ewinter's blog

The interspecific interaction I will be documenting is between humans and grass.  Despite the fact that there are an abundance of paved sidewalks on campus, students still feel the need to walk over the grass in very specific patterns.  This leads to the grass being matted down and even uprooted, leaving distinct dirt paths cutting through otherwise grassy fields. A good example of this is a position just north of the student union and just east of the parking garage.  Just looking at a map, one can tell that the humans benefit because they save time. However, the grass does seem to have one mechanism of defense. If it is saturated with water, it will be muddy, and humans will be less inclined to trample it because doing so would damage their shoes.

The average A260/280 ratio of 2.05 for the untreated samples is consistent with RNA, indicating presence of RNA in these samples, as expected.  The average A260/280 ratio for RNase treated samples was 1.60, while the accepted ratio for pure DNA is approximately 1.8.  This is slightly low to be convinced that the RNase treated sample is pure DNA.  The 260/230 ratio average of 0.60 for RNase treated samples further justifies the impurity of the samples, because a pure nucleic acid should have a 260/230 ratio that is higher than the 260/280 ratio [7].  On the gel, the consistent presence of bands slightly above the 10,000 base pair marker indicates the presence of genomic DNA in the samples.  In the RNase untreated samples, the fields of discoloration below the 500 base pair marker indicate the presence of RNA.  The RNase treated samples did not show these fields, indicating that the RNase worked to degrade the RNA to small enough lengths such that it was unnoticeable on the gel.  Although the RNA did not appear on the RNase treated gel, small RNA fragments were still in the solution even after RNase treatment because nothing was done to remove them.  This fact likely helps to explains the NanoDrop results indicating impurities for the RNase treated samples, although it is possible that there were additional cellular components left over as well.  The calculated amount of nucleic acids extracted was 15,720 ng.  The gel indicated the presence of both DNA and RNA in the untreated samples of the extract, and NanoDrop measurements indicated impurities that include RNA and possibly other cellular components.  Therefore, the actual amount of genomic DNA extracted from B. distachyon is likely far less than this value. 

In order to treat both pathways, we are planning to use elements of CRISPR gene editing techniques.  In order to target the HGSOC cells, two hallmarks of the cancer have been identified: CA-125 and MAGE-A10.  One idea we have is to insert mRNA for the BH3 protein into cancerous cells. The cell could then translate this pro-apoptotic protein.  For the cell proliferation pathway, c-Myc is likely amplified in part due to the extremely common loss of function mutations in p53 (tumor suppressor).  If we could inject mRNA coding for p21 into the cells, then the cell would transcribe p21, the CDK inhibitor that promotes apoptosis, that normal p53 acts as a transcription factor for.  We could also use CRISPR to edit the genome itself, possible looking at promoter/enhancer regions for the c-Myc gene, or just cutting the gene out entirely. More research is needed into how to target this therapy, because once that is sorted out, we view this mechanism as a pretty comprehensive solution to many cancers.

The average 260/280 ratio for RNase treated samples was 1.60, while the accepted ratio for pure DNA is approximately 1.8.  This is slightly low to be convinced that our RNase treated sample is pure DNA.  The 260/230 ratio average of 0.60 for RNase treated samples further justifies the impurity of the samples, because a pure nucleic acid should have a 260/230 ratio that is higher than the 260/280 ratio [6].  On the gel, the consistent presence of bands slightly above 10,000 b.p. indicates the presence of nondegraded genomic DNA in the samples.  In the RNase untreated samples, the fields of discoloration at less than 500 b.p. indicate the presence of RNA.  The RNase treated samples did not show these fields, indicating that the RNase worked to degrade the RNA to small enough lengths such that it was unnoticeable on the gel.  Although the RNA did not appear on the RNase treated gel, small RNA fragments were still in the solution even after RNase treatment.  This fact likely explains the NanoDrop results indicating impurities; because RNA was cut into smaller fragments by RNase does not entirely mean it cannot absorb.

The average concentration of the two trials of RNase treated samples is 336.75 ng/uL, the average 260/280 ratio is 1.60, and the average 260/230 ratio is 0.60 [Fig. 1].  The average concentration of the two trials without RNase treatment is 292.05 ng/uL, the average 260/280 ratio is 2.05, and the average 260/230 ratio is 1.57 [Fig. 2].  The average concentration of all four trials is 314.4 ng/uL.  This means in the original 50 uL of solution, there were 15,720 ng of DNA. 

Looking at Figure 3, lane 1 contains a visualized DNA ladder while lane 2 does not.  Lanes 3 and 4, the RNase treated samples, show two bright bands of slightly over 10,000 base pairs in length and no difference in intensity between the diluted and undiluted samples.  Lanes 5 and 6, the untreated samples, show these same two bands, although in the ½ dilution this band is noticeably fainter.  In lanes 5 and 6, there are large discolorations spanning the range of 0 to 500 base pairs, and in the diluted sample, it is noticeably fainter.

A B. distachyon leaf was frozen using liquid nitrogen and then ground up.  DNA extraction buffer (DEB) was added.  DEB contains 1% sodium dodecyl sulfate (SDS), a detergent which binds and solubilizes lipids found in the cell membrane.  DEB also contains 25 mM ethylene diamine tetra-acetic acid (EDTA), which chelates metal ions that enzymes, such as DNases, require to function, thereby inhibiting their function.  DEB also contains b-mercaptoethanol, a disulfide bond cleaver to further inhibit protein function [1].  Potassium acetate (KOAc) was used to precipitate proteins and carbohydrates while nucleic acids remained soluble.  100% isopropanol was used to precipitate the nucleic acids from the salty solution.  The pellet was rinsed with 70% ethanol to remove sodium.  The pellet was resuspended and stored in 50 mL T₁₀E₁ buffer, which contains 10 mM Tris and 1 mM EDTA.

The 50 uL of solution was split evenly, and one sample was digested with RNase A, an endonuclease that cleaves the phosphodiester bonds of single stranded RNA at C and U residues [2].  ½ dilutions of both RNase treated and RNase untreated DNA were prepared using 10 mM Tris.  DNA samples were prepared for gel electrophoresis using glycerol, Bromophenol blue, and Xylene cyanol.  Glycerol is a heavy molecule that ensures DNA sinks in the gel wells.  Bromophenol blue migrates faster than Xylene cyanol, which allows visualization to ensure the samples run adequately but not off the gel [5]. A 0.9% agarose gel was run at 100 volts for 30 minutes.  Agarose forms pores suitable for size separation of nucleic acids [3].  SYBR Safe DNA Gel Stain was used in the gel as a safe alternative to ethidium bromide [4].    

A B. distachyon leaf was frozen using liquid nitrogen and then ground up.  DNA extraction buffer (DEB) was added.  DEB contains 1% sodium dodecyl sulfate (SDS), a detergent which binds and solubilizes lipids found in the cell membrane.  DEB also contains 25 mM ethylene diamine tetra-acetic acid (EDTA), which binds metal ions that enzymes, such as DNases, require to function.  Potassium acetate (KOAc) was used to precipitate proteins and carbohydrates while nucleic acids remained soluble.  100% isopropanol was used to precipitate the nucleic acids from the salty solution.  The pellet was rinsed with 70% ethanol to remove sodium.  The pellet was resuspended and stored in 50 mL T₁₀E₁ buffer, which contains 10 mM Tris and 1 mM EDTA.

https://neurosciencenews.com/memory-flow-neurons-10671/

This study aims to shed light on how the brain remembers certain important locations that contribute to remembering a route of travel.  The study design was a behavioral intervention. The researchers used a two-photon microscope to observe vasoactive intestinal polypeptide-expressing (VIP)-cell activity as mice ran on treadmills while being presented with various sights and sounds.  The researchers focussed on area CA1 of the hippocampus, the area that has previously been shown to be responsible for animal location. The hippocampus at large is responsible for long term memory. It is understood that as an animal learns its route, various excitatory and inhibitory neurons are fired.  What I am left wondering is how the information is stored latently, to be used at a later time.

The 2018 Nobel Prize in Physiology and Medicine was awarded to James P. Allison for his work in tumor immunology.  In 1977, Allison found evidence that leukocytes, also known as white blood cells, the key player of the innate immune response, were prevented from interacting with cancer cells due to the cancer cells having additional proteins.  His work in later years investigated the factors that prevented the immune system from working against cancer cells. He was one of the first to isolate the T cell antigen receptor complex protein. He showed that cytotoxic T-lymphocyte-associated protein 4 (CTLA-4) is a protein receptor in T cells that downregulates the immune response.  Cancer has the ability to induce upregulation of this protein, giving the tumor the ability to avoid inducing immune responses. Allison showed that antibody blockade of CTLA-4 can lead to enhanced tumor immune response. This concept lead to the creation of Ipilimumab, a monoclonal antibody that interacts with CTLA-4.

Caspases are enzymes that are involved in apoptosis.  They are naturally transcribed and translated in the cell, but their unmodified form includes a domain that keeps them inactive.  Once apoptosis is triggered, modifier proteins cleave this domain, allowing the caspases to tetramerize into functional units. A caspase cascade is initiated, in which caspases sequentially cleave the inhibitory domains of their downstream caspases.  In order to kill the cell, caspases work primarily by destroying membranes. They form pores in mitochondrial membranes, allowing Cytochrome c release. This dissipates the membrane potential, so the mitochondria are unable to produce energy for the cell.  Caspases can also perforate the cell membrane, allowing the osmotic pressure to run to equilibrium, which spells disaster for the cell.