Fall 2015 Week 8

This week was the club Halloween fundraiser thingy so I wasn't in lab much.
On Tuesday 10/27 I did do one sterilization experiment, using the endophyte infected grass I've been growing in the plant incubator. The sterilization procedure I used for my last experiment was formulated specifically for grass species, and if I know for sure that my grass has been grown from endophyte infected seeds, then I should be able to grown something besides gray mold.

When looking at the silver leaved nightshade under the microscope there are dense trichomes on the surface of the leaf. Trichomes are fine growths on the surface of many plants that can serve many functions. They look like silver hairs protecting delicate plant tissue from harsh sunlight. They can also help prevent loss of water by creating a small bubble of moister air near the surface of the leaf. This is clearly on advantage in the desert where silver leaved night shade lives. Incidentially it is the dense mat of trichomes on SNS that gives it its name. Here is a photo of the SNS leaves under the microscope. The triangular snowflake structures are the trichomes.

I figure If I can get something besides contamination to grow on my grass I know that the procedure works and I simply need to let the silver leaved night shade soak in bleach a little longer to get it to penetrate between the trichomes of the leaf.

Fall 2015 Week 7

Some of the literature about endophytes I've read in the past have said more diversity can be found if plant metabolites are on the petri plates. So a few weeks ago I ran an experiment where I blended up some S. eleaegnifolium leaves, strained the slurry and poured the liquid on petri plates filled with PDA, rose bengal dye and streptomycin to see if I could grow anything other than the usual fungi.

Unfortunately I forgot to surface sterilize the leaves I blended up but I did grow something really interesting. On my plates were small, rounded, sticky looking, creamy white colonies that could have been either bacteria or yeast. I isolated it to a few different media but it only grew well on the TSA plate. Here's what it looked like after a few days: 

But it's way cooler now: 

I did a gram stain and found out the organism is a gram positive bacillus bacterium. This is especially interesting because the streptomycin on the plates I make is supposedly broad spectrum enough to kill most bacteria. The plates were made in April so maybe the antibiotics degrade, I thought. I made a streak plate of 4 different bacteria species I knew to be susceptible to streptomycin on one of the last plates and nothing grew. I assumed both that my antibiotics were working and the bacterium was immune to streptomycin. Then I did an antibiotic test plate with four different antibiotic disks: Streptomycin, vancomycin, penicillin and chloraphenicol. Here's what I found:

Everything except penicillin has a zone of inhibition. WHAT? Not what I expected. The concentration of the antibiotic disk was 10 micrograms, about what the concentration in the agar should have been. I looked back in my notebook and guess what? I diluted my streptomycin to about 1/10 of the strength I thought it was. Decimals are hard to keep track of. So now I know those plates were not exactly what I thought but they did their job well enough while I used them. Interesting. 

Fall 2015 Week 6

This week in lab I ran another sterilization experiment. This time I followed a protocol exactly. I soaked the nightshade leaves in 75% ethanol for three min then in 4% bleach for 5 minutes then ethanol again for 45 seconds. I made nine plates, the more specimens the more data to look at, and have begun to keep careful track of the how things grow on the plates using an Excel spread sheet.

I've found that, once again, within a few days I'm seeing significant fungal growth on all my plates. Its early to know for sure, but I'm pretty sure its the same green-grey mold stuff I've been growing all semester. I'm going to try a few more experiments to attempt to isolate the source of the contamination. I will try plating the tap water I'm rinsing the leaves in as well as test my glassware. I will also think about increasing either the concentration of the bleach or the amount of time spent soaking in the bleach solution.

Another thing I want to try is sterilizing the grass I'm growing that, according to the package, has endophytes in the leaves. If I can isolate endophytes from there I know I'm on the right track.

This is a picture of all the plates I made Spring 2015. They are arranged by decreasing times of sterilization, the first row being 10 minutes and the last being unsterilized. This photo shows that many different kinds of fungi are growing on unsterilized leaves and almost nothing is growing on the longer sterilization times.

Fall 2015 Week 5

This week in lab I did a lot of little tasks to maintain my experiments. I made more media (I simultaneously feel that I don't use my media fast enough and that I spend all my time making media), I replanted endophyte infected grass seeds, took some photos off the lab camera and diluted alcohol (though I now realize I should have made more bleach too).

The plates I made last week testing the sterility of different bleach concentrations were over run with contamination as well. All of the plates had the same green/black mold as before, no matter how high the bleach concentration. I'm beginning to feel frustrated.

Next week I plan on using a sterilization technique I found using 75% ethyl-alcohol and 4% bleach solution-way stronger than any I've used before. They say to dip the plant tissue in alcohol, then bleach, then alcohol again. Apparently the second alcohol dip removes some of the bleach residue, which could be one reason my original experiment with 1.25% bleach did not grow anything at longer sterilization times.

Contamination or not?

Fall 2015 Week 4

Lab has been so productive this week!

On Thursday (9/24/2015) I ran an experiment using the alcohol sterilization method I've been working with all summer on silver leaved nightshade leaves (S. elaeagnifolium). Four days later I knew there was a problem. On Monday I came back to lab and noticed that quadrants A-D all had 1-2 cm sized green, hilly fungal colonies growing on them, which is far too rapid of growth for endophytes.  Quadrant H also had a fungi growing on it that appeared similar to the one that digested my dead grasshopper. All the fungi were growing directly on the leaf suggesting it wasn't contamination introduced because of me but living on the surface of the plant. Based on the physical characteristics of these fungi, I believe they are species I've grown before.

I decided I needed to take a look at my chosen sterilant, 100% ethyl alcohol. When I asked Cori she said that, contrary to assumption, alcohol is a disinfectant not a sterilant, and even then it actually works better with a little water. Thus 70% alcohol is a better disinfectant than 100%. Clearly this isn't going to work the way I thought. I went back to the literature and decided to use bleach again, in the same manner as most current research. Last semester I had a problem with 1.25% bleach killing everything, including potential endophytes, so to find a concentration to work with in the future, I decided to make multiple strength bleach solutions and test which concentration would surface sterilize without penetrating the interior of the leaf. I await conclusive results but nothing has grown on any plates yet, a good sign.

I need to take more photos of my plates so I can remember what these fungi look like, even if they are contaminates. If I remember correctly the microscope photos I took were of spores from green contaminates I grew over the summer.

I made some photos of my plates. This is what I keep growing that looks like contamination:

This is what looks like the same thing that ate the grasshopper: It has the same black spores that bunch in heads and white mycelium.