The following are collection and transplant details for Porites astreoides specimens collected during my November 2015 Belize trip.

I arrived at Carrie Bow Cay late in the day on 11/11 and quickly got to work setting up an experiment attempting inhibition of DNA methylation in corals using 5-aza-2′-deoxycytidine, also known as Decitabine. Once incorporated into DNA, this cytosine analog inhibits de novo methylation. The goal is to identify a causative role of DNA methylation in coral thermal tolerance plasticity. If corals treated with Decitabine do not achieve the same level of thermal tolerance plasticity as untreated corals, this would indicate that DNA methylation may play a role in thermal tolerance plasticity (or buffering, as may be the proper term).

For the experiment, I chose the branching coral Porites porites because branch tips (“nubbins”) can be easily and cleanly cut from colonies and glued to a substrate of choice. I cut four replicate nubbins from four different colonies in the backreef (1-2 m depth) at Carrie Bow Cay. I glued the nubbins to small petri dish lids with A-788 compound, then buoyant weighed and photographed them. This morning, I placed the nubbins in their respective treatments. There are two water baths, one filled with flow-through ambient seawater, and the other with a much smaller amount of flow through water, plus a heater and a water pump for mixing. Thus the experiment is a factorial arrangement with treatments ambient (A), heated (H), Decitabine (D), and control (C).

A stock solution of Decitabine (FW = 228.21) at a concentration of 219 uM was made by dissolving 10 mg of the compound in 200 ml filtered seawater. This stock solution is kept refrigerated and I anticipate making a new stock approximately every other day. For a final concentration of 10 uM in the experimental beakers (based on the concentration used by Amarasinghe et al. 2014 in bumble bee apiary syrup), I added 3.6 ml of the stock into beakers filled with 80 ml seawater. I will do water changes twice daily.

I placed the corals in the experimental beakers today at 9:30 AM, and added Decitabine at around 10:30. The beakers were covered with a glass plate to reduce evaporation and rain water entry. This will also attenuate some light which is fine since the experiment is outdoors.

Here is a photo of the setup, with ambient on the left and heated on the right, plus a closeup of a beaker with coral nubbin.

Temperature record

Yesterday and today I made a salt saturated DMSO solution for coral DNA preservation in the field. I found support for this solution as a preservative for corals based on Gaither et al. 2011 and Concepcion et al. 2014. There is also an older article by Seutin et al. 1991 that appears to have initially promoted this solution for animal DNA preservation.

On the web I found this protocol for making salt saturated DMSO, and this is what I followed.

First I made 1L 0.5M EDTA:

900 ml nanopure H20
EDTA (FW 292.24) 146.12 g
NaOH pellets to pH EDTA to 8.0
Brought final volume to 1L

I ended up adding LOTS of NaOH pellets to bring the pH up since I did not have disodium EDTA. This took a few hours.

Then for the salt saturated DMSO:

0.5M disodium EDTA 1L
Dimethyl Sulfoxide (DMSO) 400mL
Nanopure water 500mL
NaCl – Enough to saturate the solution.

First I added 300 g NaCl, then another 100 g , then another 20 g. I brought the volume up to 2 L with nanopure water. Some solid NaCl remained, which was left out when I transferred the solution to a container. I then autoclaved the solution.

Continuing the extraction from yesterday, samples were centrifuged at 4,000 x g for 1.5 min to pellet DNA. The ethanol was aspirated and the samples were air dried under the hood for about an hour (they were watched to ensure they were not over dried). DNA was resuspended in 0.2 ml nanopure water.

Here is the electrophoresis protocol I used:

1. 75 ml TAE with 0.65 g agarose was microwaved for 1 min (with cap resting on 100 ml pyrex jar, watching for boil over) to heat and facilitate dissolution of the agarose. The jar was swirled with cap on. Solution was re-heated for a few more seconds to further facilitate particulate dissolution.

2. Solution left to cool without lid for 1-2 min

3. 7.5 ul EtBr added and swirled to mix

4. Solution left to cool for another minute before slowly pouring into the gel tray. The gel tray included an 8 slot comb, and the gel solution was poured in the opposite end from the comb.

5. Gel was left for 30 min to cool and harden.

6. Comb was removed and gel tray was rotated 90 degrees to align with electrodes.

7. Gel chamber filled with enough 1x TAE to just cover gel (~1 mm depth over gel).

8. Gel loaded with 5 ul green ladder (GeneRuler 100 bp) on each end. 20 ul of each sample was mixed with 3 ul purple loading dye on a sheet of parafilm and loaded into gel.

9. Electrodes plugged in and power supply set to 120 V.

10. Gel was run for 55 min, when ladder reference dye was ~1cm from edge of gel.

There were no bands present for either sample. Perhaps the DNA was degraded; the samples were collected 5 months ago. I did notice that both DNA pellets were quite brown in color.

Today I extracted DNA from the Porites astreoides samples I collected back in May. The samples consisted of a single coral colony that I relocated from the lagoon to the backreef, and I sampled the coral on the day of transplantion (t = 0) and three days later (t = 3). The samples were kept under refrigeration in ethanol. I extracted the DNA using DNAzol.

The DNAzol protocol was as follows:

1. The surface layer of each coral core was scraped with a razor blade and the tissue/skeletal material was homogenized with a sterile plastic pestle in a microfuge tube with 0.5 ml DNAzol.

2. The homogenate was sedimented by centrifugation at 10,000 x g for 10 min. The supernatant was then transferred to a new tube.

3. DNA was precipitated by the addition of 0.25 ml 100% ethanol. Samples were mixed by inversion and I was able to spool the DNA and transfer to a new tube.

4. DNA was washed twice with 1 ml 75% ethanol. A quick spin was used each time to pellet the DNA before removing the supernatant.

5. Samples were stored under refrigeration in 95% ethanol.

Well, it’s the start of another quarter here, so what better time to jot down some goals for the next month? I head to Belize in November for some field work, so my primary October goal is to prepare for this trip. This includes having a solid list of tasks to make the best of my time there, which I hope to have my committee review for any feedback they might have. This month I would also like to take a stab at RADseq, and possibly EpiRADseq, two methods I hope to employ to characterize the population genetics and epigenetics of Porites astreoides in Belize.

Two days ago I submitted a revised manuscript titled “Germline DNA methylation in reef corals: patterns and potential roles in response to environmental change” to Molecular Ecology. This manuscript has taken up much of my time for the past year or so, but I have not posted anything about it until now. This is because I was using a GitHub repository to document all of my work. For the revision, Steven suggested that we create a newer, cleaner version of this working repository. It can be found here.

Yesterday 7/13/15 I extracted RNA from the two Porites astreoides samples I collected in Belize (see previous post Belize pilot experiments). I used RNAzol RT and followed the total RNA protocol. The 25 mm coral cores were removed from their tubes of RNAlater and placed in a porcelain mortar that had been chilled in the -80C freezer for about 20-30 min before starting (note: this worked well, but the mortar and pestle were not chilled by the time I homogenized the second sample). I scraped off the top layer of tissue and skeleton using a razor blade, removed the rest of the core, and ground the tissue and skeleton in 1 ml RNAzol with a pestle. I did not do the optional phase separation step (#4). I left the RNA pellets in 75% ethanol in the refrigerator. The remaining homogenate in RNAzol was placed in the -80C freezer.

My big goal for this month is to complete and submit the coral CpG ratio manuscript. All sections of the manuscript need some work, but the Discussion has not been started, so that is going to be where most of the effort goes. The manuscript has a submission deadline of July 1st, so I’ve got my work cut out for me.

Also, I need to process the samples I collected in Belize. The RNAlater samples take precedence and ideally the RNA should be extracted by next week, which will be one month from collection.

Despite a very busy sampling schedule for the South Water Cay Marine Reserve biannual monitoring program, I was able to fit a little bit of my own work in at Carrie Bow Cay. On 5/17/2015, we had some down time due to some very rough seas that kept us off the forereef, so I took the opportunity to snorkel in the lagoon and collect a small Porites astreoides colony. My primary goal was to sample some tissue in order to assess my ability to obtain high quality DNA and RNA simply by fixation in ethanol and RNAlater. I was also interested in assessing the efficacy of my new corer tool, and the survival of a colony following transplantation and sampling.

The colony was approximately 25 cm2 and was located at approximately 3-4 m depth. It was the green morph of P. astreoides. I collected the colony and moved it to the backreef in front of CBC. I positioned the colony just east of the edge of the Thalassia bed in approx. 1 m depth. I did not epoxy the colony in place, opting instead to simply buttress it between pieces of coral rubble. I then used my new Elora 25 mm diameter corer to collect two cores from the colony. A shout out to Collin for helping me locate one of these hard-to-find tools on the internet. If found the corer to do a fairly decent job of extracting a piece of the colony, but not without some additional damage to the colony around the periphery of the corer. However, I do think using the corer is a better option than a hammer and chisel. I brought the cores back to the lab and fixed one core each in 95% ethanol and RNAlater within about 15 min of collection. I then stored the samples in the refrigerator.

Three days later, on 5/20/2015, I resampled the colony and again collected two cores for fixation in ethanol and RNAlater, respectively. Again, I was able to fix the cores within about 15 min of collection. The only difference this time was that the RNAlater I used was obtained on the island in the chemical storage room. I have no idea how old the reagent was; it was not dated.

I was also able to collect two colonies from the forereef on 5/19/2015, with a goal of transplanting them in the backreef to assess their survival the next time I’m on the island. I collected one colony from 10 m depth near the SWCMR transect at CBC-S-30, and another from 7 m depth near the CBC-S-15 transect. The deeper colony was about 25 cm2 and was the brown morph, while the shallower colony was also about 25 cm2 but the green morph. These colonies spent the night in buckets with flow through seawater until I was able to transplant them in the backreef the following morning. I placed the colonies adjacent to the earlier one, and also drove in a piece of rebar with flagging tape so I can relocate them in the future. The deeper, brown morph was placed behind, or west, of the rebar stake, while the shallower, green morph was placed east of the stake. Again, they were not glued, simply buttressed by rubble. I hope they survive and stay put!

Fig. 1. Colonies transplanted from the forereef to the backreef at CBC. (Left) Green morph of P. astreoides from 7 m depth. (Right) Brown morph of P. astreoides from 10 m depth.