available by permeabilizing the sample. Then you incubate the sample with multiple probes which each has to be uniquely recognizable. Then you do the data acquisition. The probes have a barcode called a DSP (digital spatial profiler) which has only 2 colors, a linker and a target complimentary sequence which is RNA target-specific. The barcode part can also be attached on a Ab which will enable the multiplex analysis even by tagging the target (also proteins) with Ab. It is also possible to incubate the same samples with both Ab-barcode and RNA complementary sequence-barcode allowing you to have a lot of information at once and the reader at the end will be able to recognize weather the probe was a sequence or an Ab at the start. Procedure: Incubate sample with fluorescent antibodies which will recognize specific structures of the sample - > you select visually the area you want to treat (ROI: region of interest) Add probes with each a specific barcode to detect target mRNA in the same sample Machine will shine UV lights on the ROI -> this allows the probes to be broken in 2 separating the target complimentary sequence with a DSP barcode The machine will aspirate/collect the barcodes in the ROI and analyze all the barcodes Then there will be a mRNA profiling for the region of interest by decoding all the barcodes-> have a list of all barcodes with the corresponding mRNA (2nd generation) With this type of technology it is possible to analyze the barcodes directly onto the sample without detaching them from the probe. The tags are different because they are made of a readout domain which makes multiple barcoding possible and a recognition domain. The recognition site of the tag can be reverse complementary to a target mRNA or an Ab linked to the tag. The recognition site will recognize a big part of for eg an mRNA and in this way cover more than one gene. Once the tag bound to its target a fluorescent reporter will be added that will be to the readout domain. The reporter has a fluorescent tag and also carries the same sequence that is being recognized. The reporter can be cleaved using UV light on its PC-cleavable site and the machine then will detect the fluorescent label. After this you add anther reporter that that will bind to another motif of the readout domain. This reporter will be cleaved again to be scanned and all this is repeated many times. The combination of the fluorescence created cycle after cycle of the reporters will generate the barcode. With this technique you can look at multiple transcripts at once on the same spot because each reporter will only recognize one gene. Also this technique makes it possible to see the spatial distribution of marker in a sample and associate these markers with structural information of the sample. At each individual cycle the machine will scan every spot that you have selected in a specific area (10mmx10mm) and form a map of the signals; the time of acquisition of these type of sample takes 1 day to complete. (For now (2023) it has been possible to map 1000 genes in a single tissue at once using the RNA-tags.) Key points: C.RNA MARKERS - SINGLE-CELL TECHNIQUES -in which cell type is the transcript expressed? 1. Single-cell RNA-seq Procedure of Bulk RNA-seq: Want to make library starting from the RNA material. Fragment the mRNA and transcribe in cDNA. Amplification of cDNA with PCR using oligos that contain an adaptor sequence to facilitate sequencing. The reads are then mapped to a reference genome and reconstitute a whole gene sequence/transcript. In the end i identify all the transcripts and quantify te expression level of each fragment. The advantage of doing cell-by cell is that you then know the exact expression of the gene of interest for each cell type making it also easier to compare the level of expression of different cells. Procedure: Get your cells of your sample in suspension where all the cells are dissociated from one another  Separate the different cell types of your sample and tag them Do RNA-seq (concept written before, just like bulk RNA-seq) for each cell of your sample Analyse The separation of the cells is made by running the reaction in droplets, where in each droplets there will be only once cell at the time. The way this is done is by doing droplets which contain 1 cell each and then add reagents to that droplet that are able to tag each RNA present in that one cell in a unique way. The droplets are made by a device made like a tube. In one entry of the tube you put the beads and on the other (lateral) entry you will put the cells and the reagents. These 2 will be encapsulated in single nanodroplet in an oil emulsion called GEM (Gel Bead-in-emulsion) when they are pushed towards the exit of the tube by a stream. [a GEM is a Gel Bead-in-emulsion partition that encapsulates each tiny micro- reaction within the chromium system]. To be sure that each droplet contains no more than one 1 cell, the ratio is adjusted to droplets >> cells. Each bead is covered by many copies of the barcode. The barcode is gradually released in the droplets thanks to the reagents and then it will attach to the mRNA of a single cell and tag it The binding to the mRNA is done by a string of polyT (oligodT) which will recognize the polyA in the mRNA. Each mRNA in the droplets will share the same barcode. So even if you then put all the cells of the different droplets together for the sequencing you will be able to distinguish the mRNA by their unique barcodes. During the RNA-seq the mRNA will be transcribed in cDNA and during this transcription the barcode will be maintained. During the replication with the PCR all the generated cDNA from individual cells will share a common barcode. What happens if there are 2 cells in a droplet? With algorithm called DoubletDecon will remove the doublets and eliminate the data that shows very different barcodes with the same barcode. The doublets get identified and then removed. LECTURE 7 — 18/10 File 8 >CELL MIGRATION ANALYSIS: Cell migration is influenced by the cell phenotype and also by the neighbouring cells present in a tissue. It is very complex and involves also the environment and the cytoskeleton. It involves the protrusion of the cell membrane and involved he reorganisation of the cytoskeleton. Traction forces are generated by the formation of new adhesions at the cell front and enable movement with disassembly of adhesions at the cell rear. The migration relies on the Assemble / Maturation / Disassembly cycles couples to actin polymerization and actin-myosin contraction. There are different migration modes (depending on substrate, cell contractility, etc). Adhesion dynamics are regulated by actin polymerization and actin-myosin contraction involving Rho GTPases and protein Tyrosine kinases. Cell movement plays a key role in many physiological processes such as Embryogenesis, Gastrulation, Neurulation, Tissue homeostasis, wound repair, immune response, angiogenesis, metastasis, inflammation, cancer, etc. Embryonic development: In gastrulation the cells migrate to populate/form the body axis. This type of movement is coordinated and forms the basic structures of the embryo such as the formation of the nervous system (neural crest cell migration). During gastrulation (the formation and the organization of the 3 embryonic cell layers /endoderm, mesoderm, of ectoderm) the formation of nervous system come from the ectoderm, from the sequence of conformation changes in the embryo. The invagination of the ectoderm will form the nervous system à a flat surface form an invagination and then this invagination close itself to form the neural tube (in blue). When the tube closes, at the very top of the tube a population of cells is present (neural crest) that migrate on the dorsal side of the neuronal tube and then, during development, they will migrate through the head basically to form some new tissue. Moreover, they contribute to forming a lot of tissue in the head. If migration is compromised, the formation of the head of the embryo will not be good. Tissue homeostasis: Cells need to migrate to go where they are needed. White blood cells need to chase the pathogens. Endothelial cells self organise naturally to make blood vessels (angiogenesis). In wound healing the cells migrate to close the wound Pathologies: in cancer there is metastasis which is a pathological process where cells change their characteristics to be able to migrate. Typically cells on the edge of the tumour will start expressing mesenchymal genes instead of endothelial cells and they will start to migrate. This is called Epithelial mesenchymal transition (EMT). This is induced by a specific signal such as TGFß signalling which enables them to acquire the mesenchymal identity and the migratory capacity. >CELL ADHESION: The cell membrane proteins will come in contact with the extracellular matrix. The typical proteins that can bind to extracellular matrix are specific receptor proteins called integrins. So, integrins are able to sense and bind to ECM proteins. They don’t only recognise and bind to ECM but they also activate a set of events intracellularly to signal the cells that they found an anchorage point. So integrins are creating the link between the cell environment (ECM) and inner cell, to adapt to the extracellular environment. They are connected to actin filaments in the cytoplasm. In adhesions there are different components involved which regulate the mechanisms of cell adhesion and are important anchorage points of the cell; these are: Integrins: protein which will do the adhesions of the cell, the integrins are read out what happens in the extracellular space and inform the inside of the cell how to adapt and coordinate the contact point, in order to apply traction to be able to move Actin: is the filament which will bind the integrin on the intracellular part. It mediates the anchoring point of the integrin Intermediary proteins: talin and vinculin that serve to bridge the gap between the integrins and the actin cytokeleton Integrins are transmembrane proteins able to recognize ECM proteins (Extracellular matric proteins such as collagens, lamimins, vitrinectin). This link needs to be maintained long enough to create the force to move but also be able to detach because the cells will need to stop eventually. The integrins can transmit the information of their link with ECM in the intracellular space and activate others component in the cell (talin, linkelin, vinculin…). All these proteins are recruited by the activated integrins and able to bind to the actin to adapt the cytoskeleton to the environment. The type of binding between cells can be: E-cadherin Occludin/claudin Gap junctions >SEEDING SURFACES: The nature of what the cell is resting on will affect the cell behaviour. The chemistry of the surface will decide what type of chemical entities the cell will get in contact with. The wettability is how wet a surface is. The swiftness and the topography (if it has nooks/cracks) will also condition the cell and the viability of the cell. Some cell are heavily dependent on having an anchorage point and having no surface or a wrong substrate will cause cell death. Cells which are seeded on different type of support will have different appearance. Only some cells can grow in suspension and form aggregates. With the right type of surface most cells will attach to the surface and start to grow in 2D. The structure of a single cell can change when they are attached/ when they are suspended; some cells get longer once their start to develop their architecture on 2D surfaces. The cells organize themselves based on the topography on which they are seeded The polystyrene used in lab is usually treated to remove some of the phenyl groups of the polystyrene (PS) monomers to have an increase hydrophilicity, facilitate cell adhesion and modulate extracellular matrix composition. >HOW TO MEASURE CELL ADHESION? 1. Static adhesion assay Cells are incubated with a fluorescent dye (eg calcein AM). A set number of cells from each experimental sample is seeded in a dish with different experimental conditions. Cells are left to adhere for a set amount of time and then see how many cells in that time have been able to adhere to that surface. Suspended cells get washed away and with a fluorescent microscopy you can then see the ones that remained attached. You can quantify the fluorescence by taking imagines and then have a parameter to work with. 2. Adhesion measurement Using a measurable lifting force to measure the adhesion between the cell and the surface. The more force you need to detach the cell the more the cell adhered in a good way to the surface. Either with an atomic force macroscopy (AFM) or with a micropipette aspiration assay. 3. Washing technique Cell adhesion is measured by exposure to shear force applied by the stirrer. You have a stirrer which makes the fluid flow and will make the cell detach from their substrate. 4. Spinning disk assay Cell adhesion is measured in a culture dish subjected to different centrifugal force by spinning disk. 5. Centrifugation technique You can also centrifugate the cells: cell adhesion is measured by exposure to centrifugate force applied to the vessel. You are putting the dish of seeding cells on the speeding platform and you apply centrifugation force on that. The stronger you need to spin for the cell to detach, the stronger thew were attached to the plate. The cells that are not very attached to the plate will be thrown out more quickly. If they are very attached, they will resist for more time, until they reach a speed that lift off the cells. That’s another way off applying a force to diagnose how strong the adhesion is. 6. Flow chamber assay Cell adhesion is measured in a rectangular microchannel device, where adhering cells are measured after exposure to different flow speeds. 7. Impedance measurement The cells are seeded on electrodes. In an empty well with two electrodes the electrons will flow from one electrode to another. If cells are seeded on those electrodes they isolate the electron flow making it possible to measure the difference of electrons that are passing from one electrode to the other. Usually there is a plate with integrated electrodes on the bottom. Increased cell adhesion and spreading will lead to increased electrode impedance. This assay is not damaging but the monitoring must be done over many days. At given intervals the machine will record the impedance which translates how many cells are attached to the electrodes on the plate. The impedance is the measure of the opposition to electrical flow. >COLLECTIVE CELL MIGRATION: In the collective migration cells will move together in what looks like a coordinated way. Different cell types will migrate as a single cells but other will coordinate themselves together to migrate together. There are different situations: Migrating leader-follower: Some cells at the front will lead the cells that are on the back. The cells on the front edge will have a slightly different phenotype from the ones in the back Contact inhibition of migration: All cells are migrating and once they touch another similar cells they will collide and migrate in the other direction by changing their migration angle Contact stimulation of migration: Cell with a high cell density near them will migrate while the ones which have less cell density will gets isolated from the pack and stop >CELL MIGRATION Cells respond to a pro-migratory factor by directed movement. Chemotaxis is the capacity of the cell to go into a direction where there is a source of a biological signal. Different signals act as chemo-attractants for example cytokines, chemokines, growth factors, bioactive lipids, extracellular nucleotides, H+ ions. The cell become polarized in presence of signal (the signal activates the receptor on one side of the cell, the signal is concentrated only on one side of the cell). So, from a round morphology it starts to extend on the front end, and it also extend protrusion to exploring the environment. This also causes changing in the forces within the cell’s cytoskeleton. There is a sequential movement form extending the front edge of the cell, extended the membrane on one side. While it migrates the cell will lose its integrity. Usually the leading part will extend and the tail will reduce itself to keep the overall volume of the cell. The end will adjust and keep up with the front of the cell. >CELL MIGRATION ASSAYS 1. Scratch assay You have a plate full of cells. Then you scratch the plate and see if the cell migrate by closing the gap or if the gap remains meaning the cells will not migrate. Over time you take out the plate and take pictures of it. Then you calculate the closure speed and extrapolate the cell migration speed. The migration speed is the time it takes to close the gap which translates if cut by half in the time it takes for each side to migrate and fill the gap. This scratch assay is also called wound healing assay because it basically does the same thing as healing over a wound on the skin. With different treatments we can see how much the closure speed/migration speed is and if it gets affected. The limitation of this technique is that by doing the scratch by hand it will not always result in the same diameter of scratch. To overcome this limitations you can do Standardised scratch assay. Comb: In this method a device is used to scratch looks like a comb and is useful to create the same parallel scratches contemporarily. Cell-exclusion zone assay — Silicon chambers: these chambers are put on the bottom of the dish. In the middle of these chamber there is a silicon wall. Cells will be put on both surfaces on the chambers but not where the wall is. Then you remove the silicon chamber and see how the cells grow to fill the silicon gap. Cell-exclusion zone assay — cylindric seal: You put a cylinder on the plate to create an exclusion zone on the plate where the cells don't grow. Then you remove/lift the cylinder and let the cells migrate. 2. 2D migration track monitoring Track cells with fluorescence and track their migration with the fluorescence on a set period of time. See over time where they have migrated from a fix starting point. The you put a circle with a fixed radius around the starting point and see how much it migrated by recording the trajectory of the cell by taking images over time. The measurable parameters include the net displacement, max displacement, global turning angle, relative turning angle, movement speed, ect. 3. 2D-migration track monitoring assay I measure how much displacement the cell is exorting on the medium and see how much it moved.When the cell is moving, there is a pressure on the extracellular matrix/or on the elastic substrate and also the network (intended as all substrate) is moving. We can calculate how much displacements there are onto the substrate to understand where the cell is moving we can do that thanks to some imaging techniques, one of them provides to seed the cells into a substrate that is transparent (or formed by gel) that contains fluorescent beads. Thanks to the fluorescent beads in the gel, we can visual the displacement of the elastic gel and the cells’ moving express in fluorescence. The images coming from the use of microscopy. We can observe the most important traction of the cells and its intensity. Bead displacement migration model: We can see/monitor the force of the cell's movement with the microscopy by looking on how the deform the substrate. The beads in the gel are displaced via the traction forces transmitted from the integrins.  Micropillar: The migration is seen via flexible pillars of known dimension and flexibility. We calculate the force based on the pillar displacement 4. Microfluids-based chemotactic migration assay These are systems which allows you to study cells in different medium in separate micro-wells. The plate is composed of a series of identical chambers with each two micro-wells and a channel in the middle of them which makes it possible to visualize the sample on the microscope. The cells are seeded in the channel between the micro-wells. First you seed the cells in the channel then you add the medium in both micro-well one on each side. The cells will then migrate towards one or the other medium and see the results under the microscope. You will need controls. In one channel you put no molecules/medium in the micro-wells; this will be a negative control. Then we need a positive control where we have the same medium with the molecule in both micro-wells so that the cells will not move because there is no difference in the concentration of the mediums. >TRANSMIGRATION ASSAY It's the analysis of the invasion capacity of the cells that migrate through structures. 1. Capillary migration assay I put cells in different capillaries which are one of their cell and one with a smaller diameter than the dimension of the cell. I can then see which cells can reach the other side of the capillary. 2. Membrane migration assay Involves a chamber with 2 medium-filled compartments separated by a microporous membrane. Cells are seeded in the upper compartment. You analyse how many cells can cross the membrane and reach the lower compartment. Usually in the bottom compartment you but a chemoattractant to stimulate the cells to move through the membrane. Invading cells will migrate and attach to the bottom of the membrane. The non invading cells will remain above and not anchor on the membrane. Boyden chamber system: It is made up by two chamber with a connective channel. You put a membrane in between the two chambers to close them. In the bottom chamber there will be the medium with the chemoattractant. Then after some days you measure how many cells are present in the bottom chamber. Trans-well system: There is only one chamber that will sit on the top of a well. You put the cells on the chamber and the chemoattractant on the bottom in the well. 3. Invasiveness assay You use the same experiments as shown in 2 but you also put a semi-permeable membrane and see how many cells cross the line/detect the signal from the chemoattractant in the bottom chamber. (usually the simplest chemoattractant will be serum (=energy)). If I put, in the precedent assay, an extra layer on the other side of the semi-permeable membrane composed by a gel (Matrigel) that imitate the extracellular matrix, I can evaluate the capacity of the cells to create a space for themselves in the gel. At the end of the experiment, if I find cells on the other side of the well, I understand that my cells are able to cross the pore-membrane and also to reorganize the “extracellular matrix” to pass on the other side. Non-invasive cells remain between the semi-permeable membrane because they can cross restricted space, but they can’t create a space in the matrix. I can also evaluate the capacity of a specific drug to increase/decrease the capacity of the cells to migrate or remain in the membrane. LECTURE 8 — 23/10 File 9 Resolution: minimal distance you can detect objects are sitting next to each other’s to consider them separated, is the minimum distance between 2 distinct points that still enables them to be seen as separate entities (resolves) >TYPE OF MICROSCOPES Microscopes can be divided essentially in 3. 1. Stereomicroscope It uses the light reflected from the object.. It is used for large samples because the tray is very large. It is used for the examination of organs, embryos, tissue slices or large tissue structures. You get a 3D effect because you'll get 2 different imagens in the binocular. The range of magnification is 20-100X. 2. Compound microscope Higher magnification than 1st one (40X-1000X). The light shines from the bottom onto the sample. There are 2 lenses which magnify the sample. In the upright microscope the light is coming trough the bottom of the sample and the ocular is positioned on top of the sample. This type of microscope is not recommended for live imaging of culture cell, but it is suggested for samples of slices. 3. Inverted compound microscope For cell culture we use the inverted microscopes. The light will come from above and illuminate the sample from the top. This microscope allows to have a small distance between the objective and the sample allowing the imaging of the culture dishes/plates used to grow the cell cultures in. >IMAGING TECHNIQUES 1. Phase contrast It allows to see the edges of a sample by augmenting the contrast of the sample. It is based on the amplification of the small shifts caused when the light encounters the sample present in the light path. The delay of the light to ge