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Cell bio
Midterm
| Question | Answer |
|---|---|
| What is Chromosome Territory (CT) | A distinct, three-dimensional region within the nucleus occupied by a single chromosome during interphase. |
| What is Chromosome intermingling | The process where chromatin from different chromosomes overlaps and mixes, potentially leading to interactions between chromosomes. |
| What is the significance of Interchromosomal interactions | These interactions can influence gene regulation, chromosome organization, and genome stability, potentially contributing to translocations. |
| What is Fluorescence In Situ Hybridization (FISH) | A technique used to detect and localize the presence or absence of specific DNA sequences on chromosomes. |
| What is Cryo-FISH | A high-resolution FISH technique applied to ultrathin cryosections (~150 nm thick) to better preserve chromatin nanostructure. |
| Microscopy methods to visualize chromosome intermingling. | Light microscopy (LM) and electron microscopy (EM) |
| How was Transcription activity measured in relation to chromosome intermingling | Immunolabeling of serine-2 phosphorylated RNA polymerase II (PolII), indicating active transcription sites. |
| Correlation between chromosome intermingling and translocations | The study found a significant correlation |
| Transcription's effect on chromosome organization | Transcription-dependent interactions influence the extent of intermingling, with active transcription stabilizing associations between specific loci. |
| What was observed about Less compact chromosome regions | Gene-rich and less-condensed chromosomal regions exhibited higher levels of intermingling. |
| Issue with standard 3D-FISH | It provides lower spatial resolution and can disrupt chromatin organization at the local level. |
| How did researchers confirm cryo-FISH did not disrupt chromatin structure | By comparing histone H2B distribution before and after the procedure, showing no significant changes. |
| Experimental control for transcription role in intermingling | Treatment with α-amanitin, an inhibitor of RNA polymerase II, to observe changes in chromosome intermingling. |
| Unified model proposed by authors about chromosome intermingling | The Interchromosomal Network (ICN) model, suggesting that chromosomes occupy territories that intermingle significantly, challenging the Interchromatin Domain (ICD) model, which posits separated compartments. |
| Proposed Impact of intermingling on genome stability | The proximity of double-strand breaks (DSBs) in intermixed regions increases the likelihood of interchromosomal rearrangements. |
| Main steps in the FISH procedure | Sample preparation, DNA denaturation, probe hybridization, and fluorescence visualization. |
| How is Cryo-FISH different from standard FISH | Cryo-FISH uses ultrathin cryosections (~150 nm) and preserves chromatin nanostructure by embedding cells in sucrose and freezing them. |
| Why is cryo-FISH more suitable for interchromosomal interactions | Because it preserves chromatin conformation, allowing for high-resolution visualization of overlapping chromosome territories. |
| Microscopy methods for visualizing cryo-FISH results | Light microscopy (for fluorescence) and electron microscopy (for immunogold labeling). |
| What is fluorescence anisotropy? | Fluorescence anisotropy tells us how much light stays lined up in one direction after a special kind of light (polarized light) shines on it. It helps measure how molecules move or change direction between absorbing and releasing light. |
| What is the excitation axis? | |
| What is polarization? | Polarization measures how much of the emitted light remains aligned in a particular direction after excitation and reflects how much rotational motion occurred during the excited state. |
| How is polarization mathematically defined? | I∥ is the intensity of light emitted parallel to the excitation axis. I⊥ is the intensity of light emitted perpendicular to the excitation axis. P |
| What causes fluorescence anisotropy in molecules? | Anisotropy arises from the specific orientation of the absorption and emission transition dipole moments within a molecule's structure. |
| How is fluorescence anisotropy utilized in biochemical research? | It is used to study molecular binding, protein dynamics, membrane fluidity, and molecular interactions by measuring changes in rotational diffusion. |
| What is a major limitation of fluorescence anisotropy? | Anisotropy can decrease due to rotational diffusion of molecules, energy transfer, or depolarization from reabsorption, making the interpretation of data complex. |
| Which factors can affect the anisotropy measurement? | Temperature, viscosity of the solvent, molecule size and shape, and the fluorophore's lifetime can all influence anisotropy. |
| What does the Perrin equation describe? | The Perrin equation describes how fluorescence anisotropy decreases as a function of molecular rotational diffusion. Helps study the size, shape, and dynamics of molecules by measuring how their rotational movement affects their fluorescence anisotropy. |
| What does the rotational correlation time (θ) tell us? | θ represents the time it takes for a molecule to rotate significantly during the excited state and depends on solvent viscosity, molecular size, and temperature. |
| What is the theoretical maximum value of anisotropy for a single-photon excitation? | The maximum anisotropy for colinear absorption and emission dipoles is r |
| How does rotational diffusion affect anisotropy? | Faster rotational diffusion during the fluorescence lifetime causes depolarization, decreasing anisotropy values close to zero for small molecules in low-viscosity solutions. |
| What methodology was used to analyze the nuclear pore complex (NPC)? | stochastic super-resolution microscopy and single-particle averaging to determine the average positions of labeled proteins within the NPC. |
| How were NPC components visualized? | Super-resolution fluorescence microscopy was used to observe labeled nucleoporins (Nups) in whole cells. |
| Why was single-particle averaging used? | It allowed the combination of thousands of NPC images to achieve higher resolution and determine protein positions with subnanometer accuracy. |
| What control was used to ensure fluorescence labeling did not alter NPC structure? | Researchers compared electron microscopy (cryo-EM) data with fluorescence-based measurements and found consistent molecular placements. |
| What contributes to uncertainty in super-resolution microscopy? | Photon counting noise + background noise |
| What is Point Spread Function(Airy Disc)? | The response of an optical system to a point source emitter |
| What is the Shannon-Nyquist sampling theorem? | A signal must be sampled at least twice its highest frequency to accurately reconstruct it without losing information. |
| What is pulse-chase labeling? | A technique to track protein synthesis and degradation over time using radioactive labeling. |
| How is protein synthesis labeled in pulse-chase experiments? | Cells are briefly exposed (pulsed) to [³⁵S] methionine, which gets incorporated into newly made proteins. |
| What happens after the pulse in pulse-chase experiements? | The radioactive media is washed out, and cells are incubated (chased) in normal media to track how long labeled proteins last. |
| How is protein degradation detected in pulse chase experiements? | Using SDS-PAGE or Western blotting, followed by film or camera detection of emitted radiation. It is shown by a smear showing degradation over time |
| What was the goal of the HCMV US11 study? | To investigate how the US11 gene of human cytomegalovirus (HCMV) causes degradation of MHC class I molecules, helping the virus evade immune detection. |
| Why does HCMV target MHC class I molecules? | MHC class I molecules present viral antigens to cytotoxic T cells. By removing these molecules, HCMV prevents immune recognition and destruction of infected cells. |
| What technique was used to track the fate of MHC class I molecules in US11-expressing cells? | Pulse-chase labeling with [³⁵S] methionine, followed by immunoprecipitation and SDS-PAGE, was used to observe the rapid degradation of class I molecules. |
| What control experiments were used in the US11 study? | -Uninfected cells to compare MHC class I stability. -Proteasome inhibitors (LLnL, Cbz-LLL, lactacystin) to confirm degradation was proteasome-dependent. -Brefeldin A (BFA) to block ER-to-Golgi trafficking and test if degradation happened before Golgi pr |
| How was MHC class I localization and degradation confirmed? | -Electron microscopy (EM) showed that US11 is localized in the ER. -Subcellular fractionation confirmed degraded MHC class I heavy chains moved into the cytosol, separate from US11. |
| What role does the proteasome play in US11-mediated MHC class I degradation? | The proteasome degrades class I heavy chains after they are removed from the endoplasmic reticulum (ER) to the cytosol, preventing their transport to the cell surface. |
| What effect did proteasome inhibitors have on MHC class I degradation in US11 cells? | When LLnL, Cbz-LLL, or lactacystin were added, MHC class I degradation was blocked, leading to the accumulation of a 40 kDa breakdown intermediate. |
| What evidence showed that MHC class I heavy chains undergo N-glycan removal before degradation? | -The 40 kDa breakdown intermediate lacked N-linked glycans. -N-glycanase activity was responsible for removing the glycans, as confirmed by endoglycosidase H (Endo H) resistance. |
| How did subcellular fractionation support the ER-to-cytosol dislocation of MHC class I? | -In normal cells, MHC class I heavy chains were found in membrane-bound fractions. -In US11-expressing cells, degraded class I molecules appeared in the 100,000 g supernatant (cytosol), proving they were extracted from the ER before degradation. |
| What are the 4 proposed mechanisms of US11-mediated MHC class I degradation? | 1. US11 binds newly synthesized MHC class I molecules in the ER. 2. Class I heavy chains are extracted from the ER into the cytosol. 3. N-glycanase removes N-linked sugars from class I heavy chains. 4. Proteasomes degrade the class I heavy chains, prev |
| What is subcellular fractionation? | A technique used to separate cellular components based on their size and density using centrifugation. |
| Why is subcellular fractionation used? | To isolate organelles (e.g., nucleus, mitochondria, ER, cytosol) for studying their functions, protein composition, and interactions. |
| What is Endoglycosidase H (Endo H)? | An enzyme that cleaves high-mannose N-linked glycans from glycoproteins in the endoplasmic reticulum (ER). |
| How does Endo H help track protein maturation? | If a protein is Endo H-sensitive, it means it is still in the ER.If it is Endo H-resistant, it has passed through the Golgi and acquired complex sugars. |
| What is a key limitation of Endo H? | It cannot remove complex N-linked glycans, so it only provides information about proteins that have not fully matured. |
| What was the goal of the ER-mitochondrial study? | To investigate how ER tubules mark mitochondrial division sites and actively participate in mitochondrial fission. |
| Why is ER-mitochondrial interaction important? | It helps regulate mitochondrial structure and distribution, which is crucial for energy production and cellular function. |
| How were ER-mitochondrial interactions visualized? | Using fluorescence microscopy, electron microscopy (EM), and tomography in yeast and mammalian cells. |
| What experimental controls were used in the ER-mitochondrial study? | -Cells lacking ER tubule-shaping proteins (Rtns, Yop1) to test if ER-mitochondrial contact still occurred. -RNAi knockdown of Drp1 and Mff to see if ER contact formed independently of fission machinery. |
| What did fluorescence microscopy reveal about mitochondrial division? | 87% (yeast) and 94% (mammalian) of mitochondrial division events happened at ER-mitochondrial contact sites. |
| How did electron microscopy support findings in ER-mitochondrial study? | 3D EM reconstructions showed ER tubules wrapping around mitochondria at sites of constriction before division. |
| How does the ER promote mitochondrial division? | -ER tubules wrap around mitochondria and constrict them before fission. -This constriction reduces the mitochondrial diameter, making it easier for Drp1/Dnm1 to complete division. |
| What is Drp1's role in mitochondrial fission? | Drp1 is a dynamin-related GTPase that forms helices around constricted mitochondria, driving membrane fission. |
| What happened to mitochondria when Drp1 was knocked down? | -Mitochondrial constriction still occurred at ER contact sites, but fission was incomplete. -This suggests ER defines division sites before Drp1 recruitment. |
| What is Mff's role in mitochondrial fission? | Mff is a mitochondrial outer membrane protein that recruits Drp1 to division sites. |
| What happened to mitochondria when Mff was knocked down? | -ER-mitochondrial contact sites still formed, proving ER defines division sites before Mff recruitment. -Mitochondria remained elongated, confirming Mff is needed for final fission. |
| What was the main goal of the RUSH study? | To develop a method for synchronizing and studying secretory protein trafficking in live cells using a retention and release system. |
| What are the three key components of the RUSH system? | 1. Reporter Protein - The protein of interest that moves through the secretory pathway. 2. Donor Compartment - The starting location (e.g., ER or Golgi) where the reporter protein is retained. 3. Streptavidin Hook - A fixed anchor that holds the reporte |
| How was the RUSH system visualized? | By tracking the ER-to-Golgi movement of a Golgi enzyme (ST-SBP-EGFP) using fluorescence microscopy and immunoelectron microscopy. |
| What experimental controls were used in RUSH? | -Non-biotin-treated cells to confirm that proteins remained in the donor compartment. -Cells expressing only the hook to ensure retention was specific to the RUSH system. |
| What did fluorescence microscopy reveal about secretory trafficking in RUSH? | -Reporter proteins remained in the ER without biotin. -Upon biotin addition, they moved synchronously to their target locations. |
| How did electron microscopy confirm trafficking in RUSH? | -Before biotin, reporters were seen in ER tubules. -After biotin addition, reporters appeared in the Golgi apparatus, confirming successful transport. |
| What was observed about different types of cargo in RUSH? | -Golgi enzymes (ManII, ST) reached the Golgi quickly and stayed. -Plasma membrane proteins (TNFα, E-cadherin) exited post-Golgi at different rates. |
| How does RUSH compare to other synchronization methods? | -Unlike temperature-based methods, RUSH works at physiological temperatures. -Allows precise timing and real-time tracking of protein movement. |
| What is the advantage of using biotin in the RUSH system? | Biotin is small, non-toxic, and diffuses freely, allowing controlled protein release without disrupting cellular function. |
| How does the RUSH system work? | 1. The reporter protein is fused to an SBP tag and a fluorescent protein (e.g., GFP or mCherry). 2. The hook protein is anchored in the donor compartment (e.g., ER, Golgi) and fused to streptavidin. 3. Without biotin, the SBP-tagged reporter binds tight |
| How is the donor compartment selected in RUSH? | ER Hook: Uses STIM1-NN, Ii-streptavidin, or KDEL-streptavidin to keep the reporter in the ER. Golgi Hook: Uses Golgin-84-streptavidin to retain proteins in the Golgi. |
| What does the Stable Compartments as Cisternal Progenitors model propose? | - The Golgi consists of stable compartments that do not undergo full cisternal maturation. -Cargo moves through Rab GTPase-mediated domain shifts instead of vesicular transport or cisternal progression. -Rab conversion allows compartments to transform, |
| What is Rab conversion, and how does it relate to the Golgi model? | Rab conversion is the gradual transformation of a compartment's identity through changes in Rab GTPases. |
| How does this model explain cargo movement in Golgi? | Cargo does not move between cisternae by vesicles or maturation. Instead, Rab domains shift, allowing cargo to transfer to different compartments without full cisternal progression. |
| What evidence supports the Stable Compartments as Cisternal Progenitors model? | -Rab GTPases create distinct Golgi domains with defined biochemical properties. -Endosome studies show Rab-driven domain conversion, supporting a similar process in the Golgi. -Megavesicles (large vesicular carriers) may facilitate bulk cargo transport. |
| What are the weaknesses of this model in the Golgi? | -Lack of evidence for megavesicles—they have not been widely observed in intra-Golgi transport. -Fails to explain transient Golgi structures in yeast and plants, which contradicts the idea of stable compartments. -Does not integrate COPI vesicle functio |
| Advantages for fluorescence microscopy | Allows live-cell imaging with fluorescent tagging. Can track mitochondria fission/fusion events, Golgi and ER morphology at a broad level, protein trafficking pathways |
| Spatial limitations for fluorescence microscopy | ~200-300nm (diffraction limit of LM) Cannot resolve details other than in larger vesicles |
| Temporal limitations for fluorescence microscopy | Allows to see movement in real time but photobleaching limits long term trafficking |
| Advantages of Super-Resolution Microscopy (STORM, PALM, Airyscan, etc.) | Breaks the diffraction limit of light microscopy, reaching ~10-50 nm resolution. Can provide high spatial precision for subcellular structures (Golgi, NPC, microtubules). |
| Temporal limitations of super resolution microscopy | No real-time monitoring at all due to slow image acquisitions and the phototoxicity damages the cells |
| Advantages of transmission electron microscopy | Ultra-high resolution (~1-2 nm), can visualize sub-organelle structures. Used to examine protein complexes, Golgi, ER-mitochondrial contacts, NPCs. |
| Temporal limitations of transmission electron microscopy | Only static images because the sample prep is destructive requiring thin sectioning (~50-100nm) and heavy metal staining. Also only provides 2D projections which loses spatial relationships |
| What are the advantages of electron tomography | Produces 3D reconstructions of subcellular structures at ~2-5 nm resolution. Useful for Golgi, NPC, ER-mitochondrial contacts. |
| Temporal limitations of electron tomography | Time consuming and requires many tilted images to reconstruct 3D volume which only works on fixed samples with no live imaging. |
| Advantages of immunoelectron microscopy | Combines high-resolution EM (~1-2 nm) with specific protein labeling using antibodies. Useful for tracking proteins inside Golgi, mitochondria, NPC, and ER. |
| Temporal limitations of immunoelectron microscopy | Only provides static images and only a fraction of the target proteins are labeled. Because it uses transmission electron microscopy its resolution is limited to ~1-2nm but labeling reduces efficiency |
| What is the diffraction limit of light? | wavelength(of X&Y)/2 Z |
| What would the resolution limit be for 500nm of excitation light? | 250nm for X and Y 500nm for Z |
| What is the wavelength and resolution of immunogold staining in transmission electron microscopy? | Wavelength - 0.004nm resolution - 0.1-0.01nm |
| How do you tag a protein with GFP? | Requires alpha-linker from the C- terminus of protein to N-terminus of GFP. Must not block functionality or folding |
| What is an angstrom? | 10^-10 m or 0.1nm |
| What are cons to particle average? | Not amenable to non-symmetric structures and 2D particle averaging does not assign 3D topology because there could be unsymmetrical structure in the z dimension |
| You are studying how a new drug affects protein degradation. How would you track the degradation rate of a specific protein over time? | Use pulse-chase labeling with radioactive amino acids (e.g., [³⁵S] methionine) to monitor protein turnover, followed by immunoprecipitation and SDS-PAGE. |
| You want to determine if a newly discovered protein is secreted through the Golgi. What experimental approach would you use? | Express the protein with a fluorescent tag, use live-cell imaging to track movement, and treat cells with Golgi transport inhibitors (e.g., Brefeldin A) to confirm Golgi involvement. |
| You are testing whether a protein moves from the ER to the Golgi before secretion. How would you confirm its trafficking route? | Use RUSH system with a biotin-dependent release mechanism, then track the protein's movement with fluorescence microscopy at different time points. |
| You hypothesize that mitochondria and lysosomes physically interact during cellular stress. What experiment would you design to test this? | Perform live-cell confocal microscopy using mitochondria- and lysosome-specific fluorescent dyes, followed by colocalization analysis. |
| You suspect that a mutation in a vesicle transport protein disrupts Golgi function. How would you test if Golgi trafficking is affected? | Express wild-type vs. mutant transport protein, track cargo movement using fluorescently labeled cargo proteins, and analyze the effect using super-resolution microscopy. |
| You are studying how a protein is degraded in the cytoplasm. What would be a strong positive control? | Treat cells with a known proteasome inhibitor (e.g., MG132) to block degradation, ensuring the accumulation of degradation intermediates. |
| You suspect a protein is involved in mitochondrial division. How would you confirm your results are specific to this protein? | Perform an RNAi or CRISPR knockout, then rescue the phenotype by reintroducing a wild-type or mutant version of the protein. |
| You believe a Golgi enzyme is required for proper glycosylation of a secreted protein. How do you test this? | Knock out the enzyme and use Endo H digestion to determine if the glycan structures remain immature (ER-type) instead of being processed in the Golgi. |
| You need to visualize nuclear pores at high resolution. What technique should you use? | Super-resolution microscopy (e.g., STORM, PALM) or Electron Microscopy (EM) for subnanometer resolution. |
| You want to track real-time protein movement in living cells. What method should you choose? | Live-cell fluorescence microscopy using GFP-tagged proteins. |
| You need to analyze subcellular structures at nanometer resolution but cannot use live-cell imaging. What technique should you use? | Transmission Electron Microscopy (TEM) or Electron Tomography for ultrastructural details. |
| You suspect a protein is moving between organelles and want to visualize interactions. What is the best imaging approach? | Colocalization analysis using dual-color fluorescence microscopy, or proximity labeling assays (e.g., APEX, BioID) for biochemical validation. |
| You want to measure how fast a protein moves within a membrane. What technique should you use? | Fluorescence Recovery After Photobleaching (FRAP) to measure diffusion rates. |
| You are testing whether a new drug promotes cell proliferation. What is a good and bad control? | |
| You designed a new fluorescent probe to label lysosomes. What is a good and bad control? | |
| You knock out a gene suspected to regulate Golgi vesicle trafficking. What is a good and bad control? | Good Control: -Use a wild-type (WT) cell line as a comparison. -Perform a rescue experiment by reintroducing the gene to see if trafficking is restored. Bad Control: -Using a completely different mutation unrelated to Golgi function as a control. -On |
| You treat cells with a new stress-inducing compound and observe mitochondrial fragmentation. What is a good and bad control? | Good Control: -Use an untreated control to establish baseline mitochondrial structure. -Use a known stressor (e.g., H₂O₂) as a positive control to compare fragmentation levels. Bad Control: -Using a different stressor that affects mitochondria differe |
| You suspect a protein shuttles between the Golgi and ER. What is a good and bad control? | Good Control: -Use a Golgi retention mutant to see if trafficking is disrupted. -Use Brefeldin A (BFA), which blocks Golgi transport, to confirm Golgi dependence. Bad Control: -Using cytosolic proteins as a control when they never localize to the Golg |
| You develop a new GFP-like protein and test its brightness in cells. What is a good and bad control? | Good control: -Compare against a well-established fluorescent protein (e.g., EGFP, mCherry) under identical conditions. -Ensure all samples are imaged with the same laser intensity and exposure time. Bad Control: -Comparing to a completely different f |
| You treat cells with a drug suspected to alter actin filament structure. What is a good and bad control? | Good Control: -Use an untreated control to compare baseline actin structures. -Use a known actin-disrupting drug (e.g., latrunculin A) as a positive control. Bad Control: -Comparing actin staining in two different cell types without accounting for nat |
| You mutate a nuclear pore protein and suspect it disrupts nuclear import. What is a good and bad control? | Good Control: -Use wild-type nuclear pore protein as a comparison. -Test with a known import-defective mutant as a positive control. Bad Control: -Using a random cytoplasmic protein as a control instead of an import-dependent protein. -Measuring nucl |
| You add a growth factor to cells and test for phosphorylation of a key signaling protein. What is a good and bad control? | Good Control: -Use untreated cells to measure baseline phosphorylation levels. -Use a kinase inhibitor to block signaling and confirm specificity. Bad Control: -Using overexpressed protein levels in the control, making it hard to detect activation dif |
| You suspect your drug induces cell death via apoptosis. What is a good and bad control? | Good Control: -Use untreated cells to establish baseline apoptosis levels. -Use a known apoptosis inducer (e.g., staurosporine) as a positive control. Bad Control: -Measuring total cell death without distinguishing between apoptosis and necrosis. -Us |
| You are comparing protein expression between treated and untreated cells using Western blot. What is a proper loading control? | Good Control: -Use a housekeeping protein (e.g., β-actin, GAPDH, or tubulin) to confirm equal protein loading across samples. Bad Control: -Using a nuclear protein (e.g., lamin A/C) as a loading control when analyzing cytoplasmic proteins. -Not normal |
| You overexpress a kinase and observe increased phosphorylation of a target protein. How can you confirm this is biologically relevant? | Good Control: -Include a dose-dependent overexpression series to ensure physiological relevance. -Express a catalytically inactive mutant to distinguish between kinase activity vs. non-specific overexpression effects. Bad Control: -Overexpressing the |
| You treat cells with a drug and observe phosphorylation of a key signaling protein. How do you ensure specificity? | Good Control: -Use a specific kinase inhibitor to block the pathway and confirm phosphorylation is drug-induced. -Use a genetic knockout or RNAi to test if the pathway still activates without the target protein. Bad Control: -Using total protein level |
| You suspect that overexpression of a protein alters its function compared to endogenous levels. How can you confirm this? | Good Control: -Use Western blot or qPCR to compare endogenous vs. overexpressed protein levels. -Use fluorescence microscopy to confirm localization does not change with overexpression. Bad Control: -Assuming that the overexpressed protein behaves ide |
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