Archive for the ‘experiments’ Category

A quick and dirty microbiology attempt

July 3, 2012

I recently felt compelled to do some microbiology work.  My goal was to make a crude growth media for microorganisms, swab my mouth for these buggers, and spell my name across a few petri dishes.

edit:  Forgive the bright spot from my lamp, it was the only way to get the letters to show.


Above you can see the letter “J”.  If you look to the right and bottom of the J you can see clumpy looking gunk – that is egg chunks.

I started out by boiling 300 g of sliced potatoes and 1 raw egg  in ~500 mL distilled water for 30 minutes.  Next I filtered the solution through numerous coffee filters and then filtered the solution a second time through a 0.45 micron membrane.  10 g of sucrose and 10 g of agar-agar (hardening agent) were added and stirred for ~10 minutes.  Next I sterilized the media and some glass petri dishes (~$1.50 per dish) in a pressure cooker for 25 minutes at 15 psi.  After sterilization I poured the media into the dishes and allowed them to cool and harden.

My open dishes

My open dishes

After I finished sterilizing the media I noticed the solution had clumpy-chunky junk floating all about.  I assume this was leftover egg that got through the filters, though it could also have been impurities from the agar-agar (least likely of the two).

I used a cotton swab to collect bacteria from my teeth and gums and I spelled my name (a letter per dish) across the plates using the swab.  On one plate I put nothing and on the last plate I spit into it to cover the whole thing.



I was happy to see that everything worked perfectly.  The dish with no bacterial additions had no growth.  The dish I spit into had growth all over and the dishes I wrote letters on only grew in the shape of the letter itself!0



Unfortunately condensation leaked all over the plates when I flipped them and blocked some letters from being photographed.

For future experiments I bought a “fitness multivitamin” that contains all of the amino acids and I am going to try and use it as an egg/tryptone/peptone replacement.

Lab grade vs. home grade electrophoresis buffers

June 8, 2012

(note:  wordpress sucks so it screwed up my images some.)

I wanted to find a cheap way to make electrophoresis buffer from easily accessible ingredients and there were two pieces of knowledge that led me to this experiment.  First I had read about some labs using sodium borate (SB) buffer because it gave great results.  Secondly, I knew that borate and boric acid were easy to find at stores in the forms of roach poison (boric acid) and borax (sodium tetraborate).

Originally I had intended to compare a large number of buffers; TAE, TBE, SB (molecular biology grade), and SB (home grade).  Due to my own personal shortages of lab-grade materials I had reduce down to a comparison between TBE, SB (home grade), and SB with EDTA (home grade).

All gels were 1% agarose.  Gel images were made with Foto/Phoresis I transilluminator and gel images were recorded with an iPhone 3GS camera.

Tris-Borate-EDTA (TBE)

Purchased from the “Online Science Mall” as a 5X concentrate.  Without a doubt this worked better than the buffers I cobbled together.


SB Buffer (home-grade)

SB buffer made by adding 1.91g of Sodium borate decahydrate (Borax by 20-mule team) to ~800mL distilled H2O (Target), pH was adjusted with 0.4M Boric Acid (Roach Away by Enoz) and then diluted to final volume with more distilled water.

SB Buffer with EDTA (home-grade)


A 600 mL aliquot of the SB buffer, made previously, had 1.2 mL of 0.5 M EDTA (molecular grade) added to reach 2mM final concentration.

I decided to test the SB buffer with EDTA to see if there were any nuclease related problems occurring.  2/3rds of the way through the project it occurred to me I was using Target brand distilled water instead of nuclease free water when mixing the 2-log DNA.  It just simply slipped my mind.  So this third gel buffer test was done with EDTA and nuclease free water to see if the results would be drastically different or not and they were not.  I believe any differences in the images is due to my lack of a proper gel documenting system.

Given the choice between the three buffers, TBE is the superior.  I do not mean to say that one could not use the SB home grade buffers and get usable results, but rather they just are not as good.  Being that store bought TBE and TAE are not crazy-expensive and buffers can be reused a couple of times, there is not a lot of impetus to use home-grade SB buffers.

My results for the SB buffer tests do not even come close to mirroring the results from the Brody paper I linked earlier, I suspect that my preparation or formulation of SB may be flawed.

While unfortunate that I couldn’t test more buffers, I was happy to figure out that TBE will likely work for my PCR projects in the future and if I am ever mismanage my supply of TBE I now know I can make up some SB buffer to get by.

Comparison of agarose, agar-agar, and cleaned agar-agar for gel electrophoresis of DNA

January 31, 2012

(cross posted here and at Citizen Science Quaterly)

Gel electrophoresis is a core technique used in molecular biology laboratories. The gels used for electrophoresis are almost always agarose which is a polysaccharide purified from agar-agar. Agar-agar is obtained from red algae and contains a variety of impurities along with the agarose. Agar-agar is often used as a substitute for gelatin in vegetarian dishes and is used to make some cool culinary creations. Pure agarose is obtained by separating it from agar-agar. I have heard from variety of sources that agar-agar could be used in place of agarose for gel electrophoresis of DNA. Given that agarose costs around $1.00 per gram and agar-agar costs around $0.05 per gram I thought it was worthwhile to check the efficacy of agar-agar. My results show that agar-agar is not an acceptable substitute for agarose (see the image above).

For more details and thoughts read the rest of this post.

Cleaning the Agar-agar

The night before I ran the experiment I decided that it would be nice if I tried to “clean” the agar-agar of some impurities and run this as an additional treatment group. Since I did this last minute I did not completely think my plan of attack through and actually made the agar-agar worse. I used 50% isopropanol and distilled water to rinse the agar-agar through a coffee filter.

After running the experiment and seeing my results I did the research I should have done prior to the experiment. A quick google search showed me a technique for purifying agarose from agar-agar using propylene glycol. Click here for the site in question.

Failed Purification Protocol:

  1. Weighed out 11.7g agar-agar into a coffee filter (2 thick).
  2. Saturated agar with distilled H2O (dH2O).
  3. Poured 100mL of dH2O through the slurry and allowed it to drain.
  4. Stirred slurry with a spatula.
  5. Pourd 100mL of dH2O through the slurry and allowed it to drain.
  6. Stirred slurry with a spatula.
  7. Poured 100mL of 50% isoproanol into the slurry.
  8. Stirred slurry with a spatula.
  9. Once the majority of fluid drained out I placed the agar-agar and coffee filter into an oven at 90C for 5 hours to dry.

Experiment Parameters

  • 1.2% gel
  • TAE Buffer
  • 2.5 hours
  • 70 volts
  • Biotium GelRed Stain (used as a precast additive)
  • Promega 1kb DNA Ladder (left lane)
  • NEB 2-Log DNA Ladder (right lane)

Pictures and Comments

Agarose gel atop the UV transilluminator.

I had minor issues taking pictures of the gels because too much light was reaching the camera lens so I made a aluminum foil frame to block out extra light.

Close-up of the agarose gel’s bands (leftside = (+) pole, rightside = (-))

All 3 gels deformed on the end closest to the (-) electrode and I am not sure why. I suspected temperature at first but it subjectively seemed constant throughout the gel rig. Next I thought it was the gels positioning, however when checked I found each end to be equally distant from both electrodes.

Compare the picture above to the reference gel below (in black and white). This reference gel was the first gel I ran with my gel rig and it ran for 80min at 70V using the same TAE buffer used in this experiment and there was no deformation of the gels wells. The two differences between the reference gel and the newest gels were time and staining compound (2.5 hours vs. 80min and ethidium bromide vs. GelRed). If this problem persist I will have to run a bunch of gels to figure this out.

See blurb above for information about this reference gel.

All 3 experimental gels on the UV transilluminator. From left to right; agar-agar, cleaned agar-agar, and agarose.


Based on my results I do not recommend purchasing agar-agar for gel electrophoresis. The only exception I would make is if the intent is to use propylene glycol to separate the agarose from the agar-agar (click here for agarose separation with propylene glycol). Keep in mind that there could be nuclease and cation contamination in the propylene glycol purified agarose.

While agarose is a lot more expensive than agar-agar, it is not expensive relative to everything else needed for conducting molecular biology at home. For the sake of simplicity, reliability, and time I intend on using only agarose going forward.

What is next

My next post will compare running buffers between gels – specifically molecular biology grade Tris-Acetate-EDTA (TAE buffer), Molecular Bio grade sodium borate buffer (SB Buffer), and sodium borate buffer made from borax and cockroach poison (boric acid). If there is any simple household substances you want me to try as a running buffer, let me know. I have a lot of school and work projects going on so this upcoming post may be a few weeks out.

Finally got around to testing some gel electrophoresis.

January 16, 2012

It took much longer for me to get a round to testing out my gel electrophoresis equipment than I thought it would.  For now I have merely got it to work.  Next I will try and fine tune it to increase the quality of the gels.  More on that below.

This isn’t the most informative post but I was kind of frustrated by a lack of information when I was troubleshooting so I figured I would throw some data out and hope it helps someone.

Note:  The cameras on my phone and iPad both captured more wavelengths of light than I could see, so these images look worse than the gel actually is.

Zoomed out gel.

Zoomed out gel.

Gel closeup

Gel closeup

Gel Parameters:

  • GEL = 1.5% food-grade agar-agar gel (not agarose)
  • DNA LADDER= New England Biolabs 2-log DNA ladder
  • STAIN = GelRed Stain (Vendor; Biotium) (Approx. Equiv. to Ethidium Bromide, except safe).  Stain was used in the precast gel (1x) context.
  • TRANSILLUMINATOR = Fotophoresis I (Fotodyne)
  • BUFFER = TAE (MB grade reagents)
Failed gel.

Failed gel.

The image above is what the first two gels looked like – no fluorescence at all.  I still do not know for sure why they failed but I narrowed it down to either the composition of the DNA ladder or the staining method.

My set-up worked when I used GelRed in the molten agar-agar and composed the DNA ladder per the manufacturers instructions.  I used 5-10uL of ladder and 1-2uL of loading buffer on the failed gels whereas I used 1uL of ladder, 4uL H2O, and 1uL loading buffer on the successful gel.  During the failed gels I tried to use the 3x post-electrophoresis stain procedure with staining times between 0.5-1.5 hours – all with no luck.  Obviously changing two variables at once confounds the results – but at least I have a baseline now.

The setup

The setup

This was the gel box I built and used.  You can read my instructions for how to build it here.

Muh lab.

Muh lab.

This is ~90% of my apartment lab.

My next goal is to work out how to fine tune the procedure.  I am going to compare the gel quality in cases where I use reagent grade vs. industrial and food grade chemicals.  I am hoping that borax (sodium tetraborate) and roach poison (boric acid) buffer will work as good as its reagent grade cousin.

Electrophoresis electrodes.

October 5, 2011

I have been trying to determine which materials work for electrophoresis electrodes.  My hope was that I could find something cheaper than platinum (I know it will work but it cost a lot).  Below are some things I have tried and the results.

  • Copper – Dissolves in solution and makes the solution blue-green (based on reports from various websites, only listed material I did not test).
  • Generic wire (22 gauge) – Mix of copper and iron.  Works as an anode but not a cathode.  As an anode copper leaked into solution and the wire became green and red striped, presumably because the braided strands were different materials.
  • Sterling silver (24 gauge) – Mix of silver (~90%), copper (~5%), and other elements.  Does not work for anode and I have not tested it as a cathode.  As an anode it leaked copper into solution, a black precipitate formed (steel was the cathode in this test) on top of both electrodes and the majority of the wire disintegrated.
  • Braided steel cable (around 1mm in diamter) – Intended as a “steel leader” for fishing.  Works as an anode but not as a cathode.  If it is used as a cathode it will disintegrate and break.
The failures I had have finally convinced me to purchased platinum wire.  The platinum will work for sure but I do not know if what I bought was thick enough so I will have to wait until it arrives in the mail.

DIY centrifuges, used centrifuges, and a fun experiment.

September 3, 2011

The background that led to the experiment

I had been thinking about building a centrifuge out of a blender but first I decided to research what other people had done before I tried my own hand at it.  I ran across a number of designs which included  “Dremmelfuge“” and a handheld centrifuge which will eventually damage something.  The best of centrifuges was one that used mixing bowls and a blender.

My own thought was to attach the caps of dry-erase board markers to a blender rotor using wire and gorilla glue.  In my design I wanted to use the blender pitcher as a safety vessel for in the likely event something went flying – but this provided a lot of design challenges.  Ultimately I decided that my idea was bad and most of the other designs were too risky.  If I were to ever revisit centrifuge construction definitely would make a variant of the blender which used mixing bowls.

All of this ended with me surfing eBay and discovering it was not too expensive to just buy a used centrifuge (from $100-150) – which I did.  I manged to find the centrifuge depicted below.  I find the aesthetic design to be quite pleasing and it is a bit sad that this style is not used anymore.  Now that I have a centrifuge, it is time to use it (see below for more)!

My little buddy. Cheap and effective.

A quick centrifuge explanation

Centrifuges spin samples around extremely fast (often >10,000 RPM).  Doing so causes centripetal force to be exerted upon the samples.  In the case of my centrifuge, it spins fast enough for the samples to have the force of 13000 times earths gravity exerted upon them.


What happens when ketchup, milk, sriracha sauce, and russian salad dressing are exposed to 13,000 x earth gravity for 30 minutes?


The source material precentrifugation.

Here are the specimens, precentrifugation.  From left to right we have non-fat milk (Trade Joe’s), Ketchup (Heinz), Sriracha Sauce (Huy Fong Foods) and Russian Saland Dressing (Wish Bone).

First we load the machine…

Rotor with tubes.

Next we run the machine for 30 minutes and we get….

sriracha sauce 13000 x g 30min

Separated out kind of like blood  (serum on top, cells on the bottom).  I would guess that the bottom layer is chili pepper fragments.

russian salad dressing 13000g 30min

This salad dressing has A LOT of ingredients and I will even try to guess as to which layer is what.

nonfat milk 13000g 30min

Milk proteins should be what has collected at the bottom.

heinz ketchup 13000g 30min

The ketchup surprisingly did not separate into layers.

My take home message from this experience was that sometimes the effort and danger in building something myself may not be worth it when one considers the cost and value of used equipment.