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.


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.

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

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.


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).


Colony PCRs – Clam RLO 16s, EHR, EUB

Colony PCRs were performed on each of the three transformations from yesterday (16s, EHR, and EUB primers) using the M13F/R vector primers. Colonies were picked form the transformation plates with pipette tips, re-streaked on a secondary, gridded, numbered LBAmp50+x-gal plate and then used to inoculate the respective PCR reactions. Six white colonies (positive clones) and a single blue colony (negative clone) were selected from each transformation.

Restreaked plates were incubated @ 37C O/N and then stored @ 4C (Parafilmed).

Master mix calcs are here: 20150227 – Colony PCR Clam RLO

30μL of each reaction was run on a 1% agarose 1x Low TAE gel, stained w/EtBr.


Ladder: Hyperladder I (Bioline)

Upper Left: 16s colonies 1 – 7

Upper Right: EHR colonies 1 – 6

Lower Left: EUB colonies 1 – 7

Based on the PCRs used for cloning, all white colonies screened exhibit the expected product sizes. Additionally, each of the blue (negative) colonies, produced the expected band size that are indicative of an empty plasmid.

Will select a positive colony from each set for mini prep and Sanger sequencing.

PCR – Ireland Clam RLO DNA S/6/14 #19

This is an exact repeat of the PCR from yesterday, but with a brand new vial of Apex Red Master Mix, in an attempt to eliminate the contamination previously seen in the NTCs.


Ladder: Hyperladder I (Bioline)

Well, for some reason there are still bands in the NTCs. However, they appear to be of different sizes than the bands in the clam DNA samples. I think they’re OK to use and the cloning/sequencing is cheap enough these days, that I’ll just get these sequenced and see what we have.

I excised each of the bands in the clam DNA samples (16s = ~2000bp; EUB = ~2100bp) and purified them using Ultrafree-DA spin columns (Millipore) in preparation for cloning.

PCR – Ireland Clam RLO DNA S/6/14 #19 from 20150130

After the last PCR continued to exhibit products in the no template controls (NTC) for most of the primer sets I was using, I ordered new primers. They arrived today so, I re-ran the PCR on the clam RLO DNA isolated 20150130 with the following new primers:

Master mix calcs are here: 20150223 – cPCR Universal Primers Apex Red MM

Cycling params were:
1 cycle of:

  • 95C – 10mins

40 cycles of:

  • 95C – 15s
  • 50C – 15s
  • 72C – 1min

Samples were run on 1.0% agarose, low TAE gel stained w/EtBr.


Ladder: Hyperladder I (Bioline).

Crazy; contamination still present in the NTCs. Primer stocks were steriley reconstituted with Low TE Buffer (IDT) and working stocks were created steriley, so I’m not really sure why this is continuing to happen. Possibly the polymerase is contaminated?  Will try again with previously unopened polymerase and see how that plays out.

No bands were excised since I can’t be certain that the bands present in the Clam DNA samples are from the actual sample and not from the apparent contamination.

PCR – Universal Primers w/New Master Mix

Since the previous check of the various universal primers with abalone DNA (sample 09:8-20) failed to amplify, even with withering syndrome primers, I’m testing repeating that PCR using a newer/different PCR master mix.

Template DNA is: 09:20-08 (from tissue)

Background info for template DNA is here: Red/Pink/Pinto

Primers being used are:

  • 16s/23s-F/R
  • 27F, 1492R
  • EHR16D, EHR16R (universal ehrlichia)
  • EUB-A/B
  • 18s EUK 581 F, 18s EUK 1134 R
  • WSN1 (withering syndrome)

Master mix calcs are here: 20150212 – cPCR Universal Primers 09:8-20 Apex Red MM

All samples were run in duplicate.

Cycling params were:
1 cycle of:

  • 95C – 10mins

40 cycles of:

  • 95C – 15s
  • 50C – 15s
  • 72C – 2mins

Ran samples out on a 0.8% agarose,  1x TBE gel w/EtBr


Well, this is a good result.  It demonstrates that the previous reagents that I had been using are no good. The primers work.  However, it does appear that all of the universal primers (excluding the 18s and EHR) are contaminated.  All of these primer sets were stocks that were prepared by other people and none of them were marked as being sterile (which they should be).  Regardless, I’ll re-run the Ireland clam DNA with all the primer sets and see how it turns out.  In the meantime, I’ll also order new universal primer sets to replace the existing, non-sterile sets.

PCR – Ireland Clam DNA 18s

Since the previous PCR didn’t amplify anything using four different universal 16s (prokaryote) primers, I am testing to verify that the two extractions (QIAamp Fast DNA Stool Mini Kit & the DNeasy Kit; Qiagen) actually isolated any amplifiable DNA using universal 18s (eukaryote) primers.

First, quantified the samples to verify that DNA actually exists in these two samples:


Sample Concentration (ng/uL)
Clam DNeasy Kit 4.432
Clam Stool Kit 6.184

The yields are surprisingly low, particularly for the DNeasy Kit sample.  In a total elution volume of 200uL, that means I only extracted 800ng…

Due to low DNA concentrations, I used 10uL of each sample in the PCRs.

Master mix calcs are here: 20150129 – cPCR Clam Universal 18s

Samples were run in duplicate.

Cycling params:

  • 1 cycle of 10mins
  • 40 cycles of:
    95C – 15s
    50C – 15s
    72C – 2mins


Ladder used was Hyperladder I (Bioline).

Neither sample produced any amplification.  The blurry “bands” that correspond to ~100bp are likely RNA carryover from the DNA isolation procedure.  They are not the amplicon we are looking for.  Additionally, they are not primer dimers, as these “bands” do not appear in the NTC.

I believe there is a small quantity of tissue debris in the original EtOH sample tube.  I will attempt to isolate some DNA from this debris and will repeat both the 16s and 18s PCRs.

Restriction Digestion – Withering Syndrome Phage ORF25 Plasmid from 20140926

Performed restriction digest using NcoI (NEB) to linearize plasmid for use as qPCR standard curve. Reactions were run for 1hr @ 37C and then heat inactivated @ 65C for 20mins.

Each reaction contained:

pCR2.1/Phage ORF25 (~1ug) – 17uL

10x Buffer 3 – 5uL

NcoI – 1uL

H2O – 27uL

Total: 50uL

After inactivation, 5uL from the reaction was run on a 1% TBE gel to confirm digestion. 5uL of undigested plasmid was run along side the digest.


Gel Loading Guide:

Lane 1 – Hyperladder I (Bioline)

Lane 2 – pCR2.1/ORF25 (undigested)

Lane 3 – pCR2.1/ORF25 (NcoI)

Vector size (bp): 3929

Insert size (bp): 483

Total size (bp): 4412

The linearized plasmid (which contains a single NcoI recognition site) runs between the 5000 and 4000bp standards, which is what we expect. Will quantify and generate dilution series for use as a qPCR standard curve.

Restriction Digests – Withering Syndrome Clone Plasmids from 20120718

Performed restriction digest on all four clones using NcoI. Reactions were run for 1hr @ 37C and then heat inactivated @ 65C for 20mins.

Each reaction contained:

Plasmid – 5uL

10x Buffer – 5uL

NcoI – 1uL

H2O – 39uL

After inactivation, 5uL from each reaction were run on a 1% TBE gel to confirm digestion. 1uL of undigested plasmid was run along side the corresponding digest.

The four clones are referred to as:

  • pWC8
  • p16RK3
  • p16RK7
  • p18RK7


Gel Loading (left to right):

Lane 1 – Hyperladder I (Bioline)

Lane 2 – pWC8 (Und.)

Lane 3 – pWC8 (NcoI)

Lane 4 – p16RK3 (Und.)

Lane 5 – p16RK3 (NcoI)

Lane 6 – p16RK7 (Und.)

Lane 7 – p16RK7 (NcoI

Lane 8 – p18RK7 (Und.)

Lane 9 – p18RK7 (NcoI)

Lane 10 – Hyperladder I (Bioline)

All digests are complete. All clones reveal the same restriction digestion pattern, producing two bands: ~2500bp and ~3000bp. The band sizes total ~5500bp, which is in line (5432bp) with the full withering syndrome 16s clone in the pCR2.1 TOPO vector (Invitrogen). Will quant and prepare a dilution series for qPCR.

PCR – Linearized pCR2.1/AF133090 Plasmid from 20120430

Ran PCRs using both the AF133090 full-length primers (WS_16s_1_F, WS_16s_1501_R) and the WSN primers (WSN1_F, WSN1_R). This time used the linearized pCR2.1/AF133090 plasmid as template instead of the bacterial colonies that were selected from cloning. Master mix calcs are here. Cycling params are as follows:

95C – 10m

40 cycles of:

  • 95C – 10s
  • 55C – 10s
  • 72C – 1.75m


Gel Loading:

Lane 1 – Hyperladder I (Bioline)

Lane 2 – 16s primers

Lane 3 – 16s primers NTC

Lane 4 – WSN primers

Lane 5 – WSN primers NTC

Essentially, this is the same result as the colony check from 20120503. The appropriate sized band (~1500bp) is generated using the 16s primers, but no band is generated using the WSN primers. Yes, there is contamination present in the 16s NTC sample, but this point is moot since the WSN primers still fail to generate a PCR product.

So, I have absolutely no idea what is cloned into this vector, despite the fact that primers designed on GenBank AF133090 produce a band.

I’m essentially running out of ideas on what else to do to get this working again. Will ask Lisa to make a curve from the linearized plasmid stock from DATE (not the pCR2.1/AF133090 plasmid that is being tested here) and see if she can get it to work. Also, Carolyn has suggested trying to run some of the more recent curves on a different qPCR machine. Will contact the guy in Health Sciences who has a Stratagene that we are allowed to use.