LAB TEST Flashcards
(99 cards)
1
Q
What are the 3 things we looked at throughout the labs?
A
- Morphology - what does the entire cell look like?
- Protein expression - what happens to the expression of a particular protein?
- Protein localization - what happens to the location of a particular protein?
Lab 1 guide
2
Q
Primary cells
A
- taken directly from a tissue
- unless modified, have a finite life span => may divide a few times but eventually die
- closely resemble in vivo physiology (advantage)
- trickier to grow, ultimately die (disadvantage)
Lab 1 guide
3
Q
Immortal (‘continuous’) cell lines
A
- divide indefinitely
- originated as cancer cells or were transformed in lab to give them this ability
- easier to grow and divide over and over (advantage)
- less representative of in vivo systems and, with time, require increased levels of genetic modifications
Lab 1 guide
4
Q
Adherent vs suspension cell lines
A
adherent
- attach to a substrate
- grow and divide to produce a solid monolayer, with minimal space between cells (100% confluency)
- some cells stop growing at this point, others (like cancer cells) continue growing on top of each other (depends on cell line)
substrate
- do not attach to a substrate
Lab 1 guide
5
Q
Common immortal cell lines used in labs
A
- HeLa => first human immortal cell line, derived from cervical cancer tumor
- HEK 293 => human embryonic kidney epithelial cells
- SF9 => insect cells that can be grown in suspension
- MCF-7, Saos-2, PC3 => cancer cells
Lab 1 guide
6
Q
Risk Group (RG) classifications
A
All microorganisms, proteins, and nucleic acids are assessed to determine their risk to the individual/ animal and public health
RG 1: very low risk (e.g., saccharomyces cerevisiae, Ptk2 cells
RG 2: pathogens are capable of causing serious disease in human but are unlikely to do so (e.g., listeria, HEK293, Norwalk virus, Sars-CoV-2 RNA)
RG 3: pathogens are likely to cause serious disease in humans or animals (e.g., mycobacterium tuberculosis, SARS-CoV-2 whole live virus)
RG 4: pathogens can produce highly contagious, serious or fatal disease for which there are no treatments or vaccines (e.g., ebola virus)
Lab 1 guide
7
Q
Containment Facilities
A
Containment levels refer to the minimum physical containment and operational practices required for safe handling of infectious materials and toxins
CL1: ‘regular’ type of teaching lab, no special design features other than a functional working space and cleanable work surfaces. Open bench work is acceptable and Biological Safety Cabinets are not required (Us!)
CL2: common type of facility in hospitals and universities for either diagnostic, health-care work or for research purposes. All RG2 pathogens are contained here. Rooms have BSCs equipped with HEPA filters
CL3: require additional primary and secondary barriers to minimize the release of infectious organisms into the environment, including sealed windows, use of BSCs for all work, and strictly controlled access (used for COVID research)
CL4: provide max level of biosafety and biosecurity. Max containment is ensured via a complete seal of the facility perimeter (includes sealing any conduits crossing the containment barrier, like electrical conduits and plumbing). Lab workers must wear full coverage, positive-pressure suits with their own breathing supply. Must go through a chemical shower before removing protective clothing
Lab 1 guide
8
Q
Personal Protective Equipment (PPE) in the lab
A
- Lab coats - also needed to not track contaminates out of lab
- Proper footwear
- Gloves
Lab 1 guide
9
Q
Ptk2 cell line
A
- epithelial kidney cells from a male potoroo (marsupial)
- isolated cells in 1962 were placed in culture dishes where they continued to replicate
Lab 1 guide
10
Q
Potoroo (why use their cells??)
A
- they have a small number of chromosomes
- Ptk2 cells stay relatively flat in cell culture, making it easy to see their large chromosomes
- these characteristics made this cell line ideal for studying genetics and the cell cycle (+ they grow nicely in culture and are a great cell line to use in BIOL 2020)
- RG1 cells => can be used in our lab
Lab 1 guide
11
Q
What cell culture equipment was mentioned in Lab 1? (No specific details)
A
- Biological Safety Cabinet
- CO2 incubator
- Inverted microscopes
- Media
- Culture vessels
12
Q
What is a biological safety cabinet?
A
NOT A FUME HOOD
- needed for a CL2 lab
- protects us from infectious materials or toxins AND protects specimens from contamination
13
Q
How does a biological safety cabinet work?
A
- creates an air curtain across the front opening to prevent aerosols from escaping out the front
- prevents unfiltered air from entering the the working area
- inside: air moves in a constant, streamlined speed and direction => creates a laminar flow that contains airborne infectious agents
- air from the cabinet is exhausted through a HEPA filter
- also maintains a sterile work environment by filtering incoming air through the HEPA filter before it blows across the working surface
Lab 1 guide
14
Q
What is a CO2 incubator used for?
A
- required for short-term storage of growing cells
- provides a clean, humidified environment with a constant temp
- supplied with 5% CO2, which maintains the pH at physiological level
Lab 1 guide
15
Q
How are cells stored long-term?
A
cells are cryopreserved in liquid nitrogen
- cryoprotectants, such as DMSO, are added to the media used for freezing cells to reduce intracellular ice formation and prevent cell death during freezing
Lab 1 guide
16
Q
Inverted microscopes
A
have the lens on the bottom of the microscope and light source above the specimen
- needed as cells are usually growing on the bottom of the flask and there is often condensation on the top
- also, flasks and dishes are bulky and the large stage provides an adequate space to view the vessel
Lab 1 guide
17
Q
Media (for cell growth)
A
used to provide nutrients the cell requires
- typically contains about 5-10% fetal bovine serum (FBS) => provides nutrients, hormones and trace elements necessary for cell growth and proliferation
- antibiotics often added to minimize risk of bacterial growth following contamination
- phenol red is added to monitor pH
18
Q
What does a change in the colour of the pH indicator mean for cell cutures?
A
its a sign that there is excess of metabolic by-products and that it si time to either split the cells (also referred to as ‘passaging’ or ‘subculturing’) if they are confluent, or change the media to replenish depleted nutrients
Lab 1 guide
19
Q
What are mammalian cells grown in?
A
specialized culture vessels that have been treated to allow adherent cells to attach to the bottom surface
- e.g., flasks with screw tops that can be vented or non-vented, plates with individual wells
20
Q
Many experiments in cell biology test the effect of a _______ _______ on some sort of measurable ________
A
single variable; phenotype
21
Q
What kind of pipette should be used for volumes greater than 1 mL?
A
a pipet boy
22
Q
What are the two types of stressors we used?
A
- Oxidative stress; from hydrogen peroxide
- pH stress (i.e., acetic acid shock); from vinegar
23
Q
What was the trick with stressing the cells? (i.e., what was the point?)
A
to induce apoptosis, but not to over stress the cells to cause necrosis
24
Q
Apoptosis vs necrosis
A
apoptosis: organized cell death
necrosis: less orderly (splat!)
delicate balance to prevent necrosis
25
What needed to be done to the negative control group in lab 1?
the same volume of water, to match the volume of stressor, had to be added to ensure everything besides the stress was the same
26
Total magnification is the product of what?
the ocular and objective lenses
- ocular is 10X
- objective is 10X or 40X
27
Why was phase contrast microscopy used in lab 1?
small, unstained specimens (like living cells) can be hard to see using brightfield
- phase contrast converts differences in light phase shifts into differences in light intensity
- made possible by aligning a phase plate that sits at the back of each objective with a special filter in the turret
28
Trypan Blue
a common stain used to determine cell viability
- dead or dying cells with compromised cell membranes allow the blue dye to leak into the intracellular space
- often called an "exclusion test" => living cells with intact cell membranes will exclude the dye and remain a clear/grayish colour
29
What was the procedure of determining cell viability in lab 1?
- retrieve the 6-well plate after 30 minute incubation in stressor (oxidative stress or acetic shock)
- retrieve two glass microscope slides and label one "T" and one "C"
- put 25 microliters of trypan blue on each slide
- pick up coverslip from well and place it cell side down on drop of trypan blue
- remove any surrounding liquid and put slide on microscope stage
- observe cells at 10X in both brightfield and phase contrast
- use the best setting for you and increase to 40X (if phase contrast, change to Ph2!!)
30
What were we looking for when we were determining cell viability?
- membrane blebbing: once apoptosis signaling pathways have been initiated, the cell will start to fragment and small vesicles will bud off from the membrane (one of the most defining characteristics of apoptotic cells
- cell shrinkage: as cell fragments, it will become smaller in size (necrotic cells will swell and appear larger than normal)
31
Hemocytometer and determining cell concentration
hemocytometer: standard way to count cells; has a specialized slide with an etched grid; pipet cells into chamber on top of the grid
- count the cells in the 4 corners (unless cell concentration is super high or low)
- cells per 0.1 microliters = (numer of cells you counted)/ (number of large squares containing counted cells)
- multiply this by the dilution factor ( sample volume plus diluent volume all over the sample volume) to get it back to the actual concentration (without trypan blue)
- finally, multiply by 10^4 to get it in per mL
32
How are proteins extracted from cell? What are the 2 methods?
proteins are extracted by disrupting the membranes and collecting the resulting liquid supernatant (i.e., cell lysate)
- physical methods of lysis
- chemical methods of lysis
* preparation must stay cold so the proteins do not denature
33
Physical methods of protein extraction (how they work and what they do)
use external forces to physically break down the membrane to release the cellular components
- liquid homogenization: apparatus shears cells by forcing them through a narrow space. Can be done with a plunger and vessel or in an automated fashion
- sonication: machine uses pulsed, high frequency sound waves to agitate and lyse cells. Sound waves are delivered using an apparatus with a vibrating probe that is immersed into the cell suspension
- manual grinding (mortar + pestle): technique works well to isolate proteins from plant cells. Freeze tissue in liquid nitrogen and then smash to release proteins
34
What is the challenge with physical methods of protein extraction?
- it's often difficult to have consistent preparations from one day to the next
- less reproducible
- these methods can generate heat, which will cause proteins to denature
35
Chemical methods of protein extraction
use detergents and/or hypotonic solutions (low salt) to disrupt the membrane and extract proteins
- RIPA is a commonly used lysis buffer => contains detergent Triton X-100
- Ripa buffer typically includes protease and phosphatase inhibitors
36
What are the key steps in extracting proteins from adhering cells using RIPA buffer?
1. grow cells in flask
2. remove media and wash cells with buffer (like PBS) to remove residual media
3. add RIPA buffer to flask => ensure all cells come into contact with buffer
4. incubate cells in RIPA buffer on ice for 10+ minutes
5. transfer lysed cells to a tube on ice and centrifuge to pellet any membranes
- supernatant will contain a soup of the proteins (= cell lysate)
37
Marion Bradford
developed a quick and easy way to find the total concentration of protein (expressed as microgram per microlitre) in a solution by using a colorimetric reaction
- technique remains one of the most common ways of testing protein concentration
38
How does the Bradford assay work?
when a dye in the bradford reagent binds to protein, there is a colour change from brown to blue
- intensity of the blue colour is measured using a spectrophotometer set at 595 nm
- assay uses BSA (bovine serum albumin) as a standard to which the unknown samples will be compared
39
BSA
bovine serum albumin
- used in the Bradford assay technique as a standard to which the unknown samples will be compared
- BSA isolated from cow blood and is often used in research when a generic protein is needed
40
How did we perform a Bradford Assay in Lab 2?
1. first created a dilution series from stock BSA
- colour change is linear only in the range of 0.20 - 1.0 micrograms/microlitres => BSA must be diluted with PIPES buffer to make a series of standards in this range
2. set up cuvettes for cell lysates (2 for negative control, 2 for test sample) => also dilute with buffer
3. add 5mL of bradford reagent to each cuvette, mix and incubate for 10 min
4. measure absorbance with spectrophotometer
41
Task 1 of lab 2
create a serial dilution of BSA stock solution
1. use C1V1 = C2V2 to determine how to serially dilute the BSA stock solution
2. label cuvettes with tape A - J
3. pipette PIPES buffer into cuvettes based on calculated values
4. pipette BSA stock solution into cuvettes A-F (use calculated values)
5. pipette cell lysates into remaining 4 cuvettes based on calculated values
6. add 5mL of bradford assay to each cuvette, stretch parafilm over the cuvettes and mix by inverting several times
7. let reaction take place for 10 minutes
42
Spectrophotometer
use light of a specific wavelength that is generated from suitable light sources
- absorbance of light by the sample is measured by a photocell that converts the received energy into electrical energy so that a reading is displayed on the unit
- degree to which light is absorbed by a solution (i.e., absorbance) is related to the concentration of the absorbing solution
43
Why do we prepare a cuvette to "blank" the spectrophotometer?
to account for any absorbance due to the solvent in which the sample is dissolved (i.e., brown background colour of bradford reagent)
- the cuvette adjusts the machine to read zero absorbance
- it contains only PIPES buffer and bradford reagent
- ensures that any positive absorbance readings by the spectrophotometer in the remaining cuvettes is the result of the blue colour produced by the bradford reagent reacting with the proteins
44
Why did we make a graph in lab 2? Did the graph include the cell lysate cuvettes?
- using a dilution series of BSA, a linear graph of BSA concentration (x-axis) vs BSA absorbance (y-axis) is made
- when using a solution of unknown concentration, we can use the resulting equation of the line (from the graph) to determine the concentration of the unknown solution
- then multiply by the dilution factor to get the actual concentration
45
Electrophoresis
- a common method for separating charged molecules in an electric field
- molecules are driven through a cross-linked matrix (= gel), which acts like a molecular sieve, allowing differential migration of molecules based on size
46
What type of gel is used for protein electrophoresis? How are they separated?
gels made of polyacrylamide (PAGE)
- separated based on molecular weight
47
What type of gel is DNA electrophoresed through? How is DNA separated?
- agarose gels
- separated according to number of base pairs
48
How are proteins prepared prior to electrophoresis?
- cell lysates are mixed with sample buffer (most contain SDS)
- SDS disrupts protein folding, which causes the proteins to denature and become rod-shaped - meaning that movement through gel does not depend on protein shape
- SDS also coats polypeptides with negative charges to increase speed of movement towards positive electrode
- sample buffer can also contain a reducing agent to break strong disulfide bonds with the protein
- samples are boiled briefly to ensure all proteins have fully denatured
49
Sample buffer (SDS-PAGE)
- exact composition varies, most contain the negatively charged detergent sodium dodecyl sulfate (SDS)
- causes protein denaturation
- coats proteins in a negative charge
- sometimes contains a reducing agent, beta-mercaptoethanol or DTT, to break the strong disulfide bonds within the protein
- normally prepared as a 2x stock solution and is mixed in equal volumes with the protein sample
50
SDS-PAGE vs Native-PAGE
- SDS ensures the proteins are denatured and all the same shape
- Native PAGE omits denaturing chemical in the sample buffer and the gel to conserve the interactions between polypeptide subunits
- Native PAGE provides more info about protein structure and folding
51
Molecular weight standards
- proteins of known weights ("standards", "ladders" or "markers") are run alongside the protein samples in the same gel
- migration distances can be plotted as a function of log molecular weight to produce a straight line (= standard curve)
- can then compare distances migrated by sample proteins to the graph to accurately estimate their MWs
- ours was pre-stained so we could follow the protein migration in real time as gel is running
52
Name the 5 tasks we did in lab 3
1. prepare the protein samples (partly done over the summer)
2. position the acrylamide gel in the electrophoresis tank
3. load marker and cell lysates into the gel & run the gel
4. remove the gel from between the plastic plates
5. transfer proteins from gel to nitrocellulose
53
What happened over the summer to prepare for lab 3? Be specific
- Ptk2 cells were grown and the experiment was set up by stressing the cells (vinegar or H2O2) or preparing unstressed cells as negative control
- cells were incubated in appropriate buffer for 2hr and were then lysed in RIPA buffer
- cell lysates were collected and the concentrations were determined
- cell lysates were then mixed with 2X Sample buffer (containing SDS, mercatoethanol, glycerol, and tracking dye) aliquoted into microcentrifuge tubes and frozen
54
In lab 3, why did we need to further dilute our samples in 1X sample buffer?
to ensure that the stressed and unstressed cells were at the same concentration of 1 microgram/ microlitre
- this ensured that when we ran the gel, we were running the same amount of protein and could make a fair comparison => also ensured that any observed differences in protein expression are due to experimental conditions and not because one sample had more protein loaded than another
55
What was the first task we did IN LAB for lab 3?
- diluted our cell lysates using 1x sample buffer into new microcentrifuge tubes
- boiled the tubes of newly diluted protein (not the marker) to ensure proteins were denatured
- centrifuged the samples (and the marker tube) to remove bubbles and bring the solution back to the bottom
56
How do you position the acrylamide gel in the electrophoresis tank?
1. place a black clamp (labelled 'L') into the left chamber of the tank
2. pour running buffer into the chamber with your clamp, so that it is level with the silver electrode
3. at a sink, use scissors to cut open the packaging and remove the gel. dispose of packaging into garbage. remove plastic comb and throw out. wipe away any stringy acrylamide bits with a kimwipe
4. hold gel upright and inspect well tails to ensure they are standing upright. fix if they are not. peel off the white strip at the bottom and throw out
5. drop the gel into the chamber in front of the black clamp. position gel so that the lower slanted plastic plate faces you and the front of the tank (running buffer will get into the wells)
6. pull the lever on the clamp forward to lock the gel into place. make sure the running buffer is slightly above the silver electrode
57
Why did we load the samples in duplicate for the electrophoresis?
to demonstrate authentic experimental approach so we can eventually probe for 2 dif proteins in lab 4
58
Why did we load 1x buffer into the empty lanes?
to ensure the samples ran evenly
59
Why could we cut off part of the gel after it had ran?
because the parts that were cut off did not contain proteins
60
How did we prepare for the Western Blot in lab 4?
by transferring the proteins on the gel to a piece of nitrocellulose membrane
- stack gel on nitrocellulose and use a Bio-Rad Trans-Blot Turbo Machine to apply a current
- current moves negatively charged proteins out of the gel onto nitrocellulose membrane
61
Why are bubbles bad when transferring the gel to nitrocellulose?
they prevent efficient transfer of proteins
62
What was the 'sandwhich' we had to make to transfer the gel to nitrocellulose?
1. thick filter paper
2. nitrocellulose
3. gel => use roller to remove bubbles
2. thick filter paper
63
What solution did we store the nitrocellulose membrane in between labs 3 & 4?
TBS-Tween
64
Coomassie Blue
a stain that could be used to stain the gel immediately after electrophoresis
- allows us to see ALL proteins in cell lysate
- not super useful: impossible to know what ban belongs to what protein of interest (looks very blurry too)
65
What are anitbodies?
- proteins produced by an animal's immune system
- powerful tools in cell biology research b/c they have high degree of specificity against target protein (antigen)
- antibodies against a given protein can be grown and collected from animal hosts (i.e., mouse, rabbit, goat)
- typically belong to IgG class
66
Antibody structure discussed in lab
Y-shaped
- base of Y = constant domain (identical for all IgG antibodies from 1 species)
- branches = variable regions => complement shape of antigen to allow binding
67
What are the 2 approaches to generating antibodies for use in research?
1. Polyclonal antibodies: made by injecting animal host with purified antigen (or protein of interest), immune sys. produces many antibodies in response => harvested from animal's blood => end up with mixture of antibodies each specific for dif epitopes on POI
2. Monoclonal antibodies: specific for same epitope on antigen, produced by injecting animal with antigen triggering immune response => remove some antibody-producing WBCs & fuse to immortal cells => produce hybridomas that continuously divide in culture => 1 hybridoma can be isolated & cultured to produce ++ copies of exact same antibody
68
Advantages of monoclonal antibodies
highly specific and less likely to bind to other proteins
69
Reporter molecules
must be able to visualize antibody binding to antigen
- reporter molecules are attached (conjugated) to antibodies
- reporter molecule could fluoresce under a microscope or could be an enzyme that catalyzes a rxn to produce a colour change or burst of light
70
Direct visualization with antibodies
uses a single antibody with a reporter molecule attached
71
Indirect visualization with antibodies
uses two antibodies (primary and secondary), primary recognizes protein, secondary recognizes primary antibody and has reporter molecule attached
72
How do you name antibodies?
primary: animal produced in + anti- protein of interest
secondary: animal produced in + anti- animal primary was produced in
reporter molecule is added to name by saying: labelled with ______ (FITC, Cy3.5...)
73
What were the antibodies we used in lab 4 conjugated to?
horseradish peroxidase (HRP)
- in presence of chemical luminol and oxidizing reagent, enzyme will catalyze reaction that releases light
74
What info did we gain based on the intensity of the light? What about the location? (Lab 4)
intensity: expression level of protein
location: molecular weight of protein
75
Why did we do a procedural check for lab 4? What is it?
uses "house-keeping" proteins => expression is always constant (i.e. stressor is not expected to cause up- or down-regulation of expression of protein)
used to be confident that any noticeable differences in expression are valid, and not the result of an issue along the way (i.e., luminol/oxidizing could be expired, total amount of protein loaded in gel could have been different, transfer from gel to nitrocellulose might've been bad...)
band intensity for control and test sample should be almost identical for a procedural check
76
What antibodies did we use in lab 4, what protein did they target, and why did we use them?
mouse anti-tubulin conjugated to HRP: targets tubulin, procedural check
mouse anti-Hsp27 conjugated to HRP: targets Hsp27, determine if target protein is up or down-regulated in response to experimental conditions
rabbit anti-p53 conjugated to HRP: targets p53, same as Hsp27 target
77
Ponceau Red
stain used in lab 4 to determine if proteins transferred to nitrocellulose successfully
78
Describe the process leading up to the antibody incubation in lab 4
1. stain nitrocellulose with ponceau red to see if proteins transferred well from gel
2. rinse membrane many times to remove ponceau red from membrane
3. examine intensity of each lane => are they all roughly the same?
4. cut membrane into 2 pieces so there's a test sample, negative control, and marker lane on each piece
5. use a pen to label pieces with bench number, protein being probed for and sample loaded into each lane (M, C, T)
6. place membranes in blue dish and pour 5% milk solution to submerge membranes (they should float)
7. set dish on shaker for 25 min => casein (protein in milk soln.) will coat nitrocellulose to reduce non-specific binding of antibody to membrane
8. pour out milk solution into sink
9. pour TBS-Tween onto membrane, swirl, and pour it out
79
Why did we wash the membranes with TBS-Tween and then TBS buffer after the antibody incubation?
we did 3 2min washes with TBS-Tween to remove any unbound antibodies
then washed with TBS buffer (3 min) b/c the TBS detergent in TBS-Tween interferes with the chemiluminescence procedure
80
Chemiluminescence procedure
pipette equal amounts of luminol and oxidizing agent into same microcentrifuge tube
- place membranes side-by-side on plastic wrap and pipette above mixture onto membranes
HRP on antibody catalyzes oxidation of luminol => release of light (not visible to naked eye)
81
How did we visualize the light reaction in lab 4?
with blue autoradiography film; in darkroom, light signal from HRP is captured on film => results in black lines on film
82
What are the 3 things we did when analyzing the western blot?
1. compared the film to nitrocellulose to see location of the bands in relation to positions of pre-stained molecular weight markers on nitrocellulose
2. look at band intensity of procedural checks to determine if we are confident in integrity of experiment
3. look at band intensity of target protein in negative control to get baseline => then look at band in test sample to determine if protein expression was changed (i.e., up or down-regulated)
83
Fluorochromes
- molecules that emit light in visible spectrum
- each has a different excitation and emission wavelengths
- absorb light for short period of time and re-emit light at a slightly shifted wavelength (20-50 nm)
84
How do we visualize light emitted by fluorochromes?
re-emitted light passes through a barrier filter in microscope on its way from stage to eyepiece => filter allows only light of desired wavelength to pass through it
85
FITC and Hoechst
FITC: fluorochrome that emits green light => is conjugated to an antibody
Hoechst: chemical compound used to label DNA => binds directly to DNA (i.e., no antibody needed), emits blue fluorescence when hit with UV light
86
Why did we incubate the coverslips in ice cold methanol?
to fix and permeabilize the cells
fix: cell structures can stay intact
permeabilize: antibodies can enter cell and bind target protein
87
How did we prepare the humidity chamber? Why was it necessary?
- squirt water to dampen filter paper in glass petri dish
- place parafilm on top of the 2 toothpicks
- pipette antibody onto parafilm
- needed to keep the cells from drying up
88
What did we rinse the coverslip in after the methanol incubation?
PBS buffer
89
Antibody incubation (lab 5)
- place coverslip cell side down on antibody drop
- put lid on petri dish (humidity chamber) and cover in aluminum foil
- incubate => put coverslip in a jar with fresh PBS to wash away unbound antibody
- do this wash 3 times
90
What was REALLY important to keep track of during lab 5?
what side of the coverslip the cells were on
91
Why did we cover the petri dish with aluminum foil?
to prevent the fluorochrome from being quenched by the light in the room
92
Why did we have to be cautious with Hoechst solution?
because it binds DNA => is a potential mutagen and carcinogen
93
Did we wash the coverslip in PBS buffer after the Hoechst incubation?
Yes, 3 times
94
Why did we put anti-fade medium on the glass slide before putting the coverslip on it?
95
Alternative fluorescent probes, i.e., GFP
- a method to visualize structures in living cells
- uses gene editing to fuse DNA that encodes for GFP to a gene encoding protein of interest
- recombinant DNA = introduced into cells where it is expressed to produce a protein with a GFP tag
- if protein is exposed to blue light, GFP will glow green => provides valuable info about location and function of proteins in living cells
96
Confocal microscopy
centers light through a pinhole to focus on a single plane => when out-of-focus light is blocked, researcher can obtain a high-quality image that can be optically sectioned to provide further info
97
FRET (Forster/ Fluorescence Resonance Energy Transfer)
uses fluorescence to measure interactions between proteins
- 2 dif proteins are labeled with dif fluorochromes
- key feature: emission wavelength of 1 fluorochrome overlaps with excitation wavelength of other
- if proteins = in close proximity, energy from one is passed to other + emission of second fluorochrome is detected
98
Flow Cytometry
- used to gain info from a large population of cells
- passes cells in a liquid suspension through a narrow channel
- laser scans cells as they flow by => based on light scatter computer generates info about size and shape of cells + detects fluorescence emitted by any antibody-tagged proteins or other fluorochromes being used
- commonly used to analyse apoptosis, detect proteins on cell's surface or intracellularly, or to determine which cells are in a particular stage of cell cycle
99
Fluorscence-Activated Cell Sorting (FACS)
- variation of flow cytometry
- isolate specific cells for future culturing and downstream applications
- cells are separated into droplets as they pass through apparatus => one cell per drop
- instrument monitors each cell's info (size, shape, fluorescence, etc.)
- can physically separate cells into different containers for future use
- e.g., label DNA with fluorochrome, software can use info regarding amount of DNA in nucleus to determine phase of cell cycle
