Tag Archives: pics

Tissue Sampling – Crassostrea virginica Tissues for Archiving

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.


  • Upper mantle (avoided area that was near gonad/white-ish)
  • Ctenidia
  • Lower mantle
  • Muscle
  • Gonad

Samples were transferred to 1.7mL snap cap tubes, frozen on dry ice, and stored @ -80C in Rack 13, Col 1, Row 5.


Sample Annotation – Olympia oyster histology blocks (from Laura Spencer)

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:

NF-10 22

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)















Troubleshooting – Synology NAS (Owl) Down After Update

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:


  1. Updated DSM via web interface in “Update & Restore”. Did NOT perform manual install.
  2. System became inaccessible via web interface and Synology Assistant.
  3. The physical unit showed blue, flashing power light and green flashing LAN1 light.
  4. No other lights were illuminated (this includes no lights for any of the drive bays).
  5. The attached expansion unit (DX513) showed steady blue power light, steady green lights on all drive bays, and steady green eSATA light.
  6. I powered down both units via the DS1812+ power button.
  7. I turned on the both units via the DS1812+ power button.
  8. Both units returned to their previous status and were still inaccessible via the web interface and Synology Assistant.
  9. I powered down both units via the DS1812+ power button.
  10. I removed all drives from both units.
  11. I turned on the both units via the DS1812+ power button.
  12. I connected to the DS1812+ via Synology Assistant. A message indicated “No Hard Disk Found on 1812+”.
  13. I powered down both units via the DS1812+ power button.
  14. I added a single HDD to the DS1812+.
  15. I turned on the both units via the DS1812+ power button.
  16. I connected to the DS1812+ via Synology Assistant. I was prompted to install the latest DSM. I followed the steps and created a new admin account. Now the system shows 7 drives in the DS1812+ with a message: “System Partition Failed; Healthy”. Disk 1 shows a “Normal” status; this is the disk that I used to re-install DSM in Step 14. Additionally, the system shows one unused disk in the DX513.
  17. I powered down both units via the web interface.
  18. I removed Disk 1 from DS1812+.
  19. I turned on the both units via the DS1812+ power button.
  20. The DS1812+ returns to its initial state as described in Step 3.
  21. I powered down both units via the DS1812+ power button.
  22. I returned Disk 1 to its bay.
  23. I turned on the both units via the DS1812+ power button.
  24. There’s an option to “Repair” the Volume, but I’m not comfortable doing so until I discuss the in/outs of this with Synology. Submitted a tech support ticket with Synology.

Below are pictures of the entire process, for reference.


Server status when I arrived to lab this morning.


Pulled the HDDs from both units, in an attempt to be able to connect via Synology Assistant.


Units w/o HDDs.


No HDDs in units made the server detectable via Synology Assistant, but it indicates “Not installed” in the “Status” column…


Successfully connected, but the DS1812+ indicates no HDDs installed.



Added a single HDD back to the DS1812+. Notice, the drive light is green and the “Status” light is amber. This is actually an improvement over what I saw when I arrived.


Added back a single HDD to the DS1812+ and now have this setup menu.


I’m prompted to install the Synology DSM.


Installing DSM. This “Formatting system partition” message might be related to the eventual error message that I see (“System Partition Failed”) after this is all set up…









Prompted to create an admin account. This does not bode well, since this is behaving like a brand new installation (i.e. no record of the previous configuration, users, etc.).


Continuing set up.


All set up…



Added all the HDDs back and detected via Synology Assistant.


This shows that there are no other users – i.e. previous configuration is not detected.


After putting all the HDDs back in, got this message after logging in.


Looking at the Storage info in DSM; seems bad.



Physically, the drives all look fine (green lights on all drive bays), despite the indication in the DSM about “System Partition Failed” for all of them (except Disk 1). The expansion unit’s bay lights are actually all green, but were actively being read at the time of picture (i.e. flashing) so the image didn’t capture all of them being green. Amber light on expansion unit reflects what was seen in the DSM – the middle drive is “Not initialized”. Note, the drive is actually inserted, but the handle has been released.


This is how I left the system. Notice that after rebooting, the expansion unit no longer shows that “Not initialized” message for Disk 3. Instead, Disk 3 is now detected as installed, but not used…



Curriculum Testing – Viability of Using Dry Ice to Alter pH

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 1.5 <0.5
3 Tap H2O 200 0.5 >5 Doesn’t trigger full color change and not much bubbling (not very exciting)
5 Tap H2O 1000 12 <1
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.



  • Universal Indicator amount doesn’t have an effect. It’s solely needed for ease-of-viewing color changes. Use whatever volume is desired to facilitate easy observations of color changes.
  • Larger solution volumes should be used in order to slow the rate of pH change, so that it’s easier to see differences in rates of change between different solutions.
  • 1M solutions of Na2CO3 and Tris Base have too much buffering capacity and will not exhibit a decrease in pH (i.e. color change) from simply using dry ice. May want to try out different dilutions.
  • Use of water + NaOH to match starting color of Na2CO3 and/or Tris Base is a good way to illustrate differences in buffering capacity to students.
  • Overall, dry ice will work as a tool to demonstrate effect(s) of CO2 on pH of solutions!

Some pictures (to add some zest to this entry):





Cart Repair


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.







Troubleshooting – Green Screen Using SPOT Camera

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.


In-situ Hybridization (ISH) – RLOv Membrane Gene 1, Tail Fiber Gene: Day 3

All washes/rinses were performed in cylindrical glass slide incubators at room temp (30mL):

  • Slides were briefly rinsed in dH2O three times.
  • Slides were counter stained with 0.05% aqueous Bismark Brown Y for 3mins.
  • Slides were briefly rinsed in dH2O, then 70% EtOH, then 100% EtOH.
  • Slides were air-dried in the fume hood.
  • Coverslips were added to each slide with three drops of Permount.
  • Permount was allowed to dry O/N at RT.

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.

08:1-7 (RLOv)

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.


In-situ Hybridization (ISH) – RLOv Membrane Genes 1 & 2, Tail Fiber Gene: Day 1

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:


  • 08:1-12-2
  • 08:1-12-3
  • 08:1-12-4


  • 08:1-7-7
  • 08:1-7-8
  • 08:1-7-9


  • 08:1-15-7
  • 08:1-15-8
  • 08:1-15-9

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.


  • All slides were deparaffinized with three changes of xylene (SafeClear II; Fisher) for 10mins each.
  • Slides were hydrated with a graded ethanol series (100%, 100%, 80%, 70%, 50%) for 3mins each.
  • Slides were rinsed with molecular grade H2O.


  • Tissue sections were equilibrated in Tris Buffer (0.2M Tris-HCl, 2.0mM CaCl, pH = 7.2) for 5mins.
  • Tissues were permeabilized for 1.5hrs in preheated 50ug/mL Proteinase K (Qiagen) in Tris Buffer @ 56C.
  • Slides were rinsed with 1x PBS three times, 10mins each.
  • Slides were incubated 30mins in 30mL Prehybridization Buffer (50% deionized formamide, 4x SSC) @ 53C in a cylindrical glass slide incubator due to limited volume of deionized formamide available:

  • Prepared probes by boiling 3mins and immediately incubating in ice water bath for 30mins.
  • Slides were rinsed with 2x SSC and air dried for 5mins.
  • Probes were diluted 1:300 in 1000uL of Prehybridization Buffer. All three negative control probes (indicated by “-C” in subsequent labeling) were combined into a single dilution.

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!


  • 300uL of probe solutions and cover slip were added to the following slides:

  • The three groups of slides were placed into separate slide cases and a 1mL of Prehybridization Buffer was added to each case (to maintain high humidity during incubation).
  • The cases were incubated on their sides O/N @ 53C.