Oly Genome: Redundans run finished.

Redundans finished over the weekend, but the results were a little… odd.

Stats for the Illumina only Platanus and the completed Redundans run are below. We increase the N50 from 105 to 251, but in doing so lose 75% of the total number of bases, and 80% of the contigs from the initial assembly. Something’s wonky in the pipeline.

I did some more research and I realized I may have made a faulty assumption that Redundans would take the final output of the Platanus pipeline, a gap closed scaffold assembly. It looks like it actually wants the initial assembled contigs.

I’ve started a run again supplying the contig assembly, and we’ll see if that yields better results! I’m a little skeptical though because the contig assembly only has 16mm base pairs, which still seems awfully low.

Redundans Output can be found: here/

The final assembly is: scaffolds.reduced.fa

I’ve also installed Falcon on Hyak. Falcon is de novo assembler for PacBio only reads.

Some notes on the assembly of Falcon.

  • Don’t load the anaconda2_4.3.1 module. Falcon is nice scripts that download and install things inline, so you don’t have the ability to specify where things are installed (user directory vs anaconda module directory).
  • Everything has to be done on a build node, rather than an interactive node, since the scripts both download and install packages
  • Install networkx v 1.10 prior to starting using easy_install, specifying an install directory via something like easy_install --install-dir /gscratch/srlab/programs/networkx networkx==1.10

Katie’s Notebook: First day at Manchester

I spent my first day at Manchester helping Laura screen and count her new larvae. After an initial walk through, Laura taught me how to screen, sample and count the contents of each bucket.

Sampling:
We started off sampling larvae that had been screened through 160um, 140um, 120um, and 100um with the goal of collecting growth rate data. Laura did the screening and I did the sampling and counting.
-I was given tripours containing the larvae from each size screening (4 tripours/bucket) that I filled to 200mL
-I took triplicate 0.5mL samples (dense amounts of larvae) or triplicate 1.0mL samples (less dense).
-Fixed wells with Lugols and recorded counts.

After a couple buckets we changed tactics and Laura decided to just screen to see if larvae were above or below a designated size. Since they all are pretty small right now we just screened the buckets through 100um screens to get total counts. Almost no larvae in the initial buckets were over 160um.

Screening:
-Screened new larvae through 224um screen onto sorting table filled with FSW.
-Placed 2x100um screens under sorting table- top one clean and vortexed.
-Drained sorting table onto 100um screen.
-Collected larvae in tripour.
-Labeled with tape, left on bench while I collected the other groups.
-Vortexed sorting table & screens between groups.
-Sampled triplicates from each group, but did not fix with Lugols because we wanted to make sure we were able to distinguish between alive and dead larvae!

Yaamini’s Notebook: DNR Sample Preparation Round 2

Slice and dice

Laura and I are going to start our proteomics pipeline with a second set of samples tomorrow, so today Grace and I prepped samples. Like last time, we had to split the samples we wanted in half.

Identified samples for protein extraction

Table 1. Samples for protein extraction, from this lab notebook entry. Originally I wanted to use O111 for extractions but I could not find it in the sample box. I used O117 instead.

Site Condition 1 2 3
PG B O25 O83 O54
PG E O31 O71 N/A
FB B O36 O70 N/A
FB E O64 O46 O32
WB B O129 O126 O135
WB E O140 O145 O147
SK B O117 O120 N/A
SK E O103 O101 N/A
CI B O11 O13 O16
CI E O01 O08 N/A

Labelled snaptop centrifuge tubes

  • Tubes for the sample I would extract proteins from (“1/2”)
  • Tubes after sonication (“11 µL”)

Cut samples in half

The reason for cutting each sample in half is to use one half for protein extractions, and another half for corresponding DNA extractions later on. Our protocol:

  • Obtained dry ice from Biochemistry J Wing
  • Placed sample centrifuge tubes (original vials and vials labelled “prot”) in dry ice to keep them from thawing
  • Created a 10% bleach solution (4 mL Clorox bleach in 40 mL nanopure water) to sanitize equipment
  • Obtained tweezers, weigh boats and razor blades
  • Placed weigh boat in dry ice to keep chill
  • Using tweezers, removed gill tissue from centrifuge tube and placed in weigh boat
  • Cut gill tissue in half with razor blade
  • Disposed of razor blade
  • Placed one gill tissue half back in original tube, the other in the tube labelled “1/2”
  • Put both tubes back in dry ice to stay cold
  • Tweezers cleaned by dipping in 10% bleach solution, and then rinsing in nanopure water
  • Repeated for all samples

When cutting the samples, I realized we forgot the step where we rinsed tweezers with nanopure AFTER we cleaned them with bleach! :open_mouth: I called Emma, and she said to dispose of those samples since they were possibly contaminated by bleach. Additionally, she said we should replace the bleach with ethanol, since trace elements of bleach in our samples would be problematic. Since I was dealing with all of my bare condition samples first, I replaced them with different samples.

Table 2. Revised samples for protein extraction. I wanted to use O127 for extractions but again, I couldn’t find it in the sample box. I replaced with O124.

Site Condition 1 2 3
PG B O26 O60 O90
PG E O31 O71 N/A
FB B O66 O39 N/A
FB E O64 O46 O32
WB B O128 O124 O137
WB E O140 O145 O147
SK B O118 O100 N/A
SK E O103 O101 N/A
CI B O12 O14 O17
CI E O01 O08 N/A

The revised protocol:

  • Using tweezers, removed gill tissue from centrifuge tube and placed in weigh boat
  • Cut gill tissue in half with razor blade
  • Disposed of razor blade
  • Placed one gill tissue half back in original tube, the other in the tube labelled “1/2”
  • Put both tubes back in dry ice to stay cold
  • Tweezers cleaned by dipping in ethanol, and then rinsing in nanopure water
  • Repeated for all samples

I noticed that the amount of gill tissue in the original DNR sample vials varied between samples. Some had large chunks of tissue that were easily split, some barely had any tissue. I ran into something similar last time, so I don’t think it’s going to be much of an issue.

Made 50 mM NH4HCO3 + 6M urea solution

Protocol used can be found here, or viewed below. This solution must be used no later than 24 hours after it is made, or it is no longer viable.

  • Measured 10 mL of nanopure water in a graduated cylinder, and poured into falcon tube
  • Weighed out 79.06 mg of ammonium bicarbonate (NH4HCO3) (0.0793g measured)
  • Added NH4HCO3 to falcon tube, vortexed until mixed
  • Weighed out 7.21g Urea (7.21g measured)
  • Added Urea to falcon tube, vortexed until mixed
  • Poured falcon tube contents into graduated cylinder
  • Topped of contents in graduated cylinder with nanopure water up to 20 mL
  • Poured contents of graduated cylinder into falcon tube

Checked amount of reagents in freezer

I double checked that we had at least 10 aliquots of DTT and IAA, and 2 aliquots of TCEP left in the freezer from Rhonda. All’s good to go for sonication tomorrow!

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