Tag Archives: Hyperladder I

PCR – RLOv In-situ Hybridization (ISH) Probes

Ran probe-labeling PCRs to use in in-situ hybridization (ISH) using the PCR DIG Probe Sysnthesis Kit (Roche). Generated PCR probes for using the following BamHI-linearized plasmids:

  • pCR2.1/RLOv_membrane_gene_1
  • pCR2.1/RLOv_membrane_gene_1
  • pCR2.1/RLOv_tail_fiber

The Roche protocol recommends using only 10pg of plasmid DNA for probe labelling. As such, all three probes were diluted 1:10,000. A 1:1000 (999μL H2O + 1μL of plasmid) was made first. Then a 1:10 dilution was made (90μL H2O + 10μL from 1:1000 dilution of plasmid).

Additionally, I ran half reactions to conserve kit components. Roche recommends 50μL reactions; I ran 25μL and scaled all components appropriately.

All reactions were set up on ice and run in 0.2mL strip-cap PCR tubes.

Reaction calculations are here (Google Sheet): 20151109 – RLOv ISH Probe PCRs

Cycling params:

  1. 95C – 5mins
  2. 95C – 15s
  3. 55C – 15s
  4. 72C – 30s
  5. Go to Step 2, repeat 39 times.
  6. 72C – 10mins

After the PCR, 5μL of each reaction was run on a gel.

Results:

Hyperladder I (Bioline)

PCR DIG probe labelling products run on 1.1% agarose 1x TBE gel stained w/EtBr. A ‘+’ indicates DIG reaction, while a ‘-‘ indicates no DIG in reaction.

Two reactions were run for each plasmid: one with the DIG label (indicated by a ‘+’) and one without (indicated by a ‘-‘). If the labeling was successful, the PCR products from those reactions containing DIG will be larger (i.e. migrate slower) than those without. That is exactly what we see in each of the three potential ISH targets.

So, we now have three ISH probes ready for action! Will proceed with making fresh ISH buffers and ISH.

Probes were transferred to 0.5mL snap cap tubes and stored in my -20C box.

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Restriction Digestions – pCR2.1/RLOv Plasmids

Set up restriction digestions to linearize the pCR2.1/RLOv plasmids in preparation for ISH probes and qPCR standard curves. Used BamHI (NEB), since it doesn’t cut in any of the RLOv sequences and cuts one time in pCR2.1-TOPO (Invitrogen).

PLASMID Vol for 1.5μg (μL) H2O to 40μL
pCR2.1/RLOv_DNA_helicase 21.4 18.6
pCR2.1/RLOv_head_to_tail 11.1 28.9
pCR2.1/RLOv_membrane_gene_1 12.2 27.8
pCR2.1/RLOv_membrane_gene_2 14.3 25.7
pCR2.1/RLOv_tail_fiber 20 20

 

Digestion Master Mix

REAGENT SINGLE REACTION (μL) x 5.5 (μL)
Plasmid 40 NA
10x Buffer 3.1 (NEB) 5 27.5
BamHI (NEB) 1 5.5
H2O 4 22
TOTAL 50 Add 10μL to each tube

Digests were incubated at 37C for 1hr in PTC-200 thermal cycler (MJ Research); no heated lid.

Ran 3μL of undigested plasmid and 10μL of linearized plasmid on 0.8% agarose 1x TBE gel stained w/EtBR.

Results:

Hyperladder I (Bioline)

U = Undigested; Bam = BamHI digest

Besides the funky way this gel ran, the digests look to be complete.

Will quantify remaining linearized plasmids with a dye-based method for accurate quantification and then proceed with the making ISH probes (membrane genes and tail fiber gene) or qPCR standard curves (DNA helicase and head-to-tail).

 

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PCR – New C. gigas COX Primers for Sequencing of Isoforms

Used new primers for obtaining bands for additional sequencing of both COX isoforms in C. gigas. Master mix calcs are here. Master mix shorthand (MM##) is described below:

MM07 – Cg_COX_416_F (SR ID: 1193) + Cg_COX1_qPCR_R (SR ID: 1191) Expected band size (if no intron) = ~1540bp

MM08 – Cg_COX_416_F (SR ID: 1193) + Cg_COX2_454align1_R (SR ID: 1190) Expected band size (if no intron) = ~1540bp

MM09 – Cg_COX1/2_qPCR_F (SR ID: 1192) + Cg_COX1_qPCR_R (SR ID: 1191) Expected band size (if no intron) = ~225bp

MM10 – Cg_COX1/2_qPCR_F (SR ID: 1192) + Cg_COX2_454align1_R (SR ID: 1190) Expected band size (if no intron) = ~225bp

MM11 – Cg_COX_1519_F (SR ID: 1146) + Cg_COX2_454align1_R (SR ID: 1190) Expected band size (if no intron) = ~275bp

MM12 – Cg_COX_982_F (SR ID: 1151) + Cg_COX2_454align1_R (SR ID: 1190) Expected band size (if no intron) = ~812bp

Results:

Ladder is Hyperladder I from Bioline.

Master mixes are indicated underneath each group by the labels MM##. The order within each MM group (from left to right) is: template, NTC, NTC.

All bands boxed with green were purified using Millipore’s Ultrafree-DA spin columns. Samples were stored @ -20C in “Sam’s Misc. -20C Box”.

MM07 – Fails to produce any bands of any size. Suggests the presence of intron(s) causing the size of the potential amplicon to exceed the capabilities of the polymerase under these cycling conditions.

MM08 – Produces a band of ~400bp which is well below the expected 1540bp (if no introns) size. Due to the faintness of the band, the band was not excised. Will consult with Steven to see if he thinks it worth repeating to produce sufficient product for sequencing.

MM09 – Produce a ~500bp band. The band was excised. This band size is ~275bp larger than the expected size of 225bp. This implies the presence of an intron in this region. This band size differs from that produced by MM10, which suggests that this primer set can be used for qPCR AND distinguish between the COX1 and COX2 isoforms.

MM10 – Produced a ~700bp band. The band was excised. This band size is ~475bp larger than the expected size of 225bp. This implies the presence of an intron in this region. This band size differs from that produced by MM09, which suggests that this primer set can be used for qPCR AND distinguish between the COX1 and COX2 isoforms.

MM11 – Produced multiple bands, of which two were excised; a ~3000bp band and a ~600bp band. These bands were excised solely based on their intensity and their immediate useability for sequencing. Will discuss with Steven on whether or not this should be repeated and the other bands excised for sequencing purposes. Both bands that were excised exceed the expected band size of ~275bp, suggesting the presence of multiple introns. Additionally, the presence of so many products suggests that the primers are not very specific, in regards to their target.

MM12 – An extremely faint band of ~350bp can be seen, however, due to it’s faintness and it’s small size (expected size was ~812bp), the band was not excised. Will discuss with Steven to see if this warrants repeating to accumulate sufficient product for sequencing purposes. No amplification of any larger products suggests the presence of introns, causing the size of the potential amplicon to exceed the capabilities of the polymerase under these cycling conditions. This is also confirmed by the MM11 PCR results in which a 3000bp band was produced. Since the primer set in MM12 has an additional 600bp at the 5′ end, this has already exceeded the abilities of the polymerase, even if this addtional 600bp does NOT include an additional intron. However, it is curious that the MM12 primer set does not produce smaller, spurious PCR products as is seen in the MM11 primer set (these two primer sets both use the same forward primer).

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Genomic PCR – C.gigas cyclooxygenase (COX) genomic sequence

Attempt to obtain full genomic sequence for C.gigas COX. PCR set up/cycling params/etc are here. Primer set combinations(master mixes) are as follows:

MM01 – Cg_COX_5’UTR_3_F (SR ID: 1150) + Cg_COX_1009_R (SR ID: 1147) Band size w/o intron = ~1000bp

MM02 – “” + Cg_COX_1545_R (SR ID: 1148) Band size w/o intron = ~1540bp

MM03 – “” + Cg_COX_2138_R (SR ID: 1149) Band size w/o intron = ~2135bp

MM04 – Cg_COX_982_F (SR ID: 1151) + Cg_COX_1545_R (SR ID: 1148) Band size w/o intron = ~550bp

MM05 – “” + Cg_COX_2138_R (SR ID: 1149) Band size w/o intron = ~1130bp

MM06 – Cg_COX_1519_F (SR ID: 1146) + Cg_COX_2138_R (SR ID: 1149) Band size w/o intron = ~620bp

Results:

Bioline Hyperladder I used for marker. Gel is loaded with template samples at the far left of each master mix group with two no template controls (NTC) in the remaining two wells of each master mix group. All NTCs on the gel are clean.

All bands surrounded by a green box were excised from the gel.

MM01, MM02 and MM03 – The smallest expected band (i.e. no intron present) would have been 1000bp in MM01. Instead, we see faint banding of multiple sizes less than 1000bp in both MM01 and MM02. MM03 fails to produce any bands. This potentially suggests a couple of things. Firstly, the multiple banding produced in MM01 and MM02 suggests that the PCR conditions lead to some non-specific priming and should be optimized. Secondly, the fact that no bands were produced that are equal to or larger than the “no intron size” suggests that intron(s) may exist in the 5′ region of the COX gene and are large enough that the polymerase had insufficient time/ability to amplify.

MM04 – Three distinct bands were produced: 2000bp, 1500bp and 550bp. The size of band that would have been produced had an intron NOT been present would have been ~550bp. A band of this size was produced in this PCR reaction. However, two additional bands were produced. The presence of these two larger bands lends additional evidence for the existence of multiple isoforms of COX (which is also supported by the fact that multiple isoforms of COX are known to exist in most other species). The 2000bp band was excised and purified with Millipore Ultra-free DA spin columns and stored @ -20C until a sequencing plate is readied.

MM05 – A distinct band of ~5000bp was produced. This is significantly larger than the “no intron size” of ~1130bp, suggesting the presence of an intron. This band was excised, but not purified due to the low concentration of DNA in the gel. The gel slice was stored @ -20C until this PCR reaction could be repeated to accumulate sufficient product for sequencing.

MM06 – A distinct band of ~2200bp was produced. This is significantly larger than the “no intron size” of ~620bp, suggesting the presence of an intron. The band was excised and purified with Millipore Ultra-free DA spin columns and stored @ -20C until a sequencing plate is readied.

The PCR reactions reveal the presence of intron(s) in the COX gene we’re investigating as well as providing evidence for the existence of multiple isoforms in C.gigas. Since the PCR products that have been excised for sequencing are so large, additional primers will need to be designed closer to the introns in order to generate smaller products that can be fully sequenced. Additionally, all reactions using the primer designed to anneal in the 5’UTR of COX (SR ID: 1150) failed to produce useful results. This is either due to poor performance of the primer under these reaction conditions or due to the presence of a large intron in the 5′ region of the gene. Additional reverse primers will be designed that anneal closer to the 5′ portion of the COX gene in hopes of characterizing the 5′ genomic sequence better.

After speaking with Steven today about the potential existence/”discovery” of multiple isoforms, he decided to map the newly-released C.gigas 454 NGS data to the existing COX coding sequence in GenBank (FJ375303). The alignment is shown below.

The two 454 reads that map closest to the 5′ end of the COX coding sequence match up nearly perfectly, with periodic SNPs. The remaining 454 reads that map to the COX coding sequence are very different and provide very good evidence of a previously unidentified isoform of COX in C.gigas. Primers will be designed from both the existing COX sequence in GenBank (FJ375303) and the other potential isoform. These primers will likely be used in both qPCR and for sequencing purposes, in order to be able to distinguish and characterize both isoforms. Additionally, BLASTing will be performed with the sequences from both isoforms to evaluate how they match up with existing COX isoforms in other species.

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gDNA Sonication – SB/WB gDNA pools (prep for MeDIP) from 20100618

The previous attempt at sonication (see 20100618) failed, likely due to no using the correct equipment (tubes and Covaris adapter). The two gDNA pools, which had previously been unsuccessfully fragmented on 20100618 (SB and WB) were sonicated using a Covaris S2. Used the guidelines of the manufacturer (listed below) for shearing gDNA to a desired target size (500bp):

Duty Cycle: 5%

Intensity: 3

Cycels per Burst: 200

Time (seconds): 90

Temp (water bath): 4C

Power Mode: Frequency Sweeping

Sample Volume: 120uL

Buffer: TE

DNA Mass: ~8ug

Starting Material: >50kb

AFA Intensifier tubes and associated Covaris adapter.

After shearing, ran 250ng of each pool on a 2% TAE agarose gel for fragmentation verification.

Results:

Lane 1 – Hyperladder I

Lane 2 – R37

Lane 3 – R51

Looking at this gel, the samples have been successfully fragmented and I would estimate have generated and average fragment size of ~400bp (going from bottom to top of the Hyperladder: 200bp, 400bp, 600bp, 800bp, 1000bp). So, this looks great! Can proceed with remainder of MeDIP procedure at any time.

Additionally, I will confirm a more accurate assessment of average fragment size by running these two samples on the Agilent Bioanalyzer.

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gDNA Sonication – SB/WB gDNA pools (prep for MeDIP) from earlier today

The two gDNA pools (SB and WB) were sonicated using a Covaris S2. Used the guidelines of the manufacturer (listed below) for shearing gDNA to a desired target size (500bp):

Duty Cycle: 5%

Intensity: 3

Cycels per Burst: 200

Time (seconds): 90

Temp (water bath): 4C

Power Mode: Frequency Sweeping

Sample Volume: 120uL

Buffer: TE

DNA Mass: ~8ug

Starting Material: >50kb

To be noted, the Covaris guidelines list the use of an “AFA Intensifier” tube, which I did not use (because we don’t have them).

After shearing, ran 250ng of each pool on a 2% TAE agarose gel for fragmentation verification. Also ran 250ng of pre-sonication DNA from each pool as controls.

Results:

Lane 1 – Hyperladder I

Lane 2 – R37, Un-sonicated

Lane 3 – R37, sonicated

Lane 4 – R51, Un-sonicated

Lane 5 – R51, sonicated

Sonication with the Covaris did NOT produce the desired fragmentation (500bp smear) in either sample, although the R37 sonicated samples shows a significantly greater degree of fragmentation than the R51 sonicated sample. Not sure how to explain this difference, other than the R51 sample has a greater amount of DNA. Additionally, the results could be explained by the fact that we did not use the AFA Intensifier listed in the Covaris guidelines…

Am consulting with a person in Genome Sciences who has used a Covaris for DNA fragmentation in the past to see if the AFA Intensifiers are indeed necessary and, if so, we can use two of them. Hopefully have an answer soon and be able to proceed with additional fragmentation next week.

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gDNA Isolation – Mac gigas gill samples (continued from yesterday)

Continued with gDNA isolation from yesterday’s samples. Additionally, isolated gDNA from R51 01, but homogenized the tissue (using disposable 1.5mL mortar/pestle) in 0.5mL of DNAzol and topped off to 1.0mL. All 3 samples were gently pipetted up and down to further dissolve the tissue. For those samples that were subjected to Proteinase K digestion, I transferred 100uL of the solution to a new tube containing 1mL of DNAzol, as described in the DNAzol protocol (see “Notes, #5″ part of protocol). Tubes were incubated 10mins @ RT.

Pelleted residual tissue 10mins @ 10,000g @ 4C. Transferred supe to new tubes. Precipitated DNA with 0.5mL 100% EtOH. Incubated 3mins @ RT. DNA was pelleted 5mins @ 5000g @ 4C. Supe was removed, pellets were washed with 1mL 75% EtOH (x2). Supe was fully removed and the pellets were resuspended in 8mM NaOH (made by Amanda Davis 5/20/10). See below for volumes:

R51 11 – 50uL

R51 01 – 100uL

R51 01 homogenized – 200uL

1M HEPES (provided with DNAzol) was added at a 1:100 dilution to achieve a pH = 8.0. This was based on the DNAzol protocol calculations (For 1mL of 8mM NaOH, use 101uL of 0.1M HEPES).

Samples were spec’d on NanoDrop 1000. Used a sample with 8mM NaOH and 1M HEPES to match the pH = 8.0 of the samples.

Results:

The 260/280 ratios for all samples are great. Yields are significantly lower than I was expecting. However, for R51 11 and R51 01, only 100uL (1/5th) of the digestion was used as described in the protocol. I may recover the remainder of the R51 11 gDNA since remaining tissue could be a limiting factor. The homogenized sample had the highest yield (9.35ug), suggesting this may be a more efficient approach to obtaining gDNA (faster procedure and higher yields).

These results also demonstrate that gDNA can be successfully isolated from samples stored in RNA Later (gel pending).

Due to low yields of the R51 11 and R51 01 samples, these will not be run on a gel until the rest of the DNA is isolated.

Isolated the remainder of the gDNA from R51 11 and R51 01. Added an additional 0.5mL of DNAzol to each tube and pipetted up and down to further dissolve the remaining tissue. Then, proceeded as described above. Samples were resuspended in 200uL of 8mM NaOH with 2.02uL of 1M HEPES added.

Results:

Both samples have great yields and excellent 260/280 ratios. Digestion, combined with gentle pipetting to disrupt undigested tissue, appears to lead to higher yields. However, the process is more time consuming than just basic, physical homogenization of tissue.

Gel Loading (0.5ug of each DNA was loaded in a volume of 25uL)

Lane 1. Hyperladder

Lane 2. R51 11

Lane 3. R51 01

Lane 4. R51 01 homogenized

The R51 11 sample looks perfect. The other two samples however, appear degraded, with the homogenized sample looking the worst. Interestingly, those are both from the same source tissue, possibly suggesting that the DNA in this particular gill sample is actually degraded and is independent of the preparation, since the R51 11 (Lane 2) and R51 01 (Lane 3) were prepared simultaneously and in the same way. The gel also seems to show that physically homogenizing the sample leads to greater degradation than performing the Proteinase K digestion.

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DNA Isolation – Qiagen Kit Comparison

Note: This information was added 20140407. Yes, you read that correctly.

Someone had noticed that gDNA isolated using a Qiagen DNeasy Blood & Tissue Kit we rec’d in April 2010 seemed to be yielding degraded DNA.

The two samples used for the comparison were a single tail (split in two equal weight pieces) from a juvenile salmon that was snap frozen, without preservatives, at the time of its collection. The samples were prepped. 0.5ug of eluted DNA was then run on a 1.2% agarose-TAE gel containing 0.1ug/mL of ethidium bromide. 5uL of Bioline’s Hyperladder I was loaded for size assessment (see link for marker layout).

Results:

Clearly, there’s significant quality difference! A free replacement kit was sent by Qiagen.

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