Archive for the ‘staining’ Category

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.

Two tissue staining tools for 24-well culture plates.

June 29, 2011

This post highlights two simple tools I made to make tissue staining faster.  I was driven to create these devices after I had to drain and fill over 1000 wells one by one.  The tissue staining these tools are used for are of 300 micron thick coronal sections of embryonic mouse brains that are stored in  24-well plates.

Vacuum Suction Tool



The vacuum suction device can drain 4 wells of a 24-well plate at the same time. These plates are used for many things such as cell culture and tissue staining. Draining the wells of the plate one by one is quite laborious and so this make-shift device greatly reduces the time spent. Disposable pipette tips are placed over the ends of 6″ glass pasteur pipettes to help keep the device clean, vacuum suction keeps the tips on (though as you will see halfway through the video, a small piece of agarose clogged one tip and this caused a loss of suction which led to a tip coming off ).

The technique shown in the video is important in avoiding damage to the tissue and a description of it follows. The plate is held at an angle and the suction device drains fluid from the highest side of the wells. Gravity helps keep the tissue from floating towards the pipette tips. Next the plate is slowly brought level again and the rest of the fluid is removed. Since the level of the fluid is reduced before the plate is brought level, most tissue samples will not be able to be pulled towards the pipette – thus reducing the risk of damage.

How to make one

Materials:  6″ glass pipettes, 3/8″ vinyl tubing, 1/4″ vinyl tubing, hot-glue gun and hot-glue sticks, aquarium air-splitter( I do not know the real name for these), and an attachment substrate (I used the lids of Ikea brand plastic containers, though anything really should work).

How:  Use a 24-well plate as a guide to mark the substrate with a pen.  This is how you will establish the proper spacing for the pipettes.  Once you have the spacing marked, apply a few drops of hot glue and attach each glass pipette one after the other.  At this point the small amount of glue will be strong enough for you to check the spacing but not strong enough for active use.  Once you know the pipettes are spaced evenly horizontally and vertically add copious amounts of glue around and between the pipettes and then finish by adding a second substrate to the top (such that each substrate is like sandwich bread between some meat).

Cut four 1.5″ pieces of 3/8″ tubing and attach these on the ends of each glass pipette.  Next cut four 6″ pieces of 1/4″ tubing and then attach one end of these tubes to the aquarium-air splitter and push the other end of the tubes inside the 3/8″ tubing that is fixed to the pipettes.  Attach the aquarium air-splitter to whichever vacuum line you typically use for draining samples.

Well Filling Tool



When you have over a thousand wells that need to be quickly filled with solution the use of a 3mL plastic pipette simply sucks.  Originally I started using 50mL syringes because I did not have to refill them as often as the 3mL pipettes.  But as my mind wandered during the mindless filling of countless wells I decided I wanted a way to fill more than one at a time.  I had tubing and drip-system fittings laying around and so I decided to build something.

While I failed in my goal was to make something that filled all 24-wells at once, I did succeed at multiplying the number of wells that could be filled.  Ultimately the humongous size of the drip-system fittings prevented me from making a robust filling system because the spacing needed exceeded the size of the 24-well plate.

How to make one

Materials: 50mL syringe, 1/4″ tubing, 1/4″ drip system fittings (1 “T” intersection” and two right angle turn pieces), razor blade

How:  Some of the tips of the 1/4″ drip system fittings need to be truncated in order for them to align with the plate’s wells.  Cut the tips off the fittings as indicated by the red “*” in the figure below.  Use 1/4″ vinyl tubing to attach the fittings together with the tubes as indicated by the placement of blue lines in the figure below.  Lastly, use 1/4″ vinyl tubing to connect the fittings to the 50mL syringe, the placement is indicated by the green line in the figure below.

Note:  The right angle turn fittings will directly touch the T-intersection piece – the tubing merely holds them together.