This instructable was based off a ScienceHackDay SanFrancisco team that I was part of along with Patrik D’haeseleer, Bonnie Barrilleaux, Lily Lew, Joseph Elsbernd, and Michelle Peters. It is really easy to do and a fun way to incorporate molecular biology into your social life!
Posts Tagged ‘DIYbio’
(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.
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.
Cross-posted here and at Citizen Science Quarterly.
I wanted to purchase a pipette stand but it seemed like such a frivolous purchase. Why should I spend between $20 (eBay) and $60 (venders) to hold my pipettes vertically? As much as I wanted a pretty acrylic stand I just could not pull the trigger and buy one (I’d rather buy DNA ladders). This post will briefly go over how I built a pipette stand out of PVC pipe.
These instructions are for a linear pipette stand. The circular/revolving stands are an annoyance and an example of bad design. If you are suffering through one of the revolving stands I urge you to abandon it for a linear stand.
Saw or PVC cutter – Either work but the PVC cutter is easier to use and costs $10-20.
Marker and Tape measure
(above) All of the supplies needed for the stand.
Pipe is usually sold in 10 foot lengths. Most stores will cut it down smaller for free if asked. Some stores even sell shorter segments. Cut the pipe into four 8″ lengths, two 6″ lengths, and six 2″ lengths.
Purchase four T-shaped fittings and six right-angle fittings.
T-shaped fittings (4) = $0.80
90deg fittings (6) = $1.02
½” pipe (10 feet) = $1.65
Total = $3.47
The construction of the stand is simple. All that has to be done is to cut the pipe and then assemble the structure.
As mentioned in the methods, cut the pipe into four 8″ lengths, two 6″ lengths, and six 2″ lengths.
Next connect the fittings together with all of the 2″ lengths of pipe as shown below.
Now add the 8″ and 6″ lengths of pipe – the stand is now complete.
(above) A view of the stand from the bottom.
(above) Fully assembled stand.
(above) A view of my home-lab with the finished pipette stand.
I needed to pellet a saline solution containing cheek cells for genomic DNA isolation but I did not have a centrifuge that could hold 50mL tubes. The solution was to tie the centrifuge tube to my ceiling fan with shoestring. The ceiling fan centrifuge was ghetto, scary-looking, and effective (it had 3 different speed settings!). I made a video, check it out below.
Before using the fan I first tried to spin the shoelace (with tube) by hand – but this did not work out at all as the solution kept getting stirred.
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:
- Weighed out 11.7g agar-agar into a coffee filter (2 thick).
- Saturated agar with distilled H2O (dH2O).
- Poured 100mL of dH2O through the slurry and allowed it to drain.
- Stirred slurry with a spatula.
- Pourd 100mL of dH2O through the slurry and allowed it to drain.
- Stirred slurry with a spatula.
- Poured 100mL of 50% isoproanol into the slurry.
- Stirred slurry with a spatula.
- 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.
- 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.
The protocol I use for collecting human DNA samples requires tubes to be vortexed for 10 minutes. Standing around for a sixth of an hour is not my idea of fun so I decided to get a foam tube holder. Unsurprisingly a piece of foam with holes in it costs 50 dollars. As per my usual I wanted to be a cheapass and thus I built my own foam tube holder with some things I had lying around. If you want to see the DIY tube holder in action, watch the youtube video below and scroll further down for instructions for building your own.
Before attempting this guide make sure your vortexer will work with this method. Check the type of head piece present on your vortexer, consult the diagram below for an example of two different types (#12+13 and #14). This guide works for vortexers with head pieces that match #14. New vortexers usually come with both of these pieces and used vortexers (like the one I bought) may only come with one type.
The tools and consumables I list are not absolute. Use your noggin and substitute if necessary.
Dremel (a drill can be used for some steps but not for carving foam)
- Sanding bits are needed for foam carving (see image below)
- Cutting tool
- Drill bit
- Sanding bits are needed for foam carving (see image below)
- Ruler or measuring tape
- Hand saw
- Compass (optional)
- Foam block
- plastic box (I used a large pipette tip box that was 4″x5″)
- Rubber bands
- Foam block
These are the two dremel tips I used to carve the foam.
Two types of modifications need to be done to the plastic box. First, a hole needs to be made in the center that will allow the vortexer head to poke through. This alteration will prevent the tube holder from wobbling off of the vortexer head. For my vortexer the size of the required hole was 1-1/4″. The second modification is the addition of 4 grooves to the top of the box so that the rubber bands do not slip.
Prior to drilling and cutting grooves. Yellow circles indicate approximate target locations for the grooves.
After drilling and cutting grooves
Close-up of the grooves
How the box fits onto the vortexer head.
Now it is time to carve the block of foam. My box was not perfectly square; the top of the box (the opening) is larger than the bottom of the box. This odd shape can be ignored but do make sure to use the top of the box for determining the foam block size. I made a block that was 4-1/4″ x 5-1/4″ x 2″. It is very important that the width and length match or slightly exceed the size of the box so that the foam sits tightly inside the box.
My original block was too tall. Rather than cut grooves in the foam for the rubber bands and have a tall block (which is preferable for large tubes), I decided to cut the foam down so that it was flush with the box.
Carving and cutting the holes in the foam can be frustrating. Make as few cuts and holes as possible because with each tear the foam becomes more likely to get caught on the Dremel bit and then it will twist and tear the foam block. Also note that the edges of the foam block are likely to get caught by the Dremel bits.
I used sanding bits to make the holes you see below. I started with the cylindrical dremel bit I pictured in the materials section. I went into the block about 0.5″ with the cylindrical drill bit (this made the cleanest looking hole opening) and then I switched to the conical bit which I used to go straight down to the bottom of the tray. Use tubes to test each hole you make to ensure they fit. A snug fitting tube is better than a loose fitting tube.
When the Dremel grabs the foam and twists some areas of the block will be torn.
Another view of the finished block.
All that is left is to attach the box to the vortexer and for that we just need rubber bands. The way the rubber bands rest on the vortexer head and on the box is important, consult the images below.
The first rubber band is hooked under the left side of the vortexer head and hooked over the right side of the box.
The second rubber band is a mirror of the first rubber band.
That is it, the attachment is finished.
If you build a vortexer attachment send me a picture and let me know how it turns out.
Quick Update: The more practice I had with the foam, the better I got making a cleaner/nicer looking piece.
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.
- 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)
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.
This was the gel box I built and used. You can read my instructions for how to build it here.
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.
Sterling silver wire does not work as a cathode or anode for gel electrophoresis.
Our lab was given an old hand-me-down water bath but it didn’t quite meet our needs. Sure, it thaws our samples precisely the way we want it to but the way the lid is set-up is just a pain.
For whatever reason, the designer who made this water bath (back during the Precambrian mind you) decided that a flat lid with no hinge and vent holes was just swell. But to be fair, the designer had the foresight to leave two holes for easy 50mL tube access (see purple lids in the above picture).
Unfortunately for us, we hardly use this water bath for 50 mL tubes – we mainly use it for 15 mL tubes and eppendorfs.
With this in mind I decided to enlarge the vent holes with a dremel so that they could accommodate 15 mL tubes.
I used the yellow tape to hold the lid tightly onto the water bath as I carved the vents larger and to be a visual guide to help me carve as straight as I could.
After cleaning up the huge mess I made I tested the holes with water. This was important because the amount of air in the tubes may have been an issue. You will note the tubes on the right of the image are floating higher than the left because they have more air. But the lid still keeps them in place and adequately submerged.
I also tried it out with just one tube because this is a more common situation. While not perfectly vertical, the lid still keeps the tube upright and in the water.
The whole project took me like 15minutes. Writing about it took longer than actually doing it.
With just two small alterations the water bath became way more user-friendly and cost the lab $0 (unless you count 15 minutes of person-hours).