Kaitlyn’s notebook: Quantifying geoduck histology

These are possible measurements that can be taken on the geoduck histology slides based on tools I’ve found:

Males:

  • pixel classification: acini vs connective tissue
  • average acinus size (measure widest portion of 25 acini/individual)

Females:

  • oocyte length
    • measure only round eggs with visible nucleus
  • average follicle size (measure widest potion of 25 follicles/individual)

Unfortunately I can’t use a pixel classifier for the females because the oocytes and connective tissue stain is very similar in color and there are significant white space between the tissue and in the nucleus. 

I have been using the Padilla-Gamiño micrscope (Nikon Eclipse Ni-U with the Nikon Elements BR [basic research] program). I have been exploring the program to find features useful to our histology slides. I can get the % area using the pixel classifier.

 

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Acini are deeply stained. Using the pixel classifier I can make these pixels red. The percent area for each phase (pixel stain) is shown to the right.

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Connective tissue is shown with the green pixels.

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A possible third phase in blue…

Alternatively, I can do just two phases-

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this is using the manual method (the bayes method has you pick sample pixels for each phase rather than showing a graph for the wavelengths).

However, the inability to remove the phase overlapping on the image makes it very difficult to find another spot to measure on the slide, and I surmise that at least 3-4 measurements throughout the slide should be taken to accurately determine the % area of the acini.

Other possible option are thresholding in Fiji (Fiji is just ImageJ [but updated]), color deconvolution in ImageJ (but I have not been able to find a way for the program to give me % area), or pixel classifier qith Qupath (a new addition to the program that still has soem kinks and I haven’t been able to use yet).

It is easy to take measurments and images with a scale burned onto the image so measuring oocytes, collicles and acini using either this program or ImageJ is easy! I am working on females first, and during any down time, i.e., someone else needs to use the scope, I am trying to learn how to sue Fiji, Qupath, or the Element BR program. 

Sam’s Notebook: Sample Submission – Tanner Crab Infected vs Uninfected RNAseq

After reviewing our options for sample pooling, we decided to do a comparison of Infected vs. Uninfected crabs.

I pooled equal amounts of RNA from each individual in a given set (e.g. Day 9 Infected) to achieve 1000ng. Samples were concentrated using the Friedman Lab SpeedVac to target the “required” concentration specified by the sequencing facility (60ng/uL – UW’s Northwest Genomics Center). I put “required” in quotes because, it turns out, that the amount of total RNA and the concentration are not actually required! Here’s an email exchange when I asked if there was any wiggle room:

Hi Sam,

Thank you for your questions. 1000ng is our “ideal” amount with a built-in 2nd run along the way just in case. We have received some samples with their volume too low to work with (regardless of concentration), but we can take how much ever you can give us.

I also tried to get them to commit to an absolute minimum for input RNA, but the correspondent just kind of talked around the question. Regardless, I submitted the sample manifest (see below) and they accepted it with samples below the “required” concentration and minimum amount of input RNA…

Samples will be sequenced on the NovaSeq S2 200 cycle flow cell with ~50M reads per sample.

Here’s the manifest sample sheet I submitted:

UDF/Investigator Sample Name UDF/Replicate Sample Id UDF/Organism UDF/Gender UDF/Race UDF/Concentration (ng\/uL) UDF/Total Volume (uL)
D9_infected Chionoecetes bairdi 75 10
D9_uninfected Chionoecetes bairdi 81 10
D12_infected Chionoecetes bairdi 57 10
D12_uninfected Chionoecetes bairdi 74 10
D26_infected Chionoecetes bairdi 59 10
D26_uninfected Chionoecetes bairdi 70 10

from Sam’s Notebook http://bit.ly/2Jxr74S
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