Lab Exam 1

What is accuracy? What is precision?

• accuracy is a measure of how close a measurement comes to the actual value of what’s being measured

• precision is a measure of how close a series of measurements are to one another (irrespective of the actual value)

What is error? How is it calculated? How do we calculate percent error and what is it?

• error is the difference between the experimental value and the accepted value

  • error = experimental value - accepted value

• percent error is the absolute value of the difference, divided by the accepted value and multiplied by 100%

  • percent error = [ |measured - accepted| / accepted] x 100%

What is repeatability? What is reproducibility?

• repeatability is the variability inherent in the measurement system under constant conditions

• reproducibility is variability among measurements made under different conditions

What is the dilution equation? What is it used for?

• the dilution equation is used to calculate volumes or concentrations needed to get a desired dilution

• C1V1 = C2V2

  • looks at initial concentrations and volumes and final concentrations and volumes

How do you do weight per volume calculations?

• % weight per volume = weight of solute (g) / volume of solution (mL)

  • can rearrange to solve for anything (just convert %w/v to a decimal)

What is transformation? How was this shown in Griffith’s experiment? What is conjugation?

• transformation is the genetic change in a cell due to the intake of DNA from the environment

• when Griffith injected mice with heat-killed S-strain and live R-strain, the mice died and when he analyzed them, he observed live S-strain bacteria → there must have been transformation to change the nonvirulent strain (R) into the virulent one (S)

• conjugation is “bacterial transformation” in which bacteria take in plasmids from other bacteria

What are plasmid vectors? What are recombinant plasmids?

• plasmid vectors are plasmids used for experimental purposes to transform bacteria with foreign DNA

• recombinant plasmids are vectors that have DNA fragments inserted into them with the use of restriction enzymes

What are the components that recombinant plasmids usually have?

• origin of replication: where replication begins

• antibiotic resistance gene: allows the bacteria that take in the plasmid to survive on plates in the presence of an antibiotic → positive selection

• multiple cloning site (MSC): aids in DNA insertion by containing multiple sites for restriction enzymes to get the plasmid

• promoter: drives transcription of the inserted gene

• selection marker: helps determine which bacteria contain the plasmid (fluorescent protein or additional antibiotic resistance gene)

• gene of interest: codes for the desired protein for expression

What is gateway cloning?

• gateway cloning is a new method of transformation that allows the insertion of multiple DNA fragments into different vectors as well as transfer of DNA sequences between plasmids

What is competency? What is natural vs artificial competency?

• if a cell is competent, it means it’s capable of taking in exogenous DNA molecules from their environment

  • natural competence: bacteria have cellular machinery to take up DNA from the environment

  • artificial competence: cells are made competent in the lab, allowing them to take up DNA

How do we accomplish artificial competence? What are the two main approaches?

• calcium chloride + heat shock approach:

  • cells are incubated on ice in a calcium chloride solution, which causes cells to become permeable to DNA molecules (weakens the cell wall)

  • cells are pelleted by centrifugation, supernatant is removed, and plasmid DNA is added

  • the mixture is briefly heated to 42 C for 50-60s, followed by rapid cooling on ice → this heat shock causes the DNA to transfer across the cell’s wall and membranes

  • the cells are placed at 37 C to allow them to re-seal their membranes

• electroporation: cells are subject to electric shock to perforate the membranes and plasmids enter through the temporary holes

  • note: this method allows for efficient transformation of large plasmids

How does the heat shock method work?

• by exposing cells to a sudden increase in temperature (heat shock), a pressure difference between the outside and inside of the cell is made

  • this induces the formation of pores, which plasmid DNA can enter through

• after cells return to a normal temperature, the cell wall will self-heal

How do we induce ampicillin resistance?

• we insert the bla gene into a plasmid to make bacteria resistant to ampicillin

• the bla gene encodes for beta lactamase, which breaks down the beta-lactam ring in ampicillin, inactivating it

Describe the GLO plasmid. What does it allow cells to do? How is it regulated?

• the GLO plasmid (pGLO) encodes for GFP and bla gene (ampicillin resistance)

• pGLO also includes a gene regulation system that allows us to control when GFP is expressed

  • when arabinose sugar is added to the medium, GFP expression is turned on (cells appear green when arabinose is in medium under UV light)

How is the arabinose operon activated? How is it repressed?

• activation: arabinose binds the activator protein (AraC) and this complex helps RNA polymerase bind to the promoter, turning on the ara operon

  • note: activation alos depends on cyclic AMP

• repression: without arabinose, AraC protein binds araI and araO regions and this forms a loop that blocks transcription

What is solid media used for? What does streak culturing involve?

• solid media is used to generate discrete colonies (collections of bacteria originating from a single bacteria cell)

• streak culture consists of spreading source material over an agar surface with a metal loop until one organism at a time falls off of the loop, then the medium is incubated until colonies arise

  • each colony represents a single type of microorganism that originated from a single cell

  • note: source materials contain a large number of microorganisms so separated colonies are obtained over the last series of streaks (first ones yield a confluent lawn of growth)

What is liquid media used for? What does liquid culturing involve?

• liquid media is often used for diffuse growth → once we’ve isolated discrete colonies with solid medium, we can expand a population in liquid medium

  • liquid cultures are also used for blood cultures and sterility tests

• liquid cultures are inoculated by touching colonies with a metal loop and then touching the media, or by adding inoculum with pipettes or syringes

  • disadvantage: doesn’t provide a pure culture from mixed inocula

What is pour plate culturing used for? How does it work?

• in pour plate cultures, agar media is melted and cooled to 45C and inoculum is added to molten agar and mixed

  • after poured and set into a petri dish, we incubate the plate at 37 C to allow colonies to appear throughout the depth of the medium

• this culture type gives us an estimate of the viable bacterial count in a suspension and it’s used for tests like quantitative urine cultures

What are the ways we can categorize media?

• by consistency: solid, liquid, or semi-solid medium

• by constituents/ingredients: simple, complex, synthetic or defined, or special (enriched, selective, differential, etc.)

• by oxygen requirements: aerobic or anaerobic media

What is serial dilution? How does it work and why do we do it?

• serial dilution is a process through which the concentration of an organism is systematically reduced through successive resuspension in fixed volumes of liquid diluent

  • we have 10 mL of solution with cells and medium → 1 mL of this is added to another tube of 9 mL of medium → repeat

• serial dilution is the key to enumeration of bacteria since mixed samples contain an unknown, often large, number of bacteria

What is streak plating? What is spread plating? How do each work?

• streak plating enables the isolation of bacteria within a sample

  • we introduce a diluted sample to a section of solid medium and then we spread the inoculum over each third of the plate in a zig-zag pattern → as different sections are streaked, the sample is spread more thinly (only need to streak from one dilution to achieve individual colonies in later sections)

  • after incubation, the streaked plates allow for observations of colony morphology so we can differentiate between bacterial species

• spread plating allows us to enumerate the bacteria in a sample

  • an aliquot of a sample is spread evenly over the entire plate

  • after incubation, enumeration can be performed (count colonies)

    • note: counts under 30 or over 300 should be discarded

Generally, how do genome editing techniques work?

• these techniques induce double-stranded breaks in the DNA with nuclease enzymes that target specific sites (with the help of proteins or RNA guides)

• when a cell attempts to repair the damage, mutations can be introduced in the target region of DNA — this allows us to induce mutations in targeted regions

How does homologous recombination work?

• homologous recombination is a classic method for introducing genetic changes to specific sequences

• here, we can incorporate homologous sequences into the targeting construct → this results to a swap of the endogenous gene for the altered one

How do cells repair double-stranded breaks? What are the two repair types?

• cells use endonucleases to repair double-stranded breaks

• in non-homologous end joining, broken DNA ends are resealed directly — note: not usually perfect and could result in bases being added or delated (mutating the target sequence)

• in homology-directed repair, damage is fixed by copying from a homologous template — allows researchers to add a template to direct specific changes to the target site

What are the three major genome editing techniques? How do they work generally?

• zinc fingers: we can fuse DNA-binding zinc finger domains of transcription factors to the DNA-cleaving domain of the FokI endonuclease — linking the two together gives us zinc finger nucleases that target unique DNA sequences

• TALENs (transcription activator-like effector nucleases): these nucleases join FokI to the variable DNA-binding domains of bacterial transcription activator-like effectors (TALEs) — each TALE domain recognizes a single, unique DNA base and this allows TALENs to have sequence specificity

• CRISPR-Cas9 system: pieces of invading foreign DNA (protospacers) are incorporated into a CRISPR locus, which is transcribed and processed by protein-RNA machinery into small RNAs called crRNAs

Why are TALENs and zinc finger nucleases not used as much now? Why did CRISPR-Cas9 gain popularity?

• TALENs and zinc finger nucleases are complex and time consuming to engineer, which limits their use

• the customizability of the RNA sequence-based CRISPR-Cas9 system provides a clear advantage over the protein-based genome editing methods

  • CRISPR-Cas9 is simpler, more versatile, and cost effective, so it is more accessible and widely used

What is the origin of CRISPR-Cas9?

• in bacteria and archaea, the CRISPR-Ca9 system serves as an adaptive immune response against invading viruses

• after infection, they capture short sequences of viral DNA to create a library of segments (CRISPR arrays)

• when re-exposed to the virus, CRISPR arrays are used to transcribe small RNA segments that help guide Cas9 to viral invaders for destruction

What are the three stages of CRISPR Cas9?

• in the acquisition stage, the protospacer region of viral DNA is cleaved by CRISPR systems — the protospacer to be cleaved is identified by the protospacer adjacent motif (PAM) present in the target viral DNA — the cleaved protospacer region is incorporated into the bacterial CRISPR locus

• in the expression stage, the CRISPR and Cas genes are transcribed to make pre-CRISPR RNA (crRNA) and Cas9 mRNA, then the pre-crRNA is processed to get mature crRNA

• in the interference stage, crRNA and the translated Cas9 protein form a ribonucleoprotein complex that targets and cleaves the viral DNA in a sequence-specific manner

What are the three types of CRISPR-Cas9 systems?

• in Type I systems, Cas3 has helicase and nuclease activity and many other Cas proteins make double-stranded DNA breaks in viral DNA

• in Type II systems, Cas9 acts alone to cleave the DNA

  • on top of crRNA, these systems have trans-activating CRISPR RNA (tracrRNA), which is required to mature the crRNA

• in Type III systems, Cas10 has unknown function but needs many proteins for DNA cleavage (like type I)

  • can also target RNA for cleavage

• note: type I and III are in bacteria and archaea whereas type II is only in bacteria

How is CRISPR-Cas9 technology used in the lab commonly?

• we use this system to remove DNA and insert a new sequence

• we create a guide RNA with a guide sequence that binds to a sequence on the target DNA

• we administer the guide RNA and Cas9 protein so that the guide RNA identifies the target sequence and Cas9 cleaves it

• the cell can then either repair the broken strands by inserting or deleting random nucleotides (makes the gene inactive)

  • OR we can introduce a customized DNA sequence to the cell with the RNA and Cas9 protein that can serve as a template for repair machinery

What are the three key components of the CRISPR-Cas9 system? What does each do?

• the Cas9 endonuclease cuts DNA at a specific site within the target site — recognizes a sequence called the protospacer (determined by a guide RNA bound to the enzyme)

• the signal guide RNA is an engineered form of guide RNA that is a fusion of two regions that occur as separate RNAs in nature

  • the guiding region is part of the crRNA in nature and it is complementary to the target DNA sequence (guides where Cas9 cuts)

  • the scaffold region is the tracrRNA in nature and forms a multi-hairpin loop structure that binds tightly to Cas9

• the protospacer adjacent motif (PAM) is a sequence motif immediately downstream of the protospacer sequence that’s required for Cas9 function — Cas9 recognizes the PAM sequence 5’-NGG and when bound, it separates the DNA strands to allow binding of the sgRNA (if it is complementary, Cas9 cuts the DNA)

What are some advantages and challenges of using CRISPR-Cas9?

• advantages: efficient, cost-effective, relatively easy to implement, high precision to target specific sequences

• disadvantages: risk of off-target effects, delivery challenges for certain cell and tissue types, length limitations for DNA insertions, and ethical concerns

What are the four stages of a typical bacterial growth curve? What happens in each?

• lag phase: when an organism is introduced to fresh medium, it takes a little while to adjust, so we see cellular metabolism accelerating and cells increasing in size, but bacteria can’t replicate so cell mass doesn’t change — length of the lag phase directly depends on previous growth conditions

  • rich medium → poor medium: organism takes longer time to adjust due to need for synthesizing proteins, co-enzymes, etc.

  • poor medium → rich medium: easy to adapt so cells can divide with little to no delay

• log (exponential) phase: bacteria divide exponentially (rapid growth and division) so the growth curve is linear and cells are metabolically most active

• stationary phase: nutrients get exhausted, waste builds up, and growth rate decreases almost to equal the death rate, so viable cell number remains the same (plateau)

  • note: if a cell is moved from this phase to fresh medium, it will move to exponential phase again

• death (decline) phase: viable cell population decreases exponentially as the amount of waste products is increased — death of cells exceeds formation of new cells and this continues until the population is diminished to resistant cells (or whole population dies)

What are the two techniques that can quantify the amount of bacteria in a culture? How do they work?

• to obtain colony forming units (CFUs), we need to do a one to ten series of nine dilutions (last tube has dilution of 1:1 billion)

  • 100 microliters of each dilution is plated, incubated, and counted

  • the dilution plate that grows between 30-300 colonies is used to calculate the CFUs per milliliter for that time point

• we can also measure bacterial concentration with optical density

  • a spectrophotometer measures the turbidity or optical density (amount of light absorbed by the bacterial suspension) and the degree of turbidity is directly related to the number of microorganisms present (dead or alive)

  • note: less precise than CFUs but can be obtained instantaneously and requires less reagents

How do we determine growth rate constant, generation time, and growth rate from a bacterial growth curve?

• growth rate constant (μ) = slope x 2.303

• generation/doubling time (g) = 0.693 / μ

• growth rate = 1 / generation time

What do most protein assays tell us? Generally, how do they work?

• various protein assays are quantitative assays that allow us to determine protein concentration

• typical protein assays determine protein concentration by comparing an assay response of the unknown concentration sample to that of a standard with a known concentration

  • protein samples and standards are mixed with an assay reagent and their absorbances are measured with a spectrophotometer — the reading gives us a standard curve that can tell us protein quantity

What are some of the spectrophotometric methods for quantifying proteins? What are advantages/disadvantages?

• Biuret assays are quicker than others, but have low sensitivity and destroy the sample

• Lowry assays have high sensitivity, but take longer and destroy the sample

• Bradford assays are quick and sensitive, but destroy the sample

• BCA assays are sensitive, but take long and destroy the sample

• Warburg christian assays don’t destroy the sample, but they’re only moderately sensitive

What is specificity and sensitivity when it comes to assays?

• the specificity of an assay is a measure of how good it is at discriminating between the requested analyte and interfering substances

• the sensitivity of an assay is a measure of how little of the analyte the method can detect

What factors should you consider when choosing a protein assay method?

• sensitivity of the assay

• the presence of interfering substances in the sample

• time available for the assay

How do BCA assays work?

• bicinchonic acid assays (Smith assays) are copper-based colorimetric assays for protein quantification

• it relies on the formation of a Cu2+ -protein complex in a basic environment, followed by the reduction of Cu2+ to Cu+

• a purple colour is formed when two BCA molecules chelate with one cuprous ion (bound to the protein of interest)

  • this complex exhibits strong absorbance at 562nm and is linear with increasing protein concentration

What are the two fundamental principles of standard curve assays?

• identically assayed samples are directly comparable — variance in protein quantity is the only possible cause for absorbance differences if samples are processed the same (same buffer and stock solution of assay reagent)

  • note: different proteins generate different absorbance values even at the same concentration (issue of this approach)

• units in equals units out — the unit of measure used to express the standard is the same unit associated with the calculated value of the unknown sample

Why is the amount of protein per well irrelevant in protein assays?

• the amount of protein per well isn’t what we want to know — we' want the concentration of the original sample

  • we can use the relationship between the absorbance values to determine the concentration of the whole sample (will be the same as the relationship between the values in the wells anyways)

• note: same goes for the protein concentration in the assay reagent (gets diluted in reagent, but we want to know about the source)

How are samples prepared for SDS-PAGE? What needs to happen prior to running your proteins on a gel?

• prior to running the gel, samples need to be boiled in a sample buffer that has many essential components

  • SDS is an anionic detergent where the hydrophobic chain blankets proteins relative to their mass and the hydrophilic group gives every protein a net negative charge (separation now occurs based on size alone)

  • EDTA is a preservative that chelates divalent cations, reducing the activity of proteolytic enzymes that require ion cofactors

  • Tris acts as a buffer to give a specific pH

  • glycerol makes the sample denser so that the sample settles in the well and doesn’t float out

  • DTT is a reducing agent that helps denature the sample

  • bromophenol blue is often added as a tracking dye

• SDS, DTT, and heat are all responsible for the denaturation of the sample

What is the gel in SDS-PAGE made of? What are the layers?

• polyacrylamide is a polymer that forms a matrix that proteins can move through during gel electrophoresis

• the gels have two layers

  • the separating/resolving gel is lower and responsible for separating proteins by size

  • the stacking gel is above and includes the sample wells

• note: the gel concentration (%T) dictates the size range of proteins that a gel can separate

What are the steps in SDS-PAGE?

1. the proteins need to be isolated from other cellular components, then concentration is measured to allow equal loading of samples (via assays)

2. loading buffer is added to denature the proteins, make the solution denser, and add tracking dye

3. the solution is boiled to help break down disulfide bonds

4. the samples are spin while the gel system is assembled

5. the inner and outer chambers are filled with buffer with the same ion concentration used for the gels

6. molecular weight ladders and samples are loaded into wells

7. positive and negative terminals are connected to a power source to get a voltage in the gel

8. once electrophoresis is complete, the cassette is removed and the gel is stained with Coomassie stain (typical protein stain)

• note: can transfer proteins to a membrane and use antibodies to locate specific proteins

How does two dimensional gel electrophoresis work?

• in 2D gel electrophoresis, samples are separated by isoelectric point and size

• first, samples are loaded onto a gel with a pH gradient and they separate based on their isoelectric points

• the proteins are denatured and moved to a polyacrylamide gel for SDS-PAGE separation

How does Coomassie staining work?

• a common stain is 0.1% Coomassie Blue dye in 50% methanol, 10% glacial acetic acid

  • the acidified methanol precipitates the proteins and the dye sticks to them

• the dye penetrates the whole gel, so excess needs to be removed with acetic acid/methanol so that only proteins are stained

• note: Coomassie blue may not stain some proteins, especially those with high carbohydrate contents