Tag Archives: equipment

Molecular Biology, sequences, Shotgun DNA Mapping

Anchor DNA Sequences

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See here for the background behind everything contained below. Note: For now I’m going to link sequences from OWW here. I was going to put the entire sequence, but that would make this page sorta sloppy and it could get lost. So I’m going to make a page that contains all the sequences necessary for Shotgun DNA Mapping.

  • pRL574– This is a non-commercial plasmid provided by Robert Landick. We have a very small supply so I will have to do some cloning to make an infinite supply!
    • primers – according to notes that I have on OWW and Google Docs I’ve had success with F834-dig as the forward primer (and might be the only primer I have in the lab), R2008 and R1985 as the reverse primers. The difference between the two reverse primers is the length of the PCR sequence, which turns out to be a difference of 23bp.
  • pALS– designed by me, purchased and built by DNA 2.0. I’ should have enough for a few PCR reactions, but I may need to clone to replenish my stocks.
    • primers – primer R4500 would bind in two places on the plasmid so I made R4000 to fix this issue. I’ll have to check my paperwork to see which primer has the dig. I think it is supposed to be on the reverse end, but I can’t be sure.
KochLab Stuff

Using cuvettes in the nanodrop

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I did a mini-study this morning to find out what the minimum volume needed is to get an accurate reading in the nanodrop. I have 2 different cuvettes (semi-micro and micro) and I wanted to impact the cultures the least so I would like to use the micro cuvettes.

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From left: 400ul in semi-micro cuvette, 200ul in micro cuvette, 500ul in semi-micro cuvette

I used 5 semi-micro cuvettes and 2 micro cuvettes. I put increments of 100ul starting at 100ul in each cuvette (100 -> 500ul in the semi-micro and 100 and 200ul in the micro cuvette). At and above 400ul the nanodrop was able to effectively and consistently read the absorbance of the semi-micro cuvette (verified because the readings for 400 and 500ul were identical). For the micro cuvette, I looked at the profile (image above) and saw by eye that the height of the meniscus of the liquid media was roughly equivalent to the height in the semi-micro cuvette with 500ul. So I put this in the nanodrop and got an equivalent absorbance reading.

So in summary:

  • For semi-micro cuvettes (and the Thermo Nanodrop 2000c), volumes of at least 400ul or more are sufficient for consistent readings.
  • For micro cuvettes, volumes of 200ul are sufficient for consistent readings.

The End.

KochLab Stuff

Experiments’ Product Page Updated

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I’ve been slacking with keeping up with the equipment that I use in the lab, but I have updated the product page and it should be all up to date with the latest string of experiments. Let me know if you notice I’m missing something.

KochLab Stuff

Fixing the incubator

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Warning: My phone takes some bewilderingly bad pictures, and unfortunately all images in this post were taken with said phone. I have a Droid Bionic, so if you are on the fence about phones and want a good camera, this is not the phone for you.

Onto the protocol…

So I noticed that our incubator/shaker (Innova 4300) wouldn’t shake. The display would read “LID” indicating that the lid was open even though it was closed. Usually I just lean on the top and presto! But this time no amount of force would register as closed. So I had to figure out what part of the shaker registers the lid as closed. Here is what I noticed:

It seems there are two sensors that must make contact between the base of the machine and the lid. Here are pictures of the pieces and their contact points:

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sensor on lid on right side.
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sensor contact on base on right side
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sensor on base on left side
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sensor contact on lid on left side

So no matter how much pressure I put on the lid, one of these contacts was not making contact. I checked and it was the one on the right side:

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see the space?

And if you see in that picture, there are screws on each side of that cylinder. So I unscrewed each one and was able to pull the plate off and push the sensor down some. I put the plate and the screws back on and put the lid down. Then the machine seemed to work.

The caveat is that the sensor thing right now isn’t making contact between the lid and the base, and the machine still works, so I’m thinking that even though what I did was mighty, it really had no effect. But my question is, if that thing is not a sensor, then why are there wires that attach to it? (I didn’t mention that before, but that is why I assumed this thing was a sensor.)

Anyways, the shaker gives me no issues now. So for your viewing pleasure here are some other pictures I took to that I sent from my phone in case I needed them, don’t really, but want to show you anyway:

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the inside of our shaker
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blurry and dark image of the left sensor making contact with the lid
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another blurry image of the same sensor.

Hopefully this post is useful to someone, or someone can see that I clearly have no idea what I’m doing and can explain to me why the lid would fail to register as closed and then I adjust a seemingly meaningless piece and now it registers as closed, even though it isn’t touching anything.

DDW5, Water Type Effects on Organism Growth

DDW5: Taking pictures and manipulating them in bulk

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A lot of this project is managing how I take pictures of the plants. I’ve talked a lot about my setup and I’ve talked some about how I take pictures and the software I use. Well I’ve had to make some changes to the process and now I’m letting you know so I can continue to be a good open notebook scientist.

camera setup

Above is a picture of my current setup. I noticed that some seedlings will float in the sample and I couldn’t fit the entire cell in the frame of the camera so I had to find an alternative to my previous method. My solution? Rotate the camera! Simple!

I just found another optical post (4″ in this case) and a 90 degrees post holder. I put the two together and used the post holder from the previous setup (2″ long) to get the current setup. All equipment used in the photo above can be purchased from ThorLabs (and if you buy a bunch you get some lab snacks with your order!).

Also since last time, I added a black backdrop (the one I’m using is a big piece of thick paper). The black provides excellent contrast to the white root hairs.

Finally I’d like to talk about the software that I use. In the previous trial of the DDW experiment (Category: DDW4) I used some software called JPEGCrops to crop the photos in bulk, and I used a program named Rename Master (both open source software) to rename the photos in bulk for organizational purposes.

With this new camera setup, rotating the frame actually allows me to minimize the amount of the other samples in the frame. But I do need to rotate the images in bulk. Windows actually does something right in this regard. If you select a bunch of images and right click, there is an option to rotate the images either clockwise or counter-clockwise in bulk. Using this handy feature saved me a ton of time. Then I can use Rename Master to rename the images in bulk as well.

And that is my whole process, well up until I upload them to my notebook. But from there you can add captions and do some other minor editing features (that I almost never use). Hope this helps someone somehow.

KochLab Stuff

-80 Freezer Down

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I can’t remember if I posted this yesterday or not (and I’m too lazy to look it up myself) but yesterday when I got into lab, the -80C freezer malfunctioned and there was an alarm alerting me to a “high system failure.” After calling VWR and receiving next to no help, I unplugged the machine and plugged it back it after waiting a few minutes. Problem solved. The freezer is working like normal again.
I have no idea what caused the malfunction, but everything in the freezer was thawed. Koch and I filled some styrofoam containers with LN2 (liquid nitrogen) and put everything in the freezer into the LN2. We then stuck the containers in the freezer to add an extra layer of insulation. I’ll remove the samples tomorrow and place them back in their respective boxes and back in the freezer like normal.

KochLab Stuff

Reusing Drierite

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We were running low on Drierite (we use the indicating variety) in the lab, which is a desiccant, and so I ordered some more. Fairly straight forward.

In my search for the stuff we currently use I came across this page talking about regenerating the dessicant. This would have been awesome to know a few years ago, as someone has been throwing out old dessicant. In the case of our indicating drierite, the chips turn from blue to pink, so the pink chips end up discarded.

I decided to order more because I didn’t want to deal with leaving chemicals that are in the desiccator in there for an extended period of time without desiccating material.

Anyways if you (like me) are too lazy to click the link that talks about desiccant regeneration then read below for the drierite protocol:

For the regeneration of Indicating DRIERITE and small lots of Regular DRIERITE , the granules may be spread in layers one granule deep and heated for 1 hour at 210° C or 425° F. The regenerated material should be placed in the the original glass or metal container and sealed while hot. The color of the Indicating DRIERITE may become less distinct on successive regenerations due to the migration of the indicator into the interior of the granule and sublimation of the indicator.

Note: I’m creating a new notebook category for stuff like this, that has little to do with my research but could be vital one day for future students in the KochLab. I am cleverly calling it “KochLab Stuff.”

Water Type Effects on Organism Growth

Macro Photography

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Koch asked me to look into getting a macro lens for the DDW experiments. So today I started and it dawned on me… how am I going to differentiate a good lens from a bad one, besides cost? I started looking at specs and I realized I know almost nothing about technical specs of camera systems. And (un)surprisingly macro photography is pretty finicky when it comes to how the camera reacts to the environment.

So now I write to blab about what I’ve learned and what I should look for in a good lens:

  • Aperture: In my experience because I’m working in such close proximity the focal plane depth is rather shallow. By this I mean that there isn’t very much in focus unless the subject(s) are all at the same distance. In the case of my seeds, as things grow, stuff comes in and out of focus depending on it’s location. This is why I push all the seeds to the front of the sample chamber so they are all the same distance from the camera. I think this means that for macro photography the aperture is relatively large indicated by a smaller F/(-number). So when it comes to buying a lens from what I’ve read I want to get a lens that has a low F/(-number) where the (-number) is a number that is as low as possible. On the lens that we are using now, this number is something like 5.6
  • Shutter Speed: From what I can tell, typically the shutter should be set to slow. I’m not fully understanding why but it just is. This means that you need to have an abundance of light to ensure a good quality photo. Flash is not an option (if it’s on your camera) because the light won’t be able to bounce off your subject because of the proximity of the camera to the subject. I don’t think I’ve been using a slow speed for my pictures, but I suppose that will change when I get a macro lens.
  • Focal length: Obviously this is important. The more you can zoom the closer and larger you object will appear. I’m limited in this regard now because the lens can only zoom so much and I’m using a magnification lens that actually gives me less focal play. According to the internet, using an extension column (basically moving your lens further from the ccd) gives you more magnification. And I’ve quickly verified this.

Surprisingly I’ve been doing extensive research on this and only now have I just found the wikipedia article on this topic. The article is very informative (as always) and more useful than what I’ve written here.

Anyways, with my advanced knowledge that I now have, I’m looking into this lens: Sigma 50mm f/2.8 EX DG Macro Lens. It seems to have everything I need. Apparently having a zoom-lens-like-macro-lens (confusing) basically just gives you the opportunity to take pictures from a distance.  I’m choosing the one above because I don’t need distance. I’m actually working in pretty close quarters so I would prefer to keep the camera as close as possible.