I replaced the alarm battery for each of our -80C freezers – Revco and Sanyo.
Revco battery – Fisher NC1397422
Sanyo battery – Fisher NC1406075
I figured it’d be prudent to collect some Eastern oyster (Crassotrea virginica) to have around the lab.
I used one of the C.virginica oysters that I picked up Taylor on 20171210 for sampling.
Samples were transferred to 1.7mL snap cap tubes, frozen on dry ice, and stored @ -80C in Rack 13, Col 1, Row 5.
I’ve been asked to isolate RNA from some paraffin-embedded Olympia oyster gonad tissue.
Despite some excellent documentation by Laura Spencer (images of tissue layouts in histology cassettes and a corresponding cassette mapping key file), the histology facility seems to have flipped some things around and/or repositioned/split the contents of each cassette. This makes ID-ing the proper tissues tedious and, at times, difficult.
The list of tissues that needs to be processed is listed in this GitHub Issue #648. I’ve also added the list below:
Prior to beginning RNA isolations, I have annotated images of the histology blocks and will be waiting for Laura to confirm that my annotations are correct. I will be posting a link to this notebook entry in the GitHub issue listed above for her to view and wait for her confirmation.
UPDATE 201700707 – Laura has indicated that many of my annotations are incorrect. Katie has gone through and made proper identification: https://github.com/sr320/LabDocs/issues/648#issuecomment-313792588
Additionally, as indicated in the GitHub Issue above, histology block “Oly 14″ does not have a corresponding tissue cassette photo (containing sample NF-10 26). Without the original image, I don’t think I can make an accurate guess on how the tissues are oriented in the resulting two histo blocks (see below).
BLOCKS 14 (unable to annotate at time of posting)
TL;DR – Server didn’t recover after firmware update last night. “Repair” is an option listed in the web interface, but I want to confirm with Synology what will happen if/when I click that button…
The data on Owl is synced here (Google Drive): UW Google Drive
However, not all of Owl was fully synced at the time of this failure, so it seems like a decent amount of data is not accessible. Inaccessible data is mostly from individual user directories.
All high-throughput sequencing is also backed up to Amazon Glacier, so we do have all of that data.
Here is what happened, in chronological order:
Below are pictures of the entire process, for reference.
Ran some basic tests to get an idea of how well (or poorly) the use of dry ice and universal indicator would be for this lesson.
Instant Ocean mix (per mfg’s recs): 0.036g/mL
Universal Indicator (per mfg’s recs): 15μL/mL
Played around a bit with different solution volumes, different dry ice amounts, and different Universal Indicator amounts.
|Indicator Vol (mL)||Solution||Solution Vol (mL)||Dry Ice (g)||Time to Color Change (m)||Notes|
|3||Tap H2O||200||0.5||>5||Doesn’t trigger full color change and not much bubbling (not very exciting)|
|3||Instant Ocean||200||1.5||<0.5||Begins at higher pH than just tap water. Full color change is slower than just tap water, but still too quick for timing.|
|2||1M Na2CO3||200||5||>5||No color change and dry ice fully sublimated.|
|2||1M Tris Base||200||5||>5||No color change and dry ice fully sublimated.|
|2||Tap H2O + 20 drops 1M NaOH||200||5||2.75||~Same color as Na2CO3 and Tris Base solutions to begin. Dry ice gone after ~5m and final pH color is ~6.0.|
Some pictures (to add some zest to this entry):
Our lab scale managed to become horribly misused and left in an unacceptable condition, so I cleaned it. Before and after pics below.
We’ve had a disabled cart sitting in lab for months that has locked up wheels and all of the bolts attaching the wheels to the cart are so rusted, the bolts and nuts cannot be separated by normal means (believe me, I’ve tried numerous methods over the last few months to no avail). Since we’re planning a significant lab cleanup next week, having this cart repaired will be good, so that it’s not sitting upside down in the lab any more AND fixing it will provide us with a second cart for cleanup day!
In any case, I ended up having to drill out the rusted bolts/nuts on three of the four wheels. After that, I put on the new wheel hardware and it’s good as new!
Some before and after pics below.
For some reason, sometimes when we use the SPOT camera for capturing images on the microscope, we see this:
For some reason, the default filter setting has been set to “Green” instead of “RGB.”
Change the setting by following the next three steps:
1. Click on the Image Settings button:
2. Click on the Filter Color drop-down menu:
3. Select the “RGB” option:
We’re back to the normal view! Granted, the image appears a bit washed out (and a bit on the yellow side), but this can be taken care of with a white balance adjustment.
All washes/rinses were performed in cylindrical glass slide incubators at room temp (30mL):
Images were captured using Nikon BR Essentials.
The same section of each slide (within an accession number set) was captured at 4x, 10x, and 20x magnifications for comparisons. Auto white balance adjustment was the only image manipulation performed. All images (see Results below) are as they were captured by the software.
Quick summary: Both probes appear to be functional! With that being the case, I will proceed to run ISH on black abalone samples (these test ISHs were with red abalone) for the proper assessment of RLOv localization.
All images are here (Dropbox): 20151204_ISH_RLOv
Tryptic images of 10x magnifications are presented below showing the H&E staining, negative control and RLOv probe.
ISH staining is expected to appear as dark brown staining.
H&E – RLO inclusions are seen as the deep purple oblong structures.
Negative Control – RLO inclusions exhibit no staining and appear as oblong empty regions. These regions also no have any apparent cell wall/membrane around them. This is in contrast to the two other accession groups (08:1-12 & 08:1-15).
RLOv tail fiber – Staining is noticeable surrounding the RLO inclusion locations, but not within the inclusions. The staining is similar to the 08:1-12 negative control. So, it’s difficult to say if the staining in this sample is binding to its intended target (RLOv tail fiber) or if the difference seen is simply due to the particular section of this tissue.
RLOv membrane gene 1 – Not shown due to this region of tissue not being present on the slide.
08:1-12 (RLO CLASSIC)
H&E – RLO inclusions are seen as the deep purple oblong structures.
Negative Control – RLO inclusions appear similar to air bubbles, with no staining within them.
RLOv probes – Both probes stain within the RLO inclusions.
08:1-15 (RLO STIPPLED)
H&E – RLO inclusions are seen as light purple bulbous structures.
Negative Control – RLO inclusions are actually stained brown and are very noticeable. This is not expected.
RLOv membrane gene 1 - The staining is in the same locations as the negative control RLO inclusions. Intensity-wise, the staining seen from this probe is not much different than the negative control. However, the way the staining appears within the inclusions is different than the negative control. Not sure if this indicative of the probe working or if the different appearance is due to difference in tissue sections.
RLOv tail fiber – The staining is in the same locations as the negative control RLO inclusions. However, the intensity of the staining with the RLOv tail fiber probe is a much deeper brown, suggesting that the probe is binding within the RLO inclusions.
To test out the viability of these RLOv ISH probes (from 20151109) and not waste black abalone slides if this doesn’t work, I selected three unstained red abalone post-esophagus sections:
RLO – NO PHAGE
RLO STIPPLED – NO PHAGE
All slides were processed in a single, horizontal glass slide incubator (200mL), unless otherwise noted.
All steps were conducted at room temperature (RT), unless otherwise noted.
DEPARAFFINIZATION & REHYDRATION
NOTE: RLOv Membrane Gene 2 probe was ruined because boiling water got into the tube during denaturation. This didn’t happen to any of the other tubes that were all boiled at the same time. Not sure what happened. However, this may have worked out OK because I did not pull enough slides to accomodate the negative control probes. So, now that I’m not able to test three probes, I can use the negative control probes!