February 11, 2014 in Posts
February 5, 2014 in Posts
Pictured below is the fungal endophyte we isolated from a Ginko tree fermenting in 300mL of Sabouraud dextrose broth. We performed an ethyl acetate extraction on the liquid partition of the broth, after filtering out the mycelial mass through cheesecloth. We dried the combined ethyl acetate partitions over sodium sulfate and concentrated on a rotovap. Due to an analytical balance malfunction, we were not able to get an accurate mass of recovered crude extract, but by eye it appeared to be a few mg.
The extract was brought up in 200uL DMSO and a small analytical sample was taken to run on an LCMS. (Chromatograms to be posted soon)
We wanted to do a quick disc-diffusion test to see if there were any compounds we had extracted in the fermentation broth that had antibacterial properties, so we plated some E. Coli and Staph. Aureus and put 2,10, and 20uL respectively on filter paper discs and let incubate at 37C overnight. Much to our excitement, we saw a concentration dependent zone of inhibition around the 10uL and 20uL discs for E. Coli.
This was a really quick and really crude disc diffusion test (we know we were missing the concentration of our original extract) but we still wanted to be able to prove to ourselves that we could isolate an endophyte, subculture it, liquid culture it, and extract the broth and test for bio-active compounds.
Notice on the 20uL disc that the DMSO appeared to “drip down” when the plate was flipped, probably because not enough time was given for it to soak into the agar, leading to a skewed portion of inhibited growth. Nevertheless, it is clear that as the concentration of the doped discs increases, so to does the ring of inhibition. Now comes the hard part…figuring out what compound(s) are responsible for the activity in a mixture of many!
Another interesting observation is that around the Staph A. discs, there appears to be a ring of where the DMSO diffused, with perhaps a little less growth of the organism, but not clear inhibition.
August 12, 2013 in Posts
A few samples were selected for hyphal tipping and pure culture isolation. Shown, in no particular order, are some of the samples to be taken and re-plated on fresh PDA. Small pieces of the mycelia were excised with a sterile scalpel and placed in the center of fresh plates. In some instances, like the dark red pigmented colony growing within a few others, hope of a perfectly pure inoculum wasn’t expected, and instead we were just interested to see what might grow on the plate.
There were instances like the first picture, in which a very homogeneous mycelial mass was obtained, which looks like a pure culture isolation. We will perform DNA isolation and ITS amplification on some of the select organisms, as well as observe some of them under the microscope. This week, agar plugs of pure strains will be put into liquid broth to ferment, with the downstream goal of processing the fermentation broth to isolate a crude extract of secondary metabolites to be used for bio-assay guided fractionation.
August 6, 2013 in Posts
We’ve done enough PCR on fungal cultures and samples to know that our PCR program, enzymes, and primers are working as intended. In order to give us confidence in the ITS amplification of unknown endophytic fungal samples, we wanted to amplify a known species and confirm that we can get clean sequencing reads.
We performed a DNA extraction using our homemade buffers and an isopropanol precipitation followed by a 50uL PCR reaction.
PCR reaction conditions are as follows:
2 uL ITS master mix (ITS1F, ITS4R – see older posts for primer sequences)
25uL NEB Taq Master Mix
After the reaction was run, a 1% agarose gel was cast, and 5uL of the PCR reactions were loaded, alongside a 1kb NEB ladder. https://www.neb.com/products/n3232-1-kb-dna-ladder
Lanes were as follows:
1 – NEB 1kb ladder
2 – + control
3 – Sample 1
4 – Sample 2
5 – Sample 1 genomic DNA (5uL load)
~40 uL was withdrawn from the sampletube taking care not to get mineral oil, and processed with an Epoch Life Science PCR purification spin kit.
Eluted DNA and a fresh set of primers were sent for sequencing, although due to very odd circumstances, only Lane 4 (faint band at ~700 bp) arrived.
Sequencing was provided by Wyzer Biosciences in Cambridge, MA https://www.wyzerbio.com/wiser/#aboutUs
We highly recommend them! They went above and beyond in order to assure we got quality sequencing data as well as went out of their way to personally call us and discuss sequencing parameters after our primers did not arrive, even with a sample order size of two. The company is run by fantastic people and is well worth supporting.
Here are the FASTA files as follows.
Put them into
Our sequencing chromatograms looked good, with one giving a read of over ~700 bases, while the other was cut short at ~400 bases. Nevertheless, for a first try the very crude attempt at sequencing went better than expected, and enough data was retrieved to give a greater than 99% confirmation on the sample sequenced via BLAST. We did indeed do ITS amplification of Agaricus Bisporus, and the sequencing data confirms that our protocols for DNA isolation, amplification, and purification are working as intended, and lend confidence to future endeavors in the identification of unknown fungal endophytes.
August 5, 2013 in Posts
In the quest to isolate new endophytic fungi and build a strain library for screening, one of the main issues we face is still being unable to afford a laminar flow hood. Sterility is a big concern in endophyte isolation, as it is very important to assure that any microorganisms found in the petri dish are coming from within the plant tissue, not on it, or outside of it. A quick google search will return many home-built laminar flow hoods as well as more simple versions such as a “sterile hood” which consists of a fish tank or plastic container turned on its side and cleaned thoroughly with bleach/isopropyl alcohol.
In this experiment, we wanted to test the bare minimum (ie, no laminar flow hood, no make-shift sterile hood) and see if we could get a control with no growth and samples that looked like endophytic fungi were growing from them.
Samples were taken from:
English Ivy http://en.wikipedia.org/wiki/Hedera_helix
Rhododendron http://en.wikipedia.org/wiki/Rhododendron (only the fresh leaf was plated, the damaged batch of small leaves and stems were discarded)
Ginko http://en.wikipedia.org/wiki/Ginkgo and a few leaves with small branches/stems were selected from each.
Samples were washed for 30s under tap water, then cut into small sections and placed in 5% sodium hypochlorite for 30s, followed by 70% ethanol for 30s, followed by two sterile water washes for 30s each. 100uL of the final water wash was plated as the control.
Afterwards, samples were plated on PDA (potato dextrose agar) supplemented with chloramphenicol to inhibit bacterial growth,and left in a dark cabinet at room temperature after being sealed with parafilm.
Fast forward a week and here are how the plates look
We’ll work on a follow up post on how the samples were re-plated via hyphal tipping to try to get pure cultures, but the good news is, the control doesn’t have any growth! Also, on the Rhododendron leaf stem (the last two pictures) you can see the white mycelium growing out of the vascular tissue, pretty cool! In some of the cases, like leaving the outer bark on the ginko stem, contamination from epiphytes/ectophytes will occur, but it is still interesting observing the organisms grow.
There was at least one yeast colony on one of the ginko leaves, and a few samples were selected to re-plate to try to get pure cultures. Ideally, we’d be observing the growth of the organisms daily and taking note of their growth rates, coloration, and other features, but we both have 9-5′s and sometimes cannot always get to the lab every day. The Ivy sample was interesting to look at, but mostly discarded for further isolation because of the density of growth of the organisms.
Needless to say, it seems that with a little bit of practice and a very crude aseptic technique, you can isolate some endophytes of your own.
June 11, 2013 in Posts
After many gels that looked like the picture above, with only primer dimers, we had to reconsider our approach to PCR.
Often times in the sciences, experiments don’t go as planned, and what we’ve come to learn is that it is incredibly important how you handle those situations, as neglecting sources of error can lead you on a hunt for variables that may or not be important. You could opt to ignore the results as a fluke and blindly repeat the same experiment, wasting time and consumables, or approach the problem in a scientific manor.
In our particular case, the one thing we were varying perhaps a little too much was the way we went about genomic DNA isolation.
We had had success with a plant genomic DNA spin kit on fungal samples, but with a ~20-30 minute processing time per sample and cost of ~$1.50 we looked to something faster and less expensive.
Previously, we had had success with a crude colony PCR approach mentioned in one of our previous posts, which basically involved grinding mycelia in 50mM NaOH. The buffer the Taq came in seemed to handle the pH change fine and we got amplification. On subsequent tries though, our results were inconsistent, with some colonies yielding successful amplification, and others yielding nothing but primer dimers. The mass of initial mycelia to be ground as a sample seemed to have a big impact on the results of amplification. A petri dish covered in what looks like a mass of mycelium results in little more than a few milligrams when scraped into a concentrate.
We had a few more failed gels, at times blaming old polymerase, bad primers, freeze thaw cycles, a poor PCR program, and even the water we were using. After doing a few runs where we caught ourselves varying too many things, thus making the task of narrowing down the source of error that much harder, we finally came to our senses and realized we couldn’t brute force our way into successful PCR, and that instead, a little extra time spent planning the experiment would save time, materials, and frustration in the long run.
We were finally able to point the finger at poor template DNA (or lack there-of in some cases) and realized that although colony PCR might work pretty consistently for bacteria, it was not giving reproducible results for non-yeast filamentous fungi.
After doing some research online for protocols involving rapid DNA isolation from fungal colonies, we happened upon a paper which seemed to present good results while using cheap and readily available materials, and gave it a shot.
We kept all other variables the same (PCR program, polymerase, primers) and did extractions with our new solutions on multiple fungal samples, with the results below.
Lanes are as follows:
- 1kb ladder
- Porchini mushroom
- Portabella mushroom
- Shitake mushroom
- Maitake mushroom
- Enoki mushroom
- King Oyster mushroom
- Crimini mushroom
- White button mushroom
- 1:2 primer dilution
- 1:4 primer dilution
- 1:2 template DNA dilution
- 1:4 template DNA dilution
- positive control
- negative control
- plant samples PCR (failed)
- plant sample PCR (you can see a faint band)
This was one of our most successful PCR reactions and we were quite happy with it as it was a validation of our efforts in trying to optimize conditions for fungal DNA amplification as well as fungal DNA isolation. The DNA isolation procedure worked on seven of the eight mushroom samples, though judging by the intensity of the primer dimers on the one lane that didn’t have an amplification product, it looks like template DNA was simply missing in that particular case, rather than just a poor isolation. We were also able to see just how big of an impact primer concentration can have on PCR, as the bands lose much of their intensity at just a 1:2 dilution, and disappear with a 1:4 dilution, yet diluting by the same amount with the template DNA almost looks as if it improved amplification in the 1:2 dilution, and band intensity seemed unchanged in the 1:4 dilution.
Not bad for a couple of chemists.
April 23, 2013 in Posts
We have successfully shown that we are able to barcode an unknown fungal sample, but the question now is, can we reproduce it? We have gathered the necessary tools and now face the task of making sure that our protocols are thorough, easy to follow and most importantly, reproducible. Over the past week we have ran ~28 reactions (DNA extraction, PCR, isolation). Unfortunately, what we have found is that our protocol is not as robust as we were hoping.
Samples 1 to 3 are DNA extractions from the same fungal colony using 3 different DNA extraction solutions. All used ITS primers (see below for primer sequences).
Sample 1 used an extraction solution of Tris-HCl, EDTA and SDS.
Sample 2 used 20mM NaOH
Sample 3 used 20mM KOH.
Samples 4 and 5 are DNA extractions from the same plant sample using Epoch Life Sciences Plant Genomic DNA Extraction Kit (Cat # 1560050).
Sample 4 used primer set LEP
Sample 5 used primer set rbcLA
Sample 6 used primer set rbcLA and a plant template extracted using 80mM NaOH
Sample 7 used fungal DNA extracted using 80mM NaOH and ITS primers
So what do we think is going on?
As for Sample 1 , we initially thought our extraction solution would be effective for lysing cells and extracting gDNA. Unfortunately, at the time we were not thinking of the impact SDS and EDTA would have on the polymerase/overall PCR reaction. SDS is a strong detergent that is often used to lyse cells and extract DNA, but is also very good at unfolding proteins. That is not good news for our polymerase. The other problem with sample 1 was the EDTA. Metal ions (specifically Mg2+) are essential co-factors for our polymerase to work correctly. Having too little or too much free Mg2+ can be devastating to a PCR reaction (as we are finding). In our case, the presence of excess EDTA and SDS rendered our reaction, unreactive…
Sample 2 and 3 seemed to work very well with 20mM NaOH and KOH, respectively. The simple colony PCR protocol was a tip given to us by an experienced molecular biologist. We figured that the base would be aggressive enough (but not too aggressive) to lyse cells and make DNA available for extraction. We were right! It felt good to know that that method worked since it’s so cheap and fast.
Sample 4 was expected to fail as the primers were designed for insect DNA amplification.
Sample 5 worked as anticipated. Again, this sample used the Epoch kit, so started with clean plant genomic template DNA.
Sample 6 was a plant prep using 80mM NaOH. It was encouraging to see enough plant DNA could be prepared by such a simple, fast, and cheap method and give comparative amplification to Sample 5, which was prepared using a spin kit costing ~$1.50 per prep.
Sample 7 was a failed ascomycete fungal sample prepared by grinding the sample in 80mM NaOH.
Standard 25uL reaction
1uL template DNA
0.5uL 10uM forward primer
0.5uL 10uM reverse primer
12.5uL NEB Taq 2x Master Mix
Overall, these 7 samples showed us that out procedure was somewhat reproducible, but we do have some variables to consider looking into. Primer design and PCR conditions for each primer set are definitely worth checking out. Also, maybe our Taq is starting to go bad; we have taken it in and out of the freezer dozens of times, same goes for our primers. We will definitely look into this further.
April 19, 2013 in Posts
This is a simple disc diffusion assay that we conjured up. The aim here is to see the effects an antibiotic has on a bacterial colony. In this case, we plated E.coli on PDA plates with no antibiotics in the media. We made a stock solution of 1mg/mL chloramphenicol in absolute ethanol and soaked small pieces of filter paper in various amounts (0ug, 5ug, 10ug, 20ug and 50ug) and placed them on plates that were freshly coated with E.coli. We incubated at 37C overnight.
The plate on the left is the before shot, prior to incubation. On the right is the plate after incubating at 37C overnight. You can clearly see the growth of E.coli around the 0ug disc (containing no chloramphenicol) and the lack of growth around the discs that do contain chloramphenicol. You can also see that the inhibition of growth is larger next to the 50ug disc versus the 5ug disc, as expected.
It’s also worthy to note that chloramphenicol is a bacteriostatic antibiotic, meaning that it does not kill bacteria, but prevents them from reproducing. If we took a plate that already had a full lawn of E.coli, we wouldn’t see complete absence of colonies after incubation, just less growth from those colonies exposed to the antibiotic. On the other hand, a bactericidal antibiotic (such as penicillin) will in fact kill bacterial colonies.
Overall, this nicely demonstrates how a disc diffusion assay works. In the future, we plan to isolate our own compounds and use this same setup to test for bioactivity and potency of our isolates. Stay tuned…
Below you can see another plate which had no antibiotics in the media and was not exposed to a disc diffusion assay.
December 30, 2012 in Posts
We’ve finally acquired enough equipment from ebay, suppliers, and junk piles to conduct our own full scale test, so to speak, of our current makeshift DIY laboratory. The whole experiment involved DNA isolation, PCR, and a gel run with visualization in order to confirm that our equipment and reagents worked as intended. We managed to complete the entire experiment using (mostly) all of our own equipment, with great results.
Our equipment/reagents we were testing:
Idaho Tech RapidCycler PCR machine
Eppendorf 5415C high speed microcentrifuge
Epoch Life Sciences plant genomic DNA isolation kit
GelGreen, agarose, TAE
Gel Electrophoresis box purchased from IO Rodeo
Electrophoresis power supply from junk pile
Fungal ID primers
Purpose of experiment – To isolate genomic DNA from a fungus, then amplify a barcoding sequence in the ITS region of its genome via PCR, followed by verification with a gel and ultimately resulting in the PCR product being sequenced and having the results BLAST searched in the NCBI database to confirm phylogeny/species ID.
Experimental overview – A button mushroom (Agaricus Bisporous) was purchased from the store for 16 cents, and a plant genomic DNA prep kit from Epoch was used on a small sample from both the cap and the stem of the mushroom (freshly cut in half).
We used two different pairs of primers: ITS1 / ITS4 and NLB4 / NSI1
These primers target an area of DNA (known as rDNA) which codes for ribosomal RNA comprising the large and small ribosomal sub-units (LSU, SSU), and the internal transcribed spacer (ITS) which has become a “standard” for barcoding and species identification of fungi.
NSI1 (forward) GAT TGA ATG GCT TAG TGA GG
NLB4 (reverse) GGA TTC TCA CCC TCT ATG AC
ITS1 (forward) TCC GTA GGT GAA CCT GCG G
ITS4 (reverse) TCC TCC GCT TAT TGA TAT GC
We expected products of around ~700 for ITS and ~900-1kb for NLB/NSI but these can vary for different mushroom varieties (ie. basiodiomycetes vs ascomycetes etc)
We tested our PCR machine, an Idaho Technologies Rapid Cycler vs a peltier effect based machine. The Idaho Tech machine uses a halogen lamp and a tornado-like air circulation effect for fast ramp times. Neither of the machines had heated lid capabilities, so mineral oil was used in each PCR reaction to avoid evaporation.
After gDNA isolation, PCR reactions were setup as follows for a final reaction volume of 25 uL in 200uL thin walled plastic PCR tubes.
10uM forward primer 0.5uL
10uM reverse primer 0.5uL
Template DNA 2uL
OneTaq 2x Master mix 12.5 uL
ddH2O 9.5 uL
Our PCR program was as follows:
94C for 30s
54C for 45s
72C for 45s
30 cycles (we wanted 35 but time didn’t allow for it)
Even at 30 cycles and those program times, the Idaho Tech machine finished ~30-45 minutes before the peltier machine, boosting confidence in the PCR machine purchased off ebay for $50.
10uL of our PCR product was run on a 1% agarose gel pre-stained with GelGreen. Product sizes were compared to 1kb NEB ladder, and single bands were observed for all wells, minus the presence of some primer dimers.
Optimization of PCR is needed, but this is a great first step which proves that with a little ingenuity and some patience in acquiring decent equipment on ebay, one can do some decent molecular biology on short funds.
The samples are being prepped for sequencing and another post will follow giving our findings.
This is an update post on the quest to isolate a Penicillium strain from citrus fruit, in the hopes of at some point along the journey finding some penicillin. A video was recently uploaded to our youtube channel that briefly explained the experimental conditions. Basically, three different liquid cultures (not pure or sterile- they were grown in coffee cups) were grown for a few weeks in the hope that some would produce pencillin, and we could test for its presence by a simple disc diffusion assay. Much simpler said than done, as is often the case in science!
The video covers how the experiment was setup, but does not cover the results, which didn’t boast well for our novel coffee cup penicillin cultivation technology.
As you can see, none of the experimental homemade disc diffusion assays showed any signs of inhibition at 5, 10, 15, 20 uL concentrations against Bacillus subtilus - the only gram positive bacterial strain we could get our hands on. The “experimental” discs had to be hand cut from new filter paper, so weren’t perfect circles, but rather an assortment of rhomboids created by a pair of scissors.
The one semi-perplexing thing was that even the control plate showed no rings of inhibition with the 10 unit penicillin discs. The reasons for that could be; bad penicillin, resistant bacterial strain, the fact that it took me 3 days to check the plates after the initial plating. If I had to make an educated guess, I’d go with #3. Plates are usually checked daily, and the fact that it took 3 days before the plates could be checked probably meant whatever penicillin had been present had been “used up” in a sense, allowing the bacteria to colonize the area after the initial 10units were gone.
After doing some googling, I came across this pretty nifty image, which showed just how innefective pencillin is against B. subtilis. (Not my photo! Thanks docpilgrim!) Notice how the P10 disc shows the smallest ring of inhibition, and that’s the same P10 disc we were using in our control.
So back to the drawing board! For the time being our goal is to use our ITS primers to try to ID the strains we did isolate. On the upside, this has been a pretty pleasant mold cultivation project, as everything smells like lemons.