Notice for this trial there is no sample with no seeds. Here’s some nomenclature for you:
DOI: 10.15200/winn.142802.20701 provided by The Winnower, a DIY scholarly publishing platform
At 11:30am MST I removed the samples from the refrigerator and sealed them with vacuum grease (see Experiment Product Page). This step is always so time consuming and I would really like to figure out a better way to do this. I basically use a small cylindrical tool (it’s a plastic scraper tool for plating cells) and just smear the grease all over the interface between the top and bottom of the analyslide.
After applying the sealant, I perturb the dish enough so that most of the seeds end up in the middle of the sample. This involves me shaking the “slide” around until I feel I have a good distribution in the viewable area of the dish.
I’ll make a video of my methods and put it on BenchFly when I get some extra hands in the lab. Hopefully that will clear things up.
Pictures of Day 0 seeds to come in about 30 minutes.
DOI: 10.15200/winn.142802.20695 provided by The Winnower, a DIY scholarly publishing platform
It’s time to track seed germination, and eventually growth rate (once we get some automation software going), and that means another Repeating Crumley style experiment. I really don’t like calling it that because I’m no longer repeating Crumley, but the fact remains the experimental setup is identical so I’m still going to refer to it as the RC experiments. This time I’m doing only DI water (our commercial waters compared against building purified waters), hence naming it RC6 DI Water and categorizing it RCW. Ok with that said lets get into the setup:
Tomorrow I will remove the samples, seal them with vaccuum grease, and then take day 0 pictures.
DOI: 10.15200/winn.142802.20656 provided by The Winnower, a DIY scholarly publishing platform
I removed the seeds from the fridge and decided to keep them in a styrofoam cuvette case. This should keep light and temperature pretty stable in the lab. I then put tape over like groups of seeds so I can easily remove the cuvettes and set them up for picture taking.
Believe it or not, that little trick saves me a load of time.
Wow, that sucked. I had to crop, rename, and label 42 pictures for this notebook. Stupid open science movement. Why do I strive to be a good scientist? Why can’t I just let myself be lazy and allow myself to be a bad scientist? Who cares about accurate data and organization?
I do unfortunately.
Anyways here are the Day 0 pictures. Luckily I don’t have to do this every day, that would suck. From now on I may label the water types as follows:
Since the last time I did this, I have learned a ton about proper exposure with a camera. And I actually know what aperture, ISO, and shutter speed are for. I’ll do a post that explains all that but in the mean time here are the settings I used for these pictures and those in the future:
I’m sure these settings are saved with the images so if you download the pics you’ll see the camera settings in photo software.
After 4 trials and talking with Steve, there is no way to ensure that the results we are getting have anything to do with deuterium content. We can’t tell at all that our DI water is just not the same kind of “pure” that the DDW is. So I bought deionized water from Sigma (same people who make our DDW and D2O) to compare with our water and to compare to the DDW results to determine if the root hairs grow based on water purity or if it has something to do with deuterium content.
Here is the setup of the experiment:
Today’s protocol was really easy. I used my macro cuvettes (see Experiment Product Page) and set up 4 cuvettes for each water type giving me a total of 24 cuvettes. I did this so I could set up samples for one seed type at a time, and because I only had enough racks for this many cuvettes. I also prepared the D2O/DDW mixture (29.7mL of DDW, 0.3mL of D2O).
I poured 5-7 seeds in each macro cuvette until all the setup cuvettes had seeds in them. Then I added 3mL of water to the samples (again making 4 samples per water type). Then I sealed the cuvettes with PE caps.
I repeated this setup for the next seed type (24 cuvettes, 5-7 seeds per sample, 4 samples per water type, with 3mL of water per sample) and sealed and labeled all cuvettes.
I placed the cuvettes in the 4C fridge to synchronize the growth among all the samples. Tomorrow I’ll remove the seeds and take Day 1 pictures.
Steve and I are trying to come up with ways to quantitatively measure differences between seeds grown in ddw vs di water. Via the slideshows it is obvious that length is one such observable. Is mass another?
Perhaps, but it’s not easy to extract a number from such small samples. I gave it a try nonetheless. 2 days ago I opened all my sample chambers and allowed the water from each sample to evaporate and the plants to dry out. I used seeds from both the Repeating Crumley experiments and the DDW Effects experiments.
I realized quickly that I can’t weigh the seeds in the analyslides because the vacuum grease that I use to seal the chambers isn’t uniform on any of the dishes. That means that I can’t take a baseline weight of one slide and use that as a zero for all the dishes. I then threw out the seeds without realizing I could try and scoop out the plants and weight them on weigh paper or something. Next time.
The seeds in the DDW Experiments are contained in macro cuvettes. I took four empty (and new) cuvettes and weighed one and zeroed the scale. Then I weighed each of the others and found that two others were identical in mass and the fourth cuvette had a mass that was 0.0004g less than the original cuvette. Not bad.
I then began to weigh each of the cuvettes that contained dried plants.
BAD IDEA!
It seems the tobacco seedlings are so slight that every sample I measured was around 0.0003g lighter than the blanked cuvetter! There are on average 5 seedlings in each sample. So that means that 5 seedlings have less mass combined then the error associated with inequalities in the mass of different cuvettes. Interesting but sucky.
It is pretty hard to remove the seedlings from the cuvettes without destroying them so I’ll have to return to the drawing board. I think I will setup another RC experiment and scrape the seedlings out of there (weigh the seeds before adding them to the batch) at the conclusion of the data acquisition and weigh them via some other means (again probably on weigh paper).
Ideas?
Note: I tried to do this with the arabidopsis but realized that those seeds are definitely too light to be noticed by our supposedly super sensitive balance.
Here are some more notes from today using the latest batch of slideshows. Some interesting observations were made.
DOI: 10.15200/winn.142801.10410 provided by The Winnower, a DIY scholarly publishing platform
I’m not sure if these are useful but in the spirit of openness here are Steve’s notes during the blind seed analysis experiment. The slideshow analysis is first then the day 11 picture analysis and on image 2 is day 4 notes with a graph that Steve draws a lot that I have yet to fully grasp.
DOI: 10.15200/winn.142801.10395 provided by The Winnower, a DIY scholarly publishing platform