Aging is characterized by the time-dependent deterioration of multiple cellular processes that ultimately increases the probability of organismal death. Within a population, this process is variable and a number of genetic and environmental factors can affect it. One of the best studied and understood variables is the role of diet on lifespan.
Dietary restriction is an intervention that ultimately delays the onset of aging and aging-associated changes. This increases an organism's lifespan and health span, or the length of time when an organism has a normal, healthy life. Our current understanding has led to the identification of a distinct set of cellular changes that have been termed the hallmarks of aging, such as the accumulation of mutations, dysregulation of telomeres, a loss of protein homeostasis, respiration problems, and reactive oxygen species, among others.
Now much of this understanding comes from studies that use many different model organisms. There've been many genetic screens that have identified genes that both shorten and extend lifespan. And these include studies in the budding yeast Saccharomyces cerevisiae, which is the focus of this methodology, as well as studies in more complex multicellular organisms, like C.elegans and the murine system.
Of these though, the budding yeast is particularly amenable to aging studies due to its short natural lifespan and a number of genetic approaches that can unravel the complex genetic relationships, making it an excellent system for functional dissection of the genetics of aging. Yeast can be utilized to study several different aspects of aging. And we will be focusing on chronological lifespan, which is the duration that a cell can survive.
The objective of this work is to present a straightforward genetic approach to perform an over-expression suppressor screen. We will utilize a genetic background that results in an abnormally shortened chronological lifespan. And we'll utilize a targeted approach to identify genes that can rescue or suppress the shortened lifespan phenotype, attempting to identify genes that restore a normal lifespan.
The first genetic background is an autophagy-deficient strain of yeast. Autophagy, which loosely translates as self-eating, is an intracellular degradation system to deliver cytosolic products, such as proteins and organelles, to the lysosome. This helps maintain cellular homeostasis by eliminating damaged proteins, as well as those that have outlived their life.
We will be working with a mutant strain that has a deletion of the ATG1 gene. ATG1 codes for a protein serine/threonine-protein kinase that is required for vesicle formation and autophagy in the cytoplasm-to-vacuole targeting pathway. The ATG1 null mutant has a phenotype of having an abnormally short chronological lifespan.
In this aging lab, we will design and test for genetic factors that can rescue, or suppress, the abnormally shortened lifespan characteristic of the ATG1 null mutant. However, this can be expanded to other short-lived genetic backgrounds as well. The first step in this process is to identify a gene that is linked to extending lifespan.
We're going to focus specifically on the genes that extend the chronological lifespan when they are over-expressed. To do this, we are going to use the publicly-available data that is accessible from the yeast genome database. We're going to go up here.
Search by phenotype. And about midway down, we will see we can select the chronological lifespan. To look for genes that increase the lifespan, what you can see is the wealth of data available that has genes linked to this particular phenotype.
Today, we will be studying SIR2, a histone lysine deacetylase that is linked to extended chronological lifespan when it is over-expressed. The first thing we will do is access the DNA sequence for the coding region, as well as the regulatory regions that flank either side of this gene. We will take 400 base pairs upstream and downstream to be sure that we encompass all the regulatory regions.
We will use this DNA sequence to design PCR primers that will allow us to clone this gene into our plasmid vector. We will use PCR to amplify our gene and to clone it into the pRS315 vector. In order to do this, we will design primers that contain restriction digestion sites that incorporate HindIII and SacII restriction sites.
As you can see, the plasmid contains both restriction sites, HindIII and SacII, that will facilitate our cloning. You should design your PCR primers to contain about 21 or 22 nucleotides of sequence complimentary to the region that you want to amplify. In addition to the sequence, you will add onto the five prime end of them the appropriate restriction site, either HindIII on the forward primer, or SacII on the reverse primer, shown here in red and purple respectively, and further upstream of that, add several nucleotides that will allow the restriction enzyme to bind and create the restriction digestion site at a higher efficiency.
Grow a five milliliter culture of the wild type yeast overnight to post-log phase in enriched media, such as YPAD. Harvest genomic DNA using a commercially available kit that is specific for fungal genomic isolation. It is important that the kit is specific for yeast, as they have a very rigid and tough cell wall to penetrate.
Zymolyase treatment is effective at digesting the cell wall and greatly increases the genomic yield. Use a spectrophotometer to determine the quantity and the quality of the DNA. To produce an amplicon that is suitable for cloning, it is necessary to utilize a high fidelity PCR polymerase to avoid mutations during the amplification process.
There are many different high fidelity PCR kits that are commercially available. To facilitate the optimization of the reaction, we recommend choosing a kit that provides multiple buffering systems, one that is a standard high fidelity buffer, and one optimized for high GC content in complex amplicons. We are going to add 200 nanograms of our genomic DNA.
We're going to add five microliters of our buffer. We're going to add 2 1/2 microliters of our forward and our reverse primers. We're going to add one microliter of our polymerase, two microliters of D NTPs.
And lastly, we're going to QS the whole reaction to 50 microliters with sterile distilled water. We'll run the reaction according to the protocol specific for the enzyme we're using, as well as for the primers that we have designed in this experiment. Cleanup and concentrate the PCR reaction.
Grow a five milliliter culture of E.coli overnight in selective media. Pellet the culture by centrifugation and discard the supernatant. Resuspend and lyse the cells in the provided chaotropic buffer for five minutes.
Neutralize the reaction and mix the two by inversion five or six times. Centrifuge your sample for 10 minutes at maximum speed. Transfer the supernatant and to a silica column and wash with the provided buffer.
Discard the flow through and repeat the wash with the other appropriate buffers, with 30 second centrifugations in between, discarding your flow through after each. At the end, centrifuge for two minutes more to remove any residual ethanol and elute your plasmid into 20 microliters elution buffer and determine the quantity and quality by spectrophotometer. Now it is time to set up a restriction digestion of the vector and the insert.
Add 625 nanograms of DNA, either the vector or the insert, five microliters of CutSmart buffer, one microliter of SacII, one microliter of HindIII, and QS the reaction with water to bring the final reaction volume to 50 microliters. Incubate the restriction digests at 37 centigrade for three hours, followed by 80 centigrade for 20 minutes to heat and activate the enzymes. At this point, digests can be stored at four degrees prior to proceeding to the next step.
Next, we're going to set up a 15 microliter ligation using our digested genomic fragment, as well as the vector that we just completed. We're going to start by adding six microliters of water. We are going to add two microliters of our digested vector.
This is our pRS315 plasmid. We're going to add four microliters of our insert, the SIR2 gene. We're going to add two microliters of our ligase buffer, case the 10X buffer.
And lastly, we're going to add one microliter of the ligase. We're going to incubate this at 16 degrees overnight, and then heat kill the enzyme at 80 degrees centigrade for 20 minutes. To screen for our successfully created plasmid, we're going to transform it into E.coli.
We're going to use 50 microliters of chemically competent cells. We're going to thaw them on ice, and as soon as they thaw, we are going to add our ligation reaction, we're going to add the entire contents of it. So we're going to add 15 microliters of the ligation with the insert, and we're going to add 15 microliters of our no-insert control as well to a separate reaction.
Once it's been added, flick the tube a few times in order to mix it. And we're going to incubate this on ice. After completing the 30 minute incubation on ice, we're going to add our samples into a 42 degree heating block where we will incubate and heat shock for 20 seconds, which point we will take them out, and we will add recovery media.
After the heat shock, we're going to allow for a recovery period. We are going to take our samples and we will add 450 microliters of SOC media. We will incubate these at 37 degrees centigrade to allow the plasmids to be picked up and to allow the ampicillin resistance gene to begin to be expressed.
We will allow these samples to recover at 37 degrees for 45 minutes. After that incubation, we set up a series of dilutions, one to 10, and one to 100, and we're going to plate these on LB/Amp media and allow the cells to grow up overnight. We will plate them using sterile technique.
We'll take the appropriate plate, which we have labeled accordingly and contains our selectable marker. We will take 150 microliters of our transformed E.coli. Using a sterile inoculating loop, we will then gently spread the cells throughout and across the entirety of the plate so we get a nice even distribution.
Once we have finished plating all our dilutions, we will incubate these plates overnight at 37 degrees centigrade to see what grows. After about a day, we should see colonies growing up that hopefully contain the plasmid we were attempting to create. They should look something like this.
Using sterile technique, inoculate potential transformants that grew from your transformation into five milliliters LB plus ampicillin following the previously outlined procedure. Isolate the plasmids from every potential transformant and run a restriction digestion as previously described. Run the reaction products on an agarose gel, comparing the potential transformants to the empty vector.
Save any transformants that result in excision of a properly-sized insert for further study. Having successfully created our plasmid, we are going to transform it into the ATG1 null yeast background. We're going to take 15 mils of cells and we're going to pellet them, which I have already done right here.
After they've pelleted, remove the YPAD media that they were growing in and wash the cells with distilled water. After that, we're going to take the cells and we're going to resuspend them in 100 millimolar lithium acetate. We're going to resuspend them in 200 microliters of that.
Resuspend the cell pellet by gentle pipetting up and down. Split the cells into separate microfuge tubes for each of the transformations that you will perform, using 50 microliters of this mix per transformation. To each of the transformations, you will need to add 240 microliters of 50%PEG.
PEG is very viscous, pipette and measure very carefully. Mix by pipetting after the addition of the PEG and after every one of the additional components. Next add 36 microliters of one molar lithium acetate, five microliters of salmon sperm DNA, or other carrier DNA, that has been boiled for five minutes and placed on ice, and finally five microliters of the plasmid for each transformation you will be performing.
Vortex each tube thoroughly to mix. Incubate your samples at 30 degrees centigrade for 45 minutes. Heat shock them at 42 centigrade for 10 minutes, and then wash the cells once in sterile water, finally resuspending them in 200 microliters of sterile water.
Set up one to 10 and one to 100 dilutions of each of the transformed strains of yeast and plate 150 microliters onto SC minus leu plates to select for the plasmids. Spread the cells uniformly and evenly, and allow the plates to dry before inverting and incubating them at 30 degrees centigrade, allowing them to grow for 48 to 72 hours. Once we have successfully cloned our gene to test, we'll test the effect on the chronological lifespan of the organism.
We will grow the yeast, monitoring the number of viable colony forming units that remain as a function of time. Grow the yeast first for 72 hours in SC minus leucine liquid media with shaking at 30 degrees centigrade. Using a hemocytometer, determine the dilution necessary to allow you to plate 500 cells.
Using sterile technique, plate the cells onto an SC minus leu plate, allowing it to dry and incubate inverted for three days at 30 degrees centigrade, at which point they will grow up and you can score the colony growth and record that time point. After making the plate, continue to incubate your yeast at 30 degrees centigrade without shaking. Initially, we repeat the plating at weekly intervals, taking more frequent time points if our preliminary data suggests a suppression of the aging phenotype.
In order to determine the percent viability, we normalize to the day one time point for analysis. Be sure to plate the same dilution at every interval. Continue with this process until you no longer see any colonies growing up.
It's helpful to enter your information into an Excel spreadsheet. This will allow you to quickly determine the viability of each strain at the conclusion of the experiment.
