Week 2: Methods in Cell Biology

Statistical Significance vs. Biological Significance

• Statistical Significance: A measure of the probability that an observed result occurred by chance. A low p-value (e.g., $p < 0.05$) suggests the result is unlikely due to random chance.

• Biological Significance: Refers to whether a statistically significant result has a meaningful impact or relevance in a biological context. A statistically significant result is not always biologically significant, and vice-versa.

• Example 1:

  • Sample sizes: $2000$ for both groups.

  • Means: $66$ inches and $65.5$ inches.

  • Standard deviation: $6$ inches.

  • p-value: $p = 0.0006305$.

  • Despite a very small p-value, indicating statistical significance, a $0.5$ inch difference in height with a standard deviation of $6$ inches might not be considered biologically meaningful.

• Example 2 (Outlier):

  • Sample sizes: $3$ for both groups.

  • Means: approximately $81$ inches and $71.5$ inches.

  • Standard deviation: approximately $6$ inches.

  • p-value: $p = 0.06858$.

  • This p-value is above the common threshold for statistical significance ($p > 0.05$), suggesting the observed $~9.5$ inch difference in means might be due to chance, likely influenced by the extremely small sample size despite a large mean difference. This highlights how small sample sizes can lead to a lack of statistical significance even with a large biological effect.

• Spurious Correlations: Not all observed correlations imply a meaningful relationship. An example given is the correlation ($r=0.980$, $r^2=0.961$, $p<0.01$) between the popularity of the first name 'Stevie' and Lululemon's stock price from $2008-2022$. This is a statistically significant correlation but has no biological or logical meaning.

Types of Cell Biology Research Questions

Cell biology aims to answer fundamental questions about cellular function and behavior, including:

• How does a specific cell, cell type, or subcellular structure behave?

• How does a biological process occur at the cellular level?

• What is the specific function of a protein within a cell?

• Which molecules (e.g., proteins, nucleic acids) does a particular protein interact with inside a cell?

• How do cells regulate various cellular processes?

• Where are proteins localized within a cell?

• How are genes or gene products influenced by changes in a cell's environment?

• How and why do cells interact with one another?

• Many other diverse questions.

Introduction to Microscopy

Fluorescence

• Concept: Fluorescence is the emission of light by a substance that has absorbed light or other electromagnetic radiation. It is a form of luminescence. In fluorescence, the absorbed light has a specific wavelength (excitation), and the emitted light has a longer wavelength (emission).

• Discovery: Discovered in creatures like Aequorea Victoria (a jellyfish) which led to the isolation of Green Fluorescent Protein (GFP).

• Fluorescent Proteins: Many fluorescent proteins exist, derived from organisms like jellyfish and coral, or evolved through methods like directed mutagenesis. They vary in excitation and emission wavelengths, producing different colors (e.g., EGFP, Citrine, Cerulean, mCherry, mPlum, TagBFP).

Green Fluorescent Protein (GFP)

• Isolated from Aequorea Victoria.

• Mechanism: Blue excitation light (e.g., $488 \text{ nm}$) is absorbed by a chromophore within the GFP structure. This causes the chromophore to emit green light (e.g., $509 \text{ nm}$).

• Applications: Bacteria expressing fluorescent proteins can create visual scenes (e.g., "San Diego beach scene"). EGFP (Enhanced GFP) is a commonly used variant.

Fluorescence Microscopy

• Principle: Cells are labeled with fluorescent molecules (e.g., fluorescent proteins, fluorescent dyes, or immunofluorescence). The sample is illuminated with a specific wavelength of light (excitation), which the fluorescent molecules absorb. These molecules then emit light at a different, longer wavelength (emission), which is visualized by the experimenter.

• Examples: Osteosarcoma cells stained with fluorescent phalloidin (an actin-binding toxin) can show actin filament distribution.

• Types (discussed for group work):

  • Epifluorescence Microscopy: Standard wide-field technique where excitation light is passed through the objective lens, and emitted light is collected back through the same objective.

  • Confocal Microscopy: Uses a pinhole to block out-of-focus light, providing optical sectioning and improved resolution in the z-axis, creating clearer 3D images.

  • Light Sheet Fluorescence Microscopy (LSFM): Uses a thin sheet of light to illuminate only a thin section of the sample, minimizing phototoxicity and photobleaching, and allowing for rapid 3D imaging of live, larger samples.

• General Considerations for all Fluorescence Microscopy:

  • Autofluorescence: Naturally occurring fluorescent molecules in the experimental system (e.g., quinidine in tonic water, vitamin B12, aromatic amino acids in beer like tyrosine, tryptophan, phenylalanine) can obscure specific signals.

  • Phototoxicity: Shining bright lights on cells can be toxic, altering their behavior or even killing them.

  • Photobleaching: Fluorescent proteins and dyes degrade over time when exposed to light, leading to a diminished or ceased fluorescent signal.

  • Perturbation: The act of imaging itself might cause cells to behave differently than they normally would. Confirming this requires careful experimental design.

  • Sample Compatibility: The cells or sample must be compatible with the physical and environmental constraints of the microscope.

Electron Microscopy (EM)

• Uses electrons instead of light to create images, offering much higher resolution.

• Transmission EM (TEM): Electrons pass through a very thinly cut sample. Provides detailed internal structure (e.g., poliovirus).

• Scanning EM (SEM): The sample's surface is coated with a conductive material and scanned with electrons. Provides high-resolution 3D surface images (e.g., pollen grains).

• Cryo-EM: The sample is plunge-frozen to super-cooled temperatures and imaged with electrons. Preserves biological structures in their native states without chemical fixation, allowing for high-resolution imaging of macromolecules (e.g., ARMAN cell).

Working with, Identifying, and Visualizing Proteins

Introducing Fluorescently Tagged Proteins into Cells

• Purpose: To visualize protein localization and dynamics within living cells.

• Methods:

  • Genome Integration (e.g., CRISPR-mediated):

    • Advantages: Preserves the native chromosomal environment of the gene, leading to more physiological expression levels. Minimally perturbs the target cell compared to overexpression systems.

    • Disadvantages: Generally more complicated and slower than plasmid-based methods (though CRISPR technology has made it faster and easier).

  • Plasmid Expression:

    • Advantages: Relatively fast and straightforward in many organisms. Easier than extensive genome engineering.

    • Disadvantages: Difficult to match the native expression level of the protein (can lead to overexpression artifacts). Expression levels can vary significantly from cell to cell, making quantitative analysis challenging.

• Process: A promoter drives transcription of a construct containing the gene for a favorite protein fused to a fluorescent protein (e.g., GFP). RNA polymerase transcribes the favorite protein-GFP mRNA, which is then translated into the favorite protein-GFP fusion protein, making the protein visible by fluorescence.

Protein Domains

• Definition: Autonomous parts of a protein with distinct structural, functional, and evolutionary characteristics. They are often compact folded units.

• Example: Pyruvate Kinase is a three-domain protein.

Immunofluorescence (IF)

• Problem it solves: Identifies specific proteins within complex cellular structures. For example, distinguishing microtubules from other filaments in a Transmission EM image of an epithelial cell periphery.

• Antibodies:

  • Definition: Immunoglobulins are a class of glycoproteins present in the serum or tissue fluids of all mammals. They are produced by plasma cells in response to an immunogen and function as antibodies, recognizing and binding to a specific region of an antigen called an epitope or antigenic determinant.

  • Classes: There are five distinct classes of antibody in higher mammals: IgG, IgA, IgM, IgD, and IgE. They differ in size, charge, amino acid composition, and carbohydrate content, but share a similar basic structure.

  • Structure: The basic structure of an antibody consists of two identical short polypeptides (light chains) and two identical long polypeptides (heavy chains), connected by covalent and non-covalent bonds.

  • Light Chains: Consist of two distinct regions: CL (Constant Light chain) in the C-terminal half and VL (Variable Light chain) in the N-terminal half, which shows much sequence variability.

  • Heavy Chains: Also composed of distinct constant and variable regions. The constant region of the heavy chain defines the class (IgG, IgA, IgM, IgD, and IgE) and subclass of the antibody.

  • Function & Bifunctional Nature: Antibodies exhibit highly specific binding to target antigens. The molecule is bifunctional: the Fab component is used to bind antigen, while the Fc region mediates the biological effect and is designated the effector region.

• Mechanism:

  • Fixation: Cells are fixed (e.g., with formaldehyde) to preserve cellular structures and prevent degradation.

  • Primary Antibody: An antibody directed against the protein of interest (antigen). This antibody (e.g., from a rabbit) binds specifically to an epitope on the antigen.

  • Secondary Antibody (Indirect Immunofluorescence): A fluorophore-coupled antibody that recognizes the primary antibody (e.g., an antibody against rabbit antibodies). The secondary antibody is from a different species than the primary to prevent cross-reactivity.

  • Direct Immunofluorescence: A less common method where the primary antibody itself is directly conjugated to a fluorophore.

• Advantages of Immunofluorescence:

  • High Specificity: Antibodies can specifically target and visualize individual proteins within a complex cellular environment.

  • High Sensitivity: Indirect immunofluorescence amplifies the signal because multiple secondary antibodies can bind to a single primary antibody.

  • Resolution of Fixed Structures: Provides excellent spatial resolution for fixed cellular structures.

  • Compatibility: Applicable to a wide range of sample types.

• Disadvantages of Immunofluorescence:

  • Fixed Cells Only: Cells must be fixed, meaning dynamic processes cannot be observed in real-time.

  • Potential for Artifacts: Fixation and permeabilization can sometimes alter cellular structures or protein localization.

  • Antibody Availability/Quality: Requires high-quality, specific antibodies, which are not always available for all proteins.

  • Background Signal: Non-specific binding of antibodies can lead to high background.

• Advantages of Immunofluorescence over Live-Cell Imaging:

  • Can use more robust dyes/fluorophores that are too toxic for live cells.

  • Allows for long-term storage of samples.

  • Often provides higher signal-to-noise ratio due to fixation and permeabilization.

• Advantages of Live-Cell Imaging over Immunofluorescence:

  • Observes dynamic cellular processes in real-time.

  • Avoids fixation artifacts.

  • Studies physiological conditions and cellular responses to stimuli.

Immunoprecipitation (IP)

• Purpose: To isolate a specific protein (antigen) and its interacting partners from a cell lysate, leveraging the natural antibody/antigen affinity. This method enables the purification of proteins, concentration of difficult-to-detect proteins (e.g., for Western blotting), and identification of protein-protein, protein-DNA, or protein-RNA interactions by immunoprecipitating their complexes.

• Workflow:

  1. Cell Lysis: Cells are lysed using a non-ionic denaturant to release cellular proteins while largely preserving protein complexes.

  2. Antibody Incubation: A protein mixture or cell lysate containing the protein of interest is incubated with a specific antibody solution. The antibodies will recognize and specifically bind to the corresponding antigens (proteins of interest).

  3. Capture: Antibodies are either previously immobilized on a solid-phase support (e.g., super-paramagnetic beads or non-magnetic agarose beads) or will be bound to Protein A or G, which has been immobilized on a solid-phase support. This solid support is sedimentable and thus indirectly precipitates the antibody/antigen complexes.

  4. Washing: The beads are washed to remove unbound proteins and cellular debris.

  5. Elution & Analysis: The captured protein complex is eluted from the beads by dissociating the complexes, and then analyzed using techniques like Western blot or mass spectrometry to identify the precipitated protein and its interacting partners.

• Types of Immunoprecipitation:

  1. Individual Protein Immunoprecipitation (IP): A general method involving an antibody to isolate and purify a particular protein of interest from a mixture (e.g., cell or tissue lysate).

  2. Co-immunoprecipitation (Co-IP): Also known as protein-complex immunoprecipitation, this powerful technique analyzes protein-protein interactions. It uses an antibody to target a specific protein that is a member of a protein complex, thereby isolating the entire complex and identifying unknown interacting members.

  3. Chromatin Immunoprecipitation (ChIP):

     • Enables the study of protein-DNA interactions, determining the location of DNA binding sites on the genome for a given protein of interest.

     • Mechanism: Relies on cross-linking between primary amines on amino acids and bases on DNA molecules using formaldehyde. Following cross-linking, chromatin is sheared, and a specific antibody precipitates the protein-DNA complex. Cross-links are reversible, allowing DNA fragments of interest to be dissociated from binding proteins. The DNA location can then be determined (e.g., by PCR).

  4. RNA Immunoprecipitation (RIP):

     • Targets RNA binding proteins to identify RNA bound by a specific protein.

     • Mechanism: RNA bound by a specific protein can be identified by RT-PCR (reverse transcription-PCR) from purified RNA in protein-RNA complexes.

  5. Tag-based Immunoprecipitation: Researchers can fuse tags (e.g., Glutathione-S-Transferase (GST)) onto either the C-terminal or N-terminal of the protein of interest. This approach allows a single, generic antibody against the tag to be used for different proteins, eliminating the difficulty of generating different specific antibodies each time.

Western Blotting

• Purpose: Detect and quantify specific proteins in a sample based on size and antibody specificity.

• Workflow:

  1. Protein Extraction: Cellular proteins are extracted from different experimental conditions.

  2. SDS-PAGE: Proteins are unfolded and coated with a negative charge by SDS (sodium dodecyl sulfate), then separated by size using gel electrophoresis. Smaller proteins migrate faster through the polyacrylamide gel.

  3. Transfer (Blot): The separated proteins are transferred from the gel onto a solid membrane (e.g., nitrocellulose or PVDF).

  4. Blocking: The free binding sites on the membrane are coated with a generic protein (e.g., milk powder or BSA) to prevent non-specific antibody binding.

  5. Primary Antibody Binding: The membrane is incubated with a primary antibody specific to the protein of interest.

  6. Washing: Excess primary antibody is washed away.

  7. Secondary Antibody Binding: The membrane is incubated with a secondary antibody that binds to the primary antibody and is conjugated to a detectable marker (e.g., enzyme, fluorophore).

  8. Washing: Excess secondary antibody is washed away.

  9. Visualization: The detectable marker is activated to visualize and quantify the specific protein (e.g., by chemiluminescence).

Mass Spectrometry (MS)

• Purpose: Measures the mass-to-charge ratio ($m/z$) of ions to identify and quantify molecules (e.g., proteins, peptides).

• Standard Mass Spectrometry (MS):

  1. A peptide mixture is introduced into an ion source, where molecules are ionized.

  2. Ions are sent to a mass analyzer, which separates them based on their $m/z$ ratio.

  3. A detector measures the abundance of ions at each $m/z$ ratio, producing a mass spectrum (relative abundance vs. $m/z$).

• Tandem Mass Spectrometry (MS/MS):

  1. A peptide mixture is ionized (MS1).

  2. A specific precursor ion (peptide) is selected by the first mass filter.

  3. The selected precursor ion is fragmented (e.g., by collision with an inert gas) into smaller product ions.

  4. The product ions are analyzed by a second mass analyzer (MS2).

  5. The fragmentation pattern (mass spectrum of product ions) is unique to a specific peptide, allowing its identification and sequencing.

Comparing Western Blotting and Mass Spectrometry

• Western Blotting vs. Mass Spectrometry (MS):

  • When Western Blotting is preferred: When a specific antibody to a protein is available, and you need to detect or quantify a single known protein, especially for routine analysis or to confirm presence and approximate size. It's often more affordable and simpler for targeted detection.

  • When Mass Spectrometry is preferred: For identifying unknown proteins, complex mixtures of proteins (proteomics), post-translational modifications, and quantifying proteins with high accuracy without requiring specific antibodies. It's more comprehensive for discovery-based research.

• Co-immunoprecipitation (Co-IP) False Positives:

  • Why proteins identified might not actually interact in the cell (three possibilities):

    1. Non-specific binding to the antibody or beads: Proteins might bind non-specifically to the antibody or the Protein A/G beads, leading to their co-purification.

    2. Transient or weak interactions: Proteins might interact in a very transient or weak manner that is amplified and stabilized under lysis conditions, or they might not be direct interactors but part of a larger complex.

    3. Indirect interactions: The identified protein might not directly interact with the protein of interest but interacts with another protein that does interact with the target protein, thus being pulled down as part of a larger complex.

  • Why false positives might vary between cell types (three possibilities):

    1. Differential expression of interacting partners: The false positive protein might be highly expressed in one cell type but not another, leading to its accidental co-purification in the high-expression scenario.

    2. Different cellular environments/crowding: The overall protein composition or cellular crowding might differ, influencing non-specific protein interactions with the antibody or beads.

    3. Altered protein complex composition: The bait protein might be part of different multiprotein complexes in different cell types, leading to varying indirect associations or stabilization of non-specific binding partners.

Western Blot theory overview

Similar to ELISAs, western blot uses antibodies to detect and analyze specific proteins in a given sample of tissue or cell extract. This technique uses gel electrophoresis to separate a mixture of proteins, followed by transfer to a PVDF or nitrocellulose membrane. The membrane then serves as a platform on which antibodies can then be used to probe for target proteins. This guide will illustrate several major steps in western blot, including tissue preparation, gel electrophoresis and transfer, and immunoblotting.

Western Blot procedures:

1. Tissue preparation

   • Protein samples could be obtained from various sources, such as whole tissue or cell culture. Bacterial, viral or environmental samples could also be a source of proteins used in western blot.

   • In general, tissues or cells are firstly lysed by sonification or by using a homogenizer to release the proteins of interest.

   • Assorted detergents, salts, and buffers are required for lysis of cells and solubilization of proteins.

   • In addition, protease and phosphatase inhibitors are often added to prevent protein denaturation and degradation.

2. Gel electrophoresis

   • The separation of molecules is dependent on differences in their isoelectric points, molecular weights and/or electric charges.

   • In western blotting, the most commonly used separation methods are denatured gel electrophoresis such as SDS-PAGE and native gel electrophoresis such as PAGE.

   • SDS is a negatively charged anionic detergent which binds proteins, denaturing them and giving them a uniform negative charge. Therefore, separation of proteins in SDS-PAGE only depends on their molecular weights.

   • On the other hand, native PAGE separates proteins by their charge-to-mass ratio.

   • The choice between denatured and native gel electrophoresis depends on which epitope the antibody recognizes. If the epitope resides within the native conformation of the protein, protein denaturation is required to enable access to the antibody. By contrast, if the epitope is made up of non-contiguous amino acids and these amino acids are close to each other in the native structure of the target protein, a native gel electrophoresis should be used.

3. Transfer

   • In order to facilitate access of the antibody to the target protein, separated proteins should be transferred from the gel onto a membrane such as nitrocellulose or PVDF (polyvinylidene difluoride).

   • The transfer can be done in wet or semi-dry conditions and by capillary action or applying electric current.

4. Immunoblotting
   4.1. Blocking
   - Blocking of unoccupied sites prevents non-specific binding (high background) of primary and/or secondary antibodies to membrane.
   - Bovine serum albumin (BSA) or non-fat milk solution is commonly used.
   - Addition of a dilute detergent, such as Tween 20, may also help in improving recognition of antigens by antibodies.
   4.2. Detection
   - Normally, secondary antibodies are conjugated to a reporter enzyme and the detection method used is based on the type of enzyme.
   - The most commonly used enzyme for detection in western blotting is horseradish peroxidase (HRP), which reacts with its substrates to produce light. A photographic film is placed against the membrane and an image is created from the reaction.