My (finished) master’s project or this is why I haven’t posted much.

May 10, 2012

I finished my master’s project two weeks ago and it was accepted by the committee, and the school administration!  If anyone feels like checking it out, follow the link below.

Investigation of the role of fragile x mental retardation protein in embryonic neocortex using RNA interference.

What I did, in brief, was attempt to recreate a fragile-x-syndrome-related disease in neural stem cells by reducing fragile x mental retardation protein (FMRP) experimentally in normal mice.  I was able to get some aspects of the disease state to manifest but I did not completely recreate it.

The write-up and final touches really sucked up all my free-time (but was totally worth it).  So now I should have more time for cheapass science.

My project was supported by the California Insitute for Regenerative Medicine (CIRM), California State University Sacramento, and the Noctor Lab at the UC Davis MIND Institute.

 

Buffer preparation.

May 10, 2012

I just made the last of the buffers I needed for my electrophoresis testing. Hopefully next week, after my final, I will start posting the comparison results.  For fun, here are a couple of pictures of my reagent preparation set-up in my apartment.

pH adjustment

pH adjustment

Filtering

Filtering

Human cheek cells with methylene blue

April 16, 2012

I have not done much home lab work on account of finishing up my master’s.  But toady I squeezed in some time for microscopy.  Below is a link to some photos of my cheek cells stained with methylene blue.

 

http://www.flickr.com/photos/codonaug/sets/72157629826384173/

I used “aquarium grade” methylene blue for the staining – obviously because it was cheap.

Build a pipette stand from PVC pipe for $3.47+tax

March 18, 2012

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.

Tools

Saw or PVC cutter – Either work but the PVC cutter is easier to use and costs $10-20.

Marker and Tape measure

Materials

(above) All of the supplies needed for the stand.

PVC Pipe

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.

PVC Fittings

Purchase four T-shaped fittings and six right-angle fittings.

Cost breakdown

T-shaped fittings (4) = $0.80

90deg fittings (6) = $1.02

½” pipe (10 feet) = $1.65

Total = $3.47

Instructions

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.

Ceiling fan centrifuge.

March 11, 2012

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.

 

 

 

 

An auction site success – $155 DNA Hybridiser.

February 26, 2012

My cost: $155 (150= auction, 5=box of rivets, only needed one)

Retail cost (new): $2000-4000

I came across a ‘working’ but damanged DNA hybridiser (which is essentially an incubator with optional rotating racks) on an auction site.  The damaged components, according to the seller, were the two parts of the door’s hinge.  I thought that I could fix it (it is just a hinge afterall).  I figured the worse case scenario was that if I couldn’t repair it, I would end up using masking tape and plastic sheeting across the front to seal it.  As is the case with life, things were a bit more complicated when I got the unit.

(Above) Hinge pre-repair.  I set a 1/4″ rivet into the hole and that allowed the door to pivot perfectly.

(Above) Hinge post-repair

(Above) Post-repair bottom hinge.  I neglected to have a picture of the bottom hinge but suffice to say it was in terrible shape.  The bottom hinge is a triangular piece of metal with a protrusion for a pivot point.  When I received it, the plate was bent two different directions.  I was surprised a lab would abuse equipment so badly!  The metal piece was small enough to fit into my vice and so I smashed it flat and reattached it with great success! The feet on the hybridiser are adjustable by rotating them clockwise or counter clockwise.  One of the feet was tilted something like 20-30 degrees.  The fix to this was to remove the foot, use pliers to bend the screw back into a straight position, and reattached the foot.

 

Going forward from this point things got frustrating.  When I turned the unit on it heated up to maybe two degrees above ambient and I immediately thought I bought a lemon.  I found the manual online and discovered there is a safety reset on the back of the unit that sometimes gets set off when the unit is moved and fortunately this fixed the problem! The second issue was the moving components of the hybridiser (see above circles within plastic brackets) – they didn’t move or make any sound.  On a whim I decided to take off some of the panels to see if maybe I could figure out the issue.  After having every other screw head strip completely and having to use a Dremel to saw off one of the heads I finally got into the unit.  Sure enough the motor that moves the components had fallen apart sometime in the past. Unfortunately I didn’t take pictures of the inner workings because I was tired and frustrated with the screws stripping and mostly I was totally focused on fixing the hybridiser.

 

On the inside there are two interconnected belt systems that feed onto 1 motor.  The belts had come off their rollers and the motor – connected to the frame only by flexible rubber spacers – was barely attached to the frame.  I used some screw spacers (the kind used to attach PC motherboards to cases) and some long screws and I remounted the motor and reattached the belts.

 

Now I have a hybridiser worth substantially more than it cost me.  My best guess is the previous owner put up with the damaged hinge up until the point that the inner-workings of the hybridiser broke and they then tried to sell it.  At any rate, I am glad they were unwilling to explore and repair it. As I have posted before, I recommend checking out old lab equipment on auction sites (i.e. eBay) but always be cautious as most sellers have never been to a laboratory before.

 

Note:  DNA hybridisers circulate heated air with a fan and while they are in essence an incubator, they can’t be used just like a typical incubator because agar plates with bacteria would get cross contaminated easily.  If I ever need to grow something I think I will try putting them into zip-loc bags before putting them into the incubator to avoid this problem.

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.

Conclusions

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.

DIY Tube Holder for Vortex Mixers

January 20, 2012


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.

Guide

Compatibility

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.


Materials

The tools and consumables I list are not absolute. Use your noggin and substitute if necessary.

  • Tools
    • 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
    • Ruler or measuring tape
    • Hand saw
    • Marker
    • Compass (optional)
  • Consumables
    • Foam block
    • plastic box (I used a large pipette tip box that was 4″x5″)
    • Rubber bands


These are the two dremel tips I used to carve the foam.

Instructions

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 block


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.


Before


After


Glamour shot


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.

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.

Update: Sterling silver as an electrode

January 14, 2012

Sterling silver wire does not work as a cathode or anode for gel electrophoresis.