CMBI 6617 - Lab Notebook Flashcards 2.0

This set of flashcards covers key vocabulary and concepts from the CMBI 6617 course, focusing on various lab techniques and terminology related to cell culture and experiments conducted during the semester.

What types of cells are commonly used in cell culture and what are the general methods for growing them?

Commonly used cells include established cell lines like MDA-MB-231 (human breast cancer cells), HeLa, HEK293, and primary cells. Methods for growing cells in cell culture involve:

1. Sterile Environment: Performed in a laminar flow hood to prevent contamination.

2. Cell Culture Media: Contains nutrients, growth factors, salts, and buffers (e.g., DMEM, RPMI-1640).

3. Supplements: Fetal Bovine Serum (FBS) provides growth factors, antibiotics prevent bacterial contamination (e.g., penicillin/streptomycin), and L-glutamine is an essential amino acid.

4. Incubation Conditions: Maintained at 37\degree C in a humidified incubator with 5\%\nCO_2 to regulate pH.

5. Culture Vessels: Tissue culture flasks, dishes, or plates treated for cell adhesion.

6. Regular Feeding: Replacing old media with fresh media to provide nutrients and remove waste products.

What is cell passage and what is its purpose?

Cell passage, also known as subculturing, is the process of transferring cells from one culture vessel to another. This is necessary when cells reach confluency (covered the available surface) or when the media is depleted, to provide fresh nutrients and space for continued growth and propagation of the cell line.

How do you estimate cell counts and concentration during cell passage?

Cell counts and concentration are estimated using a hemocytometer. After detaching cells (e.g., with Trypsin) and resuspending them, a small volume of cell suspension is loaded into the hemocytometer. Cells within specific squares are counted under a microscope, and the following formula is used:

\text{Cell Concentration (cells/mL)} = \frac{\text{Average count per square} \times \text{Dilution factor}}{ \text{Volume of 1 square (mL)} }

Where the volume of one large square is 10^{-4} mL.

How is cell viability determined during cell passage?

Cell viability is commonly determined using the Trypan Blue Exclusion assay. Trypan Blue dye enters cells with compromised cell membranes (dead cells) but is excluded by intact membranes (live cells). Live cells appear bright and refractile, while dead cells appear blue. The percentage of viable cells is calculated as:

\text{Viability (\%)} = \frac{\text{Number of viable cells}}{\text{Total number of cells (viable + dead)}} \times 100

Explain cell doubling time and its formula.

Cell doubling time (T_d) is the time it takes for a cell population to double in number during its exponential growth phase. It is a key parameter for characterizing cell line growth. The formula for doubling time is:

D = ln(2)/k

Where:

k = growth rate constant (k = ln(N/No)/t), and N is the final cell number, No is the initial cell number.

Describe general cell staining techniques and procedures.

Cell staining involves using dyes or fluorescent probes to selectively color or label specific cellular components (e.g., nuclei, mitochondria, cytoskeleton) to visualize them under a microscope. Procedures typically involve:

1. Cell preparation: Cells are cultured and often fixed (e.g., with paraformaldehyde) to preserve their structure, followed by permeabilization (e.g., with Triton X-100) if intracellular components need to be stained.

2. Staining: Cells are incubated with the specific stain or fluorescent probe.

3. Washing: Excess stain is removed by washing with buffer (e.g., PBS).

4. Imaging: Stained cells are observed using brightfield or fluorescence microscopy.

How is cell density determined using Crystal Violet Staining?

Crystal Violet staining is a colorimetric method used to quantify adherent cell density. The procedure involves:

1. Fixation: Cells are fixed to the culture plate using a solution like methanol or formaldehyde.

2. Staining: Fixed cells are stained with Crystal Violet solution, which binds to DNA and proteins, primarily staining cell nuclei.

3. Washing: Excess stain is thoroughly washed away, leaving only the stain associated with the fixed cells.

4. Solubilization: The bound stain is then solubilized (e.g., with a detergent like SDS or acetic acid).

5. Quantification: The absorbance of the solubilized dye is measured spectrophotometrically (typically at 540-590 nm), where absorbance is directly proportional to the number of cells. Standard curve ( x = abs - y / m )

What are common types of cell contamination and how can they be avoided?

Cell culture contamination refers to the presence of unwanted biological agents or substances that compromise the purity and viability of cell cultures. Common types include:

• Bacterial/Fungal Contamination: Identified by turbidity in media, sudden pH changes, and spores/mycelia. Often transmitted through aerosols, non-sterile handling, or contaminated reagents.

• Mycoplasma Contamination: Microorganisms lacking cell walls, difficult to detect visually, often causing chronic insidious effects on cell growth and morphology. Detected by PCR or specific assay kits.

• Viral Contamination: Often latent, identified by specific CPE (cytopathic effects) or molecular detection.

• Cross-Contamination: Mixing of different cell lines, leading to misidentification.

Procedures to avoid contamination:

1. Aseptic Technique: Strict adherence to sterile procedures in a laminar flow hood.

2. Sterile Reagents and Equipment: Use of sterile media, buffers, plastics, and autoclaved glassware.

3. Disinfection: Regular cleaning of the laminar flow hood and incubator with 70\%\nethanol.

4. Quarantine: Quarantining new cell lines or reagents until proven contamination-free.

5. Regular Monitoring: Visual inspection of cultures for signs of contamination.

6. Antibiotics/Antifungals: Inclusion in cell culture media, though overuse can mask contamination.

Describe cryopreservation methods, freezing media, and protocols for cell maintenance after cryopreservation.

Cryopreservation is the process of preserving cells by cooling them to very low temperatures (typically -196\degree C in liquid nitrogen) to halt metabolic activity and maintain viability for extended periods.

• Freezing Media: Typically consists of cell culture medium, a cryoprotective agent like DMSO (Dimethyl Sulfoxide, 5-10\% volume/volume) or glycerol, and serum (e.g., FBS). Cryoprotectants reduce ice crystal formation and protect cells from damage during freezing and thawing.

• Freezing Protocol: Cells are suspended in freezing media and subjected to a slow, controlled freezing rate (e.g., -1\degree C/minute) to minimize intracellular ice formation. This can be achieved using a "Mr. Frosty" container placed in a -80\degree C freezer, before transfer to liquid nitrogen.

• Thawing Protocol: Cells are rapidly thawed (e.g., in a 37\degree C water bath) to minimize osmotic shock and then immediately transferred to warmed growth media, centrifuged to remove DMSO, and resuspended in fresh media. Rapid thawing is crucial to prevent the growth of ice crystals.

• Cell Maintenance After Cryopreservation: After thawing, cells are seeded into culture vessels, monitored for attachment and growth, and the media is changed frequently in the initial days to remove residual cryoprotectant and replenish nutrients. Viability and growth rate should be assessed.

Explain the electroporation technique and its basic protocol parameters.

Electroporation is a technique that uses short, high-voltage electrical pulses to transiently permeabilize cell membranes, creating pores that allow the uptake of foreign molecules, such as DNA, RNA, or drugs, into the cells.

Basic Protocol Parameters:

1. Voltage: The strength of the electric field applied (e.g., in Volts/cm). Higher voltage increases pore formation but can also increase cell death. 160 V were used in this lab.

2. Pulse Duration: The length of time each electrical pulse is applied (e.g., microseconds to milliseconds). 15 msec in this lab.

3. Number of Pulses: The total count of electrical pulses delivered.

4. Waveform: The shape of the electric pulse (e.g., square wave, exponential decay wave).

5. Cell Density: The concentration of cells in the electroporation cuvette, which affects efficiency and viability.

6. Electroporation Buffer: A specialized buffer designed to maintain cell viability and conductivity during the process.

Describe immunocytochemistry methods of detection, focusing on NucBlue, Mitotracker, and ActinGreen 488 staining.

Immunocytochemistry (ICC) uses specific antibodies to visualize cellular components. When these antibodies are directly or indirectly conjugated to fluorescent dyes, it's called immunofluorescence.

Methods of Detection for Specific Stains:

• NucBlue (Hoechst, DAPI): These are nuclear stains (fluorescent dyes that bind to DNA). They are cell-permeable and are used to label the nuclei of live or fixed cells, making them useful for visualizing cell density, nuclear morphology, or DNA condensation. They emit blue fluorescence when bound to DNA.

• Mitotracker: This is a family of fluorescent dyes that selectively stain mitochondria in live cells. They accumulate in active mitochondria due to their membrane potential. Different variants are available (e.g., MitoTracker Green FM, Deep Red FM), allowing specific visualization of mitochondrial structure and assessing mitochondrial health.

• ActinGreen 488 (Phalloidin-conjugated): Phalloidins are toxins that bind with high affinity and selectivity to filamentous actin (F-actin), a major component of the cytoskeleton. ActinGreen 488 is a phalloidin conjugate that fluoresces green when excited, allowing for the visualization of actin filaments in fixed and permeabilized cells. This helps study cell morphology, motility, and cytoskeletal organization.

How is cell death assessed using immunofluorescence, and what dyes are commonly used to visualize dead cells?

Cell death can be assessed using immunofluorescence by detecting specific markers associated with different cell death pathways (e.g., cleaved caspases for apoptosis). Additionally, specific dyes are used to visualize dead cells based on membrane integrity.

Common Dyes to Visualize Dead Cells:

1. Trypan Blue: (Already covered, stains cells with compromised membranes).

2. Propidium Iodide (PI): A fluorescent nucleic acid stain that is impermeable to live cells but enters cells with damaged membranes (dead or dying cells), binding to DNA and fluorescing red. It is often used with Annexin V for distinguishing early apoptotic, late apoptotic, and necrotic cells.

3. 7-Aminoactinomycin D (7-AAD): Similar to PI, it is a fluorescent dye that binds to DNA but is excluded from live cells with intact membranes. It enters dead or dying cells and fluoresces red/far-red.

4. Ethidium Homodimer-1 (EthD-1): Another membrane-impermeant nucleic acid stain that strongly fluoresces upon binding to DNA, commonly used in live/dead assays alongside a live-cell dye (e.g., calcein AM) to identify non-viable cells.

5. NucBlue and NucGreen Dead 488: Blue nucleus stain on all cells, Green nucleus stain only dead/dying cells due to compromised membrane.

What is the purpose of performing a cytotoxicity assay, why use AlamarBlue, how are drugs diluted, and what mathematical formulas are used?

A cytotoxicity assay is performed to determine the capacity of a substance (e.g., a drug, chemical, or treatment) to cause damage to cells or to kill them. It helps assess drug efficacy, screen potential therapeutics, and evaluate the toxicity of compounds.

• Why AlamarBlue? AlamarBlue (Resazurin) is a colorimetric and fluorometric indicator used to measure cell viability and metabolic activity. Metabolically active cells reduce the non-fluorescent, blue resazurin to highly fluorescent, pink resorufin. It is preferred because it is non-toxic to cells, allowing for kinetic measurements over time on the same sample, and is highly sensitive and quantitative.

• Drug Dilution: Drugs are typically diluted from a concentrated stock solution to obtain a series of desired concentrations. A common method is serial dilution, where a stock solution is diluted stepwise to achieve decreasing concentrations.

• Mathematical Formulas: The primary formula for preparing drug stocks and dilutions is:

  C1V1 = C2V2

  Where:

  • C_1 = initial concentration (stock concentration)

  • V_1 = initial volume (volume of stock to use)

  • C_2 = final desired concentration

  • V_2 = final desired volume

  For preparing a stock solution from a solid compound, the mass can be calculated using:

  \text{Mass (g)} = \text{Concentration (mol/L or M)} \times \text{Molecular Weight (g/mol)} \times \text{Volume (L)}

What is the purpose of performing a migration assay, and what reagents are involved?

The purpose of performing a migration assay (e.g., Wound Healing Assay or Transwell Assay) is to study cell motility and migration capabilities, which are crucial processes in wound healing, immune responses, embryonic development, and cancer metastasis. It helps investigate the effects of different treatments or genetic manipulations on cell movement.

Reagents Involved:

• Cell Culture Media: Standard growth media, often serum-free or with reduced serum to minimize proliferation effects over migration.

• Growth Factors/Chemoattractants: Substances that stimulate or inhibit cell migration (e.g., EGF, PDGF, specific inhibitors).

• Wounding Tool: For wound healing assays, a sterile pipette tip or specialized scratch tool is used to create a gap in a confluent cell monolayer.

• Imaging System: A microscope for capturing images at different time points to track cell movement.

Describe the Chromosome Spread Assay and its important reagents.

The Chromosome Spread Assay (also known as metaphase spread) is a cytogenetic technique used to visualize and analyze chromosomes for numerical and structural abnormalities. It involves arresting cells in metaphase, swelling them, and then spreading their chromosomes on a microscopic slide for examination.

Important Reagents:

• Colchicine (or Colcemid): A microtubule inhibitor used to arrest cells in metaphase, where chromosomes are condensed and visible.

• Hypotonic Solution (e.g., 0.075 M KCl or 0.06 M Sodium Citrate): Used to swell the cells, which helps in dispersing the chromosomes evenly when cells are lysed on the slide.

• Fixative (e.g., Methanol:Acetic Acid, 3:1 ratio): Used to fix the cells and lyse the cell membrane, allowing the chromosomes to spread out from the nucleus.

• Dye/Stain (e.g., Giemsa, DAPI, Hoechst): Used to stain the chromosomes to make them visible and to reveal banding patterns for detailed analysis.

Provide general information about chromosomes and types of chromosomal defects.

Chromosomes are thread-like structures located inside the nucleus of eukaryotic cells. They are made of DNA tightly coiled many times around proteins called histones that support its structure. Chromosomes carry genetic information in the form of genes.

• Structure: Each chromosome typically consists of two sister chromatids joined at a centromere, especially during metaphase.

• Number: Humans generally have 46 chromosomes (23 pairs): 22 pairs of autosomes and 1 pair of sex chromosomes (XX for females, XY for males).

Types of Chromosomal Defects:

1. Numerical Aberrations (Aneuploidy): Involve an abnormal number of chromosomes.

   • Trisomy: Presence of an extra copy of a chromosome (e.g., Trisomy 21 or Down syndrome with 3 copies of chromosome 21. Klinefelter syndrome in males extra X chromosome XXY).

   • Monosomy: Absence of one copy of a chromosome (e.g., Turner syndrome).

2. Structural Aberrations: Involve changes in the structure of chromosomes.

   • Deletions: A segment of a chromosome is lost.

   • Duplications: A segment of a chromosome is copied, resulting in extra genetic material.

   • Inversions: A segment of a chromosome is reversed end to end.

   • Translocations: A segment of one chromosome breaks off and attaches to another chromosome.